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Identification and characterization of regulatory genes associated with secondary wall formation in Populus… Li, Eryang 2009

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IDENTIFICATION AND CHARACTERIZATION OF REGULATORY GENES ASSOCIATED WITH SECONDARY WALL FORMATION IN POPULUS AND ARABIDOPSIS THALIANA by  ERYANG LI  M.Sc., University of Northeast Forestry, China & Universität für Bodenkultur Vienna, Austria 2003  B.Sc., University of Northeast Forestry, China & Universität für Bodenkultur Vienna, Austria 2001   A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY  in  THE FACULTY OF GRADUATE STUDIES  (Botany)  THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)  April 2009  © Eryang Li, 2009  ii ABSTRACT  Transcript profiling has the potential to reveal transcriptional networks operating during development and provides expression data for genes of unknown function. Based on previous studies employing global transcription profiling of Arabidopsis thaliana inflorescence stem development and A. thaliana root cell type-specific expression, ten candidate transcription factor (TF) genes potentially associated with secondary wall formation and lignification were identified. Similar transcript profiling experiments had revealed gene expression patterns associated with secondary wall formation during secondary xylem formation in Populus species. I verified gene expression patterns of the poplar putative orthologs of the A. thaliana candidates, and the combination of the A. thaliana and poplar data identified a subset of conserved MYB and homeodomain TFs that behaved similarly in both plants. I analyzed the expression patterns of promoter-GUS fusions of all candidate genes in A. thaliana, most of which showed expression in xylem and cortex cells adjacent to interfascicular fibers in the stem, and in the root stele. T-DNA insertion mutants for most candidate genes were characterized, but only homeodomain transcription factor KNAT7 (At1g62990) T-DNA insertion mutants exhibited an obvious phenotype in inflorescence stems. The knat7 phenotype is characterized by irregular xylem (irx), interfascicular fibers with thicker cell walls, and defects in secreted seed mucilage. I also assayed for changes in gene expression in the knat7 background, and the results suggested that KNAT7 directly or indirectly regulates lignin biosynthesis genes. KNAT7 is known to interact with members of the Ovate Family Protein (OFP) transcription co-regulators. I confirmed the KNAT7-OFP1 and KNAT7-OFP4 interactions in planta, and showed that the interaction enhances KNAT7 transcriptional repression activity. Furthermore, an ofp4 mutant exhibits similar phenotypes as knat7, and the pleiotropic effect of OFP1 and OFP4 overexpression depends upon KNAT7 function. A knat7/ofp4 double mutant showed a very similar phenotype to the single mutants, supporting the hypothesis that the two  iii proteins work in the same pathway. KNAT7 thus appears to form a functional complex with OFP proteins, and may directly or indirectly regulate lignin biosynthesis through interaction with OFP family members. These investigations provide new insights into the regulatory network(s) governing secondary wall biosynthesis in A. thaliana and poplar.   iv TABLE OF CONTENTS  ABSTRACT ..................................................................................................ii TABLE OF CONTENTS ................................................................................iv LIST OF TABLES ........................................................................................vii LIST OF FIGURES..................................................................................... viii ABBREVIATIONS .........................................................................................x ACKNOWLEDGEMENTS ............................................................................xii COLLABORATION STATEMENT ................................................................xiv  Chapter 1. General introduction.............................................................1 1.1 Vascular development and interfascicular fiber differentiation........................ 2 1.1.1 Cell differentiation in xylem.................................................................................... 2 1.1.2 Secondary cell walls ................................................................................................ 7 1.1.3 Lignin and cellulose biosynthesis ............................................................................ 9 1.1.4 Mutants in secondary wall formation..................................................................... 10 1.1.5 Overview of transcriptional genes regulating secondary cell wall biosynthesis.... 12 1.2 Global transcript profiling of vascular development and fiber differentiation18 1.2.1 Transcriptional profiling of stem development in Arabidopsis thaliana ............... 18 1.2.2 Transcriptional profiling of vascular development in Populus .............................. 20 1.3 Transcription factor families in Arabidopsis and Populus ............................. 23 1.3.1 Comparative analysis of selected Arabidopsis and Populus TF families .............. 25 1.4 Homeodomain proteins................................................................................... 27 1.4.1 TALE homeodomain proteins in plants ................................................................. 28 1.4.2 KNOX family and functions.................................................................................. 30 1.4.3 Protein-protein interactions with BELL and OFP families .................................... 31 1.5 Research objectives ........................................................................................ 33 1.6 Background data ............................................................................................. 33 1.6.1 Global expression profiling of fiber development in Arabidopsis ......................... 34 1.6.2 Global expression profilings of secondary xylem development in poplar............. 38 1.7 Thesis outline.................................................................................................. 41   v Chapter 2. Materials and methods........................................................44 2.1 Plant materials and growth conditions............................................................ 44 2.1.1 Arabidopsis growth conditions .............................................................................. 44 2.1.2 Plant materials........................................................................................................ 45 2.1.3 Arabidopsis transformation.................................................................................... 47 2.2 Gene expression analysis................................................................................ 47 2.2.1 Semi-quantitative reverse transcriptase (RT)-PCR ................................................ 48 2.2.2 Quantitative real-time RT-PCR.............................................................................. 48 2.2.3 GUS expression assay............................................................................................ 53 2.3 Complementation analysis and subcellular localization................................. 55 2.4 Microscopy ..................................................................................................... 56 2.4.1 Bright-field microscopy ......................................................................................... 56 2.4.2 Resin embedding for bright-field and TEM........................................................... 56 2.4.3 Scanning Electron Microscopy .............................................................................. 57 2.4.4 Confocal laser scanning microscopy...................................................................... 57 2.5 Arabidopsis protoplast transfection assays..................................................... 58 2.5.1 Protoplast isolation ................................................................................................ 58 2.5.2 Protoplast transfection assays ................................................................................ 58 2.5.3 Bi-molecular fluorescence complementation analysis using protoplasts............... 60 2.6 Database searches, sequence alignments and phylogenetic analysis.............. 62  Chapter 3. Validation of candidate transcription factors identified from Populus and Arabidopsis microarray expression profiling experiments 64 3.1 Introduction .................................................................................................... 64 3.2 Results ............................................................................................................ 65 3.2.1 Confirmation Arabidopsis microarray experiment data on differential expression65 3.2.2 Organ and tissue expression profiles of candidate genes in Arabidopsis tissues ... 68 3.2.3 Cell and tissue expression patterns of candidate gene promoter-GUS fusions. ..... 73 3.2.4 Confirmation of T-DNA insertion sites in potential knock-out mutants ................ 76 3.2.5 Potential poplar orthologues of candidate genes.................................................... 78 3.2.6 Expression profiling of potential poplar orthologues of candidates in poplar arrays80 3.3 Discussion....................................................................................................... 86  Chapter 4. Arabidopsis KNAT7, a regulator of secondary wall formation 91 4.1 Introduction .................................................................................................... 91 4.2 Results ............................................................................................................ 92 4.2.1 Identity of SALK T-DNA insertion alleles ............................................................ 92  vi 4.2.2 Phenotypic characterization of knat7 mutants ....................................................... 97 4.2.3 knat7 seed coat phenotype ................................................................................... 102 4.2.4 Over-expression of KNAT7 .................................................................................. 106 4.2.5 Subcellular localization of AtKNAT7 and PoptrKNAT7 and complementation analysis 108 4.2.6 Gene expression changes in knat7 ....................................................................... 112 4.3 Discussion..................................................................................................... 115  Chapter 5. Interaction of Arabidopsis OFP proteins with KNAT7 forms a regulatory KNOX-OVATE complex ...................................................128 5.1 Introduction .................................................................................................. 128 5.2 Results .......................................................................................................... 133 5.2.1 KNAT7 interacts with OFP1 and OFP4 in vivo ................................................... 133 5.2.2 KNAT7 and OFPs are transcriptional repressors ................................................. 135 5.2.3 OFP1- and OFP4-GUS expression patterns ........................................................ 140 5.2.4 OFP4 is involved in secondary wall formation.................................................... 142 5.2.5 Ectopic expression of OFP4 causes abnormal development ............................... 146 5.2.6 Analysis of double mutants.................................................................................. 148 5.3 Discussion..................................................................................................... 151  Chapter 6. Conclusions and future directions ...................................156 References..............................................................................................166    vii LIST OF TABLES  Table 1.1 Transcription factors involved in regulation of secondary wall biosynthesis and assembly in Arabidopsis thaliana....................................................................... 14 Table 1.2 Predicted number of genes in selected TF families in the Arabidopsis and Populus genomes. ..................................................................................................... 24 Table 2.1 List of mutant alleles and oligonucleotides used in study................................ 46 Table 2.2 Primer sequences used for real-time PCR analysis and semi-quantitative RT-PCR....................................................................................................................... 51 Table 2.3 Primer sequences used to amplify Arabidopsis promoter sequences for GUS fusions. ........................................................................................................................ 54 Table 2.4 Primer sequences used to generate GFP fusion constructs. ............................. 55 Table 2.5 Primer sequences to generate YFP fusions.......................................................... 61 Table 3.1 Summary of histochemical assays of GUS activity in transgenic lines expressing candidate gene promoter-GUS fusions. ........................................... 75 Table 3.2 Summary of T-DNA insertion lines and mutants used for phenotypic analyses. 77 Table 3.3 Proposed poplar homologues of Arabidopsis transcription factors associated with fiber and secondary wall development and differential expression in two microarray profiling experiments.......................................................................... 82 Table 4.1 Secondary cell wall thickness in wild type and knat7-1 stems ..................... 101 Table 5.1 A summary of interaction partners of KNAT7 identified from Hackbusch et al. (2005). ....................................................................................................................... 132   viii LIST OF FIGURES  Figure 1.1 Organization of the primary and secondary vascular tissues in Arabidopsis shown schematically................................................................................................ 5 Figure 1.2 Xylem cell differentiation. ...................................................................................... 6 Figure 1.3 Schematic diagrams of plant cell wall structure................................................. 8 Figure 1.4 Model of the transcriptional network proposed to regulate secondary wall biosynthesis................................................................................................................ 17 Figure 1.5 General features of KNOX and BLH homeodomain proteins. ..................... 29 Figure 1.6 Primary stem development in Arabidopsis. ...................................................... 36 Figure 1.7 Hierarchical clustering of expression profiles of 10 transcription factors. . 37 Figure 1.8 Experimental designs for poplar microarray expression profiling conducted in parallel with my thesis research............................................................................. 40 Figure 3.1 Validation of candidate transcription factor gene expression over the course of Arabidopsis inflorescence stem development. ................................................... 67 Figure 3.2 In silico analysis of relative expression levels of ten candidate transcription factor genes in Arabidopsis. ................................................................................... 71 Figure 3.3 Experimental analysis of expression of ten Arabidopsis candidate transcription factor genes in different organs. ............................................................................ 72 Figure 3.4 Histochemical localization of GUS activity in transgenic Arabidopsis lines expressing promoter-GUS fusions. ....................................................................... 74 Figure 3.5 Phylogenetic reconstructions of selected transcription factor gene families in Arabidopsis and poplar. ........................................................................................... 79 Figure 3.6 Analysis of differential expression of selected Populus transcription factor genes using quantitative real time RT-PCR......................................................... 83 Figure 3.7 Public microarray data on relative expression levels of poplar orthologs of Arabidopsis candidate genes in different tissue. ................................................ 85 Figure 4.1 Characterization of the knat7-1 allele................................................................. 94 Figure 4.2 Additional KNAT7 T-DNA insertion alleles and their mutant phenotypes in stems............................................................................................................................ 95 Figure 4.3 Histochemical analysis of GUS activity of promoterKNAT7-GUS fusions.99 Figure 4.4 Anatomy at the bases of wild type and knat7-1 inflorescence stems......... 100 Figure 4.5 Transmission electron microscopy of wild-type and knat7 inflorescence stem fiber and vessel secondary walls. ........................................................................ 101 Figure 4.6 knat7 seed coat mucilage phenotype ................................................................ 104 Figure 4.7 Analysis of wild type and knat7 mutant seed coat epidermal cells. ........... 105 Figure 4.9 Complementation of the Aknat7 phenotype with 35S::GFP-AtKNAT7 and subcellular localization of GFP-AtKNAT7. ...................................................... 109 Figure 4.10 Complementation of knat7 by PoptrKNAT7 and subcellular localization of PoptrKNAT7-GFP................................................................................................... 111  ix Figure 4.11 Quantitative real-time RT-PCR analysis of gene expression in knat7 seedlings relative to wild type seedlings............................................................ 114 Figure 4.12 In silico analysis of relative expression levels of KNAT7 in Arabidopsis.121 Figure 4.13 Proposed model of KNAT7 roles in the transcriptional network regulating secondary wall biosynthesis. ................................................................................ 125 Figure 5.1 Phylogenetic analysis of the Arabidopsis thaliana ovate family protein (OFP) gene family............................................................................................................... 131 Figure 5.2 Bimolecular fluorescence complementation analysis of OFP1, OFP4 and KNAT7 interactions in vivo. ................................................................................. 134 Figure 5.3 OFP4 and KNAT7 are transcriptional repressors that interact in vivo....... 137 Figure 5.4 KNAT7 transcriptional repression domains. ................................................... 139 Figure 5.5 Histochemical assay of OFP4-GUS and OFP1-GUS expression............... 141 Figure 5.6 Analysis of OFP1 and OFP4 insertion mutants............................................. 144 Figure 5.7 Phenotypic characterizations of loss-of-function mutants of OFP1, OFP4 and ofp1/ofp4 double mutant. ...................................................................................... 145 Figure 5.8 Phenotypes of transgenic plants overexpressing 35S:OFP4 in Arabidopsis.147 Figure 5.9 Anatomical characterization of knat7/ofp1 and knat7/ofp4 double mutants.149 Figure 5.10 Phenotypic characterization of 35S:OFP1/knat7, and 35S:OFP4/knat7 plants. .................................................................................................................................... 150 Figure 6.1 A proposed model of the function of KNOX-OVATE complex in the SND1-mediated pathway. ..................................................................................... 163    x ABBREVIATIONS  ABRC Arabidopsis Biological Resource Center AFLP Amplified fragment length polymorphism AP2/EREBP APETALA2/ETHYLENE RESPONSE ELEMENT BINDING PROTEIN BAC Bacteria artificial chromosome BAR Bio-Array Resource for Arabidopsis Functional Genomics BiFC Bi-molecular fluorescence complementation BLAST Basic Local Alignment Search Tool BLH BEL1-like homeodomain (protein/gene) BP BREVIPEDICELLUS bp  DNA base pairs CaMV35S Cauliflower mosaic virus 35S (constitutive) promoter CesA Cellulose synthase cDNA Complementary DNA reverse transcribed from messenger RNA (mRNA) ChIP Chromatin immunoprecipitation CYFP/NYFP C terminus / N terminus of yellow fluorescent protein DIC Differential interference contrast DNA Deoxyribonucleic acid EST Expressed sequence tag EYFP Enhanced yellow fluorescent protein GA Gibberellin GD Gal4 DNA binding domain GFP Green fluorescent protein GR Glucocorticoid receptor GTs Glycosyltransferases GUS β-glucuronidase hr Hours IFF Interfascicular fiber IFL1 INTERFASCICULAR FIBERLESS 1 irx Irregular xylem kb kilo bases (1000 base pairs) KNAT Knotted 1-like (protein/gene) of Arabidopsis thaliana KNOX Knotted 1-like homeobox (protein/gene) LB Luria-Bertani bacterial growth medium forumulation MEIS Myeloid ectropic viral integration site mRNA Messenger RNA MUG 4-methylumbelliferyl-β-D-glucuronide NPA N-(1-naphthyl) phthalamic acid NST1 NAC secondary wall-thickening-promoting factor 1  xi OFP OVATE FAMILY PROTEIN ORF Open reading frame PCD Programmed cell death PCR Polymerase chain reaction qRT-PCR Quantitative real-time RT-PCR RNA Ribonucleic acid RNAi RNA mediated interference RT-PCR Reverse-transcriptase PCR SAM Shoot apical meristem SD Standard deviation SEM Scanning electron microscopy SND1 Secondary wall-associated NAC domain protein 1 STM SHOOTMERISTEMLESS TAIR The Arabidopsis Information Resource; http://Arabidopsis.org/ TALE Three amino acid loop extension TEM Transmission electron microscopy TFs Transcription factors TW Tension wood UV Ultra violet VND VASCULAR-RELATED NAC-DOMAIN XETs Xyloglucan endotransglycosylases X-gluc 5-bromo-4-chloro-3-indolyl-β-D-glucuronide XTH xyloglucan endotransglucosylase hydrolases YFP Yellow fluorescent protein   xii ACKNOWLEDGEMENTS  I would like to express my gratitude to all those who helped and inspired me during my doctoral study and gave me the possibility to complete this thesis. I owe sincerest thanks to Prof. Carl J. Douglas, my supervisor for the last five years, for his guidance and encouragement. His incredible patience, energy, and passion for science are inspiring my pursuit of graduate studies. I would like to deeply thank Prof. Jingbo Jia, my supervisor in Northeast Forestry University, Harbin, China, who has been my mentor since I was an undergrad. He inspired me to the science world and recommended me to go to Vienna in 2000 because of “Asia-Europe Forestry Exchange Program”. I should also thank my master supervisor Prof. Marie-Theres Hauser at Universität für Bodenkultur Vienna, Austria for her supervision during October 2000 till March 2003 in the Center of Applied Genetics. Her energy and enthusiasm in research had initially motivated me on plant genetics and encouraged me to carry on scientific career. Drs. Xin Li, Brian Ellis, and Geoffrey Wasteneys deserve special thanks as my thesis committee members and advisors for their encouragement and suggestions on my project and critical comments on my thesis. In particular, I would like to thank Dr. Ellis for his NSERC Green Crops network (GCN) funding support during my study. The continuous funding helped me to complete my thesis. I am very thankful to Drs. George Haughn, Fred Sack, Lacey Samuels, Jin-Gui Chen, and Shawn D. Mansfield for their practical advice and suggestions for my project. Great thanks goes to all of my current and former colleagues in Department of Botany, especially those in Douglas lab for valuable advice and a pleasant environment to work. Michael Friedmann, Juergen Ehlting, Bjoern Hamberger, Clarice Souza, Bahram Soltani, June Kim, David Johnston Monje, Lee Johnson, Apurva Bhargava, Sung Soo Kim, Yuanyuan Liu, and Teagen Quilichini had inspired me in research and life through our interactions during the long hours in the lab. Thanks. My great gratitude to the various colleagues in the field who gave me clones, advice, seeds,  xiii and shared unpublished data and technical experiences. These are addressed in the collaboration statement for their contributions. Most of images were obtained in UBC bioimaging facility and Fred Sack Lab. Specially I would thank to Garnet Martens and Dr. Jie Le for their technical assistant and valuable advice for the microscopy work, to Dr. Shucai Wang to share his experience, data and provide OFPs materials, and to Sarah Mckim to provide YFP vectors for BiFC analysis. I thank to staff and faculty of Department of Botany, UBC. Thanks to the awards of UBC university of Graduate Fellowship (UGF) given by the Faculty of Graduate Studies and Frances Chave Memorial Scholarship given by Department of Botany. Special thanks to my dear friends in UBC Songhua Zhu, Jian Guo, Ye Wang, Jim Guo, Hardy Hall, Qingning Zeng, Miki Fujita, Minako Kaneda, Michael Friedmann, Jun Huang, Yunkun Dang, Jun chen and Echo Yu, they are the source of happiness and wisdom, I am grateful to their friendship and good times during my study in UBC. Family friends Mr.&Mrs Yan, Mr.&Mrs Liu and their family have offered love and warmest kindness to me which support my life in Vancouver and make me feel like family. Especially, my deepest gratitude goes to my family for their unflagging love and support throughout my life, this thesis is impossible without them. My perfect parents, Jing-Ran Li and Min Yang have been constantly supportive and understanding in every stages of my life without any complaints. Thanks largely to the everlasting love from my family. My most heartfelt thanks go to my husband Haitao Sun for his constant dedication. This study is the result of his understanding, encouragement, and love.   xiv COLLABORATION STATEMENT Chapter 1 Preliminary data from the Douglas Lab and the Treenomix I project from transcriptional profiling microarray were generated by Ehlting et al. (2005), Dr. Lee Johnson Ph.D. thesis (2006), Mrs. Margaret Ellis and Dr. Michael Friedmann (unpublished data). The results of this work are summarized in a manuscript in preparation: Eryang Li, Michael Friedmann, Lee A. Johnson, Margaret Ellis, Rick White, Steven Ralph, Jun Zhang, Natascha Forneris, Lindsey Rawding, Weiya Qiang, Rodney A. Savidge, Brian E. Ellis, and Carl J. Douglas Identification of common regulatory genes associated with secondary wall formation in poplar (Populus sp.) secondary xylem and Arabidopsis inflorescence stems. (First authorship is equally contributed).  Chapter 3 Mrs. Margaret Ellis contributed the phylogenetic reconstructions of selected transcription factor gene families in Arabidopsis and poplar. A former visiting scholar, Dr. Weiya Qiang from Lanzhou University in China, and former post-doc fellow Dr. Jun Chen contributed to preliminary work on recombinant DNA constructs, GUS assays and phenotypic observations of T-DNA insertion mutants in candidate genes. Part of my data on verification of Arabidopsis candidate transcription factor gene expression from this chapter was published in: Ehlting, J., Mattheus, N., Aeschliman, D.S., Li, E. Hamberger, B., Cullis, I.F., Zhuang, J., Kaneda, M., Mansfield, S.D., Samuels, L., Ritland, K., Ellis, B.E., Bohlmann, J., and Douglas, C.J. (2005). Global transcript profiling of primary stems from Arabidopsis thaliana identifies candidate genes for missing links in lignin biosynthesis and transcriptional regulators of fiber differentiation. Plant J 42, 618-640.  Chapter 4 I prepared fixed stem and root material from knat7 stems and PromKNAT7:GUS plants, and processing and sectioning were done in the UBC BioImaging Facility by Garnet Martens. Dr. Jun Chen contributed with D35S and 4CL promoter Gateway destination vectors used for overexpressing KNAT7.  Chapter 5 Drs. Shucai Wang and Jin-Gui Chen provided Arabidopsis seeds of the following genotypes: ofp1, ofp4, ofp1/ofp4, PromOFP1:GUS, and PromOFP4:GUS. Dr. Wang was involved in designing the transient protoplast expression experiments, helped me carry out and interpret the experiments, and provided images of OFP protein family tree and ofp1, ofp4, ofp1/ofp4 mutant morphology at rosette leaves stage. The collaboration work was summarized in the following manuscript in preparation: Eryang Li, Shucai Wang, Jin-Gui Chen, and Carl J. Douglas A KNOX-OFP transcription factor complex regulates secondary wall formation (in preparation).  1       CHAPTER 1  General introduction  2 Chapter 1. General introduction  1.1 Vascular development and interfascicular fiber differentiation Wood (secondary xylem), fiber formation and vascular differentiation are important plant-specific processes that are critical to the life histories of both woody perennial plants such as trees, and shorter-lived herbaceous plants, such as Arabidopsis thaliana (hereafter Arabidopsis). Vascular tissues include phloem and xylem. The secondary xylem or wood provides mechanical strength and allows long-distance transport of water and nutrients, which enables shoots of woody plants to grow in height and diameter.  1.1.1 Cell differentiation in xylem The vascular system is composed of two basic tissues, xylem and phloem. Xylem is composed of conducting tracheary elements and nonconducting elements such as xylary parenchyma cells and xylary fibers. Vascular tissues can be formed from two different meristematic tissues, the procambium and vascular cambium. During primary growth of stems and roots, procambial initials derived from apical meristems produce primary xylem and primary phloem. Vascular cambium initials, which originate from procambium and other parenchyma cells when plants undergo secondary growth, give rise to secondary xylem, commonly called wood, and secondary phloem (Evert, 2006). Both primary and secondary xylem are water and nutrient conducting tissues in plants (Figure 1.1). Xylem tracheary elements (vessels, tracheids) are made up of lifeless tubes with thick secondary cell walls. The function of xylem requires strong and lignified secondary cell walls in the tracheary elements. In addition, the shape of cell walls in the primary xylem is distinctive. Primary xylem develops unique annular, spiral or reticular cell wall thickenings in tracheary elements (McCann, 1997). Therefore, the primary cell wall is reinforced at some, but not all, of its surfaces. In contrast, secondary xylem has heavy secondary cell walls throughout the tracheary elements and fiber cells. Cells of the secondary xylem in angiosperm and gymnosperms each have unique modifications to their walls. Vessels, large tracheary elements characteristic of angiosperms, have perforated end walls with simple pits. Around them, there are fibers, also lifeless at maturity, which give  3 mechanical support, and xylem parenchyma cells, some of which are long-lived and provide metabolic support (Figure 1.1) (Chaffey et al., 2002). Xylem cells with secondary cell wall thickenings undergo common developmental steps: differentiation begins with cell expansion, followed by secondary cell wall deposition of polysaccharides and lignin, and finally programmed cell death (PCD, Figure 1.2) (Fukuda, 1996; Roberts and McCann, 2000).  Cell expansion The first step in differentiation is cell expansion. During primary xylem development, procambium cells derived from the apical meristem, start to expand longitudinally, differentiating into tracheary elements. In secondary xylem, different cell types have distinct expansion patterns, in which fibers elongate, doubling in length from cambium initials by intrusive growth (Lev-Yadun, 1997). In contrast, radial expansion dominates in the vessel elements of the secondary xylem where cell length does not change but width is greatly increased. During the expansion phase, the cell has to loosen the existing wall matrix and synthesize new wall material. Expansins and xyloglucan endotransglycosylases (XETs) have been identified as cell wall-loosening proteins in Arabidopsis (Im et al., 2000; Cosgrove, 2001; Bourquin et al., 2002). Expansins disrupt hydrogen bonds between xyloglucan chains and cellulose microfibrils, thereby loosening microfibrils that could then slide past each other (Cosgrove, 2001). XETs, also called xyloglucan endotransglucosylase hydrolases (XTH), can cleave and rejoin xyloglucan chains, a process which is important in control of cell wall expansion (Bourquin et al., 2002; Vissenberg et al., 2005).  Secondary wall thickening At the end of the cell expansion phase, the protoplast begins to produce a thickened secondary cell wall, a three-layered structure (S1, S2, S3) made of cellulose microfibrils (Donaldson, 2001) (Figure 1.3C). In Arabidopsis, a generally accepted model of cellulose synthesis is that a cellulose synthase complex (rosette) composed of CESA (cellulose synthase) subunits and other proteins is the functional unit, producing cellulose microfibrils (crystalline arrays of cellulose polymers) outside of the plasma membrane. The pattern of  4 secondary wall thickening is regulated through deposition of cellulose microfibrils in controlled orientations, a process that is apparently regulated by the patterns of cortical microtubules located underneath the plasma membrane. During primary cell wall deposition, microtubule disorganization or complete depolymerization does not alter the orientation of cellulose microfibrils, but growth anisotropy is lost (Wasteneys and Fujita, 2006) (Figure 1.3B). Microfilaments also appear to be important for the normal patterning of secondary walls in tracheary elements (Kobayashi et al., 1988). A recent study has shown the relationship between the microtubules and cellulose synthase complex during secondary wall formation (Wightman and Turner, 2008). They observed that the cellulose synthase complexes form bands beneath sites of secondary wall synthesis, and the maintenance of these sites is dependent upon underlying bundles of microtubules, which localize the cellulose synthase complex to the edges of developing cell wall thickening (Wightman and Turner, 2008) Subsequent to cellulose deposition, lignin accumulates in secondary cell walls of tracheary elements in xylem. Initiation of lignification starts in the middle lamella then is deposited throughout the primary and secondary cell wall layers (Donaldson, 2001).  Programmed cell death The final stage of xylem differentiation in cells such as vessel elements, fibers, and tracheids, is autodigestion of their living protoplast due to programmed cell death (PCD), resulting all cell contents are degraded (Fukuda, 1996). Cell death during xylogenesis has been extensively studied in the Zinnia in vitro tracheary element differentiation system, where the major events of PCD are vacuole swelling followed by tonoplast rupture and rapid nuclear degradation (Fukuda, 1996).     5     Figure 1.1 Organization of the primary and secondary vascular tissues in Arabidopsis shown schematically.  (A) Whole plant, longitudinal view, (B) Shoot apex, cross section, (C) Leaf cross section, (D) Root tip cross section, (E) Organization of vascular tissues in the basal region of the inflorescence stem during the secondary phase of vascular development. Inside the secondary xylem, the position of layers associated with cell expansion, cell wall deposition and cell death has been indicated. The image was adapted from Nieminen et al. (2004); used with permission.  6      Figure 1.2 Xylem cell differentiation.  The steps of differentiation are cell expansion, secondary cell wall deposition, followed by lignification and programmed cell death (PCD). The image was modified from Samuels et al. (2006); used with permission.  7  1.1.2 Secondary cell walls Plant cell walls are characterized as being either primary or secondary. Primary walls are found in growing plant cells and contribute to controlling cell shape (Figure 1.3A). In contrast, secondary cell walls are laid down once the cell has stopped growing, and contribute to many of the structural properties associated with plant cells. Secondary walls are the major components of tracheary elements and fibers in the plants. Typically, they are composed of cellulose (~50%), lignin (~20%) and hemicelluloses (~30%) (Balatinecz et al., 2001), including xylans and glucomannans (Figure 1.3B).  8       Figure 1.3 Schematic diagrams of plant cell wall structure.  (A) A primary cell wall containing cellulose microfibrils, hemicellulose, pectin, and soluble proteins. (B) Diagram of a secondary cell wall relative to the primary wall and plasma membrane, which contains cellulose synthase enzymes as integral membrane proteins. (C) The secondary cell wall S1, S2 and S3 layers are based on different microfibril arrangment. Images were adapted from Sticklen (2008); used with permission.  9  1.1.3 Lignin and cellulose biosynthesis  Lignin biosynthesis Lignin impregnated in the cellulose and hemicellulose network provides additional mechanical strength to the secondary wall and also renders the secondary wall waterproof owing to lignin’s hydrophobic nature. Lignin is a polymer of phenylpropanoid monolignols that cross-link together. These are synthesized through the phenylpropanoid pathway and exported into the secondary cell wall. Lignin is composed of three types of units, p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) units, derived from three monolignols: p-coumaryl alcohol, coniferyl alcohol and sinapyl alcohol, respectively (Higuchi, 1985). H and S units are deposited at the initial stage of lignification within middle lamellae, and later in the secondary walls of xylem cells (Terashima et al., 1993). G and S are the major units found in the lignin of angiosperms, but gymnosperm lignin is devoid of S units. Our understanding of the monolignol biosynthetic pathway has undergone major revisions over the past decade, as a result of in vitro kinetic studies on lignin biosynthetic enzymes and genetic studies of mutants and transgenic plants with altered expression levels of phenylpropanoid pathway genes (Humphreys and Chapple, 2002; Boerjan et al., 2003). Ten enzymes are required for H, G, and S monolignol biosynthesis starting with the amino acid phenylalanine: phenylalanine ammonia lyase (PAL), cinnamic acid 4-hydroxylase (C4H), 4-(hydroxy)cinnamoyl CoA ligase (4CL), hydroxycinnamoyl CoA:shikimate hydroxycinnamoyl transferase (HCT), p-coumaroylshikimate 3'-hydroxylase (C3H), caffeoyl CoA O-methyltransferase (CCoAOMT), (hydroxy)cinnamoyl CoA reductase (CCR), ferulic acid 5-hydroxylase (F5H), caffeic acid/5-hydroxyferulic acid O-methyltransferase (COMT) and (hydroxy)cinnamyl alcohol dehydrogenase (CAD). In the cell wall, dehydrogenative polymerization of the monolignols is thought to be catalyzed by peroxidases (PER) and laccases (LAC) (Boerjan et al., 2003); however, the exact roles of these polymerization enzymes in the regulation of lignin biosynthesis in planta have not been defined.  Cellulose synthesis Cellulose, a simple polymer of unbranched β-1,4-linked glucan chains, is a central  10 component of plant cell walls, and is especially abundant in the secondary cell wall. Hexagonal complexes on the plasma membrane, known as rosettes, are cellulose synthase complexes and consist ofβ-glucosyltransferase (cellulose synthase) subunits encoded by CESA (cellulose synthase) genes (Doblin et al., 2002). The Arabidopsis genome contains a complement of 10 CESA genes (Richmond, 2000), and analysis of the Populus trichocarpa genome reveals the presence of 18 CESA genes (Taylor, 2008). Three CESA genes (CESA1, CESA3 and CESA6) are required for primary cell wall formation. Two recent studies demonstrate that CESA2, CESA5 and CESA9 are partially redundant with CESA6 at different stages of growth (Desprez et al., 2007; Persson et al., 2007). Therefore, this has led to the idea that CESA1, CESA3 and a CESA6-related protein are all essential for cellulose synthesis in primary cell walls.  1.1.4 Mutants in secondary wall formation Arabidopsis inflorescence stems develop a substantial amount of xylem and interfascicular fibers, making it a good system in which to study secondary wall synthesis. Several Arabidopsis mutants affecting secondary wall formation have been isolated, including the irx (irregular xylem) mutations that dramatically reduce secondary wall thickening of vessel elements in the xylem. This reduction in secondary wall thickening is due to a decrease in cellulose deposition (Turner and Somerville, 1997; Brown et al., 2005). The Arabidopsis irx1, irx3 and irx5 mutants are caused by lesions in the CESA8, CESA7 and CESA4 genes, respectively (Taylor et al., 1999; Taylor et al., 2000; Taylor et al., 2003). CESA7 and CESA8 were also named as FRAGILEFIBER5 (fra5) and FRAGILEFIBER6 (fra6) mutants, which were isolated by screening mutants for interfascicular fibers of reduced strength (Zhong et al., 2003). Due to the similarity of many aspects of secondary cell wall deposition in both xylem and interfascicular fibers, it is likely that some regulatory genes, such as those encoding proteins involved in signal transduction or transcriptional regulation, will be common to both cell types (Turner and Hall, 2000). A gapped xylem (gpx) mutant of Arabidopsis was identified as lacking proper co-ordination of secondary cell wall deposition in both xylem and interfascicular cells, resulting in some cells with very thick cell walls and some that  11 apparently undergo little or no secondary cell wall deposition (Turner and Hall, 2000). Xylan, composed of a (1,4)-linked β -D-xylopyranose backbone substituted with α-glucuronic acid (GlcUA) or 4-O-methyl-α-D-glucuronic acid (4-O-Me-GlcUA), is a major hemicellulosic constituent and the second most abundant polysaccharide in dicot wood. Galactoglucomannans are major components of the cell walls of the woody tissues of both angiosperms and gymnosperms, and also occur in fern and moss cell walls (reviewed in (Popper, 2008). Although genes participating in the biosynthesis of cellulose and lignin have been well characterized, much less is known about genes involved in hemicellulose biosynthesis (Ye et al., 2006; York and O'Neill, 2008). A number of glycosyltransferases (GTs) associated with xylan biosynthesis have now been identified in Arabidopsis and poplar, suggesting that many of these GTs are likely to be conserved and involved in polysaccharide synthesis during wood formation (Aspeborg et al., 2005; Ye et al., 2006; Brown et al., 2007; Persson et al., 2007). They include members of the GT8 (PARVUS [At1g19300], IRX8 [At5g54690]), GT43 (IRX9 [At2g37090] and IRX14 [At4g36890]), and GT47 (FRA8/IRX7 [At2g28110], IRX10 [At1g27440] and IRX10-like [At5g61840]) families (Ye et al., 2006; Brown et al., 2007; Lee et al., 2007; Persson et al., 2007; Brown et al., 2008; Wu et al., 2008; York and O'Neill, 2008). Mutants in each of these genes show decreased xylan in stems, and an increase in the proportion of 4-O-Me-GlcUA side branches relative to the non-methylated GlcUA (Brown et al., 2007). Further analysis of the xylan reducing end structure and xylan chain length indicates that FRA8/IRX7, IRX8 and PARVUS are involved in the synthesis of the reducing end structure, whereas IRX9, IRX10, IRX10-like and IRX14 may function in xylan backbone chain elongation (Bauer et al., 2006; Brown et al., 2007; Lee et al., 2007; Wu et al., 2008). Alternatively, it has been suggested FRA8 may add GlcUA onto the xylan backbone (Zhong et al., 2005). In order to fully understand cell wall biosynthesis, it is essential to identify all of the proteins that are responsible for xylan synthesis, and further elucidate both the function and interaction between the different components of the xylan biosynthetic machinery.   12 1.1.5 Overview of transcriptional genes regulating secondary cell wall biosynthesis Due to the importance of secondary cell walls, plants must have evolved mechanisms to turn on biosynthetic pathways for synthesis of cell wall components in various cell types in the right amounts for assembly in the right spatial-temporal patterns, according to their functions in specific cell types (Zhong and Ye, 2007). The identification of transcription factors that regulate overall secondary cell wall synthesis is a major step in this direction.  Secondary cell wall transcriptional regulation network There have been important recent discoveries in transcriptional regulation of secondary wall biosynthesis (Table 1.1). Several NAC, MYB, homeodomain, and AP2/ERF proteins have been shown to be key players in regulating secondary cell wall biosynthesis (Table 1.1). XND1 (xylem NAC domain 1) has been shown to affect tracheary element growth through regulation of secondary wall synthesis and programmed cell death and it is highly expressed in xylem (Zhao et al., 2008a). Arabidopsis ERF38 (ETHYLENE RESPONSE FACTOR 38) was found to be co-expressed with several genes involved in secondary wall thickening and is a potential regulator in secondary wall metabolism (Lasserre et al., 2008). SND1 (secondary wall-associated NAC domain protein 1), acting redundantly with NST1 (NAC secondary wall-thickening-promoting factor 1), is specifically expressed in interfascicular and xylem fibers. Expression of dominant negative version of SND1 in transgenic plant drastically reduces the secondary wall thickening in fibers (Zhong et al., 2006). NST1 and NST2 are redundant in regulating secondary cell wall formation in the endothecium cell layer of anthers, where a secondary wall is required so that rupture can occur to release pollen grains (Mitsuda et al., 2005). A MYB transcription factor, MYB26, is also essential for secondary wall thickening in the endothecium (Yang et al., 2007). It has been shown that MYB46 is a direct target of SND1, and dominant repression of MYB46 resulted in a lack of secondary cell wall thickening in fiber and xylem vessels in stems (Zhong et al., 2007a). A recent study investigated several SND1-regulated transcription factors. Of these, dominant repression of SND2, SND3, MYB103, MYB85, MYB52, MYB54, and KNAT7 (Knotted 1-like protein of Arabidopsis thaliana 7) significantly reduced secondary wall thickening in fiber cells (Zhong et al., 2008). In  13 addition, this study revealed that SND3, MYB103, and KNAT7, together with MYB46, are direct targets of SND1 and of its close homologs NST1, NST2, and of the vessel-specific VND6 (Vascular-related NAC domain 6) and VND7 proteins (Zhong et al., 2008). These data all suggest that all three major pathways for secondary cell wall synthesis (cellulose, lignin and xylan) are coordinately regulated at the transcriptional level. It is likely, however, that there is a cascade of transcription factors regulating secondary cell wall synthesis, and that further dissection of these networks may identify transcription factors involved in activation of the individual pathways for the three major secondary cell wall polymers, including cellulose. Zhong et al. (2008) proposed a network that regulates secondary wall synthesis (Figure 1.4). In this network model SND1 and its homologs act as master switches at the top of a hierarchy that regulates secondary wall biosynthesis in different cell types. According to this model, SND1 and NST1 activate the secondary wall biosynthetic program in fibers; VND6 and VND7 specifically regulate secondary wall biosynthesis in vessels; and NST1 and NST2 act together in regulating secondary wall biosynthesis in the endothecium of anthers. SND1 exerts its functions through activating several direct target transcription factor genes as mentioned above. The other SND1-regulated transcription factors appear not to be direct targets of SND1 and are likely positioned further downstream in the transcriptional network (Figure 1.4). SND2 is able to induce the expression of cellulose biosynthetic genes and MYB85 can induce the expression of lignin biosynthetic genes. In addition, MYB52 and MYB54 appear to slightly induce the expression of genes in all three major secondary wall biosynthetic pathways. Determining the positions of other transcription factors in the SND1-mediated transcriptional cascade will contribute to our understanding of how the expression of secondary wall biosynthetic genes is coordinately regulated. The models discussed above provide a possible framework for understanding the secondary cell wall transcription factor regulatory network, but when we consider that a few hundred genes are probably involved in the biosynthesis of secondary walls in plants, we are still far from fully understanding the transcriptional network and how regulation of multiple target genes in a spatially and temporally defined manner is achieved.   14  Table 1.1 Transcription factors involved in regulation of secondary wall biosynthesis and assembly in Arabidopsis thaliana Gene Gene product Loss-of function or dominant repression phenotype Tissue/cell expression patterns Reference SND1 /NST3/ At1g32770 NAC domain protein Repression of secondary wall thickening in fibers; SND1/NST3 and NST1 function redundantly in the activation of secondary wall thickening in fibers Interfascicular fibers, xylary fibers (Zhong et al., 2006; Mitsuda et al., 2007; Zhong et al., 2007b) SND2 /At4g28500 NAC domain protein Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers Developing protoxylem and metaxylem, elongating interfascicular fibers, interfascicular fibers, secondary xylem in root (Zhong et al., 2008) SND3 /At1g28470 NAC domain protein Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers Developing protoxylem and metaxylem, elongating interfascicular fibers, interfascicular fibers, secondary xylem in root (Zhong et al., 2008) NST1 /At2g46770 NAC domain protein Repression of secondary wall thickening in endothecium Anthers, xylem, interfascicular fibers, other organs (Mitsuda et al., 2005) NST2 /At3g61910 NAC domain protein Repression of secondary wall thickening in endothecium; NST1 and NST2 function redundantly in the activation of secondary wall thickening in endothecium Anothers, other organs (Mitsuda et al., 2005) VND6 /At5g62380 NAC domain protein Repression of secondary wall thickening in metaxylem Metaxylem in primary roots (Kubo et al., 2005) VND7 /At1g71930 NAC domain protein Repression of secondary wall thickening in protoxylem Protoxylem in primary roots (Kubo et al., 2005) XND1 /At5g64530 NAC domain protein Affects tracheary element growth through regulation of secondary wall synthesis and Xylem (Zhao et al., 2008a)  15 Gene Gene product Loss-of function or dominant repression phenotype Tissue/cell expression patterns Reference programmed cell death MYB26 /At3g13890 MYB transcription factor Loss of secondary wall thickening in endothecium Anthers (Steiner-La nge et al., 2003; Yang et al., 2007) MYB46 /At5g12870 MYB transcription factor Repression of secondary wall thickening in fibers and vessels Interfascicular fibers, xylary fibers, vessel (Zhong et al., 2007a) MYB52 /At1g17950 MYB transcription factor Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers Developing protoxylem and metaxylem, elongating interfascicular fibers, interfascicular fibers, secondary xylem in root (Zhong et al., 2008) MYB54 /At1g73410 MYB transcription factor Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers Developing protoxylem and metaxylem, elongating interfascicular fibers, interfascicular fibers, secondary xylem in root (Zhong et al., 2008) MYB69 /At4g33450 MYB transcription factor Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers Developing protoxylem and metaxylem, interfascicular fibers, secondary xylem in root (Zhong et al., 2008) MYB85 /At4g22680 MYB transcription factor Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers, deformation of vessels Developing protoxylem and metaxylem, interfascicular fibers, secondary xylem in root (Zhong et al., 2008) MYB103 /At1g63910 MYB transcription factor Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers Developing protoxylem and metaxylem, interfascicular fibers, secondary xylem in (Zhang et al., 2007; Zhong et al., 2008)  16 Gene Gene product Loss-of function or dominant repression phenotype Tissue/cell expression patterns Reference root IFL1 (INTERFAS CICULAR FIBERLESS 1) /At5g60690 Homeodomain leucine zipper protein Reduced secondary xylem differentiation in fibers Interfascicular fibers and vascular region (Zhong and Ye, 1999) KNAT7 /At1g62990 Homeodomain protein Reduction in secondary wall thickening in both interfascicular fibers and xylary fibers (Dominant repression) Developing protoxylem and metaxylem, interfascicular fibers, secondary xylem in root (in situ hybridization (Brown et al., 2005; Zhong et al., 2008) ERF38 /At2g35700 AP2/ERF type transcription factor No mutant phenotype, co-regulated with several secondary wall thickening genes, a candidate regulator of secondary wall metabolism Seed coat outer integument, cells between phloem and cortex in stems, endodermal cells of young roots (Lasserre et al., 2008)   17       Figure 1.4 Model of the transcriptional network proposed to regulate secondary wall biosynthesis.  Adapted from Zhong et al. (2008); used with permission.  18 1.2 Global transcript profiling of vascular development and fiber differentiation Global transcript profiling across developmental gradients using cDNA or oligonucleotide-based microarrays has emerged as a powerful approach to reveal genome-wide developmental gene expression patterns in several organisms. Here I provide an overview of transcriptional profiling studies related to xylem and fiber differentiation and maturation in Arabidopsis, Populus species (hereafter Populus or poplar), and Zinnia sp.  1.2.1 Transcriptional profiling of stem development in Arabidopsis thaliana To classify the expression profiles of individual genes in terms of their preferential expression site along the developing Arabidopsis inflorescence stem, Ehlting (2005) and Imoto (2005) used tissue sections representing certain developmental stages along a gradient of maturation. They performed whole genome microarray analysis to study the transcriptional events associated with interfascicular fiber and xylem development. These studies identified various families involved in phenylpropanoid biosynthesis, secondary cell wall deposition and transcriptional regulation (Ehlting et al., 2005; Imoto et al., 2005). Demura et al. (2002) employed expression profiling of an in vitro Zinnia cell culture in which leaf mesophyll cells trans-differentiate into tracheary elements to identify genes whose expression is closely associated with morphological events during secondary cell wall biosynthesis. Another expression profiling experiment of xylem cell-differentiation-related genes in Arabidopsis used an in vitro xylem vessel element-inducible system in Arabidopsis suspension culture cells (Kubo et al., 2005). These authors identified a variety of proteins closely associated with secondary cell wall formation (cellulose synthases, xylanases, and laccases), programmed cell death (nucleases and proteases) and transcription factor candidates for regulating these processes (Demura et al., 2002; Kubo et al., 2005). Arabidopsis has been shown to undergo secondary vascular growth induced either when it is kept from flowering by repeated removal of inflorescences (Zhao et al., 2000; Oh et al., 2003) or by applying a weight load on the stem (Ko et al., 2004; Yokoyama and Nishitani, 2006), and can produce secondary xylem in the inflorescence stems (Oh et al., 2003) and in  19 the root-hypocotyl junction (Chaffey et al., 2002). Using this system, Oh et al., (2003) utilized microarrays to profile genes with preferred expression in Arabidopsis secondary xylem, and during the transition from primary to secondary growth in Arabidopsis stems. This led to the identification of candidate transcription factors and other regulatory genes that may be involved in xylem development. Ko et al. (2004) demonstrated that the application of artificial weight induces the production of significant amounts of secondary xylem tissues in A. thaliana inflorescence stems and identified 700 genes that were differentially expressed during the transition from primary growth to secondary growth (Ko et al., 2004). These results suggest that the weight carried by aerial plant parts may generate a signal for the formation of xylem secondary cell walls with greater mechanical strength. It was demonstrated that the primary signal from the applied weight not only induces cambium differentiation, but also perturbs auxin levels, a potential down-stream regulator of this process (Ko et al., 2004). A recent study used the auxin transport inhibitor N-(1-naphthyl) phthalamic acid (NPA) to induce vascular overgrowth and study global changes in gene expression patterns associated with differentiation of cambial tissues, xylem, phloem and fibers (Wenzel et al., 2008). Microarray analysis revealed a large number of genes whose expression was up-regulated in shoots developing vascular overgrowth and identified 40 genes having vascular-related expression patterns in Arabidopsis included cambial tissues and differentiating xylem, phloem and fibers (Wenzel et al., 2008). An analysis combining fluorescent-activated cell sorting of green fluorescent protein (GFP)-marked cell populations with microarray analysis described gene expression profiles for five tissue types in three developmental zones in the Arabidopsis root, including the stele, where xylem and phloem differentiate (Birnbaum et al., 2003). These profiles revealed a greater transcriptional complexity than profiles of the whole root organ alone, and generated information on gene sets expressed in cell-type and development specific contexts. Recently, this experimental approach was extended at higher resolution by the Benfey group (Brady et al., 2007) to reveal a complex transcriptional network that underlies root spatio-temporal development. This study provided evidence for fluctuating gene expression over developmental time and considerable expression variation between individual roots (Brady  20 et al., 2007). These cell type-specific profiles could contribute important insights into the network of transcriptional interactions that regulate Arabidopsis root development. The large number of Arabidopsis gene expression analyses available in Affymetrix GeneChip databases, allow in silico-correlated gene expression studies to be carried out. Two examples of the use of such existing microarray data as a guide for discovery of genes involved in secondary wall formation and cellulose biosynthesis are from Brown et al. (2005) and Persson et al. (2005). These authors identified genes co-expressed with secondary wall-specific CESA genes as a guide to identify candidate genes involved in secondary wall formation. A similar approach was taken to identify candidate genes involved in secondary xylem formation in Arabidopsis (Ko et al., 2006). Arabidopsis offers an outstanding model system to study the molecular mechanisms in secondary growth. However, limitations still exist for this model system. First, while the anatomy of secondary xylem in Arabidopsis closely resembles the wood of a poplar tree (Chaffey et al., 2002), Arabidopsis stems do not have the ray parenchyma cells that are present in poplar. Another limitation is the lack of a perennial growth of Arabidopsis stem (Oh et al., 2003). Therefore, important aspects of secondary growth, such as the seasonal cycle of cambial activity cannot be studied in Arabidopsis, but are better addressed in tree species such as Populus, for which excellent genomics resources exist (Jansson and Douglas, 2007).  1.2.2 Transcriptional profiling of vascular development in Populus Poplar has been increasingly used as a model tree for understanding wood formation and other tree-specific processes with a large number of EST (expressed sequence tag) sequences (Dejardin et al., 2004; Sterky et al., 2004) a complete genome sequence (Tuskan et al., 2006), and other genomic tools (Jansson and Douglas, 2007). The relatively large region of developing secondary xylem in poplar wood can be divided into different developmental zones: meristematic cells, early expansion, late expansion, secondary wall formation and programmed cell death (Hertzberg et al., 2001). Hence, poplar wood offers a useful system for discovering genes involved in various developmental stages of xylem differentiation. The first reported poplar transcript profiling experiment to search for genes involved in  21 wood formation (Hertzberg et al., 2001) assessed the expression profile of 2,995 ESTs in wood-forming tissues. This experiment confirmed that known lignin and cell wall-related genes were up-regulated, but also found interesting upregulation of previously unstudied transcription factors. Tangential cryosectioning of developing poplar wood combined with use of a high-density poplar cDNA microarrays has allowed high-resolution global gene expression profiling in developing poplar secondary xylem. Aspeborg et al. (2005) performed microarray analysis using RNAs isolated from different xylem zones and found that 25 genes encoding glycosyltransferases (GTs) are highly expressed in the secondary wall formation zone of poplar xylem and play potential roles in hemicellulose synthesis during wood formation (reviewed in (Ye et al., 2006)). Using this approach, Schrader et al (2004) generated a high-resolution transcriptional map covering the cambial region of developing aspen (Populus tremula) secondary xylem. This provided sets of marker genes for different stages of xylem and phloem differentiation and identified potential regulators of cambial meristem activity. A significant recent advance was the use of a microgenomic analysis to reveal cell type-specific gene expression patterns within cambial meristem of Populus (Goue et al., 2008). In this study, microdissection and RNA amplification were carried out to document the differences in the transcriptomes of neighboring meristematic cells between ray cambial cells, which give rise to rays and fusiform cambial cells, which give rise to the axial cell system (i.e. fibers and vessel elements). Notably, genes involved in pectin metabolism and xyloglucan metabolism were overrepresented in ray cambial cells and fusiform cambial cells, respectively. In addition, many cell wall-related genes showed cell type-specific expression patterns. This approach of microgenomics revealed the differences in biological processes in neighbouring meristematic cells, and identified key genes involved in these processes (Goue et al., 2008). A study aimed at understanding the regulation of cell differentiation in Populus apical and vascular cambium meristems analyzed the poplar KNOX homeobox gene, ARBORKNOX1 (ARK1), which is orthologue to Arabidopsis SHOOT MERISTEMLESS (STM). STM is expressed in the shoot apical meristem (SAM) and the vascular cambium, and is down-regulated in the terminally differentiated cells of leaves and secondary vascular tissues  22 that are derived from these meristems (Groover et al., 2006b). Microarray analysis of ARK1-overexpressing poplar plants showed a set of genes up-regulated in the stems, including proteins involved in cell identity and signaling, cell adhesion, or cell differentiation. The results revealed that the SAM and vascular cambium are regulated by overlapping genetic programs (Groover et al., 2006b). Young poplar trees exhibit a transition from primary growth at the apex, characterized by a individual bundles, to secondary growth characterized by secondary xylem and lignified woody tissue within a few internodes below the apex (Savidge, 2000). This is the result of formation and activation of the vascular cambium. Two cDNA-AFLP (amplified fragment length polymorphism) -based transcriptome analyses along this developmental transition were carried out in hybrid aspen and identified genes expressed during primary growth (top), transition from primary growth to secondary growth, and secondary growth (bottom) (Prassinos et al., 2005; van Raemdonck et al., 2005). Moreover, two poplar AP2/ERF-like transcription factors were confirmed to be expressed specifically in either phloem or ray, which suggested a potential role in vascular tissue development (van Raemdonck et al., 2005). Temperate woody perennials, including trees, have adapted to freezing temperatures and limited water during winter by alternating between active growth and vegetative dormancy, which is a complex developmental process controlled by interactions between environmental and internal factors (Lang et al. 1987, Dennis 1994). To gain insights into the underlying molecular mechanisms, Park et al. (2008) carried out a global transcriptional profiling of xylem and bark from field-grown Populus deltoides trees at several times over the annual growth cycle. They compared these expression patterns with profiles of trees grown under various controlled environmental conditions and identified gene sets that are differentially regulated during the growth cycle, providing novel insights into the seasonal regulation of woody plants (Park et al., 2008). Stems and branches of angiosperm trees form tension wood (TW) when exposed to a gravitational stimulus, in which the normal development of secondary wall biosynthesis is greatly modified (Mellerowicz et al., 2001; Hellgren et al., 2004; Pilate et al., 2004). One of the main characteristics is the formation of fibers with a thick inner gelatinous cell wall layer  23 (G-layer) mainly composed of crystalline cellulose. Hence TW cell walls are enriched in cellulose, and deficient in lignin and hemicelluloses. Data from transcript profiling using microarray and metabolite analysis were obtained during TW formation in Populus tremula (Andersson-Gunneras et al., 2006). They found that expression of secondary cell wall associated CESA genes (orthologs of Arabidopsis CESA8, CESA7 and CESA4) was differently affected in TW, suggesting that different CESA isoforms might be present in the biosynthetic rosettes of TW compared with normal wood. In addition, the data suggested decreased activity in the lignin biosynthesis pathway, and in biosynthesis of cell wall matrix carbohydrates. Several differentially expressed transcription factors were also identified, eg. the homeodomain transcription factor, KNAT7 (Andersson-Gunneras et al., 2006). Following the extensive synthesis of the secondary cell walls, the final phase in maturation of both vessel elements and fibers is cell death and autolysis of the cell contents. Moreau et al (2005) studied gene expression during the process of fiber death by in silico analysis of a “fiber death library” and by microarray analysis using a Populus 25K cDNA microarray. They identified several novel candidate regulatory genes for the biological process of xylem programmed cell death, including two novel extracellular serine proteases, nodulin-like proteins and an Arabidopsis thaliana OPEN STOMATA 1 (AtOST1) homolog in signaling fiber-cell death, as well as the Populus homolog of Arabidopsis XYLEM CYSTEINE PEPTIDASE 2 thought to be responsible for protein degradation and remobilization of nutrients in the dying fibers (Moreau et al., 2005). These experiments have demonstrated the potential for transcriptome profiling as a strategy for gene discovery in poplar trees, specifically in the field of vascular development, and confirm that transcriptome-level profiling is an extremely powerful tool for identifying previously unknown genes involved in developmental processes in this newly emerged model system (Jansson and Douglas, 2007).  1.3 Transcription factor families in Arabidopsis and Populus “Transcription factors” is a term usually used to describe proteins that recognize DNA in a sequence-specific manner as either activator or repressor, and that regulate the frequency of  24 initiation of transcription upon binding to specific sites in the promoter of target genes (Pabo and Sauer, 1992). Transcription factors (TFs) play important roles in plant cellular and developmental processes by controlling gene expression. With the completion of the Arabidopsis (The Arabidopsis Genome Initiative, 2000) and Populus trichocarpa (Tuskan et al., 2006) genome sequences, it became possible not only to study the function of TFs on a genome-wide scale but also to compare the structural and functional similarities and differences between the TF families in these two species.   Table 1.2 Predicted number of genes in selected TF families in the Arabidopsis and Populus genomes.   TF family Arabidopsis TAIRa Populus DPTFb MYBs 133 216 MADS 109 111 (1051) AP2/EREBP 138 212 (2002) ARF 24 37 (393) AUX/IAA 293 33 (353) a Numbers of TFs reported in TAIR Arabidopsis gene family information (http://www.arabidopsis.org/browse/genefamily/index.jsp). b Numbers of TFs predicted in DPTF, a database of poplar transcription factors (http://dptf.cbi.pku.edu.cn/index.php) (Zhu et al., 2007). 1 Reference from (Leseberg et al., 2006) 2 Reference from (Zhuang et al., 2008) 3 Reference from (Kalluri et al., 2007)  25 1.3.1 Comparative analysis of selected Arabidopsis and Populus TF families The analysis of the Arabidopsis genome sequence indicates that it codes for at least 1,572 transcription factors, which account for ~6% of its estimated ~26,000 genes (Riechmann et al., 2000; The Arabidopsis Genome Initiative, 2000). Their systematic functional characterization can be pursued with a variety of reverse genetic methods (Riechmann and Ratcliffe, 2000). On the basis of similarities in the DNA-binding domains, which are commonly defined by 50 to 70 amino acids with multiple residues being highly conserved, transcription factors have been classified into families (Pabo and Sauer, 1992). The Arabidopsis genome contains several large TF families that include more than 100 members, such as MYB, MADS, basic helix-loop-helix (bHLH), APETALA2 (AP2)/ETHYLENE RESPONSE ELEMENT BINDING PROTEIN (EREBP) and Auxin Response Factors (ARFs) (Guilfoyle et al., 1998; Lee and Schiefelbein, 1999; Riechmann and Ratcliffe, 2000; Bailey et al., 2003; Kirik et al., 2005; Guilfoyle and Hagen, 2007). The poplar genome is the first to be sequenced among woody plants (Tuskan et al., 2006), and is quite similar in size to the rice genome and about four times larger than that of Arabidopsis (Brunner et al., 2004). As a result of computational prediction and manual curation, a total of 2576 putative transcription factors were identified and classified into 64 families in Populus and organized in a public database of poplar TFs (DBTF) (Zhu et al., 2007). Some MYB transcription factors were functionally characterized in hybrid aspen (Karpinska et al., 2004). Comprehensive genome-wide analyses of some TF families identified in Populus trichocarpa, have been carried out, including MADS-box, AUX/IAA, ARF and AP2/ERF gene families (Leseberg et al., 2006; Kalluri et al., 2007; Zhuang et al., 2008). Two selected TF gene families are discussed below, and homeodomain TFs are discussed in more detail in section 1.4.  MYB superfamily The MYB superfamily is one of the largest TF families known in plants, consisting of three families, the R2R3, R1R2R3 and MYB-related families, on the basis of the number and position of the MYB repeats. Members of the MYB family have been found to be involved in diverse processes, including developmental control and determination of cell fate and  26 identity (Oppenheimer et al., 1991; Lee and Schiefelbein, 1999; Wang et al., 2008), plant responses to environmental factors and hormones (Jin and Martin, 1999), signal transduction in plant growth processes (Gubler et al., 1995; Iturriaga et al., 1996; Shin et al., 2007), abiotic stress (Magaraggia et al., 1997; Hoeren et al., 1998; Jung et al., 2008), pathogen defence (Yang and Klessig, 1996) and regulation of phenylpropanoid metabolism and lignin biosynthesis (Bender and Fink, 1998; Borevitz et al., 2000; Jin et al., 2000; Patzlaff et al., 2003b; Patzlaff et al., 2003a; Newman et al., 2004; Goicoechea et al., 2005; Dubos et al., 2008). However, few studies have addressed the role of MYB transcription factors in transcriptional regulation of secondary growth in Populus. In a study of MYB genes in hybrid aspen (Populus tremula L.X tremuloides Michx.), three MYB genes differentially expressed in developing secondary xylem were identified, suggested potential roles in the regulation of secondary vascular tissue development (Karpinska et al., 2004).  AP2/EREBP family The Arabidopsis genome contains 138 members of the AP2/EREBP family, which were classified into five groups—the AP-2 subfamily, the RAV subfamily, the DREB subfamily, the ERF subfamily, and others (Sakuma et al., 2002). All 145 AP2/EREBP TF genes have been successfully amplified and cloned from various types of Arabidopsis tissues or organs or after different treatments, opening the door to comprehensive functional analysis (Feng et al., 2005). Many gene family members are involved in responses to different stresses: drought, freezing and pathogen invasion (Gutterson and Reuber, 2004). In poplar, a number of studies identified AP2/EREBP genes. Examination of a full-length enriched cDNA library from leaves of Populus nigra var. italica subjected to environmental stress treatments yielded thirteen candidates containing the AP2/ERF domain that showed stress-responsive gene expression (Nanjo et al., 2004). Two other DREB-like genes from Populus were isolated using the yeast one-hybrid system (Han et al., 2007). Four paralogs of Arabidopsis C-repeat binding factor (CBF or DREB1) subfamily genes, PtCBFs, are cold inducible in leaves and increase freezing tolerance in Populus (Benedict et al., 2006). Two ERF-related transcription factors were expressed during the transition from growth to  27 dormancy in apical and axillary buds (Rohde et al., 2007). Genome-wide analysis revealed that the total number of AP2/ERF genes in the DREB, and ERF subfamilies in Populus trichocarpa is about 1.4 fold higher than in Arabidopsis (Zhuang et al., 2008) (Table 1.2). This ratio is very similar to the overall ratio of protein-coding genes in Populus relative to Arabidopsis; Populus has an average of 1.4-1.6 putative homologs for each Arabidopsis gene (Tuskan et al., 2006).  These and other gene families remain largely uncharacterized in forest tree species. The genome-wide investigation of similarities and differences in such transcriptional regulators between woody and herbaceous plants (e.g. poplar versus Arabidopsis) should open the way to better understanding of the functions of unknown TFs and their conservation in plants.  1.4 Homeodomain proteins Homeodomain TFs are characterized by a highly conserved 60-amino acid motif DNA-binding domain called the homeodomain (Gehring et al., 1994a).  The 180-bp DNA sequence that codes for this domain is called a homeobox (Gehring et al., 1994a). The homeodomain folds into a characteristic three-helix structure. Helices I and II are connected by a loop, while helices II and III are separated by a turn, resembling prokaryotic helix-turn-helix TFs. However, unlike helix-turn-helix-containing proteins, most homeodomain proteins are able to bind DNA with high affinity as monomers, through interactions made by helix III (the so-called recognition helix) and a disordered N-terminal arm located beyond helix I (Wolberger et al., 1991; Gehring et al., 1994b). Amino acid sequence comparison studies have subdivided the homeodomain proteins into two classes of proteins based on the presence of extra amino acids within the homeodomain. The typical homeodomain class has 60 amino acids, while the TALE (Three amino acid loop extension) class has an extra three amino acids (Pro-Tyr-Pro) inserted between helices I and II (Bertolino et al., 1995; Burglin, 1997). These three residues form part of a hydrophobic loop that is thought to be involved in protein-protein interactions between TALE and other homeobox proteins.   28 1.4.1 TALE homeodomain proteins in plants Five TALE protein classes (PBC, Myeloid ecotropic viral integration site [MEIS], Iroquois, TG-interacting factor and Mohawk) have been identified in metazoans based on the conservation of the homeodomain and associated domains, but only two classes exist in plants (KNOTTED1-like homeobox [KNOX] and BEL1-like homeobox [BLH]) (Hake et al., 2004a; Mukherjee and Burglin, 2007). The KNOX and BLH proteins share significant sequence similarities in the homeodomain region but differ greatly in the N terminal regions of the proteins (Figure 1.5). All plant KNOX proteins have an ELK domain and a KNOX (MEINOX) domain (Burglin, 1997). The ELK domain consists of 24 amino acids and has been postulated to be involved in nuclear localization, protein-protein interactions or suppression of gene activation (Meisel and Lam, 1996; Nagasaki et al., 2001). The MEINOX domain name is an acronym derived from animal MEIS and plant KNOX proteins and both MEIS and KNOX domains are conserved upstream of homeodomains (Burglin, 1997; Mukherjee and Burglin, 2007). The plant MEINOX domain consists of two smaller domains, KNOX1 and KNOX2, separated by a poorly conserved linker sequence. Two-hybrid assays showed that the MEINOX domain is necessary and sufficient for interacting with BELL proteins (Bellaoui et al., 2001; Muller et al., 2001; Smith et al., 2002; Bhatt et al., 2004; Kumar et al., 2007). A strong functional relationship exists between the homeodomain and the MEINOX domain, but a recent observation indicates that the plant MEINOX domain can also function in a homeodomain-independent fashion: the Arabidopsis KNATM gene contains a MEINOX domain, but lacks a homeodomain (Magnani and Hake, 2008). Perhaps related to this finding is the earlier observation that the human TALE protein MEIS2 has an alternate splice form that does not include the homeodomain, and this form acts as a dominant-negative competitor with homeodomain proteins (Yang et al., 2000). The BLH proteins have a conserved SKY domain and a BELL domain upstream of their homeodomain (Burglin, 1997; Bellaoui et al., 2001). The VSLTLGL box is a conserved sequence of unknown function present in some of the BLH proteins, but absent from others (Chen et al., 2003). The conserved SKY and BELL domains of BLH proteins and the conserved MEINOX domain of KNOX proteins are required for the heterodimerization of  29 the BLH and KNOX proteins (Bellaoui et al., 2001; Muller et al., 2001) (Figure 1.5). TALE homeodomain proteins play key roles in developmental programs in plants, animals, and fungi (Burglin, 1997; Mukherjee and Burglin, 2007). In plants, KNOX proteins have been found in dicots, monocots, gymnosperms and algae and mosses (Reiser et al., 2000; Hake et al., 2004b; Sakakibara et al., 2008).         Figure 1.5 General features of KNOX and BLH homeodomain proteins.    30 1.4.2 KNOX family and functions In plants, the first homeobox gene identified was the maize Knotted1 (KN1) gene (Vollbrecht et al., 1991). Dominant mutations in KN1, which is normally active only in meristematic cells, affect leaf development due to its aberrant expression in these organs (Smith et al., 1992). Knotted-like homeobox (KNOX) genes have been isolated from maize and other monocot and dicot species reviewed in Chan et al. (1998). Liu et al. (2008) indicating that this class of genes constitutes a family throughout the angiosperms. In Arabidopsis, the KNOX family of eight genes can be subdivided into two phylogenetic classes, I and II, based on sequence similarity and expression patterns (Kerstetter et al., 1994). STM, BREVIPEDICELLUS (BP)/KNAT1, KNAT2, and KNAT6 belong to class I and are characteristically expressed in the shoot apical meristem (SAM), while class II KNAT3, KNAT4, KNAT5, and KNAT7 are more broadly expressed (Hake et al., 2004a). Some of the class I genes have been shown to play roles in proper development of the SAM (Long et al., 1996; Douglas et al., 2002; Belles-Boix et al., 2006). SHOOT MERISTEMLESS (STM) is required for the initiation of the meristem during embryogenesis and its maintenance during postembryonic development (Long et al., 1996). In Arabidopsis, bp mutant is reduced in size because of irregularly shortened internodes and their pedicels point down (Douglas et al., 2002; Venglat et al., 2002). Like BP, KNAT6 contributes redundantly with STM to SAM function, as the inactivation of KNAT6 abolishes the meristematic potential of the weak allele stm-2 (Belles-Boix et al., 2006). In addition, a specific role for KNAT6 in boundary maintenance is indicated by a severe defect in cotyledon separation observed in the stm-2 knat6 double mutant. KNAT2, the gene most closely related to KNAT6, is expressed in the SAM, but a knock-out mutation in KNAT2 gene failed to produce a notable phenotype (Belles-Boix et al., 2006; Ragni et al., 2008). The functions of the class II KNOX genes are less well characterized than those in class I, but based on cell type specific expression patterns in Arabidopsis roots, KNAT3, KNAT4, KNAT5 are proposed to play roles in Arabidopsis root development (Truernit et al., 2006). KNAT7 (IRX11) was identified as a transcriptional regulator involved in secondary wall formation (Brown et al., 2005; Persson et al., 2005; Zhong et al., 2008). Besides class I and II of the KNOX subfamilies, a recent study identified and characterized  31 KNATM, a novel Arabidopsis KNOX gene that encodes a MEINOX domain but lacks the homeodomain, and defines KNATM as a new KNOX family subgroup that is conserved in eudicots (Magnani and Hake, 2008). KNATM is expressed in the proximal-lateral regions of organ primordia and the leaf hydathode, and plants overexpressing KNATM show multiple leaf pattern defects, including longer petioles, curled-down leaves, and shorter leaf lamina, as well as delayed bolting. In addition, yeast two-hybrid assays show that KNATM interacts with other plant BELL TALE proteins through its MEINOX domain and also with BP through an acidic coiled-coil domain, raising the possibility that KNATM might regulate transcription through interaction with a DNA-binding protein. (Magnani and Hake, 2008). Populus KNOX family genes have not been well studied. However, the Populus homeobox gene ARBORKNOX1 (PoptrARK1), which is an ortholog of Arabidopsis STM, is expressed in the shoot apical meristem (SAM) and in the vascular cambium in Populus (Groover et al., 2006a).  1.4.3 Protein-protein interactions with BELL and OFP families It is well documented that KNOX and BLH proteins interact in many plant species (Bellaoui et al., 2001; Muller et al., 2001; Smith et al., 2002; Chen et al., 2003). KNOX-BLH interactions form heterodimers that are thought to constitute functional complexes that regulate plant development (Bellaoui et al., 2001; Muller et al., 2001). In Arabidopsis, there are 13 BLH (BEL1-like homeodomain proteins) genes (Smith et al., 2004), and the functions of six of them have been characterized. BEL1, the founding member of the BLH family, is involved in the development of ovule integuments (Reiser et al., 1995). Mutations in a BLH gene PENNYWISE (PNY) (also known as BELLRINGER, REPLUMLESS or VAAMANA) disturb the Arabidopsis spiral phyllotactic pattern, while double mutations in PNY and its close paralog POUND-FOOLISH (PNF) arrest the transition from vegetative to reproductive phase (Byrne et al., 2003; Roeder et al., 2003; Smith and Hake, 2003; Bhatt et al., 2004; Smith et al., 2004). The BLH protein ARABIDOPSIS THALIANA HOMEOBOX1 (ATH1) also controls floral competency by transcriptionally activating the flower repressor gene FLOWERING LOCUS C (Proveniers et al., 2007). Two BLH genes, SAWTOOTH1 (SAW1) and SAW2, negatively regulate the  32 KNOX gene, BP, and a saw1 saw2 double mutant has increased leaf serration (Kumar et al., 2007). In yeast two-hybrid experiments, it has been demonstrated that interactions between the Arabidopsis proteins display some specificity since BEL1 interacts with STM, KNAT1 KNAT2 and the class II gene KNAT5, but not with KNAT3, KNAT4 and KNAT7 (Bellaoui et al., 2001). The BLH protein PNY interacts specifically with class I KNOX proteins STM, BP/KNAT1, and KNAT6 and is important for inflorescence stem growth and meristem function in Arabidopsis (Bhatt et al., 2004). PNY interaction with BP was also shown to be required for regulation of inflorescence patterning and fruit development (Smith and Hake, 2003; Alonso-Cantabrana et al., 2007). Members of the KNOX family also interact with the previously uncharacterized family of Arabidopsis OVATE proteins (Hackbusch et al., 2005). The OVATE gene was originally was found to encode a protein with a putative nuclear localization signal and an ≈70-aa C-terminal domain that is conserved in tomato, Arabidopsis, and rice (Liu et al., 2002). A single mutation leading to a premature stop codon in the tomato OVATE gene causes the normally round fluit to become pear-shaped (Liu et al., 2002). There are 18 genes in the Arabidopsis genome that encode proteins with a conserved C-terminal OVATE domain, and most members of this OVATE FAMILY PROTEIN (OFP) family contain a predicted nuclear localization signal (Hackbusch et al., 2005; Wang et al., 2007). However, little is known about the functions of OFP proteins in plants, and their molecular mechanisms of action are unknown. So far, only the Arabidopsis OFP protein OFP1 has been well characterized. OFP1 functions as an active transcriptional repressor that has a role in regulating cell elongation by controlling the expression of GA20ox1, a gene that encodes a key enzyme in gibberellin biosynthesis (Wang et al., 2007). OFP1 overexpression leads to pleiotropic phenotypes in Arabidopsis, including severe dwarfism and defective organ morphogenesis (Wang et al., 2007). When the yeast two-hybrid system was used to systematically analyze the interaction networks among Arabidopsis TALE homeodomain proteins, this revealed a densely connected network of interactions between and within the two TALE families (Hackbusch et al., 2005). According to this analysis, every member of the KNOX family has at least one  33 interaction partner amongst the BELL proteins. In addition, homo- and heterodimerization of several proteins within each TALE family, including dimerization of BELL proteins not previously described (eg. BLH9/KNAT3 and BLH9/KNAT5, was demonstrated in Hackbusch et al (2005)). Significantly, the network also involved nine OFP members (Hackbusch et al., 2005). OFP1, OFP2, and OFP4 interacted with four BELL and two KNOX proteins (BLH1, BLH3, BLH4, BLH10, KNAT5, and KNAT6), while OFP5 shows the ability to interact with a number of BLH proteins, but does not interact with any of the KNOX proteins. Certain KNOX proteins, including KNAT7, are capable of interaction with numerous OFPs in this assay system. These data provide the outlines of a possible TALE protein interaction network and provide evidence for a close functional link between TALE proteins and the OFP family. Such interactions could reflect specific roles for these complexes in the regulation of plant development.  1.5 Research objectives The primary goal of this research was to functionally characterize candidate transcription factors that were identified in a previous study as potential regulators of interfascicular fiber and/or secondary wall development in Arabidopsis (Ehlting et al., 2005). One objective was to use reverse genetics to study the functions of ten Arabidopsis candidate TFs. A second objective was to identify poplar homologs and potential orthologs of the Arabidopsis candidate TFs, and to determine whether expression of the poplar orthologs was associated with secondary xylem development and secondary wall formation in poplar. The latter work took advantage of the release of the poplar genome sequence, and cDNA microarray gene expression profiling experiments carried out by the Treenomix I project and Douglas lab, while I focused primarily on most of the functional analyses of the potential candidate gene(s) that appeared most likely to be involved in a conserved transcriptional network regulating secondary wall formation.  1.6 Background data Here I provide background data from microarray transcription profiling experiments  34 carried out in Arabidopsis (Ehlting et al., 2005) and poplar (unpublished) by the Treenomix I project and Douglas Lab. These data provided the foundation for my research.  1.6.1 Global expression profiling of fiber development in Arabidopsis Ehlting et al. (2005) designed a near full-genome Arabidopsis spotted 70-mer oligo array and used this to profile global changes in gene expression along the developmental gradient of stem maturation, interfascicular fiber differentiation and secondary wall formation in the Arabidopsis inflorescence stem. In order to characterize different stages of vascular and interfascicular fiber differentiation along the axis of bolting stems in Arabidopsis, the following samples were collected by Ehlting et al. (2005) as shown in Figure 1.6. a) 0-2 cm from the apex of 5 cm stems (developing vascular bundles, no interfascicular fibers), b) 0-3 cm from the apex of 10 cm stems (vascular bundle differentiation almost completed, no interfascicular fibers), c) 2-4 cm from the apex of 5 cm stems (late stage of vascular bundle differentiation, early stage of interfascicular fiber differentiation), d) 3-5 cm from the apex of 10 cm stems (early to middle stages of interfascicular fiber differentiation), e) 5-7 cm from the apex of 10 cm stems (middle to late stages of fiber differentiation), and f) 7-9 cm from the apex of 10 cm stems (fiber differentiation almost completed). In this experiment, about 5000 genes were found to be differentially expressed over the course of stem development, including 270 transcription factors (Ehlting et al., 2005). In order to filter the transcription factor dataset and enrich it for candidate transcription factor genes most likely to be involved in fiber or vascular differentiation, Ehlting et al. (2005) compared the data to those from an expression profiling experiment that focused on differential gene expression in various tissues, cell types, and developmental stages of the Arabidopsis root, including the stele (Birnbaum et al., 2003). Among transcription factors found in up-regulated gene expression clusters in the stem study, nine were also expressed in a stele-specific manner, and were up-regulated during the course of stele development Birnbaum et al. (2003). This group of candidate fiber development transcription factors, indicated in Figure 1.7, has members from six different families: three MYB (MYB20, At1g66230; MYB43, At5g16600; and MYB63, At1g79180) two bHLH (bHLH68, At4g29100 and bHLH144, At1g29950), one bZIP (bZIP9, At5g24800), one  35 AP2-EREBP (At5g0758), one C3H (At5g42200), and one homeodomain protein (KNAT7, At1g62990). One other transcription factor gene (bZIP47, TGA1, At5g65210) that was found in an up-expression cluster but was not stele-specific was also included in the candidate list. These 10 transcription factors were the candidates for regulators of vascular development, fiber differentiation and secondary wall formation. At the time, no functional information was available for any of these 10 genes, and verification and characterization of the candidates was the subject of my research.  36   Figure 1.6 Primary stem development in Arabidopsis.  Nine-week old Arabidopsis Ler plants were harvested when primary stems reached a total length of 4-6 cm (A) or 9-11 cm (B). Stems were divided into different segments as indicated in lays (marker unit is cm). Serial hand sections of the primary stem were performed at different distances from the tip and samples were analyzed using bright field (C, E, G, and I) and UV-fluorescence (D, F, H, and J) light microscopy. From 10 cm stems, sections were analyzed at 2 cm (C and D), 4 cm (E to H), 6 cm (G and H), and at 8 cm from the tip (I and J). All micrographs were taken at a magnification of 200x. The figure is adapted from Ehlting et al. (2005), and original images have no scale bar, but with magnification.  37     Figure 1.7 Hierarchical clustering of expression profiles of 10 transcription factors.  Ten candidate genes were differentially expressed as shown in the heatmap. Mean expression ratios [log2 (sample / 0-2 cm sample)] of these genes were used for hierarchical cluster analysis with average linkage. Expression ratios are shown as a color coded heatmap with more than 2.5 fold higher expression indicated in red, and more than 2.5 fold lower expression in a sample compared to the 0-2 cm reference indicated in blue. Locus identifier and gene names are given to the right of the heatmap. The figure is adapted from data in the Ehlting et al. (2005) experiment.  38 1.6.2 Global expression profilings of secondary xylem development in poplar A 15, 496-element poplar cDNA microarray (Treenomix 15.5K array; (Ralph et al., 2006) was employed in two experiments to identify poplar genes whose expression was 1) differentially regulated during the transition from primary to secondary growth and during the onset of secondary xylem development at the poplar shoot apex and 2) differentially regulated in cells at different stages of development from the cambial zone to cells in the maturing secondary xylem in wood taken from mature field-grown trees (Figure 1.8, Douglas lab unpublished data). For the first experiment, the samples were collected from four internodes at the apices of 4-month old poplar (P. trichocarpa X P. deltoides, clone H11) saplings, calibrated to the internode subtended by the first fully expanded leaf that was designated internode “T1”. Internodes above were numbered sequentially from T2 to T5, which was reproducibly the youngest macroscopic node that could be isolated. Hand sections from multiple ramets were taken from the centers of nodes T2 to T5 and examined under a microscope with UV excitation to detect autofluorescence associated with lignification. Rings of extensively lignified secondary xylem were apparent in internode T1 and older nodes, and the second internode above the fully expanded leaf (T2, Figure 1.8A) was the first internode in which secondary xylem was clearly obvious. Sections from internodes T4 and T5, nearest the shoot apex, showed only individual vascular bundles without a vascular cambium or secondary xylem (Figure 1.8), while in section T3 an incipient vascular cambium was observed, but little if any secondary xylem was obvious. Thus, the onset of the developmental change from primary to secondary growth occurred mainly between internodes T3 and T4. In the second experiment, the samples were from developing xylem after spring re-activation of cambial zone activity in field-grown Populus balsamifera trees (R. Savidge, personal communication). In these trees, renewed progression of xylem cell differentiation from immediate cambial derivatives to radially expanding fibers and vessel elements, through to fibers and vessels with lignified secondary cell walls was observed (Figure 1.8B). By sampling developing xylem from trees at different times after cambial reactivation, secondary xylem tissues enriched for cambial derivatives at different stages of development could be isolated. The Samples included: G, recent cambial derivatives with primary walls  39 only; O, cambial derivatives with active radial expansion of primary walls; M, vessels with secondary cell walls developed in vessels but fibers without secondary walls and no obvious lignification; CC, secondary cell walls formed in fibers, with vessels fully lignified, primary walled radially expanded cambial derivatives also present (Figure 1.8B). In both experiments, the samples represent different stages in the progression of secondary xylem development, either from the youngest (T1) to oldest (T5) internode examined, or from the earliest (G) to latest (CC) stages of secondary xylem development in mature trees. Consequently, a large proportion of the elements on the array showed differential expression between one or more tissue types. An objective of my research was to identify the poplar orthologs of candidate Arabidopsis transcription factor genes identified by Ehlting et al. (2005), and to further investigate those genes whose expression was up-regulated over the progression of secondary xylem and fiber development in both experiments. I hypothesized that the subset of genes whose expression is associated with secondary wall formation in both species would be potentially the most important in the regulation of secondary xylem and secondary wall development.  40  Figure 1.8 Experimental designs for poplar microarray expression profiling conducted in parallel with my thesis research.  (A) Progression of vascular system development in Populus trichocarpa X P. deltoides hybrid H11 saplings. Left, the most apical five internodes from a sapling are shown, with the arrow indicating the first fully expanded leaf. The internode-naming scheme is depicted. Right, hand sections from the youngest internodes, T5-T2, are shown under UV light illumination to reveal secondary cell wall autofluorescence. A ring of vascular cambium is visible in node T3, but not T4. Figure adapted from Lee Johnson, PhD Thesis, UBC (Johnson, 2006). (B) Progression of secondary xylem formation tissues isolated from Populus balsamifera after release from winter dormancy. Top, cross sections of secondary xylem from dormant and re-activated cambial regions. Low, depiction of approximate stages in secondary xylem development of samples used for expression profiling (G), early cambial derivatives with beginnings of primary wall expansion; (O), cambial derivatives in the process of primary wall radial expansion; (M), secondary wall formation but no detectable lignification; (CC), secondary wall formation well advanced with lignification of vessel elements. (From unpublished data in Douglas lab).  41 1.7 Thesis outline  The work in my thesis is divided into three parts.  Functional analysis of Arabidopsis TF candidate genes (Chapter 3) Expression patterns of the ten candidate transcription factor genes that were identified by Ehlting et al. (2005) were confirmed using quantitative real-time RT-PCR with the same samples used in the microarray. To test the expression of these genes in different Arabidopsis organs, RT-PCR, promoter-GUS, and in silico expression assays were used. Phenotypic evaluation of T-DNA insertion mutants (from the SALK, SAIL or other collections) of these genes for changes in morphology and growth and inflorescence stem anatomy and development relative to wild type were carried out. By using data from phylogenetic analysis of poplar homologues of the candidates, and data from poplar cDNA microarrays, the orthologs of these genes in poplar were identified and expression patterns determined using quantitative real-time RT- PCR.  Detailed analysis of KNAT7 (Chapter 4) Since KNAT7 was the only one in ten Arabidopsis candidates with an obvious mutant phenotype that affected secondary wall development, phenotypic evaluation (morphology, growth, and vascular organization) of loss-of function lines of KNAT7 was carried out in detail relative to wild type. To investigate potential target genes downstream of KNAT7, expression of a suite of structural genes involved in lignin biosynthesis, secondary wall formation and fiber/xylem development was assayed in the knat7 mutant background. To determine their subcellular localization, constructs were designed to fuse GFP to both Arabidopsis KNAT7 and PoptrKNAT7 and then the gene constructs were transformed into the Arabidopsis knat7 background. Transgenic lines were also tested for the ability of the fusion proteins to complement the knat7 mutant phenotypes.  Analysis of KNAT7 interaction with OVATE family proteins (OFPs) (Chapter 5) KNAT7 is known to interact in yeast two hybrid assays with members of the Ovate Family  42 Protein (OFP) transcription co-regulators. To confirm the interactions in planta, a protoplast transient assay system was used to verify interactions and test if the interaction affects KNAT7 activity. The mutant phenotypes of the interaction partner were examined to test similarity with those of knat7. A series of double mutants was generated, combining overexpression of OFP with the knat7 mutant background, and combining the knat7 mutation with ofp mutations. The phenotypes of these double mutants helped clarify the functional consequences of KNAT7-OFP interaction.  43      CHAPTER 2  Materials and methods   44 Chapter 2. Materials and methods  2.1 Plant materials and growth conditions 2.1.1 Arabidopsis growth conditions Arabidopsis thaliana (Heynh; hereafter Arabidopsis) ecotype Columbia (Col-0) was used as wild type throughout, and all mutants and transgenic lines are in this background. To produce seedlings for use in phenotypic and genotypic analysis, or GUS activity assays, seeds were sterilized and sown on MS plates or selection MS plates consisting of Murashige & Skoog (MS) basal medium with vitamins (PlantMedia, http://www.plantmedia.com), cold-treated at 4°C in the dark for at least 48 hours, then moved to 22°C, with a 16/8 hr (light/dark) photoperiod, unless specified (8/16 hr dark/light) under relatively weak light (60 µmol m−2 sec−1) for 7-10 days. In some cases seedlings were transfered into soil to allow plant maturation, crossing, and seed harvesting. For plant transformation and protoplast transfection, approximately 20 Col-0 or appropriate mutant seeds were germinated and grown in 2×2 inch pots containing a moistened of fertilizer and Sunshine Mix #4 (SunGro Horticulture Canada Ltd, http://www.sungro.com) with a 16/8 hr (light/dark) photoperiod at approximately 120 µmol m−2sec−1 at 22°C, unless specified otherwise (8/16 hr dark/light). Plants that were about 6 weeks old with several mature flowers in the main inflorescence were cut off the primary stems and the plants would be ready for plant transformation when multiple bolting stems with opening flower buds. Leaves from plants that were approximately 3–4 weeks old were used for protoplast isolation.  MS agar medium: 4.3 g/L Murashige and Skoog (MS) salt mixture (Murashige and Skoog, 1962), 1% sucrose (w/v) and 0.6% agar (w/v) dissolved in 1000 mL dH2O. And the pH was adjusted to 5.7 with 1M KOH or 1N HCl. Selection medium: The sterile stock solution of appropriate antibiotic is added to autoclaved medium. eg. Kanamycin medium: 100 µg/mL kanamycin; Hygromycin medium: 50 µg/mL Hygromycin.   45 2.1.2 Plant materials All T-DNA insertion mutants alleles for selected Arabidopsis genes were identified using SIGnal database (http://signal.salk.edu/) and were obtained from the Arabidopsis Biological Resources Center (ABRC, Columbus, OH, USA), the GABI-kat collection (http://www.gabi-kat.de/; (Li et al., 2003; Rosso et al., 2003), the FLAGdb/FST (http://genoplante-info.infobiogen.fr) (Samson et al., 2002), or from the Versailles Genetics and Plant Breeding Laboratory Arabidopsis thaliana Resource Centre (INCR, France) and are listed in Table 2.1. AtOFP1 transposon insertion mutant ofp1-1, AtOFP4 T-DNA insertion mutant ofp4-1 (SALK_022396) and both OFP1 and OFP4 overexpression lines were obtained from Dr. Shucai Wang (Chen lab, UBC). All insertion mutants were examined and confirmed by PCR genotyping using flanking gene-specific primers (Table 2.1) and SALK line T-DNA left border LBb1 (5’-TCAAACAGGATTTTCGCCTGCT-3’), SAIL line LB1 (5’-GCCTTTTCAGAAATGGATAAATAGCCTTGCTTCC-3’), Wisconsin line Wiscp745 (5'-AACGTCCGCAATGTGTTATTAAGTTGTC-3'), FLAGdb/FST line LB4 (5'-CGTGTGCCAGGTGCCCACGGAATAGT-3') or GABI-LB (5’-GGGAATGGCGAAATCAAGGCATCG-3’) primers. Homozygous T-DNA knock out line for MYB61 line (SALK_106556) was a gift from Eric Johnson (Wasteneys Lab, UBC). The ofp1-1 allele was verified by gene-specific primers and primers specific for the transposon element. To verify the T-DNA insertion locations and effects on gene expression, sequencing of PCR products and RT-PCR analysis of target gene full-transcript expression levels were performed. Homozygous lines were identified for phenotype characterization. Double mutants were generated by crossing the two individual homozygous lines, and double mutants were identified in the F2 generation by PCR-aided genotyping. Materials for study of Arabidopsis inflorescence stem development were grown until the inflorescence stem with two or three fully expanded siliques (6–8 weeks). Stems at different ages of development were sampled at 0-3 cm, 3-5 cm, 5-7 cm and 7-9 cm from the inflorescence stem apex as described (Ehlting et al., 2005), and for some phenotypic analyses were sampled 5cm from the bottom of 6-8 week old stems.  46 Table 2.1 List of mutant alleles and oligonucleotides used in study.  Gene alleles T-DNA insetion primers knat7-1/irx11 SALK_002098 UK2098F, AAGTTTGGGCTTGGGCTTGAC UK2098R, TTGCCTTGTCATCTTCCTGTTCA Knat7-2 WiscDsLox367A 5 LP, GATAATTCCGGCGTTGATTTT RP, CGACCTAAAATTTGCAATTGC Knat7-3 SALK_110899 LP. GTCTTGAAAAATTGGTGGCAA RP, GCTTCAAAGAACAGCTGCAAC Knat7-4 SAIL_757 LP, CTTTGGACCGATCAATGAAAG RP, TCTTGAAAAATTGGTGGCAAC AtKNAT7/At1g62990 Knat7-5 WiscDsLox461 LP, CTTTGGACCGATCAATGAAAG RP, TCTTGAAAAATTGGTGGCAAC AtMYB20/At1g66230 myb20-1 GABI_109C11 109C11R, CGA GGA AGT ATG GAA CCC AAT 66230F,TGGCAAGAGTTGCAGACTTCGTTG AtMYB43/At5g16600 myb43-1 SALK_030146 T16600 F: CGAGACTATTGCGGTGTGG T16600 R: TCATGTGGATTGAGCAATGG AtMYB63/At1g79180 NA NA NA TGA1/AtbZIP47/At5g65210 tga1-2 SALK_028212 T65210 F: TCTTTTGGTGCTCTCCAAGG T65210 R: TGAGGAGTTCCTGCTGTTCC AtbZIP9/At5g24800 bzip9-1 SALK_093416 093416-LP TTTTGCTTAGCCATTAAGAAGC 093416-RP TTGTGTTGCGTCTATGAGCTG bHLH068/At4g29100 bhlh068-1 SALK_059682 T29100 F: AGGACCCTGGTCAGGTACG T29100 R: AAATGATGAAACGCCAAACG bHLH144/At1g29950 bhlh144-1 SALK_147291 147291-LP AAGGGAAACCGATCATTGAAG 147291-RP GAGAGCATCGATCTCATTTGG AP2 EREBP/At5g07580 At5g07580-1 SALK_097771 LP, TCAGCATCAAAGGTAAAAGACAAG RP, TGCTAAAATGCATCACAGGAAG C3H/At5g42200 At5g42200-1 FLAG_505G05 505G05LP, CATAATTGGTGAAACCCATGG 505G05RP, TCGTGCTCGATCACATAATTC AtOFP1/At5g01840 Atofp1-1 transposon insertion CDSOFP1F,ATGGGTAATAACTATCGGTTTAAG CDSOFP1R,TTATTTGGAATGGGGTGGTGGAA Tran-element: TACGAATAAGAGCGTCCATTTTAGAGTGA AtOFP4/At1g06920 Atofp4-1 SALK_022396 CDSOFP4F, ATGAGGAACTATAAGTTAAGATTG CDSOFP4R, CTACTTCG ATGCAAATGTAGAG SALK LBb1   TCAAACAGGATTTTCGCCTGCT SAIL LB1   GCCTTTTCAGAAATGGATAAATAGCCTTGCTTCC Wisc P745   AACGTCCGCAATGTGTTATTAAGTTGTC GABI LB   GGGAATGGCGAAATCAAGGCATCG FLAG LB4   CGTGTGCCAGGTGCCCACGGAATAGT  47 2.1.3 Arabidopsis transformation All transgenic lines were generated by transformation of Arabidopsis Col-0 using the Agrobacterium tumefaciens-mediated floral dip method (Clough and Bent, 1998). The Arabidopsis plants were grown in 8-inch pots at a density of 8-16 plants per pot. Single colonies of Agrobacterium strain GV3101 containing a binary vector with the proper construct were inoculated into 5 mL LB (Luria-Bertani) medium and grown overnight at 28º C, and after a PCR verification that the appropriate construct was present, the culture was transferred to 25 mL of LB medium and cultured at 28ºC overnight. The Agrobacterium cultures were transferred to sterile centrifuge tubes and spun for 20 minutes at 4ºC in a SORVALL RC-5C centrifuge with a GS-3 rotor at 3000 rpm. The pellet was resuspended with infiltration medium. The plant inflorescence were dipped upside down into the medium for 5 minutes and then covered with plastic wrap and stored horizontally overnight at RT in the dark. The plants then were placed under long day conditions to allow seed maturation and harvesting. Seeds were harvested and sown onto MS medium containing kanamycin (100 µg/mL) or hygromycin (50 µg/mL) depending on the transgenic constructs. Only the healthy plants which formed healthy true leaves were selected and planted into soil and grown under long days to get obtain the next generation (T2). T2 seeds were selected on MS plates containing 50 µg/ml kanamycin or 25 µg/ml hygromycin, and transgenic plants were transferred to soil. Phenotypes, GFP or GUS expression analysis of transgenic plants were identified in the T2–T4 generations. The expression levels of the transgene were determined by RT-PCR as described below.  Infiltration medium: 2.15 g MS salts (0.5 x) and 50 g sucrose (5%) in 1 L sterile water, pH adjusted to 5.70 with KOH, 300 µL Silwet L-77 (0.02%).  2.2 Gene expression analysis All RNA samples used during this project were extracted from healthy fresh tissues using the Qiagen RNeasy plant mini kit (Qiagen, Valencia, CA) according to the manufacturer’s instructions. During the isolation, RNA was treated with RNase-Free DNase (Qiagen) to avoid contamination with DNA. RNA concentrations were qualified using a  48 spectrophotometer to measure absorbance at 260 nm and 280 nm. After quantification, 2 µg of total RNA was used for transcriptase synthesis either by Omniscript RT reverse transcriptase Kit (Qiagen, USA) or by Superscript II reverse transcriptase (Invitrogen life technology, Canada) according to the manufacturer’s instructions.  2.2.1 Semi-quantitative reverse transcriptase (RT)-PCR Template concentrations and RT-PCR cycle numbers were optimized for all fragment amplified in order to equalize cDNA concentrations in the samples and to avoid saturation of cDNA amplification. The expression level of the ACTIN (ACT/At2g37620) or TUBULIN (TUB9/At4g20890) genes was used as reference. The gene-specific primers were designed for RT-PCR such as that their predicted Tm was at least 55-60°C and that they flanked an intron (to differentiate between cDNA and genomic DNA amplification). General PCR conditions were 94°C for 2 minutes, followed by the number of cycles optimized for the particular primer pair of denaturation at 94°C for 15 s; annealing at 55°C or 60°C for 30 s; and extension at 72°C for 30 s. PCR products were resolved by electrophoresis in a 1% (w/v) agarose-TAE gel containing 5 µg/mL of ethidium bromide and visualized and photographed using the AlphaImager 1220 UV transilluminator equipped with a digital camera. The primers used for RT-PCR was as described in Table 2.2.  50 X TAE: 250 mM NaOAc, 2 M Tris and 50 mM EDTA, use HOAc adjust pH to 7.8. Loading buffer: 100µl 1M pH 8 Tris/HCl, 5 mL 87% (w/v) Glycerol, 0.02 % (v/v) Bromphenolblue (BPB), 0.02 % (v/v) ylene cyanol (XC). X% agarose gel: X g agarose resuspends in 100 mL 1xTAE buffer, 5 µL EtBr stock (0.5 µg/mL) is added, for use immediately or store at 4°C.  2.2.2 Quantitative real-time RT-PCR Real-time PCR methods are powerful analytical tools for the quantification of defined nucleic acid sequences using a fluorescent dye (SYBR® Green) to monitor and qualify the formation of PCR products in solution. Formation of PCR products is quantified after each cycle of the reaction and the kinetics of product formation can be used to determine the relative amount of the target transcript in RNA samples. The basic principle of real-time  49 PCR is the recurring measurement of a fluorescent signal, which is proportional to the amount of amplification product. The amplification cycle at which the emission intensity of the amplification product rises above an arbitrary threshold level (CT cycle) is inversely proportional to the log of the initial number of target sequences (Karsai et al., 2002). For the quantitative analysis of certain gene transcript levels in different tissues/organs of Arabidopsis wild-type Col-0 plants, mutants or wild type of Arabidopsis plants, or Populus trichocarpa (poplar) plants, quantitative real-time PCR was performed using gene-specific primers (Table 2.2). Analyses were performed using an MJ MiniOpticon real-time PCR system (Bio-Rad, http://www.biorad.com). 2 μl cDNA (the cDNA equivalent of 200 ng total RNA) was incubated with 10 µl QuantiTect SYBR Green Mastermix (Qiagen) or IQ SYBR Green Supermix (Bio-Rad) and 30 nmol of each forward and reverse primer in a total volume of 20 µl. Oligonucleotide sequences of all primers are given in Table 2.2. After an initial denaturation at 95°C for 15 min, 35 cycles at 95°C for 30 sec, 60°C for 30 sec, and 72°C for 25 sec followed by a fluorescence reading were performed. After a final incubation at 72°C for 5 min, a melting curve was generated ranging from 95 to 52°C. Threshold cycles (CT) were adjusted manually, and the resulting CT were subtracted from CT values obtained for an ACTIN or TUB9 probe amplified in parallel on each plate thus generating normalized ΔCT values. ΔCT values obtained for RNA from the control (either wild type, certain cell type or certain stem section) was subtracted from ΔCT values for each sample thereby generation the equivalent of log2-ratios comparing expression levels in each sample with the control sample as a reference. For surveys of expression patterns in different organs, RNA was isolated from the inflorescence stems of 6-8 week-old Arabidopsis Col-0 plants at 0-3 cm and 5-7 cm from the apex, cauline leaves, mature rosette leaves, flowers, and mature roots. The expression levels in the 0-3 cm inflorescence stem sample were set to one. For testing gene expression in the knat7 background, RNA samples were isolated from 14-day old seedlings homozygous for the knat7-1 mutation and from wild-type Col-0 plants grown in parallel. The relative mRNA levels were determined by normalizing the PCR threshold cycle number of each gene with that of the Tub9 as reference gene. The expression level of each gene in the wild-type control was set to 100. Poplar RNA samples were isolated from the same sets of pooled tissues  50 harvested from poplar (Populus trichocarpa X P. deltoides) sapling internodes or poplar (P. balsamifera) developing secondary xylem used in poplar microarray hybridization analysis (L. Johnson, M. Friedmann, Douglas lab and Treenomix I project, unpublished). Threshold detection cycles (CT) were normalized using the reference gene C672 (Ralph et al., 2006) and CT values to generate ΔCT values. ΔCT values for each gene were compared to the CT value obtained for xylem sample “G” or internode sample “T-5” to generate ΔΔC(T). Each qRT-PCR assay result reported here was reproduced at least three times, and the standard deviation was from these replicates.  51 Table 2.2 Primer sequences used for real-time PCR analysis and semi-quantitative RT-PCR. Gene Primer pairs    5'- 3' AtKNAT7/At1g62990 62990F, TGCAATCCTGTCTCCTCCACCAAT 62990R, GCAAATGGCCTTACCCTACGGAA For Figure 4.1: CDSAtKNAT7F, ATGCAAGAAGCGGCACTAGGTA CDSAtKNAT7R, TTAGTGTTTGCGCTTGGACTTCA T2098-1, GAGAAATCCATCGGAAGATCAT AtMYB20/At1g66230 66230F, TGGCAAGAGTTGCAGACTTCGTTG 66230R, TGACCACCTGTTTCCAAGCTGG AtMYB43/At5g16600 16600F, TGGGGAGGCAACCATGTTGTGA 16600R, CGCAATAGTCCAGAAAGCTTGGGA AtMYB63/At1g79180 79180F, TGGGCTATTGAGGTGTGGGAAGA 79180R, TCCTCCTCTGAAGTGAAGTTGCCA TGA1/AtbZIP47/At5g65210 65210F, TCTTCGTCATGTCAGGGATGTGGA 65210R, AACCTTGAGAAGATCGGAGGGTCG AtbZIP9/At5g24800 24800F, CGAAAAGGTCCAGCCGGAAACAAT 24800R, TCGACCTCATGAACCGGGATTACA bHLH068/At4g29100 29100F, CGTTGCCACCACATATGACTCC 29100R, GCAAAGACGGTGAAGAAGAAGAGG bHLH144/At1g29950 29950F, TCCAGGGATTATGGAAACACCAC 29950R, GCACTGCCCGATAAACTCTGC C3H/At5g42200 42200F, GGAATGTGCTGTTTGCCTTGA 42200R, AACTGGACAAACCGTGTGGTTAGA PoptrKNAT7/estEXT_fgenesh1_pg_v1.C_LG_I0964 POPKNAT7 F: CCAACTGAAGATGACAAAGCA POPKNAT7 R: CTTGGACTTCAAGGATGTCA PoptrMYB028/fgenesh4_pg.C_LG_V000361 Pop00361rt F, TCCAAGTCCGAAGATGATCAA Pop00361rt R, TGGCATGCTTCAACGTTTTCC PoptrMYB192/eugene3.00070799 Pop70799rt F, CAGACCAGTGGAATTCAGCT Pop70799rt R, GACTATTGTACTCTCCGCAG PoptrC3H1/grail3.0003013001 Pop13001rt F, CTTTAACAGCGACGCTTCCT Pop13001rt R, ACCGGTAAAGGAATGTTCTCT PoptrTGA1/gw1.V.3010.1 PopTGA1rt F, TCTGGTCCGCTTTGTGCAG PopTGA1rt R, AATTGCCCAAAGTGAACTCAG PoptrC3H2/ eugene3.00051495 Pop51495rt F, CCAAAGAGAGGCTGGAGGA Pop51495rt R, TCCGATTGTTTTGCCTACCAA PoptrERF/eugene3.00010657 Pop10657rt F, CAGGAAGAGGAGAAGAGTGA Pop10657rt R, TACTGACACGTAAAGCAAAACT PoptrMYB018/grail3.0038010201 Pop10201rt F, CCTCTTCATTAACATCAGCATC Pop10201rt R, CTTGCTTCCTGCCACTGTC PoptrMYB152/eugene3.00002261 Pop0002261rtF, TCCACTAATATCGTATCTGAAC Pop0002261rtR, TAGCGGAACTTCATCTATGCA PoptrC3H/ fgenesh4_pg.C_LG_V001589 Pop001589rtF, CAGAGATTCGTTTGCCTGTAA  52 Gene Primer pairs    5'- 3' Pop001589rtR, AGTATTGTTCTCTTGAGCAGGA NST2/ At3g61910 NST2F: TCACCCAACCGAGGAAGAGC NST2R: CATGATCGCCACACGAGGAG IRX4 (CCR)/ At1g15950 IRX4F: CGTTATCTCCTAGCCGAGAGTGCTC IRX4R: TGCCATTTTCCACGGATTCTTGCGATGC IRX8/ At5g54690  IRX8F: ACTATCGACGGCGATCCCTCT IRX8R: GCCGCTGCGTTTATCGAGTG IRX13/ At5g03170 IRX13F: AGGTCCAACGAACATAACCGC IRX13R: CAAACCCGAATCCAGTCCTCTC MYB32/ At4g34990 MYB32F: CCTAGATCCGCCGGTCTTCA MYB32R: TCTCGTCCCCGAAATTTGCT MYB61/ At1g09540 MYB61F: AGACTGCAGTTCTTCTTCTCTTTTACTGTT MYB61R: GTCGGATCCTCATTGTTTCAGTTTCTTCT C4H / At2g30490 C4HF: TTCACCGGATCTAACCAAGG C4HR: CGTTGATTTCTCCCTTCTGC COMT/ At5g54160 COMTF: ATGCTCCTTCTCATCCTGGTATTG COMTR: GCAATGTTCGTCACTCCAGTCA PAL2/ At3g53260 PAL2F: GAGGCAGCGTTAAGGTTGAG PAL2R: TTCTCGGTTAGCGATTCACC IRX12/ At2g38080 IRX12F GGTGGATGGGTCGTCATGAGATTC IRX12R CGTGGCGTGATGTTGATATGTCGCCC ATHB-8/ At4g32880 ATHB8-F, CTCAACATCAGCCTCGTGATG ATHB8-R, TCCAGAGATCTGCAATCACGC CCoAOMT/ At4g34050I CCoAOMT-F, TCGTTGATGCTGACAAAGACA CCoAOMT-R, ACTGATCCGACGGCAGATAG 4CL1/ At1g51680 4CL1-F, GGTTACCTCAACAATCCGGCA 4CL1-R, CAAATGCAACAGGAACTTCAC FRA8/ At2g28110 FRA8 F, GACTTGTTGAATCGGTGGCTC FRA8 R, GAAAGAGTTTGACCTTCTAAC BFN1/ At1g11190 BFN1-F, CGTGGACAGAATGCAACGATC BFN1-R, ACCAGCAATAGCATGATCGTC IFL1/ At5g60690 IFL1-F, CCAAGCTGTGAATCTGTGGTC IFL1-R, CGATCTTTGAGGATCTCTGCA CesA8/IRX1/ At4g18780 CesA8 F, TGAGCTTTACATTGTCAAATG CesA8 R, GCAATCGATCAAAAGACAGTT CesA7/IRX3/ At5g17420 CesA7 F, TTGTTGCAGGCATCTCAGATG CesA7 R, GCAGTTGATGCCACACTTGGA CesA4/IRX5/ At5g44030 IRX5F: GCTCAGTGTACCTCGCCAT IRX5R: TTGGACGCCATTGCTGCTTA IRX10/ At1g27440 IRX10-F, CCGAAGGTGGATATTATGCAAGAG IRX10-R, GTATGTTGTCGGGTGATCTGTTGA TUBLIN9/ At4g20890  TUB9F, GTACCTTGAAGCTTGCTAATCCTA  53 Gene Primer pairs    5'- 3' TUB9R, GTTCTGGACGTTCATCATCTGTTC ACTIN1/ At2g37620 ACTIN1F, GCGACAATGGAACTGGAAT ACTIN1R, GGATAGCATGTGGAAGTGCATACC C672 QPCR-F-c672, GACGGTATTTTAGCTATGGAATTG QPCR-R-c672, CTGATAACACAAGTTCCCTGC     2.2.3 GUS expression assay Arabidopsis Col-0 genomic DNA fragments containing 2 kb of DNA 5’ to the start codons of candidate transcription factor genes were generated by PCR using gene-specific primers (as described in Table 2.3), and cloned into the binary vector pCambia 1305.1 (Genbank #AF354045, CAMBIA, Australia) to create Promoter::GUS constructs. The constructs were transformed into wild type Arabidopsis plants by means of Agrobacterium tumefaciens–mediated transformation. The transgenic plants were selected on hygromycin and multiple independent transgenic lines of each construct were examined for GUS activity. PromAtOFP4::GUS transgenic plants were from Dr. Shucai Wang (Chen lab, UBC). GUS activity was assayed in the T2 or T3 generation in 4-day old seedlings and 6-8 weeks old plants by incubating tissues in a solution containing 100 mM sodium phosphate buffer, pH 7.0, 0.1% Triton X-100, 1 mM substrate 5-bromo-4-chloro-3-indolyl-ß-D-glucuronide (X-Gluc; Rose Scientific Ltd, http://www.rosesci.com), and 0.5 mM potassium ferricyanide at 37°C for 1-12 hours. Tissues were cleared with 70% ethanol and placed in 50% glycerol for analysis. For analysis root expression of Prom-AtKNAT7::GUS, the root from stained 4-day seedling was imbedded in Spurr’s resin and sectioned with a microtome (Leica Microsystems GmbH, Germany) to obtain 10-µm-thick sections. All light microscopic observations were performed with OLYMPUS AX70 bright field microscopy.   54 Table 2.3 Primer sequences used to amplify Arabidopsis promoter sequences for GUS fusions.  Gene Forward primer Restriction site Reverse primer Restriction site AtKNAT7/At1g62990 TCTGCAGGTATCGTGT CACTAATAGTATCTC PstI GTTCCATGGGGCTC CCATCATACCTAGTGC NcoI AtMYB20/At1g66230 TCTGCAGGAAGCAAA GAAAAGTGGACGAAG PstI GTTAGATCTGGTTGT CTCCCCATTTCTCTC BglII AtMYB43/At5g16600 GAGCTGCAGAATGAAT TAATTGAGGTAC PstI CACCATGGTTGCCTC CCCATCTCTC NcoI AtMYB63/At1g79180 GTTCTGCAGCTTTGGC GATGAGTTGTTTTC PstI GTTAGATCTGTCTTGT CACAACAAGGTGCTC BglII AtbZIP47/At5g65210 GTTCTGCAGCGTTAC TACGTCACCAGAATCG PstI GTTCCATGGCTCGGT GGCACAAAATGTGTC NcoI AtbZIP9/At5g24800 TAACTGCAGTTGGAG ATTTTCTCTTG PstI GATTATCCATGGTCTT TGAATGTGAACAC NcoI bHLH068/At4g29100 GTTCTGCAGGTATGCG TCATATGACACAATGC PstI GTTCCATGGCTCCAA CACACCTCTATTCATC NcoI bHLH144/At1g29950 GTTCTGCAGCAAGTAG ATTCACGCACGAAAAG PstI GTTCCATGGCTGAGAA GTGAGGAAACTGATTG NcoI AP2 EREBP/At5g07580 GTTCTGCAGGTATTGTA GCTAACTTTGAACTG PstI GTTAGATCTCAGAGCTT TCCTCAAAACTCG BglII C3H/At5g42200 GTTCTGCAGCAACCGCA AACATTGTATAATACG PstI GTTCCATGGGTGTGTAG TGCATCTTTAC NcoI   55 2.3 Complementation analysis and subcellular localization The full-length open-reading frames of AtKNAT7 (At1g62990) and PoptrKNAT7 (EstEXT_fgenesh1_pg_v1.C_LG_I0964) were amplified from Arabidopsis Col-0 cDNA and poplar (P. trichocarpa) cDNA, which prepared from RNAs, respectively, with primers described in Table 2.4, cloned into the pCR8/GW/TOPO TA-cloning vector (Invitrogen, http://www.invitrogen.com), and then subcloned into Gateway plant transformation destination binary vector pGWB6 by LR recombination reactions according to the manufacturer’s instruction (Invitrogen). In these constructs, AtKNAT7 or PoptrKNAT7 was fused in frame at a N-terminal GFP driven by the cauliflower mosaic virus (CaMV) 35S promoter. Binary vectors were transformed into Atknat7 mutants by Agrobacterium-mediated transformation. Multiple independent transgenic lines were selected from each transformation, and two to four representative lines were used for further studies. The expression of transgenes was examined by RT-PCR. Stem phenotypes were examined in hand cross sections prepared from inflorescence stems. Transgenic seedlings 4-7 days old were examined for green fluorescence as described below.  Table 2.4 Primer sequences used to generate GFP fusion constructs.  Gene model Primers AtKNAT7/At1g62990 CDSAtKNAT7F, ATGCAAGAAGCGGCACTAGGTA CDSAtKNAT7R, TTAGTGTTTGCGCTTGGACTTCA PoptrKNAT7/estEXT_fgenesh1_pg_v1.C_LG_I0964 CDSPtKNAT7 F, ATGCAAGAACCAAACTTGGGCA CDSPtKNAT7 R, CTACCTTTTGCGCTTGGACTTC   56 2.4 Microscopy All micrographs were acquired digitally. Adobe Photoshop and ImageJ software (http://rsb.info.nih.gov/ij/index.html, Maryland, USA) were used for image processing.  2.4.1 Bright-field microscopy To characterize certain phenotypes and GUS activity, inflorescence stems of 6-8 week old plants were sectioned either by hand or microtome for resin embedded samples (see 2.4.2). Stem sections about 200 µm in thickness were cut with a razor blade and stained in aqueous 0.02% toluidine blue O (Sigma) for 1 to 2 min, rinsed briefly in distilled water, and mounted in water. Alternatively, the sections were stained and mounted in phloroglucinol (saturated solution in 2 M HCI) and viewed immediately. Samples were viewed using an Olympus AX70 light microscope. Autofluorescence under UV excitation was also performed using this microscope. For seed analysis, after staining with 0.2% w/v aqueous ruthenium red (Sigma) solution or Indian ink, seeds were photographed with Leica MZ6 dissecting microscope. This microscopy was also used to visualize seedlings stained for GUS activity.  2.4.2 Resin embedding for bright-field and TEM For tissue embedding and for light and transmission electron microscopy (TEM), tissue was taken 5 cm from the base of inflorescence stems from 8-week-old plants. Gluteraldehyde fixed stems were placed in 1% osmium tetroxide, 0.05M sodium cacodylate, pH 6.9 for a minimum of 30 minutes under 30 inches of mercury vacuum. Stems were rinsed twice with distilled water then dehydrated through an alcohol series as follows: 30, 40, 50, 70, 80, 90, 95, 100, 100, 100% for a minimum of 15 minutes for each alcohol dilution. Fully dehydrated stems were placed in anhydrous acetone (2X) to facilitate better resin solubility. Pure medium hardness (low-viscosity) Spurr’s resin was added one drop at a time to the stems in acetone, rotated for a minimum of 15 minutes, until a 15% (w/v) resin:acetone mixture was obtained. The resin acetone mixture was then replaced of 25% resin for a minimum of 3 hours, with rotation at room temperature, exchanged for 50% resin, then 75% resin and finally three changes of 100% resin. Samples were oriented for sectioning in flat  57 bottom BEEM capsules and polymerized at 60°C for a minimum of 24 hours. 0.5 µm sections were cut using a Leica Ultracut T and a Druuker diamond Histoknife and stained with 1% Toluidine Blue for bright field and TEM microscopies. TEM images were viewed on Hitachi H7600 PC-TEM (Hitachi Ltd., Tokyo, Japan) at an accelerating voltage of 80kV. Photographs were taken using an ATM Advantage HR digital CCD camera (Advanced Microscopy Techniques, USA). 0.5-µm longitudinal sections and 1-µm cross sections were observed under bright-field using an Olympus AX70 microscope. For cell wall thickness determinations, secondary cell wall measurements were taken from 50 separate cells in light micrograph images from 1-µm inflorescence stem cross sections of wild-type and knat7-1 inflorescence stem bases, using high magnification images. In ImageJ software, I first set the scale with pixels/µm according to the magnification of images, and then drew lines across each cell wall and measured the length of the lines. Finally the measurement was automatically converted into EXCEL. Measurements of cell wall thickness were subjected to statistical analysis using the Student's t test program (http://www.graphpad.com/quickcalcs/ttest1.cfm). The quantitative differences between wild type and mutant in all data sets were shown to be statistically significant (P < 0.0001).  2.4.3 Scanning Electron Microscopy Samples were dry-mounted on stubs, coated with gold or gold-palladium in a sputter coater (SEMrep2, Nanotech, Machester, UK), and observed using Hitachi S4700 Scanning electron Microscope with an accelerating voltage of 30 kV. Photographs were manipulated with ImageJ software.  2.4.4 Confocal laser scanning microscopy To visualize GFP fluorescence, freshly dissected roots from Arabidopsis 7-day seedlings were stained for 1–2 min in an aqueous solution containing 2 mg/mL propidium iodide to stain the cell walls. The roots were then rinsed and mounted in distilled water. The roots were viewed using a Zeiss LSM5 PASCAL confocal laser-scanning microscope equipped with 488 nm argon and 543 nm helium–neon lasers. Images were managed with ImageJ software.   58 2.5 Arabidopsis protoplast transfection assays 2.5.1 Protoplast isolation Arabidopsis leaf mesophyll protoplasts were isolated according to the procedure of (Abel and Theologis, 1994; Tiwari et al., 2006a) with some modifications. Rosette leaves (~1 g) from plants that were 3 to 5 weeks old were collected, washed with deionized water, dried with a paper towel, and cut into 0.5- to 1-mm strips with a razor blade. Leaf sections were transferred to a Petri-dish containing 20 to 25 mL of enzyme solution, vacuum infiltrated for 20 min, and gently shaken (40 rpm on a platform shaker) in darkness for 90 min. After shaking at 80 rpm for an additional 1 min, the protoplasts were filtered (200-µm nylon mesh; Spectrum Laboratories, Rancho Dominguez, CA) and diluted by adding one-third volume 200 mM CaCl2. The protoplasts were pelleted at 180g for 3 min in a Heraeus Multifuge 3S-R centrifuge (Germany), washed once with 25 mL of prechilled W5 solution, repelleted, resuspended gently in 25 mL of prechilled W5 solution, and incubated on ice for 30 min. During the period of incubation, protoplasts were counted using a hemacytometer under a light microscope. The protoplasts were then repelleted and resuspended in prechilled MMg solution at 3 × 105 protoplasts per milliliter for experiments requiring transfected effector genes. The protoplasts in MMg solution could be kept on ice for several hours without affecting the transfection results. Enzyme solution: 1% cellulase R10 (SERVA Electrophoresis, Heidelberg, Germany), 0.25% macerozyme R10 [SERVA Electrophoresis], 0.4 M mannitol, 80 mM CaCl2, and 20 mM Mes, pH 5.7. W5 solution: 154 mM NaCl, 125 mM CaCl2, 5 mM KCl, 5 mM glucose, and 1.5 mM Mes, pH 5.7. MMg solution: 0.4 M mannitol, 15 mM MgCl2, and 4 mM Mes, pH 5.7.  2.5.2 Protoplast transfection assays All effector gene constructs used for protoplasts transfection were generated by amplifying the full-length open-reading frame (ORF) or truncated version of the corresponding genes (primers as described in Table 2.5), then digesting the PCR fragments with NdeI/ClaI and cloning them into a pUC19-based expression vector described by Wang et al. (2007). This placed the fragments in-frame to an amino terminal hemagglutinin (HA) epitope tag or Gal4  59 DBD (GD) tag under the control of the double CaMV35S promoter. Effector AtOFP1 and AtOFP4 genes, transactivator LD-VP16 and reporter LexA(2x)-Gal4(2x):GUS were obtained from the Chen lab, UBC (Wang et al., 2007). All reporter and effector plasmids used in transfection assays were prepared using the EndoFree Plasmid Maxi Kit (Qiagen, Valencia, CA). Ten µg of effector and/or reporter plasmid DNA and 200 µL of the protoplast suspension were mixed with an equal volume of 40% (w/v) PEG3350 (Sigma-Aldrich), and the mixture was incubated at room temperature for 20 min. After incubation, 0.8 mL of W5 solution was added slowly without mixing and incubated for another 10 min. Then the solution was fully mixed and protoplasts were pelleted by centrifugation at 180 g for 3 min. The protoplasts were resuspended gently in 1 mL of WI solution and were incubated at room temperature for 20 to 22 hr in darkness. After incubation, cells were centrifuged at 180 g for 3 min, and the supernatant was removed. The cells were resuspended in 100 µL 1 X cell culture lysis reagent (Promega Corp., Madison, WI; Cat # 153A) and immediately followed with MUG assay. Ten µL of lysed protoplasts were added to a sterile 96 well cell culture cluster plate (Corning Inc.) and 100 µL of MUG Assay Solution was added to each well. The plate was covered and incubated for 0.5-1 hr at 37 °C. After incubation, 100 µL of MUG Stop Solution was added to each well and fluorescence measured using a microplate fluorometer. Expression of 35S:luciferase (Luc) was used to normalize the expression of the GUS reporter. Luciferase activities were measured using a microplate luminometer (Turner Designs, http://www.turnerdesigns.com) together with the Promega Steady-Glo luciferase assay system (http://www.promega.com/). All transfection assays were performed as three replicates, and assays were repeated on at least two separate occasions.  40% PEG [polyethylene glycol] Solution: 40% PEG [avg. mol. wt. 3,350] (Sigma) in 0.1 M Ca(NO3)2 and 0.4 M mannitol solution, pH 10 with 1 N KOH. Filtered and store at –200 C. WI solution: 0.5 M mannitol, 20 mM KCl, and 4 mM Mes, pH 5.7. MUG Buffer: 50 mM sodium phosphate buffer (pH7.0), 10 mM EDTA (ethylenediaminetetraacetic acid), 10 mM DTT (dithiothreitol), 0.1% w/v sodium-lauryl sarcosine and 0.1% v/v Triton X-100.  60 MUG Assay Solution: 1 mM  4-methylumbelliferyl-β-D-glucuronide [MUG] (Acros Organics from Fisher Scientific) in MUG Buffer. MUG Stop Solution: 600 mM Na2CO3, Filtered and store at room temperature.  2.5.3 Bi-molecular fluorescence complementation analysis using protoplasts For BiFC analysis, we used a set of cloning vectors including pSAT6-EYFP-N1 (full enhanced-YFP, EYFP), pSAT6-EYFP-N1 (N-terminal of EYFP) and pSAT4A-cEYFP-N1 (C-terminal of EYFP) in which constructs were placed under the control of the CaMV 35S promoter using multiple cloning sites (Tzfira et al., 2005). We introduced XhoI and HindIII restriction sites into PCR primers used to amplify AtKNAT7, AtOFP1 or AtOFP4 full-length open-reading frames (ORF) without stop codons (Table 2.5) and cloned restriction enzyme digested fragment in-frame into the pSAT6-EYFP vectors resulting in an N-terminal EYFP fusion. Fusions to N-terminal and C-terminal truncated EYFP variants were generated in the same manner. For transient expression using Arabidopsis leaf protoplasts, 10 µg of plasmid DNA was transfected into protoplasts as described above, and incubated for 20–22 hr. After incubation, cells were concentrated by centrifugation to examine them for YFP fluorescence. Two or three drops of cell solution was deposited on a microscope slide and between two cover slips, and covered with another cover slip, which can give enough space to protect protoplasts from damage. Slides were examined immediately using a Leica DM-6000B fluorescent microscope. The yellow fluorescent protein (YFP) excitation was examined and photographed using phase and differential interference contrast (DIC) and a Leica FW4000 digital image acquisition and processing system (Leica Microsystems).  61 Table 2.5 Primer sequences to generate YFP fusions.  Gene Constructs Primers 35S-GD:AtKNAT7 CAATCATATGATGCAAGAAGCGGCACTAGGTA CAATATCGATTTAGTGTTTGCGCTTGGACTTCA 35S-GD:AtKNAT-KNOX1 CAATCATATGATGCAAGAAGCGGCACTAGGTA CAAATCGATTTAGGAAGCGTAAGAACGGAG 35S-GD:AtKNAT-KNOX2 CAACATATGCTCCGTTCTTACGCTTCC CAAATCGATTTAGCCGGAATTATCTGACGAGAA 35S-GD:AtKNAT-Meinox CAATCATATGATGCAAGAAGCGGCACTAGGTA CAAATCGATTTAGCCGGAATTATCTGACGAGAA 35S-GD:AtKNAT- Homeodomain CAACATATGCACGATATGACGGGATTTGGT CAATATCGATTTAGTGTTTGCGCTTGGACTTCA 35S: AtKNAT7- pSAT6-EYFP-N1 AtKNAT7/At1g62990 35S: AtKNAT7- pSAT1A-nEYFP-N1 CAACTCGAGATGCAAGAAGCGGCACTAGGTA CAAAAGCTTCCGTGTTTGCGCTTGGACTTCA 35S:AtOFP1- pSAT6-EYFP-N1 AtOFP1/At5g01840 35S:AtOFP1- pSAT4A-cEYFP-N1 CAACTCGAGATGGGTAATAACTATCGGTT CAAAAGCTTCCTTTGGAATGGGGTGGTGGAA 35S:AtOFP4- pSAT6-EYFP-N1 AtOFP4/At1g06920 35S:AtOFP4- pSAT4A-cEYFP-N1 CAACTCGAGATGAGGAACTATAAGTTA CAAAAGCTTCCCTTCGATGCAAATGTAGAGTT   62 2.6 Database searches, sequence alignments and phylogenetic analysis The 2-kb promoter fragments of candidate Arabidopsis transcription factor genes were obtained from Arabidopsis BAC sequences at the Arabidopsis Information Resource (TAIR; http://www.arabidopsis.org/). For alignment or checking restriction sites, sequences were assembled into a local database using BioEdit (http://www.mbio.ncsu.edu/BioEdit/BioEdit.html) and alignments carried out using Genomatix (http://www.genomatix.de/cgi-bin/dialign/dialign.pl). The functional genomics tools for in silico gene expression analysis in Arabidopsis and poplar were accessed at the Bio-Array Resource for Arabidopsis Functional Genomics (BAR, http://www.bar.utoronto.ca/) (Winter et al., 2007a) and Genevestigator V3 (https://www.genevestigator.ethz.ch/gv/index.jsp) (Hruz et al., 2008). Protein sequences of Arabidopsis transcription factors were obtained from TAIR based on literature, gene family annotations referenced at TAIR, or by BLASTP searches using target proteins sequences as queries. Populus trichocarpa (poplar) protein sequences related to Arabidopsis transcription factors of interest retrieved by BLASTP searches of Jamboree Gene Models (proteins) at the JGI Populus trichocarpa v1.1 database (http://genome.jgi-psf.org/Poptr1_1/Poptr1_1.home.html). Arabidopsis and poplar protein sequences were aligned with Clustal IX, v1.8. The resulting alignments were submitted to PHYML Online at http://atgc.lirmm.fr/phyml/, and trees viewed with TreeView (http://taxonomy.zoology.gla.ac.uk/rod/treeview.html ) Default parameters were used except for number of substitution rate categories (4); the number of bootstrap data sets was 500.   63       CHAPTER 3.  Validation of candidate transcription factors identified from Populus and Arabidopsis microarray expression profiling experiments  64 Chapter 3. Validation of candidate transcription factors identified from Populus and Arabidopsis microarray expression profiling experiments  3.1 Introduction Transcript profiling has the potential to reveal transcriptional networks operating during development, as well as providing expression data for genes of unknown function. Previous studies on global transcript profiling of Arabidopsis inflorescence stems (Ehlting et al., 2005) and root development (Birnbaum et al., 2003) identified transcription factors up-regulated over the course of interfascicular fiber and xylem development (Ehlting et al., 2005). Bioinformatic analyses of genes whose expression is strongly correlated with cellulose biosynthesis in secondary wall development in Arabidopsis (Brown et al., 2005; Persson et al., 2005) also highlighted a number of transcription factor and other genes whose correlated expression patterns suggest that they could play roles in regulating secondary wall development. Ehlting et al. (2005) identified ten candidate transcription factors that are up-regulated in tissues with cells undergoing secondary cell wall formation in at least two independent global expression profiling experiments. Although each experiment had a different focus, in all cases gene expression in samples enriched for cells undergoing secondary cell wall biosynthesis were compared with samples that are mainly characterized by cells with primary walls. Some of the genes identified by Ehlting et al. (2005) are in common with those identified by Brown et al (2005) and Persson et al (2005). Therefore, we believe that these genes are strong candidates for transcriptional regulators of secondary cell wall formation, and are worthy of further investigation. Species of the genus Populus (poplar) are important for a large variety of wood-based products. Since the poplar genome was the first to be sequenced among woody plants (Tuskan et al., 2006), and due to its relative rapid growth and development of genomic tools and resources, it has become a suitable model for plant biology, for example in the study of wood formation (Jansson and Douglas, 2007). A 15.5K-element poplar cDNA microarray has been developed and used to study different aspects of poplar biology including response  65 to insect and pathogen attack (Ralph et al., 2006; Miranda et al., 2007). This microarray has also been used to profile global changes in gene expression associated with the onset of secondary xylem formation at the poplar shoot apex and secondary xylem development in mature trees (Treenomix I project and Douglas lab, unpublished). In parallel, many poplar genes represented on this array potentially involved in cellulose and lignin biosynthesis deposition have been annotated based on similarity to Arabidopsis homologues and by analysis of phylogeneic trees, and gene expression data (Tuskan et al., 2006; Hamberger et al., 2007). These data opened the door to investigations of the expression patterns of the putative poplar orthologs of Arabidopsis genes differentially expressed during secondary wall formation. In this study, we confirmed the differential expression of ten candidate transcription factor genes, which were identified by Ehlting et al., (2005) using quantitative real-time RT-PCR to assay expression in the same samples used in the microarray experiment. Additional expression assays, as well as in silico analyses, were performed to examine tissue and organ specific expression of these genes in Arabidopsis. We conducted initial phenotypic evaluation of T-DNA insertion mutant lines for these genes to identify potential functions in fiber and secondary wall development. In addition, the putative poplar orthologs of these genes were identified and their expression patterns examined during poplar stem and secondary xylem development using real-time PCR.  3.2 Results 3.2.1 Confirmation Arabidopsis microarray experiment data on differential expression Global expression profiling of Arabidopsis genes over the course of Arabidopsis inflorescence stem development, using a longmer microarray representing 25,792 Arabidopsis genes, identified differentially regulated transcription factors that could be involved in regulating processes of fiber differentiation and maturation (Ehlting et al., 2005). This group of transcription factors has members from six different families: three MYB (MYB20, At1g66230; MYB43, At5g16600; and MYB63, At1g79180), two bHLH (bHLH68, At4g29100 and bHLH144, At1g29950), two bZIP (bZIP9, At5g24800 and bZIP47, TGA1,  66 At5g65210), one AP2-EREBP (At5g0758), one C3H (At5g42200), and one homeodomain protein (KNAT7, At1g62990) (Ehlting et al., 2005), and each gene was up-regulated in association with interfascicular fiber development (Ehlting et al., 2005) as well as in association with xylem development in the stele of Arabidopsis roots (Birnbaum et al., 2003). Quantitative real-time RT-PCR was used to confirm expression of the candidate genes during the course of stem development. cDNAs prepared from stem RNA samples were used in microarray analysis to perform real-time PCR (Figure 3.1). As a reference standard for expression, the ACTIN1 gene was amplified in parallel with each of the target genes. The ∆∆CT values normalized to ACTIN1 expression values were calculated for each gene relative to the expression level in the RNA sample from the top of the stem (0-2 cm sample) as described in Materials and Methods (chapter 2.2.2) and are shown as Figure 3.1. Each RT-PCR reaction was reproduced twice and results from these duplicates are shown for each gene. The RT-PCR results show clear evidence for differential expression of nine of the ten transcription factor genes tested. Only for bHLH144 (At1g29950) was no clearly reproducible increase in transcript abundance relative to the 0-2 cm sample observed.  67   Figure 3.1 Validation of candidate transcription factor gene expression over the course of Arabidopsis inflorescence stem development.  Data are from quantitative real-time RT-PCR (qRT-PCR) assays conducted with QuantiTect SYBR Green Mastermix (Qiagen) and gene specific primers. Threshold detection cycles (CT) were normalized using the corresponding ACTIN CT values to generate ∆CT values. ∆CT values for each gene were compared to the ∆CT value obtained for the 0-2 cm sample, which was used as a reference in microarray experiments and has therefore also been used as a reference here. The horizontal axis shows origins of different stem samples used for RNA isolation (in cm from the top of the stem). The vertical axis indicates ∆∆CT values relative to RNA from the 0-2 cm stem section (set at 0). Two duplicate RT-PCR results from the same cDNA sample are shown with the two individual bars for each sample.  68  3.2.2 Organ and tissue expression profiles of candidate genes in Arabidopsis tissues Previous microarray experiments focused on differential gene expression in various tissues, cell types, and developmental stages of the Arabidopsis root (Birnbaum et al., 2003) showed that the ten transcription factor candidate genes are expressed in a stele-specific manner, and are up-regulated during the course of stele development (Birnbaum et al., 2003). To further investigate the expression patterns of these candidate transcription factor genes in different tissue/cell types, I used data mining tools to query gene expression data from publicly available microarray experiments. I used both the GENEVESTIGATOR V3 (Hruz et al., 2008); https://www.genevestigator.ethz.ch) and BAR (Winter et al., 2007a); http://bbc.botany.utoronto.ca/) sites for these analyses, and retrieved the data that are summarized in Figure 3.2. Figure 3.2A shows a graphical summary of the in silico analysis of relative expression levels of the ten transcription factor genes in numerous organs, cells, and developmental stages retrieved from the ANATOMY expression set in Genevestigator V3 site based on data from 3110 Affymetrix ATH1 22K arrays, with the number of arrays used for each expression analyses shown. According to this analysis, the expression in root, stem, xylem and cork was generally associated with the microarray data from stem and root development in the inflorescence stem and root studies (Birnbaum et al., 2003; Ehlting et al., 2005). Figure 3.2B shows an in silico analysis of the expression levels of the ten transcription factor genes in different organs from the “Developmental Map” in Arabidopsis eFP Browser at BAR (http://bbc.botany.utoronto.ca/efp/cgi-bin/efpWeb.cgi). A comparison of the relative expression levels in Figure 3.2B shows that the transcripts of all ten candidate genes are more abundant in roots and in the stem 2nd internode compared to the 1st node, which is consistent with the criteria used to select the candidates. A further in silico analysis of the tissue/cell specific expression patterns of the candidates is shown from the “Tissue Specific” in eFP Browser at BAR is shown in Figure 3.2C. According to this analysis, the genes have generally higher expression in root stele, xylem and cork, again consistent with the criteria used to select them as candidate regulators of secondary wall formation. To experimentally examine the expression of the candidate regulatory genes, relative  69 transcript levels of all ten Arabidopsis candidates were analyzed in different organs using quantitative real-time RT-PCR. RNA samples were isolated from wild type (Col-0) inflorescence stems at 0-3 cm and 5-7 cm from the tip, cauline leaves, mature leaves, flowers and mature roots. CT values were normalized using the corresponding ACTIN1 reference gene CT values to generate ∆CT values relative to the 0-3 cm samples, which was set at 1. Consistent with the microarray data (Birnbaum et al., 2003; Ehlting et al., 2005), most genes exhibited relatively high expression levels in both roots and mature stems (5-7 cm), and low expression in flowers and leaves. The AP2/EREBP gene At5g07580 also showed a high expression level in mature leaves (Figure 3.3), which is consistent with the public microarray data on expression of this gene (Figure 3.2).   70   A  B     71 C     Figure 3.2 In silico analysis of relative expression levels of ten candidate transcription factor genes in Arabidopsis.  The data was retrieved from Genevestigator and BAR public databases. (A) Graphical overview of relative expression from Anatomy in Genevestigator V3 https://www.genevestigator.ethz.ch). The number of arrays from which the data was obtained in each organ, tissue, and/or developmental stage is given. (B) Relative expression different organs; data from Developmental map in BAR (http://bbc.botany.utoronto.ca/ (C) Relative expression different tissue/cell types; data from tissue specific in BAR (http://bbc.botany.utoronto.ca/.  72      Figure 3.3 Experimental analysis of expression of ten Arabidopsis candidate transcription factor genes in different organs.  Quantitative real-time RT-PCR analyses were carried out using cDNA samples isolated from the organs indicated. The ACTIN1 gene (At2g37620) was used as a reference gene. Expression levels of each gene in the 0-3 cm stem sample were set to 1.0, and expression in other organs is given relative to this value. The values are the means of reactions run in triplicate from the same cDNA.  73 3.2.3 Cell and tissue expression patterns of candidate gene promoter-GUS fusions In order to investigate the cell-type expression patterns controlled by the promoters of the ten candidate genes during root and stem development, approximately 2-kb promoter fragments upstream of the ATG start codons from most of the candidate genes (except At5g42200/C3H, for which a promoter fragment was not isolated for technical reasons) were fused to the GUS reporter gene and introduced into Arabidopsis by Agrobacterium-mediated transformation. GUS activity in multiple lines (4-8 independent transgenic lines for each construct) was surveyed by histochemical assays. Samples from transgenic generation T2 plants were analyzed in seedlings 4 days after germination and at 7-9 cm from apex of mature inflorescence stems. A summary of expression patterns of all candidate genes except At5g42200 is given in Table 3.1. Images from representative lines for prom-AtMYB20::GUS, prom-AtMYB43::GUS, prom-AtMYB63::GUS and prom-AtKNAT7::GUS are shown in Figure 3.5, stained for GUS activity. Expression patterns specified by all candidate MYB gene promoters promAtMYB20::GUS (Figure 3.4A-C)  promAtMYB43::GUS (Figure 3.4D-F) and prom-AtMYB63::GUS (Figure 3.4G-I) showed strong activity in vascular systems of young seedlings, especially in roots, where it was mostly restricted to the vascular cylinder (Figure 3.4B, E, H). Expression was also observed in the vascular system of hypocotyls and cotyledons. At the 7-9 cm of inflorescence stems, GUS activity was observed in cortex cells adjacent to interfascicular fibers, and associated with vascular bundles in the phloem or cambial regions and in adjacent xylem, where it appeared to be mostly in developing vessels of protoxylem (Figure 3.4C, F, I). Expression of the promKNAT7::GUS transgene in seedlings was similar to that of the MYB candidates, except that expression was stronger and more restricted to vascular tissues in the hypocotyl and cotyledons (Figure 3.4J). In the root, cross sections revealed stele-preferred expression, consistent with previous data (Birnbaum et al., 2003) (Figure 3.4K). Expression appeared to be stronger in the differentiation and maturation zone of the root and at the root-hypocotyl junction (Figure 3.4J). In the cross sections of inflorescence stem, quite high GUS expression was observed in the cortex adjacent to interfascicular fibers, as well as in vascular bundles, in phloem or cambial regions adjacent to the xylem, and like MYB genes,  74 activity was associated with developing vessel elements in the protoxylem (Figure 3.4L).     Figure 3.4 Histochemical localization of GUS activity in transgenic Arabidopsis lines expressing promoter-GUS fusions.  promAtMYB20::GUS (A, B, C), promAtMYB43::GUS (D, E, F), prom-AtMYB63::GUS (G, H, I) and promAtKNAT7::GUS (J, K, L) are showing 4-day old seedlings (A, D, G, J); 4-day old seedling root at higher magnification (B, E, H) and 10 µm-thick root cross section from a 4-day-old seedling showing expression in the stele (K). C, F, I, L are cross-sections of inflorescence stems at 7-9 cm from the apex (C, F, I from 6-week old plants; L from 8-week-old plant). Bars 50 µm for C, F, I, J and K; 100 µm for B, E, H.  75  Table 3.1 Summary of histochemical assays of GUS activity in transgenic lines expressing candidate gene promoter-GUS fusions.  Gene Location of GUS activity  4 day seedling Stem section At4g29100 Hypocotyl/root junction, root/stele, leaf veins Vascular bundles/metaxylem At1g29950 Hypocotyl; root cylinder Cortex; vascular bundles At5g16600 Vascular system; root cylinder cortex cells adjacent to interfascicular fibers; Vascular bundles/metaxylem At1g79180 Vascular system; root cylinder cortex cells adjacent to interfascicular fibers; Vascular bundles/metaxylem At1g66230 Vascular system; root cylinder cortex cells adjacent to interfascicular fibers; Vascular bundles/metaxylem At1g62990 Hypocotyl/root junction; root/stele Cortex adjacent to interfascicular fibers; cambial region At5g65210 Hypocotyl/root junction, stronger in root/cylinder compare to cortex, leaf veins Vascular bundles/metaxylem At5g24800 Root cylinder; root/hypocotyl junction; cotyledon Vascular bundles/metaxylem; cambial regoin At5g07580 Root cylinder; root/hypocotyl junction; vascular system Epidermis, cortex, stronger in vascular bundle At5g42200 NA NA    76 3.2.4 Confirmation of T-DNA insertion sites in potential knock-out mutants We identified T-DNA insertion lines for all ten of the Arabidopsis transcription factor candidates using the SIGnal database http://signal.salk.edu/cgi_bin/tdnaexpress), focusing initial analysis on insertions in exons or introns where possible. Primers flanking the insertion sites were used in conjunction with a primer from left border of T-DNA to identify lines homozygous for the T-DNA insertion in each line. To verify the location, sequencing of T-DNA/genomic DNA junctions was performed. Homozygous lines were then used for further phenotype characterization. These data are summarized in Table 3.2. While this work was underway, analyses of T-DNA insertion lines for KNAT7/At1g62990 and MYB43/ At5g16600 were published (Brown et al., 2005; Persson et al., 2005), and we used the same alleles as those authors. We conducted phenotypic analyses of all homozygous mutant lines by observing basic growth and development, and by sectioning developing inflorescence stems to view any possible changes in anatomy or cell morphology. Most homozygous mutants exhibited no obvious different phenotypes, and showed no apparent changes in interfascicular fiber or xylem vessel development, or in secondary wall formation, relative to wild type. Because two candidates genes MYB20 and MYB43 are closely related paralogs (Stracke et al., 2001) and could have overlapping functions, we generated a myb20/myb43 double mutant by crossing homozygous lines. However, the double mutant also appeared phenotypically normal (Table 3.2). In contrast to most lines that could not be distinguished from the wild type, the plants homozygous for the knat7-1 allele showed in irregular xylem (irx) xylem morphology phenotype. While wild type xylem is characterized by vessel elements with a relatively round shape, the vessels from knat7-1 are irregular and frequently collapsed inwards (shown in further detail in Chapter 4, Figure 4.2), consistent with previous reports on the initial phenotypic characterization of this mutant (Brown et al., 2005; Persson et al., 2005). We confirmed the location of the knat7-1 T-DNA insertion line SALK_002098 and used RT-PCR to show the apparent complete loss of KNAT7 transcripts in homozygous plants (shown in Figure 4.1). Because of the strong association of KNAT7 expression with secondary wall formation and a knat7 phenotype suggesting a functional role in secondary wall formation, we looked in more detail at knat7 phenotypes, presented in Chapter 4.  77  Table 3.2 Summary of T-DNA insertion lines and mutants used for phenotypic analyses.  AGI code Gene name T-DNA insertion Insertion site Allele Phenotype At4g29100 bHLH068 SALK_059682 intron bhlh068-1 WT At1g29950 bHLH144 SALK_147291 exon bhlh144-1 WT At5g16600 MYB43 SALK_030146 intron myb43-1 WT At1g79180 MYB63 NA NA NA NA At1g66230 MYB20 GABI_109C11 exon myb20-1 WT At1g62990 KNAT7 SALK_002098 intron knat7-1/irx11 irx At5g65210 bZIP47/TGA1 SALK_028212 exon tga1-2 WT At5g24800 bZIP9 SALK_093416 exon bzip9-1 WT At5g07580 AP2-EREBP SALK_097771 exon At5g07580-1 WT At5g42200 C3H FLAG_505G05 300-5’ UTR At5g42200-1 WT At1g66230/ At5g16600 MYB20/MYB43   myb20-1 /myb43-1 WT   78  3.2.5 Potential poplar orthologues of candidate genes To investigate conservation of the Arabidopsis candidate transcription factor genes in other plants, we first determined if potential poplar orthologs of these Arabidopsis candidate genes exist in the fully sequenced poplar genome, and then determined whether the potential orthologs were also differentially regulated in concert with secondary xylem and secondary wall formation in this model woody plant. The closest poplar and Arabidopsis homologues to each of the 10 candidate transcription factors were identified by separate BLASTP searches of the translated Arabidopsis and poplar genomes using each of the candidate genes as queries (analyses performed by Margaret Ellis, Treenomix I project and Douglas lab). After alignment of the amino acid sequences, Ms. Ellis generated rooted maximum likelihood trees of the combined sets of poplar and Arabidopsis proteins. The results for bZIP, KNOX, MYB, and C3H zinc finger classes of candidate genes are shown in Figure 3.5. For each of the classes shown, it was possible to identify potential orthologs based on phylogenetic relationships. Within the KNOX family of homeodomain encoding genes, this analysis confirmed that a single poplar gene is a likely orthologue of KNAT7, PoptrKNAT7 (poplar gene model estExt_fgenesh1_pg_v1.C_LG_I0964) within the class II KNOX genes (Figure 3.5C). In another case, the duplicated poplar MYB paralogs PoptrMYB108 and PoptrMYB152 are both potential orthologs of one or both of the Arabidopsis candidate paralogs MYB20 and MYB43 (Figure 3.5B). For poplar homologues of Arabidopsis bZIP and zinc finger C3H candidates (Figure 3.5A and D), one or two potential orthologs were evident, but were less clear due to the presence of duplicated poplar genes. The potential poplar orthologs of each of the 10 Arabidopsis candidates are summarized in Table 3.3, giving the P. trichocarpa gene model names.  79   Figure 3.5 Phylogenetic reconstructions of selected transcription factor gene families in Arabidopsis and poplar.  BLAST searches were used to identify poplar and Arabidopsis genes most closely related to candidate Arabidopsis transcription factor genes up-regulated in association with interfascicular fiber differentiation and root stele development shown in red (Ehlting et al., 2005); See Materials and Methods for details. Protein sequences were aligned with Clustal X, v1.8, and rooted, bootstrapped trees generated using PHYML (http://atgc.lirmm.fr/phyml/). Asterisks indicate branches with 80% or higher bootstrap support. Poplar genes are designated according to the annotated gene model at the JGI Populus trichocarpa v. 1.1 genome browser (http://genome.jgi-psf.org/Poptr1_1/Poptr1_1.home.html). Putative poplar orthologs of Arabidopsis candidate genes are shown in red (up-regulated in association with secondary wall formation in two poplar microarray experiments; Treenomix I and Douglas lab unpublished data) or in blue (no up-regulation in association with secondary wall formation in either one poplar microarray experiment; Treenomix I and Douglas lab unpublished data). A) bZIP family; B) MYBs; C) KNOX; D) C3H. Phylogenetic data shown in this figure were generated by Margaret Ellis, (Treenomix I project and Douglas lab).  80  3.2.6 Expression profiling of potential poplar orthologues of candidates in poplar arrays Data from two experiments performed using the Treenomix 15.5K poplar cDNA microarray (Ralph et al., 2006) were used to identify poplar genes differentially regulated in association with secondary xylem development at the poplar shoot apex, and differentially regulated over the course of secondary xylem development in wood taken from mature field grown trees (Figure 1.8; Treenomix I project and Douglas lab, unpublished data). We determined if there were elements presenting on the 15.5K cDNA microarray corresponding to the potential poplar orthologs of the ten Arabidopsis candidate genes (Figure 3.5 and Table 3.3), and if present determined whether the poplar genes were differentially regulated in one or both of the two microarray experiments. As indicated in Table 3.3, potential poplar orthologs of bHLH144 (poplar bHLH gene fgenesh4_pm.C_LG_XI0001300), bHLH068 (poplar bHLH gene estExt_fgenesh4_pg.C_LG_XVIII0694), KNAT7 (PoptrKNAT7), MYB20 and MYB43 (PoptrMYB018), MYB63 (PoptrMYB028 and PoptrMYB192), bZIP9 (poplar bZIP eugene3.01470069), TGA1/bZIP47 (poplar bZIP gene gw1.VII.1997.1), and C3H/At5g42200 (PoptrC3H1, grail3.0003013001), were represented by cDNA elements on the array. Of these poplar genes, PoptrKNAT7 was clearly differentially expressed in both poplar microarray experiments in association with secondary wall formation. In addition, both PoptrMYB028 and PoptrMYB018 were also differentially expressed in both experiments, although the fold-change values were more modest compared to PoptrKNAT7. In contrast, the predicted poplar orthologs of the Arabidopsis bHLH, bZIP and C3H candidates were either not differentially expressed or the differential expression was not evident in one of the two experiments. For example, while both gw1.VII.1997.1 (TGA1/bZIP47 ortholog) and grail3.0003013001 (C3H ortholog) were differentially expressed during internode maturation, this differential expression was not observed during secondary xylem development in the second experiment for both genes (Table 3.3). These results support a potential role for a subset of the poplar and Arabidopsis orthologs in secondary wall formation in both species. To validate the microarray expression data, and to obtain data on candidate gene orthologs not represented by elements on the 15.5K  81 microarray, I used quantitative real-time RT-PCR to assay expression of selected poplar genes. Gene-specific primers for poplar genes (Table 2.2) were used with cDNA prepared from RNA isolated from the same tissues used for microarray analysis. The results presented in Figure 3.6 and summarized in Table 3.3 confirmed differential expression of PoptrMYB028, PoptrMYB018, and PoptrKNAT7. For each gene, increases in expression in coordination with the onset of secondary wall formation were observed in both experiments. The differential expression of the C3H ortholog grail3.0003013001 over the course of internode maturation, but not during over the course of secondary xylem development, was also confirmed. In addition, PoptrMYB152, a potential ortholog of Arabidopsis MYB20 and MYB43 but not represented on the array, as showed differential expression in the transition to secondary wall formation in secondary xylem (e.g., O to M and CC transition; see Figure 1.8), and differential expression in the early stages of internode maturation (T5 to T4 transition; see Figure 1.8). However, one of the potential ortholog of MYB63 (PoptrMYB192/eugene3.00070799), and potential poplar orthologs of At5g07580 (AP2-EREBP, PoptrERF/eugene3.00010657) and TGA1 (PoptrTGA1/gw1.V.3010.1) showed no clear differential expression in association with secondary wall formation in either experiment. PoptrC3H2/ fgenesh4_pg.C_LG_V001589, a second possible ortholog of C3H/ At5g42200 also not presenting on the array, showed an expression pattern very similar to its paralog PoptrC3H1 (grail3.0003013001), with increasing expression during internode maturation, but no differential expression in developing secondary xylem (Figure 3.6).  82 Table 3.3 Proposed poplar homologues of Arabidopsis transcription factors associated with fiber and secondary wall development and differential expression in two microarray profiling experiments.  Arabidopsis Gene Poplar homologue gene model Poplar annotation FC1 T5-T2   G-CC RT- PCR2 At1g29950/ AtbHLH144 fgenesh4_pm.C_LG_XI000130 bHLH 1.0      0.74* NT At4g29100/ AtbHLH068 estExt_fgenesh4_pg.C_LG_XVIII0694 bHLH 0.97     1.1 NT At1g62990/ AtKNAT7 estExt_fgenesh1_pg_v1.C_LG_I0964 KNOX 4.1*     3.9* + At1g66230/ AtMYB20 grail3.0038010201 PoptrMYB018 2.5*     1.5* + At5g16600/ AtMYB43 eugene3.00002261 PoptrMYB152 NA      NA + At1g79180/ AtMYB63 fgenesh4_pg.C_LG_V000361 eugene3.00070799 PoptrMYB028 PoptrMYB192 2.0*     1.4* NA      NA + - At5g07580/ AP2-EREBP eugene3.00010657 ERF NA      NA - At5g24800/ AtbZIP9 eugene3.01470069 bZIP 0.87     0.68* NT At5g65210/ AtbZIP47/TGA1 gw1.VII.1997.1 gw1.V.3010.1 bZIP bZIP 1.4*     1.07 NA      NA NT - At5g42200/ C3H grail3.0003013001 fgenesh4_pg.C_LG_V001589 C3H1 C3H2 1.7*     0.9 NA      NA +/- +/- 1 Fold-change values in the two microarray experiment comparisons (Figure 1.8). FC values with stars have p-values <0.01; NA, not available (no element on array). 2 Expression data from quantitative RT-PCR experiments (Figure 3.6). +, increased expression associated with secondary xylem maturation in both experimental systems; +/-, increased expression associated with secondary xylem maturation in stem development but not in seasonally activated secondary xylem development; -, no significant expression levels changed in both experiments; NT, not tested.  83     Figure 3.6 Analysis of differential expression of selected Populus transcription factor genes using quantitative real time RT-PCR.  Real time RT-PCR using SYBR Green with gene specific primers for the genes indicated at top in each graph was performed with cDNA derived from RNA of the two poplar experiments (Figure 1.8), as indicated at bottom of each graph. The reference gene used was C672 (Ralph et al., 2006). Threshold detection cycles (CT) were normalized using the corresponding C672 CT values to generate ΔCT values. ΔCT values for each gene were compared to the CT value obtained for samples G or T5 to generate ΔΔC(T) values. Each reaction was technically replicated at least three times, and values are shown as the mean +/- SD.  84 A publicly available database on poplar developmental series based on Affymetrix expression data generated by Campbell Laboratory has recently become available (http://bbc.botany.utoronto.ca/efppop/cgi-bin/efpWeb.cgi), and allows in silico analysis of poplar gene expression patterns. The data I retrieved from BAR showed the absolute signal intensities of potential poplar orthologs of the ten Arabidopsis candidate genes except AtMYB20/At1g66230, for which no poplar ortholog was identified. A comparison of relative expression levels of 9 poplar orthologs of the ten Arabidopsis candidate genes, annotated according to the Arabidopsis AGI code name, is shown in Figure 3.7. Except At5g16600/PtpAffx.203.1.S1_at, At4g29100/PtpAffx.141217.1.A1_at and At5g42200/PtpAffx.66132.1.S1_a_at, the relative expression levels were low in all tissues, and some genes exhibited specific expression in certain tissues. Interestingly, At5g65210/PtpAffx.21973.1.A1_at and At5g07580/PtpAffx.42822.1.A1_at exhibited specific expression in roots and leaves respectively, which is consistent with the expression patterns of their orthologs in Arabidopsis (highest expression also in root and in leaves respectively). Therefore, At5g65210/TGA1 and At5g07580/AP-EREBP might have conserved functions in root or leaf development in both species. At1g62990 (KNAT7)/PtpAffx.18687.1.A1_at has an extremely high expression level in xylem compared to other tissues and other genes (Figure 3.7), in agreement with our expression data (Table 3.3; Figure 3.6).  85      Figure 3.7 Public microarray data on relative expression levels of poplar orthologs of Arabidopsis candidate genes in different tissue.  Poplar orthologs of the Arabidopsis candidate genes shown, and their expression data from poplar Affymetrix microarrays (array identifier given) were obtained from the BAR eFP site (http://bbc.botany.utoronto.ca/efppop/cgi-bin/efpWeb.cgi). Relative expression levels of each gene is in young leaf, mature leaf, root, male catkins, female catkins and xylem are shown.  86 3.3 Discussion The secondary cell wall of Arabidopsis, like other vascular plants, is composed predominantly of cellulose, lignin, and xylan, making it an attractive model for the study of the mechanism of secondary wall deposition. Genetic screens, based upon the irregular xylem (irx) mutant phenotype in stem sections, have led to the isolation of several important genes required for secondary cell wall biosynthesis, including those involved in cellulose and lignin biosynthesis (Turner and Somerville, 1997; Jones et al., 2001; Brown et al., 2005). Global analysis of changes in gene expression associated with secondary wall deposition during organ and tissue differentiations, using microarray expression profiling, has also been broadly used to identify genes of unknown function that could also be involved in secondary wall formation (Birnbaum et al., 2003; Oh et al., 2003; Ko et al., 2004; Ehlting et al., 2005; Kubo et al., 2005; Ko et al., 2006; Yokoyama and Nishitani, 2006; Wenzel et al., 2008). Ten candidate transcription factor genes were identified from microarray analysis of the developmental transitions from the top to the bottom of Arabidopsis inflorescence stems, during which interfascicular fiber is a prominent event (Ehlting et al., 2005). These candidates include bZIP, MYB, C3H, AP2-EREBP, bHLH and HD family members. For most of these ten genes, there is little or no functional information. For example, the bZIP9 (At5g24800) is an uncharacterized member of the basic leucine zipper motif (bZIP) transcription factor class. Interestingly, a second bZIP gene, TGA1 (At5g65210) together with other TGA partner proteins, is known to play a key role in defense signaling by interaction with the regulatory protein NPR1 (Despres et al., 2003) and was previously found to be more than two-fold more highly expressed in Arabidopsis secondary xylem relative to bark (Oh et al., 2003). Our data and publically available microarray data show that this gene is remarkable for its high expression level in roots relative to other organs (Figures 3.2A, and 3.3), suggesting a potentially important role in root development. The data presented in this chapter help to evaluate the functions of these genes as regulators of secondary wall formation. Developmental expression patterns (RT-PCR and in silico) showed, as predicted, that most of the candidate genes are more highly expressed in stems and roots, with preferential  87 expression in older portions/xylem. These data are consistent with their roles as candidates selected from microarray data (Birnbaum et al., 2003; Ehlting et al., 2005). Most of the nine candidate genes (except C3H/At5g42200) were predominantly expressed in cells undergoing secondary wall thickening in stems and specific expression in root stele of young seedlings using promoter-GUS fusions. Some of the candidates are not specifically expressed in cells associated with secondary wall formation, but also expressed in other types of cells as cortex or epidermis, which implies that these genes are also associated with other developmental processes of stem in addition to secondary wall formation, or contribute to secondary wall formation in a non cell-autonomous manner (Figure 3.4, Table 3.1). The poplar orthologs of the ten Arabidopsis candidate genes were identified, and tested for expression in association with secondary wall formation in poplar. This showed that KNAT7 and three MYBs (MYB20, MYB43, and MYB63) are conserved in poplar and Arabidopsis, and associated with secondary wall formation in both species. But, the potential role of MYB63 is not clear due to lack of T-DNA insertion mutant lines. It is interesting that MYB58 and MYB63 is a paralog pair in Arabidopsis, and both exhibit similar differential expression patterns in Arabidopsis and poplar, for example, AtMYB63 is up-regulated in association with secondary wall formation in the inflorescence stem, but AtMYB58 is not (Ehlting et al., 2005). Similarly, one poplar paralog of Arabidopsis MYB58/MYB63, PoptrMYB028 (fgenesh4pg.C_LG_V000361) is up-regulated in association with the secondary xylem development in poplar, but the second poplar paralog, PoptrMYB192 (eugene3.00070799) is not (Table 3.3; Figure 3.6). Other experiments have also shed light on potential roles of these candidate genes in xylem, fiber, and secondary wall formation. For example, in order to profile expression of xylem cell-differentiation-related genes in Arabidopsis, an in vitro xylem vessel element inducible system was established from Arabidopsis suspension cells, and changes on gene expression monitored using the Arabidopsis full-genome GeneChip array ATH1 (Affymetrix) (Kubo et al., 2005). Many genes showing up-regulated expression just when the xylem vessel elements were actively forming (6 days after induction) were identified, including MYB63, MYB43, and KNAT7 (Kubo et al., 2005). This suggests that these genes might control sets of genes required for xylem vessel formation. In addition, previous studies have revealed that  88 MYB20 and MYB43 are regulated by SND1, a master transcriptional switch activating the developmental program of secondary wall biosynthesis, and by its direct target gene MYB46 (Zhong et al., 2007a; Zhong et al., 2008). Finally, coexpression analysis based of publicly available data has identified genes that are coexpressed with cellulose synthase (CESA) genes required for either primary or secondary cell wall formation. KNAT7 was one of the genes most highly coregulated with CESA4, 7 and 8, encoding CESA subunits involved in secondary wall formation (Persson et al., 2005). As in our study, the combination of expression analysis and reverse genetics has led to the identification of many genes involved in plant processes of interest. However, estimates from the number of phenotypes found based upon the T-DNA insertion mutants suggest that perhaps mutations in only one in ten genes will give a clear phenotype (Brown et al., 2005). My results were consistent with this estimate, showing that the phenotype of most of the lines studied (including the myb20/myb43 double mutant) were indistinguishable from the wild type. However, in contrast to most lines, plants homozygous for the knat7-1 allele showed an irregular xylem (irx) phenotype, consistent with previous reports on the initial phenotypic characterization of this mutant (Brown et al., 2005; Persson et al., 2005). The mutant plants appear morphologically normal, but have significantly altered xylem morphology in stem sections. The irx phenotype is presumably caused by the collapse of the xylem due to defects in secondary wall deposition. Of the eight other genes (the T-DNA insertion line of AtMYB63/At1g79180 was not available) that did not give a mutant phenotype, the expression patterns determined by RT-PCR expression analysis suggest that they are preferentially expressed in mature stems and the root. The promoter-GUS expression pattern of MYB63, with high expression in protoxylem cells undergoing secondary wall formation, is also consistent with a role in secondary cell wall formation. There are several possibilities as to why mutant phenotypes were not observed in knock-out lines for these genes. One possibility is that they do not play a role in secondary wall, xylem, or fiber differentiation, and other potential phenotypes were not investigated in detail. Another explanation could be a combination of gene redundancy and a lack of detailed phenotypic analysis. All of the genes investigated are members of gene families, and partial functional redundancy with other gene family members cannot be  89 excluded. This was tested for the pair of MYB paralogs, MYB20 and MYB43, by generation of the double mutant. However, a mutant phenotype was still not evident. With respect to phenotypic analysis, we performed a limited screen of possible phenotypes, and subtler secondary wall phenotypes, for example cell wall polymer composition, would have been missed. For example, a T-DNA insertion in At5g54160 that encodes caffeic acid O-methyltransferase 1 gene (AtOMT1) has been described previously to be without any visible phenotype. However, chemical analysis of the mutant revealed a reduction in syringyl lignin in the vasculature of the plant (Goujon et al., 2003). Thus, biochemical analysis of lignin, cellulose, and hemicellulose content of candidate gene mutant lines could reveal secondary wall phenotypes. In summary, our results on poplar gene expression profiling together with the results in Arabidopsis (Birnbaum et al., 2003; Brown et al., 2005; Ehlting et al., 2005; Persson et al., 2005), highlight four Arabidopsis genes encoding transcription factors MYB63, MYB20, MYB43, KNAT7, together with their putative poplar orthologs (PoptrMYB028/fgenesh4_pg.C_LG_V000361, PoptrMYB018/grail3.0038010201, PoptrMYB152/eugene3.00002261, and PoptrKNAT7/estExt_fgenesh1_pg_v1.C_ LG_I0964), that are differentially expressed in association with secondary wall formation in both species. Significantly, PoptrKNAT7 is also exhibited high expression level in xylem based on analysis of a public poplar microarray database, and the knat7-1 allele shows an irx phenotype in Arabidopsis. The combination of these data suggests that KNAT7 is a good target for further functional analyses as potential regulator of secondary wall development in both species.  90             CHAPTER 4  Arabidopsis KNAT7, a regulator of secondary wall formation   91 Chapter 4. Arabidopsis KNAT7, a regulator of secondary wall formation  4.1 Introduction Secondary walls are the major component of wood, and are formed during differentiation of specialized cells types such as vessel elements, tracheids and fibers in vascular plants. Secondary walls are composed mainly of cellulose, lignin, and hemicelluloses and their formation is a crucial developmental process in vascular plants. During secondary wall formation, the biosynthesis of these cell wall components is highly coordinated (Turner and Somerville, 1997; Donaldson, 2001; Zhong and Ye, 2007). Although a number of genes involved in the biosynthesis of secondary wall components have been characterized, the regulation of these genes is still poorly understood. (Brown et al., 2005; Persson et al., 2005; Zhong and Ye, 2007). Genomic studies in both Populus and Arabidopsis have been used to identify the regulatory genes expressed in coordination with secondary wall formation. Ten candidate transcription factor genes were shown to be differentially expressed during fiber differentiation in Arabidopsis inflorescence stems (Ehlting et al., 2005) and in the developing stele of Arabidopsis roots (Birnbaum et al., 2003). Moreover, the poplar orthologs of a subset of these genes were associated with secondary xylem formation in two different experiments (Chapter 3). These data provide a small set of potential candidate genes that may play important roles in the regulation of secondary xylem formation. Due to its strong secondary cell wall-associated expression pattern and loss of function irx phenotype in Arabidopsis (Brown et al., 2005; Chapter 3), KNAT7 is a strong candidate for a regulator of secondary wall formation in both species. The KNOX family in Arabidopsis contains the STM gene and seven KNAT (KNOTTED1-like homeodomain protein) genes and can be divided into two classes based on phylogenetic analysis (Dean et al., 2004; Truernit et al., 2006). KNAT7 falls within the poorly characterized Class II clade, with some members that may play roles in regulation the development of the root (Birnbaum et al., 2003; Truernit et al., 2006). Further studies  92 demonstrated that Medicago truncatula KNOX4 could be the putative orthologue of the Arabidopsis KNAT7 (Di Giacomo et al., 2008), however further analysis is needed to assess the similarities of Medicago KNOX4 to KNAT7. In a previous study that combined expression data with reverse genetics, KNAT7 (designated IRX11) was classified as an IRREGULAR XYLEM (IRX) gene based on a clear collapsed xylem (irx) phenotype characteristic of mutants with a secondary cell wall defect (Brown et al., 2005). In this chapter, I more fully characterized the knat7 mutant phenotype. In accordance with previous results (Chapter 3), KNAT7 appears to be a key transcriptional regulator associated with secondary wall formation. The mutant phenotypes associated with independent knat7 loss of function alleles include both irx morphology and thicker interfascicular fiber walls. Furthermore, knat7 mutants display changes in seed coat mucilage. Overexpression of KNAT7 results in a reduction of cell wall thickness in interfascicular fibers. We show that PoptrKNAT7 as well as Arabidopsis KNAT7 can complement the Arabidopsis knat7 mutant phenotypes, that both proteins are nuclear localized, and that expression of lignin biosynthetic genes is directly or indirectly regulated by KNAT7. Together, the results indicate that KNAT7 is a key homeodomain transcription regulator involved in secondary wall formation in both Populus and Arabidopsis.  4.2 Results  4.2.1 Identity of SALK T-DNA insertion alleles In order to analyze the function of Arabidopsis KNAT7, we used a reverse genetics approach to isolate and characterize loss-of function KNAT7 mutant alleles. Five independent T-DNA insertion mutant alleles of AtKNAT7 (At1g62990) were identified using the SIGnAL database (http://signal.salk.edu/cgi-bin/tdnaexpress), including SALK_002098 (knat7-1), WiscDsLox367A5 (knat7-2), SALK_110899 (knat7-3), SAIL_757 (knat7-4), and WiscDsLox461 (knat7-5). Seed stocks were obtained, and T-DNA insertions were confirmed by PCR with T-DNA border flanking sequences primers. After generation of plants homozygous for the different alleles, the T-DNA insertion sites were confirmed by sequencing of T-DNA-genomic DNA junctions. Except insertions in  93 knat7-2 and knat7-3, located at the second intron of AtKNAT7, the T-DNA insertions in other alleles including knat7-1 were located in the fourth intron, and the relative positions of the T-DNAs in these five alleles are shown in Figure 4.1A and 4.2A. The results from RT-PCR analysis using a primer pair that amplifies the full-length coding sequence indicated that the full transcript of KNAT7 was undetectable in plants homozygous for the knat7-1 allele (Figure 4.1B) and other alleles (Figure 4.2B). For knat7-1, a second primer pair 5’ to the T-DNA insertion (CDSAtKNAT7F and T2098-1; Table 2.2) also failed to detect mRNA transcript, this suggests that these five alleles are probably loss-of-function mutant alleles of KNAT7. Like knat7-1, all other plants homozygous for additional knat7 alleles showed an irx phenotype (Figure 4.2C), This irx phenotype, defined on the basis of collapsed vessels elements, is obvious in all vascular bundles and plants examined and always resulted in severely irregular vessels as previously described for other irx mutants (Turner and Somerville, 1997; Brown et al., 2005). While the knat7 (irx11) irx phenotype was clear (Figure 4.2C), it was less severe than that observed for irx3, in which vessels are irregular and frequently collapsed inwards such that some appear almost completely occluded (Brown et al., 2005). Because knat7-1 was the first well-characterized allele, knat7-1 homozygotes segregated at an approximately 1:3 ratio (12:62, n = 74; chi-square indicated no significant difference from the expected 1:3 ratio; http://www.graphpad.com/quickcalcs/chisquared1.cfm) when backcrossed to wild type Col-0 indicating the presence of a single T-DNA locus in knat7-1, and other knat7 alleles had a phenotype similar to that of knat7-1, knat7-1 was used for most of the further phenotypic analyses.  94     Figure 4.1 Characterization of the knat7-1 allele.  (A) Schematic diagram of KNAT7 gene structure and the position of the T-DNA in knat7-1 (SALK-002098). Numbers are nucleotide positions relative to start and stop codons; boxes represent exons; arrows present primer pair (CDSAtKNAT7F/R; Table 2.2) used in RT-PCR. (B) RT-PCR analysis of KNAT7 expression in knat7-1 and in myb61-2 (SALK_106556) (see below Section 4.2.6). No transcript was detected in the knat7-1 mutant, but a reduction in transcript levels was observed in the myb61-1 mutant. ACTIN2 was used as a positive control.   95  (C)    Figure 4.2 Additional KNAT7 T-DNA insertion alleles and their mutant phenotypes in stems.  (A) Predicted T-DNA insertion sites in a schematic diagram of the KNAT7 gene. T-DNA  96 insertion sites in the WiscDsLox367A5 (knat7-2), SALK_110899 (knat7-3), SAIL_757 (knat7-4), and WiscDsLox461 (knat7-5) lines are shown. Arrows present primers used in RT-PCR analyses. The relative positions of T-DNA insertions are based on sequence analysis of T-DNA-genomic DNA junctions. (B) RT-PCR analysis of KNAT7 expression in knat7-2, knat7-3, knat7-4 and knat7-5 mutants showing undetectable levels of transcripts. ACTIN2 was used as positive control. (C) Hand sections from wild type (a) and knat7 mutant plants (b-f) taken from the bases of 8-week old inflorescence stems and stained with toluidine blue. (b) SALK_002098 (knat7-1), (c) WiscDsLox367A5 (knat7-2), (d) SALK_110899 (knat7-3), (e) SAIL_757 (knat7-4), and (f) WiscDsLox461 (knat7-5). Red arrows indicate the irx phenotypes in each mutant. Bars 20 µm.  97 4.2.2 Phenotypic characterization of knat7 mutants Histochemical analysis of GUS activity in an Arabidopsis line expressing a KNAT7 promoter-GUS fusion, as shown in Figure 4.3 indicates strong activity of the KNAT7 promoter in the vascular system of seedling roots, hypocotyls, and cotyledon veins, where secondary walls of the primary xylem differentiate (Figure 4.3). Despite this pattern of expression, no defects in root or shoot development or growth were observed in knat7 mutant seedlings. As previously described (Brown et al., 2005), mature (6-week old) knat7 plants grown under long day conditions (16 h light/8 h dark) appeared morphologically normal. In order to view in detail and at high resolution the anatomy and morphology of developing inflorescence stems from 6-week old plants, thin sections of embedded wild type and knat7-1 mutant plants were generated. As previously documented (Brown et al., 2005), knat7/irx11 xylem vessels at the base of inflorescence stems exhibit a collapsed morphology (Figure 4.4C, D), relative to the normally round shapes of such vessels in wild type plants (Figure 4.4A, B), suggesting defect(s) in secondary wall deposition or composition. In addition to the xylem vessel irx phenotype, however, knat7-1 mutant xylary fibers also appeared to have a collapsed morphology (Figure 4.4D), a phenotype not previously described. Furthermore, comparison of cross sections taken from knat7-1 and wild type plants revealed an apparent increase in the thickness of interfascicular fiber secondary cell walls in the mutant (Figure 4.4D). To observe potential changes in cell wall thickness and morphology at higher resolution, we prepared inflorescence stem cross sections from wild type and knat7-1 mutant plants for transmission electron microscopy. These results, shown in Figure 4.5, confirmed the knat7 phenotypes observed using light microscopy (Figure 4.4 A-D). Both vessel elements and xylary fibers had obvious collapsed morphologies in the knat7 mutant (Figure 4.5, E-H), and the xylary secondary cell walls appeared thinner. In contrast, interfascicular fibers showed no evidence of changes in morphology other than significantly enhanced secondary cell wall thickness. In order to quantify interfascicular fiber, xylary fiber, and vessel cell wall thicknesses, we took measurements from multiple cells in light microscopy images taken from thin sections of wild type and knat7 mutant plants at high magnification. This analysis (Table 4.1) demonstrated a nearly 60% increase in cell wall thickness in the knat7-1 interfascicular  98 fibers relative to wild type, while xylary fiber walls were about 30% thinner than in wild type. Despite the collapsed vessel phenotype of the knat7-1 mutant, vessel cell wall thickness showed little if any change in the mutant relative to wild type. Re-examination of stem cross sections of plants homozygous for additional knat7 alleles (Figure 4.2C) revealed that they also exhibited thicker interfascicular fiber cell walls compared to wild type. Finally, to get a more complete picture of cell morphologies, we prepared longitudinal sections from the bases of wild type and knat7-1 inflorescence stems, focusing on the interfascicular fiber (Figure 4.4 E, G) and vascular bundle (Figure 4.4 F, H) regions of the stem. In these sections, knat7 interfascicular fiber cell walls also appeared thicker, fiber cells appeared to be less regular in organization, and the lengths of longitudinally sectioned fibers appeared in general to be shorter. In vascular bundles, knat7-1 xylary fibers also had apparently thicker cell walls. However, spiral secondary cell wall thickenings were apparent in knat7-1 xylem vessels, despite the predominant mutant irx phenotype of the mutant, suggesting that secondary wall formation is qualitatively similar to wild type.  99      Figure 4.3 Histochemical analysis of GUS activity of promoterKNAT7-GUS fusions.  (A) 4-day old seedling, (B) cotyledon of a 4-day old seedling, (C) 10 µm-thick root cross-section from a 4-day-old seedling showing expression in the stele, (D, E) cross-sections of inflorescence stems at 7-9 cm from the apex of an 8-week-old plant. Bars 20 µm.  100  Figure 4.4 Anatomy at the bases of wild type and knat7-1 inflorescence stems.  Thin sections taken from embedded material were stained with toluidine blue and images obtained using an OLYMPUS AX70 light microscope (A-D) cross sections from wild type (A, C) and knat7 (B, D) plants. Higher magnification views in (B) and (D) show sections having thicker interfascicular and xylary fibers, but collapsed vessels with thinner cell walls. (E-H) longitudinal sections from interfascicular fiber regions (E, G) and vascular bundle regions (F, H) from the bases of wild-type (E, F) and knat7 (G, H) inflorescence stems. If, interfascicular fiber; ph, phloem; ve, vessel; xf, xylary fiber. Bars 20 µm.  101   Figure 4.5 Transmission electron microscopy of wild-type and knat7 inflorescence stem fiber and vessel secondary walls.  (A-D) sections from a wild-type plant, (E-F) sections from a knat7 plant. (A, B, E, F) interfascicular fiber walls showing thicker secondary cell walls in knat7 plants. (C, D, G, H) The collapsed vessels with reduced secondary cell wall thickening in vessel xylary fiber cells of knat7. If, interfascicular fiber; ve vessel; xy, xylary fiber. Bars, 2 µm.    Table 4.1 Secondary cell wall thickness in wild type and knat7-1 stems  Genotype  Interfascicular fiber wall thickness (µm)1 Vessel wall thickness  (µm)1 Xylary fiber wall thickness (µm)1 Wild type 0.95 ± 0.17  0.97 ± 0.21 0.61 ± 0.15 Knat7-1 1.50 ± 0.17  0.63 ± 0.15 0.80 ± 0.17 1Data are means ± SD from at least 50 cells measured from light micrograph images taken from the bases of primary inflorescence stems. The quantitative differences were shown to be statistically significant (P<0.0001).  102 4.2.3 knat7 seed coat phenotype The Arabidopsis seed coat is the site of extensive secondary cell wall deposition, including the secretion of abundant pectin-rich mucilage during the later stages of seed coat development. Following mucilage secretion, the epidermal cells synthesize a thick cellulosic secondary cell wall, which is laid down across the apical surface of the cytoplasm (Western et al., 2000; Western, 2006; Young et al., 2008). Thus, we examined knat7 seeds for possible mucilage defects, which can easily be observed following imbibition (Western et al., 2000). In wild type Arabidopsis, upon contact with water, seeds release mucilage from the seed coat epidermis, which swells to form a gelatinous coating over the seed. This can be visualized by staining with ruthenium red, which stains negatively charged biopolymers such as pectin and DNA (Koornneef, 1981). Wild type seeds stained with ruthenium red (in the absence of shaking, which aids in mucilage release) displayed both an outer layer of diffuse staining outside the seeds, and an inner layer, dense capsule of mucilage directly surrounding the seeds as described by Western et al., (2000) (Figure 4.6 E). However, the shaking of wild type seeds in water prior to staining with ruthenium red results in a seed surrounded by only the inner capsule of mucilage (Western et al., 2001) (Figure 4.6 A). When stained with ruthenium red after shaking in water, knat-1 seeds were surrounded by a barely visible layer of unstained mucilage (Figure 4.6 B). However, when knat7 seeds were placed directly in water, then stained without agitation, the inner, dense layer of mucilage was apparent. To determine if the knat7 mucilage defect is due to loss of mucilage after agitation, we stained seeds with India ink, which is a colloidal liquid whose molecules are too large to penetrate the mucilage gel. When knat7 and wild type seeds were shaken (Figure 4.6 C, D), or left without shaking (Figure 4.6 G, H) in water, then placed in India ink, the seeds from both genotypes showed mucilage capsules (Figure 4.6 C, D, G, H), with no apparent difference in knat7 seeds. This indicates that the knat7 mutant mucilage defect results from altered staining properties with respect to ruthenium red following agitation. This experiment was also performed with lines homozygous for other knat7 mutant alleles and similiar phenotypes were observed (data not shown). To further analyze the defect in the knat7 seed coat testa, we examined dry seeds by scanning electron microscopy (SEM). Wild type seeds, when viewing with SEM, exhibited a  103 typical reticulate appearance with the radial cell walls and a raised columella in the center of each epidermal cell (Figure 4.7 A, B). SEM images from knat7-1 mutant seeds revealed that the epidermal cells were the same size and shape compared with wild type, but that the radial cell walls were thicker (Figure 4.7), and the columella was slightly reduced in height (Figure 4.7 C, D) when compared with wild type. Measurement of the epidermal cell walls in wild type and mutant seed coats showed that the walls were almost two-fold thicker in the mutant (Figure 4.7 lower). These data suggest that KNAT7 plays a role in regulating secondary cell wall formation in specialized seed coat epidermal cells, but further work will be necessary to determine if other knat7 mutant alleles have similar phenotype and if so, what is the relationship between reduced mucilage cohesiveness and thicker radial cell walls in the knat7 mutant.  104  Figure 4.6 knat7 seed coat mucilage phenotype  Ruthenium red staining (A, B, E, F) and India ink staining (C, D, G, H) of wild type (A, C, E, E, G) and knat7 mutant (B, D, F, H) seeds with or without shaking. (A) Wild type seeds with shaking. Inner layer of mucilage is stained with ruthenium red. (B) knat7 mutant seeds with shaking. No ruthenium red stained mucilage is apparent. (C) Wild type seeds and (D) knat7 mutant seeds with shaking stained with India ink. A capsule of mucilage surrounds the seeds of both genotypes. (E) Wild type seeds without shaking. Two layers of ruthenium red-stained mucilage are present, an outer (cloudy) and an inner layer (intensely stained). (F) knat7 seeds without shaking. Ruthenium red staining of mucilage appears only with inner layers. (G) Wild type seeds and (H) knat7 seeds stained with India ink without shaking. The capsules of mucilage is in both genotypes appear the same. Scale bar = 0.50 mm.  105   Figure 4.7 Analysis of wild type and knat7 mutant seed coat epidermal cells.  Upper panel (A, B) Scanning electron micrographs of a wild type seed showing radial cell walls and central columella in each cell. (C, D) Scanning electron micrographs of a knat7 mutant seed showing thickened radial cell walls and slightly flattened columella. Bars = 10  µm Lower panel. Seed coat epidermal radial cell wall thickness of wild type and the knat7 mutant. Standard deviation was calculated from measurements of 30 cells from SEM images.  106  4.2.4 Over-expression of KNAT7 To further investigate KNAT7 functions in the regulation of secondary wall formation, we next studied the effects of overexpressing KNAT7. Two overexpression constructs were made: double-35S:KNAT7, and prom4CL1:KNAT7. prom4CL1:KNAT7 was generated by fusing the promoter of the parsley 4CL1 gene, known to direct expression of lignifying cells (Hauffe et al., 1991), to KNAT7. Both constructs were transferred to Arabidopsis by Agrobacterium-meditated transformation, and phenotypes related to stem development assayed by observation of growth and development of T2 lines relative to wild type plants, and examination of inflorescence stem cross sections. Among at least three lines examined, there were no obvious phenotypes in the double-35S:KNAT7 expressing lines. However, in at least five individuals from each of two different prom4CL1:KNAT7 transgenic lines a pronounced anatomical phenotype was observed relative to wild type (Figure 4.8). Plants of both prom4CL1:KNAT7 transgenic lines showed wild type morphology, but examination of stem sections showed an apparent decrease in the secondary wall thickness of interfascicular fibers compared with the wild type (Figure 4.8B, D), which was supported by wall measurement from hand sections (wild type 1.84 ± 0.29 µm vs. Prom4CL1:KNAT7 0.80 ± 0.21 µm). No dramatic changes in thickness were observed in the secondary walls of xylary fibers and vessels (Figure 4.8A, C). This phenotype is the opposite that of KNAT7 loss of function mutants, which have increases in interfascicular fiber wall thickness, and is consistent with a role for KNAT7 in regulation of interfascicular fiber secondary wall properties.  107      Figure 4.8 Effects of overexpression of KNAT7 on interfascicular fiber secondary wall thickening. Prom4CL1:KNAT7 was transferred to Arabidopsis plants to overexpress KNAT7. Cross sections of the basal internodes of 8-week-old transgenic plants were examined for alterations in secondary wall thickness. (A, B) Cross sections of wild type stems, showing vascular bundles with vessels, xylary fibers (A) and interfascicular fibers (B). (C, D) Cross sections of a stem from a prom4CL1:KNAT7 overexpression line, and showing decreased secondary wall thickening in interfascicular fibers (D) but no obvious differences in wall thickness of xylary fibers and vessels (C). Stem sections were stained (blue) with toluidine blue and images obtained using an OLYMPUS AX70 light microscope. Similar results were obtained from multiple cross sections from the same Prom4CL1:KNAT7 line, and from a second independent Prom4CL1:KNAT7 line. if, interfascicular fiber; ve, vessel; xf, xylary fiber. Bars 40 µm.  108 4.2.5 Subcellular localization of AtKNAT7 and PoptrKNAT7 and complementation analysis To provide further evidence that the knat7 mutant phenotypes are caused by a lesion in the Arabidopsis KNAT7 gene, a GFP-tagged KNAT7 construct was generated and used to test complementation of the knat7 phenotypes. A genomic fragment containing the KNAT7 coding sequence was fused at its C terminus to GFP, placed under the control of the 35S promoter. The 35S::GFP-KNAT7 construct was then transformed into the knat7-1 mutant background by Agrobacterium mediated transformation (Figure 4.9A). Several transgenic lines were examined for the ability of this construct to complement the mutant phenotype. Six out of ten transformed lines examined were rescued and showed wild type-like phenotypes in stem sections (Figure 4.9C shows a representative line). This indicates that the observed knat7 mutant phenotypes are linked to the KNAT7 locus. The ELK domain is conserved amongst all the KNOX proteins including AtKNAT7, and is proposed to be involved in KNOX protein nuclear import (Meisel and Lam, 1996). To determine if KNAT7 is indeed nuclear-localized, the root of a GFP-KNAT7 expressing line was examined for GFP fluorescence using confocal microscopy. This analysis showed that GFP fluorescence in the root was restricted to the nucleus (Figure 4.9 B), indicating that KNAT7 is a nuclear localized transcription factor.  109     Figure 4.9 Complementation of the Arabidopsis knat7 phenotype with 35S::GFP-AtKNAT7 and subcellular localization of GFP-AtKNAT7.  (A) Schematic representation of the 35S::GFP-AtKNAT7 construct and transformation into the Atknat7-1 mutant line. (B) Laser confocal images from Arabidopsis Atknat7 roots expressing 35S::GFP-AtKNAT7. Cell walls were stained with propidium iodide (left). Monitoring of GFP fluorescence showed that AtKNAT7 is nuclear localized (middle). The two confocal images from different channels were merged (right). Scale bar: 20 µm. (C) Cross sections from the bases of inflorescence stems taken from one representative transgenic line that recovered phenotypes (3) compared to wild type (Col-0) (1) and the Atknat7 mutant (2). They were stained with toluidine blue (1 and 2) or phloroglucinol (3). Scale bar: 20 µm  110 To establish whether PoptrKNAT7 can replace the AtKNAT7 function, as predicted if they are orthologs, and to establish the subcellular localization of PoptrKNAT7, we prepared an N terminal fusion of GFP to PoptrKNAT7 and placed it under control of the 35S promoter to generate 35S:PoptrKNAT7-GFP. This gene was introduced into the knat7-1 mutant background by Agrobacterium-mediated transformation, and T1 lines expressing different levels of the 35S::PoptrKNAT-GFP mRNA identified by RT-PCR using PoptrKNAT7-specific primers. Figure 4.10A shows the expression levels in four representative lines, with line 3 showing the highest expression level, and expression barely detectable in line 1. T2 plants from the four lines were identified that were homozygous for knat7-1 and contained the 35S::PoptrKNAT7- GFP transgene. Phenotypic analysis of mature parts of inflorescence stems taken from such individuals from lines 1 and 3 are shown in Figure 4.10B. While no rescue of the knat7 irx phenotype was observed in line 1, the irx phenotype was abolished in line 3, which expresses the transgene at higher levels. Thus, PoptrKNAT7 appears to be capable of replacing the AtKNAT7 function, suggesting that the gene is the true ortholog. Furthermore, the PoptrKNAT7-GFP-fusion retained KNAT7 activity. In order to determine the subcellular localization of PoptrKNAT-GFP, we monitored GFP fluorescence by confocal microscopy in the roots tips of wild type plants transformed with this construct. Figure 4.10C shows that GFP fluorescence was restricted to the nucleus of root cells, indicating that the PoptrKNAT7 transcription factor is nuclear targeted and functions in the nucleus.  111   Figure 4.10 Complementation of knat7 by PoptrKNAT7 and subcellular localization of PoptrKNAT7-GFP.  (A) Expression of 35S::PoptrKNAT7- GFP in the Atknat7 background in 4 independent transgenic lines. Expression was assayed in wild type (Col-0) as a negative control. (B) Stem cross section from the bases of inflorescence stems taken from transgenic lines 1 and 3. v, vessel. Scale bar: 20 µm (C) Laser confocal images from root tips of Arabidopsis Col-0 expressing of 35S::PoptrKNAT7-GFP. Cell walls were stained with propidium iodide (1). GFP fluorescence signal from 35S::PoptrKNAT7- GFP, showing PoptrKNAT7 is nuclear localized (2). Two optical series from two channels were merged to create (3). Scale bar: 10 µm.  112 4.2.6 Gene expression changes in knat7 The formation of thickened and lignified secondary cell walls during xylem and fiber development requires the coordinated regulation of a number of complex metabolic pathways, including those for cellulose, lignin, and hemicellulose (e.g. xylan) biosynthesis and deposition. As a first step towards understanding the biochemical and regulatory networks that may be affected by KNAT7 activity, I targeted two sets of genes for expression analysis in the knat7 background relative to wild type, using RNA isolated from young seedlings. The first set of genes chosen for this analysis (Figure 4.11 A) included a suite of structural genes involved in secondary wall formation and fiber/xylem development (for example, CESA genes involved in secondary cell wall biosynthesis (Taylor et al., 2003) and selected co-expressed IRX genes (Brown et al., 2005; Persson et al., 2005), a xylan synthase (FRA8), as well as known regulators of lignification and xylem or fiber differentiation (MYB61, ATHB8, NST2, IFL1/REV, MYB32) (Zhong and Ye, 1999; Baima et al., 2001; Newman et al., 2004; Preston et al., 2004; Zhong et al., 2005). Phenylpropanoid and lignin biosynthetic genes (PAL2, 4CL1, C4H, COMT, CCR and CCoAOMT) are known to be up-regulated along fiber development and/or secondary wall formation and lignification (Ehlting et al., 2005), and all of them and their orthologs from other species (eg. poplar and rice) are likely to have actual enzyme functions associated with monolignol biosynthesis (Raes et al., 2003; Hamberger et al., 2007). In general, expression of genes involved in secondary cell wall cellulose biosynthesis was not strongly affected by loss of KNAT7 function. However, IRX12 (which encodes a laccase) (Brown et al., 2005) expression was about 1.5 fold above wild type levels, while IRX10 (Brown et al., 2008; Wu et al., 2008), which encodes a glycosyl transferase of unknown specific function, expression was reduced by about half. In contrast, expression of a suite of genes encoding lignin biosynthetic enzymes (PAL2, 4CL1, C4H, CCoAOMT, COMT, CCR) was down regulated by at least 50% in the mutant, suggesting that a primary function of KNAT7 may be to directly or indirectly regulate genes in this pathway. The BIFUNCTIONAL NUCLEASE 1 (BFN1) gene, shown to be associated with programmed cell death during xylem differentiation (Ito and Fukuda, 2002) was the most highly down regulated gene, suggesting some connection of KNAT7 function to the last stages of fiber  113 and xylem differentiation. Expression of the ATHB8 HD-ZIP regulatory gene was not affected by the knat7 mutation, while modest to pronounced decreases in expression of the NST2 and MYB32 regulatory genes were observed. We also profiled the expression of ten candidate transcription factors identified from gene expression profiling. Figure 4.11B shows, as expected, that KNAT7/At1g62990 transcripts were greatly reduced in the knat7 background. In the knat7 background, dramatic reductions in expression of two MYB candidate genes, MYB20 and MYB63 were also observed. Reduction in expression of the remainder of the genes was modest. Finally, we conducted a survey of publicly available microarray gene expression data to determine if KNAT7 expression is altered in any known Arabidopsis mutants or mis-expression lines, using BAR (Winter et al., 2007a). Interestingly, according to these data, Arabidopsis lines overexpressing MYB61, a known regulator of lignin and mucilage deposition (Penfield et al., 2001; Newman et al., 2004) showed consistent up-regulation of KNAT7 expression. My expression data suggest that MYB61 expression was not affected in knat7 (Figure 4.11A). This indicates that MYB61 could act upstream of KNAT7. To further analyze this potential regulatory interaction, we assayed KNAT7 expression in the myb61-2 (SALK_106556) mutant background by semi-quantitative RT-PCR. Figure 4.1B shows that KNAT7 expression was significantly down regulated in myb61-2 seedlings. This suggests that KNAT7 could be a direct or indirect downstream target of MYB61 in lignin and mucilage formation.  114  Figure 4.11 Quantitative real-time RT-PCR analysis of gene expression in knat7 seedlings relative to wild type seedlings.  RNA was harvested from 14-day old seedlings for expression analysis. The expression level of each gene in the wild type background was set to 100, and expression in the knat7 background is expressed relative to wild-type (Col-0) expression for each gene. TUB9 (At4g20890) was used as a reference gene. (A) Relative expression levels of secondary wall–related biosynthetic and regulatory genes. Phenylpropanoid genes:  PAL2 (At3g53260) CCoAOMT (At4g34050), 4CL1 (At1g51680), C4H (At2g30490), CCR1 (At1g15950), COMT (At5g54160). CesA and other IRX genes: IRX1/CesA8 (At4g18780), IRX3/CesA7 (At5g17420), IRX5/CesA4 (At5g44030), IRX8 (At5g54690), IRX10 (At1g27440), IRX12 (At2g38080), IRX13 (At5g03170). Xyloglucan synthesis: FRA8  (At2g28110). Programmed cell death: BFN1 (At1g11190). Transcriptional regulation: NST2  (At3g61910), ATHB8 (At4g32880), IFL1/REV (At5g60690), MYB32 (At4g34990), MYB61 (At1g09540). (B) Relative expression levels of ten transcription factor encoding genes identified as candidates for regulating secondary cell wall deposition (Ehlting et al., 2005).  115 4.3 Discussion The regulation of secondary wall deposition in plants is important to understand due to its importance in wood and fiber formation, which provide the most abundant sources of biomass for potential bioenergy production and other purposes. Arabidopsis inflorescence stems develop cells with massive secondary wall thickening in vessels and fibers, making this an excellent model to study the transcriptional control of secondary cell wall synthesis. Previous studies have suggested that a network of transcription factors is involved in the regulation of secondary wall biosynthesis in Arabidopsis, including the closely related Arabidopsis NAC domain proteins and MYB transcription factors (Zhong and Ye, 2007; Zhong et al., 2008). Our findings that Arabidopsis KNAT7 and its poplar orthologue PoptrKNAT7 are required to regulate secondary wall formation highlight this transcription factor as an important component of the complex network that regulates secondary wall synthesis.  KNAT7 expression pattern We have demonstrated that KNAT7 encodes a transcription factor involved in the regulation of secondary wall formation in Arabidopsis, because KNAT7 loss-of function mutants exhibit irx morphology in vessels as well as thicker interfascicular fiber cell walls. Consistent with this function, data from GUS expression of promKNAT7-GUS fusions expressed in transgenic Arabidopsis suggests that KNAT7 is specifically expressed in the vascular system in roots and shoots of seedlings (Figure 4.3A), and in the stele of roots (Figure 4.3C). In inflorescence stems where the mutant phenotype is visible, however, PromKNAT7-GUS expression is mainly evident in cortex cells adjacent to interfascicular fibers and cambial regions near xylem in stems, although expression is also associated with the protoxylem in vascular bundles (Figure 4.3D, E). Thus, the expression data is not fully consistent with the mutant phenotypes. One possible explanation for this discrepancy is that the KNAT7 gene is expressed in different cells from the site of protein accumulation, for which there is precedence. For example, previous studies on KNOX family members including maize KNOTTED1 (KN1) gene, its Arabidopsis (KNAT1/BREVIPEDICELLUS), and rice (OsKN1) orthologs, and the  116 Arabidopsis SHOOTMERISTEMLESS (STM) gene suggest that the activities of KN1, STM and related homeobox proteins are maintained by intercellular trafficking of the proteins to cells adjacent to those in which the genes are expressed, and that trafficking may be required for their normal developmental function (Kim et al., 2003; Kuijt et al., 2004; Winter et al., 2007b). Further investigation demonstrated that the microtubule-associated and viral movement binding protein AtMPB2C interacts with KN1/STM, regulating the cell-to-cell trafficking of these homeodomain proteins through plasmodesmata (Winter et al., 2007b; Bolduc et al., 2008). Thus, PromKNAT7-GUS expression patterns may not indicate the cells in which the KNAT7 protein accumulates, and it could be trafficked, for example from cortex cells to adjacent fiber cells. It is also possible that KNAT7 affects the expression of target genes that themselves act to control events in adjacent cells, for example by cell-cell signaling. A second possibility is that the KNAT7 promoter used to generate the promKNAT7::GUS fusion construct does not contain all the information required to fully replicate the KNAT7 expression pattern since the nearest upstream gene (At1g62981) is located approximate 2.6 kb away from KNAT7, however, we only used the 2 kb upstream fragment for the GUS fusion. Recent in situ hybridization data on KNAT7 shows that KNAT7 mRNA in fact accumulates in interfascicular fiber cells and xylem (Zhong et al., 2008).  KNAT7 regulates secondary wall formation Considering the fact that loss of KNAT7 function not only affects the morphology of vessels, but also causes a dramatic increase in the thickness of interfascicular fiber secondary walls, it appears that secondary wall synthesis in both fibers and in vessels is regulated by KNAT7. This is consistent with the previous characterization of the knat7-1 mutant as an irx mutant defective in secondary wall formation, with collapsed vessels (Brown et al., 2005), although the effect on fiber cell wall dimensions was not described by those authors. Based on the observations that loss-of function of KNAT7 causes an increase in secondary wall thickening (Figure 4.4) and that overexpression of KNAT7 results in a reduction in secondary wall thickening (Figure 4.8), it is tempting to propose that KNAT7 is required for specific aspects of secondary wall biosynthesis in both vessel and fiber cells.  117 Other work has highlighted the potential role played by KNAT7 in xylem and secondary wall formation. In a transcriptional profiling study of genes up-regulated during tracheary element differentiation in cultured Arabidopsis cells, KNAT7 expression was induced more than eight-fold (Kubo et al., 2005). In that study KNAT7 was revealed to be up-regulated during vessel element formation in coordination with NAC-domain proteins VND6 (At5g62380) and VND7 (At1g71930), which are transcription switches for plant metaxylem and protoxylem vessel formation (Kubo et al., 2005). Moreover, a recent study on SND1 (secondary wall-associated NAC domain protein 1, At1g32770) -regulated transcription factors, a dominant repressor KNAT7 mutation was found to reduce secondary wall thickness in both interfascicular fibers and xylary fibers in inflorescence stems (Zhong et al., 2008). These authors did not find the increased fiber cell wall thickness phenotype we observed in knat7 loss of function mutants. However, it is difficult to compare their results to ours, since the dominant negative construct was expressed in a wild type background that could contain residual KNAT7 function, using the 35S promoter, which we found ineffective in generating a KNAT7 overexpression phenotype. Also, since KNAT7 is likely to work with interaction partners in a protein complex (Hackbush et al., 2005; Chapter 5), it is difficult to predict the nature of such complexes that would form with the KNAT7 dominant negative protein, and we consider the knat7 loss of function phenotype to be more definitive. The vessel and fiber cells in Arabidopsis are known to have similar structures to these cells found in the secondary xylem of poplar (Chaffey et al., 2002), and conserved regulatory pathways are thought to be involved in xylem formation in herbaceous plants such as Arabidopsis, as well as in trees (Groover, 2005). Previous work (Chapter 3) showed that putative poplar orthologs of the Arabidopsis candidate secondary cell wall regulators KNAT7, MYB20, MYB43, and MYB63 are differentially expressed in poplar developing secondary xylem. My work now confirms that PoptrKNAT7 is a true orthologue of AtKNAT7 based on the ability of the PoptrKNAT7-GFP fusion to complement Atknat7 loss-of function mutant phenotypes. Our results suggest that KNAT7 is also a key regulator of secondary wall formation in woody plants and could provide tools for the genetic engineering of wood and its derivatives.   118 Role of KNAT7 in secondary wall formation Brown et al. (2005) measured the cellulose content of stem cell wall material in several irx mutants relative to wild type. While a number of irx mutants had significant decreases in cellulose content, knat7 stems did not exhibit a significant decrease, consistent with our findings that expression of genes encoding secondary cell wall CESA subunits is not affected in the knat7 mutant (Figure 4.11A). Similarly, stems of the wild type and the knat7 mutant had similar amounts of noncellulosic sugars including xylose (Brown et al., 2005), characteristic of secondary walls (Turner and Somerville, 1997; Brown et al., 2005), suggesting that KNAT7 does not play a major role in regulating hemicellulose biosynthesis and deposition. Since there is no direct correlation between KNAT7 expression and expression of a number of key genes involved in cellulose biosynthesis, the increase in fiber secondary cell wall thickness may not be due to increases in cellulose biosynthesis, consistent with the findings of Brown et al. (2005) that the cellulose content of knat7 inflorescence stems does not increase relative to wild type. However, our expression profiling was carried out in young seedlings, in which a limited number of cell types may be affected by KNAT7 activity, and in which fiber cells are absent. Thus, definitive judgment of the function of KNAT7 on the regulation of cellulose biosynthesis in inflorescence stems will require further experiments. In contrast to the lack of effect on cellulose biosynthetic genes, there was a strong effect of the knat7 mutation on genes encoding lignin biosynthetic enzymes (Figure 4.11A). Thus, it appears more likely that the effect on secondary wall formation in the knat7 mutant is related to the biosynthesis of this secondary wall polymer. Lignin is derived from the dehydrogenative polymerization of hydroxycinnamyl alcohols (monolignols) (Boerjan et al., 2003). Biosynthesis of monolignols requires enzymes of the phenylpropanoid pathway, and the genes encoding lignin biosynthetic enzymes in Arabidopsis have been well annotated (for example Raes et al., 2003; Ehlting et al, 2005). The expression of Arabidopsis lignin biosynthetic genes PAL2, CCoAOMT, 4CL1, C4H, and CCR1 were decreased dramatically compared with wild type in the knat7 background (Figure 4.11A). Additionally, expression of MYB32, which has been shown to regulate genes in the phenylpropanoid pathway (Preston et al., 2004), was also dramatically decreased.  119 These data suggest that KNAT7 may be capable of regulating a whole set of genes involved in lignin biosynthesis during the course of secondary wall formation, directly or indirectly. Preliminary analyses using histochemical stains for lignin did not reveal any striking defect in lignin deposition in the knat7 mutant (data not shown), but future biochemical analyses should clarify whether lignin content or composition is altered.  KNAT7 is a direct or indirect target of MYB61 and is involved in seed coat mucilage and wall formation Analysis of publicly available gene expression databases indicated that KNAT7 is up-regulated in Arabidopsis lines overexpressing MYB61. Mutations in MYB61 (At1g09540) have been found to specifically affect both mucilage and columella in the Arabidopsis seed coat (Penfield et al., 2001), regulate the closure of stomata (Liang et al., 2005), and regulate lignification (Newman et al., 2004). However, the links between lignification, testa mucilage formation, and changes in stomatal physiology have not been clearly described. Our gene expression data further demonstrate that MYB61 acts upstream of KNAT7, since KNAT7 expression is decreased in the myb61 mutant background (Figure 4.1B). The seed mucilage phenotype we observed in the knat7 mutant is consistent with the role of MYB61 in regulating mucilage production (Penfield et al., 2001). However, our analysis suggests that the knat7 defect results from altered mucilage structure or composition rather than reduction in quantity, since the amount of mucilage observed by India ink exclusion was not affected in knat7, and a difference in staining by Ruthenium red was only observed when seeds were subjected mechanical agitation in water prior to staining. Since ruthenium red staining is dependent on the distance between galacturonic acid residues of homogalacturonan (HG), a constituent of pectin (Sterling, 1970), our data suggest that the mucilage in knat7 mutant has weakened connections within the pectin network that are disrupted by shaking. Pectin exists as a complex with other polysaccharides, including hemicelluloses in the plant secondary cell wall matrix (Western et al., 2001). These results suggest that KNAT7 may be involved in the regulation of mucilage composition, affecting mucilage cohesiveness, and potentially the interaction of pectin with other polysaccharides in secondary walls. It would be interesting to assay changes in expression of genes encoding  120 enzymes involved in pectin biosynthesis in the knat7 mutant (Egelund et al., 2006; Jensen et al., 2008; Mohnen, 2008). Scanning electron microscopy (SEM) of mature Arabidopsis seeds demonstrated that the wild type Arabidopsis seed coat is comprised of five- or six-sided cells with central protrusions, the columella, which develops through active cytoplasmic rearrangement and synthesis of a secondary cell wall, as previously described (Western et al., 2000). Strikingly, the mature seeds of the knat7 mutant exhibit two-fold thicker radial cell walls, as well as slightly reduced columella height (Figure 4.7). The radial cell wall measurements based on SEM in wild type are from the primary cell wall, not secondary cell wall. However, the thicker radial cell wall in the knat7 mutant could result in a secondary cell wall that extends far enough up the primary cell wall to make it thicker at the top of the cell. In addition, columella formation results from the deposition of secondary cell wall material around a cytoplasmic column (Western et al., 2000; McFarlane et al., 2008). To fully characterize the potential effect of the knat7 mutation on secondary wall formation in the seed coat testa, sectioning of seed coats from mature seeds and light microscope or TEM observations would be required. This evidence suggests a possible link between the regulation of mucilage composition and secondary cell wall formation, and suggests that both processes might be regulated by a system involving KNAT7. Thus, in addition to regulating secondary wall formation in stems, KNAT7 appears be part of a MYB61-based transcriptional complex network involved in mucilage and seed coat wall formation. Further in silico analysis of KNAT7 developmental expression using the eFP Browser at the The Bio-Array Resource for Arabidopsis Functional Genomics (BAR; http://bar.utoronto.ca/) shows that KNAT7 expression is significantly up-regulated during the course of seed and silique development, and the levels of expression in developing seeds approach that seen at the base of inflorescence stems (Figure 4.12). This provides further support the function for KNAT7 in seed coat development.  121     Figure 4.12 In silico analysis of relative expression levels of KNAT7 in Arabidopsis.  The data of relative expression level of different tissue types was retrieved from “Developmental map” in BAR (http://bbc.botany.utoronto.ca/.  122 KNAT7 affects expression of secondary cell wall candidate transcription factor genes Several MYB transcription factors have been linked to the regulation of phenylpropanoid metabolism and lignin deposition in Arabidopsis (Newman et al., 2004; Preston et al., 2004). Our data profiling the expression of candidate transcription factors associated with fiber differentiation (Ehlting et al., 2005) indicate that expression of the transcription factor genes MYB20 and MYB63 are especially strongly down regulated in the knat7 mutant background (Figure 4.11B). This result indicates that AtKNAT7 is situated upstream of these two genes in a network regulating secondary wall formation, and or indirectly regulating MYB20 and MYB63. MYB20 and its paralog, MYB43, have been identified by others as being part of the secondary cell wall regulatory network (Brown et al., 2005) downstream of the NAC domain master transcriptional regulators (Zhong et al., 2006; Zhong et al., 2007; Zhong et al., 2008). By contrast, the expression of the well-characterized fiber-associated homeobox gene INTERFASCICULAR FIBERLESS1 (IFL1) and HD-ZIP gene that is involved in the initiation of fiber differentiation (Zhong and Ye, 1999) was less strongly affected in the knat7 mutant background. Similarly, no alternation in the expression of another homeobox gene ATHB8, which is involved in the early stage of xylem differentiation (Baima et al., 2001), or the candidate gene AP2-EREBP/At5g07580 was observed. However the expression levels of the remainder of the candidate regulators identified by Ehlting et al. (2005) were down regulated by approximately 50% in the knat7 mutant background (Figure 4.11B). This suggests that there may be general developmental and/or metabolic changes in the knat7 mutant background that have global effects on gene expression. Future work aimed at identifying direct targets of KNAT7 will aid in determining its role in the network that regulates fiber differentiation.  KNAT7 and the transcriptional network regulating secondary wall biosynthesis A recent series of studies on NAC and MYB domain transcription factors has provided a view of a complex transcriptional network that appears to regulate secondary wall biosynthesis. This network includes a group of closely related Arabidopsis NAC domain proteins that are master regulators of xylem and secondary wall formation NAC secondary wall thickening promoting factor -NST1; NST2; SND1; VND6 and VND7 (Kubo et al.,  123 2005; Zhong et al., 2006; Zhong et al., 2007a; Zhong et al., 2007c; Zhong and Ye, 2007; Zhong et al., 2008) as well as MYB transcription factors and KNAT7. A body of evidence indicates that SND1 is a master transcriptional switch activating the developmental program of secondary wall biosynthesis (Zhong et al., 2008). While SND1 and NST1 function redundantly in the activation of secondary wall biosynthesis in fibers (Zhong et al., 2006; Mitsuda et al., 2007; Zhong et al., 2007b), VND6 and VND7 are proposed to regulate the differentiation of metaxylem and protoxylem, respectively (Kubo et al., 2005). Overexpression of SND1 up-regulates the expression of a group of secondary wall associated transcription factor genes, including MYB20 and KNAT7 (Zhong et al., 2006, 2007c). Furthermore, MYB46 was shown to be a direct target of SND1, and MYB46 overexpression activates the expression of KNAT7 (Zhong et al., 2007a; Zhong and Ye, 2007). The most recent findings demonstrate that KNAT7 is a direct target of SND1 and of its close homologs, NST1, NST2 and vessel-specific VND6 and VND7 (Zhong et al., 2008). Interestingly, in silico analysis of KNAT7 developmental expression shows that KNAT7 expression is also more strongly up-regulated in stamens during flower stage 12, which is when the development of secondary thickening in the anther endothecium occurs, compared to other stages of flowering development (Figure 4.12). This provides further suggestion that KNAT7 has a potential function in regulation of anther endothecium lignification. It is known that NST1 and NST2 are regulated by MYB26/MALE STERILE35 (MS35), and act together in regulating secondary wall biosynthesis in the endothecium of anthers (Yang et al., 2007; Zhong and Ye, 2007). It is apparent that KNAT7 regulates secondary wall biosynthesis in different types of secondary wall–containing cells downstream of certain cell type–specific, functionally redundant NAC genes. Our work on KNAT7 helps to place it in this regulatory network since KNAT7 regulates both interfascicular fiber cells and vessels based on mutant phenotypes in both cell types. This provides additional information on how it might control the gene expression required for xylem vessel and fiber formation, as well as the potential function in endothecium as part of this transcriptional network. Our findings support the hypothesis that KNAT7 is part of a transcription network in the regulation of secondary wall formation, in mucilage biosynthesis and potentially in endothecium lignification, as outlined in Figure 4.13. In this network, we propose that  124 KNAT7 acts as an essential regulator downstream of SND1/NST1, NST2 and VND6/VND7 which directly regulate KNAT7 expression (Zhong et al., 2008). Therefore, upstream of KNAT7, SND1 and NST1 activate the secondary wall biosynthetic program in fibers, and VND6 and VND7 specifically regulate secondary wall biosynthesis in vessels, and NST1 and NST2 regulate secondary thickening in the anther endothecium. Based on our data, KNAT7 may work primarily to regulate the expression of lignin biosynthetic genes in vascular tissue and fibers, and possibly in the endothecium. In addition, MYB20, MYB43 and MYB63 are likely positioned downstream of KNAT7 in the transcriptional network, since we found that MYB63 and MYB20 are down-regulated in the knat7 background. This is consistent with the previous findings that MYB20 is induced by overexpression of SND1 (Zhong et al., 2007a), and that MYB43 and MYB63 are also up-regulated during tracheary element differentiation in cultured Arabidopsis cells (Kubo et al., 2005). Thus, we propose that KNAT7 might regulate secondary wall synthesis partially via these MYB genes as part of the transcriptional network regulating secondary wall biosynthesis (Figure 4.13). Interestingly, in addition to SND1/NST1 and its homologs, the regulator of lignification and seed mucilage deposition MYB61 also appears to be involved in the regulation of KNAT7 and this regulatory node seems to be important in regulating seed mucilage composition, and perhaps seed coat cell wall biosynthesis (Figure 4.12). It is still unclear if this node operates during secondary wall formation in association with xylem and fiber cell differentiation, but the role of MYB61 in the regulation of lignin deposition (Newman et al., 2004) suggests that it could.    125    Figure 4.13 Proposed model of KNAT7 roles in the transcriptional network regulating secondary wall biosynthesis.  Based on our data and data from Zhong et al. (2008), a hierarchy of transcription factors involved in regulation of secondary wall biosynthesis in seed coat, fibers, vessels, and endothecium is shown. The solid arrows indicate a directly regulated target gene. The dashed arrows and question mark show hypothesized affects (direct or indirect) on genes and processes in the biosynthesis of lignin, cellulose and xylan.   126 In conclusion, we have demonstrated that the conserved transcriptional regulator, KNAT7 plays a role in secondary wall formation. Our finding that KNAT7 is required for normal expression of lignin biosynthesis genes and other potential secondary wall-associated transcription factors suggests that KNAT7 coordinates the expression of other transcription factors, which in turn might directly or indirectly regulate lignin biosynthetic genes. Our analyses have revealed that both KNAT7 and its poplar orthologue PoptrKNAT7 are targeted to the nucleus. Other studies also suggest that KNAT7 homologues are present in other plant species, for example in Medicago truncatula (Di Giacomo et al., 2008), indicating that regulation of secondary wall formation by KNAT7 genes might be a common mechanism in plants. Homologues of SND1 and SND1-regulated transcription factors are also found in tree species, such as poplar, pine, eucalyptus, and spruce (Patzlaff et al., 2003a; Karpinska et al., 2004; Goicoechea et al., 2005; Bedon et al., 2007), suggesting that they may also regulate secondary wall biosynthesis during wood formation in trees. Therefore, it is likely that the transcriptional network regulating lignin biosynthesis and secondary wall formation is conserved between herbaceous Arabidopsis and woody tree species. Since secondary wall deposition is important in the formation of biomass such as wood, the identification and characterization of the KNAT7 homologue-mediated transcriptional network in trees could be very important for future tree biotechnological applications.  127             CHAPTER 5  Interaction of Arabidopsis OFP proteins with KNAT7 forms a regulatory KNOX-OVATE complex   128 Chapter 5. Interaction of Arabidopsis OFP proteins with KNAT7 forms a regulatory KNOX-OVATE complex  5.1 Introduction KNAT7 is a 3-aa loop extension (TALE) homeodomain transcription factor, one of seven Arabidopsis KNAT proteins and among four Arabidopsis Class II KNOX paralogs (Kerstetter et al., 1994). Although the functions of class II KNOX genes are poorly understood, KNAT7 has been identified as a candidate regulator of secondary wall formation (Brown et al., 2005; Ehlting et al., 2005; Persson et al., 2005). Recent work has shown that KNAT7 is part of a transcription factor regulatory network required for secondary cell wall biosynthesis (Chapter 3; Zhong et al., 2008). Class I KNOX proteins are well-characterized as developmental regulators, and often work via protein-protein interaction with other KNOX proteins and with BELL and BELL-LIKE (BLH) proteins. For example, BEL1 interacts with SHOOT MERISTEMLESS (STM), BREVIPEDICELLUS (BP)/KNAT1, and KNAT5 to maintain the indeterminacy of the inflorescence meristem (Bellaoui et al., 2001). The physical association of the BELL homeodomain protein PENNYWISE (PNY) and BP/KNAT1 protein suggested the role of this complex in regulating internode patterning in the inflorescence meristem (Smith and Hake, 2003). Two BLH proteins, SAWTOOTH1 (BLH2/SAW1) and SAWTOOTH2 (BLH4/SAW2) act redundantly to establish leaf shape by repressing expression of BP/KNAT1 or more of the other KNOX genes (Kumar et al., 2007). Many methods to detect protein-protein interactions rely on in vitro affinity-based selections such as the yeast 2-hybrid system (Field and Song, 1989; Stephens and Banting, 2000). However, in vitro selected candidates do not necessarily interact in vivo, since the proteins may not be present in the same cells, or cellular conditions may not be appropriate for complex formation in the context of the original host. Therefore, in vivo methods have been developed based on the two-hybrid approach to study interaction in native host cells (Tiwari et al., 2004; Tiwari et al., 2006b; Wang et al., 2007), or to visualize protein interaction in living cells with fluorescent protein variants (Hu et al., 2002a; Walter et al.,  129 2004). Systematic analysis of protein interactions of Arabidopsis TALE homeodomain proteins using the yeast two-hybrid system revealed a highly connected, complex network of interacting class I and II KNOX, BLH, and ovate family proteins (OFPs) (Hackbusch et al., 2005). OFPs are a novel plant-specific and poorly characterized family of regulatory proteins (Hackbusch et al., 2005). The founding member of the OFP gene family, OVATE previously was found to encode a protein with a putative nuclear localization signal and was identified by mutation in tomato, in which a mutation in the OVATE gene causes the tomato fruit to become pear-shaped instead of round (Liu et al., 2002). OFP proteins in tomato, Arabidopsis, and rice are characterized by a conserved ~70-aa C-terminal OVATE domain (Liu et al., 2002). There are 18 OFP genes in Arabidopsis that encode proteins with the conserved OVATE domain (Figure 5.1), and most members of this family contain a predicted nuclear localization signal but lack recognizable DNA binding domains (Hackbusch et al., 2005; Wang et al., 2007). However, little is known about the functions of OVATE-containing proteins in plants. Only the Arabidopsis OVATE-containing protein OFP1 has been functionally characterized so far. No obvious morphological phenotypes are seen in loss-of-function ofp1 mutants, but a dominant, gain-of-function mutant in which OFP1 is overexpressed displayed reduced lengths in all Arabidopsis aerial organs including hypocotyl, rosette leaf, cauline leaf, inflorescence stem, floral organs and silique. OFP1 is an active transcriptional repressor and plays a role in regulating cell elongation by controlling the expression of the gene encoding a key enzyme in gibberellin biosynthesis, GA20ox1 (Wang et al., 2007). Another study on Arabidopsis OFP5 could not detect any ofp5 homozygous plants, suggesting that OFP5 might be required for essential processes in gametophyte development, consistent with the proposed role for OFP5 as a key negative regulator of BELL-KNOX (BLH1-KNAT3) activity during early embryo sac development in Arabidopsis (Pagnussat et al., 2007). Hackbusch et al. (2005) found that the conserved OVATE domain mediates the interaction of OFPs, with the homeodomains of both BELL and KNOX proteins (Hackbusch et al., 2005). KNAT7 was shown to interact with BELL family members BLH5, BLH7; KNOX family members STM, KNAT1, KNAT4, KNAT5, KNAT6 and OFP family members OFP1,  130 OFP2, OFP3, OFP4, OFP6 as summarized in Table 5.1. A BLH1-KNAT3-OFP5 protein complex is essential for Arabidopsis embryo development (Pagnussat et al., 2007), and KNAT-BLH-OVATE complexes may play other yet undiscovered regulatory roles in plants. Here, we report the functional characterization of Arabidopsis OFP4, an OFP family member relatively closely related to OFP1. We demonstrate that OFP1 and OFP4 interact with KNAT7 in vivo and that both OFP4 and KNAT7 are transcriptional repressors. We further show that the pleiotropic OFP1 and OFP4 overexpression phenotypes depends upon KNAT7 function, and that an ofp4 mutant and ofp1/ofp4 double mutant have irx phenotype similar to that of knat7. These results indicate that OFP4 plays a role in regulating secondary cell wall formation through its interaction with KNAT7, and that KNAT7 and OFP4 form a regulatory complex important in secondary cell wall biosynthesis.  131     Figure 5.1 Phylogenetic analysis of the Arabidopsis thaliana ovate family protein (OFP) gene family.  The protein sequences of 18 members of the OFP family were retrieved from the Arabidopsis genome. A multiple sequence alignment and dendrogram were generated by Dr. Jin-Gui Chen (UBC) using CLUSTAL W (1.83) (http://clustalw.genome.jp). The red box highlights the closely related OFP1 and OFP4 genes.  132  Table 5.1 A summary of interaction partners of KNAT7 identified from Hackbusch et al. (2005).  Locus Gene Description Expression information 1 At2g27220 BLH5  BEL1-like homeodomain 5 Up-reguated At2g16400 BLH7  BEL1-like homeodomain 7 No change At3g47730 ATH1  BEL1-like homeodomain 1 No change At4g08150  KNAT1  homeobox protein knotted-1 like 1 Not on array At5g11060 KNAT4  homeobox protein knotted-1 like 4 Up-regulated from top to 3-5cm, then down-regulated At4g32040 KNAT5  homeobox protein knotted-1 like 5 No change At1g23380 KNAT6  homeobox protein knotted-1 like 6 Down-regulated At1g62360 STM  homeobox protein SHOOT MERISTEMLESS No change At5g01840 AtOFP1  ovate family protein 1  Up-regulated  At2g30400 AtOFP2  ovate family protein 2  No change At5g58360 AtOFP3  ovate family protein 3  No change At1g06920 AtOFP4  ovate family protein 4  Slightly change At3g52525 AtOFP6  ovate family protein 6  Not on array At1g11910  aspartyl protease family protein  Up-regulated 1 indicates the differential expression information of each gene observed during inflorescence stem maturation, based on microarray expression profiling from the top 0-3 cm, to the bottom 7-9 cm of stems (Ehlting et al., 2005).  133 5.2 Results 5.2.1 KNAT7 interacts with OFP1 and OFP4 in vivo Investigation of a KNAT-BELL-OFP protein interaction network by yeast two-hybrid analysis showed the potential for KNAT7 to interact with a number of partner proteins including OFP1, OFP2, OFP3, OFP4, and OFP6 (Hackbusch et al. 2005) (Table 5.1). Review of microarray data over the course of inflorescence stem development (Ehlting et al. 2005) showed that OFP1 is differentially regulated, with strongest expression in the oldest part of the stem (7-9 cm; up-reglated), suggesting that it could work together with KNAT7 to regulate secondary wall formation (Table 5.1). Among the 18 Arabidopsis OFP family members, OFP2, OFP3, and OFP4 are the most closely related to OFP1, but insertion mutants were only available for OFP1 and OFP4. Furthermore, while OFP4 is slightly up-regulated over the course of inflorescence stem development (Ehlting et al., 2005), OFP2 and OFP3 show no change (Table 5.1). Thus, we chose to focus on OFP1 and OFP4 as potential interaction partners with KNAT7 in the control of secondary wall formation. To demonstrate and confirm OFP-KNAT7 protein-protein interactions in vivo, we applied the biomolecular fluorescence complementation technique (BiFC) (Hu et al., 2002b; Tzfira et al., 2005; Shyu et al., 2006). OFP1 and OFP4 were fused to a C-terminal (C-EYFP) and KNAT7 to an N-terminal (N-EYFP) fragment of the enhanced yellow fluorescent protein (EYFP), neither of which is capable of fluorescence alone. Using an Arabidopsis leaf mesophyll protoplast transient expression system (Tiwari et al., 2006b), we transformed different combinations of fusion constructs. Fusions of OFP1, OFP4 and KNAT7 to a complete EYFP generated fluorescence localized to the nuclei of transformed protoplasts (Figure 5.2 A-C), consistent with the results reported by Wang et al. (2007) on nuclear localization of OFP1, and the results of Zhong et al. (2008) on KNAT7 nuclear localization. Co-transformation of truncated EYFP fusions to these genes revealed that co-expression of OFP1:C-EYFP with KNAT7:N-EYFP, as well as OFP4:C-EYFP with KNAT7:N-EYFP generated nuclear localized fluorescence (Figure 5.2D, E). However, no fluorescence was detected when either of the KNAT7:N-EYFP or OFPs:C-EYFP constructs was co-expressed with RACK1:C-EYFP and RACK1:N-EYFP, a non-interacting protein (RACK1, Receptor for Activated C Kinase 1; (Chen et al., 2006; Guo and Chen, 2008) that served as a negative  134 control (Figure 5.2F; data not shown). These data show both OFP1 and OFP4 can interact with KNAT7 to form a protein complex in vivo.     Figure 5.2 Bimolecular fluorescence complementation analysis of OFP1, OFP4 and KNAT7 interactions in vivo.  Proteins fused to the full EYFP (A,B,C) (F-EYFP) and N- or C-terminal fragments (D,E,F) (N-EYFP or C-EYFP) were coexpressed transiently in protoplasts. Yellow fluorescence indicating reconstitution of YFP was analyzed by confocal laser scanning microscopy. (A) A protoplast transfected with 35S:KNAT7-F-EYFP. Left: DIC image; middle: image with YFP channel; right, merged images. (B) Images of a protoplast transfected with 35S:OFP1-F-EYFP. (C) Images of a protoplast transfected with 35S:OFP4-F-EYFP. (D) Images of coexpressing OFP1-C-EYFP and KNAT7-N-EYFP, yellow fluorescence indicates that the proteins interact in vivo in the nucleus. (E) Images of coexpressing OFP4-C-EYFP and KNAT7-N-EYFP indicate that the proteins interact in vivo in the nucleus as well. (F) Images of coexpressing the construct KNAT7-N-EYFP with negative control RACK1-C-EYFP (construct from Chen lab, unpublished) revealed no fluorescence.  135 5.2.2 KNAT7 and OFPs are transcriptional repressors Previous data on OFP1 protein indicated that OFP1 functions as a transcriptional repressor (Wang et al., 2007). To test the possibility that OFP4, a closely related paralog of OFP1 in the OFP gene family (Figure 5.1) could also function as a transcriptional repressor, we employed the protoplast transfection system previously used to demonstrate the transcriptional repression function of OFP1 (Wang et al., 2007). In this system, illustrated in Figure 5.3A, a ß-glucuronidase (GUS) gene under the control of both LexA and Gal4 DNA binding sites (LexA[2X]-Gal4[2X]:GUS reporter gene) is used as a reporter, and cotransfected with two effector plasmids. The first contains a chimeric protein consisting of the LexA DNA binding domain (DBD) fused to the Herpes simplex virus VP16 transcription activation domain (LD-VP16), driven by the CaMV 35S promoter. A second effector plasmid contains a chimeric gene encoding a protein consisting of the Saccharomyces cerevisiae Gal4 DBD fused to the gene of interest, for example OFP4 (GD-OFP4), also under control of the CaMV 35S promoter. A control second effector plasmid contains a 35S-driven GD domain only. Co-transfection of the LD-VP16 transactivator gene and the effector gene encoding only the Gal4 DBD (GD) resulted in strong activation of the GUS reporter gene, measured by GUS enzyme activity (Figure 5.3A). Co-transfection of LD-VP16 with either GD–OFP1 or GD–OFP4 effectors resulted in a nearly complete repression of GUS activity expression of the reporter gene (Figure 5.3A), demonstrating that, like OFP1 (Wang et al., 2007), OFP4 functions as a strong transcriptional repressor. Since KNAT7 interacts with OFP1 and OFP4 in vitro (Hackbusch et al., 2005) and in vivo (Figure 5.2), we tested KNAT7 for similar transcriptional repression activity using same protoplast transfection experimental design. Figure 5.3B shows that co-transfection of the GUS reporter plasmid with the LD-VP16 effector gene plasmid and a third plasmid containing the GD domain alone resulted in strong activation of GUS expression, as expected. However, co-transfection with a plasmid containing a GD-KNAT7 fusion (GD–KNAT7) resulted in a strong repression of the expression of GUS activity, relative to protoplasts transfected with the GD control. These results indicate that both OFP4 and KNAT7 can repress gene expression when targeted to cis-acting elements upstream of the target gene.  136 To determine if in vivo interactions between KNAT7 and OFP1 and/or OFP4 affect KNAT7 transcriptional activity, we again used the protoplast transfection system described above. We fused both OFP1 and OFP4 to an N-terminal HA tag and placed the gene under the control of the CaMV 35S promoter (HA-OFPs, Figure 5.3B). A plasmid containing this fusion was then co-transfected with the GUS reporter plasmid [LexA(2x)-Gal4(2x):GUS], an effector plasmid containing the activator fusion gene (LD-VP16), and a second effector plasmid containing the GD-KNAT7 fusion protein (GD–KNAT7). Since OFP1 and OFP4 lack predicted DNA binding domains, any effect of their co-transfection on GUS activity would require OFP-KNAT7 protein-protein interaction. Figure 5.3B shows that both OFP1 and OFP4 enhanced the repression of LexA(2x)-Gal4(2x):GUS reporter gene expression compared to KNAT7 repression activity alone. This suggests that both OFP1 and OFP4 bind to KNAT7 in vivo in this protoplast transient assay system, which is consistent with the biomolecular fluorescence complementation results (Figure 5.2D, E) and the in vitro interactions identified by Hackbusch et al. (2005). It also suggests that the KNAT7-OFP complex acts as a strong transcriptional repressor.  137  Figure 5.3 OFP4 and KNAT7 are transcriptional repressors that interact in vivo.  A protoplast transfection system was used to assay the ability of OFP1, OFP4 and KNAT7 to repress VP16-activated transcription of the GUS reporter gene. (A) Left, effector and reporter constructs used in the transfection assays. Right, GUS activity in protoplasts derived from Arabidopsis rosette leaves co-transfected with an LD-VP16 transactivator plasmid and one of three effector plasmids (GD-OFP1, GD-OFP4, or GD alone) together with the reporter plasmid. The expression of 35S:luciferase (Luc) was used to normalize the expression of the GUS reporter gene. Error bars represent the standard deviation of three replicate transfections. (B) Left, effector and reporter constructs used in the transfection assays. Right, GUS activity in protoplasts derived from Arabidopsis rosette leaves co-transfected with an LD-VP16 transactivator plasmid and different combinations of effector plasmids (GD-OFP1, GD-OFP4, GD-KNAT7, or GD alone) together with the reporter plasmid. The expression of 35S:luciferase (Luc) was used to normalize the expression of the GUS reporter gene. Error bars represent the standard deviation of three replicate transfections.  138 Besides the conserved TALE homeodomain, KNAT7 contains the conserved KNOX (MEINOX) domain, consisting of KNOX1 and KNOX2 sub-domains that are conserved in all KNOX family members (Burglin, 1997). To investigate which KNAT7 domains are required for the transcriptional repression function, KNAT7 effector gene fusions consisting of the GD fused to truncated versions of KNAT7 were generated (Figure 5.4A) and tested for ability to repress of VP16-activated GUS reporter gene expression in protoplast co-transfection assays (Figure 5.4B). First, we fused an N-terminal fragment containing only the KNOX1 to GD (GD–KNAT7-KNOX1). We found that this truncated N-terminal portion of KNAT7 protein failed to repress expression of the reporter gene. However additional effector gene fusions containing either the KNOX2 domain (GD-KNAT7-KNOX2) or the KNOX1+KNOX2 domains (GD-KNAT7-MEINOX) caused significant repression of the VP16-activated transcription (Figure 5.4C), suggesting that the KNOX2 domain within the MEINOX domain is involved in repression. Finally, a fusion of the KNAT7 homeodomain region including the ELK motif to GD (GD–KNAT7-homoedomain) caused repression at a level near to that of the full fragment KNAT7 protein (Figure 5.4C). These results indicate that there are two major transcriptional repression domains within KNAT7, one within the fragment containing the KNOX2 domain and adjacent C-terminal sequences, and one within the C-terminal portion of the protein including the ELK motif and homeodomain.  139     Figure 5.4 KNAT7 transcriptional repression domains.  A protoplast transfection system used to assay for the location of the KNAT7 transcriptional repression domains. (A) Schematic diagram of KNAT7 with four different truncated versions, KNOX1, KNOX2, MEINOX (KNOX1+KNOX2), and Homeodomain. (B) Effector and reporter constructs used in the transfection assays. (C) GUS activity in protoplasts derived from Arabidopsis rosette leaves co-transfected with an LD-VP16 transactivator plasmid and one of the effector plasmids together with the reporter plasmid. The expression of 35S:luciferase (Luc) was used to normalize the expression of the GUS reporter gene. Error bars represent the standard deviation of at least three replicate transfections.  140 5.2.3 OFP1- and OFP4-GUS expression patterns To test the tissue and organ expression patterns of OFP4, Dr. Wang (Chen lab, UBC) generated transgenic plants transformed with a 643-bp genomic fragment upstream of the OFP4 coding region. Since the orientation of OFP4 is opposite that of the adjacent upstream gene At1g06923 in Arabidopsis genome, the two genes share an approximately 1000-bp putative promoter region. Thus, Dr. Wang chose 2/3 of the shared region (643 bp) upstream of the OFP4 start codon as the OFP4 promoter, and fused it to the GUS reporter gene to generate promOFP4:GUS. Four independent transgenic Arabidopsis lines were analyzed for the location of GUS activity using a histochemical assay. Consistent expression patterns were found in all four lines, and representative expression is shown in Figure 5.5. In 7-day-old seedlings (Figure 5.5D), promOFP4:GUS was expressed mainly in the root, at the root-hypocotyl junction, and in cotyledons (Figure 5.5A-C). In roots, expression was especially strong in the vascular cylinder (stele) (Figure 5.5C) and at the root tip (Figure 5.5E), while in cotyledons expression was observed both in veins and other tissues (Figure 5.5A). In mature plants, we assayed promOFP4:GUS activity in cross sections of inflorescence stems. Here, expression was strong in xylem and in cambium cells, as well as in the cortex adjacent to interfascicular fiber cells (Figure 5.5F, G). In accordance with the previously reported tissue and organ expression patterns of the 1383-bp promOFP1:GUS reporter construct (Wang et al., 2007), we found highest levels of promOFP1:GUS expression mainly in the roots of seedlings, especially in the vascular cylinder, of the representative lines assayed (data not shown). We also assayed promOFP1:GUS expression in cross sections of inflorescence stems. Here, GUS activity was detected from xylem, cambium cells and the cortex cells close to interfascicular fibers, in a pattern similar to that of promOFP4:GUS (Figure 5.5H, I). These results demonstrate that the expression of OFP1 and OFP4 is correlated partially with secondary wall formation in interfascicular fibers and the vascular system. The expression patterns of these genes in inflorescence stems is very similar to that of KNAT7, consistent with the hypothesis that in vivo interaction of OFP1 and OFP4 with KNAT7 could help to regulate secondary wall deposition.   141        Figure 5.5 Histochemical assay of OFP4-GUS and OFP1-GUS expression.  (A-E) Expression of promOFP4:GUS in 7-day-old seedlings. (A) cotyledon, (B) junction of hypocotyl and root, (C) root, (D) whole seedling, (E), root tip. (F, G) Expression of promOFP4:GUS in inflorescence stem cross sections. (H, I) Expression of promOFP1:GUS in inflorescence stem cross sections at higher magnification. Bars, 20 µm for A, B, C, E and 40 µm for E, G, H, I.  142 5.2.4 OFP4 is involved in secondary wall formation To further investigate the potential roles of OFP1 and OFP4 in regulating secondary wall formation as interaction partners with KNAT7, we analyzed the phenotypes of OFP1 and OFP4 T-DNA insertion lines. Previous results showed that plants homozygous for a transposon insertion loss-of-function mutant allele of OFP1, ofp1-1 (Figure 5.6A) did not display any obvious morphological phenotypes compared to wild type (Wang et al., 2007), and this was confirmed in rosette-stage ofp1 mutants (Figure 5.6C). We carried out additional phenotypic analyses of the anatomy of cross sections taken from the bases of ofp1-1 inflorescence stems (Figure 5.7B), but no differences in xylem or interfascicular fiber morphology were evident, relative to wild type (Col-0). In order to further analyze the function of OFP4, we characterized loss-of-function mutant alleles of OFP4. The ofp4-1 (SALK_014905) allele used by Pagnussat et al. (2007) contains a T-DNA insertion located in the 3-UTR and was proposed in their study to be a loss-of-function mutant allele of OFP4. However, based on studies in the Chen lab (unpublished data), ofp4-1 is not a loss-of-function allele for OFP4 (data not shown). Moreover, the Chen lab obtained plants homozygous for SALK_022396, an OFP4 allele we designated as ofp4-2, which has a T-DNA insertion located 274 bp downstream of the OFP4 start codon  (Figure 5.6A). RT-PCR analysis confirmed a full transcript of OFP4 was undetectable in the homozygous line, suggesting that ofp4-2 is probably a loss-of-function mutant allele of OFP4 (Figure 5.6B). At the rosette stage, the morphology of ofp4-2 homozygous plants appeared normal compared to wild type Col-0 (Figure 5.6C). However, examination of cross sections taken from the basal part of inflorescence stems revealed an irx phenotype, with vessels in vascular bundles collapsed and irregularly shaped compared to those of the wild type (Figure 5.7A, C). We also measured the wall thickness of ofp4 interfascicular fibers and found that they were slightly thicker than wild type. These data suggest an essential role of OFP4 in secondary wall formation function, similar to that of KNAT7. OFP1 and OFP4 are very closely related among the 18 Arabidopsis ovate family proteins, and functional redundancy has been suggested among family members (Wang et al., 2007). Therefore, the Chen lab generated an ofp1/ofp4 double mutant by crossing ofp1-1 with  143 ofp4-2. I examined the double mutant and found that it had wild type morphology at the rosette stage (Figure 5.6C), but sections from the bases of inflorescence stems showed a collapsed vessel (irx) phenotype and thicker interfascicular fiber cell walls, similar to the single ofp4-2 mutant phenotype (Figure 5.7D).  144    Figure 5.6 Analysis of OFP1 and OFP4 insertion mutants.  (A) Transposon and T-DNA insertion sites in OFP1 (ofp1-1) and OFP4 (ofp4-2), respectively. The ofp1-1 allele has been described (Wang et al., 2007); the ofp4-2 insertion site was confirmed by the Chen lab (personal communication). (B) RT-PCR analysis of OFP4 expression in wild type Col-0; ofp4-2 mutant and 35S:OFP4 plants. ofp4-2 homozygous plants showed no detectable transcripts, while 35S:OFP4 had significantly stronger expression relative to wild type. Actin was used as a control. (C) Morphology of ofp1, ofp4, and ofp1/ofp4 double mutants at the rosette stage compared to wild type (Col-0) (Image from the Chen lab, unpublished).  145   Figure 5.7 Phenotypic characterizations of loss-of-function mutants of OFP1, OFP4 and the ofp1/ofp4 double mutant.  Cross sections of the basal internodes of 6-week-old wild type (Col-0) (A), ofp1 (B), ofp4 (C) ofp1/ofp4 double mutant plants (D). The purple arrows indicate the irx morphology in vessels; black arrow shows the general locations of interfascicular fiber (IFF) cell wall thickness measurements indicated below (n= 40). Bars, 40 µm.  146 5.2.5 Ectopic expression of OFP4 causes abnormal development Previous data showed that plants overexpressing the closely related OFP genes OFP2 and OFP7 showed similar phenotypes to the OFP1 overexpressor, including dwarfism and ovate-shaped organs (Wang et al., 2007). The OFP4 protein not only has a conserved OVATE domain near the C-terminus as other family members, but also has a relatively higher similarity to OFP1 than to other OFP proteins in the Arabidopsis OFP family (Figure 5.1). In addition, OFP4 functions similarly to OFP1 as a transcriptional repressor (Figure 5.3A). To determine if overexpression of OFP4 results in a similar phenotype to OFP1 overexpression, a construct containing the OFP4 open reading frame under the control of the 35S promoter was generated by Dr. Wang (Chen lab, unpublished data) and transferred into wild-type Arabidopsis plants by Agrobacterium-meditated transformation. I germinated the 35S:OFP4 line and characterized its phenotypes. RT-PCR analysis demonstrated that OFP4 transcripts accumulated at high levels in the seedlings of 35S:OFP4 transgenic plants (Figure 5.6B). Transgenic Arabidopsis plants expressing 35S:OFP4 exhibited dominant pleiotropic phenotypes (Figure 5.8) similar to those of 35S:OFP1 (Wang et al., 2007). The seedlings exhibited ovate-shaped cotyledons and curled leaves (Figure 5.8A). In older plants, the prominent visual phenotype was a small rosette size and aberrant growth of leaves with severely curling ovate-shaped blades (Figure 5.8B). In mature plants, stunted inflorescence stems delayed in development were observed, and plants were characterized by irregular ovate-shaped leaves with curved surfaces, thicker and shorter stems, short anthers with little pollen, and uneven-sized short siliques with very few seeds (Figure 5.8C, D). Examination of cross sections taken from the bases of 35S:OFP4 inflorescence stems revealed no dramatic differences from wild type, except delayed interfascicular fiber and vascular bundle differentiation, which was likely associated with the stunted growth of stems (Figure 5.8E, F). Thus, plants overexpressing OFP4 have similar pleiptropic phenotypes to those overexpressing OFP1 as previously described (Hackbusch et al., 2005; Wang et al., 2007) in agreement with the view that OFP proteins including OFP4 may have overlapping functions in the regulation of plant growth, and organ and cell morphogenesis. In contrast, the ofp4-2  147 loss of function mutant phenotype is much more subtle, and appears to be restricted to xylem and interfascicular fiber differentiation, suggesting that OFP4 plays a more specific role in the differentiation of these cells, as a KNAT7 interaction partner.      Figure 5.8 Phenotypes of transgenic plants overexpressing 35S:OFP4 in Arabidopsis.  (A) Ten-day-old wild-type (Col-0) and 35S:OFP4 seedlings. (B) Rosette of wild type (left) and 35S:OFP4 (right) plants. (C) 8-week-old 35S:OFP4 plants (left) compared with WT plants of the same age (right). Inset, higher magnification view of the 35S:OFP4 plant. (D) The morphology of an 11 week-old 35S:OFP4 plant. (E) Cross section from the basal internode of an 8-week-old wild type (Col-0) plant. (F) Cross section from the basal internode of an 11 week-old 35S:OFP4 plant. Bars, 100 µm for E, and F. Images A-B generated by the Chen lab, UBC.  148 5.2.6 Analysis of double mutants To further test whether OFP1 and OFP4 are required in a KNOX-OVATE complex with KNAT7 to regulate secondary wall formation, we generated double mutants by crossing the knat7-1 mutant line with both ofp1-1 and ofp4-2. Double mutants (homozygous for both mutant alleles) were identified in the F2 population by PCR-aided genotyping, but had no morphological differences compared with wild type (data not shown). Light microscopic analysis of cross-sections from inflorescence stems of double mutants revealed that both of knat7/ofp1 and knat7/ofp4 double mutants exhibited similar phenotypes to knat7, with collapsed vessels and increased thickness of interfascicular fiber cell walls (Figure 5.9A-C). The lack of an additive phenotype suggests that KNAT7 and OFPs function through the same pathway, consistent with a role for KNAT7-OFP4/1 complexes in regulating secondary cell wall formation. If OFP1 and OFP4 functions are manifested through a KNOX-OVATE regulatory complex, we might expect that loss-of function of the corresponding KNOX gene - KNAT7 - would at least partially suppress the OFP overexpression phenotypes. To test this possibility, we generated double mutants by crossing knat7-1 mutant line to 35S:OFP1 and 35S:OFP4 over-expression lines, respectively. While both double mutants retained the knat7 irx phenotype as expected (Figure 5.10C. D), mature double mutant plants were similar in morphology to wild type plants, with normal inflorescence stem development (Figure 5.10A). At the seedling stage, formation of ovate-shaped cotyledons and leaves by OFP4 overexpression was suppressed in the knat7 background (Figure 5.10B).  149     Figure 5.9 Anatomical characterization of knat7/ofp1 and knat7/ofp4 double mutants.  Images show cross sections of internodes at the bases of 8-week-old plants. (A) Wild type Col-0 (B) knat7/ofp1 double mutant (C), knat7/ofp4 double mutant. Double mutants both showed collapsed vessels and thicker interfascicular fiber cell wall phenotypes similar to knat7. Arrows show vessels with irx morphology. Bars, 40 µm.  150    Figure 5.10 Phenotypic characterization of 35S:OFP1/knat7, and 35S:OFP4/knat7 plants.  (A) Morphology of 8 week-old plants transgenic for 35S:OFP1 and 35S:OFP4 in the knat7 background. Plant morphology is similar to wild type (Col-0, shown at left). (B) Morphology of seedlings transgenic for 35S:OFP4 in the knat7 background relative to the wild type (KNAT7/KNAT7) background. (C), (D) Cross sections from basal internodes of 8-week-old 35S:OFP1/knat7 (C) and 35S:OFP4/knat7 (D) stems. Arrows show vessels with irx morphology. Bars, 40 µm.  151 5.3 Discussion Previous data from a yeast two-hybrid screen indicated that some members of the Arabidopsis ovate family protein (OFP) family could have a close functional connection to the TALE homeodomain proteins BELL and KNOX, with which they interact in this system (Hackbusch et al., 2005). Since most of the OFP proteins themselves do not contain predicted DNA binding domains, and OFP1 acts as a strong transcriptional repressor (Wang et al., 2007), a model for OFP functions in developmental control suggests that they participate in a KNOX-BELL-OFP network of protein-protein interactions, acting to repress transcription in association with KNOX and BELL transcription factor complexes (Pagnussat et al., 2007; Wang et al., 2007). In this chapter, I used several approaches to test the hypothesis that OFP1 and OFP4 interact with KNAT7 as part of a transcription factor complex regulating secondary wall formation in the Arabidopsis inflorescence stem.  KNAT7 interacts with OFP1 and OFP4 in vivo To demonstrate that KNAT7 interacts with OFP1 and OFP4 in vivo, we used several different approaches based on protoplast transfection assays. Transfection of OFP1-YFP and OFP4-YFP fusions into protoplasts demonstrated that OFP4 localizes exclusively to the nucleus, similarly to OFP1 (Figure 5.3), consistent with the results of Wang et al. (2007) for OFP1. We also used BiFC (split YFP) in combination with the protoplast transfection system to confirm the interaction of both OFP1 and OFP4 with KNAT7, showing that these interactions can occur in vivo. OFP1 has already been demonstrated to be a potent transcriptional repressor (Wang et al., 2007), and we showed that OFP4 and KNAT7 can also suppress the expression of both a constitutively expressed reporter gene and a reporter gene activated by the VP16 transcriptional activator in a protoplast transient transfection system (Figure 5.3A). Thus, OFP4 and KNAT7 are both indeed transcriptional repressors. In the presence of added 35S:OFP1 or 35S:OFP4, transcription repression by KNAT7 in transfected protoplast cells was clearly enhanced (Figure 5.3B). These results provide further evidence that KNAT7 interacts with OFP1 and OFP4 in vivo, and furthermore suggest a biological function for this  152 interaction, the enhancement of the transcriptional repression by KNAT7. This suggests that transcriptional repression is important in the biological activity of the KNAT7-OFP complex. KNAT7 contains a conserved MEINOX domain, which contains KNOX1 and KNOX2 domains and is known to be required for heterodimerization with SKY and BELL domains of BLH proteins (Bellaoui et al., 2001; Muller et al., 2001) and a homeodomain that functions by directly interacting with target DNA or other proteins (Gehring et al., 1994a). OFPs are characterized by a conserved C-terminal OVATE domain, and most members of this family contain a predicted nuclear localization signal (Hackbusch et al., 2005). The interaction domain of Arabidopsis TALE proteins and OFPs was identified using deletion constructs in the yeast two-hybrid system, and the conserved OVATE domain of the OFPs was found to mediate the interaction with the homeodomains of both BELL and KNOX proteins (Hackbusch et al., 2005). In terms of transcriptional repression, Wang et al. (2007) found that the middle region of OFP1 protein probably represents the major repression domain of OFP1, while the conserved OVATE domain contributed little to its transcriptional repression function. To our knowledge, transcription activation and/or repression domains have not been identified in TALE homeodomain proteins. To identify potential KNAT7 transcription repression domains, we used truncated versions of KNAT7 and found that the N-terminal region containing the KNOX1 domain alone could not function in transcriptional repression. However, since the KNOX2, MEINOX, and homeodomains were all effective in transcriptional repression (Figure 5.4C), repression domains may be present in both the KNOX2 and homeodomain portions of the protein, or reside in the center of KNAT7 between KNOX2 and the homeodomain.  A KNOX-OVATE complex is involved in Arabidopsis secondary wall formation Little information is available about the functions of TALE homeodomain protein-OFP complexes, with exception of a report that a BLH1-KNAT3-OFP5 complex regulates egg development in Arabidopsis (Pagnussat et al., 2007). Given that KNAT7 plays a key role in secondary wall and vascular development, the in vivo interaction between KNAT7 and OFP1 and OFP4 suggested that one or both of these OFP proteins could work with KNAT7 as a  153 part of a KNOX-OVATE complex to control secondary wall formation. If this were the case, OFP1 and OFP4 would be expected to have expression patterns that overlap with KNAT7. OFP1 had previously been shown to be expressed at the root and root-hypocotyl junction and multiple organs (Wang et al., 2007). Further analysis of expression patterns of OFP1 and OFP4 (Figure 5.5) showed that promOFP1:GUS and promOFP4:GUS are expressed mainly in root vascular cylinder-stele and xylem and cambial cells in stem sections of inflorescence stems, which is in accordance with the expression pattern of KNAT7. These results demonstrate that, based on the OFP1 and OFP4 expression patterns, OFP1 and OFP4 could associate with KNAT7 to contribute to regulation of secondary wall formation during the course of root and stem development. In agreement, phenotypic characterization of the OFP4 loss-of function mutant ofp4-2, and the ofp1-1/ofp4-2 double mutant showed that the mutants exhibited similar phenotypes in inflorescence stems to knat7, with both thicker interfascicular fiber walls and irregularly shaped vessel elements (the irx phenotype) (Figure 5.7). However, we failed to observe any irx or secondary wall phenotype for the previously described mutant ofp1-1 (Wang et al., 2007). This suggests that, although it can interact with KNAT7 in vivo and shares a similar expression pattern, OFP1 may not function together with KNAT7 in a KNOX-OVATE complex regulating secondary wall formation, or may play a minor role. This is supported by the fact that the double ofp1/ofp4 mutant had a phenotype very similar to the single ofp4 mutant (Figure 5.7). In addition, to confirm the potential functions of OFP genes, we also made the double mutants of ofp1 and ofp4 with knat7. Both knat7/ofp1 and knat7/ofp4 presented similar phenotypes in inflorescence stems as the knat7 mutant, eg. irx and thicker interfascicular fiber cell walls (Figure 5.9A-C). The lack of an additive phenotype indicates that KNAT7 and OFP4 might work in the same regulatory pathway, consistent with a role for a KNOX-OVATE complex in regulating secondary wall formation. Consistent with the existence of biologically active KNOX-OVATE complexes involving both OFP1 and OFP4, KNAT7 appears to be essential for the pronounced and pleioptropic phenotypic effects of OFP1 and OFP4 overexpression. In the knat7 mutant background, these phenotypes appeared completely rescued and the inflorescence stems phenotypes of these double mutants exhibit knat7-like phenotypes (Figure 5.10). Thus, both of these ovate  154 family proteins might work primarily in partnership with KNAT7, and the regulatory activities might extend beyond secondary wall formation. Overexpression of OFP1, for example, has been shown to have general effects on cell growth and to affect GA homeostasis (Wang et al., 2007). By binding directly or indirectly to other proteins to form a complex, transcription factors regulate the expression of certain target genes. Our results suggest that OFP1, OFP4 and KNAT7 could function together to form a complex that functions as a transcription repression unit to regulate the expression of target genes. OFP1 and OFP4 proteins do not contain apparent DNA binding domains, therefore, the current data support a model in which OFP1 and OFP4 protein can enhance the repression level of certain target genes involving in secondary wall formation through binding to KNAT7. Future work on identifying target genes of KNAT7 and the KNAT7-OVATE complex will be of interest. This regulator module might work in a similar manner to other functional modules linking TALE proteins and OFPs first described by Hackbusch et al. (2005). As one example, Wang et al. (2007) identified OFP1 as a regulator of a key enzyme in GA biosynthesis GA20ox1, and speculated that OFP1 could interact with KNAT1 in repressing GA20ox1 expression (Wang et al., 2007). In a second example, OFP5 appears to act as a key negative regulator of the activity of BELL-KNOX complex between BLH1 and KNAT3 during early embryo sac development in Arabidopsis (Pagnussat et al., 2007). Given the highly cross-linked, yet selective position of OFPs in an interaction network involving regulatory BELL and KNOX TALE homeodomain proteins, these complexes are likely to play fundamental roles in plant development, but a high degree of functional redundancy is also possible. Our work suggests that a KNOX-OVATE transcription repression complex containing KNAT7, OFP4, and likely other interacting TALE or OVATE proteins, plays a key role in plant vascular development and secondary wall formation.   155                CHAPTER 6  Conclusions and future directions    156 Chapter 6. Conclusions and future directions  In this thesis, I investigated the roles of candidate transcription factors in regulating secondary wall formation. The major findings from each chapter are summarized below. 1. Ten Arabidopsis transcription factor genes and their poplar homologs play potential roles in secondary wall formation (Chapter 3). 2. KNAT7 is a regulator of secondary wall formation in Arabidopsis and poplar (Chapter 4). 3. KNAT7 and certain Ovate Family Proteins (OFPs) transcriptional repressors form a functional complex, which may play key role in secondary wall biosynthesis (Chapter 5).  Ten candidate transcription factor genes (Chapter 3) A combination of expression analyses and reverse genetics allowed us to systematically identify new genes required for the development of physical properties of the xylem secondary wall. Based on the work of Ehlting et al. (2005) who followed gene expression over a gradient of Arabidopsis stem developmental stages by microarray analysis, ten up-regulated transcription factors from different families were identified as candidate regulators. Although my analysis of the T-DNA insertion mutants suggested that mutations in only one of the 10 genes gave a clear mutant phenotype, based on their expression patterns others still remain as having possible functions in secondary wall formation. Explanations for the lack of phenotypes in the T-DNA insertion mutants of the other 8 genes for which insertion lines were available could be either functional redundancy with other closely related family members, or for the presence of more subtle phenotypes for which I did not assay. Further experiments to examine if functional redundancy contributes to the lack of apparent phenotypes would be to obtain loss of function mutants in the most closely related gene members and generate double or triple mutants. Moreover, the RNAi and dominant repression approaches have been effectively used to study the redundant functions of transcription factors in regulation of secondary wall biosynthesis (Zhong et al., 2006; Zhong  157 et al., 2007b; Zhong et al., 2008), and could be employed for these genes. Dominant negative repression was especially successfully applied to the study of NAC family genes, since such variants inhibit the function of not only the target TF but also any homologs with redundant function. Other phenotype screening methods could be used, including biochemical analysis of cell wall composition. Such analyses could reveal changes in cellulose, hemicellulose sugars, and lignin content and composition in the mutants, which might not contribute to an obvious irx or other cell wall phenotypes. Screening on the basis of irx phenotypes alone may also limit observation of other secondary cell wall properties, such as cell–cell adhesion. It has been suggested that a network of transcription factors are involved in the regulation of secondary wall biosynthesis and many TFs are required to coordinately regulate the network (Zhong et al., 2008). The study of interactions between transcription factors could reveal potential functions of some candidate genes of unknown function. Five of the candidate gene members are in the larger MYB and bHLH transcription factor families. Interestingly, MYB and bHLH transcription factors have the potential to interact with each other to regulate target gene transcription, such as a TTG1-bHLH-MYB complex that can regulate trichome cell fate and patterning (Zhao et al., 2008b). If these MYB and bHLH candidate genes prove to play roles in secondary wall biosynthesis, it is possible that they work together with partner candidate transcription factors (eg. MYB20, MYB43, or MYB63 with bHLH144 or bHLH068). To test for these specific interactions in the future, the yeast 2-hybrid system could be used. It is also possible that interacting partners are not among the candidates identified. To identify potential additional interacting proteins, MYB and /or bHLH candidate genes can be used as bait to screen for potential new interacting proteins in the yeast 2-hybrid system. Based on the phylogenetic relationships between Arabidopsis and Populus TF gene family members, we identified the potential poplar orthologs of these ten candidate genes. The two experiments conducted by Treenomix I using the 15.5K poplar cDNA microarray to identify poplar genes differentially regulated secondary xylem development at the poplar shoot apex and differentially regulated development of maturing secondary xylem in wood of mature trees, have led us to a set of candidate poplar genes up-regulated in secondary xylem  158 maturation and in secondary wall formation over the course of development in both experiments. Microarray expression profiling and RT-PCR verification showed the poplar orthologs of the Arabidopsis MYB candidates, PoptrMYB028, PoptrMYB018, PoptrMYB152, and the poplar orthologue of AtKNAT7, PoptrKNAT7, were up-regulated in coordination with the onset of secondary wall formation. Therefore, the identification of the potential functions of these genes in poplar wood formation will be a future important goal. Previous study on the poplar ortholog of Arabidopsis SHOOT MERISTEMLESS (STM) provides the approach to study poplar ortholog genes using poplar system (Groover et al., 2006a). Our potential poplar orthologs can be cloned into a 35S promoter-driving vector, and introduced into poplar using Agrobacterium-mediated leaf disk transformation to determine the effects of overexpression in poplar wood formation.  KNAT7 functions in secondary wall formation (Chapter 4) Data from the characterization of knat7 mutants suggest that KNAT7 is a key transcriptional regulator associated with secondary wall formation. Interestingly, there is only one mutant published that exhibits an interfascicular fiber (IFF) phenotype similar to that of knat7. The gapped xylem (gpx) mutant exhibits gaps in the xylem in position where xylem elements would normally be located, resulting plants having fewer functional xylem elements. gpx also shows a highly irregular pattern of secondary cell wall thickening in interfascicular fiber cells, with some cells apparently undergoing little or no secondary cell wall deposition, but some cells with thicker cell walls (Turner and Hall, 2000). Unfortunately, so far there is no information on cloning of the GPX gene. A possible explanation of both the collapsed vessels and thicker IFF cell walls in the knat7 mutant arises from considering both the similarities and differences of these cell types. Both vessel and IFF cells form a secondary cell wall that contains a variety of different polymers, including cellulose, hemicellulose and lignin. However, the secondary cell walls of these cell types show some differences, such as in lignin composition. In cellulose measurements of stem cell wall material reported by Brown et al (2005), knat7 stems exhibited a slight decrease in cellulose content compared with wild type, but noncellulosic carbohydrate composition measurements showed that both wild type and the knat7 mutant have similar  159 proportion of sugars and similarly high xylose content, a characteristic of secondary walls (Turner and Somerville, 1997; Brown et al., 2005). Therefore, we might suggest that another component of secondary walls, lignin, may differ in amount in the vessel and IFF cell walls of the knat7 mutant relative to wild type. Thus, an essential future area of investigation would be on the chemical analysis of lignin composition in the mutant. If the interfascicular fiber cells and xylem cells of Arabidopsis stems can be separately analyzed using a laser capture microdissection system, the lignin content of knat7 in xylem and in IFF, relative to wild type, could be measured and could better explain the two different knat7 phenotypes seen in vessels and IFF. Overexpressing genes of interest could also help to reveal the potential roles of KNAT7 in secondary wall biosynthesis. Zhong et al (2008) reported that when KNAT7 was placed under the control of the CaMV 35S promoter and expressed in wild type Arabidopsis plants, no ectopic deposition of secondary walls, and no changes in fibers and vessels were observed. We observed a similar lack of phenotypes in plants expressing KNAT7 under the control of the double 35S promoter, but placing the KNAT7 gene under the control of the parsley 4CL1 promoter generated a striking cell wall phenotype, in which IFF cell walls were much thinner than wild type. This highlights the need to target overexpression to appropriate cell types in which the process of interests (e.g. lignification) is taking place. Because activation of the secondary wall biosynthetic program may require all the components in the transcriptional complex, overexpression partners that interact with the TF (e.g. OFP4 in the case of KNAT7) could enhance any phenotypes related to the secondary wall deposition. An interesting feature of KNOX protein function is the movement between cells demonstrated for some members. This was first observed in maize homeodomain protein KN1, and is known that KN1 protein (and mRNA) is translocated through the plasmodesmata (Lucas et al., 1995). Furthermore, in Arabidopsis, the KNAT1 protein shows a limited mobility (Kim et al., 2003). The activity of some KNOX homeodomain proteins is dependent on intercellular trafficking for their normal developmental function. Since promKNAT7:GUS expression patterns are not at the location where the mutant phenotypes exhibit, it is possible that, promKNAT7:GUS expression patterns may not indicate the cells  160 in which the KNAT7 protein accumulates. We may further test this hypothesis by protein immunolocalization using a KNAT7 antibody.  KNOX-OVATE complex (Chapter 5) KNAT7 was known to interact in yeast two hybrid assays with members of the Ovate Family Protein (OFP) transcription co-regulators (Hackbusch et al., 2005). In our study, we used various methods based on a protoplast transient assay system to further investigate this interaction. I showed that KNAT7 and both OFP1 and OFP4 are transcriptional repressors, and in addition that KNAT7 can interact with OFP1 and OFP4 proteins in planta, which accentuates KNAT7 repression activity. In addition, the GUS expression patterns of OFP1 and OFP4 promoters are similar with that of the KNAT7 promoter. Moreover, an ofp4 mutant exhibited similar irx and cell wall phenotypes as knat7, and pleiotropic OFP1 and OFP4 overexpression phenotypes depend upon KNAT7 function. To investigate in more detail functions of the OFP and KNAT7 interactions, I generated double mutants (ofp1/knat7; ofp4/knat7), which had similar phenotypes to the single mutants. To extend this analysis, a triple mutant (ofp1/ofp4/knat7) could also be made to help test whether potential KNAT7-OFP4 and KNAT7-OFP1 complexes work in different pathways. In further experiments, we could cross the prom4CL1:KNAT7 overexpression line into the ofp1 and ofp4 mutant backgrounds, to determine if the KNAT7 overexpression phenotype depends on OFP4 or OFP1. prom4CL1:OFP1/4 overexpression constructs could also be transferred into plants, alone or in combination with prom4CL1:KNAT7 to determine if similar or enhanced phenotypes can be generated by manipulating OFP1/4 overexpression in lignifying cells. KNAT7 loss-of function leads to the specific repression of lignin biosynthetic genes, which suggests that KNAT7 directly or indirectly regulates lignin biosynthesis. To confirm KNOX-OVATE complex is essential to regulate downstream target genes, we can perform the same experiment using ofp4 loss-of function mutants compared with wild type to investigate if OFP4 is involved in the regulation of lignin biosynthesis as well. BELL and KNOX proteins are members of the TALE superfamily that function as heterodimers and are known to form functional complex regulating developmental processes in plants (Bellaoui et al., 2001; Muller et al., 2001; Smith et al., 2004). OFP  161 proteins have been shown to interact with both KNOX and BLH homeodomain proteins, which indicates a close functional connection to TALE homeodomain proteins (Hackbusch et al., 2005). It is likely that OFP, KNOX, and BELL proteins form transcription complexes to regulate plant development by controlling the transcription of target genes. A recent study confirmed that the complex of BLH1-KNAT3-OFP5 regulates cell specification in Arabidopsis embryo sac development (Pagnussat et al., 2007). Interestingly, BLH5, a member of BELL homeodomain protein family that is so far poor characterized was shown to interact with KNAT7 and also OFP1 (Hackbusch et al., 2005). Both BLH5 and OFP1 are differentially regulated over the course of inflorescent stem development according to microarray analysis (Ehlting et al., 2005). Preliminary data from the Douglas lab shows that two blh5 T-DNA insertion mutants have no obvious cell wall or other mutant phenotypes compared to wild type, similar to the ofp1 mutant. Thus, the role, if any, of BLH5 in the KNAT7-OFP1/4 KNOX-OVATE complex I identified remains unclear. Further studies on the interactions between KNAT7, OFP1, and BLH5 in planta and the identification of target genes of a potential KNOX-BLH-OVATE complex are promising areas that are worth further investigation.  Transcriptional network of secondary wall biosynthesis A network of transcription factors involved in the regulation of secondary cell wall biosynthesis has been reported including several NAC and MYB domain genes. SND1 and SND1-related transcription factors are placed at the top of the network as master switches to activate the developmental program of secondary cell wall biosynthesis. MYB46, SND3, MYB103 and KNAT7 are direct targets of SND1 and its homologs (Zhong et al., 2008). Combined with our detailed characterization of KNAT7, the identification of KNAT7 as a direct target of SND1 moves us a step closer to elucidating the cascade of transcription factors involved in regulation of secondary wall biosynthesis. Our study revealed that KNAT7 can interact with OFPs to regulate secondary wall formation, and the KNAT7-OFP interaction complex could enhance the transcriptional repression activity of KNAT7. Therefore, summarizing all the data, we propose a model of for the function of this KNOX-OVATE complex in the SND1-mediated pathway (Figure  162 6.1). Expression of KNAT7 is induced by both SND1 and MYB46, and both MYB46 and KNAT7 regulate lignin biosynthetic genes in a direct or indirect way. In addition, OFP1 and OFP4 do not have predicted DNA binding domain, so their role in regulation of downstream targets must be through interaction with KNAT7. Since KNAT7, together with OFP4, functions as a repressor, but lignin biosynthetic genes are repressed in the knat7 mutant, our model suggests that the target of the KNAT7-OFP4 complex is an inhibitor of lignin biosynthetic genes (Figure 6.1).  163     Figure 6.1 A proposed model of the function of KNOX-OVATE complex in the SND1-mediated pathway.  (Left) KNAT7 binds to DNA binding domain and directly or indirectly represses the expression of lignin biosynthetic inhibitor genes, while the OFPs have no predicted DNA binding domain. (Right) The interaction complex of KNAT7-OFPs can enhance the repression of the lignin biosynthetic inhibitor genes.  164 Having determined that KNAT7 acts as an active transcriptional repressor, and that KNAT7 induces the expression of lignin biosynthetic genes directly or indirectly, the investigation of the direct target genes whose expression is regulated by KNAT7 will help to complement the information obtained on the secondary wall formation transcriptional network. To test it directly, chromatin immunoprecipitation (ChIP) can be used to determine the association of KNAT7 with the cis-acting regulatory sequence of potential lignin biosynthetic genes. In addition, dexamethasone (DEX)-inducible system uses the construct of the mammalian glucocorticoid receptor (GR) fusing to KNAT7 under either native promoter of KNAT7 or 35S promoter driven and transform the construct into knat7 mutant plants. Treatment of 35S: KNAT7-GR lines with DEX is expected to mobilize KNAT7-GR from the cytoplasm to the nucleus, activating KNAT7 target genes. Microarray analysis will be used to identify downstream differentially expressed genes using control transgenic lines of the empty vector containing GR alone. Transfer of information from Arabidopsis to other plants of economic value is becoming an important area of research. Based on the genome sequence and genomics tools available in poplar, and the high degree of gene conservation between Arabidopsis and poplar, this is an attractive system for defining regulatory genes such as transcription factors. Such poplar genes could be used for genetic improvement and also as functional tools for transgenic research. In our study, we have identified that four Arabidopsis transcription factor encoding genes (MYB63, MYB20, MYB43, KNAT7), together with their Populus putative orthologs, are differentially expressed in association with secondary wall formation (PoptrMYB028/fgenesh4_pg.C_LG_V000361, PoptrMYB018/grail3.0038010201, PoptrMYB152/ eugene3.00002261, and PoptrKNAT7/estExt_fgenesh1_pg_v1.C_LG_I0964) in both Arabidopsis and Populus. In addition, the close homologs of SND1 and SND1-regulated transcription factors were identified in tree species, and some of them were shown to be expressed in developing wood (Patzlaff et al., 2003b; Patzlaff et al., 2003a; Karpinska et al., 2004; Goicoechea et al., 2005; Bedon et al., 2007). Therefore, it is likely that the proposed SND1-mediated transcriptional network regulating secondary cell wall biosynthesis is conserved between the herbaceous plant Arabidopsis and trees such as poplars. Since Arabidopsis OFP proteins were the first time identified to have a functional  165 interaction with KNOX homeodomain proteins in our study, this offers an opportunity to directly transfer this finding to poplar. Therefore, future study of this network should focus on identification of the potential Populus orthologs of Arabidopsis OFPs, testing protein-protein interaction in Populus, and investigating the transcriptional network underlying regulation of secondary cell wall biosynthesis in Populus. This will undoubtedly provide insights into the molecular mechanisms underlying wood formation in tree species, and maybe provide functional gene resources for transgenic approaches to manipulate the biosynthesis of cellulose, xylan and lignin in this and other trees.    166 References Abel, S., and Theologis, A. (1994). Transient transformation of Arabidopsis leaf protoplasts: a versatile experimental system to study gene expression. Plant J 5, 421-427. Alonso-Cantabrana, H., Ripoll, J.J., Ochando, I., Vera, A., Ferrandiz, C., and Martinez-Laborda, A. (2007). Common regulatory networks in leaf and fruit patterning revealed by mutations in the Arabidopsis ASYMMETRIC LEAVES1 gene. Development 134, 2663-2671. Andersson-Gunneras, S., Mellerowicz, E.J., Love, J., Segerman, B., Ohmiya, Y., Coutinho, P.M., Nilsson, P., Henrissat, B., Moritz, T., and Sundberg, B. (2006). 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