CHARACTERIZATION OF

FASCICLIN-LIKE -

PROTEINS IN

Edgar Zhipeng Liu

Submitted in total fulfilment of the requirements of the degree of

Doctor of Philosophy

December 2017

School of Biosciences

University of Melbourne

ABSTRACT

All cells are surrounded by an elastic primary and, in some specialised cell types/tissues such as the vasculature, the cell deposits a secondary wall that is capable of withstanding enormous compressive forces important for water conducting tissues and plant support (Doblin et al., 2010). The primary wall largely consists of polysaccharides such as cellulose, hemicelluloses and pectins and a minor portion of components. An abundant superfamily of plant-specific wall are the -rich (HRGPs), which includes the arabinogalactan- proteins (AGPs). AGPs are characterised by a protein backbone rich in non-contiguous (P) residues that direct O-linked with type II arabino-(3,6)- (AGs) (Schultz et al., 2002; Johnson et al., 2003; Basu et al., 2015a). AG glycans have been proposed to be important in secretion and trafficking of proteins, and in modulating their biological function (Ellis et al., 2010; Kovall and Blacklow, 2010). Within the AGP family, chimeric AGPs, with recognised PFAM domains other than the AGP regions, include an important class known as the fasciclin-like arabinogalactan-proteins (FLAs) (Schultz et al., 2002; Johnson et al., 2003). In FLAs, AGP regions are found adjacent to fasciclin (FAS) domains (Johnson et al., 2003). FAS domains are known to be conserved in proteins from a broad range of living organisms playing important biological roles related to adhesion (Elkins et al., 1990a; Johnson et al., 2003; Hamilton, 2008; Moody and Williamson, 2013). The combination of O-linked glycans and FAS domains is proposed to confer unique biological roles to FLAs. Bioinformatic studies have shown that FLAs belong to multigene families in all plant species, for example, 21 FLAs have been identified in the Arabidopsis genome (Johnson et al., 2003). The function of some FLAs has been inferred through investigation of their tissue-specific expression profiles and phenotypes of the respective fla mutants. Based on these studies, FLAs are proposed to play roles in cell expansion, plant growth and development (Shi et al., 2003; Seifert and Roberts, 2007; Li et al., 2010; MacMillan et al., 2010). In particular, a subset of FLAs specifically expressed in stems have been shown to influence stem biomechanical properties. MacMillan et al. (2010) demonstrated that loss of FLA11 and FLA12 resulted in changes in cell wall composition and the microfibril angle of cellulose in Arabidopsis stems. No obvious growth phenotypes were observed in fla11 fla12 mutants suggesting further I redundancy with other FLA members. FLA16 was also identified as having stem-specific expression yet its role in stem biomechanics was not explored. In this study, multi-disciplinary approaches, including molecular biology, microscopy, biochemistry, biomechanics and genetics, have been employed to investigate the biological function(s) and genetic relationships of FLA11 (AT5G03170), FLA12 (AT5G60490) and FLA16 (AT2G35860). We developed molecular resources to further explore the role of Arabidopsis FLA11 and FLA12 during stem development and characterise FLA16 as a novel regulator of stem development. Chapter 1 provides an overview of AGP/FLA structures, classification and known biological roles. Chapter 2 provides a detailed description of the experimental materials and research methods used in this thesis. This is followed by Chapter 3, an investigation of FLA11, FLA12 and FLA16 at the protein level. Fluorescent protein fusions of FLA11, FLA12 and FLA16, driven by either their endogenous promoters or the 35S promoter were generated and transformed into either Arabidopsis or tobacco (), respectively. The tissue/cellular distributions of the FLA proteins in different plant organs as well as their sub-cellular location was determined. In addition, several enrichment methods, including: chelation, immunoprecipitation (IP) and hydrophobic interaction chromatography (HIC), were explored in preparation for structural analyses and identification of potential interacting partners of the FLAs (see Chapter 3, Appendices A3.1). FLA11, FLA12 and FLA16 were shown to be predominantly expressed in stems, siliques and branches with lower abundance in cotyledons and . The location of FLAs was largely confined to cells with secondary walls, such as interfascicular fibres and cells in stems/branches, endocarp b and replum cells in the siliques, guard cells in the and vasculature in the main roots. Membrane fractionation studies suggest the FLAs are present at the plasma membrane (PM) and in the wall and are glycosylated. Enrichment of detergent-solubilised FLAs showed IP and chelation are the most effective methods, however, improved methods for enriching FLAs from Arabidopsis are needed given the difficulty of extracting proteins from stems due to the abundance of secondary walls. These data suggest that FLA11, FLA12 and FLA16 are likely to function at the PM and/or wall of cells with secondary walls in inflorescence stems. In Chapter 4, the function of FLA16 in stem development is explored through characterisation of a fla16 mutant at different growth stages and tissues. Phenotypic II analyses of the fla16 mutant shows a delay in growth during early seedling development, early bolting and reduced stem length and thickness. Morphological analysis of stems using light microscopy shows a reduced number of pith cells with no obvious changes in fibre and xylem cells. Cell wall compositional analyses of the stem showed that fla16 has reduced cellulose compared to wild-type (wt). Similar to FLA11 and FLA12, loss of FLA16 altered biomechanical properties of the stems, with increased tensile stiffness in the middle stems and decreased flexure strength in the basal stems. Quantitative RT-PCR (Q-PCR) was used to study the expression of primary and secondary wall-specific cellulose synthases (CesAs) in stems and showed changes of the secondary- wall active CesA7 occurs in fla16. Our results suggest FLA16 plays a role in maintaining biomechanical properties in the stem, likely via regulating cellulose in the secondary walls. In Chapter 5, a series of comparative and functional analyses are undertaken for FLA11, FLA12 and FLA16, based on analysis of the double fla11 fla12, fla11 fla16, fla12 fla16, and triple fla11 fla12 fla16 mutants compared to the respective single fla11, fla12 and fla16 mutants and wt. Through comparative analyses of growth parameters such as the length, thickness and cellulose content of the stems, we found all fla mutants show similar phenotypes, suggesting they act in the same genetic pathway. Q-PCR analyses of the expression of CesAs active in either primary or secondary wall cellulose synthesis show CesA7 expression is consistently increased in fla16, fla11 fla12 and fla11 fla12 fla16 mutant stems. Our results suggest FLA11, FLA12 and FLA16 likely act in similar pathways to regulate stem growth and cellulose biosynthesis during secondary wall formation, and the genetic relationship of these FLAs seems to be epistatic. In Chapter 6 we summarise the implications of these data on plant growth and development and speculate on the roles and properties of FLAs. We pose questions as to whether they function as wall biosynthesis regulators, architectural components, or both. This research provides valuable insights into the biology of FLA11, FLA12 and FLA16, and provides future directions in the study of these cell wall glycoproteins. Greater knowledge of how FLAs influence cellulose deposition and biomechanics will underpin future applications in the forest industry and the agri-food sector.

III

DECLARATION

This is to certify that

i. The thesis comprises only my original work, except where indicated to the contrary ii. Due acknowledgement has been made in the text to all other materials used iii. The thesis is less than 100,000 words in length, exclusive of tables, maps, bibliographies and appendix

Print Name: EDGAR ZHIPENG LIU

Signature:

Date: December 21, 2017

IV

ACKNOWLEDGEMENTS

This PhD would not be achieved without the generous support and warm- hearted assistance from many people whom I will never forget. I would like to give my first thanks to my supervisor Prof Tony Bacic. I was struggling in the intersection whether to continue pursuing my PhD in a new venture or to regretfully admit being beaten by the disadvantageous circumstance in my previous university. Tony encouraged me with his respect and care for students and showed me the scientist’s altitude in studying. He gave me his firm support for my PhD entry application, throughout my PhD course and most importantly he gave me two remarkable supervisors, Dr Kim Johnson and Dr John Humphries. I am always grateful to Kim who has spent a significant portion of her time on the supervision of my project. Kim lights me up on the path towards a PhD and guides me from an outsider to a qualified researcher. She is a passionate and knowledgeable scientist always refuels me when I was in lack of idea and experiencing fatigue in work with a will. I was touched that Kim corrected my works in late nights and even during her pregnancy, on playful weekends and gave me great understanding and mental support when I was down after a traumatic accident. I need to thank John who has been very helpful in my project and is always supportive when I experience difficulties no matter in studies or personal life. It was wonderful time I spent with John when we go out for drinks and foods. I am appreciated to John’s sacrifice of his weekend time correcting my works and giving me valuable opinions. I need to thank Ms Kristina Ford who is a skilful and helpful expert in proteomics, giving me very kind supervision and patient training in my project. I need to thank my course advisors, A/Prof Ed Newbigin and Dr Monika Doblin for their constructive input and valuable discussion in the guidance of my study. I need to thank the collaborators involved in my project. These brilliant people are Dr Colleen MacMillan, Prof Staffan Persson, Prof Shawn Mansfield, Dr John Golz and Dr Ingvil Austarheim. I am appreciated to their kind supports in offering technical supervision, research materials and scientific discussions. I need to thank A/Prof Joshua Hazaelwood for his informative advices in my study and funny jokes brought to crow. I need to thank the post-doctoral research fellows from multiple groups. They are Drs Wei Zeng, Edwin Lampugnani, Rene Schneider and Berit Ebert who have provided me great support on bench and at desk during my project. I must acknowledge I have received substantial V technical support from the great technicians Ms Cherie Terese Beahan and Mr Roshan Chetamum who provided excellent assistance in my analyses, and Ms Kelsey Picard and Jessica Jia Zhao for her kind assistance in the lab. I need to thank Mr Andrew Cassin and Andrew Lonsdale, the Andrew-Andrew, who assist me in my bioinformatics. I thank Ms Lisa Wittick is a very kind technician always allows me to borrow experimental apparatus from the Burney campus. I need to thank Ms Cassie Watt, Andrea Gallo, Ouda Khammy, Kirsty Turner and Mr Steve Elefteriadis who have been supporting me by providing a well-organised study and work environment. I need to thank Prof Paul Dupree who has been very generous to host my visit in the University of Cambridge and given me constructive opinion in my project. I certainly need to thank my previous and current colleagues, Drs Xingwen, Shane Emanuelle, Tony Chin and Joan Narciso and future Drs Yingying, Ting Guo, Yingxuan Ma and Melisa Bain for your companion during my study in the University of Melbourne. Last but not least, I truly appreciate to my family and my best friend Jimmy Tan for their endless support and to those haven’t been mentioned in the limited space but made contribution to my work. You made me!

VI

ABBREVIATIONS

abscisic acid ABA -Hydroxyproline-Alanine-Hydroxyproline AOAO alcohol-insoluble residue AIR Arabidopsis transcription activation factor ATAF Arabidopsis Biological Resource Centre ABRC Arabidopsis fragile fibre fra1 Arabidopsis Functional Genomic Consortium AFGC arabinogalactan-protein AGP ara BRI1-EMS-Suppressor BES by prolyl 4-hydroxylase P4H cell adhesion molecule CAM cellulose synthase CesA cellulose synthase complex CSC chitinase-like CTL Columbia Col cup-shaped cotyledon CUC cyan fluorescent protein CFP day old d.o. ER fasciclin FAS fasciclin-like AGP FLA FLA promoter pFLA Gal Galactosyltransferase GalT galacturonic acid GalA Genomic DNA gDNA gibberellin A GA Glucose Glc glucuronoarabinoxylans GAX glyceraldehyde 3-phosphate dehydrogenase GAPDH glycosylphosphatidylinositol GPI GT histidine HIS homogalacturonan HG hydrophobic interaction chromatography HIC hydroxyproline Hyp hydroxyproline galactosyltransferase HPGT hydroxyproline-rich HRGP immunoprecipitation IP interfascicular fibres IF isobaric peptide tags for relative and absolute quantification iTRAQ

VII

Knotted1-like homeodomain protein KNAT liquid chromatography mass spectrometry LC-MS Luria broth LB Manose Man mass spectrometry MS microfibril angle MFA mixed linkage glucan MLG mixed membranes MM monoclonal antibodie mAb Myeloblastosis MYB NAC secondary wall thickening NST NAC master switches SWN NAM, ATAF1,2 and CUC2 NAC no apical meristem NAM Nottingham Arabidopsis Stock Centre NASC nuclear magnetic resonance NMR Phospholipase PL plasma membrane PM plasma membrane intrinsic protein PIP polymerase chain reaction PCR post-translational modification PTM quantitative PCR Q-PCR receptor-like kinase RLK rhamnogalacturonan RG Rha room temperature RT root hair specific RHS salt overly sensitive SOS secondary wall-associated nac domain protein SND -Hydroxyproline-Hydroxyproline-Hydroxyproline SOOO Serine-Hydroxyproline--Hydroxyproline SOTO Serine-Hydroxyproline-Serine-Hydroxyproline SOSO SP sodium dodecyl sulphate SDS standard deviation SD standard error SE super optimal broth with catablolite SOC transcription factor TF transforming growth factor beta induced protein TGFBIp Tris-acetic-EDTA TAE ultra violet UV vascular endothelial growth factor receptor VEGFR vascular related NAC-domain VND wall associated kinase WAK Xylose Xyl

VIII xyloglucan XyG yellow florescent protein YFP

IX

TABLE OF CONTENTS

Abstract I Declaration IV Acknowledgements V Abbreviations VII Table of contents X List of tables XIII List of figures XIV

Chapter 1 Literature review 1 1.1 The plant cell wall 2 1.1.1 Primary cell wall 3 1.1.2 Secondary cell wall 4 1.1.3 Wall polysaccharides 5 1.2 The wall (glyco)proteins 6 1.3 Arabinogalactan-proteins (AGPs) 7 1.3.1 AGP structure and classification 8 1.3.2 AGP biosynthesis 8 1.3.3 Glycosylphosphatidylinositol (GPI) anchor 10 1.3.4 Roles of AGP 11 1.4 Fasciclin-like AGP (FLAs) 12 1.4.1 The fasciclin (FAS) molecules and FAS domains 12 1.4.2 Identification of FLAs in 14 1.4.3 Biological roles of AtFLAs 15 1.4.4 Prediction of FLA functional domains 17 1.5 Research aims 18

Chapter 2 Materials and methods 27 2.1 Plant material and growth conditions 28 2.2 Crossing of plants 28 2.3 Surface sterilization of Arabidopsis (ethanol method) and growth on MS plates 28 2.4 Dark grown hypocotyl and isoxaben treatment 29 2.5 Genomic DNA (gDNA) isolation 29 2.6 Preparation of plasmid DNA 29 2.7 Polymerase chain reaction (PCR) 29 2.8 RNA extraction, cDNA synthesis and quantitative real time PCR (Q-PCR) 30 2.9 DNA Electrophoresis 30 2.10 Generation of constructs using a NEBuilderRHiFi DNA Assembly kit 31 2.11 Total protein extraction 31 2.12 Isolation of mixed membrane from plant tissues 31 2.13 SDS-PAGE protein gel electrophoresis 32 X

2.14 Immunoblotting 32 2.15 Enrichment of FLA fusion proteins 33 2.15.1 Chelation 33 2.15.2 Immunoprecipitation (IP) 33 2.15.3 HIC 34 2.16 Bacterial transformation 34 2.17 Arabidopsis transformation by floral dip 35 2.18 Selection of transformed Arabidopsis 35 2.19 Infiltration of Nicotiana benthamiana with Agrobacterium 35 2.20 Fixation and embedding of Arabidopsis stem tissues with LR White resin 35 2.21 Toluidine blue staining 36 2.22 Mäule staining 37 2.23 Confocal Laser-Scanning Microscopy 37 2.24 Alcohol-insoluble residue (AIR) cell wall preparation and starch removal 37 2.25 Linkage analysis with carboxyl reduction 38 2.25.1 Reduction I – linkage analysis 38 2.25.2 Reduction II – linkage analysis 38 2.25.3 Methylation analysis – linkage analysis 39 2.26 Acetic/Nitric crystalline cellulose assay 39 2.27 Measurement of biomechanical properties of the stem 40

Chapter 3 Tissue and sub-cellular distribution of fla11, fla12 and fla16 in Arabidopsis 45 3.1 Introduction 46 3.2 Results 47 3.2.1 Generation and validation of FLA reporter constructs 47 3.2.2 Sub-cellular fractionation suggests FLA11 FLA12 and FLA16 are localised to the 49 plasma membrane and walls 3.2.3 Evaluation of FLA11, FLA12 and FLA16 reporters shows expression in tissues and cells 50 with secondary walls 3.3 Discussion 51 3.3.1 FLA11, FLA12 and FLA16 are largely cell surface proteins (plasma membrane and/or 52 cell wall) specific to cell types that develop secondary wall-specific 3.3.2 Sub-cellular localisation of FLA11, FLA12 and FLA16 in the PM and wall suggests 54 interaction with wall proteins and/or and roles for the determination of wall properties Appendices to Chapter 3 77 A3.1 Enrichment of FLA11 reporter proteins 77 A3.2 A YFP signal corresponding to FLA16-HV is detected in pavement cells and trichomes 78 in leaves of young seedlings

Chapter 4 Functional investigation of Arabidopsis fasciclin-like arabinogalactan-protein 16 88 (FLA16) 4.1 Introduction 89 4.2 Results 90 4.2.1 Investigation of FLA16 transcript levels in Arabidopsis 90 4.2.2 Growth is delayed in the fla16 mutant 90

XI

4.2.3 Reduced stem length and transverse area was observed in fla16 mutants 91 4.2.4 Differences in cellular morphology are observed in fla16 stems compared to wt 92 4.2.5 Carbohydrate content is altered in the fla16 stems 93 4.2.6 fla16 stems display altered biomechanical properties compared to wt 94 4.2.7 Complementation of the fla16 mutant 95 4.2.8 Expression of FLA16 in the stem overlaps with the expression of the primary and 95 secondary wall cellulose synthases (CesAs) 4.2.9 The fla16 mutant is more sensitive to isoxaben treatment than wt 96 4.3 Discussion 97 4.3.1 FLA16 potentially influences stem biomechanics by regulating cellulose microfibrils 98 Appendices to Chapter 4 122

Chapter 5 Comparative and functional investigation of FLA11, FLA12 and FLA16 through mutant studies 128 5.1 Introduction 129 5.2 Results 130 5.2.1 Generation of fla11, fla12 and fla16 double and triple mutants 130 5.2.2 Loss of FLA11, FLA12 and FLA16 causes a reduction in inflorescence stem length 130 5.2.3 Total stem area is reduced in fla mutants 131 5.2.4 Cellular morphology is altered in triple fla11 fla12 fla16 mutant 131 5.2.5 Stem biomechanical properties are not altered in double and triple fla mutants 132 5.2.6 The fla mutant stems have altered polysaccharide composition 133 5.2.7 Analysis of CesA transcript levels in wt and fla11 fla12 fla16 mutants 133 5.2.8 FLA11, FLA12 and FLA16 show an epistatic relationship for stem length 134 5.3 Discussion 135 5.3.1 FLA11, FLA12 and FLA16 likely have overlapping roles regulating stem growth in Arabidopsis 135 5.3.2 Genetic redundancy might contribute to the genetic interaction between FLAs 137 Appendices to Chapter 5 149

Chapter 6 Summary 156 6.1 Summary of current research 157 6.2 Hypothetical models of FLA functions 157

References 163

XII

LISTS OF TABLES

Table 1.1 Summary of FLA functional studies 19 Table 2.1 List of primers used for cloning constructs 41 Table 2.2 DNA templates for generation of constructs 42 Table 2.3 Primers for genotyping T-DNA insertions in FLA11, FLA12 and FLA16 43 Table 2.4 Primers for Q-PCR analysis 44 Table 3.1 Tissue-specific expression patterns of FLA11, FLA12 and FLA16 in Arabidopsis 59 Table A3.1 Mass spectrometry identification of FLA11 peptides after different enrichment 79 methods Table A3.2 Proteins identified by mass spectrometry after enrichment by Chelation, IP and 80 HIC of HV-FLA11 Table 4.1 Analysis of selected growth stages of long day grown Arabidopsis wt and fla16 plants 103 Table 4.2 Measurement of stem morphology, biomechanical properties and cellulose 104 content of long day grown wt and fla16 stems at maturity Table 4.3 Analysis of tissue and cellular morphology in mature stems of Arabidopsis wt 105 and fla16 plants grown in long day conditions Table 4.4 Biomechanical properties of wt and fla16 stems at senescence completion 106 Table A4.1 Vegetative organ growth rate of fla16 and wt 122 Table A4.2 Polysaccharide composition from stems of fla16 and wt deduced from linkage 123 analysis Table 5.1 Measurement of length, total transverse sectional area and cellulose content 140 in mature stems of fla11, fla12 and fla16 single, double and triple mutants and wt Table 5.2 Measurements of tissue area in transverse sections of mature stems from 141 fla11fla12fla16 and wt Table 5.3 Measurements of cell size and number in transverse sections of stems of 142 fla11fla12fla16 and wt Table 5.4 Measurement of biomechanical flexure properties of stems of fla11 fla12, 143 fla11 fla16, fla12 fla16, fla11 fla12 fla16 and wt Table A5.1 Calculation of polysaccharide composition based on linkage analysis of fla11 149 fla12 fla16 triple mutant and wt stems Table A5.2 Calculation of crystalline cellulose in stems of fla mutants and wt 151

XIII

LIST OF FIGURES

Figure 1.1 Plant cell wall structure and organization 20 Figure 1.2 Schematic diagram of plant cell wall polysaccharides 21 Figure 1.3 Biosynthesis of AGPs – post-translational modifications (PTMs) 23 Figure 1.4 Occurrence of AGPs in sequenced plant genomes 25 Figure 1.5 Schematic of the predicted protein structures of FLAs from Arabidopsis 26 Figure 3.1 Protein sequences and predicted domains of Arabidopsis FLA11, FLA12 and 60 FLA16 Figure 3.2 Schematic representation of FLA11, FLA12 and FLA16 constructs 61 Figure 3.3 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins extracted 62 from tobacco and Arabidopsis Figure 3.4 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins in sub- 63 cellular fractions of proteins extracted from mature Arabidopsis stems Figure 3.5 HV-FLA11, HV-FLA12 and FLA16-VH are co-localised with plasma membrane 65 intrinsic protein 2 (PIP2) in turgid and plasmolysed tobacco cells Figure 3.6 Transcript abundance of FLA11, FLA12 and FLA16 in Arabidopsis tissues 67 Detection of YFP in plants expressing pFLA11:HV-FLA11, pFLA12:HV-FLA12 Figure 3.7 and pFLA16:FLA16-VH reporter proteins in transverse sections taken at the 69 first internode of mature Arabidopsis stems Figure 3.8 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins in the first 71 branch of mature Arabidopsis stems Figure 3.9 Detection of HV-FLA11, HV-FLA12 and FLA16-VH in transverse sections of 73 Arabidopsis siliques Figure 3.10 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins in the 75 guard cells of 7 d.o. Arabidopsis cotyledons Figure 3.11 Detection of HV-FLA11, HV-FLA12 and FLA16-VH in the Arabidopsis root 76 vasculature of 4 d.o. seedlings Figure A3.1 Enrichment of HV-FLA11 from tobacco leaves using a chelation method 81 Figure A3.2 Enrichment of HV-FLA11 from tobacco leaves using a hydrophobic interaction 82 chromatography (HIC) method Figure A3.3 Detection of FLA16-VH in the epidermal pavement cells of Arabidopsis 83 cotyledons and true leaves Figure A3.4 Detection of FLA16-VH in the Arabidopsis true leaves 84 Figure A3.5 Expression pattern of FLA11 during Arabidopsis development 85 Figure A3.6 Expression pattern of FLA12 during Arabidopsis development 86 Figure A3.7 Expression pattern of FLA16 during Arabidopsis development 87 Figure 4.1 Expression levels of FLA16 transcripts in tissues of Arabidopsis wt and a fla16 107 T-DNA insertion line Figure 4.2 The fla16 mutant shows a reduced stem length phenotype at maturity 108 Figure 4.3 Cellular morphology of Arabidopsis fla16 and wt in transverse sections of 109 fresh mature stems Figure 4.4 Lignin distribution in transverse sections of fresh mature stems of fla16 and 111 wt Figure 4.5 Cellular morphology derived from transverse sections of mature stems of 113 Arabidopsis fla16 and wt Figure 4.6 Measurement of branch angle relative to the main stem at the first node of 115 mature Arabidopsis wt and fla16 plants Figure 4.7 Cell wall polysaccharide composition derived from linkage analysis of mature 116 stems of fla16 and wt Figure 4.8 Complementation of the fla16 mutant with pFLA16:FLA16 and FLA16:FLA16- 117 VH

XIV

Figure 4.9 Q-PCR analysis of CesA transcript abundance in top-, mid- and basal- 119 segments of mature Arabidopsis wt and fla16 stems Figure 4.10 Isoxaben treatment of dark grown Arabidopsis fla16 and wt seedlings 121 Figure A4.1 Division of stem transverse area into three main tissues of interest for total 125 area analysis in wild-type (wt) and fla16 plants Figure A4.2 Biomechanical properties of dry wt and fla16 basal stems 126 Figure A4.3 Tensile properties of fla16 and wt stems at senescence completion 127 Figure 5.1 Stem length phenotypes of fla11, fla12 and fla16 single, double and triple 144 mutant plants and wt Figure 5.2 Light micrographs of transverse sections showing cellular morphology of 145 stems of wt and fla11fla12fla16 mutant Figure 5.3 Analysis of cell wall polysaccharide composition (A) and crystalline cellulose 146 (B) levels in mature stems of fla11 fla12 fla16 mutants and wt Figure 5.4 Q-PCR analyses of CesA transcript abundance in the top, middle and basal 147 stems of wt, fla16, fla11fla12 and fla11fla12fla16 plants Figure 5.6 Double and triple mutant analyses of fla11, fla12 and fla16 total stem length 148 Figure A5.1 Schematic diagram of T-DNA insertions in FLA11, FLA12 AND FLA16 152 Figure A5.2 Genotyping of homozygous fla11, fla12 and fla16 mutants by PCR 153 Figure A5.3 Phenotypes of dark-grown wt and fla mutant seedlings treated with isoxaben 154 Figure 6.1 Schematic representation of hypothetical mechanism(s) of action of FLA11, 161 FLA12 and FLA16

XV

CHAPTER 1

LITERATURE REVIEW

1

1.1 The plant cell wall

Plant cells are surrounded by a cell wall that largely determines the properties of the cell (Cosgrove, 2005). It is estimated that there are approximately forty different cell types in plants, each having individual size, shape, cellular componentry and location (Demura et al., 2002). By combining tensile strength with extensibility, the wall is key to regulating the shape, direction and rate of growth (Carpita and McCann, 2000). The plant cell wall allows transport of metabolites and signal transduction between cells. It also plays critical roles in both abiotic and biotic stress responses, and is an important determinant of plant morphology. There are two main types of plant cell walls, primary wall and secondary wall (Doblin et al., 2010). The primary wall, laid down during cell expansion, is present in all cells and many mature plant cells are only surrounded by the primary cell wall. The primary wall largely consists of polysaccharides with a minor (glyco)protein component having structural, enzymatic and signalling functions (Fry, 2004) (Fig 1.1). In some specialized cell types both a primary wall and secondary wall (laid down after cells have stopped expanding and undergo differentiation) are deposited (e.g. xylem cells) (Harris, 2006). These two types of walls differ in a number of aspects such as the composition and structure of the matrix phase polysaccharides, as well as their arrangement, and the presence (secondary wall) or absence (primary wall) of lignin. This in turn results in different wall properties, such as rheological and mechanical (Cosgrove and Jarvis, 2012).

2

1.1.1 Primary cell walls The primary wall is hydrated (approx. 60% water), thin (≤100 nm), extensible and of sufficient tensile strength to simultaneously prevent the protoplast from rupturing and yielding to accommodate cell expansion and maintain integrity during cell growth (Fincher and Stone, 1986; Bacic et al., 1988; Cassab and Varner, 1988; McCann and Roberts, 1994; Doblin et al., 2010; Hamant and Traas, 2010). Polysaccharides account for about 90% of primary walls with (glyco)proteins comprising the remaining 10%. The major polysaccharide is cellulose which forms rigid microfibrils and is the scaffold for association of other wall polysaccharides. The cellulose microfibrils are embedded in a gel-like matrix composed of pectins and non-cellulosic polysaccharides (hemicelluloses). Wall proteins include wall loosening proteins (eg. expansins, hydrolases and endo-transglycosylases) (Nishitani and Tominaga, 1992; Purugganan et al., 1997; Steele et al., 2001; Johnson et al., 2003; Eklof and Brumer, 2010), cross-linking enzymes (eg. peroxidases and laccases) (Hatfield et al., 2017) and glycoproteins (e.g. arabinogalactan-proteins) (Josè-Estanyol and Puigdomènech, 2000; Ellis et al., 2010; Tan et al., 2012). Primary walls of flowering plants have been classified into two major groups, Type I and Type II walls (Carpita and Gibeaut, 1993; Harris et al., 1997; Harris, 2005) although in reality a continuum exists across the plant kingdoms (Bacic et al 1988; Doblin et al 2010). Type I walls are composed of a cellulose-xyloglucan (XyG) network embedded in a matrix abundant in pectic polysaccharides and are found in gymnosperms, dicots, and non- commelinid monocots (Fincher and Stone, 1986; Bacic et al., 1988; Carpita and Gibeaut, 1993; McCann and Roberts, 1994). Type II walls, found in the commelinid monocots, that include the cereals (Poales), differ in the composition of their major matrix polysaccharides that are predominantly (1,3;1,4)-β-glucans (MLGs) and glucuronoarabinoxylans (GAXs) in addition to a much lower abundance of pectic polysaccharides and XyGs (Bacic et al., 1988; Carpita and Gibeaut, 1993; McCann and Roberts, 1994; Smith and Harris, 1999). Significant advances have been made in the identification of the enzymes that make the wall polysaccharides and their post-transcriptional regulation (Ghahremani et al., 2016). Despite this, currently there is little information about the transcriptional regulation of primary walls biosynthesis. Progress is being made, for example, the brassinosteroid-

3 activated transcription factor BRI1-EMS-SUPPRESSOR 1 (BES1) is proposed to control the expression of genes encoding cellulose synthases (CesAs) in the primary wall (Nishitani and Demura, 2015). Investigation of the transition from primary walls to secondary walls has also proven a successful approach to identify master regulators of secondary wall development (Li et al., 2016a; Li et al., 2016b).

1.1.2 Secondary cell walls A secondary wall develops in specialized tissues such as the vasculature (e.g. xylem) and interfascicular fibre cells when the cell stops expanding (Bacic et al., 1988; Carpita and Gibeaut, 1993; Harris, 2006; Zhong and Ye, 2014). The secondary wall is deposited on the inside of the primary wall and mainly consists of cellulose, lignin and lowly-substituted hemicelluloses (eg. heteromannans and heteroxylans). Secondary walls provide mechanical strength to the plant to support vertical growth and for the transport of water and nutrients (Alberts et al., 2002; Farrokhi et al., 2006). Being hydrophobic, lignin is crucial to the ‘waterproofing’ of the secondary wall-possessing cells (Wang and Dixon, 2012; Prats et al., 2016). Formation of the secondary wall is complex and requires controlled expression of genes responsible for wall biosynthesis, precise secretion, deposition and assembly of the wall components (Zhong and Ye, 2014). Recently a number of transcription factors (TFs) that act as master regulatory switches to co-ordinate the expression of genes required for the secondary wall development have been identified (Zhong et al., 2010a). Members of the plant-specific NAC (for no apical meristem (NAM), such as Arabidopsis thaliana transcription activation factor (ATAF1/2) and cup-shaped cotyledon (CUC2) domain family of TFs (Lv et al., 2016), act as master regulators governing activation of the secondary wall assembly and deposition. Collectively named the secondary wall NAC master switches (SWNs) (Zhong and Ye, 2014), ten SWNs have been identified in Arabidopsis. Of the seven vascular related NAC-domain (VND1-7) members, VND6 and VND7 are specifically expressed in vessels and are responsible for vessel secondary wall thickening (Kubo et al., 2005; Yamaguchi et al., 2008; Zhou et al., 2014). Secondary wall -associated NAC domain 1 (SND1) and NAC secondary wall thickening factor 1 (NST1) are specifically expressed in the fibres and are responsible

4 for controlling secondary wall thickening of these tissues (Zhong et al., 2006; Mitsuda et al., 2007; Zhong et al., 2007; Mitsuda and Ohme-Takagi, 2008; Zhong and Ye, 2015). SWNs are proposed to activate a range of TFs including SND2 and 3, the myeloblastosis (MYB) domain TFs (MYB20, 42, 43, 46, 52, 54, 58, 63, 69, 83, 85, 103) and the Knotted1-like homeodomain protein 7 (KNAT7) that also act as important regulators of secondary wall development (Zhong et al., 2008). For example, MYB46 and MYB83 are close homologs that act downstream of both SND1 and VND6/7 (Zhong et al., 2007; McCarthy et al., 2009). MYB46 and MYB83 are responsible for the thickening of secondary walls in both fibres and vessels by regulating the expression of genes responsible for the biosynthesis of xylan, cellulose and lignin (Ko et al., 2009; McCarthy et al., 2009). These findings have enabled significant progress in our understanding of the regulatory networks controlling the assembly and deposition of the secondary walls (Sánchez-Rodríguez et al., 2010; Zhong and Ye, 2014; Schneider et al., 2016).

1.1.3 Wall polysaccharides

There is a huge diversity in the composition and structure of the wall polysaccharides which includes cellulose, XyG, MLGs, heteroxylans, heteromannans, and the pectins (homogalacturonan (HG), rhamnogalacturonan I (RGI) and rhamnogalacturonan II (RGII) (Fig. 1.2). The entire process of polysaccharide synthesis is considered to comprise four steps. These include Stage I, activation and production of nucleotide-sugar donors; Stage II, polymer synthesis initiation; Stage III, polymer elongation; and Stage IV, polymer synthesis termination (Delmer and Stone, 1988). So far, little is known about the initiation and termination processes but there has been considerable progress on the activation and production of nucleotide-sugar donors and polymer synthesis stages. Two types of enzyme are essential in the biosynthesis of plant cell walls polysaccharides, namely, glycosyltransferases (GTs), generally responsible for the synthesis of the sidechains, and polysaccharide synthases involved in backbone assembly. These enzymes are critical in the catalysis of bonds formed between contiguous monosaccharides that are delivered by activated nucleotide-sugar donors (Bar-Peled and O'Neill, 2011; Orellana et al., 2016). Detailed information about the structures and biosynthesis of these wall polysaccharides 5 can be found in major reviews (Cosgrove, 2005; Doblin et al., 2010; Gorshkova et al., 2013) and the current study focuses on the wall glycoproteins.

1.2 The wall (glyco)proteins

Wall proteins account for only a small portion of the primary wall, usually no more than 10% of dry weight in land plant (embryophyte) walls (Farrokhi et al., 2006) whereas they can be a significant component of some algal walls (up to 30%) (Roberts, 1974). These wall proteins are key components in the maintenance of the biological and physical functions of the extracellular matrix in plants. Furthermore, the wall proteins are important elements in the continuum of the wall and plasma membrane to perceive and transduce signal from the exterior to interior compartment (Johnson et al., 2003; Wolf et al., 2012) for example, the wall associated kinases (WAKs) (Kohorn, 2001). These wall proteins can be divided into structural, e.g. extensin and some arabinogalactan-proteins (AGPs), and enzymatic (e.g. esterases, glycosidases, peroxidases, proteases and hydrolases). Wall proteins can be glycosylated (Josè-Estanyol and Puigdomènech, 2000; Johnson et al., 2003) and are the main focus of the current study.

The hydroxyproline-rich glycoproteins (HRGPs) represent a large and diverse group of glycosylated structural wall proteins (Showalter, 1993). The HRGPs include three major groups, the arabinogalactan-proteins (AGPs), extensins and proline-rich proteins (PRPs) (Showalter et al., 2010; Johnson et al., 2017a; Johnson et al., 2017b). These proteins are characterized by being rich in Proline (Pro, P) residues which can be hydroxylated to Hydroxyproline (Hyp, O) depending on the surrounding amino acids, including Serine (Ser, S), Alanine (Ala, A) and Threonine (Thr, T). The ‘Hyp contiguity hypothesis’ (Kieliszewski and Lamport, 1994; Johnson et al., 2017a; Johnson et al., 2017b) predicts that when O occurs in a non-contiguous manner, for example SOTO, such as occurs in AGPs, this acts as a signal for glycosylation of large branched type II arabinogalactan (AG) polysaccharides (Tan et al., 2003). Contiguous O residues, such as SOOO, which occur in extensins, direct addition of short arabino-oligosaccharide structures (Ma and Zhao, 2010). This hypothesis is supported by experiments using synthetic peptides whereby large AG polysaccharides were added to 6

O residues in repeats of either AOAO or SOSO (Shpak et al., 1999; Shpak et al., 2001) and small arabino-oligosaccharides where found on contiguous O residues (ie. T/SOOO) as occurs in extensin-like sequences (Goodrum et al., 2000). AG glycosylation rarely occurs when the peptide sequence consists of O only (Ma and Zhao, 2010).

In order to gain insight into the diversity of HRGPs that exist, bioinformatic approaches, for example in Arabidopsis, have revealed that these proteins belong to large multigene families (Showalter, 2001; Schultz et al., 2002; Doblin et al., 2010; Showalter et al., 2010; Johnson et al., 2017a; Johnson et al., 2017b).

The wall glycoproteins play important roles in the plant growth and development. Extensins are reinforcing elements during both growth and in response to biotic and abiotic stresses (McNeil et al., 1984; JosÈ and PuigdomÈNech, 1993; Josè-Estanyol and Puigdomènech, 2000). PRPs are known to play a contributory role against infection by pathogens and physical damage and are also determinants of the wall structure in specific cell types (Fowler et al., 1999). AGPs are a particularly large gene family with more than one hundred members predicted in Arabidopsis (Johnson et al., 2017a; Johnson et al., 2017b) and are proposed to play a number of biological roles, for instance interaction between cells, and governance of cell proliferation (Sommer-Knudsen et al., 1998). The focus of this literature review will be on the AGP family of HRGPs.

1.3 Arabinogalactan-proteins (AGPs)

The protein backbone of AGPs is rich in the amino acids (aa), P, A, S and T, or ‘PAST’ (Schultz et al., 2002; Ma and Zhao, 2010; Showalter et al., 2010; Johnson et al., 2017a; Johnson et al., 2017b). AGPs have attracted considerable attention due to their highly complex structures and potential roles in signalling. In addition, they have industrial and health applications due to their chemical/physical properties (water-holding, adhesion and emulsification) (Ellis et al., 2010). AGPs have been reported in early diverging land plants and algae and, in eudicots, are widespread in seeds, roots, stems, leaves and (Gaspar et al., 2001). They have also been reported in secretions of cell culture medium of

7 root, leaf, and tissues, and some exudate producing cell types such as stylar canal cells are capable of producing large amounts of AGPs (Fincher and Stone, 1983).

1.3.1 AGP structure and classification

AGPs can be classified into several sub-groups (Showalter, 2001; Schultz et al., 2002; Gaspar et al., 2004; Huang et al., 2013; Johnson et al., 2017a; Johnson et al., 2017b) on the basis of their predicted protein structure. These are: classical AGPs consisting of a signal peptide at the N-terminus, a PAST-rich sequence of 100-150 aa and a hydrophobic region at the C-terminus that directs addition of a glycosylphosphatidylinositol (GPI)-anchor; non GPI-AGPs that lack the C-terminal GPI signal sequence, Lys(K)-rich AGPs that contain a K- rich region within the PAST-rich backbone; AG-peptide that have a short PAST-rich backbone of 10-15 aa; chimeric AGPs that include fasciclin-like AGP (FLAs) and plastocyanin AGPs that, in addition to the AGP domain have an additional PFAM domain. Other ‘non- classical’ AGPs exist, for example those having, histidine (H)-rich and cysteine (C)-containing domains as well as many hybrid HRGPs that have motifs characteristic of AGPs and other HRGPs (Baldwin et al., 2001; Gaspar et al., 2001; Showalter et al., 2010; Ma et al., 2017; Johnson et al., 2017a; Johnson et al., 2017b).

1.3.2 AGP biosynthesis

After translation, the AGP protein backbones are highly decorated with complex carbohydrates, primarily type II AG polysaccharides (Liang et al., 2013). The biosynthesis of the mature AGP involves several processes which are cleavage of the signal peptide at the N-terminus, on the P residues in some consensus regions, the addition of the GPI-anchor and arabinogalactosylation on the O residues (Liepman et al., 2010) (Fig 1.3).

All AGPs protein backbones contain an N-terminal signal peptide that directs the protein into the endoplasmic reticulum (ER) where the post-translational modification (PTM)

8 processes begin (Schultz et al., 1998). Prolyl hydroxylation of P to O is fulfilled by prolyl 4- hydroxylases (P4Hs) belonging to the 2-oxoglutarate dependant dioxygenase family (Koski et al., 2007). P4H has been identified in both the ER and (Ellis et al., 2010). Prolyl hydroxylation appears to depend on specificity of the protein backbone sequence by P4Hs (Estevez et al., 2006) apart from the preferences of some of the GTs that add the first Gal residue onto the Hyp (Showalter and Basu, 2016).

The addition of the GPI-anchor occurs in most but not all AGPs (Schultz et al., 2002). The concomitant cleavage of the C-terminal hydrophobic (GPI-anchor) signal sequence of some AGPs and the addition of the GPI-anchor converge in the ER (Youl et al., 1998; Ellis et al., 2010). The integration of a GPI-anchor enables the attachment of the protein to the membrane of the ER transiting to the Golgi apparatus leading to secretion to the outer leaflet of the plasma membrane facing the wall (Muniz and Zurzolo, 2014). As proposed by Oxley and Bacic (1999), the GPI-anchored AGPs are likely released via cleavage by some phospholipases (PLs) (C or D) and secreted into the extracellular compartment. Previously, a few GPI-anchored proteins on the plasma membrane were reported to exhibit transglycosylase like activities (Hurtado-Guerrero et al., 2009), which gives some GPI- anchored AGPs a possible pathway to be cross-linked to the wall glycan chains by transglycosylation. Tan et al. (2013) proposed AGPs could be associated with other wall polysaccharides via covalent cross-linking (e.g. AtAGP1 and wall hemicellulose and pectin).

The glycosylation initiation of AGP occurs in the ER (Oka et al., 2010) whereas the chain extension process takes place primarily in the Golgi apparatus (Kato et al., 2003). Some GTs are known to assemble the AGs on the AGP backbone and may cooperate to determine the sequence, length and density of the AG chains (Showalter and Basu, 2016). After prolyl hydroxylation, Gal and/or Ara residues are added to individual O residues, and the galactosyltransferases (GalTs)/hydroxyproline galactosyltransferases (HPGTs), two subfamilies of the CAZy GT31 family, (e.g. GalT2-6 and HPGT1-3) (Showalter and Basu, 2016) and AraTs (Porchia et al., 2002), respectively, are responsible. The rest of the AGP sugars are contributed by other GTs: for example, AtGlcAT14 (A, B and C) have GlcAT activity to β- (1,6)- and β-(1,3)- (Knoch et al., 2013; Dilokpimol and Geshi, 2014); AtGALT29A have β-(1,6)-GalT activity to β-(1,6)- and β-(1,3)-galactan (Dilokpimol et al., 2014); RAY1 has 9

β-ArafT activity for methyl-β-Gal (Gille et al., 2013); and AtFUT (4 and 6) show fucosyltransferase (FucT) activity for AGPs as found in the BY2 cells (Liang et al., 2010; Wu et al., 2010b; Tryfona et al., 2014).

1.3.3 Glycosylphosphatidylinositol (GPI) anchor

The majority of AGPs in Arabidopsis encode a C-terminal hydrophobic signal sequence for attachment of a GPI anchor (Schultz et al., 2002; Johnson et al., 2003). The GPI-anchor has a core glycan structure consisting of a trimannosyl-glucosamine tetrasaccharide linking to the C-terminal amino acid of the AGP protein through ethanolamine phosphate (Schultz et al., 1998; Youl et al., 1998; Saha et al., 2016). The early synthesis of the GPI moiety occurs on the ER cytoplasmic surface and subsequent assembly take place in the lumen of the ER. These include the assembly of tri-, galactose and ethanolamine phosphate to form the mature GPI moiety. AGPs undergo GPI-anchor addition while co-translationally migrating into the ER and these two processes finally converge. Subsequently, a transamidase complex simultaneously cleaves the core protein at the C-terminus when it recognizes the ω cleavage site and transfers the fully assembled GPI-anchor onto the amino acid residue at the C-terminus of the protein. These events occur prior to prolyl hydroxylation and glycosylation (Imhof et al., 2004; Ellis et al., 2010).

After GPI-anchor assembly and glycosylation, the mature AGPs are exported to the cell membrane by vesicular transport. AGPs are bound to the membrane facing the wall via the GPI-anchor or potentially released by the action of phospholipases (PLs) which cleave the GPI-anchor. Two classes of PLs present in the apoplast, GPI-PLC and/or GPI-PLD, are proposed to cleave the GPI-anchor according to their preferred cleavage sites on the GPI moiety (Fig 1.3) (Udenfriend and Kodukula, 1995). GPI-anchored proteins are enriched in lipid nano-domains where they are proposed to associate with other proteins, including those involved in signalling functions (Ellis et al., 2010). It has also been proposed membrane bound GPI-anchored AGPs are able to interact with either the matrix wall components or other membrane proteins (Schultz et al., 1998). An in vitro study showed low molecular weight APGs from the membrane bound fraction cause formation of the AGP

10 complexes/aggregates after treatment with a peroxidase and hydrogen peroxide. These aggregates resemble those from the sugar beet leaf extracts and suggest a mechanism by which AGPs might interact with the wall matrixpolysaccharides and/or proteins (Kjellbom et al., 1997; Schultz et al., 1998).

1.3.4 Roles of AGP

Conventional methods to study functions of AGPs include the use of β-glycosyl (usually glucosyl) Yariv reagents and monoclonal antibodies (mAbs). β-Glycosyl Yariv reagents are synthetic phenylazo glycoside probes that specifically, but not covalently, bind to AGPs and can be used to precipitate AGPs from solution. They are also used commonly as histochemical stains to probe the locations and distribution of AGPs (Yariv et al., 1962; Tang et al., 2006). A number of studies have shown that addition of β-Yariv reagents to plant growth medium can inhibit seedling growth, cell elongation, block and fresh cell wall mass accumulation (Willats and Knox, 1996; Chapman et al., 2000; Zagorchev et al., 2013). The use of mAbs that specifically bind to carbohydrate of AGPs have also been employed to infer functions based on the location and pattern of the AGP epitopes. Commonly used mAb against AGPs include CCRC-M7, LM2, JIM8, JIM13 and JIM14 (Seifert and Roberts, 2007).

The determination of fine structure of any single AGP and the establishment of its precise function are major challenges. Many roles have been attributed to AGPs, including roles in hormone signalling, cell expansion and division, embryogenesis of somatic cells, differentiation of xylem, responses to abiotic stress, plant growth and development. These studies suggest that they are multifunctional, similar to what is found in mammalian /glycoproteins (Filmus et al., 2008; Schaefer and Schaefer, 2010; Tan et al., 2012). The large number of family members also suggests functional redundancy causing difficulties for mutant analysis (Motose et al., 2004; Coimbra et al., 2009). The heavy decoration by glycans on the AGP core proteins makes structural and functional analyses for a single AGP extremely difficult.

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Despite the challenges, a number of groups have made significant progress into understanding the function of individual AGPs. For example, the presence of an Arabidopsis root-specific AtAGP30 was shown to be required for in vitro root regeneration suggesting a function in regenerating the root by modulating phytohormone activity (Van-Hengel and Roberts, 2003). In addition, studies of agp6 and agp11 mutants in Arabidopsis have demonstrated the importance of these AGPs to prevent uncontrolled generation of the -grain and for normal growth of the pollen-tube (Coimbra et al., 2010; Suzuki et al., 2017). Roles in plant development for members in the fasciclin-like AGPs (FLAs) have also been reported (see below).

1.4 Fasciclin-like AGP (FLAs)

FLAs are members of the ‘chimeric’ AGP sub-class and are characterised by the presence of fasciclin (FAS) domains in addition to the AGP motifs in their protein backbones. FASI and FASII proteins were initially identified in grasshopper growth cones and fasciculating axons (Bastiani et al., 1987). Since this discovery FAS domains have been identified in proteins across all kingdoms of living organism from a broad spectrum of taxonomic classifications suggesting they are evolutionarily ancient FAS-like molecules related to cell adhesion molecules (CAMs) which act to sense the extracellular environment and play critical roles in the structure, function and development of multi-cellular organisms (Aplin et al., 1999; Clout et al., 2003; Wilkins et al., 2003). FLAs are therefore of interest to determine if FAS domains confer similar functions in plants.

1.4.1 The fasciclin (FAS) molecules and FAS domains

FAS domains are generally 110 to 150 aa in length, and have been identified in a broad range of taxonomic classifications in all phyla (Moody and Williamson, 2013). FAS domains are commonly found in extracellular and membrane associated proteins, and are proposed to act as evolutionarily ancient cell adhesion domains (MacMillan et al., 2015). Proteins frequently contain multiple FAS domains, either in tandem or interspersed with

12 other domains, however single FAS domain containing proteins are common in plants (Johnson et al., 2003; Moody and Williamson, 2013).

Although FAS sequences show low sequence similarity, all known FAS domains share two highly conserved motifs, H1 and H2, that each consist of approximately 10 aa (Kim et al., 2000; Kim et al., 2002; Johnson et al., 2003). The removal of H1 and H2 motifs did not prevent cell adhesion of the MRC-5 cell line, a diploid human cell line, and were proposed to facilitate molecular interactions (Kim et al., 2002). Proteins with two FAS domains are common in animals and structural analysis suggests that FAS domains can interact. For example, a crystal structure of FAS domains 3 and 4 of the Drosophila FAS1 protein show they interact via a rich polar interface (Clout et al., 2003). In the single FAS domain containing protein Fdp from Rhodobacter sphaeroides, structural determination suggests the interaction with other proteins occurs at the site usually embedded within the dimer interface (Moody and Williamson, 2013).

Some FAS-containing proteins have been studied for their cell adhesion properties. FASI in Drosophila was one of the earliest proteins to be characterized and is a GPI-anchored molecule (Hortsch and Goodman, 1990) with four tandem FAS (~150 aa) domains. The cell adhesion function of FASI was based on in vitro studies using Drosophila non-adhesive Schneider 2 cells (Elkins et al., 1990a). Cells expressing FASI become aggregated and the cell aggregation could be eliminated in the presence of a FASI-specific antibody. In addition, when mixing cell lines expressing either FASI or FASIII, the mixed cells formed homogenous aggregates for either FASI or FASIII. These findings suggested that FASI facilitates homophilic interactions (Elkins et al., 1990a). In addition, FAS domains are strong candidates in protein-protein interaction as not only can FAS domains themselves interact, the FAS domain also have the potential to interact with other proteins to regulate cellular events. Another well studied example is periostin in humans, consisting of four FAS1 domains in a tandem arrangement (Litvin et al., 2004; Kudo et al., 2007). Periostin is expressed in a number of tissues such as bone, skin and periodontal ligament, and is involved in protein-protein interactions with collagen and integrins (Hamilton, 2008). The interaction of perostin with integrins is proposed to impact upon proliferation, differentiation and adhesion of cells (Kudo et al., 2007). Nam et al. (2012) reported the 13

FAS1 domain of human transforming growth factor beta induced protein (TGFBIp/βig-h3) interacts with αvβ3 integrin to prevent association of vascular endothelial growth factor receptor 2 (VEGFR2) and αvβ3 integrin.

The cell adhesion function also appears to be conserved in the plant lineage. Studies in Volvox carterii identified an algal cell adhesion molecule (Algal-CAM) required for correct embryogenesis (Sumper and Hallmann, 2003). Algal-CAM consists of an extensin-like domain at the N-terminus (Cassab and Varner, 1988) and two tandem FAS domains (Huber and Sumper, 1994). Disruption of Algal-CAM function using antibodies resulted in embryonic defects including: interference with contact between adjacent embryonic cells at the 4-cell stage causing a hole between the cells, a marked reduction and alteration in the pattern of , and inhibited embryonic inversion. These finding suggest the FAS containing Algal-CAM is involved in cell division and embryogenesis. This pioneering study has been followed by further examples of FAS-containing glycoproteins functioning during plant development (Elkins et al., 1990a; Johnson et al., 2003; Morra and Moch, 2011; Huang et al., 2013; Moody and Williamson, 2013).

1.4.2 Identification of FLAs in plants

Bioinformatic studies have shown that FLAs belong to multigene families in all plant species investigated to date, for example, 21 FLAs in the Arabidopsis genome, 24 in rice (), 34 in wheat (Triticum aestivum) and 16 in Eucalyptus (Eucalyptus grandis) (Schultz et al., 2002; Johnson et al., 2003; Faik et al., 2007; Ma et al., 2017) (Fig. 1.4). The structures of FLAs vary in terms of the number of FAS and AGP domain(s) and the presence of a GPI-signal sequence. To date, FLAs have been best characterised in the model plant Arabidopsis (Johnson et al., 2011). By means of pair-wise sequence comparisons of the predicted proteins, the 21 Arabidopsis FLAs were grouped into 4 sub-groups (A-D) based on structural similarity (Fig. 1.5). FLAs are predicted to be post-translationally modified to include N-glycosylation in the FAS domain and O-glycosylation in the AGP domains. Support for the addition of AG-polysaccharides on these proteins can be seen in biochemical studies showing FLAs can interact with the β-Glc (Johnson et al., 2003). 14

1.4.3 Biological roles of AtFLAs

From the Arabidopsis Functional Genomic Consortium (AFGC) microarrays, expression data for 11 of the 21 AtFLAs indicates a number of FLAs have high expression in flowers compared to other tissue types. These include AtFLA1, AtFLA2, AtFLA7, AtFLA8, AtFLA9, AtFLA11, AtFLA12, AtFLA15, AtFLA16 and AtFLA18 (Schultz et al., 2002). Some FLAs show more restricted expression patterns that may indicate specialized functions. Based on promoter-GUS experiments, AtFLA11 and AtFLA12 are almost exclusively expressed in the stem (MacMillan et al., 2010). In situ hybridization indicates AtFLA11 is expressed predominantly in the sclerenchyma cells of siliques and stems (Ito et al., 2005). AtFLA3 also displays a restricted expression pattern, being specifically expressed in the pollen grains and tubes. Sub-cellular localization studies showed AtFLA3 attaches tightly to the PM, likely through its GPI-anchor (Li et al., 2010). Comprehensive tissue-specific and sub-cellular localization data is not available to date for most FLAs.

The function of some FLAs has been evaluated through phenotypes attributed to the respective fla mutants (Table 1.1). Studies of Atfla1 suggest AtFLA1 has a role in shoot development and early development of lateral roots based on a reduced ability of Atfla1 to undergo shoot regeneration in in vitro experiments (Johnson et al., 2011). A cotton GhFLA1 mutant has been investigated due to a potential role in fibre development. Huang et al. (2013) showed that down-regulation of GhFLA1 led to delayed fibre initiation and elongation whereas up-regulation resulted in promotion of fibre elongation and increased fibre length. This suggests GhFLA1 may be responsible for initiation and elongation of the fibre through influencing the matrix integrity of the primary wall (Huang et al., 2013).

A study of the Arabidopsis group C FLA, AtFLA3, revealed a role in pollen development. RNA interference experiments to knock down AtFLA3 resulted in plants with approximately half the pollen exhibiting morphological deformities, such as wrinkled and shrunken grains. In plants overexpressing AtFLA3, siliques are notably shorter with low set caused by reduced female fertility and a stamen filament elongation defect. These

15 phenotypes indicate abnormal AtFLA3 expression may cause disruption and reduction of cell growth and thus defective outcomes in these tissues (Li et al., 2010).

An Arabidopsis salt overly sensitive 5 (sos5) mutant was identified as a mutation in the gene encoding AtFLA4. When grown in the presence of high salt sos5 exhibited phenotypes such as swelling root tips and arrested growth suggesting a role for AtFLA4 in root growth and development. Intriguingly, promoter-GUS examination revealed the ubiquitous expression of AtFLA4 in all organs and tissues (Shi et al., 2003). A more recent study proposed AtFLA4 glycosylation is critical in mediating root growth and the adherence of seed through a pathway involving the cell wall receptor-like kinases, FEI1/FEI2 (Basu et al., 2016).

Expression of Eucalyptus FLA1 and FLA2 was found to be inversely correlated with the cellulose microfibril angle (Qiu et al., 2008). Overexpression of Eucalyptus FLA2 (ortholog of Arabidopsis FLA12) using the p35S promoter caused a cellulose microfibril angle reduction in the xylem by 3 degrees, whereas in transgenic tobacco expressing a p35s:EgrFLA3 construct (ortholog of Arabidopsis FLA11) caused a reduction in flexural strength in the stem (MacMillan et al., 2015).

In Arabidopsis, studies of two stem-specific FLAs, AtFLA11 and AtFLA12 also revealed a role in maintaining stem biomechanical properties. Single mutants of Atfla11 and Atfla12 show an increase in the flexural stiffness of stems compared to the wild-type and a reduction in cellulose content (MacMillan et al., 2010). Double fla11fla12 mutants show a further reduction of cellulose content, as well as reduced Gal and Ara, and reduced tensile stiffness compared to wild-type stems. These phenotypes indicate AtFLA11 and AtFLA12 play a role in altering the cell wall composition and hence architecture which in turn affect the biomechanical properties of the stem (MacMillan et al., 2010).

The function of this sub-group of FLAs in stem development also appears to be conserved in Populus trichocarpa. Knockdown of PtFLA6 (ortholog of AtFLA11 and AtFLA12) using antisense led to a change in the stem biomechanical properties, including a reduction of flexural stiffness and strength (Wang et al., 2014).

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1.4.4 Prediction of FLA functional domains

The presence of both FAS and AGP domains in FLAs opens up the potential for a range of putative interactions: protein-protein, protein-carbohydrate and carbohydrate- carbohydrate interactions. Their potential role as protein-protein interaction domains was discussed in section 1.4.1 above.

Bearing AGP domains, FLAs also have potential to interact with wall polysaccharides via their structural glycans. Bucior and Burger (2004) demonstrated the binding force of oligosaccharides between different surface proteoglycans are specific and as strong as that between antigen and antibody. This establishes the carbohydrate-carbohydrate interaction by not only the binding strength but also the specificity (Bucior and Burger, 2004). Furthermore, recent studies have shown that AGPs can be covalently cross-linked with pectins and matrix phase wall polysaccharides suggesting AGP glycans are also involved in maintenance of cell wall architecture by carbohydrate-carbohydrate interactions (Tan et al., 2013).

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Therefore, the FLAs are fascinating wall glycoproteins and their proposed roles in plant development warrants further investigation of the potential mechanism(s). To-date there is little information about their sub-cellular location, tissue expression, and their interacting partners; such knowledge would greatly assist our understanding of how these glycoproteins might function. Furthermore, in depth mutant studies whereby multiple FLA members are knocked out will also aid clarification of the role of FLAs in plant development. A multidisciplinary approach will be taken in this project to address key questions of AtFLA functions.

1.5 Research aims The aim of this study is to investigate a selection of group A and B FLAs in Arabidopsis in order to understand their function in stem biomechanics. Secondary cell wall-specific FLAs are attractive for functional studies as loss of fla11 fla12 results in measurable changes in wall content and architecture. This research will make a major contribution to our basic understanding of how wall glycoproteins influence plant growth. FLA11 and FLA12 influence the MFA in secondary walls in stems yet the mechanism is unclear. The subtle phenotype of fla11 fla12 mutants suggested other FLAs may be involved in maintaining stem biomechanics. The relatively high expression of FLA16 in the stem indicated this would be a good candidate for investigation. To gain further insight into the role of FLA11, FLA12 and FLA16 in cells with secondary walls, details of their post-translational modifications and sub-cellular location would be informative. The specific research aims are:

1. Investigation of the biological function of AtFLA16 (group B). 2. Identification and generation of FLA mutant combinations (double mutants: fla11fla12, fla11fla16, fla12fla16 and triple mutant fla11fla12fla16) to examine FLA functional redundancy. 3. Characterization of FLA (FLA11, FLA12 and FLA16) expression patterns in vivo via FLA fusion proteins.

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Figure 1.1 Plant cell wall structure and organization. A. Schematic diagram of the plant cell wall arrangement showing the primary and secondary (S1, S2, S3) walls. B. Simplified schematic of the primary wall. Figure taken from Sticklen (2008).

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Figure 1.2 Schematic diagram of the structures of representative plant cell wall polysaccharides. These structures can vary depending on the plant species. Cellulose (A), xyloglucan (XyG) (B) and (1,3;1,4)-β-glucan (MLG) (C) have a backbone made of glucose (Glcp), heteroxylan (D) of xylose (Xylp), heteromannan (E) of mannose (Manp), rhammogalacturonan I (RGI) (G) of alternating rhamnose (Rhap) and galacturonic acid (GalpA), homogalacturonan (HG) (F) of GalpA and RGII (H). Taken from (Doblin et al., 2010).

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Figure 1.3 Biosynthesis of AGPs – post-translational modifications (PTMs). PTMs include co-translational insertion of the nascent N-terminus of the core protein from the cytoplasm into ER, attachment of the fully assembled GPI anchor on the C-terminus, prolyl hydroxylation and glycosylation. The AGPs are then transported to the Golgi apparatus for further AG assembly and the mature AGPs are transported to the plasma membrane via vesicles. The AGPs may then be shed from the PM by phospholipases (GPI-PLD or GPI-PLC) (Schultz et al., 1998).

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Figure 1.4 Occurrence of AGPs in sequenced plant genomes. The numbers of AGP members in the subfamilies are indicated in columns corresponding to the species (C: classical AGP, KC: Lysine-rich classical AGP). The species tree is indicated on the left based on Phytozome (https://phytozome.jgi.doe.gov/pz/portal.html) and NCBI (http://www.ncbi.nlm.nih.gov/guide/taxonomy/). Figure taken from Ma et al. (2017).

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Figure 1.5 Schematic of the predicted protein structures of FLAs from Arabidopsis. The 21 FLAs are divided into four sub-groups and all contain N-terminal signal peptides. Group A FLAs consist of 1 FAS and 2 AGP domains, and are all predicted to be GPI- anchored. Group B contain 2 FAS domains and 1 AGP domain with the exception of AtFLA16 which predicts 2 AGP domains. All group B FLAs possess 1 intron in their genomic DNA (indicated by arrow head). Group C FLAs share less sequence similarity within the group and are all GPI-anchored. FLA1, FLA2, FLA8 and FLA10 all contain 2 FAS and 2 AGP domains whereas FLA3, FLA5 and FLA14 all have 1 FAS and 1 AGP domain. Group D FLAs share even less intra-group sequence similarity than group C. Only FLA4 is predicted to be GPI-anchored and has 2 FAS and 2 AGP domains. FLA20 also has 2 FAS domains and only 1 AGP domain. Both FLA19 and FLA21 have only 1 FAS and 1 AGP domain (Modified from Johnson et al., 2011).

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CHAPTER 2 MATERIALS AND METHODS

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2.1 Plant material and growth conditions Seeds of wild type Arabidopsis thaliana var. Columbia (Col) were obtained from the Arabidopsis Biological Resource Centre (ABRC, USA). Seeds of T-DNA insertion lines, fla11 (SALK_046976) and fla16 (SALK_131248) were obtained from the Salk Institute for Biological Studies (SALK, CA). Seeds of T-DNA insertion line fla12 (SM.15162 NASC) were obtained from the Nottingham Arabidopsis Stock Centre (NASC, UK). Seeds were sown in soil in punnet pots, jiffy pots (Garden City, Australia) or ½ MS plates (0.22% Murashige & Skoog basal salt, MS, and 0.8% phytagel), stratified at 4oC for 3 days and transferred into a controlled environment growth chamber (Thermoline, Australia). Growth conditions were 16 h light/8 h dark at 21oC. Seeds of tobacco, Nicotiana Benthamiana, were provided by the Plant Cell Biological Research Centre, University of Melbourne. Tobacco seeds were sown on soil, and grown in a glasshouse maintained at 21-25oC and humidity of approximately 50%. Tobacco seedlings were then transferred into individual pots. Young leaves were used for Agrobacterium infiltrations.

2.2 Crossing of plants Flowers were prepared by removing anthers, petals and sepals, retaining only the pistil at stage 6.5 (Boyes et al., 2001; Weigel and Glazebrook, 2002). Flowers were marked and left overnight to ensure no self-pollination had occurred. Anthers from mature flowers were removed and the convex surface used to brush against the surface of the to transfer pollen. The mature siliques were collected when yellow and dry but before splitting.

2.3 Surface sterilization of Arabidopsis seeds (ethanol method) and growth on MS plates Arabidopsis seeds were washed in seed wash solution (70% ethanol and 0.05% Triton X-100) with mixing for 10 min. The seed wash solution was discarded and seeds washed 2-3 times in 100% ethanol. The seeds were transferred onto a piece of sterile Whatman filter paper and allowed to dry. The seeds were placed on ½ MS plates using a sterile toothpick.

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2.4 Dark grown hypocotyl and isoxaben treatment Arabidopsis seeds were plated on ½ MS plates containing the herbicide isoxaben (Sigma #36138) (2nM) or plates lacking isoxaben (controls). The plates were wrapped three times in aluminium foil sheets to block light and stratified at 4oC for 3 days prior to removal of the foil and light treatment for 4 h at room temperature (RT). The plates were then re-covered in foil and transferred to a growth chamber and maintained in the dark at 21oC for 4 days.

2.5 Genomic DNA (gDNA) isolation Leaf material was harvested, snap frozen in liquid nitrogen and milled into a fine power using a TissueLyserII mixer mill (Qiagen #85300) with 5 mm stainless steel beads (Qiagen #69989) at 18 vibrations s–1 for 30 s. Extraction buffer (0.5mL, 200mM Tris-HCl, 25 mM EDTA, 250mM NaCl and 0.5% SDS) (Edwards et al., 1991) was added to the plant material. The tissue was mixed by vortexing and centrifuged for 5 min at 10,000g. DNA in the supernatant was precipitated with an equal volume of isopropanol and centrifuged at 10,000g for 10 min. The pellet was washed with 80% ethanol and air- dried before addition of 55 µL of distilled H2O (dH2O).

2.6 Preparation of plasmid DNA E.coli containing plasmids of interest were cultured in Terrific Broth (1.2%

Tryptone, 2.4% yeast extract, 17mM KH2PO4 and 7.2mM K2HPO4) with appropriate antibiotics at 37oC, 220 rpm for at least 16 h. A 500 µL aliquot of bacterial culture was mixed with 50% glycerol stock and stored at -80oC. The remaining culture was pelleted by centrifugation at 5,000g for 10 min. Plasmid DNA was extracted using the Isolate II Plasmid mini kit (Bioline #BIO-52027) as per the manufacturer’s instructions. The eluted plasmid DNA was analyzed for DNA concentration and purity with a NanoDrop 2000 spectrophotometer (NanoDrop) and stored at -20oC.

2.7 Polymerase chain reaction (PCR) PCRs were carried out to amplify DNA fragments of interest for cloning and genotyping. Primers and DNA templates are listed in Table 2.1 and 2.2. PCR reactions included a final concentration of 0.5 µM of each of the forward and reverse primers, 0.5 mM deoxyribonucleotides (dNTPs) (0.125mM each dATP, dTTP, dCTP and dGTP), 1.5 29 mM MgCl, 1× PCR reaction buffer (1.5 mM MgCl2, 10 mM Tris HCl pH8.3 and 50 mM KCl), 1 U of polymerase (either high fidelity Phusion Taq (ThermoFisher F530S) for cloning or

TM MyTaq DNA polymerase (Bioline #BIO-21105) for genotyping), and dH2O to 20 µL. PCR reactions included an initial denaturation step of 95oC for 3 min followed by 25 repeated cycles (95oC denaturation for 30 s, 50-68oC primer annealing for 30 s, 72oC extension for 1 min per 1000bp) and a final extension at 72oC for 10 min. Primers for genotyping are listed in Table 2.3 and information of the expected size of DNA products.

2.8 RNA extraction, cDNA synthesis and quantitative real time PCR (Q-PCR) Approximately 100 mg of fresh plant material was ground to a fine power using liquid nitrogen as described in section 2.4. RNA isolation was performed using an RNeasy Plant Mini Kit (Qiagen 74904) according to the manufacturer’s instruction and RNA concentration were determined by a NanoDrop 2000 spectrophotometer (NanoDrop). DNase treatment was carried out using a DNaseI kit (Invitrogen #18068- 015) and complementary DNA (cDNA) synthesised using a SuperscriptIII Reverse Transcriptase kit (Invitrogen #18080-093) as per the manufacturer’s instructions. Q-PCR was performed to examine the expression levels of FLA and CESA genes. An absolute quantitative method adapted from (Burton et al., 2004) was used with two biological and three technical replicates. Primers are listed in Table 2.4. Transcript levels were normalized with four housekeeping genes, Actin, Tublin, Cyclophylin and Glyceraldehyde 3-phosphate dehydrogenase (GAPDH). DNA standards for each primer set were kindly prepared by Dr. Neil Shirley, University of Adelaide, Australia at a concentration of 109 copies/ l. Q-PCR reactions were performed with the KAPA SYBR FAST Q-PCR Kit Master Mix (2×) Universal (KapaBiosystems, #KK4601) in 10µl reactions in a Bio-Rad CFX384 Real-Time System (BioRad).

2.9 DNA Electrophoresis DNA was separated on 1-2% agarose gels containing 1 µg/mL ethidium bromide in 1 x Tris-acetic-EDTA (TAE) (Fisher Scientific #2500060). DNA samples were mixed with loading dye (NEB, #B7024S) prior to loading into the gel. A constant voltage of 130V was used to separate DNA. The HyperLadderTM 1kb molecular weight marker (Bioline, #BIO- 33025) was run with the samples for size estimation, and the DNA was visualized using a Gel DocTM EZ Imager (Bio-Rad) or ultra violet (UV) transilluminator. 30

2.10 Generation of constructs using a NEBuilder○R HiFi DNA Assembly kit

Constructs of interests were generated by cloning of DNA fragment(s) into vectors using Seamless Cloning with a NEBuilder○R HiFi DNA Assembly kit (New England BioLabs #E5520). This was performed as per the manufacturer’s instructions. Primers and DNA templates used for molecular cloning are listed in Table 2.1 and 2.2.

2.11 Protein extraction Plant tissue was homogenized in protein extraction buffer (100mM Tris-HCl pH7.4, 150mM NaCl, 1% NP-40 and cOmplete Protease inhibitor, Roche # 11697498001, adapted from the lysis buffer recipe of a GFP-Trap ○R immunoprecipitation kit

(Chromotek) and previous AGP extraction protocols (Poon et al., 2012). Proteins were extracted in an ice bath, with a ratio of tissue to buffer of 0.5 g/mL. The homogenate was rotated end-to-end at 4oC for 1 h. The mixture was filtered through Miracloth (Merk Millipore #475855) and the liquid centrifuged at 10,000g, at 4oC for 10 min to pellet cell debris. The supernatant, containing the protein, was used for subsequent analysis.

2.12 Isolation of mixed membrane from plant tissues Mixed membranes were prepared according to Doblin et al. (2009) with the following modifications. Plant material was grounded in liquid nitrogen and mixed in a Microsomal Prep Homogenizing Buffer (100mM PBS, 20mM KCl, 500mM sucrose, cOmplete Protease inhibitor, (Roche # 11697498001)), with a ratio of plant tissue to homogenizing buffer of 0.5 g/mL. The homogenate was filtered through a double layer of Miracloth (Merk Millipore #475855) with pore size 22-25 µm prior to centrifugation at 10,000g, at 4oC for 10 min to pellet cell debris. The supernatant was transferred to an ultracentrifugation tube and centrifuged at 100,000g at 4oC for 1 h to pellet the mixed membrane microsomes. The microsomes were re-suspended in a minimal amount of 50 mM Tris-HCl (~0.5-1 mL) containing cOmplete EDTA-free proteinase inhibitor (Roche #11836170001) at the concentration recommended by the manufacturer.

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2.13 SDS-PAGE protein gel electrophoresis Protein electrophoresis was performed using a Mini-PROTEIN○R System (BioRad #1658004). Protein samples were quantified by a BCA Protein Assay Kit (Pierce #23225) as per the manufacturer’s instructions. Approximately 30 µg of protein per sample was mixed with a Laemmli Sample Buffer (BioRad #161-0737) and incubated in a thermoshaker at 70oC for 15 min before centrifugation at 10,000g at room temperature for 1 min. The samples, along with a KaleidoscopeTM prestained SDS-PAGE Standard (BioRad #161-0324), were loaded into a precast Mini-PROTEIN○R TGX Stain-FreeTM Gel (BioRad #456-8094). The gel was run in a Tris/Glycine/SDS Buffer (BioRad #161-0732) at 200 V for approximately 20 min or until the loading dye reached the bottom of the gel. Proteins were visualized using a ChemiDocTM MP Imaging System (BioRad #170- 8280).

2.14 Immunoblotting

Gels were dry-blotted on an iBlot○R 2 nitrocellulose membrane (ThermoFisher

#IB23001) by an iBlot○R 2 Gel Transfer Device (ThermoFisher #IB21001) using a blotting procedure of 20 V for 1 min, 23 V for 4 min and 25 V for 2 min. The membrane was then blocked in blocking solution (5% skim milk in Tris buffered saline, TBS) for 30 min. The blot was then incubated in primary antibody (1:1000 in blocking solution with 0.05% Tween) for 2 h. The membrane was washed five times in 0.05% Tween in TBS (TBST) for 5 min each. This was followed by application of goat anti-mouse horse radish peroxidase (HRP) conjugated secondary antibody (ThermoFisher #31430) (1:5000 in blocking solution). The membrane was washed as above and visualized using a SuperSignal West Femto Maximum Sensitivity chemiluminescent substrate (ThermoFisher #34094), and signal detected by a ChemiDocTM MP Imaging System (BioRad #170-8280) as per the manufacturer’s instructions.

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2.15 Enrichment of FLA fusion proteins FLA fusion proteins tagged with yellow florescent protein (YFP) and poly histidine (HIS) tags were enriched using chelation, immunoprecipitation (IP) and hydrophobic interaction chromatography (HIC).

2.15.1 Chelation Chelation employed the cOmplete HIS-Tag purification resin (Roche, #05893801001). Approximately 1 mL of resin was packed into a 1 mL Bio-ScaleTM Mini Cartridge (BioRad, #7324660). The resin was then equilibrated with at least 5 mL of enrichment buffer (EB) (100mM Tris-HCl pH7.4, 150mM NaCl, 1% NP-40 and cOmplete Protease inhibitor, Roche # 11697498001, adapted from the lysis buffer recipe of a GFP- Trap○R immunoprecipitation kit, Chromotek). Soluble total protein extracts (0.5 mL) were loaded into a 12 mL syringe and loaded onto the cartridge at a constant speed of 1 mL/min then both ends of the cartridge were sealed. The cartridge was incubated at 4oC for at least 2 h. Fractions were collected in a 9-well (5 mL/well) tissue culture plate on ice. The resin was washed in 3 mL of cold EB (UNBOUND) then washed with 5 mL of EB containing 10 mM imidazole (WASH). This was followed by step-wise elution in EB containing 100, 200 and 500 mM imidazole, and fractions collected and marked accordingly. All protein fractions were desalted with 5 kDa PD-10 columns (GE Healthcare Life Science, #17085101) and concentrated to 0.5 mL with 10 kDa Ultra-15 Centrifugal Filter Units (Amicon, #UFC901008). The fractions were transferred to 1.5 mL microcentrifuge tubes (Eppendorf) and examined under a Dark Reader LED transilluminator (Clare, #DR46B) emitting visible light at 4-500 nm for YFP detection.

2.15.2 Immunoprecipitation (IP) IP employed the immunoaffinity of the YFP and anti-GFP antibodies coupled to agarose beads (GFP-Trap, Chromotek, #gta-20). Approximately 30 µL of beads were washed with 1 mL of cold EB three times. Tubes were centrifuged at 2500g for 2 min at 4oC, and supernatant removed between each wash. Approximately 0.5 mL of soluble protein extract was incubated with the beads on a rotating platform for at least 2 h at 4oC. The mixture was centrifuged at 2500g for 2 min at 4oC, and supernatant removed prior to washes (5x) with EB as above. The beads and bound proteins were incubated with 100 µL of 100 mM glycine (pH2.5) and vortexed at RT for 10 min. It was then 33 centrifuged at 2550g and supernatant collected (Elution 1), then repeated (Elution 2). The eluate was neutralized with 20 µL of 1 M Tris-HCl (pH10.4) then mixed with 1.2 mL of ice cold acetone and precipitated overnight. The eluate was then centrifuged at 22000g for 10 min at 4oC, supernatant removed and airdried for 5 min at RT. The pellet was solubilized with 10 mM Tris-HCl (pH7.4) for downstream applications.

2.15.3 HIC HIC employs the hydrophobic interaction between YFP and phenyl sepharose resin. Approximately 3 mL of phenyl sepharose resin was packed into a 4 mL Econo column (Biorad, #7374006) connected to an Econo pump system (Biorad, #7318142) and flow rate set to 1 mL/min. A 24-well (3 mL/well) tissue culture plate was placed on ice for fraction collection. The pump was set to a single buffer mode and column equilibrated with 6 mL of buffer A (10mM Tris-HCl, pH8.0, 1.6M ammonium sulphate) at RT and the flow-through was discarded. Approximately 9 mL of soluble protein extract in buffer A was introduced to the column, fractions collected for reloading and repeated three times. The column was then washed with 15 mL of buffer A. The pump was set to a dual buffer/gradient mixing mode and the injecting buffer reservoirs filled with buffer A and buffer B (10mM Tris-HCl). The bound proteins were eluted in increasing concentrations of B/A. The fractions were examined on the Dark Reader LED transilluminator (Clare, #DR46B) at 4-500 nm and YFP positive fractions desalted and concentrated as outlined in 2.15.1.

2.16 Bacterial transformation Plasmids were transformed into bacteria using either electroporation or heat- shock. Electroporation was performed using a MicroPulser Electroporator (BioRad #165-2100) following the manufacturer’s instructions. Heat-shock transformation was performed at 42oC for 30 s followed by incubation on ice for 2 min using commercial competent cells (NEB, #C2987H) according to the manufacturer’s protocol. Bacteria were recovered in Super Optimal broth with Catablolite (SOC) medium (NEB #B9020S) then plated on Luria broth (LB) agar plates containing the appropriate antibiotics for plasmid selection.

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2.17 Arabidopsis transformation by floral dip Agrobacterium tumefaciens (AGL1) culture was centrifuged at 5,000g at room temperature for 10 min. The supernatant was removed and the bacterial pellet was re- suspended in 500 mL of floral-dip transformation medium (5% sucrose, 10mM MgCl2). Approximately 125 µL of Silwet L-77 was mixed with the bacterial solution and Arabidopsis flowers dipped in the solution three times for 20 s each as outlined in the floral dip method by Weigel and Glazebrook (2002). Dipped plants were placed horizontally in trays covered in plastic wrap and maintained at 4oC overnight. The following morning, the plastic wrap was removed and plants placed upright and watered.

2.18 Selection of transformed Arabidopsis Seeds (F0) collected from dipped plants were dried with desiccant at room temperature for 3 days before sowing on soil. From 7 days-old, seedlings were sprayed

○R with a solution containing 0.1% BASTA and 0.25% Silwet L-77 twice a week for 4 to 6 times. Seeds were collected from plants that survived BASTA treatment (F1) and used for subsequent BASTA screening and downstream analysis.

2.19 Infiltration of Nicotiana benthamiana with Agrobacterium

Agrobacterium culture containing the plasmid of interest and a culture containing the P19 plasmid (Qiu et al., 2002) were centrifuged at 5,000g for 15 min and the pellet re-suspended in MMA infiltration buffer (0.45% MS salt, 10mM 2-(N- morpholino) ethanesulfonic acid, MES, 200µm acetosyringone and 2% sucrose) to give an optical density (OD) reading of 4.8. Equal amounts of the bacterial solutions were mixed to give a final OD of 2.4. Young leaves of Nicotiana benthamiana were infiltrated with bacteria according to the method of (Yang et al., 2000).

2.20 Fixation and embedding of Arabidopsis stem tissues with LR White resin

Arabidopsis stem tissues were hand-sectioned to a thickness of approximately 2 mm and fixed according to the protocol outlined in Wilson and Bacic (2012). The tissue was fixed in 2.5% (v/v) glutaraldehyde (ProSciTech #C001) in phosphate buffer saline (PBS, pH 7.4, Sigma #P4417) overnight at 4oC. Samples were washed 3 x with PBS then

35 dehydrated in a graded ethanol series (10%, 20%, 30%, 50%, 70%, 90%) in dH2O then twice in 100% ethanol for 10 min each. LR White Resin (London Resin Company Ltd #AGR1281) was substituted for ethanol in a graded series (25%, 50%, 75%, 2 × 100%) for 24 h each and embedded in fresh resin in gelatin capsules (Chemist Warehouse, #56065) and polymerized at 55oC overnight. Blocks were trimmed using a Leica EM Trim trimmer (Leica Microsystems, Germany) and sectioned with a Leica Ultracut R microtome (Leica Microsystems, Germany) using glass knives.

2.21 Toluidine blue staining

Arabidopsis stems embedded in LR white were sectioned (500 µm) and placed on glass slides before staining with toluidine blue as previously described (Chateigner- Boutin et al., 2014) with the following modifications. Sections were incubated in toluidine blue (0.5% Toluidine blue O, 0.1% sodium carbonate, pH11.1) for 10 min then washed in dH2O to remove excess stain. For staining of fresh Arabidopsis stems, hand-sections (approx. 0.5 mm) were placed in five serial concentrations of Toluidine blue (0.01, 0.3, 0.5, 0.8, 0.1%) solution and incubated for 2 min prior to washing three times in dH2O to remove excess stain. Stem sections were examined under a Leica DM6000 B compound microscope (Leica Microsystems, Germany) and images captured by a Leica DFC450 C camera (Leica Microsystems, Germany). Images were analyzed using ImageJ (Schneider et al., 2012) (http://rsb.info.nih.gov/ij/). For area measurements the stem circumference was traced and area calculated using Image J plugin. For tissue/cell area, cell number and cell length measurements, more than three biological samples with two technical replicates each were used.

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2.22 Mäule staining Mäule staining (Pradhan Mitra and Loqué, 2014) was carried out to examine lignin distribution in the fresh Arabidopsis stems. Fresh stem tissue was hand-sectioned transversely with a thickness of approx. 0.5 mm prior to incubation in KMnO4 (1%) for 10 min. Sections were then washed briefly in water before acidification with HCl (37%) for 1 min. The sections were soaked in NaHCO3 (5%) for 5 min or until a red/brown colour developed then washed in water. The sections were then examined under a Leica DM6000 B compound microscope (Leica Microsystems, Germany) and images captured by a Leica DFC450 C camera (Leica Microsystems, Germany).

2.23 Confocal Laser-Scanning Microscopy

Fluorescence microscopy of fresh tissue, either whole mount or transverse sections, was carried out on a Leica SP5 microscope (Leica Microsystems, Germany). Laser beam lines exciting at 514 nm were employed for the yellow fluorescent protein (YFP), Venus, and fluorescence emitted between 520 and 530 nm was captured. Images were analysed for tissues expressing reporter fusions with the Leica LAS Lite software and images processed with Fiji (Schindelin et al., 2012) and Adobe Photoshop CS.

2.24 Alcohol-insoluble residue (AIR) cell wall preparation and starch removal AIR material was prepared as described in Pettolino et al. (2012). Mature stems were dissected out, snap frozen in liquid nitrogen and ground into fine power with pre- cooled mortar and pestle. A pinch of ground sample was placed on a microscopic slide and examined under an optical microscope to ensure few intact cell structures remained. Approximately 100 mg of sample was weighed out and transferred into a 2 mL microcentrifuge tube. About 1.5 mL of 80% (v/v) ethanol was added, mixed thoroughly and incubated on ice for 30 min. The tissue was centrifuged at 10,000g at room temperature for 5 min and the supernatant removed. Ethanol extraction of the pellet was repeated until the green colour in the pellet disappeared and appeared grey/white. Acetone (1.5 mL) was then added to the pellet and incubated at room temperature for 10 min prior to centrifugation as above. A 100% methanol extraction was performed as

37 above and the pellet was dried with desiccant at 40oC under vacuum overnight and then stored in an airtight container at room temperature until required. Starch in the AIR preparation was removed by α-amylase treatment. AIR samples were soaked in 100-200 µL of 10 mM Tris-maleate buffer for 30 min prior to incubation in boiling water for 5 min for the starch granules to gelatinize. Samples were equilibrated to 40oC for 10 min in a thermo-shaker before adding α-amylase (Sigma #A3403), 2U per microgram of AIR sample, at 40oC for 1 h. An additional 1U per microgram of AIR of α-amylase solution was added to the reaction and incubated as above for 30 min. Following this, four volumes of cold ethanol were added to the reaction and incubated in -20oC for 1 h. The solution was then centrifuged at 1,500g at room temperature for 5 min, supernatant removed and the AIR preparation dried under vacuum as above.

2.25 Linkage analysis with carboxyl reduction

Linkage analysis was performed as described in Pettolino et al. (2012), which was adapted from Sims and Bacic (1995) as well as Kim and Carpita (1992). The three steps of linkage analysis, two reduction steps and methylation are as follows:

2.25.1 Reduction I Five milliliters of 1 M ice-cold imidazole-HCl was added to 5 mg of AIR preparation before addition of 1 mL of 100 mg/mL sodium borodeuteride. The samples were vortexed, incubated on ice for 5 min followed by a further two sodium borodeuteride additions (1 mL each) for 5 min and 2 h. Glacial acetic acid (500 µL) was added slowly to neutralize the excess reductant in the samples before dialysis in dH2O overnight and followed by freeze-drying. The samples were dissolved in 1 mL of dH2O before addition of 200 µL of 0.2 M MES buffer and 400 µL of freshly prepared 500 mg/mL carbodiimide (Sigma, #C10,640-2). The samples were vortexed to be homogenate before incubation at room temperature for 3 h.

2.25.2 Reduction II After the first reduction, 1 mL of 4 M imidazole was added to the samples before incubation on ice. Each sample was divided in half and transferred to two separate 38

Falcon tubes (BD Falcon, USA). To one of the duplicate tubes, was added 1 mL of 70 mg/mL sodium borohydride whereas 1 mL of 70 mg/mL sodium borodeuteride was added to the other. They were then incubated at room temperature for 3 h. After this, 500 µL of glacial acetic acid was slowly added to the samples to destroy excess reductant and the samples then dialysed against dH2O for 24 h and then freeze-dried.

2.25.3 Methylation analysis This methylation method was adapted from Ciucanu and Kerek (1984) as described by Pettolino et al. (2012). About 100 µL of DMSO was added to the samples treated by carboxyl reduction (see above) prior to sonication for 20 min. After this, 100 µL of DMSO/NaOH slurry was added to samples with a SMI glass pipette. The samples were sonicated for 20 min before adding 20μl of methyl iodide and sonication for 10 min and then another 20μl addition of methyl iodide and sonication as mentioned. After this, 40 µL of methyl iodide was added before sonication of 20 min. milli-Q water (1 mL) and 1 mL of DCM were added and the tubes then vortexed for 40 s to be homogeneous. The samples were centrifuged at 10,000g for 1 min to allow phase separation, the upper aqueous phase removed and the CHCl3 phase washed three times with 1 mL of milli-Q water. The CHCl3 phase was dried in the presence of nitrogen stream and the samples were hydrolysed, reduced and acetylated to generate the permethylated alditol acetates as described in section 2.24. The samples were then analysed by GC-MS as described by Lau and Bacic (1993), and the molecular percentage of each polysaccharide was calculated according to Pettolino et al. (2012).

2.26 Acetic/Nitric crystalline cellulose assay The acetic/nitric protocol was adapted from Updegraff (1969) with the following modifications. AIR (10-50 mg) was transferred into a pre-weighed 2 mL microcentrifuge tube and Updegraff reagent (acetic acid : nitric acid : water, 8:1:2 v/v) added to the sample, mixed, and then heated in an oven at 100oC for 90 min. The sample was allowed to cool to room temperature and centrifuged at 10,000g at room temperature for 5 min to pellet crystalline cellulose. The supernatant was removed and the pellet was washed with 80% ethanol (2x) and acetone (1x) at room temperature. After the acetone wash, the pellet was allowed to air-dry and further dried at 40oC in a vacuum oven in the presence of desiccants overnight. The sample was allowed to cool to room temperature 39 with desiccant then weighed on a five-figure balance. The percentage of crystalline cellulose was calculated according to the following equation: Crystalline cellulose (%) = crystalline cellulose (mg) / AIR cell wall (mg) x 100

2.27 Measurement of biomechanical properties of the stem

Three-point flexural and tensile and tests were carried out by a 4500 series Instron universal testing machine (series IX automated materials testing system, http://www.instron.co.uk) as described by MacMillan et al. (2010). The modulus of elasticity (stiffness) was calculated using Hooke’s law whereas flexural three-point bending stiffness and strength were calculated via standard equations (MacMillan et al., 2010). Tensile strength was calculated as the maximum load needed to break the stem within the gauge length, and this was divided by the cross-sectional area of the stem (MacMillan et al., 2010). These measurements were kindly performed by Dr MacMillan (CSIRO, Canberra). A snapping profile was determined based on graphs assessing the response of stems to the applied load. The number of stems that gave a sharp break (‘snapped’) when the load reached the maximum were compared to those with a gradual break.

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Table 2.1 List of primers used for cloning constructs. Product Primer Sequence Template (bp)

35s Sp16 F ACGCTCGAGGAATTCATGGATTCCTCC Arabidopsis gDNA (/Sp16 R) 117

pFLA16 F TTCGGTACCAGTCAACATTC Arabidopsis genomic DNA (gDNA) 2974 Sp16 R TTATTTGACCCGGTACTGG

pMigro F TCTCGAGGAATTCGGTACC pMigro vector (pFLA16F/Sp16 R) 2984

Sp16 F CCAGTACCGGGTCAAATAA FLA16 Arabidopsis stem cDNA 2054 FLA16 R CCATGGAGTGGATCCTCA

pMigro R AGAACTCCATGGAGTGGATC pMigro vector (Sp16 F/FLA16 R) 2060

VENUS_F GTGAGCAAGGGCGAGGAG Venus 713 VENUS_R CTTGTACAGCTCGTCCATGCC

16RVH_F TAGCTTGTAGAATGATGGGGTCAC Venus F/R, 16RV_TF/TC), 873 16RVH_R ATATCTCATTAAAGCAGGACTCTAGA VHpFu_TF/TC)

35s FLA16 His R CTCTAGAGGATCCTCAATGATGATGGT PCR (16RVH_F/R) 842

HVRFLA11F1 GCATGGACGAGCTGTACAAGGGTA HVRFLA11R1 HVRFLA11R1/ HVRFLA11TF/TC 100 GGGCCTGGAGCTGGAGCC

pFu_FLA11F GAGGACACGCTCGAGGAATTCATG Sp11 His TF/TC 170 HV_R TCGCCCTTGCTCACAAGCTTAGAG

pFLA11_F TTCGGTACCGCACACAC FLA11 Arabidopsis gDNA Sp11_R CTATAACGAGGAAGAAGATGAAG 443

pMigro_F TCTCGAGGAATTCGGTACC Sp11 R/PCR (pFLA11 F/Sp11 R) 453 HVHindIII_F GTGGCTCTAAGCTTGTGAGCAAGGG

HV11HindIII_R Venus AGCCTGacgAAGCTTCTTGTACAGC 757 SP11 F CTTCATCTTCTTCCTCGTTATAG FLA11_R Arabidopsis gDNA 1508 GGAGTGGATCCTTATATCCAC

pMigro R AGAACTCCATGGAGTGGATC pMigro vector (Sp11 F/FLA11 R) 1518 CACGCTCGAGGAATTCATGGAACATTCTCT 35s Sp12 F C Sp12 R/ At gDNA 88

pFLA12CM_F GTTGCGAGTGAAATATAAACAAGAGA Arabidopsis gDNA 2087 Sp12_R AGATAGGATTCCGGGAGTGGT FLA12 TCTCGAGGAATTCGGTACCGTTGCGAGTG pMigro_pFLA12 F AAAT pMigro vector (pFLA12CM_F/Sp12 R) 2117

Sp12_His F GGAATCCTATCTGGTACCCACCATCATCAT GGAGAGGGCTGACGAAGCTTCTTGTACAG Plasmid pFLA11:HVRFLA11 914 Venus FLA12 R CTCGTCC

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Table 2.2 DNA templates for generation of constructs. Template Sequence GTGAGCAAGGGCGAGGAGCTGTTCACCGGGGTGGTGCCCATCCTGGTCGAGCTGG ACGGCGACGTAAACGGCCACAAGTTCAGCGTGTCCGGCGAGGGCGAGGGCGATGC CACCTACGGCAAGCTGACCCTGAAGCTGATCTGCACCACCGGCAAGCTGCCCGTGC CCTGGCCCACCCTCGTGACCACCCTGGGCTACGGCCTGCAGTGCTTCGCCCGCTACC CCGACCACATGAAGCAGCACGACTTCTTCAAGTCCGCCATGCCCGAAGGCTACGTC CAGGAGCGCACCATCTTCTTCAAGGACGACGGCAACTACAAGACCCGCGCCGAGGT GAAGTTCGAGGGCGACACCCTGGTGAACCGCATCGAGCTGAAGGGCATCGACTTC Venus AAGGAGGACGGCAACATCCTGGGGCACAAGCTGGAGTACAACTACAACAGCCACA ACGTCTATCTCACCGCCGACAAGCAGAAGAACGGCATCAAGGCCAACTTCAAGATC CGCCACAACATCGAGGACGGCGGCGTGCAGCTCGCCGACCACTACCAGCAGAACA CCCCCATCGGCGACGGCCCCGTGCTGCTGCCCGACAACCACTACCTGAGCTACCAGT CCGCCCTGAGCAAAGACCCCAACGAGAAGCGCGATCACATGGTCCTGCTGGAGTTC GTGACCGCCGCCGGGATCACTCTCGGCATGGACGAGCTGTACAAG 16RV_TF TAGCTTGTAGAATGATGGGGTCACGGTTTATTCCGTGTCAGCGTGGTACCGCAGCA GTGAGCAAGGGCGAGGAGCTGTTCACCGGGGTGGT 16RV_TC ATCGAACATCTTACTACCCCAGTGCCAAATAAGGCACAGTCGCACCATGGCGTCGTC ACTCGTTCCCGCTCCTCGACAAGTGGCCCCACCA VHpFu_TF ATGGACGAGCTGTACAAGGGCGGAGGTGGCTCTCACCATCATCATCATCACCATCA CCATCATCACCATCATCATTGAGGATCCTCTAGAGTCCTGCTTTAATGAGATAT VHpFu_TC TACCTGCTCGACATGTTCCCGCCTCCACCGAGAGTGGTAGTAGTAGTAGTGGTAGT GGTAGTAGTGGTAGTAGTAACTCCTAGGAGATCTCAGGACGAAATTACTCTATA acacgctcgagGAATTCATGGCTACTTCAAGAACATTCATTTTCTCTAATCTCTTCATCTT Sp11 His F CTTCCTCGTTATAGCCACTACTTATGGTGGTACCcaccatcatcatcatcaccatcaccatcatc accatcatcatGGCGGAGGTGGCTCTAAGCTTGTGAGCAAGGGC tgtgcgagctcCTTAAGTACCGATGAAGTTCTTGTAAGTAAAAGAGATTAGAGAAGTAG Sp11 His R AAGAAGGAGCAATATCGGTGATGAATACCACCATGGgtggtagtagtagtagtggtagtggt agtagtggtagtagtaCCGCCTCCACCGAGATTCGAACACTCGTTCCCG

GCATGGACGAGCTGTACAAGGGTACCGGTGGAGGCGGTTCAGGCGGAGGTGGCT HVRFLA11 F CTGGCGGTGGCGGATCGCGTCCCGGGCAGGCTCCAGCTCCAGGCCC

CGTACCTGCTCGACATGTTCCCATGGCCACCTCCGCCAAGTCCGCCTCCACCGAGAC HVRFLA11 R CGCCACCGCCTAGCGCAGGGCCCGTCCGAGGTCGAGGTCCGGG

AGGAAGTTCATTTCATTTGGAGAGGACACGCTCGAGATGGAACATTCTCTCATCATC SpFLA12 F CTCCTCTTCACCGTCCTCCTCCTCCTCACCACCACTCCCGGAATCCTATCTGAATTCca ccatcatcatcatcaccatcaccatcatGGTGGAGGCGGTTCAGGCGGAGGTGGCTCTGGCG GTGGCGGATCGGAATTCATGGTGAGCAAGGGCGAGGAGCTGTTCACC TCCTTCAAGTAAAGTAAACCTCTCCTGTGCGAGCTCTACCTTGTAAGAGAGTAGTAG GAGGAGAAGTGGCAGGAGGAGGAGGAGTGGTGGTGAGGGCCTTAGGATAGACTT SpFLA12 R AAGgtggtagtagtagtagtggtagtggtagtaCCACCTCCGCCAAGTCCGCCTCCACCGAGA CCGCCACCGCCTAGCCTTAAGTACCACTCGTTCCCGCTCCTCGACAAGTGG

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Table 2.3 Primers for genotyping T-DNA insertions in FLA11, FLA12 and FLA16. Primer Sequence Ta (oC) Product (bp) T-DNA LBb1.3 ATTTTGCCGATTTCGGAAC 52 fla11 PEL166 CCACAGAGAAGAAGAAGCAGC 55 447-747/LBb1.3 (fla11)1 PEL165 TCATTTTCAATCCTCACCCAC 55 990/PEL166 (wt)2 fla12 PEL13 TAAATTCGTTAACCGATGAGCAACAAG 59 PEL16 AAGCTTTTCATCACAAATAAACCATGC 59 468/ PEL13 (wt) PEL14 ATTACACTCATGACGCAAAACGACA 59 PEL15 CAGGGAGAAAGGAGAGAAGAAGCAC 59 589/ PEL14 (fla12) fla16 PEL163ii AAGCTTTTCATCACAAATAAACCATGC 52 794/PEL164 (wt) PEL164 AATAGGCACACAAATGGATCC 52 536-836/LBb1.3 (fla16) 1fla mutant 2wt- wild-type

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Table 2.4 Primers for Q-PCR analysis. Gene Accession Primer Sequence Product (bp) Actin AT5G43500 Actin qF GAGTTCTTCACGCGATACCTCCA 180 Actin qR GACCACCTTTATTAACCCCATTTACCA Cyclophilin AT2G36130 Cyclophilin qF TGGCGAACGCTGGTCCTAATACA 223 Cyclophilin qR CAAAAACTCCTCTGCCCCAATCAA Tubulin AT1G50010 Tubulin qF ATGTGGGTCAGGGTATGGAA 143 Tubulin qR CCGACAACCTTCTTAGTCTCCTCT GAPdH AT3G26650 GAPdH qF TGGTTGATCTCGTTGTGCAGGTCTC 262

GAPdH qR GTCAGCCAAGTCAACAACTCTCTG FLA11 AT5G03170 FLA11 qF GAAAGGCGGCTCTGTTTCAA 102

FLA11 qR ATCCCAAACCCGAATCCAGT FLA12 AT5G60490 FLA12 qF TGATGATTCTCCGGCGGATG 109 FLA12 qR TCACAAATAAAACCATGCGAGCA FLA16 qF1 GGTCGGATTTCAGTTCAGGGT 150 FLA16 AT2G35860 FLA16 qR1 CCCCATCATTCTACAAGCTACCT

FLA16 qF2 TCCCACATAAAGTGTTGGCTCAAG 213 FLA16 qR2 CTCTTCTTGATTTAGAAACTTTCTTAACG CESA1 AT4G32410 CESA1 qF TGTTCTTCTCGCCTCCATCTTC 194 CESA1 qR CGGTTCACTGGGGTTTGATG CESA3 AT5G05170 CESA3 qF ACCAATCATGGGGACCACTCTTT 192 CESA3 qR TGACTCGGCTAGTGAAGGGATCA CESA6 AT5G64740 CESA6 qF TGCCAACCATTATTGTCGTCTG 188 CESA6 qR CCTCTCTTTGATCCGTGGGAATTA CESA4 AT5G44030 CESA4 qF CACCAGCCAAAGACGCATACC 162 CESA4 qR TCTTTTCCCCATCCCGTTCA CESA7 AT5G17420 CESA7 qF CAGCCCTTGGAGGAGCAGAA 178 CESA7 qR CCGCAGCTTATGACATGGATTG CESA8 AT4G18780 CESA8 qF GAGAGGGCTTGATGGGATTCA 169 CESA8 qR GGCAGGATCTTGGGGTTGTTT

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CHAPTER 3

TISSUE AND SUB-CELLULAR DISTRIBUTION OF FLA11, FLA12 AND FLA16 IN ARABIDOPSIS

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3.1 Introduction Genes encoding fasciclin-like arabinogalactan-proteins (FLAs) are predicted to have a N-terminal signal sequence, one or two fasciclin (FAS) domains, one to several regions with AGP glycomotifs and most are predicted to contain a C-terminal GPI-anchor signal sequence. The FAS domains have the potential to either form homophilic interactions (Elkins et al., 1990a), heterophilic interactions with other proteins (Hamilton, 2008) or with glycans (Nagae and Yamaguchi, 2012). Glycomotifs present in the protein backbone of FLAs can potentially be glycosylated with either N- or O-(AG) glycans. The glycan chains themselves can display heterogeneity, thereby providing enormous potential to act in molecular recognition and signalling pathways (Baldwin et al., 2001; Tan et al., 2012). The specific functions and potential cooperation between the different domains in FLAs remains largely unexplored (Shi et al., 2003; MacMillan et al., 2010; Johnson et al., 2011). A recent study of FLA4 showed that removal of the GPI- anchor does not affect function, the C-terminal (but not N-terminal) FAS1 motif is essential for function and the O- and N-glycosylation impact molecular trafficking (Xue et al., 2017). To understand how FLAs fulfil their molecular functions, investigation of their sub-cellular location, tissue- and cell-specific expression profiles can be highly informative. A number of FLAs show tissue-specific expression profiles and roles in the corresponding tissues. For example, Arabidopsis FLA11 and FLA12 have been shown to have specific expression patterns in the inflorescence stems by Promoter:GUS (MacMillan et al., 2010) and in situ hybridization (Ito et al., 2005). Arabidopsis FLA11 and FLA12 were shown to be the most highly expressed of all FLAs in the stem, and their homologs identified in other species such as Eucalyptus (MacMillan et al., 2010), Brassica (Li and Wu, 2012), Zinnia (Dahiya et al., 2006) and Populus (Lafarguette et al., 2004) also show predominant expression in the stem. Loss of function of FLA11 and FLA12 in Arabidopsis leads to altered stem biomechanical properties (MacMillan et al., 2010). The phenotype of fla11 fla12 mutants is subtle suggesting potential redundancy with other FLAs. FLA16 was also identified as being highly expressed in the stem with low levels of expression in other tissues (Schmid, 2005; MacMillan et al., 2010), suggesting a stem function for this FLA. This Chapter outlines the generation of FLA reporter constructs to gain insight into the tissue- and cell-type specific distribution of FLA11, FLA12 and FLA16. Enrichment and visualisation of tagged proteins extends the 46 studies undertaken by MacMillan et al. (2010) by providing information of their sub- cellular location(s) and post-translational modifications.

3.2 Results

3.2.1 Generation and validation of FLA reporter constructs Analyses of the predicted protein backbones of FLA11, FLA12 and FLA16 show FLA11 and FLA12 are highly similar, with more than 80% similarity (Johnson et al., 2003). Both contain a C-terminal GPI-anchor and a single FAS domain containing 4 glycomotifs for N-linked glycans (Fig. 3.1). In the AGP-like regions, 6 and 7 glycomotifs predicting O- linked glycans in FLA11 and FLA12, respectively, are present. FLA16 has a different protein backbone structure compared to that of FLA11 and FLA12, lacking a GPI-anchor signal sequence, but possessing 2 FAS domains each with a single glycomotif for N-linked glycans and 13 predicted AGP-glycomotifs outside of the FAS domains (Fig. 3.1). To facilitate visualisation and biochemical analyses of FLA11, FLA12 and FLA16, fusion proteins were generated with reporter and affinity tags, respectively. The coding region for each of FLA11, FLA12 and FLA16 was fused to the enhanced yellow fluorescence protein (YFP) variant, Venus (V) and poly-histidine (HIS/H) sequences. Since the proteins encoded by FLA11 and FLA12, but not FLA16, are predicted to possess a C-terminal GPI-anchor this influenced the positioning of the tags (Fig. 3.1). For FLA11 and FLA12 the reporters were positioned at the N-terminus downstream of the signal peptides to avoid disruption of the C-terminal GPI-anchor signal, whereas for FLA16 the tags were placed at the C-terminus. An arginine (Arg; R) residue was incorporated between the tags and the FLA sequences to create a trypsin cleavage site for subsequent proteomic analyses. These constructs were driven by either the 35S promoter (p35S:HV- FLA11, p35S:HV-FLA12, and p35S:FLA16-VH) or their respective endogenous promoters (pFLA11:HV-FLA11, pFLA12:HV-FLA12 and pFLA16:FLA16-VH) (Fig. 3.2). The p35S driven reporter fusion proteins were validated using a tobacco transient expression system (Van Loock et al., 2010) prior to stable transformation using endogenous promoters in Arabidopsis. In addition to their use in determining the sub-cellular location of FLA proteins, the fusions were generated for future studies investigating the post- translational modifications (PTMs). Such studies require enrichment of the proteins and several enrichment methods were evaluated including hydrophobic interaction 47 chromatography (HIC), chelation affinity and immunoprecipitation (IP) (see Appendices A3.1). The efficiency of these methods varied and depended on the tissue of origin and promoters used (see Appendices Figs. A3.1, A3.2, Table A3.1). To determine if the fusion proteins were correctly translated, the p35S driven FLA reporter proteins were extracted from tobacco leaves and visualised on Western blots using antibodies raised against GFP and HIS, respectively (Fig. 3.3A, B). The size of HV-FLA11 and HV-FLA12 were similar, yielding products of approximately 52-60 kDa and a diffuse band between 70-75 kDa. The predicted sizes of FLA11 and FLA12 fusion proteins are 52.4 kDa and 53.3 kDa, respectively, suggesting the lower MW products represented immature forms of FLA11 and FLA12 fusion proteins with little or no PTMs. This was supported by the identification of peptides matching the GPI-anchor signal sequence using mass spectrometry (see Appendices Table A3.1). Since both FLA11 and FLA12 are predicted to contain multiple sites for both N- and O-linked glycosylation (Fig 3.1), the higher MW products are likely to be the mature forms of glycosylated proteins. Two bands corresponding to FLA16-VH proteins were detected at approximately 60 and 75-85 kDa, respectively. Neither of the bands appeared as a smear and the higher MW product is larger than the predicted size of FLA16-VH proteins (76 kDa). This suggests the higher MW form contains PTMs. The lower MW band is potentially a proteolytically processed form of the FLA16-VH. FLA fusion proteins driven by their respective endogenous promoters were transformed into Arabidopsis. The HV-FLA11, HV-FLA12 and FLA16-VH fusion proteins were extracted from stem tissue and detected using Western blots (Fig. 3.3C). HV-FLA11 showed an increased molecular weight compared to that observed in tobacco with a smear of 75-100 kDa. A similar sized band occurred for HV-FLA12 with a smear of approx. 70-90 kDa. A faint lower MW band of approx. 60 kDa was visible for FLA16-VH and a more abundant higher MW product at approx. 90 kDa. The fusion proteins detected in the Arabidopsis stems were larger than in the tobacco leaves for both FLA11 and FLA16 but of similar size as the higher MW product for FLA12. These results suggest that the FLA fusion proteins are expressed in planta and that FLA11, FLA12 and FLA16 are glycosylated.

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3.2.2 Sub-cellular fractionation suggests FLA11 FLA12 and FLA16 are localised to the plasma membrane and walls Due to the presence of a GPI-anchor signal, FLA11 and FLA12 are predicted to be located at the PM whereas FLA16, which is not predicted to be GPI-anchored would be expected to be located in the apoplast. To investigate the sub-cellular location of FLA fusion proteins, stem tissue expressing pFLA11:HV-FLA11, pFLA12:HV-FLA12 and pFLA16:FLA16-VH was fractionated into different cellular compartments namely, wall, cytoplasm and microsomal/mixed membranes (MMs). The wall fraction was then washed with buffer to remove cellular contaminants, and then treated with detergent (sodium dodecyl sulphate (SDS)) to extract wall proteins. An antibody raised against the PM localised H-ATPase was used to investigate PM distribution. An anti-GFP antibody was used to detect the presence of the FLA fusion proteins in each of these fractions. With equal protein loading from each fraction, HV-FLA11 and HV-FLA12 were present with comparable abundance in the wall and MM fractions, with low levels of protein also detected in the cytoplasmic fraction. FLA16-VH was present in the wall and MM fractions with relatively greater abundance in the wall and was not detected in the cytoplasmic fraction. The anti-H-ATPase antibody showed the H-ATPase PM marker was present in both the wall and MM fractions. This suggests separation of the wall from the PM was not complete due to cross-contamination of different cellular compartments, eg. between wall components and PM proteins (Fig. 3.4). The location of FLA11 and FLA12 at the PM through a GPI-anchor is further supported by plasmolysis experiments in tobacco. Transient expression of p35S:HV- FLA11 or p35S:HV-FLA12 with plasma membrane intrinsic protein 2 (PIP2) tagged with a cyan fluorescent protein (CFP) (p35S:PIP2-CFP) show co-localisation at the membrane both before and after plasmolysis (Fig. 3.5). HV-FLA11 and HV-FLA12 were not detected in the wall. This may be due to using a transient expression system and/or because tobacco epidermal leaf cells have primary walls. The biochemical and microscopy data provide an indication of the potential location of FLA11 and FLA12 at the PM and in the wall. Transient expression of p35S:FLA16-VH and p35S:PIP2-CFP yielded a similar result to p35S:HV-FLA11 and p35S:HV-FLA12 suggesting FLA16 also associates with the PM. As FLA16-HV lacks a GPI-anchor, this was an unexpected result and the factors associating FLA16 with the PM are unclear.

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These preliminary results provide evidence for FLA11, FLA12 and FLA16 being located in both the wall and PM and glycosylated. To further examine the cell types these FLAs are present in throughout Arabidopsis development, an investigation of their expression patterns was undertaken.

3.2.3 Evaluation of FLA11, FLA12 and FLA16 reporters shows expression in tissues and cells with secondary walls

Previous studies of FLA11, FLA12 and FLA16 suggest they are expressed in a range of tissues and stages of development, with highest transcript abundance in the stem and silique, and lower levels in cotyledons, true leaves and seedling roots (Fig. 3.6A). This indicated they are largely stem-specific (Schmid, 2005; Winter et al., 2007; MacMillan et al., 2010). To verify the expression of FLA11, FLA12 and FLA16 in different regions of the stem, transcript levels were determined using Q-PCR analysis in the top, middle and basal segments of mature Arabidopsis stems (growth stage 6.5; (Boyes et al., 2001)). FLA11, FLA12 and FLA16 were expressed at similar levels in all regions of the stem (Fig. 3.6B). Interestingly, the transcript abundance of FLA16 between the first and second internode of the stem was lower than FLA11/FLA12 at growth stage 6.0, but progresses to a comparable level to FLA11/FLA12 at a later growth stage (6.5). This indicates the expression of these FLAs may be developmentally regulated during stem growth (Fig. 3.6). To gain information about their tissue location at the protein level, plants expressing reporter constructs, pFLA11:HV-FLA11, pFLA12:HV-FLA12 and pFLA16:FLA16-VH, were examined using confocal microscopy to visualise the YFP reporter. In the stems and first branch, a strong YFP signal was seen in cells with secondary walls such as the interfascicular fibres, xylem vessels, sclereids within the phloem, and starch sheath cells adjacent to interfascicular fibres. No signal was present in the cell types with only primary walls, including the epidermis, cortex and pith (Figs. 3.7, 3.8). This is largely consistent with previous pFLA11/12:GUS studies in Arabidopsis stems (MacMillan et al. (2010)), with the exception that MacMillan et al., observed a positive GUS signal in the pith cell layer adjacent to the interfascicular fibres. The tissue- specific expression profile of FLA16 has not been reported and this study provides the first evidence for overlapping expression of FLA16 with FLA11 and FLA12.

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Data obtained from the eFP browser for FLA11, FLA12 and FLA16 suggests these FLAs are also expressed at low levels in siliques and in other tissues (Fig. 3.6). The expression of pFLA16:FLA16-VH as well as pFLA11:HV-FLA11 and pFLA12:HV-FLA12 was therefore investigated in other tissues. Transverse sections of mature siliques showed YFP present in the endocarp b layer and replum of all lines (Fig. 3.9). A weak YFP signal is also seen in pFLA16:FLA16-VH lines in the developing seeds (Fig. 3.9). YFP was observed in cotyledons of 7 day old (d.o.) seedlings in plants expressing pFLA11:HV- FLA11, pFLA12:HV-FLA12 and pFLA16:FLA16-VH, specifically in the stomatal guard cells (Fig 3.10). In the hypocotyl and root of 4 d.o. seedlings, a weak YFP was visualized in the vasculature of all lines (Fig. 3.11). No YFP was detected in the epidermal pavement cells of cotyledons and true leaves of plants expressing pFLA11:HV-FLA11 and pFLA12:HV- FLA12. However, a YFP signal was observed in epidermal pavement cells in cotyledons and true leaves of plants expressing pFLA16:FLA16-VH (see Appendices Fig. A3.3). In addition, in plants expressing pFLA16:FLA16-VH, YFP was detected in the trichomes of true leaves (see Appendices Fig. A3.4). In summary, through the application of reporter proteins, we show that FLA11, FLA12 and FLA16 are predominantly located in cells with secondary walls, predominantly in inflorescence stems and at low levels in other tissues/organs (Table 3.1).

3.3 Discussion

Previous studies have investigated FLA11, FLA12 and FLA16 at the transcriptional level (Ito et al., 2005; Schmid, 2005; Winter et al., 2007; MacMillan et al., 2010). In this study, translational reporters for each of these FLAs were used to investigate their expression and protein location at the tissue and sub-cellular levels. Reporter proteins have been successfully and extensively employed for research of protein location and function in the past decades (Ghim et al., 2010; Zhong et al., 2017). Fusion proteins for FLA11, FLA12 and FLA16 enabled visualisation of protein location and enrichment from stems to reveal their likely glycosylation and subcellular localisation in the wall and PM of cells with secondary walls. Confirmation of their tissue and cellular distribution is discussed in relation to their domain characteristics and potential functions.

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3.3.1 FLA11, FLA12 and FLA16 are largely cell surface proteins (plasma membrane and/or cell wall) specific to cell types that develop secondary wall-specific The expression and location of FLA11, FLA12 and FLA16 proteins are tightly correlated with tissues/cells with secondary walls. The transition from a primary walls to secondary walls involves coordinated cessation of cell growth, the process of differentiation is initiated and, as a consequence secondary wall formation is initiated (Li et al., 2016a). The secondary walls are rich in cellulose, lowly-substituted hemicelluloses (eg. heteromannans and heteroxylans) and in many cases lignin, and have very low levels of pectin and proteins compared to primary walls (Bacic et al., 1988; Lodish et al., 2000; Doblin et al., 2010; Zhong and Ye, 2014). The activities of proteins involved in secondary wall biosynthesis are likely to be highly coordinated to enable efficient biosynthesis, secretion, deposition and assembly of the wall (Zhong and Ye, 2014). In vascular tissues, this results in a cell with a thicker, dehydrated and rigid wall that ultimately undergoes (Cosgrove, 2005; Doblin et al., 2010; Cosgrove and Jarvis, 2012). The onset of secondary wall development in stems is regulated by a complex transcription factor (TF) network consisting of master regulators including NACs, ATAFs, CUCs, SCWNs, VNDs and SNDs (see Chapter 1.1.2). The induction of FLA11, FLA12 and FLA16 expression in response to induction of VND6 and VND7, master-regulatory transcription factors of secondary wall development, indicates they are involved in early secondary wall development (Yamaguchi et al., 2010; Li et al., 2016b). In addition, co- expression studies show FLA11 and FLA12 are part of the co-expression network associated with secondary wall CesAs (Li et al., 2016b). In addition, expression of secondary wall FLAs could be regulated by other signals, such as stress (Houston et al., 2016)), hormone (Wang et al., 2017) or cell wall sensing (Xu et al., 2008) pathways. Hormones involved in regulation of secondary wall development include auxin which promotes development of xylem fibres in Arabidopsis (Ranocha et al., 2013; Didi et al., 2015) and abscisic acid (ABA) which is able to upregulate the secondary wall specific TF secondary wall-associated nac domain protein 1 (SND1) (Jensen et al., 2010; Kumar et al., 2016). A number of FLAs are regulated by phytohormones. For example, Arabidopsis FLA1 expression is up-regulated by auxin (Johnson et al., 2011); FLA4 is upregulated by ABA (Shi et al., 2003); and Poplar FLA6 is down-regulated when expression of gibberellin A3 (GA3) is restrained (Wang et al., 2017). Further work is 52 required to determine if hormones are involved in regulating the expression of FLA11, FLA12 and FLA16 in relation to a role in secondary wall development. This could include investigating if the expression level and pattern of FLA11, FLA12 and FLA16 are altered in response to hormone application. If hormones can be identified that are likely to regulate FLA expression, analyses of hormone responses in fla mutants could be undertaken. Another possible role for FLAs in secondary wall development could be through modulating the TF network through sensing feedback from the wall. AGPs and FLAs are implicated in cell wall sensing pathways, largely due to the presence of AG-glycans that can be differentially glycosylated and potentially act as signals (Schultz et al., 2000; Doblin et al., 2014). For example, in ROOT HAIR SPECIFIC 10 (RHS10), a member of the proline-rich extension-like receptor kinase family that contains AGP motifs in its extracellular domains. Domain deletion analysis shows some of the AGP-containing domains of RHS10 are required for its biological function in root hair inhibition (Cho, 2016; Hwang et al., 2016). It is suggested that glycosylation of the AGP domains contributes to the ability of RHS10 to sense changes in the wall and transduce signals to the cytoplasmic kinase domain for autophosphorylation (Cho, 2016). If FLA11, FLA12 and/or FLA16 are involved in wall sensing pathways, this could either result from protein-protein, or protein-glycan interactions, for example, with receptor-like kinases (RLKs) at the PM that in turn transduce signals intracellularly. This has been proposed for the GPI-anchored FLA4 which interacts with RLKs FEI1/FEI2 at the PM (Xu et al., 2008). FLA-interacting partners implicated in cell wall sensing might include either wall- associated kinases (WAKs) (Kohorn and Kohorn, 2012) or CrRLKs (Nibau and Cheung, 2011; Hou et al., 2016). Further study is needed to confirm if such interactions exist, whether or not the interaction with FLAs would activate the kinases as part of the signalling process, and the functional significance. A number of studies have shown that Arabidopsis FLA11/FLA12 and their homologues in other species (eg. Eucalyptus FLA1/FLA3 and Poplar FLA6) have regulatory roles in the biomechanical properties of the (MacMillan et al., 2010; MacMillan et al., 2015; Wang et al., 2017). Loss of FLA11 and FLA12 affects the polysaccharide composition in stem walls, particularly the production of cellulose (MacMillan et al., 2010), a key determinant of mechanical strength in both primary walls and secondary walls (Koehler and Telewski, 2006). Roles for FLAs in cellulose 53 biosynthesis is also supported by findings that the Poplar FLA6, which shows high sequence similarity to FLA11 and FLA12, was shown to play a crucial role in the formation and differentiation of tension wood (Wang et al., 2017). Tension wood consists of fibre cells encased by thickened walls rich in cellulose (Baba et al., 2009). Downregulation of PtFLA6 in the Poplar stem caused repression of tension wood formation (Wang et al., 2017). In addition, disruption of FLA4 in Arabidopsis leads to a cellulose reduction in the inner layer of seed coat mucilage (Harpaz-Saad et al., 2011). As members of AGP superfamily, characterised by AGP glycomotifs, a signal peptide, and for many, a GPI-anchor, FLA11, FLA12 and FLA16 are predicted to travel through the endomembrane system to the wall (see Chapter 1.3.2). Knowledge of the location of these FLAs at the PM or in the wall is useful to determine how they might act to regulate secondary wall deposition.

3.3.2 Sub-cellular localisation of FLA11, FLA12 and FLA16 in the PM and wall suggests interaction with wall proteins and/or carbohydrates and roles for the determination of wall properties Our data support the sub-cellular location of FLA11, FLA12 and FLA16 at the PM and in walls of xylem and fibre cells. Although cross-contamination of PM and wall fractions occurred, and contamination by other cellular compartments cannot be excluded, there is strong support for FLAs at both the wall and PM based upon fluorescent reporter protein studies (see Fig 3.5, 3.7, 3.8). It is extremely difficult to obtain ‘pure’ cell wall or PM preparations as shown by numerous proteomics studies (Cho et al., 2015; de Michele et al., 2016; Durufle et al., 2017). Besides this, in future experiments it is desire to include cytosolic protein markers such as tubulin and GTPase so that confirmation of target protein locations could be made (Drykova et al., 2003; Abbal et al., 2008). Repeated identification of FLA11, FLA12 and FLA16 in wall and PM fractions from multiple independent experiments and labs would provide greater confidence of their location. Glycosylation of FLAs is predicted based on the presence of glycomotifs for N- glycans in the FAS domain and AGP glycomotifs for O-linked AG glycosylation in other regions of the protein backbone (Kieliszewski and Lamport, 1994; Shpak et al., 1999; Shpak et al., 2001; Johnson et al., 2017a; Johnson et al., 2017b). Previous studies suggest FLAs (Johnson et al., 2003; Basu et al., 2016; Wang et al., 2017), and other 54 chimeric AGPs (Hijazi et al., 2012; Poon et al., 2012) are indeed glycosylated. Through biochemical approaches, Johnson et al. (2003) demonstrated the presence of FLA2, FLA7 and FLA8 in samples precipitated with β-glucosyl Yariv reagent suggesting O-linked glycosylation occurs on these FLAs (Johnson et al., 2003). FLA4 is also proposed to be glycosylated based on a shifted protein size after enzymic treatment with enzymes that cleave AG-glycans (Xue et al., 2017). In support of this, Basu et al. (2016) showed that GALT2 and GALT5 are involved in AG glycan biosynthesis, glycosylate FLA4, and confirmed the glycosylation is required for seed coat mucilage adherence and root growth. Our results suggest that FLA11, FLA12 and FLA16 are likely glycosylated. A high molecular weight smear on protein blots larger than that predicted for the protein backbone alone was observed and is typical for glycoproteins (Dixon et al., 1990; Yang et al., 2015). It is currently unclear if both N- and O-linked glycosylation occurs on FLA11, FLA12 and FLA16. Confirmation of the glycosylation status of these FLAs requires further analyses such as staining of glycans using periodic acid-Schiff (PAS) staining method (Van-Seuningen and Davril, 1992) and enzyme digestions (Knoch et al., 2013; Ford et al., 2015). Ultimately the aim is to fully characterise the glycan structure on individual FLAs. To date, the glycan structures of mixed populations of AGPs have been determined (Tryfona et al., 2012). Characterisation of the glycans on an individual FLA would provide greater insight into how they function. Several enrichment methods (i.e. IP, HIC and chelation) have been trialled (see Appendices A3.1). Enrichment of these FLAs will enable more detailed analyses of their structures using mass-spectrometry based analyses (Tryfona et al., 2012; Ratnayake et al., 2016; Wilkens et al., 2016). The overall function(s) of glycosylation on FLAs is unclear. N-linked glycosylation is proposed to contribute to proteins in different ways, namely stabilising protein folding/tertiary structure and facilitating protein trafficking between the sub-cellular compartments (Lee et al., 2015). The role of O-linked glycosylation is suggested to facilitate interaction either with other extracellular molecules (Jentoft, 1990), potentially cell wall polysaccharides (Hijazi et al., 2014) or PM located proteins (Chen et al., 2010; Suzuki et al., 2017). O-linked glycosylation has also been proposed to assist secretion and trafficking of the protein so that it is delivered to its functional site (Zhang et al., 2014). The exact role of glycosylation for FLA11, FLA12 and FLA16 function(s) requires in depth functional investigation such as deletion and substitution of N- and O-glycosylation sites. Nevertheless, our data suggests that if the shift in MW is due to glycosylation, the degree 55 of glycosylation and glycan heterogeneity is differentially regulated for FLA11, FLA12 and FLA16. This may arise due to differences in the glycomotifs and/or yet to be identified regulatory domains in other regions of the protein (Shpak et al., 1999; Goodrum et al., 2000; Shpak et al., 2001). To assess how each functional domain influences the biological function of FLA11, FLA12 and FLA16, domain deletion/swap studies could be undertaken (Sugimoto et al., 2003; Xue et al., 2017). For example, deletion of the GPI-anchor signal sequences of FLA11/FLA12 and complementation of the fla11 fla12 mutant could be undertaken to determine if the GPI-anchor is important for biological function. Similarly, deletion and substitution of glycosylation sites as mentioned above or FAS domains and complementation studies would determine their importance for FLA stability, location and function. All FLAs are predicted to contain a N-terminal signal peptide (Schultz et al., 2002; Johnson et al., 2003), presumably directing FLAs to the PM/wall as occurs for other AGPs (Coleman et al., 1985; Schultz et al., 2000). A PM location for AGPs can also be predicted by the presence of a C-terminal GPI-anchor on the protein backbone (Schultz et al., 1998; Schultz et al., 2000; Schultz et al., 2002). FLA11 and FLA12 are predicted to contain a GPI-anchor (Johnson et al., 2003) and our study suggests they are located at the PM. The GPI-anchor gives an alternative means to transmembrane domains for the proteins to locate on the outer leaflet of the PM and might provide other functions such as changed mobility on the PM and signal transduction (Solomon et al., 1996; Hooper, 1997; Peles et al., 1997; Schultz et al., 1998). These functions have in part been proposed due to the affinity of GPI-anchors for lipid rafts or microdomains rich in glycosphingolipids (Simons and Toomre, 2000; Rajendran and Simons, 2005). These microdomains provide a platform for the GPI-anchored proteins and signalling proteins to associate and fulfil diverse cellular functions such as signal transduction and protein trafficking via vesicles (Simons and Toomre, 2000; Rajendran and Simons, 2005). A recent study showed that FLA4 is localised to the PM and the GPI-anchor is critical for PM location (Xue et al., 2017). Previous studies of other GPI-anchored AGPs, for example Arabidopsis AGP18 (Yang and Showalter, 2007; Zhang et al., 2011) and AGP1 (Sun et al., 2004), show they are located at the PM. The importance of the GPI-anchor on FLA11 and FLA12 for function is yet to be determined, but it is likely a GPI-anchor locates these FLAs to the PM. GPI-anchored proteins could undergo different fates including; anchor cleavage for release into the wall, degradation and/or endocytosis (Censullo and Davitz, 1994). Many 56

GPI-anchored proteins in plants are released from the PM and eventually incorporate into the wall (Kinoshita, 2016). The physical association of AGPs with flax fiber cellulose in secondary walls has led to the proposal that GPI-anchored AGPs might be secreted to the cell surface along with CesAs and then associate with cellulose in the thickening wall (Girault et al., 2000; Seifert and Roberts, 2007). The identification of FLA11 and FLA12 in the wall in the current study is potentially due to the release of these FLAs from the PM via either enzymatic (eg. phospholipase C/D) digestion of the GPI-anchor or physical detachment from the PM ((Schultz et al., 1998) and Chapter 1.3.3). FLAs could potentially bind to other wall polysaccharides such as hemicellulose and pectins as has been shown for the AGP, APAP1 (Huang et al., 2013). This might occur via transglycosylation to cross-link the AG glycans of FLAs to the wall polysaccharides (Hurtado-Guerrero et al., 2009; Johnston et al., 2013). In contrast to FLA11 and FLA12 (group A), FLA16 (group B) is not predicted to be GPI-anchored and possesses two FAS domains (Johnson et al., 2003). From the present study the final location of FLA16 is unclear. In protein extractions FLA16 was present in both PM and wall fractions, however cross-contamination of PM into the wall fraction makes it difficult to assign where FLA16 is ultimately located. Further analysis with TEM would provide more accurate conclusions to be made as to the sub-cellular location of FLA16 in planta. FAS containing proteins are commonly extracellular in a range of organisms (Kim et al., 2002; Clout et al., 2003; Johnson et al., 2011). The FAS domain is shown to have cell-adhesion properties in broad taxonomical classes (eg. algae, insects and humans) and involved in both homophilic and heterophilic interactions (Elkins et al., 1990a; Huber and Sumper, 1994; Kim et al., 2000; Kim et al., 2002; Clout et al., 2003). In plants, the role of FAS domains is poorly understood. Considering the potential for both protein-protein and carbohydrate-protein interactions, the FAS domains of FLA16 could interact with other FLAs and/or PM/wall-located proteins and polysaccharides. Further work is necessary to define both FAS domains and FLA16 function. Furthermore, the N-terminal FAS domain of FLA4 was shown to be critical in assisting PM localisation, independently from the GPI-anchor, and the C-terminal FAS is required for function in roots (Xue et al., 2017). Given the limited functional studies available for the plant FAS domains to date, this research gives valuable insight into the potential co-operation of the N- and C-terminal FAS domains within the same FLA. It is possible the N-terminal FAS domain in FLA16 might act in a similar way as for FLA4 and facilitate PM association 57 whereas the C-terminal FAS may have a different function. Visualisation of the trafficking and secretion of FLA16 as well as further investigation of its functional domains through deletion/swapping studies would be highly informative. Expression of FLA11, FLA12 and FLA16 overlaps in most of the tissues examined in the current study indicating they are present in cells with secondary walls. Our results for FLA11 and FLA12 are consistent with Promoter:GUS studies undertaken in MacMillan et al. (2010). We present novel data on the tissue-specific location of FLA16 in cells containing secondary walls and the potential overlapping location of these FLAs at the PM and wall. In comparison to FLA11 and FLA12, much less is known about the biological roles of FLA16. Therefore, we undertook further study of FLA16 and its potential role in stem development and this is reported in Chapter 4.

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Table 3.1 Tissue-specific expression patterns of FLA11, FLA12 and FLA16 in Arabidopsis. FLA11 FLA12 FLA16 Transcript abundance in stemsa Top 1.20b (± 0.10)c 1.50 (± 0.06) 1.30(± 0.43) Middle 1.80 (± 0.13) 1.00 (± 0.15) 1.40 (± 0.39) Basal 1.50 (± 0.08) 0.80 (± 0.03) 1.00 (± 0.28) FLA reporter protein distribution Stemd Xylem, Starch sheath, Xylem, Starch sheath, Xylem, Starch sheath, Interfascicular fibre, Interfascicular fibre, Interfascicular fibre, Sclereid Sclereid Sclereid Branche Xylem, Starch sheath, Xylem, Starch sheath, Xylem, Starch sheath, Interfascicular fibre, Interfascicular fibre, Interfascicular fibre, Sclereid Sclereid Sclereid Cotyledonf Guard cell Guard cell Guard cell, Pavement cell True leafg Guard cell Guard cell Guard cell, Pavement cell, Trichome Siliqueh Endocarp b, Lateral Endocarp b, Lateral Endocarp b, Lateral vascular bundle, Replum vascular bundle, Replum vascular bundle, Replum Rooti Vasculature Vasculature Vasculature a,d,e Growth stage 6.5 (Boyes et al., 2001) b Relative expression level determined by Q-PCR (see Fig 3.6) c Standard error of 2 biological replicates. f,g Growth stage 1.02 h Fully elongated I Growth stage 1.0

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Figure 3.1 Protein sequences and predicted domains of Arabidopsis FLA11, FLA12 and FLA16. Proteins encoded by Arabidopsis FLA11 and FLA12 are predicted to have a N-terminal signal peptide (grey), a fasciclin (FAS) domain (blue), a GPI-anchor signal sequence (purple) and the omega (ω) cleavage site amino acid for addition of a GPI-anchor (green). Glycomotifs for N-linked glycans (orange) occur in FAS domains whereas regions with AGP-like glycomotifs (red text) that are predicted to direct addition of large O-linked glycans occur in regions bordering the FAS domains. FLA16 is predicted to have 2 FAS domains, each with a single N-glycomotif, a central AGP-glycomotif rich region and lacks a GPI-anchor signal sequence.

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Figure 3.2 Schematic representation of FLA11, FLA12 and FLA16 constructs. Reporter constructs for FLA11 (HV-FLA11), FLA12 (HV-FLA12) and FLA16 (FLA16-VH) were engineered with an endogenous signal peptide (SP), 14x poly-histidine (HIS/H) and a variant of YFP, Venus (V). Constructs are driven by either the 35S promoter (p35S, red arrows) or their respective endogenous FLA promoters (pFLA, green arrows). An empty- vector-control construct (pFLA11:HV) was generated as a control. Constructs used to complement the fla16 mutant (see Chapter 4) included the genomic DNA sequence of the FLA16 gene (pFLA16:FLA16) or the FLA16 coding region fused to Venus and HIS (pFLA16:FLA16-VH).

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Figure 3.3 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins extracted from tobacco and Arabidopsis. HV-FLA11, HV-FLA12 and FLA16-VH fusion proteins transiently expressed in tobacco leaves under the 35S promoter were detected by anti-GFP (A) and anti-HIS (B) antibodies in protein extracts. Two products corresponding to HV-FLA11 are detected at approximately 52-60 kDa and 70-75 kDa, HV-FLA12 at 55-60 kDa and 70-75 kDa, and FLA16-VH at 60 kDa and 75-85 kDa. (C) HV-FLA11, HV-FLA12 and FLA16-VH fusion proteins driven by their endogenous promoters were detected in proteins extracted from Arabidopsis stems (growth stage 6.5) (Boyes et al., 2001) by anti-GFP antibodies at sizes of 75-100 kDa for HV-FLA11, 70-90 kDa for HV-FLA12 and about 60 kDa and 90 kDa for FLA16-VH. A total of 30 µg of protein is present in each sample.

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Figure 3.4 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins in sub-cellular fractions of proteins extracted from mature Arabidopsis stems. Proteins extracted from stems were fractionated into: Wall wash representing the soluble phase after washing the cell walls with buffer; Wall SDS representing the soluble phase after treatment of the walls with SDS; Cytoplasm; and MM SDS representing the soluble phase after treatment of microsomal membranes (MMs) with SDS. Soluble proteins (30 µg) were loaded from each fraction for separation by denaturing protein electrophoresis and Western blotting. (A) Detection of HV-FLA11, HV-FLA12 and FLA16-HV by an anti-GFP antibody shows they are most abundant in Wall SDS and MM SDS fractions and present in the Wall wash fraction with lower abundance. A much lower signal is also detected in the Cytoplasm fraction of HV-FLA11 and HV-FLA12. FLA16-VH is most abundant in the Wall SDS and to a lesser extent in the MM SDS fraction. (B) Replicate western blots probed with the anti- H-ATPase plasma membrane (PM) marker antibody shows the PM is found in both the Wall SDS and MM SDS fractions.

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Figure 3.5 HV-FLA11, HV-FLA12 and FLA16-VH are co-localised with plasma membrane intrinsic protein 2 (PIP2) in turgid and plasmolysed tobacco leaf cells. Transient expression of p35S:PIP2-CFP, coding for PM intrinsic protein 2 (PIP2) tagged with cyan fluorescent protein (CFP, blue) (A1-F1) co-expressed with p35S:HV-FLA11 (A2, B2), p35S:HV-FLA12 (C2, D2) or p35S:FLA16-VH (E2, F2) visualized for YFP (yellow) in tobacco leaves. A3-F3 show the merged images overlayed with transmitted light. HV- FLA11, HV-FLA12 and FLA16-VH signal was found evenly distributed around the periphery of the cell and overlaps with that of the PM marker PIP2-CFP in both turgid (pre-plasmolysed, Pre-p) and plasmolysed (P) cells (white arrows). Scale bar = 10 µm.

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Figure 3.6 Transcript abundance of FLA11, FLA12 and FLA16 in Arabidopsis tissues. (A) Expression levels of FLA11, FLA12 and FLA16 from the Arabidopsis developmental data set of Schmid (2005) visualised via the eFP browser in a range of Arabidopsis tissues at different growth stages (Boyes et al., 2001) as indicated in brackets. (B) Q-PCR analysis of transcript abundance of FLA11, FLA12 and FLA16 in the top, middle (mid) and basal (base) segments of Arabidopsis stems (growth stage 6.5) (Boyes et al., 2001). Transcript levels were determined relative to DNA standards of known concentration and normalized with GAPDH, tubulin and cyclophilin housekeeping genes (Czechowski et al., 2005). Data are obtained from two biological replicates with three technical replicates each. Error bars represent ± SD in (A) and ± SE in (B).

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Figure 3.7 Detection of YFP in plants expressing pFLA11:HV-FLA11, pFLA12:HV- FLA12 and pFLA16:FLA16-VH reporter proteins in transverse sections taken at the first internode of mature Arabidopsis stems. Arabidopsis transgenic plants are grown to stage 6.5 (Boyes et al., 2001). Visualisation of YFP fluorescence show HV-FLA11 (B1-6), HV-FLA12 (C1-C6) and FLA16-VH (D1-D6) reporter proteins present in cells with secondary cell walls, including interfascicular fibre (IF), xylem (Xy), starch sheath (SS) and phloem fibre sclereids (S), whereas no YFP is observed in wt (A1-A6). Autofluorescence (Auto) (A2, B2, C2 and D2) is seen in the cortex and does not overlap with the YFP signal in merged images overlaid with transmitted light (Trans) images at 10x (A4-D4) and 63x magnification (A5-D5). Scale bar = 100µm.

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Figure 3.8 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins in the first branch of mature Arabidopsis stems. Arabidopsis transgenic plants are grown to stage 6.5 (Boyes et al., 2001). YFP signal in plants expressing pFLA11:HV-FLA11 (B1-B6), pFLA12:HV-FLA12 (C1-C6) and pFLA16:FLA16-VH (D1-D6) was observed in cells with secondary cell walls, including interfascicular fibre (IF), xylem (Xy), starch sheath (SS) and phloem fibre sclereids (S), whereas no YFP is observed in wt (A1-A6). Autofluorescence (Auto) (A2, B2, C2 and D2) is seen in the cortex and does not overlap with the YFP signal as observed when YFP, auto and transmitted light (Trans) channels are merged in 10x (A4-D4) and 63x (A5-D5) magnification images. Scale bar = 100µm.

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Figure 3.9 Detection of HV-FLA11, HV-FLA12 and FLA16-VH in transverse sections of Arabidopsis siliques. Fully elongated Arabidopsis siliques expressing pFLA11:HV-FLA11, pFLA12:HV-FLA12, pFLA16:FLA16-VH and wild-type (wt) visualised for YFP fluorescence (A1-D1), auto fluorescence (Auto, A2-D2), and transmitted light (Trans) (A3-D3). Merged channels (A4-D4) of YFP, Auto and Trans at 10x. Boxed areas at 63x magnification show YFP is present in the endocarp b (En b) layer, lateral vascular bundle (LVB) (1: A5, B5, C5 and D5) and replum (R) (2: A6, B6, C6 and D6) regions, but not septum (S) and valve (V). YFP channel in left panels of A5-D5 and A6-D6, and merged channels in right panels of A5- D5 and A6-D6. Scale bar = 100µm.

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Figure 3.10 Detection of HV-FLA11, HV-FLA12 and FLA16-VH reporter proteins in the guard cells of 7 d.o. Arabidopsis cotyledons. A weak YFP signal is detected in the cotyledons of 7 d.o. seedlings expressing pFLA11:HV- FLA11, pFLA12:HV-FLA12, pFLA16:FLA16-VH with no signal observed in wt (A1-D1). Merged images (A4-D4) of YFP with autofluorescence (Auto A2-D2), overlaid with transmitted light (Trans) images (A3-D3) shows HV-FLA11 and HV-FLA12 are evenly present on the inner wall of guard cells, and FLA16-VH is present on both inner and outer sides with higher intensity at the polar end wall of paired guard cells. Scale bar = 100µm.

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Figure 3.11 Detection of HV-FLA11, HV-FLA12 and FLA16-VH in the Arabidopsis root vasculature of 4 d.o. seedlings. Roots of 4 d.o. seedlings (growth stage 1.0) expressing pFLA11:HV-FLA11, pFLA12:HV- FLA12, pFLA16:FLA16-VH are visualised under a confocal microscope using wild-type (wt) as control. (A1-D1) transmitted light (Trans) channel at low magnification (10x), (A2-D2) YFP channel at 63x magnification. (A3-D3) merged channels of YFP and BF, both at 63x magnification showing YFP is present in the vasculature of all FLA reporter lines in selected area with low intensity. Scale bar = 100µm.

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APPENDICES TO CHAPTER 3

A3.1 Enrichment of FLA11 reporter proteins In order to obtain information about the post-translational modifications and potential interacting partners of FLA11, FLA12 and FLA16, a number of enrichment methods were trialled. These included a chelation method, immunoprecipitation (IP) and hydrophobic interaction chromatography (HIC). The HV-FLA11 fusion protein was chosen for investigation and the detergent solubilized microsomal fraction was used as starting material for these enrichment trials. FLA11 was chosen as it is predicted to be GPI-anchored hence bound to the PM, and has been shown to localise to membranes (Fig 3.5). Tobacco leaf tissue overexpressing p35S:HV-FLA11 was used in this study. The chelation method employed the specific chelation bonding between the poly histidine tag and Ni2+ resin (Bornhorst and Falke, 2000). The elution of HV-FLA11 from the Ni resin was detected using fluorescence detection, denatured electrophoresis separation, Western blots and liquid chromatography mass spectrometry (LC-MS). A yellow fluorescence signal was not detected by LED transillumination in the unbound and wash fractions and eluates in lower (100 mM) and higher (500 mM) concentrations of imidazole (Fig A3.1). A signal was specifically detected in the 200 mM imidazole eluate indicating a strong interaction of HV-FLA11 with the resin (Fig A3.1). Trypsin digestion and LC-MS analysis of proteins in the 200mM imidazole eluate identified two peptides covering 8.5% of the entire HV-FLA11 sequence, ranking it the third amongst all Arabidopsis proteins identified in the search list (Table A3.1, A3.2). The IP method utilised the immuno-specificity between the YFP tag and anti-GFP antibodies conjugated to micro-spherical agarose beads (Chromotek, USA). Elution from the beads was found to have a very low abundance of all bound proteins compared to other methods. No FLA11 was detected on western blots, whereas using LC-MS analysis in high sensitivity mode, FLA11 was identified (Table A3.1). Only one peptide was identified covering 4.5% of the entire HV-FLA11 sequence but the ranking was first in the search list (Table A3.1, A3.2). The HIC method employed the hydrophobic interaction between the YFP tag and phenylsepharose under specific salt concentrations (Peckham et al., 2006). Detection

77 and analyses of proteins eluted from the sepharose used the same techniques as the chelation method mentioned above. Similar to the chelation results, a specific YFP signal was detected in the elution with no signal in the unbound and wash fractions. This was supported by Western blots (Fig. A3.2). LC-MS analysis detected two peptides covering 8.5% of the FLA11 protein sequence, and the ranking was sixteenth in the search list (Table A3.1, A3.2). A combined method using chelation followed by immunoprecipitation was also trialled. Three peptides were identified covering 11.8% of the entire HV-FLA11 sequence ranking the first in the search list (Table A3.1, A3.2). These trials suggested that chelation and immunoprecipitation in succession are the most effective enrichment method. However, when these methods were applied to the Arabidopsis stem tissue, HV-FLA11, HV-FLA12 and FLA16-VH were detected at very low rankings (40-1000) with few peptides identified, and in some cases not detectable. This is possibly due to the lower abundance of FLA proteins in the stem and/or located in the wall rather than at the PM. Further enrichment procedures are therefore required prior to the application of these methods otherwise new approaches are needed.

A3.2 A YFP signal corresponding to FLA16-HV is detected in pavement cells and trichomes in leaves of young seedlings. Visualisation of YFP in plants expressing pFLA16:FLA16-VH in Arabidopsis identified a signal in 7 d.o. seedlings. A YFP signal was found to be located in the pavement cells of cotyledons and true leaves, as well as trichomes in the true leaves (Fig. A3.3, 3.4). The distribution of FLA16-VH reporter protein aligns with the transcript distribution data by Marks et al. (2009) and Gilding and Marks (2010).

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Table A3.1 Mass spectrometry identification of FLA11 peptides after different enrichment methods.

Method Ranking Peptides (ion score) Coverage (%) GFP-Trap 1 SGTLNSLSDQQ (66)a 4.5% Ni2+ resin 3 AGQFTLFIR (62), 8.5% SGTLNSLSDQQ (58) Phenylsephrose 16 AGQFTLFIR (60), 8.5% SGTLNSLSDQQ (62) Ni2+ resin/GFP-Trap 1 AGQFTLFIR (60), 11.8% SGTLNSLSDQQ (62), TGFGFGIRb (35) Peptide coveragec MATSRTFIFSNLFIFFLVIATTYGQAPAPGPSGPTNITAILEKAGQFTLFIRLLKSTQASDQINTQLNSSSS NGLTVFAPTDNAFNSLKSGTLNSLSDQQKVQLVQFHVLPTLITMPQFQTVSNPLRTQAGDGQNGKFP LNITSSGNQVNITTGVVSATVANSVYSDKQLAVYQVDQVLLPLAMFGSSVAPAPAPEKGGSVSKGSA SGGDDGGDSTDSSDAERTGFGFGIRITTVAAIAASSSLWI aion score bGPI-anchor recognition signal sequence. cPeptides identified by mass spectrometry are shown underlined in the FLA11 protein sequence. Coloured regions are: grey=signal peptide, blue=FAS domain, green=GPI-anchor attachment site, purple=GPI-anchor recognition sequence. Coloured text are: red=O-linked glycomotifs, orange=N- linked glycomotifs.

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Table A3.2 Proteins identified by mass spectrometry after enrichment by Chelation, IP and HIC of HV-FLA11. Method Proteina Chelation (HIS) AT1G23410 Ribosomal protein s27a AT4G10340 Light harvesting complex of photosystem II 5 AT5G03170 Fasciclin-like arabinogalactan-protein 11 Immunoprecipitation (GFP-Trap) AT5G03170 Fasciclin-like arabinogalactan-protein 11 AT5G64040 Photosystem I reaction center subunit PSI-N AT1G07920 GTP binding elongation factor Tu family protein HIC (Phenylsepharose) ATCG00490 Ribulose-bisphosphate carboxylases AT1G53310 Phosphoenolpyruvate carboxylase 1 AT3G50820 Extrinsic subunit of photosystem 2 AT3G14940 Phosphoenolpyruvate carboxylase 3 AT2G42600 Phosphoenolpyruvate carboxylase 2 AT3G03780 Cytosolic methionine synthase 2 AT3G12390 Nascent polypeptide-associated complex (NAC), alpha subunit family protein AT5G28540 Luminal binding protein bip AT5G66190 Leaf-type ferredoxin:NADP(H) oxidoreductase AT5G49910 Chloroplast heat shock protein 70-2 AT1G66410 Calmodulin 4 AT3G52880 Peroxisomal monodehydroascorbate reductase AT2G21330 Fructose-bisphosphate aldolase 1 AT2G36530 Enolase AT5G13850 Nascent polypeptide-associated complex subunit alpha-like protein 3 AT5G03170 Fasciclin-like arabinogalactan-protein 11 AT5G35630 Chloroplastic glutamine synthetase Chelation and IP AT5G03170 Fasciclin-like arabinogalactan-protein 11 AT1G23410 Ribosomal protein s27a aProteins identified by mass spectrometry are ranked by abundance in the total sample.

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Figure A3.1 Enrichment of HV-FLA11 from tobacco leaves using a chelation method. (A) After chelation, YFP was detected in the 200mM imidazole elution (indicated by *) and not in unbound (UB), wash, and eluates from different imidazole concentrations using LED trans-illumination (400-500 nm). (B) Western blot of proteins corresponding to fractions in (A) and total proteins extracted from tobacco before chelation (neat), detected by an anti-GFP antibody. HV-FLA11 is shown to be enriched in the 200mM imidazole elution.

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Figure A3.2 Enrichment of HV-FLA11 from tobacco leaves using a hydrophobic interaction chromatography (HIC) method. Transiently expressed p35:HV-FLA11 in tobacco were extracted and bound to phenylsepharose resin then eluted in a salt gradient as shown in (A). YFP detected using a LED transilluminator shows a signal in fractions indicated by an (*) and is not present in the unbound (UB) or wash. (B) HV-FLA11 was detected using an anti-GFP antibody on Western blots of fractions indicated in (A).

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Figure A3.3 Detection of FLA16-VH in the epidermal pavement cells of Arabidopsis cotyledons and true leaves. Epidermal pavement cells in the cotyledons and true leaves of 7 d.o. seedlings (growth stage 1.02) (Boyes et al., 2001) expressing pFLA16:FLA16-VH examined under a confocal microscope using wild-type (wt) as a control. (A1-D1) YFP fluorescence, (A2-D2) auto fluorescence (Auto), (A3-D3) bright field (BF) and (A4-D4) merged channels. Scale bar = 20µm.

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Figure A3.4 Detection of FLA16-VH in the Arabidopsis true leaves. The first true leaves of 7 d.o. seedlings (growth stage 1.02) (Boyes et al., 2001) of wt and transgenic plants expressing pFLA16:FLA16-VH are visualised under a confocal microscope in the YFP channel for fluorescent detection. (A1) YFP signal is undetectable in wt sample (10x magnification). (B1) YFP signal is detected in trichomes (Tr) and epidermal (Ep) cells (10x). (C1) A trichome expressing pFLA16:FLA16-VH as boxed in B1 Tr (63x). (D1) Pavement cells (white arrows) and guard cells (red arrows) expressing pFLA16:FLA16-VH as boxed in B1 Ep (40x). (A2-D2) Images are merged for the YFP, auto fluorescence and transmitted light channels. Scale bar = 100µm in 10x, 10µm in 63x and 20µm in 40x.

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Figure A3.5 Expression pattern of FLA11 during Arabidopsis development. Tissue distribution and transcript abundance of FLA11 in Arabidopsis using developmental data sets of Schmid (2005) with data extraction tool by Toufighi (2005) and visualized using the eFP browser (Winter et al., 2007). Expression levels are represented in the colour gradient from yellow (lowest) to red (highest).

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Figure A3.6 Expression pattern of FLA12 during Arabidopsis development. Tissue distribution and transcript abundance of FLA12 in Arabidopsis using developmental data sets of Schmid (2005) with data extraction tool by Toufighi (2005) and visualized using the eFP browser (Winter et al., 2007). Expression levels are represented in the colour gradient from yellow (lowest) to red (highest).

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Figure A3.7 Expression pattern of FLA16 during Arabidopsis development. Tissue distribution and transcript abundance of FLA16 in Arabidopsis using developmental data sets of Schmid (2005) with data extraction tool by Toufighi (2005) and visualized using the eFP browser (Winter et al., 2007). Expression levels are represented in the colour gradient from yellow

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CHAPTER 4

FUNCTIONAL INVESTIGATION OF ARABIDOPSIS FASCICLIN-LIKE ARABINOGALACTAN-PROTEIN 16 (FLA16)

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4.1 Introduction FLA11 and FLA12 are predominantly expressed in stems and are the most highly expressed FLAs in this tissue (Schmid, 2005; Toufighi, 2005; Winter et al., 2007) (see Appendices Fig. A3.5, A3.6) (MacMillan et al., 2010). Knockout mutants of Arabidopsis FLA11 and FLA12 displayed differences in biomechanical properties, wall architecture and carbohydrate composition of the stems (MacMillan et al., 2010). fla11 and fla12 single mutants showed no major phenotypic differences whereas fla11 fla12 double mutants showed a reduction in tensile strength and stiffness, increased cellulose microfibril angle (MFA) and reduced cellulose, galactose and arabinose content (MacMillan et al., 2010). This suggests that FLA11 and FLA12 contribute to the maintenance of stem biomechanical properties via regulation of wall architecture and/or biosynthesis (MacMillan et al., 2010). Overexpression of Eucalyptus FLA2, the putative orthologue of Arabidopsis FLA12, resulted in a reduction of MFA by three degrees in the Eucalyptus xylem fibres, and the heterologous overexpression of Eucalyptus FLA3, the putative orthologue of Arabidopsis FLA11, in tobacco caused reduced flexural strength in the stem (MacMillan et al., 2015). Furthermore, suppression of Populus PtFLA6, which shows high sequence similarity to Arabidopsis FLA11 and FLA12 using antisense RNA led to a reduction of flexural strength and stiffness in the stem, suggesting PtFLA6 influences the biomechanics of the Populus stem (Wang et al., 2015). Changes in cellulose levels and cellulose MFA in stems of fla mutants suggests a relationship between FLAs and cellulose synthesis or deposition.

Cellulose provides mechanical strength to walls and is synthesised by cellulose synthases (CesA) located at the PM that are organised into cellulose synthase complexes (CSCs), likely in a heterotrimer configuration (Endler et al., 2016). For example, in primary walls CesA1, CesA3 and CesA6 isoforms constitute the catalytic core of the active CSC (Desprez et al., 2007), whereas in secondary walls, it is the CesA4, CesA7 and CesA8 isoforms (Persson et al., 2007).

Despite differences in the biomechanical properties in stems, no overall change in plant morphology was observed in fla11 fla12 mutants suggesting that redundancy with other FLA members might occur. Amongst all the FLAs, a member in group B, FLA16 (Fig. 1.6), is proposed to be a good candidate as a stem growth regulator given it is the most highly expressed FLA in the stem after FLA11 and FLA12 (MacMillan et al., 2010). 89

Based on the Arabidopsis eFP browser FLA16 is almost exclusively expressed in the stem (Schmid, 2005; Toufighi, 2005; Winter et al., 2007) (see Appendices Fig. A3.7). Unlike FLA11 and FLA12, FLA16 is predicted to contain two FAS domains and is not predicted to be GPI-anchored (Johnson et al., 2003). The biological roles of FLA16 in Arabidopsis are unknown. In this Chapter we aimed to determine if FLA16 plays a role in stem development using a fla16 mutant.

4.2 Results

4.2.1 Investigation of FLA16 transcript levels in Arabidopsis

To confirm the levels of FLA16 transcript abundance, Q-PCR was undertaken using a range of Arabidopsis organs. These included the flower, silique, cauline leaf, rosette leaf, branch and the top, middle and basal segments of the stem. Transcripts of FLA16 were detected in all tissues investigated (Fig. 4.1A). Transcripts were most abundant in the branch, followed by silique, flower, stem and root, and barely detectable in the cauline and rosette leaves (Fig. 4.1A).

A T-DNA insertion in the intron of the FLA16 gene was identified in the Arabidopsis SALK collection (SALK_131248) (Fig. 4.1B). The insertion in FLA16 was confirmed using genotyping PCR and Q-PCR of FLA16 transcripts in the stem tissue (Fig 4.1C). The expression levels of FLA16 in the fla16 mutant stem was about one-sixth of that of wt suggesting this was a severe knock-down mutant line (Fig 4.1C). Therefore, the fla16 mutant line was investigated further to determine if it showed phenotypic differences in growth and development compared to wt plants.

4.2.2 Growth is delayed in the fla16 mutant

The timing of specific growth stages was analysed in fla16 compared to wt using the method of Boyes et al. (2001). The emergence of even-numbered rosette leaves, root growth, the timing of bolting and appearance of the first flower were assessed (Table 4.1). Between seedling stages 0.7-1.2 (Boyes et al., 2001), a delay in root growth was observed in fla16 compared to wt. During early root growth, between day 2 to 5, a significantly shorter root length was observed in fla16 mutants (Table 4.1). The reduced 90 root length in fla16 was shown to result from the slower growth rate between days 1-3 with no significant difference in growth rate between days 3 to 4 or 4 to 5 (Table 4.1).

Analysis of the timing of rosette leaf appearance was carried out from the first true leaf until the first flower emerged. No significant differences in the timing of rosette leaf development was observed between fla16 and wt from the 2nd to the 8th leaf (see Appendices, Table A4.1, A4.2). The completion of rosette development was shown to occur earlier in fla16 compared to wt which had fewer rosette leaves at inflorescence emergence (Table 4.1). Although an earlier bolting (emergence of the inflorescence) time of 3 days occurred in fla16 compared to wt, no differences were seen in the timing of when the first flower was fully opened (Table 4.1).

4.2.3 Reduced stem length and transverse area was observed in fla16 mutants

Due to the expression of pFLA16:FLA16-VH in fibre cells in stems/branches (see Chapter 3, Figs. 3.8, 3.9), the stem length and area of fla16 mutants was of interest. Length of the total stem and first internode were measured at post-maturity (growth stage 6.9) of plants grown in long day conditions. For area measurements, the base and first node positions were chosen as two developmentally comparable regions (growth stage 6.5). Total area was determined from images of hand-sections of fresh stem tissue using image J software (see Chapter 2.21). The length of first internode versus total stem length and transverse area at the base versus first node area were used to assess if differences in the proportions of stem growth occurred.

The fla16 mutant showed significantly reduced total stem and first internode length (Fig 4.2, Table 4.2). The average total stem length of fla16 mutants was reduced by approximately 25%, and the length of the first internode of fla16 was approximately 46% of wt. The ratio of the first internode to total stem length showed that the first internode of fla16 was proportionally shorter than wt (Fig 4.2, Table 4.2).

In addition to changes in the stem length, fla16 mutants have thinner stems. The average stem area of fla16 mutants was reduced by approximately 14% and 31% at the base and first node, respectively, compared to wt. The ratio of the first node to basal area of fla16 (16%) was also found to significantly differ from wt (26%) (Table 4.2).

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4.2.4 Differences in cellular morphology are observed in fla16 stems compared to wt

Histological analyses of transverse and longitudinal sections of the stem (growth stage 6.5) were undertaken to determine if cellular differences in fla16 mutants contribute to the reduced stem length and area.

Regions of the base and first node of stems were transversely sectioned from either fresh or chemically fixed and resin embedded tissue of fla16 and wt. Toluidine blue stained sections revealed the fla16 pith area was smaller than wt with no other obvious differences in cellular organisation (Figs. 4.3, 4.5). Mäule staining for lignin did not show obvious differences in the distribution or type of lignin between fla16 and wt (Fig. 4.4).

To investigate the changes in stem area further, transverse sections of the stem were divided into several regions for area analysis including: 1) cortex, 2) pith and 3) intra-vascular region which consisted of interfascicular fibre, phloem, cambium and xylem cells (see Appendices Fig. A4.1). Cellular morphology was investigated with cell size analysis of the epidermis, cortex, interfascicular fibre, xylem vessel and pith cells using transverse sections. To assess if changes in cell number occurred, the total number of pith, interfascicular fibre and xylem vessel cells in transverse sections was undertaken.

Analyses of the tissue area revealed that the pith area was significantly reduced in the base (30%) and first node (38%) of fla16 compared to wt (Table 4.3). Measurement of the total number of pith cells in the base and first node revealed fla16 had significantly fewer pith cells than wt. In addition, the fla16 stem showed a significantly reduced intra-vascular region area (32%) at the first node but not the base, which likely resulted from a reduced number of interfascicular fibre cells. No significant changes were observed in the cortex region at either stem position (Table 4.3).

Analysis of cell size in the stem transverse sections showed fla16 had significantly smaller pith cells in the first node but not the base. Other cell types examined, including epidermis, xylem vessel and interfascicular fibre cells, did not show statistically significant differences in size at both stem positions, suggesting that the reduction in

92 pith cell size and number in fla16 compared to wt is the major cause of the reduced stem area in fla16 (Table 4.3).

Longitudinal sections at the base and first node of the stem were examined to determine if differences in cell length could be observed in fla16 compared to wt. No significant differences in the length of epidermal and pith cells was observed (Fig. 4.6A, Table 4.3).

During handling of stem tissue it was observed that branches detached more easily from the main stem in fla16 mutants and the mutant stems appear to be more brittle compared to wt. Investigation of the angle of the first branch in relation to the stem showed no differences between fla16 and wt (Figs. 4.6B, C; Table 4.2). Changes in the composition or properties of the cell walls in stems could explain differences in stem detachment and brittleness. To determine if cell wall compositional changes occur in the fla16 mutant stems, analyses of carbohydrates were undertaken.

4.2.5 Carbohydrate content is altered in the fla16 stems

The polysaccharide composition was determined by linkage analyses of the alcohol insoluble residue (AIR) representing the cell wall fractions extracted from fla16 and wt stems (growth stage 6.9) (Pettolino et al., 2012). Linkage analyses of the cell wall polysaccharides revealed an approx. 9% reduction of cellulose and 10% increase in glucuronoxylan in fla16 stems (Fig. 4.7A). Using an acetic/nitric digestion assay to determine the amount of crystalline cellulose (Updegraff, 1969) an approx. 16% reduction was observed in the mutant line (Fig. 4.7B, Table 4.2). Thus cellulose, which is the major component of the walls (approx. 50%) is significantly reduced and as it is known to provide rigidity and mechanical strength (Cosgrove, 2005) this led us to investigate whether this reduction affects the biomechanical properties of the mutant stems.

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4.2.6 fla16 stems display altered biomechanical properties compared to wt

Biomechanical tests were carried out for the basal and middle stem segments, whereas the top of the stem was excluded due to the reduced stem length. Differences in the hydration status of stems can influence the biomechanical properties leading to variable results, therefore dry stems were chosen for testing (Molina-Freaner et al., 1998). Flexural and tensile strength analyses for both the strength and stiffness of the stem tissues were undertaken. For flexural analysis, three-point bending tests were undertaken whereby the bending stress at the midpoint of the stem segment when the load reaches its maximum is recorded. For tensile analysis, the ends of the stem segment were fixed and a pulling force applied until the stem broke (MacMillan et al., 2010). The strength indicates the maximum load required to break the stem, whereas stiffness is a measure of the elasticity of the stem (MacMillan et al., 2010). In addition, a ‘break’ profile was included to evaluate the response of stems under load. This allowed evaluation of which stems underwent a sharp break and those with a gradual break. Since the stems were not of uniform shape, three flexure analytical models were compared covering the major potential geometrical shapes of the stem: circular, oval and rectangle (see Appendices Fig. A4.2). The circular model was chosen as the best model for tensile analyses as it catered for the broad range of stem shapes (Dr Colleen MacMillan, personal communication).

Flexure tests revealed that the basal stems of fla16 have significantly reduced flexural strength (34.3%) compared to wt (Table 4.4). None of the fla16 samples yielded a sharp break profile at the base compared to 16.7% of wt stems having a sharp break profile in the basal stems. No significant differences in flexure tests of the middle stems were found between fla16 and wt (Table 4.4 and see Appendices Fig. A4.2).

Tensile strength tests of basal and middle stems identified a significantly increased tensile stiffness in the middle stems, but not the basal-stems of fla16 compared to wt (Table 4.4 and see Appendices Fig. A4.3). No other significant differences in tensile stiffness or strength were observed between fla16 and wt stems.

In summary, the fla16 mutant exhibited a number of phenotypes during growth and development, in particular, reduced stem length and cellulose content and

94 biomechanical properties. To verify that these phenotypes resulted specifically from the loss of FLA16, complementation of the fla16 mutant line was undertaken.

4.2.7 Complementation of the fla16 mutant The endogenous FLA16 promoter (pFLA16) was used to drive two constructs that were transformed into the fla16 mutant to determine if complementation of the mutant phenotype occurred. The first construct was made using the FLA16 coding region (pFLA16:FLA16) and the second construct included reporter proteins with Venus (V) and HIS (H) tag fusions at the C-terminus (pFLA16:FLA16-VH) (see Chapter 3, Fig. 3.2). Homozygous plant lines for the transgenes in the fla16 mutant background were generated and examined for FLA16 expression by Q-PCR and complementation of the reduced cellulose content and stem length phenotypes. The expression of FLA16 transcripts in the total stem of fla16 mutants is reduced to 21% of wt levels (Fig 4.8). In fla16 pFLA16:FLA16 and fla16 pFLA16:FLA16-VH lines the transcript levels in stems were 73% and 123% of wt levels, respectively, based on three biological replicates, indicating that FLA16 expression was likely to be partially recovered in fla16 pFLA16:FLA16 lines and increased in fla16 pFLA16:FLA16-VH lines. Based on acetic/nitric cellulose assays, the stem cellulose content in fla16 pFLA16:FLA16 and fla16 pFLA16:FLA16-VH stems was 98% and 93%, respectively, of wt levels. This is a significant increase in crystalline cellulose from the fla16 line (83% of wt). Both complementation lines partially recovered the stem length phenotype compared to wt, whereas the length of the first internode was not recovered and was comparable to fla16 (Fig. 4.8). While these complementation studies show partial recovery of the mutant phenotypes, these data were gathered from only two individual transformed lines for each construct, thus screening of more individual transformants is needed to confirm the phenotypes observed in fla16 mutants are indeed caused by loss of FLA16.

4.2.8 Expression of FLA16 in the stem overlaps with the expression of the primary and secondary wall cellulose synthases (CesAs)

fla16 was shown to impact the levels of crystalline cellulose in the stem. This could either be due to reduced expression of CesA transcripts or reduced activity of cellulose synthases (CesAs), trafficking disruption of cellulose synthase complexes (CSC),

95 or altered cellulose crystallinity in the wall. To determine if the expression of CesAs involved in primary and secondary wall cellulose synthesis was altered in fla16 mutants, Q-PCR analysis was used to investigate the expression of the primary wall CesAs (1, 3 and 6) and secondary wall CesAs (4, 7 and 8) (Persson et al., 2007) at the base-, middle- and top-stem portions of fla16 and wt (growth stage 6.5).

No clear pattern of altered expression for primary wall CesAs (1, 3, 6) or the secondary wall CesAs (4 and 8) was observed in any stem segments of fla16 compared to wt. In contrast, a consistent increase in the expression of CesA7 was observed in all segments of the fla16 stems compared to wt (Fig. 4.9), suggesting FLA16 may be involved in the regulation of CesA7. The relationship between FLA16 and cellulose synthesis could occur during the transition from a primary wall to secondary wall. To determine if FLA16 is implicated in cellulose deficiency in primary walls, hypocotyls were examined in seedlings grown under conditions known to perturb cellulose synthesis.

4.2.9 The fla16 mutant is more sensitive to isoxaben treatment than wt

Transcripts of FLA16 were expressed in the hypocotyl during early development (Schmid, 2005; Toufighi, 2005; Winter et al., 2007). Compared to wt, mutants with reduced cellulose levels frequently display higher sensitivity to the herbicide isoxaben, displaying reduced hypocotyl growth and swelling (Desprez et al., 2002). Isoxaben has been shown to inhibit cellulose synthesis in plant primary walls (Desprez et al., 2002). fla16 mutants and wt were tested for sensitivity to isoxaben during dark-grown seedling development. Hypocotyl growth of fla16 mutants was found to be more sensitive to isoxaben treatment, with significantly reduced hypocotyl length and increased swelling compared to wt, indicating that cellulose synthesis in primary walls is compromised in fla16 (Fig. 4.10, Table 4.1).

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4.3 Discussion In the current study, we identified and characterised a fla16 mutant. Loss of FLA16 leads to a number of alterations in plant growth, including root growth, flowering time, reduced stem length and changes in stem cell morphology and biomechanics. These data provide insight into the potential role(s) of FLA16 during stem growth and its possible roles in wall assembly, organisation and differentiation.

Expression of FLA16 was shown to be largely specific for cells with secondary walls in stems however no obvious phenotypic changes were observed in these cell types. During stem development, growth must be carefully coordinated with maintaining structural integrity. Therefore secondary wall deposition is balanced with stem growth and likely involves feedback mechanisms to ensure tissue integrity. Although no morphological changes in fibre or xylem cells were observed in the fla16 stems compared to wt, changes in fla16 wall integrity likely occurs based on the reduction in cellulose and altered biomechanics. Changes in wall mechanics are thought to be interpreted by cell wall integrity sensors that initiate signalling pathways to regulate growth (Lodish et al., 2000). The reduction of fla16 pith area could be a result of such compensation to reduce tension in the outer layers by restricting cell division and expansion (for further discussion see Chapter 5.3.1). The current study focussed on the cellular morphology of stems at mature growth stages. Future studies could include examination of stems throughout development to obtain systematic and comprehensive data for the dynamics of stem development.

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4.3.1 FLA16 potentially influences stem biomechanics by regulating cellulose microfibrils The expression of FLA16 in xylem and fibre cells of the stem and its likely location in the wall suggests that the reduced stem length and altered biomechanics observed in fla16 are related to altered architecture and/or composition of secondary walls in stems. Changes in the fla16 stem biomechanical properties are seen in both the strength and stiffness. The reduction of flexural strength in the fla16 basal stems is consistent with the reduced cellulose content. Cellulose is a major factor influencing stem mechanical properties (Turner and Somerville, 1997; Li et al., 2003) and has been demonstrated to impact flexural strength (Wang et al., 2012). Another important wall component influencing stem biomechanics is lignin, a phenolic polymer of lignified secondary walls, particularly in the vasculature (Boudet et al., 1995; Whetten and Sederoff, 1995). Lignin has been shown to provide flexural strength and stiffness to the stem (MacKay et al., 1997). A minor increase in the lignin content was observed in the fla11 single mutant (3%) and an even smaller change in the fla11 fla12 double mutants (1.7%) by MacMillan et al. (2010) but its relevance given the dramatic changes in microfibrillar cellulose remain unclear. As a result, further analysis of lignin composition, for example, by higher-resolution techniques such as nuclear magnetic resonance (NMR) (Ghaffar and Fan, 2013; Wen et al., 2013) should be conducted, and perhaps more importantly, studies of lignin mutants in a fla16 background would more directly address if lignin contributes to these biomechanical properties in a significant manner. In addition to the reduction in strength, we also detected an increased tensile stiffness in the fla16 middle stem. Tensile stiffness in stems is influenced by several factors, including the composition of the wall matrix (i.e. lignin and hemicelluloses) (Kohler and Spatz, 2002) and wall architecture, in particular the cellulose microfibril angle (MFA) (Cave and Walker, 1994; Keckes et al., 2003). An increase in tensile stiffness in the fla16 middle stem segment could result from changes in these components. Although not statistically significant, increased tensile stiffness was also observed in the base of the stem. Given 12 samples were used for the middle and only 6 samples for the basal stem, with increased sampling it is possible that the basal tensile stiffness will also be significant. A small increase in the hemicellulose glucuronoxylan was observed from linkage analysis however it is unclear if this would be sufficient to influence biomechanical properties. To assess this, labelling of xylan epitopes using antibodies 98 directed against these polysaccharides in fla16 and wt stems could reveal if changes occur in specific cell types (Schädel et al., 2010) as well as investigating the effect of mutants with reduced glucuronxylan in the fla16 background. A likely factor influencing tensile stiffness in fla16 stems is MFA, that is, the angle between the cellulose fibrils and the longitudinal cell axis. MFA is a crucial factor determining wood properties that has shown to be regulated by FLAs (MacMillan et al., 2010). For example, in fla11 fla12 mutant stems MFA was significantly increased (MacMillan et al., 2010). A large MFA is thought to confer flexibility to younger regions of the stem and a transition to a smaller MFA occurs in older regions of the stem to confer stiffness and the ability to withstand stresses such as the weight of the organ and wind (Barnett and Bonham, 2004). During tensile tests, or the force required to break or deform the sample, a low MFA results in higher stiffness and strength (MacMillan et al., 2010). In future, analysis of MFA in fla16 mutant stems using X-ray diffraction (Sarén and Serimaa, 2005) and scattering (Saxe et al., 2014) is required to determine if changes occur and if so, how this relates to the altered stem biomechanics in fla16. Differences in the architecture of fla16 stems can also be inferred by differences in the break profile of basal stems during flexure tests (Table 4.4). The break profile might reflect the biomechanical properties of the stem (Read and Stokes, 2006), which are influenced by the uniformity of the stem material, cell organisation (Hesse et al., 2016) and cell wall integrity (MacMillan et al., 2010). The amount, degree of crystallinity and degree of polymerisation of cellulose influences stem strength and stiffness and varies depending on the species and wall types being investigated (Nookaraju et al., 2013). In this study, reduced cellulose levels are observed, however further investigation is needed to determine if changes to the cellulose itself also occurs in the mutant stems. Within cellulose microfibrils, a highly crystalline core is surrounded by amorphous forms (Fan et al., 1980). The crystallinity of cellulose can be modified by a number of factors, including proteins that associate with the CSCs and cellulose-matrix interactions. For example, mutants lacking the GPI-anchored COBRA-like proteins, that directly bind cellulose and regulate microfibril crystallinity, in Arabidopsis (Roudier et al., 2002), rice (Liu et al., 2013) and maize (Sindhu et al., 2007) have reduced mechanical strength (Ries and Pruitt, 2005). To determine if cellulose crystallinity is altered in fla16, X-ray diffraction could be undertaken to provide the crystallinity index (Liu et al., 2016) and ratio of amorphous to crystalline cellulose (Li et al., 2014). 99

While it is hard to elucidate the exact mechanism of how FLAs influence cellulose deposition, this could be either via direct contact with the microfibrils or through interaction with the CSC regulatory complex. For example, FLAs could potentially associate with proteins guiding the movement, orientation or activity of CSCs and hence influence the deposition of cellulose (Schneider et al., 2016). Proteins known to regulate cellulose synthesis include the endoglucanase KORIGAN (Vain et al., 2014), chitinase-like (CTL) proteins (CTLs1/2) (Sanchez-Rodriguez et al., 2012) and companions of CesAs (CCs) (Endler et al., 2016). Currently there is no direct evidence showing FLA16 interacts with CSCs, however, CSCs and FLAs could potentially be delivered to the same region of the PM (see Chapter 3.2.2) and/or undergo protein-protein interactions (see Chapter 1.4.1). The protein-protein interaction potential of the FAS domain has been clearly demonstrated in animal and insect cells (Bastiani et al., 1987; Kolodkin et al., 1992; Litvin et al., 2004; Kannabiran and Klintworth, 2006; Liu et al., 2009) and this needs to be investigated further for plant FAS domains. For instance, Co-IP might be a useful biochemical approach (Speth et al., 2014; Kudla and Bock, 2016; Lin and Lai, 2017) to find candidate FLA interacting proteins. Co-location of FLAs with CSCs would also provide further evidence (Liu et al., 2014; Endler et al., 2016; Schneider et al., 2016), although this is challenging for internal cells such as xylem and fibre cells. A GR-VND7 inducible system that initiates secondary walls to develop in any cell type has proven a highly effective tool to visualize CesA7 during secondary wall initiation (Watanabe et al., 2015; Schneider et al., 2017). This system could also be employed to visualise FLA16 and its potential interacting partners as well as to investigate how loss of FLA16 leads to altered expression of CesA7 (Figure 4.9). To examine if changes in CesA7 stability or trafficking are altered, the GR-VND7 inducible YFP-CesA7 system in a fla16 mutant background could be employed. Integration of the pFLA16:FLA16-HV into the VND7 system would also enable visualisation of when and where FLA16 is located during the transition to secondary walls. The expression of genes for secondary wall biosynthesis is regulated by a complex TF network (see Chapter 1.1.2) and the TF MYB46 is known to regulate expression of CesA4, 7 and 8 that are active in secondary wall biosynthesis (Kim et al., 2013). A study of the specificity and organization of the secondary wall CesAs within CSCs suggests that CesA7 has higher class specificity than CesA4 and CesA8 and therefore a more constrained position within the CSC (Kumar et al., 2017). Loss of CesA7 100 results in an irregular xylem (irx3) mutant phenotype with reduced cellulose leading to collapsed xylem cells (Taylor et al., 1999). How loss of FLA16 leads to increased CesA7 transcript levels is unclear (Figure 4.9). Since direct transcriptional regulation of CesA7 occurs intracellularly (Kumar et al., 2017) whereas FLA16 is located in the PM/extracellular matrix, this is likely to be indirect. FLAs have been proposed to be wall sensing molecules (see Chapter 3.3.1) as exemplified by the case of FLA4 cooperating with FEI1/FEI2 in the regulation of cell wall biosynthesis (Xu et al., 2008). It is possible the transcriptional regulation of CesA7 results from feedback from the wall due to the reduced cellulose content. For example, altered mechanical properties in walls of fla16 could generate signals that are transduced intracellularly leading to activation of CesA7 expression. Why CesA7 is specifically affected and not the other CesAs is unclear. CesA7 has been shown to be post-translationally regulated by phosphorylation which targets the CesA7 protein for degradation via the proteasome, hence regulating CesA7 levels and modulating activity of CSCs (Taylor, 2007). To determine if a discrepancy between CesA transcript levels and protein occurs, quantitative analysis of CesA levels using isobaric peptide tags for relative and absolute quantification (iTRAQ) by mass spectrometry (Tweedie-Cullen and Livingstone-Zatchej, 2008) could be used. In addition, quantification of CesA7 phosphorylation by phosphor-proteomics (Previs et al., 2008) could yield information regarding the post-translational status of CesA7. FLAs could directly interact with microfibrils in a mechanism similar to the GPI- anchored COBRA which is proposed to act as a ‘polysaccharide chaperone’ to regulate cellulose crystallization in the wall (Li et al., 2014; Sorek and Somerville, 2016). This is supported by the proposed role of FLA4 to align cellulose microfibrils (Harpaz-Saad et al., 2011) and interact with pectins in seed mucilage (Griffiths et al., 2016). For FLAs it is possible to envisage that either the FAS domain and/or AG moieties could interact with microfibrils (see also proposed models of interaction discussed in Chapter 6). The interaction between FLAs and cellulose might occur given FAS domains in animals have been shown to interact with extracellular matrix components (Kim et al., 2002). Higher resolution methods, such as atomic force microscopy are now capable of detecting interactions between single molecules and could be employed to study these processes (Willemsen et al., 2000). The AG glycans of some AGPs have the ability to covalently crosslink wall components, such as hemicelluloses and pectins (Tan et al., 2013). Such carbohydrate-carbohydrate interactions might play a crucial role in the spatial 101 arrangement of the wall matrix (Gorshkova et al., 2013). Furthermore, the association between cellulose and these wall components could potentially influence matrix viscosity and impact MFA. It is currently unclear which domains of FLA16 are required for its function in maintenance of cellulose and biomechanics. FLA16 consists of N- and C-terminal FAS domains (Johnson et al., 2003) and is likely to be glycosylated (see Chapter 3). The FAS domains of FLA4 have been shown to have distinct properties, the N-FAS plays a role in directing subcellular localisation whereas the C-FAS in required for protein function in roots (Xue et al., 2017). In addition, glycosylation can play a number of roles, such as stabilising protein tertiary confirmation and assisting molecular interactions (Chen et al., 2010). To verify which functional domains of FLA16 are required for function requires further investigation such as domain deletion/swap studies (Sugimoto et al., 2003; Xue et al., 2017) as discussed in Chapter 3.3.2. To summarise, FLA16 appears to play a role during stem development, similar to that described for FLA11 and FLA12. To investigate if these FLAs could play roles in overlapping pathways genetic interaction studies are undertaken in Chapter 5.

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Table 4.1 Analysis of selected growth stages of long day grown Arabidopsis wt and fla16 plantsa. wt fla16 n Root length (mm) Day 2 0.15 (± 0.02)b 0.07 (± 0.01)c 10 Day 3 0.45 (± 0.03) 0.28 (± 0.04) 10 Day 4 0.68 (± 0.04) 0.51 (± 0.05) 10 Day 5 1.05 (± 0.06) 0.84 (± 0.04) 10 Day 6 1.47 (± 0.10) 1.25 (± 0.06) 10 Day 7 1.89 (± 0.13) 1.70 (± 0.07) 10 Day 8 2.36 (± 0.16) 2.10 (± 0.08) 10 Root growth rate (mm/day) Day 1-2 0.15 (± 0.02) 0.07 (± 0.01) 10 Day 2-3 0.30 (± 0.03) 0.21 (± 0.03) 10 Day 3-4 0.23 (± 0.02) 0.23 (± 0.01) 10 Day 4-5 0.37 (± 0.02) 0.34 (± 0.01) 10 Day 5-6 0.42 (± 0.05) 0.41 (± 0.02) 10 Day 6-7 0.42 (± 0.04) 0.45 (± 0.03) 10 Day 7-8 0.47 (± 0.12) 0.40 (± 0.03) 10 Hypocotyl length (cm)d ½ MS 0.92 (± 0.04) 0.79 (± 0.03) 14-17 ½ MS + 2nM isoxaben 0.36 (± 0.001) 0.22 (± 0.009) 17-20 Rosette number at rosette growth completion 13.78 (± 0.17) 12.06 (± 0.32) 48-50 Days to first flower bud visible 26.16 (± 0.55) 23.73 (± 0.34) 48-50 Days to first flower fully open 30.48 (± 0.51) 30.08 (± 0.34) 48-50 aGrowth stages as outlined in Boyes et al. (2001), root length at day 2-8 corresponds to growth stage 0.7- 1.02, rosette number at rosette growth completion, first flower bud visible and first flower fully open correspond to stage 3.9, 5.1 and 6.0. bBracket indicates ± SE. cBold text indicates a statistically significant value at p<0.05 using student T-test. dSeedlings grown in the dark and hypocotyls measured at 4th day post-stratification.

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Table 4.2 Measurement of stem morphology, biomechanical properties and cellulose content of long day grown wt and fla16 stems at maturity a,b. wt fla16 nc Stem length (cm) Total stem 46.44 (± 0.66)d 35.02b (± 0.38) 68 First internode 11.79 (± 0.34) 5.46 (± 0.26) 68 Internode to total stem ratio 25.55 (±0.73) 15.73 (± 0.78) 68 Branch to stem angle (o) 52 (± 3.84) 57.75 (± 2.09) 4-5 Stem transverse area (mm2) Basal stem 0.87 (± 0.04) 0.75 (± 0.03) 33 First node stem 0.97 (± 0.05) 0.67 (± 0.04) 33 First node to basal stem ratio 1.19 (± 7.09) 0.97 (± 4.39) 33 Cellulose content (%)e 50.54 (± 1.38) 42.30 (± 0.18) 3 aGrowth stage 6.5 as outlined in Boyes et al. (2001). bBold text in numeral data column indicates data value statistically significant at p<0.05 using Student T test. cn presents number of biological replicates. dBracket indicates ± SE. eCrystalline cellulose content determined by the acetic/nitric digestion assay (Updegraff, 1969).

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Table 4.3 Analysis of tissue and cellular morphology in mature stems of Arabidopsis wt and fla16 plants growna in long day conditions. wt fla16 nb Base Tissue area (mm2)c Cortex 0.18 (± 0.01)d 0.17 (± 0.01) 6 Intra-vascular region 0.33 (± 0.01) 0.32 (± 0.02) 6 Pith 0.37 (± 0.03) 0.26e (± 0.02) 6 Cell area (µm2) Epidermis 238.94 (± 20.71) 226.41 (± 19.54) 5-6 Interfascicular fibre 128.76 (± 1.62) 130.65 (± 7.84) 5-6 Cortex 150.98 (± 18.77) 204.02 (± 10.74) 5-6 Xylem vessel 184.70 (± 7.49) 210.12 (± 10.53) 5-6 Pith 817.14 (± 36.15) 742.14 (± 23.42) 5-6 Cell number Pith 492.33 (± 38.19) 360.40 (± 22.80) 5-6 Interfascicular fibre 1149.75 (± 39.86) 1130.25 (± 87.10) 4 Xylem vessel 922.75 (± 49.57) 884.75 (± 54.15) 4 Cell length (µm)f Epidermis 89.47 (± 12.02) 96.54 (± 0.95) 4-5 Pith 75.42 (± 8.20) 74.40 (± 3.06) 4-5 First node Tissue area (mm2) Cortex 0.169 (± 0.01) 0.15 (± 0.01) 4-6 Intra-vascular region 0.31 (± 0.02) 0.21 (± 0.01) 4-6 Pith area 0.5 (± 0.05) 0.31 (± 0.04) 4-6 Cell area (µm2) Epidermal cell (µm2) 151.71 (± 10.41) 154.97 (± 5.00) 4-5 Interfascicular fibre cell (µm2) 142.30 (± 6.24) 140.30 (± 6.77) 4-5 Cortex cell (µm2) 138.68 (± 20.07) 124.02 (± 4.27) 4-5 Xylem vessel cell (µm2) 171.98 (± 5.79) 198.79 (± 7.91) 4-5 Pith cell (µm2) 1140.12 (± 50.49) 902.80 (± 47.11) 4-5 Cell number Pith 462.33 (± 10.30) 365.80 (± 13.31) 4-5 Interfascicular fibre 811.75 (± 47.57) 635.00 (±50.86) 4 Xylem vessel 646.75 (± 59.09) 618.25 (± 25.39) 4 Cell length (µm) Epidermis 75.52 (± 5.56) 99.25 (± 10.67) 4-5 Pith 83.43 (± 7.90) 85.44 (± 10.41) 4-5 aGrowth stage 6.5 as outlined in Boyes et al. (2001). bn presents number of biological replicates. cTissue divisions refer to Appendices Fig. A4.1. dBracket indicates ± SE. eBold text in numeral data column indicates data value statistically significant at p<0.05 using Student T test. fCell length measurements taken from longitudinal stem sections.

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Table 4.4 Biomechanical properties of wt and fla16 stems at senescence completiona,b. Stem biomechanics (N/mm2)c wt fla16 nd Flexure Basal stem flexural strength 122.26 (± 13.23)e 80.32 (± 3.52) 6-7 Basal stem flexural stiffness 7789.83 (± 755.89) 7992.82 (± 470.94) 6-7 Basal stem flexural breaking (%) 16.7 0 6-7 Mid stem flexural strength 97.59 (± 12.38) 97.44 (± 6.38) 12 Mid stem flexural stiffness 14544.73 (± 1777.28) 13462.14 (± 926.87) 12 Mid stem flexural breaking (%) 0 0 6 Tensile Basal stem tensile strength 83.13 (± 10.86) 92.54 (± 13.90) 6 Basal stem tensile stiffness 3822.34 (± 484.33) 4799.72 (± 529.22) 6 Basal stem tensile breaking (%) 83.33 83.33 Mid stem tensile strength 74.55 (± 6.19) 68.43 (± 5.37) 12 Mid stem tensile breaking (%) 100 100 12 Mid stem tensile stiffness 4098.26 (± 172.54) 4952.03 (± 170.86) 12 aGrowth stage 9.7 as outlined in Boyes et al. (2001). bBold text in numeral data column indicates data value statistically significant at p<0.05 using Student T test. cN/mm2 represents Newton/squared millimetre. dn presents number of biological replicates. eBracket indicates ± SE.

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Figure 4.1 Expression levels of FLA16 transcripts in tissues of Arabidopsis wt and a fla16 T-DNA insertion line. (A) Q-PCR analysis of FLA16 transcript levels in different Arabidopsis tissues at growth stage 6.5 (Boyes et al., 2001). N = 2, data are represented as mean ± SE (B) Schematic representation of the FLA16 gene showing regions predicted to encode the signal peptide (Sp), fasciclin (FAS) domain and arabinogalactan-protein (AGP) domains. The position of an intron (black), the T-DNA insertion in the fla16 mutant, and primers used for Q-PCR analysis (qF1/R1, qF2/R2) are indicated. (C) FLA16 transcript levels are significantly reduced in stems of a fla16 mutant compared to wt using the qF2/R2 primers. Transcript levels were determined relative to DNA standards of known concentration and normalized with GAPDH, tubulin and cyclophilin housekeeping genes (Czechowski et al., 2005). N = 3, p<0.05 (*) using Student T-test, data are represented as mean ± SE.

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Figure 4.2 The fla16 mutant shows a reduced stem length phenotype at maturity. Representative image of wt and fla16 mutants grown to post-maturity (growth stage 6.9 (Boyes et al., 2001)) shows the reduced total stem and first internode lengths of fla16 compared to wt. Scale bar=2 cm.

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Figure 4.3 Cellular morphology of Arabidopsis fla16 and wt in transverse sections of fresh mature stems. Hand sections of fresh stems at growth stage 6.5 (Boyes et al., 2001) were taken at the base and first node of stems and stained with toluidine blue. A reduction in the total stem area occurs in fla16 (B) compared to wt (A) with a reduction in pith area. No obvious differences in cellular morphology are observed at higher magnification (a, b, boxed areas from A and B, respectively). Transverse sections taken at the first node of stem shows the reduction in stem and pith area in fla16 (D) compared to wt (C). Higher magnification images (c, d, boxed areas from C and D, respectively) of cellular morphology at the first node show obvious differences in area. Pith (Pt), cortex (cr), starch sheath (SS), epidermis (Ep), cambium (Cm) and phloem (Ph). Scale bar=200 µm.

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Figure 4.4 Lignin distribution in transverse sections of fresh mature stems of fla16 and wt. Hand sections of fresh stems at growth stage 6.5 (Boyes et al., 2001) were taken at the base and first node, and stained with the Maule reagent (Pradhan Mitra and Loqué, 2014). No notable difference in lignin distribution between fla16 and wt stem sections were evident at either the base or first node. (A) Basal stem section of fla16. (B) Basal stem section of wt. (C) First node stem section of fla16. (D) First node stem section of wt. (a, b, c, d) High magnification insets of boxed areas in A, B, C, D, respectively, showing pith (Pt), cortex (Cr), starch sheath (SS), epidermis (Ep), cambium (Cm) and phloem (Ph). Scale bar=200 µm.

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Figure 4.5 Cellular morphology derived from transverse sections of mature stems of Arabidopsis fla16 and wt. Toluidine blue staining shows the reduction of pith area in fla16 (B) compared to wt (A) at the base and first node (C, D). (a, b, c, d) are high magnification insets of boxed areas in A, B, C, D, respectively. At the base (a, b) and first node (c, d) no obvious differences in cellular morphology are observed between wt (a, c) and fla16 (b,d). Pith (Pt), cortex (cr), starch sheath (SS), epidermis (Ep), cambium (Cm) and phloem (Ph). Scale bar = 500 µm in A, B, C and D and 50 µm in a, b, c and d.

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Figure 4.6 Measurement of branch angle relative to the main stem at the first node of mature Arabidopsis wt and fla16 plants. Representative image of a longitudinal stem section showing measurement of the angle of the first branch relative to the main stem. Analysis of the branch angle in fla16 compared to wt shows no significant differences. Data are presented as means ±SE, N = 4-6, p>0.05 using Student T-test. Scale bar = 500 µm.

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Figure 4.7 Cell wall polysaccharide composition derived from linkage analysis of mature stems of fla16 and wt. (A) Polysaccharide linkage analysis (see Table A4.3) shows a reduction of cellulose and an increase in glucuronoxylan in fla16 compared to wt with no difference in the other polysaccharides. (B) Acetic/nitric cellulose assay (Updegraff, 1969) shows a 16% reduction of crystalline cellulose in fla16 compared to wt. Data are presented as means ±SE.

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Figure 4.8 Complementation of the fla16 mutant with pFLA16:FLA16 and FLA16:FLA16-VH. (A) Q-PCR analysis of FLA16 transcript abundance shows fla16 with 21%, complemented fla16 FLA16:FLA16 lines with 73.3% and fla16 pFLA16:FLA16-VH with 123.8% of wt FLA16 levels. (B) Crystalline cellulose levels are largely recovered to wt levels in both fla16 pFLA16:FLA16 (98.2%) and fla16 pFLA16:FLA16-VH (93.6%) complementation lines. (C) Total stem length was partially recovered in both fla16 pFLA16:FLA16 (81.7%) and fla16 pFLA16:FLA16-VH (89.5%) complementation lines with no recovery of first internode length (N = 25-40, p<0.05 (*) using Student T-test) compared to wt. Data are presented as means ±SE.

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118

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Figure 4.9 Q-PCR analysis of CesA transcript abundance in top-, mid- and basal- segments of mature Arabidopsis wt and fla16 stems. Transcript abundance of CesAs involved in the synthesis of primary (CesA1, 3 and 6) and secondary (CesA4, 7 and 8) wall cellulose (Persson et al., 2007) in the top (A), middle (B) and basal (C) stem segments show increased levels of only CesA7 transcripts in all stem segments. Transcript levels were determined relative to DNA standards of known concentration and normalized with GAPDH, tubulin and cyclophilin housekeeping genes (Burton et al., 2004). Data are represented as mean ± SE.

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Figure 4.10 Isoxaben treatment of dark grown Arabidopsis fla16 and wt seedlings. A significant reduction in hypocotyl length (A; Table 4.1) and increased swelling (B) occurs in the fla16 seedlings compared to wt in the presence of 2nM isoxaben (see Table 4.1). Scale bar = 1cm.

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APPENDICES TO CHAPTER 4

Table A4.1 Vegetative organ growth rate of fla16 and wt. wt fla16 n Growth stagea Days 2 cotyledon 1.0 8.02 (±0.02)b 8.12 (±0.07) 48-50 2 rosette leaf 1.02 11.98 (±0.08) 12.1 (±0.11) 48-50 4 rosette leaf 1.04 14.7 (±0.13) 15.1 (±0.13) 48-50 6 rosette leaf 1.06 17.98 (±0.13) 17.84 (±0.14) 48-50 8 rosette leaf 1.08 19.1 (±0.20) 20.33 (±0.12) 48-50 10 rosette leaf 1.10 21.71 (±0.16) 22.53 (±0.32) 48-50 12 rosette leaf 1.12 23.93 (±0.21) 2 (±0.49)c 48-50 aVegetative organ development of of fla16 relative to wt according to Boyes et al. (2001). bData are presented as mean (±SE). cStudent T-test, p<0.05 as indicated in bolded data.

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A4.2 Polysaccharide composition from stems of fla16 and wt deduced from linkage analysis (see Fig. 4.7).

wt fla16

1 2 3 Mean (± SE) 1 2 Mean (± SE)

Arabinan 1,5-Ara (f) 1.1 1.8 1.0 0.8 0.7 Total Arabinan 1.1 1.8 1.0 1.3 (± 0.3) 0.8 0.7 0.8 (± 0.1)

Type I AG 1,4-Gal (p) 3.2 2.5 3.4 4.0 3.1

1,4,6-Gal (p) 1.3 1.2 1.5 0.9 1.2

t-Ara 1.1 1.2 1.1 0.9 0.8 Total Type I AG 5.6 4.9 6.0 5.5 (± 0.3) 5.9 5.0 5.5 (± 0.5)

Type II AG 1,3-Gal (p) 0.0 0.0 0.0 0.0 0.0

1,6-Gal (p) 0.1 0.0 0.0 0.0 0.1

1,3,6-Gal (p) 0.7 0.5 1.7 2.0 1.2

1,3,4,6-Gal (p) 0.0 0.0 0.0 0.0 0.1

t-Ara 0.0 0.5 0.0 2.0 0.0

t-Gal 0.6 0.0 0.2 0.0 0.6 Total Type II AG 1.4 1.0 1.9 1.5 (± 0.3) 4.0 2.0 3.1 (± 1.1)

Homogalacturonan 1,4-GalA 4.4 3.6 3.4 2.7 4.2 Total Homogalacturonan 4.4 3.6 3.4 3.8 (± 0.3) 2.7 4.2 3.5 (± 0.7)

RG I/II

1,4-GalA 0.4 0.2 0.0 0.0 0.4

1,2,4-Rha (p) 0.4 0.2 0.0 0.0 0.4 Total RG I/II 0.8 0.3 0.0 0.4 (± 0.2) 0.0 0.8 0.4 (± 0.4)

Glucuronoxylan 1,4-Xyl (p) 25.9 25.2 25.3 27.0 27.9

1,2,4-Xyl (p) 2.5 2.5 1.5 2.4 2.9

t-GlcA 2.5 2.5 1.5 2.4 2.9 Total Glucuronoxylan 30.8 30.3 28.3 29.8 (± 0.8) 31.9 33.7 32.8 (± 0.9)

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Table A4.2 continued.

Xyloglucan 1,4,6-Glc (p) tra tr tr tr tr

1,4-Glc (p) tr tr tr tr tr

1,2-Xyl (p) tr tr tr tr tr

t-Xyl tr tr tr tr tr Total Xyloglucan 0.1 0.0 0.0 0.00 (± 0.00) 0.0 0.1 0.1 (± 0.1) Cellulose 1,4-Glc 51.4 51.8 52.5 46.3 48.2 Total Cellulose 51.4 51.8 52.5 51.9 (± 0.3) 46.3 48.2 47.3 (± 1.0) Total 95.7 93.8 93.1 94.2 (± 0.78) 91.7 94.7 93.2 (± 1.5) Unassigned linkage t-Ara (f) 0.0 0.0 0.0 0.4 0.0 t-Xyl (p) 1.0 2.0 0.9 4.2 1.0

1,2-Xyl (p) 1.3 2.0 2.0 1.4 2.2

t-Glc (p) 0.1 0.0 0.0 0.0 0.2

1,3,4-Glc (p) 0.3 0.7 1.1 0.4 0.6

1,2,4-Glc (p) 0.4 0.4 0.5 0.0 0.4

1,3,4,6-Glc (p) 0.1 0.3 0.4 0.0 0.0

1,2,4-Man (p) 0.1 0.0 0.0 0.9 0.1

t-Gal (p) 0.0 0.3 0.0 0.0 0.0

t-GlcA 1.0 0.5 1.8 1.0 0.8 Total 4.3 6.2 6.7 5.7 (± 0.8) 8.3 5.3 6.75 (± 1.45) avalue is lower than 0.05 of Mol%.

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Figure A4.1 Division of stem transverse area into three main tissues of interest for total area analysis in wild-type (wt) and fla16 plants. Transverse sections of the stem at growth stage 6.5 (Boyes et al., 2001) stained with toluidine blue were divided into four main tissue types, epidermis (Ep), pith (Pt), cortex (Cr), and intra-vascular region (IVR). Pith, cortex and intra-vascular region are used for total area analysis whereas epidermis is use for cell area analysis (see Table 4.3). Scale bar = 500µm.

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Figure A4.2 Biomechanical properties of dry wt and fla16 basal stems (stage 6.9; (Boyes et al., 2001) in flexural tests. (A) No difference in stem diameter is observed in dried fla16 stems compared to wt. (B) A sharp break of stems during flexural tests was not observed for fla16 whereas 17% of wt stems snapped. There is no difference in the flexure stiffness analysed by circular (C), oval (E) and rectangle (G) models. Flexure strength of fla16 stem was significantly reduced in all models (D, F and H) compared to wt. Student T-test: P<0.01 **. Data are presented as mean ±SE.

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Figure A4.3 Tensile properties of fla16 and wt stems (stage 6.9 (Boyes et al., 2001)) at senescence completion. (A) No difference in basal stem diameter is observed in dried fla16 stems compared to wt whereas the fla16 middle stem diameter is significantly reduced (B). No differences in snapping profiles in either the basal (C) or middle (D) stems of fla16 and wt were observed. The tensile stiffness of fla16 was not different to wt in basal stems (E) whereas an increase in tensile stiffness occurred in the midddle (F) stems. No differences in the tensile strength of fla16 and wt was observed in basal (G) and middle stems (H). Student T-test: P<0.01 **. Data are presented as means ±SE.

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CHAPTER 5

COMPARATIVE AND FUNCTIONAL INVESTIGATION OF FLA11, FLA12 AND FLA16 THROUGH MUTANT STUDIES

128

5.1 Introduction The biological roles of only a few FLAs have been characterized, largely through analyses of their corresponding mutants (see Chapter 1.4.3). A number of fla single mutants either display no obvious phenotype or the phenotype is conditional (Shi et al., 2003; MacMillan et al., 2010; Johnson et al., 2011). This suggests functional redundancy between FLA members (Pickett and Meeks-Wagner, 1995; Kafri et al., 2009) and/or environmentally dependent roles for these FLAs (Rutter et al., 2017). FLA11, FLA12 and FLA16 have been shown to be involved in stem development and maintenance of stem biomechanical properties (see MacMillan et al. (2010) and Chapter 4). Due to the importance of wood quality in the forestry industry and crop losses in agriculture because of stem lodging, understanding the genetic relationships and molecular functions of these FLAs is of interest. FLA11 and FLA12 protein sequences are highly similar (57%) and they have largely overlapping expression profiles, suggesting they may act redundantly. There were no obvious morphological phenotypes and only minor changes in biomechanical properties in the single fla11 and fla12 mutants (MacMillan et al., 2010). In the double fla11 fla12 mutant decreased tensile strength and stiffness was observed as well as reduced cellulose content and altered cellulose microfibril angles (MFA) (MacMillan et al., 2010). This suggested functional redundancy exists between FLA11 and FLA12. The lack of gross morphological phenotypes in the fla11 fla12 double mutant suggests further functional redundancy might exist, for example, with other stem-specific FLAs such as FLA16. We have identified a role for FLA16 in maintaining cellulose deposition and stem biomechanics during stem growth (see Chapter 4). To examine if FLA16 has overlapping functions with FLA11 and FLA12 in the stem, all mutant combinations were generated including all double mutant combinations (fla11 fla12, fla11 fla16 and fla12 fla16) and the triple mutant (fla11 fla12 fla16). These mutant lines were then analysed for morphological differences, biomechanical properties, cell wall/cellulose composition and genetic interactions.

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5.2 Results

5.2.1 Generation of fla11, fla12 and fla16 double and triple mutants FLA16 displays functional roles in the inflorescence stem similar to those reported for FLA11 and FLA12 (see Chapter 4) (MacMillan et al., 2010). To investigate potential functional relationships between FLA11, FLA12 and FLA16, double fla11 fla12, fla11 fla16 and fla12 fla16 mutants and triple fla11 fla12 fla16 mutant plants were generated and compared to the single fla11, fla12 and fla16 mutants and wt. Single fla11 and fla12 as well as the double fla11 fla12 mutants (MacMillan et al., 2010) were crossed to the fla16 single mutant (see Chapter 4). All single, double and triple mutants were confirmed by genotyping in the resulting F3 or F4 generations (see Appendices Fig. A5.1), and were then investigated for stem phenotypes as described in Chapter 4.

5.2.2 Loss of FLA11, FLA12 and FLA16 causes a reduction in inflorescence stem length In comparison to wt, all single, double and triple fla mutants showed a consistent and significant reduction in the length of both total stem and first internode, in addition to the ratio of first internode to total stem length (Fig. 5.1, Table 5.1). Compared to wt, the fla16 single mutant had the largest reduction in stem length and was statistically shorter than all other single, double and triple fla mutant lines. There was no significant difference in the total stem length between the different combinations of double mutants. Comparisons between the double and triple mutants showed the triple fla11 fla12 fla16 mutant (36.1cm) was only statistically shorter than the double fla12 fla16 mutant (38.2cm). The length of the first internode of fla16 (4.2cm) was significantly shorter than wt (11.69cm) and shorter than fla11 fla12 (5.7cm) with no significant difference to any other mutant lines. Comparisons between fla mutant combinations showed the first internode of fla11 fla12 (5.7cm) was significantly longer than fla12 fla16 (4.5cm), however fla11 fla16 and fla11 fla12 fla16 were not significantly different to other mutant lines (Table 5.1). The ratio of first internode to total stem length showed all fla mutant lines were significantly shorter than wt, however comparisons between all fla mutant lines showed that fla11 fla12 (15%) was longer than fla12 fla16 (12%) (Table 5.1). 130

5.2.3 Total stem area is reduced in fla mutants

Stem area was analysed at the base and first node (immediately below the first branch) using sections taken from fresh tissue. Compared to wt, a consistent and significant reduction in total stem area occurred in all mutant lines, at both the base and first node, but no significant difference in the ratio between first node and basal area was observed (Table 5.1). In general, at both the base and first node, the transverse stem area ranking followed a descending order, with the largest area in wt, decreasing progressively in the single through to the fla11 fla12 fla16 triple mutants (Table 5.1). As the triple mutant generally showed more severe phenotypes in stem growth, it was therefore investigated further to determine if changes in cellular morphology could explain these phenotypic differences.

5.2.4 Cellular morphology is altered in triple fla11 fla12 fla16 mutant Transverse sections taken at the stem base and first node were examined for changes in tissue area, cell size and shape in the triple fla11 fla12 fla16 mutant compared to wt (Fig. 5.2, Tables 5.2, 5.3). In basal stem sections, a significant reduction in cortex and pith area was observed in the triple fla11 fla12 fla16 mutant compared to wt (Table 5.2). No significant changes between the triple mutant and wt were detected when measuring the area of the intra-vascular region, which included the vascular bundles (phloem, cambium and xylem) and interfascicular fibres (IFs). As a result, when comparing the ratio of the intra-vascular region to the total basal stem area it was significantly increased in the triple mutant compared to wt. In contrast, the pith area was significantly decreased in fla11 fla12 fla16 compared to wt (Table 5.2). Despite a reduction in cortex area, the ratio of cortex area to total stem showed an increase in the triple mutant compared to wt. These results suggest the changes in tissue area are differential rather than proportional. As changes in area can result from differences in cell number, cell size or both, these were also analysed. Comparison of the triple mutant cell sizes to wt showed a significant increase in xylem vessel cells, but no significant differences in the epidermis, IF, pith and cortex (Table 5.3). A count of the total number of cells present in a given

131 transverse section for selective cell types showed a significant reduction in the number of pith cells. A reduction in the apparent number of IF and xylem fibre cells was also observed, however, this was not significant compared to wt (Table 5.3). At the first node, a significant reduction in pith area was observed with no significant differences in the area of cortex and intra-vascular regions. The ratio of individual cell types to the total stem area at the first node followed the same pattern as described above for the basal stem (Table 5.2). Cell sizes were also the same in all cell types and cell number showed a reduction in only pith cells in the triple mutant compared to wt at the first node (Table 5.3). These data suggest that the reduced stem area in fla11 fla12 fla16 mutant stems is largely a result of a reduced number of pith cells at both the base and first node. In addition, at the base, a small portion of the area reduction was due to a reduced number of cells in the cortex. This is inferred from the reduction in cortex area with no significant difference in cell size At the base and first node, the intra-vascular region area in the triple mutant was comparable to wt. Given the reduced total area of the stem, this region then contributed a greater proportion in the triple mutant (Table 5.2). Other than a larger xylem cell size at the base, no differences in either cell size or number were observed. These results indicate that the loss of FLA11, FLA12 and FLA16 cause altered stem development and morphology. To determine if these differences impact on stem biomechanical properties, flexure tests were performed.

5.2.5 Stem biomechanical properties are not altered in double and triple fla mutants The biomechanical flexure properties of double (fla11 fla12, fla11 fla16, fla12 fla16) and triple fla11 fla12 fla16 mutants were examined using three-point-bending tests (see Chapter 2.27) on the basal and middle stem segments as described in MacMillan et al. (2010) (Table 5.4). The parameters measured were the breaking profile, flexure stiffness and strength. A large variation in the flexure breaking profile, categorised as having a sharp or slow/no break when the load reached the maximum, occurred in the basal stem segment. Sharp breakages occurred in approximately 11% of wt stem samples. In both fla11 fla12 and fla11 fla12 fla16, breakages were much more frequent, occurring in approximately 67% of the samples. In contrast, no breakages occurred in fla11 fla16 132 and fla12 fla16 samples. Tests of the middle stems did not yield any breaking profile for any of the samples tested, including wt (Table 5.4). At the base, only the fla11 fla12 fla16 triple mutants displayed a significant increase in flexure stiffness. Flexural strength was also slightly increased compared to wt and fla11 fla12 although this was not significant (Table 5.4). At the middle stem, only fla11 fla16 double mutants showed a significant increase in the flexure stiffness compared to wt (Table 5.4). The significant increase of flexure stiffness indicated there was a difference in the biochemical properties of the fla11 fla12 fla16 triple mutant stems compared to wt. Such differences may be caused by either altered wall organization and/or composition of the wall polysaccharides, and was therefore investigated further.

5.2.6 The fla mutant stems have altered polysaccharide composition Linkage analyses were undertaken to determine if the polysaccharide composition of fla11 fla12 fla16 triple mutant stems was altered compared to wt (Fig. 5.3A, Table A5.1). In the fla11 fla12 fla16 triple mutants, an increase in glucuronoxylan and a decrease in cellulose was observed (Fig. 5.3A). The decrease in cellulose content was confirmed by the acetic/nitric cellulose crystallinity assay (Fig. 5.3B) and was observed in all fla mutant stems (single, double, and triple) (Table 5.1). This reduction in crystalline cellulose was previously reported for the fla11 fla12 double mutant (MacMillan et al., 2010). The reduced levels of crystalline cellulose observed in fla mutant stems could result from altered expression of CesAs, and/or disruption of CesA trafficking, stability or activity. To investigate this further, the expression of selected cellulose synthases (CesAs) was examined.

5.2.7 Analysis of CesA transcript levels in wt and fla11 fla12 fla16 mutants Q-PCR was undertaken to investigate the transcript levels of CesAs in stems of fla16, fla11 fla12 and fla11 fla12 fla16 mutants compared to wt. Transcript levels of CesAs that are active in both primary (CesA1, 3 and 6) and secondary (CesA4, 7 and 8) wall cellulose biosynthesis (Persson et al., 2007) were compared in the top, middle and basal stem segments (Fig. 5.4). The expression of CesA1, CesA3 and CesA6 showed no clear differences between the fla16 single, fla11 fla12 double and fla11 fla12 fla16 triple mutants compared to wt 133 in any of the stem sections (Fig. 5.4). For CesAs active in secondary wall synthesis, the expression of CesA4 and CesA8 did not show any clear differences between the mutants and wt whereas CesA7 was consistently higher in the fla mutants in all stem sections. This was proportional to the number of FLAs missing, with the greatest increase in the fla11 fla12 fla16 triple mutant compared to wt (Fig. 5.4). These results suggest the expression of CesA7 is increased in response to loss of FLA11, FLA12 and FLA16 activity.

5.2.8 FLA11, FLA12 and FLA16 show an epistatic relationship for stem length Mutants in fla11, fla12 and fla16 were found to display a number of similar phenotypes in the stem. To investigate if these FLAs are likely to act in the same functional pathways, genetic interaction studies were performed for the most robust stem-length reduction phenotype. The interactions could be described as additive if the double mutants show a combination of the phenotypes observed in single mutants; epistatic if the double mutants present a trait similar to only one single mutant of the two; suppressive if the double mutants phenotype is close to that which occurs in wt; and synergistic when the phenotype of the double mutants exceeds that displayed by either single mutations and often indicates a functional interaction (Perez-Perez et al., 2009). It is expected that if the FLAs act in independent pathways the decreases in length that occur in the single mutants would be additive in the double mutants. That is, the same relative decrease in length that occurs between the wt and single mutant will also occur between the single mutant and double mutant. If the FLAs act in the overlapping pathways, epistasis, suppression or synergy could occur. Analyses of the double fla mutants showed no decrease in length compared to the single mutants in their respective groups (compare Figs 5.6A, 5.6B, 5.6C). Since FLA11 and FLA12 are orthologues shown to act redundantly, the single fla16 mutant was also compared to the double fla11 fla12 mutants to determine if these three FLAs are likely to act in the same pathway. The same relative decrease in stem length of fla16 (25%) compared to wt did not occur in the triple mutant compared to the double fla11 fla12 mutant (4%) (Fig 5.6D). These data indicate that FLA11, FLA12 and FLA16 act in pathways to regulate the stem length and their genetic interaction could to be epistatic.

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5.3 Discussion We endeavoured to reveal if a genetic interaction occurs between FLA11, FLA12 and FLA16. A key finding of this preliminary study is the similarity of phenotypes in single, double and triple mutants suggesting they act in similar/the same pathway(s). Some inconsistencies do occur suggesting further work is needed to fully dissect/understand the relationship between these FLAs during stem development.

5.3.1 FLA11, FLA12 and FLA16 likely have overlapping roles regulating stem growth in Arabidopsis The genetic interaction between FLA11, FLA12 and FLA16 appears to be largely epistasis based on analysis of stem length. Given stem development is regulated by many factors, including hormones, biomechanics and complex feedback pathways, it is possible the FLAs act in different gene regulatory networks. No changes in either stem length or cellular morphology have previously been observed for fla11, fla12, or fla11 fla12 compared to wt (MacMillan et al., 2010). In fla16 mutants, reduced stem length likely results from reduced cell proliferation rather than cell elongation (see Chapter 4.2.5). In future, it would be interesting to determine if the reduction in stem length observed in our study in fla11, fla12 and fla11 fla12 mutants result from altered cell proliferation. Unlike stem length, stem area appeared to be additive and in fla11 fla12 fla16 mutants results from a significant reduction in pith cell number and area (Table 5.2). An increase in intra-vascular area as a proportion of the total basal stem is also seen in the fla11 fla12 fla16 triple mutants. Changes in the biomechanical properties of IF and xylem cells in fla mutants could result in altered mechanical stress patterns in the growing stem and changes in cellular morphology. Changes in mechanical properties of walls is known to induce signalling to ensure that wall integrity is maintained (Amanda et al., 2016; Amanda et al., 2017). As cells with secondary walls, including IF and xylem cells, contribute significantly to the integrity of stem strength (Zhong et al., 2001), changes to either wall composition or architecture will have significant effects on stem development. Mechanical perturbation of plants during growth frequently results in reduced stem length and changes in cellular morphology. For example, mechanically perturbed Arabidopsis plants developed proportionally less pith and IF tissue and more cortex (Paul-Victor and Rowe, 2011). Reduced cellulose content and altered 135 biomechanics observed in fla11 fla12 fla16 stems suggest wall integrity is compromised and likely triggers compensatory changes in plant growth. This is similar to an Arabidopsis fragile fibre (fra1) mutant that has reduced mechanical strength in fibres due to changes in the orientation of cellulose MFs, with no obvious changes in fibre cell morphology. fra1 mutant stems break more easily, display reduced stem length and changes in the length of pith cells (Zhong et al., 2002). Although the role of mechanical signals in regulating organ growth in the shoot apical meristem has been elegantly demonstrated, relatively little is known about their role in stems (Hamant et al., 2008; Uyttewaal et al., 2012; Maeda et al., 2014; Sampathkumar et al., 2014). Further investigation into how FLAs may influence mechanical properties in stems and their impact on stem growth and development is needed. FLAs are known to influence the MFA in stems (MacMillan et al., 2010). MFA is an important determinant of biomechanical properties and it remains unclear how, and which, FLAs are involved in its regulation. A study of three Eucalyptus stem FLAs over- expressed in tobacco and Eucalyptus showed only EgrFLA2 caused changes in MFA (MacMillan et al., 2015). This suggest that some FLAs have become functionally specialized for MFA regulation in stems. Determining the MFA in all single, double and triple fla mutant stems in Arabidopsis, and if over-expression has the opposite effect, would be informative. This would help determine whether individual or a combination of stem FLAs are required to ‘fine-tune’ MFA. Increased flexural stiffness and a tendency towards increased flexural strength was also observed in the basal stems of fla11 fla12 fla16 mutants. Interestingly, fla11 fla12 showed a trend to decreased flexural stiffness suggesting a complex interplay between FLA11, FLA12 and FLA16 to regulate stem biomechanics. The biomechanical properties of stems are regulated by a number of traits including the number and size of cells, and the thickness and composition of secondary walls (Burgert, 2006). In future, tensile properties should also be investigated and biomechanical properties correlated to changes in cellular morphology and wall composition. Changes in either cell wall composition or architecture in stems frequently results in a collapsed cell phenotype due to the significant mechanical load from the plant body mass and pressures needed to transport water and nutrients (Hamann, 2012; Hao and Mohnen, 2014). A study by Persson et al. (2005) found that a fla11 allele, also called irx13, showed a collapsed xylem phenotype. Many of the irx mutants have defects 136 in cellulose synthesis and in addition to collapsed xylem, display reduced cell wall thickness in IFs (Turner and Somerville, 1997; Hao and Mohnen, 2014). Despite the reduction in cellulose in the total stem cell wall material of all fla mutants, no collapsing of xylem or obvious changes in IF wall thickness was observed by either MacMillan et al. (2010) or in this study, suggesting that some fla11, fla12 and fla16 phenotypes might be conditional. In triple fla11 fla12 fla16 mutants, an increased size of xylem cells at the stem base was observed in transverse sections. Further, detailed study of the cell wall thickness and composition of xylem and IF cells using TEM and immuno-labelling with antibodies recognising cellulose, glucuronoxylans, AGPs and pectins, that are known to influence the mechanical properties, is needed and would reveal changes in micro- heterogeneity of wall organisational changes at the cellular level. Reduced length of inflorescence stems is observed in mutants lacking Hyp O-galactosyltransferases involved in AGPs glycosylation (Ogawa-Ohnishi and Matsubayashi, 2015). A spatio- temporal study of cell wall changes in stems suggested certain AGP sub-classes are specifically expressed during early IF development (Hall et al., 2013). If loss of stem FLAs results in changes to wall composition it will be important to understand how this contributes to the cellular and growth phenotypes. With further detailed examination using a greater number of biological replicates and in different growth conditions of all single, double and triple mutant combinations, it should be possible to begin to clarify relationship(s) between the biomechanical, cell morphological, wall and growth changes.

5.3.2 Genetic redundancy might contribute to the genetic interaction between FLAs

Gene duplications are thought to impart phenotypic robustness to an organism resulting in genetic redundancy (Conant and Wagner, 2004; Hsiao and Vitkup, 2008; Kafri et al., 2008). From an evolutionary perspective, genetic redundancy increases the chance for organisms to survive and evolve in a competitive environment (Kirschner and Gerhart, 1998). This is achieved by different evolutionary fates of duplicated genes, namely non-functionalisation (i.e. loss of gene function), neo-functionalisation (i.e. new gene function) and sub-functionalisation (i.e. both duplicated genes sub-divide the ancestor gene’s function) (Briggs et al., 2006). FLAs belong to a multi-gene family with multiple homologous members in respective groups, potentially presenting genetic redundancy (Johnson et al., 2003; MacMillan et al., 2010). Functional redundancy of

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FLAs is exemplified by FLA11 and FLA12, which show more prominent phenotypes in the double fla11 fla12 than the individual single fla11 and fla12 mutants (MacMillan et al., 2010; MacMillan et al., 2015). In the current study, loss of FLA11, FLA12 and FLA16 results in similar biological changes to stem growth suggesting they might carry similar functions. However, the broader expression of FLA16 suggests it may also have additional functions from FLA11/FLA12 at other growth stages (see Chapter 3). It is possible that a defined combination of FLAs and/or interacting partners is needed at different developmental stages. This is supported by studies of periostin, a multi-FAS1 domain containing protein in animals involved in multiple signalling pathways including cell migration, adhesion and proliferation (Landry et al., 2017). Periostin in the extracellular matrix of animal cells has been shown to interact with cell-surface receptors in response to mechanical stress and initiate integrin-associated intracellular signalling. Periostin associated proteins include fibronectin, tenascin-C, laminin and collagens type I and V, that are responsible for determining tissue biomechanical properties, matricellular proteins and enzymes that catalyse crosslinking between matrix proteins (Kii and Ito, 2017). FAS domains are proposed to facilitate cell-adhesion (Shi et al., 2003)in a broad taxonomy of living organisms, including algae, insects and humans, via molecular interactions and/or homophylic/heterophilic interactions (Elkins et al., 1990a; Huber and Sumper, 1994; Kim et al., 2000; Kim et al., 2002; Clout et al., 2003). Potentially, different homodimer or heterodimer combinations of stem FLAs could exist in different tissues and stages of secondary wall development to modulate function. Differential glycosylation of these FLAs might also result in specific molecular interactions as supported by the heterogeneity of glycans (Strasser, 2016). If FLAs act in a similar way to Periostin, then the receptors they interact with could be involved in triggering downstream signalling pathways. This could include regulation of CesAs as indicated by up-regulation of CesA7 in fla mutants. Identification of the interacting partners of FLAs and the transcriptional changes associated with changes in FLA function is an avenue for future research (see Chapter 6 for further discussion). The genetic interaction study of fla11, fla12 and fla16 requires further investigation. This might be due to limited sampling sizes for statistical analyses and insufficient technical replicates. Further investigation should be undertaken with increasing sampling size and multiple repeated experiments. Examination of mutant 138 phenotypes under different stress and hormone conditions could also uncover additional tissue or developmental specific phenotypes. In addition, genetic redundancy with other FLA members needs to be considered. Analysis of mutants for other group A (FLA11/FLA12) and group B (FLA16) FLA members may reveal if further redundancy exists. In the future, gene editing approaches, such as CRISPR (for Clustered Regularly Interspaced Short Palindromic Repeats) (Ran et al., 2013), could be used to edit the gene activities (i.e. knock-down/out) of FLAs in the plant genome. Theoretically, CRISPR is advantageous over traditional T-DNA insertion as it is more targeted (Camporesi and Cavaliere, 2016). This might allow us to mutate multiple FLAs in the same plant line for genetic redundancy analyses. In addition, if gene editing becomes more accurate, systematic analyses of the function of specific amino acids within the FLA functional domains (FAS, N-/O-glycosylation motifs and GPI-anchor) could be undertaken.

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Table 5.2 Measurements of tissue area in transverse sections of mature stems from fla11fla12fla16 and wta. wt fla11fla12fla16 nb Tissue area (mm2) Base Cortex 0.25 (± 0.02) 0.19c (± 0.01) 4-5 Cortex/total area (%) 19 (± 0.56) 22 (± 0.64) 4-5 Intra-vascular regiond 0.47 (± 0.04) 0.39 (± 0.03) 4-5 Intra-vascular region/total area (%) 37 (± 1.06) 45 (± 0.79) 4-5 Pith 0.56 (± 0.04) 0.29 (± 0.03) 4-5 Pith/total area (%) 44 (± 1.47) 34 (± 0.76) 4-5 First node Cortex 0.24 (± 0.01) 0.23 (± 0.02) 4 Cortex/total area (%) 17 (± 0.50) 22 (± 0.34) 4 Intra-vascular region 0.45 (± 0.02) 0.41 (± 0.03) 4 Intra-vascular region/total area (%) 33 (± 0.49) 39 (± 0.46) 4 Pith 0.68 (± 0.04) 0.42 (± 0.02) 4 Pith/total area (%) 50 (± 0.47) 40 (± 0.45) 4 aStem material at growth stage 6.9 (Boyes et al., 2001). bn is biological replicates cBold text in numeral data column indicates data value presented as ± SE is statistically significant at p<0.05 using Student T test. dIntra-vascular region includes interfascicular fibres and vascular bundles.

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Table 5.3 Measurements of cell size and number in transverse sections of stems of fla11fla12fla16 and wta. wt fla11fla12fla16 n Base Cell size (µm2) Epidermis 307.26 (± 42.37) 308.23 (± 28.36) 4-5 Interfascicular fibre 217.33 (± 12.50) 232.57 (± 20.95) 4-5 Cortex 216.26 (± 23.76) 232.45 (± 36.58) 4-5 Xylem vessel 306.09 (± 20.02) 397.74b (± 12.73) 4-5 Pith (µm2) 1290.18 (± 70.68) 1279.39 (± 94.64) 4-5 Cell number Total interfascicular fibre cell number 1100.00 (± 145.34) 855.60 (± 85.10) 4-5 Total xylem vessel cell number 1269.25 (± 125.45) 953.60 (± 133.01) 4-5 Total pith cell number 473.00 (± 24.20) 247.60 (± 33.33) 4-5 First node Cell size (µm2) Epidermis 177.24 (± 12.51) 204.57 (± 21.20) 4 Interfascicular fibre 201.21 (± 14.93) 207.87 (± 15.27) 4 Cortex 157.41 (± 31.64) 170.11 (± 9.14) 4 Xylem vessel 354.96 (± 17.17) 417.77 (± 28.75) 4 Pith 1660.46 (± 99.85) 1430.04 (± 58.96) 4 Cell number Total interfascicular fibre cell number 813.00 (± 67.44) 763.25 (± 100.28) 4 Total xylem vessel cell number 838.25 (± 72.46) 941.75 (± 75.85) 4 Total pith cell number 443.00 (± 12.42) 336.00 (± 15.53) 4 aStem material at growth stage 6.9 (Boyes et al., 2001). bBold text in numeral data column indicates data value presented as ± SE is statistically significant at p<0.05 using Student T test.

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Table 5.4 Measurement of biomechanical flexure properties of stems of fla11 fla12, fla11 fla16, fla12 fla16, fla11 fla12 fla16 and wt. Unit (N/mm2)a wtb fla11fla12c fla11fla16d fla12fla16e fla11fla12fla16f n Basal stem Flexural strength 74.78 68.94 72.27 79.31 93.32 6- (± 5.02) (± 6.35) (± 5.71) (± 9.85) (± 10.49) 9 Flexure stiffness 6161.85 5304.25 7486.70 7822.10 7881.98g 6- (± 258.48) (± 528.04) (± 1194.53) (± 1238.31) (± 799.83)b,c 9 Flexure breaking (%) 11 67 0 0 67 6- 9 Mid stem Flexure strength 99.51 84.21 95.73 84.72 86.23 6- (± 17.47) (± 13.14) (± 13.65) (± 7.77) (± 17.18) 9 Flexure stiffness 11128.26 10871.24 13930.57 13271.28 14601.67 6- (± (± 2098.57) (± 2509.79)b (± 1967.35) (± 6174.64) 9 3136.80) Flexure breaking (%) 0 0 0 0 0 6- 9 aAll biomechanical strength and stiffness are recorded as N/mm2 representing Newton/squared millimetre. bwt, cfla11fla12, dfla11fla16, efla12fla16, ffla11fla12fla16 plant lines were grown to maturity and dried. gBold text indicates a statistically significant difference at p<0.05 using Student T test to the plant lines indicated next to the bracket. Data is shown as average ± SE. n is biological replicates.

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Figure 5.1 Stem length phenotypes of fla11, fla12 and fla16 single, double and triple mutant plants and wt. At post-maturity (growth stage 6.9) (Boyes et al., 2001) fla11, fla12 and fla16 single, fla11 fla12, fla11 fla16 and fla12 fla16 double and fla11 fla12 fla16 triple mutants showed reduced stem length compared to wt. Scale bar = 2cm.

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Figure 5.2 Light micrographs of transverse sections showing cellular morphology of stems of wt and fla11fla12fla16 mutant. Toluidine blue staining of transverse sections of wt (A, C) and fla11 fla12 fla16 (B, D) taken at the base and first node of stems show the reduction in stem area of fla11 fla12 fla16 mutants. Scale bar = 100 µm. a, b, c and d are images representing boxed areas in A, B, C and D showing cell types: pith (Pt), xylem (Xy), phloem (Ph), cambium (Cm), cortex (Cr), interfascicular fibre (IF), starch sheath (SS) and epidermis (Ep). Scale bar = 50 µm.

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Figure 5.3 Analysis of cell wall polysaccharide composition (A) and crystalline cellulose (B) levels in mature stems of fla11 fla12 fla16 mutants and wt. (A) Polysaccharide composition derived from linkage analysis of fla11 fla12 fla16 triple mutant stems shows an increase in glucuronoxylan and a reduction of cellulose compared to wt. (B) Acetic/Nitric crystalline cellulose assays show a reduction of crystalline cellulose in fla11 fla12 fla16 mutant stems. Error bars show ± SE. Raw data and calculations are in Appendices, Tables A5.1, 5.2, 5.3.

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Figure 5.4 Q-PCR analyses of CesA transcript abundance in the top, middle and basal stems of wt, fla16, fla11fla12 and fla11fla12fla16 plants. Q-PCR analyses of transcript abundance of CesAs involved in primary (1, 3 and 6) and secondary (4, 7 and 8) wall biosynthesis in top (A), middle (B) and basal (C) stem segments show CesA7 expression is increased in fla16, fla11 fla12 and fla11 fla12 fla16 compared to wt. Other CesAs do not show obvious expression differences between the plant lines. Error bars = ± SE.

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Figure 5.6 Double and triple mutant analyses of fla11, fla12 and fla16 total stem length. Double mutants of fla11 fla12 (A) . Double mutants of fla11 fla16 (B) and fla12 fla16 (C) show epistatic phenotypes as assessed by the total stem length. (D) Assessment of fla11 fla12 fla16 triple mutants suggest they act in the same genetic pathway. Values are present as mean ±SE (n = 18-32).

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APPENDICES TO CHAPTER 5

Table A5.1 Calculation of polysaccharide composition based on linkage analysis of fla11 fla12 fla16 triple mutant and wt stems. (Refer to Fig. 5.3)

wt fla11 fla12 fla16

1 2 3 Mean (± SE)a 1 2 3 Mean (± SE)

Arabinan 1,5-Ara (f) 1.9 2.2 2.3 1.8 1.6 1.9

1,2,5-Ara (f) 0.5 0.5 0.4 0.4 0.3 0.3 t-Ara 2.2 2.4 2.7 1.9 1.8 2.2

Total Arabinan 4.6 5.1 5.4 5.0 (± 0.2) 4.1 3.7 4.4 4.1 (± 0.2)

Type I AG 1,4-Gal (p) 0.9 2.0 1.3 1.0 1.4 1.1

Total Type I AG 0.9 2.0 1.3 1.4 (± 0.3) 1.0 1.4 1.1 1.2 (± 0.1) Type II AG 1,3-Gal (p) 0.0 0.0 0.0 0.0 0.0 0.0

1,6-Gal (p) 0.0 0.0 0.2 0.1 0.1 0.2

1,3,6-Gal (p) 0.0 0.1 0.1 0.0 0.1 0.1

t-Ara 0.0 0.1 0.1 0.0 0.1 0.1

Total Type II AG 0.0 0.2 0.4 0.2 (± 0.1) 0.1 0.3 0.4 0.3 (± 0.1) Glucuronoxylan 1,4-Xyl (p) 14. 19. 20.5 21.9 27. 29. 0 3 2 8 1,2,4-Xyl (p) 0.8 1.3 1.2 1.6 2.4 2.0

1,3,4-Xyl (p) 0.0 0.2 0.4 0.5 0.7 0.8

t-GlcA 0.8 1.5 1.6 1.8 1.9 2.4

Total 15. 22. 23.7 20.5 (± 2.5) 25.8 32. 35 31 (± 2.7) Glucuronoxylan 6 3 2 Heteromannan 1,4-Man (p) 3.0 3.6 3.1 3.6 3.2

1,4,6-Man (p) 0.2 0.5 0.4 0.3 0.3 0.4

1,4-Glc (p) 3.0 3.6 3.1 3.6 3.2 3.6

1,4,6-Glc (p) 0.2 0.5 0.4 0.3 0.3 0.4

t-Gal 0.4 0.9 0.9 0.7 0.7 0.8

t-Man Total Heteromannan 6.8 9.1 7.9 7.9 (± 0.7) 4.9 8.1 8.4 7.1 (± 1.1)

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Table A5.1 continued. Xyloglucan 1,4,6-Glc (p) 2.8 2.8 2.4 2.4 2.1 2.3 1,4-Glc (p) 2.8 2.8 2.4 2.4 2.1 2.3 1,2-Gal (p) 0.4 0.8 0.7 0.6 0.6 0.7 t-Fuc (p) 0.5 0.3 0.4 0.3 0.2 0.3 t-Xyl 1.9 2.4 2.2 1.9 1.8 2.2 Total Xyloglucan 8.4 9.1 8.0 8.5 (± 0.3) 7.6 6.8 7.8 7.4 (± 0.3) Cellulose 60. 47. 47.2 48.5 39. 36. 1,4-Glc 1 0 7 9 Total Cellulose 60. 47. 47.2 51.4 (± 4.3) 48.5 39. 36. 41.7 (± 3.5) 1 0 7 9 Assigned linkage Total 97. 97. 97.4 97.2 (± 0.1) 98.0 97. 97. 97.8 (± 0.1) 3 0 9 6 Unassigned t-Man (p) 0.0 0.2 0.1 0.1 0.1 0.2 linkage t-Glc (p) 0.9 1.7 1.3 0.8 1.0 0.9 t-GlcA 1.6 0.9 1.2 0.0 0.0 0.0 1,3,4-Glc (p) 0.3 0.5 0.5 0.5 0.5 0.6 1,2,4-Glc (p) 0.3 0.4 0.3 0.3 0.4 0.3 1,3,4,6-Glc (p) 0.0 0.0 0.0 0.0 0.0 0.1 1,2,3,4-Xyl (p)b 1.6 4.0 5.1 4.4 7.9 5.4 t-Gal (p) 1.2 0.3 0.4 0.3 0.0 0.2 Total 7.1 7.6 8.9 7.9 (± 0.5) 6.4 9.9 7.7 8 (± 1.0) aMean ± Standard error (SE) of biological replicates. b likely undermethylation

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Table A5.2 Calculation of crystalline cellulosea in stems of fla mutants and wt. Plant line Sampleb AIRc weight (mg) Cellulose (mg) Cellulosec % Mean (± SE)d wt 1 10.33 5.08 49.18 50.54 (± 1.38)

10.56 5.19 49.15

2 11.48 5.94 51.74

11.65 6.07 52.10 fla11 1 37.87 13.46 35.54 38.64 (± 1.17)

41.12 16.20 39.40

1 43.36 16.63 38.35

32.66 13.48 41.27 fla12 1 54.15 23.64 43.66 45.94 (± 0.38)

63.15 30.93 48.98

2 28.63 12.81 44.74

42.53 19.73 46.39 fla16 1 12.58 5.36 42.61 42.30 (± 0.18)

10.35 4.31 41.64

2 10.99 4.60 41.86

10.14 4.37 43.10 fla11 fla12 1 10.38 4.71 45.38 46.26 (± 0.80)

12.14 5.53 45.55

2 11.61 5.44 46.86

11.53 5.45 47.27 fla11 fla16 1 10.25 4.86 47.41 47.39 (± 0.37)

11.35 5.46 48.11

2 12.43 5.82 46.82

12.60 5.95 47.22 fla12 fla16 1 10.18 4.65 45.68 46.20 (± 1.07)

12.65 5.64 44.58

2 11.21 5.37 47.90

12.59 5.87 46.62 fla11 fla12 fla16 1 10.16 4.36 42.92 41.86 (± 0.45)

10.03 4.00 39.88

2 11.53 4.88 42.32

11.49 4.86 42.30 aCrystalline cellulose prepared from cAlcohol insoluble residues (AIR) of the stem cell wall material using acetic/nitric treatment. b2 biological replicates with 2 technical replicates each. cPercentage of crystalline cellulose content from acetic/acid pre-treated stem cell wall AIR material. dMean ± Standard error (SE) of biological replicates.

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Figure A5.1 Schematic diagram of T-DNA insertions in FLA11, FLA12 AND FLA16. Structures of FLA11, FLA12 and FLA16 with T-DNA insertions are shown in diagramatic form. Regions encoding the signal peptides (yellow), O-linked glycomotifs (red), fasciclin domains (blue), GPI-anchor recgnition sequences (green), regions of unknown function (white) and the intron in FLA16 (black) are indicated. The start (ATG) and stop (TAA/TGA) codons and location/direction of primers used for genotyping are indicated.

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Figure A5.2 Genotyping of homozygous fla11, fla12 and fla16 mutants by PCR. (A) Expected product sizes and primers used for genotyping of fla11, fla12 and fla16 T- DNA insertion lines and their respective wt alleles. (B, C, D) gDNA extracted from fla11, fla12, fla16 single mutants and wt was amplified by PCRs using mutant- and wt-specific primers as listed in (A). All single fla mutant samples yield only mutant- but not wt- specific products; alternatively all wt samples yield only wt- but not mutant-specific products showing all fla single mutant lines are homozygous. (E) Validation of homozygous double fla11 fla12, fla11 fla16, fla12 fla16 and triple fla11 fla12 fla16 mutant lines using genotyping primers as mentioned. Actin primers were used for validation of PCR efficiency and gDNA quality (yielding specific products of 180bp) showing both are sufficient. Specific fla mutant products are amplified from gDNA in respective mutant combinations and no products yielded using the wt-specific primers indicating the fla mutants are homozygous. PCR products are separated on a 1% agarose gel.

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Figure A5.3 Phenotypes of dark-grown wt and fla mutant seedlings treated with isoxaben. Dark-grown 4 d.o. seedlings of fla mutants grown in either the absence (A) or presence of isoxaben (B) compared to wt. (C) In the absence of isoxaben, fla12 and fla11fla16 show no differences in hypocotyl length compared to wt. fla11, fla12fla16 and fla11fla12fla16 show increased hypocotyl length and fla16 and fla11fla12 show reduced hypocotyl length. All fla mutants show a significant reduction in hypocotyl length compared to wt with isoxaben treatment. (D) Comparison of hypocotyl length in the ratio of presence/absence of isoxaben for each line show fla11fla12 mutants have a similar reduction to wt and all other fla mutants have a greater percent reduction in length than wt. Statistical analyses using Student T-test (n = 16 - 42, p < 0.05). Scale bar = 1cm in (A) and (B). Error bar = ± SE.

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CHAPTER 6

SUMMARY

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6.1 Summary of current research

This research has provided new insights into the understanding of FLA11, FLA12 and FLA16 in stem development. Reporter constructs have shown for the first time the location of FLA11, FLA12 and FLA16 proteins at the PM and wall of cells with secondary walls (Chapter 3). To our knowledge this study is the first time FLA11, FLA12 and FLA16 have been enriched to confirm that they display characteristics of glycoproteins. Data from the analyses of the fla16 mutant has revealed a novel role for FLA16 in stem biomechanics and regulation of cellulose levels (Chapter 4). Preliminary genetic interaction studies show a likely epistatic relationship between FLA11, FLA12 and FLA16 suggesting they act in overlapping pathways (Chapter 5). This research also provided informative suggestions for future investigations towards uncovering functional mechanism(s) of these fascinating molecules during secondary wall development. Fluorescent reporter fusions of FLA11, FLA12 and FLA16 suggest they are glycosylated to different extents, likely with both N- and O-glycans. Glycosylation is proposed to contribute to FLA function in several ways. For example, stabilising the tertiary conformation, assisting secretion and trafficking, acting as ligands in signalling pathways due to the micro-heterogeneity of glycan structures, and facilitating interaction with functional partners (Chapter 3). The location of these putative glycoproteins in the PM and wall of the same cell type suggests they may have similar functions, potentially regulating wall biosynthesis and architecture. This suggests genetic redundancy to enable robustness to environmental conditions and against a loss of function (Chapter 5).

6.2 Hypothetical models of FLA functions Given the potential for FLAs to interact with other proteins/carbohydrates (see Chapter 1.4) and their involvement in potentially regulating wall biosynthesis/integrity, we propose four alternate models (Fig 6.1) for the stem-specific FLAs to function in cell wall organisation. In model A, FLA11, FLA12 and FLA16 are suggested to interact with the cellulose synthase complexes (CSCs), either with the CesA catalytic sub-units or their associated regulatory proteins (Fig. 6.1A). Proteins associated with CSCs have been shown to modulate cellulose biosynthesis (Persson et al., 2005; Schneider et al., 2016). A number

157 of regulatory proteins have been identified through co-expression studies of the CesAs and include COBRA (Persson et al., 2005) and KORRIGAN1 (KOR1) (Bhandari et al., 2006). COBRA is a GPI-anchored protein proposed to be involved (by a yet to be defined mechanism) in controlling the orientation of cellulose microfibrils (Roudier et al., 2005). FLA11 and FLA12 could potentially interact with COBRA, as there is evidence that GPI- anchored proteins associate in lipid microdomains (Laude and Prior, 2004). KOR1 is an endo-1,4-D-glucanase integrated into the CSC complex on the extracellular face of the PM and proposed to play roles in intracellular CSC trafficking and regulating the length of the glucan chains (Vain et al., 2014). Although FLA11, FLA12 and FLA16 are co- expressed with secondary CesAs (Persson et al., 2005; Persson et al., 2007; Li et al., 2016b), it is unclear how they influence cellulose production and wall integrity. FLAs could potentially stabilize the secondary CSC complexes, which is important during the transition from a primary to a secondary wall. In model B, rather than a direct interaction with CSCs, FLAs could influence cellulose levels indirectly via interaction with wall integrity sensors (Fig. 6.1B) (also see Chapter 3). Mutants in THESEUS1, encoding a member of the CrRLK family involved in wall integrity sensing, suppress the effects of cellulose deficiency in the walls of cesa6 mutants (Hematy et al., 2007; Merz et al., 2017). Therefore, THESEUS1 is proposed to repress growth in response to reduced cellulose levels and/or cell wall damage. FLAs could interact with either THESEUS1, other members of the CrRLK family, or WAKs (Ringli, 2010) that have been implicated in signalling functions in response to changes in the wall. In support of this, FLA4 is proposed to interact with the RLKs, FEI1/FEI2, in roots (Xu et al., 2008; Basu et al., 2016). FEI1/FEI2 act independently of COBRA and CesA6 to regulate cell wall integrity. FLA4 is a putative ligand of FEI1/FEI2 as genetic interaction studies suggest they act in the same pathways. Alternatively, the GPI-anchored FLA4 could create a ‘spacer’ between the wall and PM so that wall-PM-cytoplasmic signalling involved in regulating CSCs occurs unimpeded. A role for FEI1/FEI2 in secondary walls is yet to be demonstrated, and what RLKs act in these cell types is unclear. The PM located WAKs are potential candidates as they are able to interact with structural wall proteins such as glycine-rich proteins and pectins in the wall to fulfil their role in modulating cell growth and expansion and stress responses (Ringli, 2010). FLAs could potentially associate with WAKs at the wall-PM-cytoplasmic interface and act in wall sensing (Gens

158 et al., 2000; Ma et al., 2017). Further investigation is needed to identify interacting partners of FLA11, FLA12 and FLA16 (see Chapter 3). In model C, FLAs could potentially interact with cellulose microfibrils leading to a defined microfibril angle (MFA) (Fig 6.1C). This hypothesis is based on our observations that fla16 mutants display altered stem biomechanics (Chapter 4) and previous studies showing changes in biomechanics and MFA in fla11 fla12 mutants (MacMillan et al., 2010). The change in cell wall stiffness observed in fla16 mutants is a strong indication of changes in MFA (MacMillan et al., 2010). In model C we propose the FAS domain(s) and/or AG glycans of FLAs interact with microfibrils via protein- carbohydrate and/or carbohydrate-carbohydrate interactions, respectively. FLAs could be present between microfibrils and guide alignment during the transition from primary to secondary walls (Li et al., 2016a). As the microfibrils extend and the wall thickens, GPI-anchored FLAs could be released from the PM into the wall (Neumann et al., 2007). Enzymic cleavage of the GPI- anchored FLAs could be undertaken by phospholipases C/D (PLC/D) that have been demonstrated to cleave GPI-anchored proteins from the PM in bacteria, fungi and mammals (Lehto and Sharom, 1998; Mann et al., 2004; Elortza et al., 2006; Rupwate and Rajasekharan, 2012; Fujihara and Ikawa, 2016). To date, no GPI-specific PLs have been characterized from plants although there are many candidates with sequence similarity to PLs and structural characterization of the GPI-anchor present on the soluble isoform of Pyrus communis AGP1 suggests that this was cleaved either by a PLD, or a PLC (Oxley and Bacic, 1999) . Although the dynamic release of GPI anchored proteins has not been visualised in plants, immuno-gold labelling of a COBRA-like protein in rice, BC1, found it was located at the PM in cells with thin walls, and in cells with thickened, secondary walls, present in the wall. BC1 was also shown to be PI-PLD sensitive (Li et al., 2003; Liu et al., 2013). Both the GPI-anchored FLA11, FLA12 and non-GPI anchored FLA16 were found in PM and wall extracts suggesting they might undergo release similar to BC1 and influence microfibril orientation at the PM and in other layers of the secondary wall (Fig. 6.1C).

In model D, the wall matrix polysaccharides, including pectins and non-cellulosic polysaccharides could interact through non-covalent and/or covalent association(s) with the O-glycans of FLAs (Fig. 6.1D). This could influence the wall matrix properties and in

159 turn define the arrangement of microfibrils as they are extruded into the wall (Peaucelle et al., 2015; Busse-Wicher et al., 2016). This hypothesis is also based on the findings of altered stem stiffness and MFA in the fla mutants ((MacMillan et al., 2010) and Chapter 4, 5). Cellulose is the major component of walls and is embedded in a matrix of non- cellulosic polysaccharides and proteins (Doblin et al., 2010). Therefore, the organisation of the wall cellulose, such as MFA and cellulose density, is influenced by the deposition patterns of these non-cellulosic wall components. AGPs are known to interact with pectins and non-cellulosic polysaccharides via covalent association with the AG glycans (van Hengel et al., 2001; Tan et al., 2012; Tan et al., 2013; Hijazi et al., 2014). For example, the arabinoxylan-pectin-arabinogalactan-protein1 (APAP1) is covalently bound to the wall pectins and non-cellulosic polysaccharides via its AG glycan decorations (Tan et al., 2013). FLAs are likely to be glycosylated with AG glycans (see Chapter 3 and (Johnson et al., 2003)), therefore it is possible that FLA11, FLA12 and FLA16 could form interactions with the non-cellulosic wall polysaccharides and thus modulate the wall architecture and biomechanics (Fig. 6.1D). To date, covalent interactions of AG-glycans with wall matrix components has only been shown in primary walls. Further work is needed to investigate if they can also associate with matrix polysaccharides in secondary walls.

In summary, these models provide hypotheses for further investigation of these important regulators of secondary wall development and stem biomechanics.

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Figure 6.1 Schematic representation of hypothetical mechanism(s) of action of FLA11, FLA12 and FLA16. Four hypothetical models of FLA11, FLA12 and FLA16 function. (A) FLAs potentially interact with the plasma membrane (PM) proteins responsible for cellulose biosynthesis such as cellulose synthases (CesAs), cellulose synthase complexes (CSCs), GPI-anchored proteins (GPI-AP) and/or (B) receptor-like kinases (RLKs) that act in cell wall integrity sensing pathways. (C) FLAs could interact with cellulose microfibrils either via the FAS domains or O-glycans and influence the MFA during deposition into the wall. GPI- anchored FLAs attached to microfibrils could be released into the wall either by enzymic cleavage (by phosphatidylinositol-specific phospholipase C/D (PLC/D)). (D) FLAs interact with wall matrix polysaccharides, such as pectins and non-cellulosic polysaccharides, to influence cellulose microfibril deposition. The N- and C-terminal FAS domains in FLA16 are indicated.

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Minerva Access is the Institutional Repository of The University of Melbourne

Author/s: Liu, Edgar

Title: Characterization of fasciclin-like arabinogalactan proteins in Arabidopsis thaliana

Date: 2018

Persistent Link: http://hdl.handle.net/11343/212069

File Description: Characterization of Fasciclin-like Arabinogalactan Proteins in Arabidopsis thaliana

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