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Roles of the BLADE-ON-PETIOLE genes in Arabidopsis thaliana lateral organ development McKim, Sarah Michelle 2009

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ROLES OF THE BLADE-ON-PETIOLE GENES IN ARABIDOPSIS THALIANA LATERAL ORGAN DEVELOPMENT  by  SARAH MICHELLE MCKIM BSc., The University of New Brunswick, 1999 MSc., The University of New Brunswick, 2004  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)  August 2009 © Sarah Michelle McKim, 2009  Abstract The BLADE-ON-PETIOLE1 (BOP1) and BOP2 genes encode redundant transcription factors involved in morphological patterning in the proximal regions of lateral organs in Arabidopsis thaliana. Loss-of-function bop1 bop2 mutants display several developmental defects including a loss of floral organ abscission. Abscission occurs along specialised cell files, called abscission zones (AZs), which form at the boundary between the leaving organ and main plant body. This dissertation examined the contribution of BOP1 and BOP2 to the known abscission developmental framework and determined that bop1 bop2 flowers lack anatomy associated with AZs. Vestigial cauline leaf AZs are also absent in bop1 bop2 suggesting that BOP proteins are essential to establish AZ anatomy in both leaves and flowers, the first genes identified in Arabidopsis to do so. In support of this hypothesis, BOP1/2 activity is required for both premature floral organ abscission and ectopic cauline leaf abscission promoted by the constitutive expression of INFLORESCENCE DEFICIENT IN ABSCISSION (IDA) gene. In addition, BOP1 and BOP2 were found to promote growth of nectary glands which normally develop off the receptacle at the base of stamens adjacent to AZs. Boundary, nectary and AZ specific-gene expression is relatively unperturbed in bop1 bop2, indicating that positional information is intact. Taken together, these data suggest that BOP1 and BOP2 are key downstream modulators of positional programs operating at the lateral organ-plant body interface. bop1 bop2 flowers also show a subtle loss in floral meristem identity. Genetic analyses revealed a crucial role for BOP1 and BOP2 in suppression of secondary inflorescence identity in early floral primordia only when functions either LEAFY (LFY) or APETALA (AP1) were compromised, suggesting that BOP1 and BOP2 are potentiators of LFY and AP1 function rather than floral meristem identity genes themselves. BOP1 and BOP2 belong to the NPR1-like protein family. A phylogenetic analysis in land plants found both monocot and dicot BOP ii  protein homologues that cluster independently from other NPR1-like proteins. My final contribution examined the subcellular localisation of BOP2 during development and showed that BOP protein-protein interactions in planta showed behaviour consistent with that known for NPR1.  iii  Table of Contents Abstract ..................................................................................................................................... ii Table of Contents..................................................................................................................... iv List of Tables ......................................................................................................................... viii List of Figures .......................................................................................................................... x List of Abbreviations................................................................................................................ xi Acknowledgements .................................................................................................................xiv Dedication ...............................................................................................................................xv 1  GENERAL INTRODUCTION .......................................................................................... 1 1.1 1.1.1 1.1.2  1.2  THE SHOOT APICAL MERISTEM .................................................................................. 4 Apical architecture...........................................................................................................................4 Meristem maintenance .....................................................................................................................5  LATERAL ORGAN DEVELOPMENT .............................................................................. 7  1.2.1 Primordium specification and phyllotaxy..........................................................................................8 1.2.2 Boundary specification ....................................................................................................................9 1.2.3 Organ outgrowth and polarity......................................................................................................... 12 1.2.3.1 Establishment abaxial/adaxial polarity................................................................................... 14 1.2.3.2 Establishment of proximal/distal polarity .............................................................................. 15  1.3  FLOWERING AND FLOWERS ....................................................................................... 16  1.3.1 Inflorescence architecture............................................................................................................... 16 1.3.2 Flower development ...................................................................................................................... 20 1.3.3 Execution of floral homeotic programs........................................................................................... 23 1.3.3.1 Polarity in flowers and floral organs...................................................................................... 23 1.3.3.2 Boundary specification in flowers ......................................................................................... 23  1.4  BLADE-ON-PETIOLE1 (BOP1) AND BOP2 GENES ..................................................... 24  1.4.1 Experimental rationale ................................................................................................................... 24 1.4.2 blade-on-petiole1 (bop1) and bop2 mutants.................................................................................... 25 1.4.2.1 BOP1 and BOP2 repress growth in the proximal leaf............................................................. 25 1.4.2.2 BOP1 and BOP2 repress floral bract and abaxial flower growth............................................. 27 1.4.2.3 BOP1 and BOP2 promote abscission..................................................................................... 29 1.4.3 Thesis objectives............................................................................................................................ 30 1.4.3.1 Contributions of BOP1 and BOP2 to abscission .................................................................... 30 1.4.3.2 Contributions of BOP1 and BOP2 to nectary development .................................................... 30 1.4.3.3 Suppressors of bop1 bop2 ..................................................................................................... 30 1.4.3.4 Roles of BOP1 and BOP2 in floral meristem identity ............................................................ 30 1.4.3.5 Biochemical mechanism and phylogeny of BOP1 and BOP2 ................................................. 31  1.5  ABSCISSION BIOLOGY .................................................................................................. 31  1.5.1 Biological overview of abscission .................................................................................................. 31 1.5.2 Anatomy and ontogeny of abscission zones .................................................................................... 32 1.5.3 Activation of abscission ................................................................................................................. 36 1.5.3.1 Enzymes............................................................................................................................... 37 1.5.3.2 Separation signals................................................................................................................. 38 1.5.3.3 Cellular enlargement............................................................................................................. 40 1.5.4 Abscission and protective layers..................................................................................................... 40 1.5.5 Abscission and other cell separation processes................................................................................ 41  1.6  BOP1 AND BOP2 SIGNALLING...................................................................................... 42  iv  1.6.1 1.6.2  2  NPR1 signalling pathway............................................................................................................... 42 Conserved signalling mechanisms between BOP1, BOP2 and NPR1 .............................................. 46  ROLES OF BOP1 AND BOP2 IN ABSCISSION ZONE DEVELOPMENT.................. 47 2.1  INTRODUCTION .............................................................................................................. 47  2.2  METHODS ......................................................................................................................... 48  2.2.1 2.2.2 2.2.3 2.2.4 2.2.5 2.2.6  2.3  Plant materials and growth conditions ............................................................................................ 48 BOP2 in situ hybridisation ............................................................................................................. 49 Senescence and ethylene responses ................................................................................................ 51 Petal breakstrength......................................................................................................................... 51 β-Glucuronidase activity staining ................................................................................................... 51 Microscopy.................................................................................................................................... 52  RESULTS ........................................................................................................................... 53  2.3.1 bop1 bop2 mutants completely lack floral organ abscission ............................................................ 53 2.3.2 Floral organ senescence in bop1 bop2 ............................................................................................ 56 2.3.3 Ethylene responses in bop1 bop2.................................................................................................... 57 2.3.4 Characterisation of the bop1 bop2 abscission defect ....................................................................... 57 2.3.4.1 Petal AZ breakstrength analysis ............................................................................................ 59 2.3.4.2 Scanning electron microscopy of floral organ AZs ................................................................ 59 2.3.4.3 Histological examination of floral organ AZs ........................................................................ 62 2.3.5 Vestigial AZ formation in bop1 bop2 ............................................................................................. 63 2.3.5.1 SEM of vestigial AZs ........................................................................................................... 63 2.3.5.2 Histological examination of vestigial AZs ............................................................................. 65 2.3.6 BOP1 and BOP2 expression analyses ............................................................................................. 67 2.3.6.1 BOP2 in situ hybridisation .................................................................................................... 67 2.3.6.2 BOP1::GUS expression patterns ........................................................................................... 67 2.3.7 Genetic analysis of BOP loci with abscission-related genes ............................................................ 71 2.3.7.1 Suppression of ida and 35S::IDA phenotypes by bop1 bop2 .................................................. 71 2.3.7.2 Abscission-related gene expression in bop1 bop2: IDA::GUS, CHIT::GUS and ..................... 76 BAC::GUS reporter gene analyses ........................................................................................................... 76  2.4 2.4.1 2.4.2 2.4.3 2.4.4  3  BOP1 and BOP2 act early to specify AZ anatomy .......................................................................... 79 BOP1 and BOP2 also specify vestigial AZ anatomy ....................................................................... 82 The role of BOP1 and BOP2 in proximal lateral organ development ............................................... 83 New candidates for shared function with BOP1 and BOP2 ............................................................. 85  ROLES OF BOP1 AND BOP2 IN NECTARY DEVELOPMENT .................................. 87 3.1  INTRODUCTION .............................................................................................................. 87  3.2  METHODS ......................................................................................................................... 90  3.2.1 3.2.2  3.3 3.3.1 3.3.2 3.3.3  3.4 3.4.1 3.4.2 3.4.3 3.4.4  4  DISCUSSION ..................................................................................................................... 79  Plant materials and growth conditions ............................................................................................ 90 β-Glucuronidase activity staining and microscopy .......................................................................... 91  RESULTS ........................................................................................................................... 91 BOP1 and BOP2 are necessary for nectary development................................................................. 91 Genetic interaction between CRABS CLAW and BOP1 and BOP2................................................... 94 Restoration of nectary elaboration in bop1 bop2 by superman-1 ..................................................... 95  DISCUSSION ..................................................................................................................... 98 Loss of nectary elaboration and CRABS CLAW expression in bop1 bop2......................................... 98 Organ identity, boundary position and nectary elaboration.............................................................. 99 Enhanced proliferation permits elaboration of nectary glands in bop1 bop2................................... 102 BOP1 and BOP2 pattern multiple tissues at the receptacle ............................................................ 103  SUPPRESSORS OF bop1 bop2..................................................................................... 104 v  4.1  INTRODUCTION ............................................................................................................ 104  4.2  METHODS ....................................................................................................................... 105  4.2.1 4.2.2 4.2.3  4.3  Plant materials, mutagenesis and growth conditions...................................................................... 105 Genetic screen, suppressor isolation and segregation analysis ....................................................... 105 Microscopy.................................................................................................................................. 105  RESULTS ......................................................................................................................... 106  4.3.1 4.3.2 4.3.3  Isolation of lines showing recovered abscission in sepals and petals.............................................. 106 Penetrance and expressitivity of recovered abscission phenotypes............................................... 1069 Segregation patterns of recovered abscission phenotype of rva1 and rva2 in bop1 bop2 background 109 4.3.4 Abscission zone development in rva1 bop1 bop2 and rva2 bop1 bop2 .......................................... 109 4.3.4.1 Sepal and petal floral organ AZs in rva1 bop1 bop2 show partial development and disorganised cellular expansion ................................................................................................................................. 111 4.3.4.2 Sepal and petal floral organ AZs in rva2 bop1 bop2 are partially activated........................... 114 4.3.5 Nectary development in rva1 bop1 bop2 and rva2 bop1 bop2 ....................................................... 118  4.4 4.4.1 4.4.2 4.4.3 4.4.4 4.4.5 4.4.6 4.4.7  5  DISCUSSION ................................................................................................................... 120 rva1 and rva2 restore abscission to sepals and petals in bop1 bop2................................................ 120 Recovery of sepal and petal abscission can be separated genetically from stamen abscission ......... 121 Organ specificity of abscission recovery....................................................................................... 122 Vascular abscission is not recovered in rva1 bop1 bop2................................................................ 122 rva1 bop1 bop2 and rva2 bop1 bop2 shows aberrant cell expansion.............................................. 123 Suppressors for other bop1 bop2 phenotypes were not recovered .................................................. 124 Conclusions................................................................................................................................. 124  ROLES OF BOP1 AND BOP2 IN FLORAL MERISTEM IDENTITY ........................ 125 5.1  INTRODUCTION ............................................................................................................ 125  5.2  MATERIALS AND METHODS...................................................................................... 128  5.2.1 5.2.2  5.3 5.3.1 5.3.2 5.3.3 5.3.4 5.3.5 5.3.6 5.3.7  5.4 5.4.1 5.4.2 5.4.3 5.4.4 5.4.5  Plant materials and growth conditions .......................................................................................... 128 Scanning electron microscopy...................................................................................................... 128  RESULTS ......................................................................................................................... 129 BOP1 and BOP2 expression in developing floral primordia .......................................................... 129 bop1 bop2 mutants show slight defects in floral meristem identity ................................................ 131 bop1 bop2 lfy-2 and bop1 bop2 lfy-1 triple mutants show enhanced loss of floral meristem identity 132 bop1 bop2 lfy-2 and bop1 bop2 lfy-1 triple mutants enhance lfy floral organ defects ...................... 135 bop1 bop2 ap1-1 and bop1 bop2 ap1-12 show enhanced loss of floral meristem identity............... 140 bop1 bop2 ap1-1 and bop1 bop2 ap1-12 triple mutants enhance ap1 floral organ defects .............. 141 AGL24, SOC1 and FUL mis-expression in bop1 bop2 ap1 and bop1 bop2 lfy................................ 144  DISCUSSION ................................................................................................................... 145 BOP1 and BOP2 act in parallel with LFY to specifically activate AP1 .......................................... 145 Regulation of inflorescence identity gene expression by BOP1 and BOP2..................................... 147 Other genetic players in bract formation and phyllotaxy of floral organs ....................................... 148 Floral organ identity and floral organ number ............................................................................... 150 Roles of BOP1 and BOP2 in floral meristem identity.................................................................... 151  6 PHYLOGENETIC ANALYSIS OF THE NPR1-LIKE PROTEIN FAMILY AND BIOCHEMICAL MECHANISM OF BOP1/BOP2............................................................... 153 6.1  INTRODUCTION ............................................................................................................ 153  6.2  MATERIALS AND METHODS...................................................................................... 155  6.2.1 6.2.2  Plant materials and growth conditions .......................................................................................... 155 Phylogenetic Analysis.................................................................................................................. 156  vi  6.2.3 6.2.4  6.3  Generation of pBOP1::BOP2:EYFP transgenic plants .................................................................. 156 Bimolecular Florescence Complementation.................................................................................. 157  RESULTS ......................................................................................................................... 158  6.3.1 Phylogenetic analysis of the NPR1-like protein family ................................................................. 158 6.3.2 Complementation of pBOP1:BOP2::EYFP plants ........................................................................ 164 6.3.3 Subcellular localisation of pBOP1:BOP2::EYFP during lateral organ development ...................... 164 6.3.4 BOP2 interacts with PAN and as well as itself in planta ............................................................... 169 6.3.4.1 Positive and negative controls ............................................................................................. 169 6.3.4.2 In planta confirmation of BOP and PAN nuclear interaction................................................ 172 6.3.4.3 In planta detection of BOP - BOP interaction in cytoplasm and nucleus............................... 172  6.4 6.4.1 6.4.2 6.4.3  7  DISCUSSION ................................................................................................................... 173 Distribution of BOP-like proteins across in land plants ................................................................. 173 Conservation of protein-protein interactions of BOP2 and NPR1 .................................................. 174 Biological significance of BOP and PAN interaction .................................................................... 176  CONCLUSIONS ............................................................................................................ 179 7.1 7.1.1 7.1.2 7.1.3 7.1.4 7.1.5  7.2  THESIS CONTRIBUTIONS ........................................................................................... 179 Roles of BOP1 and BOP2 in abscission........................................................................................ 179 Contributions of BOP1 and BOP2 to nectary development ........................................................... 179 Analysis of recovered abscising lines ........................................................................................... 180 Roles of BOP1 and BOP2 in floral meristem identity.................................................................... 180 Biochemical mechanism and phylogenetic analysis of BOP1 and BOP2 ....................................... 180  SYNTHESIS ..................................................................................................................... 181  7.2.1 BOP1 and BOP2 are downstream executors of positional programs .............................................. 181 7.2.2 BOP1 and BOP2 as negative regulators of growth? ...................................................................... 183 7.2.3 Relationship between boundaries and proliferation ....................................................................... 184 7.2.4 Abscission zones and nectaries proliferate from cells derived from differentiated tissues - the role of the receptacle ............................................................................................................................................ 185 7.2.5 Do BOP1 and BOP2 respond to developmentally informative redox cues?.................................... 186  BIBLIOGRAPHY.................................................................................................................. 188 APPENDIX ........................................................................................................................... 207  vii  List of Tables  Table 2.1. Characterisation of floral organ abscission in wild-type and bop1 bop2 mutants...... ............................................................................................................................................. 55 Table 3.1 Association between first and third whorl organs and elaboration of nectary growth ............................................................................................................................................. 99 Table 4.1. Phenotype characteristics and segregation of restored floral organ abscission mutants rva1 bop1 bop2 and rva2 bop1 bop2................................................................................... 107 Table 4.2 Mature silique length of rva2 bop1 bop2, bop1 bop2 and Col-0 .......................... 110 Table 5.1 Quantitative analysis of floral-meristem identity phenotypes in wild type and mutants ........................................................................................................................................... 120 Table 5.2. Quantitative analysis of floral-organ identity phenotypes in wild-type and mutants ........................................................................................................................................... 136 Table 6.1 BOP protein homologues in land plants .............................................................. 161 Table 6.2 pBOP1::BOP2-EYFP complementation of bop1 bop2 ........................................ 165  viii  List of Figures  Figure 1.1 Arabidopsis thaliana aerial development ............................................................... 2 Figure 1.2 Shoot organ boundaries in Arabidopsis ................................................................ 10 Figure 1.3 Polarities in developing leaves ............................................................................. 13 Figure 1.4 Flower development in Arabidopsis..................................................................... 18 Figure 1.5 Leaf and inflorescence phenotypes of bop1 bop2 and wild type ........................... 26 Figure 1.6. Comparison of wild-type and bop1 bop2 floral phenotypes................................ 28 Figure 1.7 Floral organ abscission zones............................................................................... 33 Figure 1.8 NPR1 signalling pathway .................................................................................... 44 Figure 2.1 Abscission phenotype and petal breakstrength of bop1 bop2................................ 54 Figure 2.2 Triple response of wild-type and bop1 bop2 seedlings ......................................... 58 Figure 2.3 Hypotheses for lack of floral organ abscission in bop1 bop2 ................................ 60 Figure 2.4 Morphology of floral abscission zones in wild type and bop1 bop2...................... 61 Figure 2.5 Morphology of vestigial abscission zones in wild type and bop1 bop2................. 64 Figure 2.6 Serial sections through vestigial abscission zones of wild-type and bop1 bop2......... ............................................................................................................................................. 66 Figure 2.7 In situ hybridisation of BOP2 mRNA .................................................................. 68 Figure 2.8 Expression of BOP1::GUS .................................................................................. 70 Figure 2.9 Genetic interaction between bop1 bop2 and ida ................................................... 73 Figure 2.10 Suppression of premature and ectopic abscission from 35S::IDA in bop1 bop2...... ............................................................................................................................................. 74 Figure 2.11 Real time quantitative RT-PCR of IDA and HAESA in wild-type and bop1 bop2 abscission zones ................................................................................................................... 75 ix  Figure 2.12 Expression of IDA::GUS, BAC::GUS and CHIT::GUS in wild type and bop1 bop2 ............................................................................................................................................. 78 Figure 2.13 Model of abscission ........................................................................................... 81 Figure 3.1 Nectary morphology and CRC::GUS activity in wild type and bop1 bop2 ......92-93 Figure 3.2 Restoration of mature nectary surface characteristics in bop1 bop2 sup-1 ............ 96 Figure 3.3 Whorl models and nectary formation ................................................................. 100 Figure 4.1 Abscission phenotypes of rva1 bop1 bop2 and rva2 bop1 bop2 ......................... 108 Figure 4.2 Scanning electron microscopy of rva1 bop1 bop2 floral organ abscission zones ............................................................................................................................... ….112-113 Figure 4.3 Scanning electron microscopy of rva2 bop1 bop2 floral organ abscission zones ............................................................................................................................ ……116-117 Figure 4.4 Nectary development in rva1 bop1 bop2 and rva2 bop1 bop2............................ 119 Figure 5.1 Floral meristem defects in bop1 bop2 ................................................................ 131 Figure 5.2 Loss of floral meristem identity and aberrant floral organ identity of lfy mutants are enhanced by bop1 bop2 ............................................................................................... 133-134 Figure 5.3 Loss of floral meristem identity of ap1 mutants are enhanced by bop1 bop2 ........................................................................................................................ ………138-139 Figure 5.4 bop1 bop2 exacerbates floral organ defects in ap1 mutants ................................ 143 Figure 6.1 Phylogenetic analysis of the NPR1-like protein family ...................................... 159 Figure 6.2 Amino acid alignment of NPR1-like proteins in Arabidopsis ...................... 162-163 Figure 6.3 BOP2-EYFP localisation in pBOP1::BOP2-EYFP bop1 bop2 plants ............167-68 Figure 6.4 In planta interaction and localisation of BOP2 and PAN fluorescently-tagged fusion proteins ....................................................................................................................... 170-171  x  List of Abbreviations  ACC - 1-aminocyclopropane-1-carboxylic acid AG – AGAMOUS AGL15 – AGAMOUS-LIKE15 AGL24 – AGAMOUS-LIKE24 ANK – Ankyrin repeat domain AP1 – APETLA1 AP2 - APETALA2 AP3 – APETALA3 AS1 – ASYMMETRIC LEAVES1 AS2- ASYMMETRIC LEAVES2 AT – Arabidopsis thaliana minimal media AtZFP2 – ARABIDOPSIS THALIANA ZINC-FINGER PROTEIN2 AZ – Abscission zone BAC – BEAN ABSCISSION ZONE CELLULASE BOP1– BLADE-ON-PETIOLE1 BOP2- BLADE-ON-PETIOLE2 BP – BREVIPEDICELLUS BTB – Bric-a-brac, tramtrack, broad-complex domain CAL – CAULIFLOWER CHIT - CHITINASE CLV – CLAVATA CRC – CRABS CLAW xi  CUC – CUP-SHAPED COTYLEDONS Cys – Cysteine CZ – Central zone EYFP – Enhanced Yellow Florescent Protein FIL - FILAMENTOUS FLOWER FT – FLOWERING TIME FUL - FRUITFULL HAE – HAESA HSL – HAESA-LIKE HWS – HAWAIIAN SKIRT IDA – INFLORESCENCE DEFICIENT IN ABSCISSION IDL – IDA-LIKE IM – Inflorescence meristem JAG - JAGGED JNT – JOINTLESS KAN – KANADI KLUH - KLU KNAT2 – KNOTTED-LIKE2 KNAT6 – KNOTTED-LIKE6 LAS – LATERAL SUPPRESSOR LBD – LATERAL ORGAN BOUNDARIES DOMAIN LFY – LEAFY LMI – LATE MERISTEM IDENTITY1 MAFFT – Multiple Alignment using Fast Fourier Transform xii  NPR1 – NON-PATHOGENESIS RELATED1 PAN – PERIANTHIA PHB - PHABULOSA PHV - PHAVOLUTA PI – PISTALATTA PNF - POUNDFOOLISH PNY – PENNYWISE POZ – Poxvirus zinc finger domain PR – PATHOGENESIS-RELATED PTL – PETAL LOSS PZ – Peripheral zone RBE – RABBIT EARS REV – REVOLUTA SA – Salicylic acid SAM – Shoot apical meristem SAR- Systemic acquired resistance SEP – SEPELLATA SOC1 – SUPPRESSOR OF CONSTANS1 STM – SHOOT MERISTEMLESS SUP - SUPERMAN SVP – SHORT VEGETATIVE PHASE TFL – TERMINAL FLOWER TGA – TGACG-BOX BINDING UFO – UNUSUAL FLORAL ORGANS xiii  YAB - YABBY WUS - WUSCHEL  xiv  Acknowledgements I gratefully acknowledge the National Science and Engineering Research Council of Canada for funding me through a Canada Graduate Scholarship and to the University of British Columbia for awarding me a University Graduate Fellowship. I am also indebted to my supervisor, Dr. George Haughn, whose insight always helped me put my work into proper focus. I would also like to thank my committee members, Drs. Xin Li, Jim Kronstad and Carl Douglas. In addition, I was greatly helped by my fellow researchers: in particular, Dr. Gill Dean, Dr. Shelley Hepworth, Dr. Robin Young and Will Iles. Furthermore, I would like to pay tribute to my longsuffering roommate Gina Choe for her encouragement throughout the dissertation writing process. Finally, I would like to acknowledge the support of my family, Dad, Mom, Andrew and Heather, and my many wonderful friends whom I’ve met along the way.  xv  Dedication  This thesis is dedicated to the memory of my mother, Jacqueline Dionne McKim (1950-2006)  xvi  1 GENERAL INTRODUCTION Multicellular organisms face serious challenges. By definition, they consist of different types of cells specialised to perform particular functions; however, proliferation of these cell types must be spatially and temporally coordinated such that the relative positions and shapes of developing tissues establish a cohesive body plan. Once a mature body has formed, the maintenance of aging tissues also must be tightly regulated. Developmental biology seeks to understand the underlying mechanisms controlling these processes, from specific cell identity to overall organism architecture. Unlike animal body plans, plant form is largely determined during post-embryonic growth and in response to developmental and environmental cues. All non-embryonic plant structures derive from central populations of stem cells called meristems, laid down during embryonic axis formation at the growing tips of the stem and the root. Meristems have a basal character state that can either differentiate into specific cell types or additional meristems (Fleming, 2005). Angiosperms, the flowering plants, initiate lateral organs as primordia off the flanks of the shoot apical meristem (SAM). Lateral primordia can differentiate into leaves, flowers or additional indeterminate flowering or vegetative shoots. Aerial plant architecture reflects the arrangement and type of lateral organs initiated from the SAM (Fig. 1.1). This dissertation describes the contributions of two homologous genes, BLADE-ONPETIOLE1 (BOP1) and BOP2, to lateral organ development in the small herbaceous crucifer, Arabidopsis thaliana. Arabidopsis, commonly known as thale cress, is a member of the Brassicaceae family which includes familiar cruciferous crops such as broccoli, cauliflower and canola. Through phenotypic characterisation, genetic analysis and biochemical approaches, I will  1  Figure1.1 Arabidopsis thaliana aerial development Following production of a vegetative rosette, Arabidopsis converts to inflorescence flowering characterised by early production of cauline leaves subtending as associated axillary meristem which develops into a seconday inflorescence shoot. The inflorescence shoots commit to an indeterminate production of floral meristems which then elaborate into flowers. (Inset) diagram of a longitudinal section through the shoot apical meristem (SAM) which forms at the tip of the growing stem. The SAM is divided into a central zone (red) surrounded peripheal zone (orange) where lateral organs arise (P). Beneath the central zone is the organising centre (merlot) and the rib meristem (green). Superimposed on this structure is the L1 and L2 tunica layers and the multi-layer L3 corpus. Diagram adapted by permission from Macmillan Publishers Ltd: Nature Reviews Genetics, Wolters and Jurgens, 2009.  www.nature.com/nrg/index.html  2  demonstrate that BOP1 and BOP2 play integral roles in both leaf and flower organogenesis, acting to execute morphological organ plans, in particular with regards to boundary establishment with the main plant body. Before further discussion of BOP1 and BOP2, general regulatory mechanisms central to aerial plant development will be described. In all seed plants, including Arabidopsis thaliana, the primary SAM is formed in the embryo and after germination generates a vegetative stem that produces leaves. In the axil between the emerging leaf and SAM, a new lateral meristem or axillary meristem develops, capable of forming additional vegetative branches. In response to environmental and endogenous cues, the SAM shifts from vegetative to reproductive identity, a major developmental phase change known as the floral transition, to become an inflorescence meristem (IM). The central plant stem is now referred to as the primary flowering shoot or inflorescence stem (Fig. 1.1). The IM continues to produce vegetative organs, called cauline leaves, which also develop associated axillary meristems. These axillary meristems elaborate into secondary inflorescence shoots. Later inflorescence nodes lack leaves but still produce lateral meristems, called floral meristems that, in turn, differentiate a determinate number of lateral floral organs which collectively form the flower. Floral organs are classically viewed as modified leaves (Goethe, 1790). Thus fate of the lateral or secondary meristems switches from vegetative to inflorescence shoot to floral, phase transitions that strongly influence the overall plant body plan (Poethig, 1990).  3  1.1 THE SHOOT APICAL MERISTEM 1.1.1 Apical architecture Most angiosperm meristems consist of isodiametric, thin-walled and weakly vacuolated cells which form a dome-shaped apex at the stem tip (Steeves and Sussex, 1989). Underlying this homogenous cellular anatomy, however, is a functional segregation between the upper central zone (CZ) of slowly proliferating cells and the surrounding peripheral zone (PZ) of more quickly dividing cells (Fig. 1.1). The CZ plays a dual role to maintain the primary meristem population, called stem cell initials, while continually contributing cells to the PZ. Cells within the PZ depart from basal meristem cell identity to form clusters of so-called founder cells which develop into lateral organs (Poethig and Sussex, 1985). Underlying the CZ and the PZ is the rib meristem (RM) whose pith growth continually propels the stem upward (Steeves and Sussex, 1989). The organising centre of the upper RM is responsible for maintaining the meristematic identity and integrity of the overlying CZ cells (Laux et al., 1996; Mayer et al., 1998; Wolters and Jurgens, 2009). Apical meristems are also categorised based on cytological stratification as described by by the tunica corpus model (Fig. 1.1; (Schmidt, 1924). Most eudicots, including Arabidopsis, have a two-layered tunica, the outer L1 and inner L2, characterised by anticlinal (at right angles) cell divisions allowing for continual surface growth , L1 and L2 layers overlie the multilayer L3 corpus cells that divide along multiple planes to yield thickened growth (Evert, 2007). The L1, L2 and L3 layers are encompassed by the CZ and extend into the PZ where the distinction amongst cell layers is lost (Steeves and Sussex, 1989). Secondary meristems formed in regions other than at the primary shoot tip such as the axillary vegetative and inflorescence shoots as well as floral meristems, that do not derive from a  4  continual descent of stem cells but rather reflect a reversion from a more differentiated state (Steeves and Sussex, 1989; Long and Barton, 2000).  1.1.2 Meristem maintenance The CZ consists of approximately 35 stem cell initials in Arabidopsis (Yadav et al., 2009), an astonishingly small number considering that these cells supply all plant aerial structures. Due to continual recruitment of cells from the CZ to the PZ, SAM integrity depends upon the retention of meristematic identity in sufficient proportion of CZ daughter cells. Balance between meristematic versus lateral organ fate is regulated by the synergistic activity of two central players, SHOOT MERISTEMLESS (STM) and WUSCHEL (WUS). Loss of function stm mutants lack apical meristems and arrest shortly following cotyledon (embryonic leaf) emergence (Barton and Poethig, 1993). STM encodes a member of the class 1 KNOX (KNOTTED1-LIKE HOMEOBOX) gene family and is expressed throughout the SAM (Long et al., 1996). In addition to specifying meristem identity, STM delays differentiation of PZ stem cell derivatives to allow for adequate PZ proliferation fueling primordium growth (Lenhard et al., 2002). STM is rapidly excluded from PZ cells following commitment to lateral organ fate by the action of ASYMMETRIC LEAVES1 (AS1), a MYB domain protein, and ASYMMETRIC LEAVES2 (AS2), a LATERAL ORGAN BOUNDARIES DOMAIN (LBD) protein (Byrne et al., 2000; Ori et al., 2002; Iwakawa et al., 2002; Lin et al., 2003; Okushima et al., 2007; Guo et al., 2008). Other members of the class 1 KNOX (hereafter referred to as KNOX) gene family, including BREVIPEDICELLUS (Colombo et al., 1995) and KNOTTED-LIKE HOMEOBOX2 (KNAT2) have been implicated in meristem maintenance (Scofield and Murray, 2006); in fact, KNOX genes appear essential for meristematic competence in all angiosperms examined (Hake et al., 2004). In general, impaired KNOX functions lead to meristem arrest while ectopic misexpression 5  of KNOX results in ectopic meristem formation and subsequent lateral organ growth (Hake et al., 2004). Like STM, WUS promotes meristem cell identity (Schoof et al., 2000; Gallois et al., 2002). Meristem cell populations are completely depleted by developing organs in wus mutants, suggesting that WUS activity balances meristem cell proliferation with lateral organ production (Laux et al., 1996; Mayer et al., 1998). WUS is expressed in the organising centre and regulates the overlying CZ cells non-cell autonomously via the CLAVATA (Brand et al., 2000) pathway (Mayer et al., 1998; Fletcher et al., 1999; Brand et al., 2000; Schoof et al., 2000; Lenhard and Laux, 2003; Reddy and Meyerowitz, 2005). Reduced CLV signalling causes an expansion of WUS expression, associated with an enlarged CZ which produces more lateral organs (Clark et al., 1993; 1995, Kayes and Clark, 1998) indicating that lateral organ production is limited, in part, by meristem size. Pathways controlled by STM and WUS are important for integrity of all apical meristems including axillary shoot and floral meristems (Long and Barton, 2000; Sablowski, 2007). Hormones have also been implicated in meristem maintenance (Hay et al., 2004; Wolters and Jurgens, 2009). Cytokinin (CK), a classic plant hormone long associated with cell division and meristematic fate, promotes the expression of KNOX genes, such as STM, which in turn enhances cytokinin biosynthesis (Rupp et al., 1999; Frank et al., 2000; Hamant et al., 2003; Jasinski et al., 2005; Yanai et al., 2005). WUS may also enhance cytokinin signalling and thus KNOX expression in the SAM via negative regulation of type A ARR transcription factors that suppress cytokinin synthesis (Leibfried et al., 2005). Another classic plant hormone, gibberellic acid (GA) associated with suppression of cell division, has been proposed to antagonize KNOX function and be actively degraded by KNOX gene activity (Sakamoto et al., 2001; Hay et al., 2002; Gallois et al., 2002; Jasinski et al., 2005). 6  1.2 LATERAL ORGAN DEVELOPMENT Both zonation and stratification categorisations of meristem architecture relate to lateral organ development. Leaves and flowers initiate as primordia from the PZ of the apical meristem. Given the small size of the CZ, the bulk of a lateral organ derives from the extensive proliferation of the PZ cells during specification and early stages of primordium outgrowth (Laufs et al., 1998; Reddy et al., 2005). Progress from meristematic precursor to determinate lateral organ involves numerous steps including primordia specification, initiation, commitment to specific lateral organ fate and morphogenesis of mature tissue. The tunica-corpus distinctions determine the type of tissue each layer differentiates into: L1 cells give rise to the epidermal tissue, the L2 to several underlying layers while the L3 contributes to the ground tissue (Sussex, 1998). These layers are clonally distinct (Satina et al., 1940) and thus, communication between layers is essential for coordinated growth (Satina et al., 1940; Satina and Blakeslee, 1941). The earliest anatomical signs of primordium initiation in the PZ are periclinal rather than typical anticlinal divisions in the L2 layer. Subsequent primordium growth reflects a combination of periclinal and anticlinal divisions in the L2 and L3 layers. The genotype of the L3 was shown to control meristem size and organ initiation position (Szymkowiak and Sussex, 1992; 1993), suggesting that primordium initiation is influenced by inner to outer control, correlating well with the non-cell autonomous signalling of the WUS/CLV network. However, superficial application of cell wall loosening enzymes can induce primordia formation in the PZ, suggesting that the mechanical properties of the L1 cell wall or the release of wall-signalling molecules may also play an important role (Fleming et al., 1997; 1998; 2006; Pien et al., 2001; Peaucelle et al., 2008; Nakayama and Kuhlemeier, 2009).  7  1.2.1 Primordium specification and phyllotaxy The zonation model of meristem architecture suggests that lateral organ cell fate is dictated by its peripheral position. Primordia are marked by the downregulation of STM and the expression of several lateral organ-specific genes, such as AS1 and FILAMENTOUS FLOWER (FIL; Yadav et al., 2007). However, not all peripheral cells generate lateral organs at any one time; rather, primordia are initiated in a regular, organised pattern, described as phyllotaxis. As in other seed plants, Arabidopsis exhibits spiral phyllotaxis off the SAM, where leaves and secondary meristems, including floral meristems, arise with consistent spacing of 137.5 (Steeves and Sussex, 1989; Kuhlemeier, 2007). A crucial player in phyllotaxis and primordia initiation is the plant hormone auxin (Tanaka et al., 2006; Benjamins and Scheres, 2008; Vanneste and Friml, 2009). Auxin foci in the L1 mark regions of presumptive primordia of leaves, flowers and floral organs (Benkova et al., 2003; Heisler et al., 2005; Scarpella et al., 2006) formed via epidermal flow of auxin driven by the activity of auxin efflux carriers called PINs (PINFORMED), auxin influx carriers and local auxin production (Steiger et al., 2002; Benkova et al., 2003; Reinhardt et al., 2003; Heisler et al., 2005; Bainbridge et al., 2008; Zhao, 2008). The current phyllotaxy model suggests that a localised maximum in the L1 acts as a sink for auxin accumulation in the SAM, leading to areas of local auxin depletion surrounding the initiating organ; thus, subsequent auxin maxima can only develop at a specified distance from the proceeding primordia (Kuhlemeier, 2007). Following initiation, PINs divert auxin flow subepidermally, establishing an acropetal gradient of auxin in the emerging primordium extending from high at the tip to low at the base (Benkova et al., 2003; Heisler et al., 2005).  8  1.2.2 Boundary specification A key event in lateral organogenesis involves the formation of boundaries between cells remaining associated with the meristem and those committed to lateral organ fate (Fig. 1.2a). Boundary formation within and between the floral whorls ensures separation between developing floral organs (Fig. 1.2b). Loss of boundary formation usually leads to the excess growth and fusion of the lateral organ tissue and of the stem tissue (Aida and Tasaka, 2006). Boundaries are traditionally considered regions of suppressed growth (Breuil-Broyer et al., 2004; Aida and Tasaka, 2006; Blein et al., 2008); however, mitotic indices show that the rate of cell division is unchanged in the boundaries versus the meristem proper (Laufs et al., 2004). Analysis of the CUP-SHAPED COTYLEDONS (CUC) genes (CUC1, CUC2 and CUC3) suggest that the boundary domain has a distinct non-lateral organ identity consisting of specialised cells whose proliferation is tightly controlled (Golz and Hudson, 2002; Aida et al., 2006; Rast and Simon, 2008). The CUC genes encode transcription factors essential for boundary formation between lateral organs and the meristem and between lateral organs themselves (Long et al., 2006; Aida et al., 1997; Aida et al., 1999; Takada et al., 2001; Vroemen et al., 2003; Keller et al., 2006; Hibara et al., 2006). The bases of the cotyledons fuse in double cuc1 cuc2 mutant seedlings, leading to a blunted stem apex lacking an SAM (Aida et al., 1999; Sakamoto et al., 2001) suggesting that, in addition to establishing organ boundaries, the CUC genes also promote meristem development. CUCs contribute to SAM establishment in part through up-regulation of STM and other KNOX genes (Aida et al., 1997, 1999; Long et al., 1996; Hibara et al., 2003). CUC2 and CUC3 are also implicated in axillary meristem formation as well as the separation of leaves and flower pedicels from the stem (Hibara et al., 2006; Raman et al., 2008). CUC expression is restricted to the boundary zones by antagonistic regulation from STM in the meristem and by AS1/AS2 in the developing organ (Aida et al., 1999; Sakamoto et al., 2001; 9  Figure 1.2 Shoot organ boundaries in Arabidopsis Scanning electron micrographs of Arabidopsis meristems and lateral organs (a) Top view of the inflorescence meristem (b) Floral meristem (c) Longitudinal section through the seedling shoot apical meristem (SM) stained with Toluidine blue. The black arrows show the boundary between the meristem and organ primordia, whereas the white arrows indicate the boundary between adjacent organ primordia. Both types of boundary are characterised by the formation of a hollow or constriction. Note that in (c), the shoot meristem (SM) consists of densely stained small cells, whereas the cotyledons (CO) contain vacuolated and elongated cells. SM, shoot apical meristem; FM, floral meristem; IM, inflorescence meristem; P1–P7, flower primordia; S, sepal primordia. Taken with permission from Elsevier: Current Opinion in Plant Biology, Aida and Tasaka, (2006).  www.sciencedirect.com/science/journal/13695266  10  Hibara et al., 2003; Xu et al., 2008). CUC1 and CUC2 mRNAs are also targets of posttranscriptional downregulation by miRNA164 (Laufs et al., 2004; Mallory et al., 2004; Baker et al., 2005; Nikovics et al., 2006; Sieber et al., 2007; Peuacelle et al., 2007). When expression domains of CUCs are enlarged, so is the corresponding boundary region (Laufs et al., 2004), suggesting that CUCs promote the proliferation of boundary cells. CUCs are expressed in areas of low auxin and their expression domains are enlarged in polar auxin transport mutants (Vernoux et al., 2000; Furutani et al., 2004; Heisler et al., 2005) indicating that polar auxin transport from the developing primordia also restricts CUC expression to the boundary zone. As the name implies, genes of the LATERAL ORGAN BOUNDARIES DOMAIN (LBD) (Okushima et al., 2007) family often have boundary-specific expression (Shuai et al., 2002; Husbands et al., 2007). In addition to AS2, which is a LBD protein, JAGGED LATERAL ORGANS (JLO) was recently shown to promote boundary identity through downregulation of auxin signalling and promotion of KNOX expression (Borghi et al., 2007). Other single lbd mutants examined do not show boundary-specific phenotypes presumably due to redundancy (Shuai et al., 2002; Okushima et al., 2007). Several regulators of floral boundary specification have been identified. PETAL LOSS (PTL) is important to separate sepals and functions in parallel with the CUCs (Brewer et al., 2004). Another flower-specific gene, RABBIT EARS (RBE) is also critical for sepal separation, in addition to promoting petal identity, and it functions downstream of the CUCs (Takeda et al., 2004; Krizek et al., 2005). The adaxial boundary between the lateral organ and the meristem is also the site of axillary meristem formation (Aida and Tasaka, 2006) underscoring the relationship between boundary formation and their promotion of meristematic competence. Recent evidence suggests a pivotal role for CUCs in promoting both axillary meristems and boundaries (Raman et al., 2008). 11  1.2.3 Organ outgrowth and polarity Due to localised anticlinal and periclinal divisions, the primordium bulges off the SAM (Fig. 1.2a). At this point, primordia for different types of lateral organs are equivalent. Similar to meristems, the fate of a lateral organ cell is dependent upon its final position (Tilney-Bassett, 1986; Irish, 1993; Szymkowiak and Sussex, 1996; Evert, 2007). Cells which retain meristematic identity maintain small, cytoplasmically-dense anatomy and proliferative capacity while those incorporated into lateral organs become larger, highly vacuolated and show slower rates of cell division (Steeves and Sussex, 1989). Subsequent lateral organ outgrowth reflects a combination of cell division and expansion coupled with cellular specialisation to form the tissues which define lateral organ cells (note the distinction between the SAM and cotyledons in Fig. 1.2c). Lateral organs develop several polarities with respect to the meristem during outgrowth: proximal/ distal, adaxial (beside)/ abaxial (away), and medial/ lateral (Golz and Hudson, 2002; Fig. 1.3). Both leaves and flowers develop particular shapes defined by the activities of genes specific to these domains. Arabidopsis leaves are connected to the stem by a narrow petiole which expands into a single, distal blade (Fig. 1.3). The side of the leaf facing the meristem (adaxial) develops characteristic jigsaw-shaped epidermal cells and contains palisade parenchyma specialised for photosynthesis whereas the abaxial side contains more stomata and spongy parenchyma which performs respiration in addition to photosynthesis. Whereas the petiole is highly radialised, the blade develops a distinct medial mid-vein and lateral lamina at whose edges the adaxial and abaxial cells converge to form long marginal cells. The lateral organ vasculature is polarised with the water-conducting xylem forming adaxially and the photosynthate transporting phloem forming abaxially underneath. Floral organs also have distinct polarities whose characteristics  12  Figure 1.3 Polarities in developing leaves Scanning electron micrographs of developing leaves in Arabidopsis (A) As the leaf primordium emerges, the adaxial face is closest to the shoot apical meristem (SAM, arrow) while the abaxial faces away from the SAM. (B) The side of the outgrowing leaf closes to the stem attachment region is proximal while the region furthest away is distal. The central region is medial. Reproduced with permision from PLoS Genetics, Byrne (2006).  www.plosgenetics.org  13  vary depending on the particular floral organ. Flower morphology and development will be discussed in detail in Section 1.4.2. 1.2.3.1 Establishment abaxial/adaxial polarity Realisation of adaxial and abaxial fates is essential for proper morphogenesis of leaves. Loss of either identity function results in the formation of adaxialised or abaxialised leaves which lack blade growth and instead are radialised organs (Byrne, 2006). A major conceptual advance in plant development was put forward by Waites and Hudson (1995) who postulated that the contact between adaxial and abaxial fates at the marginal leaf cells promotes marginal meristematic activity and blade expansion. In Arabidopsis, adaxial cell fate is specified by class III homeodomain-leucine zipper (HDZIPIII) transcription factors and the AS1/AS2 module while abaxial identity is promoted by the KANADI (KAN) and YABBY (YAB) transcription factors (Bowman et al., 2003; Lin et al., 2003; Eshed et al., 2004; Kidner and Timmermans, 2007; Fu et al., 2006). AS1/AS2 establish adaxial fate by downregulation of KNOX and abaxial factors in the adaxial domain (Fu et al. 2006, Iwakawa et al., 2007). The HD-ZIPIII gene family consists of three members, PHABULOSA (PHB), PHAVOLUTA (PHV) and REVOLUTA (REV) (McConnell et al., 2001; Otsuga et al., 2001; Emery et al., 2003) whose expression parallels the flow of auxin from the tip of the developing primordia into SAM subepidermal layers (Emery et al., 2003; Heisler et al., 2005; Prigge et al., 2005). HD-ZIPIIIs promote adaxial identity through the downregulation of the KANs in the adaxial domain (Eshed et al., 2004). They themselves are downregulated abaxially via 165/166 miRNA (Emery et al., 2003; Williams et al., 2005). The three KANs (KAN1, KAN2 and KAN3) encode transcription factors of the GARP (maize Golden 2, ARR and Psr1) family (Kerstetter et al., 2004; Eshed et al., 2004). KANs, in turn, downregulate HD14  ZIPIIIs as well as AS2 in the abaxial domain (Eshed et al., 2004; Wu et al., 2008). Another set of factors which promote abaxial identity while repressing meristematic fate in leaves and flowers include the YABBY genes such as FILAMENTOUS FLOWER (FIL), YABBY2 and YABBY3 (YAB3; Sawa et al., 1999; Seigfried et al., 1999; Eshed et al., 2004). A close association exists between the specification of adaxial identity and meristematic competence: in addition to a radialised cotyledon phenotype, phv phb rev triple mutant seedlings ectopically express KANs, at the shoot apex and lack an SAM (Emery et al., 2003) Moreover, overexpression of KANs and YABBYs lead to a loss of meristematic potential (Kumaran et al., 2002; Eshed et al., 2004) suggesting that YABBYs promote lateral organ fate. In fact, YABBYs such as FIL are considered one of the earliest markers during primordium initiation (Golz and Hudson, 2002; Gordon et al., 2007). 1.2.3.2 Establishment of proximal/distal polarity Primordia amass sufficient cell numbers through an early proliferation phase. Subsequent lateral outgrowth reflects a combination of cellular proliferation and expansion;thus, the proximal distal outgrowth of an organ is shaped in large part by genes involved in the timing of each phase (Anastasiou et al., 2007). One such gene is JAGGED (JAG) which encodes a basic helix-loophelix transcription factor which maintains cell cycle activity in the distal domain of developing lateral organs (Dinneny et al., 2004; Ohno et al., 2004; Dinneny et al., 2006). Loss of function jag plants develop smaller lateral organs due to premature cellular proliferation arrest while ectopic expression of JAG can lead to excess organ growth manifested in leaves as excess blade growth along the petiole (Dinneny et al., 2004; Ohno et al., 2004). JAG also negatively regulates expression of the boundary genes along with AS1/AS2 (Xu et al., 2008). Thus, restriction of boundary gene expression may be important to promote primordium outgrowth. In addition, a 15  cytochrome P450 discovered to be a crucial mediator of cellular proliferation growth is KLUH (KLU; Anastasiou et al., 2007; Wang et al., 2008). Interestingly, KLU was originally described a decade ago as a regulator of the lateral organ-meristem boundary domain (Zondlo and Irish, 1999) again suggesting a link between an organ’s proliferative capacity and boundary specification.  1.3 FLOWERING AND FLOWERS Angiosperms are distinguished by the presence of flowers, highly specialised reproductive structures, the evolution of which heralded a massive radiation and diversification of the angiosperms. The developmental pathway to a mature flower begins with the production of a meristem off the flanks of the SAM. First, the inflorescence meristem (IM) must be programmed to make floral meristems instead of secondary inflorescence or vegetative shoots. Secondly, a series of floral homeotic genes must be activated in the floral meristem in a spatially and temporally distinct manner. Thirdly, downstream targets of the floral homeotic genes must execute a patterning program to produce the floral organs (Krizek and Fletcher, 2005).  1.3.1 Inflorescence architecture Arabidopsis begins aerial development in a vegetative growth habit (Fig. 1.1). Little internode elongation occurs between leaves, leading to the production of a compact ground-layer rosette. In response to integrated developmental and environmental cues, the primary stem bolts into a simple, indeterminate inflorescence. At the onset of the floral transition, the IM increases the rate of cell division and produces several cauline leaves which subtend secondary inflorescence shoots (Steeves and Sussex, 1989). Subsequently, the IM irreversibly commits to a continuous production of floral meristems which, like leaves, arise in a spiral phyllotaxy, developing a 16  racemic flowering architecture (Weberling, 1989). In Arabidopsis, the switch to reproductive development is accompanied by substantial internode elongation between lateral organs such that the stem bolts up from the rosette to produce a flower-bearing structure referred to as the inflorescence (Steeves and Sussex, 1989). A complex interplay of endogenous and environmental inputs converge on a group of flowering time integrators to promote the floral transition (Glover, 2007). Three key flowering time integrators are FLOWERING-TIME LOCUS T (FT), SUPPRESSOR OF OVEREXPRESSION OF CONSTANS1 (Liu et al., 2008) and LEAFY (LFY) which are all expressed at low levels in vegetative organs but are strongly upregulated in the SAM upon transition to flowering (Parcy, 2005). The flowering-time integrators activate another set of genes known as inflorescence identity genes which include the MADS-box genes AGAMOUSLIKE24 (AGL24), SOC1, FRUITFULL (FUL) and SHORT VEGETATIVE PHASE (SVP) (Ferrandiz et al., 2000; Yu et al. 2002; Michaels et al. 2003; Yu et al. 2004; Liu et al. 2007; 2008; Lee et al. 2008). Up-regulation of these genes in the IM coincides with formation of cauline leaves and associated inflorescences (Yu et al. 2002; Michaels et al. 2003). Subsequently, the IM irreversibly switches to the production of floral meristems. Unlike the IM’s indeterminate growth habit, floral meristems differentiate determinate numbers of floral organs in a series of four concentric whorls (Fig. 1.4A,B; Krizek and Fletcher, 2005). Most angiosperm flowers are subtended by a leafy organ, known as a floral bract, and thus architecturally, floral meristems can be viewed as modified axillary meristems arising from the axil of the bract (Sablowski, 2007). However, Arabidopsis, like other Brassicaceae, develop ebracteate flowers which lack visible floral bracts (Arber, 1931). Interestingly, genetic analyses suggest that the bract is patterned but its outgrowth actively repressed (Long and Barton, 2000; Dinneny et al., 2004; Ohno et al., 2004). 17  Figure 1.4 Flower development in Arabidopsis Wild type Arabidopsis flower showing concentric whorls of sepals (se), petals (pe), stamens (st) and central carpels (ca). (C) Overlapping domains of A, B and C class functions specify the floral organs. (D) Genetic network specifying the floral transition which involves the downregulation of the inflorescence identity genes, SVP, AGL24 and SOC1 by LFY and AP1, thus liberating LFY and SEP3 to activate floral homeotic genes. (D) the floral quartet model of floral organ identity where ABC dimers and trimers are stabilised through interaction with SEPs. (A-C, E) reprinted with permission from Elsevier: Developmental Biology, Lolhmann and Weigel (2002) and (D) reprinted with permission from Elsevier: Developmental Biology, Liu et al., (2009).  www.sciencedirect.com/science/journal/00121606  18  During the shift to exclusive flower production, flowering time integrators converge on another set of genes known as floral meristem identity genes. In addition to promotion of the initial floral transition, LFY is also a key mediator of floral-meristem identity. LFY encodes a plantspecific transcription factor that is expressed in the floral anlagens at stage 0, prior to any morphological sign of floral primordia formation. Together with LATE MERISTEM IDENTITY1 (LMI; Saddic et al., 2006), LFY directly activates another crucial me diator of floral-meristem identity, APETALA1 (Hong and Tucker, 1998) and its close homologue CAULIFLOWER (CAL), both MADS-box genes (Wagner et al., 1999; Ferrandiz et al., 2000a). AP1 is first expressed in stage 1 primordia which form slight bulges in the PZ and is a hallmark for commitment to floral fate (Hempel et al., 1997). AP1 and CAL, together with FUL and LMI1, feed-forward to positively enhance LFY expression in the developing floral meristem (Ferrandiz et al., 2000b; Saddic et al., 2006). At the same time, the inflorescence identity factors (SVP, SOC1 and AGL24) are downregulated to ensure that initiating primordia develop into floral meristems rather than secondary inflorescences (Fig. 1.4D, Gregis et al., 2008). Loss of function in either LFY or AP1 leads to the conversion of floral meristems into indeterminate flowering inflorescence shoots reflecting a loss of floral meristem identity (Irish and Sussex, 1990; Schultz and Haughn. 1991; Huala and Sussex, 1992; Weigel et al., 1992; Bowman et al., 1993) while over-expression of either of these genes results in the eventual conversion of the SAM into a single terminal flower (Weigel and Nilsson, 1995; Mandel and Yanovsky, 1995; Ferrandiz et al., 2000) suggesting that AP1 and LFY are both necessary and sufficient for floral fate. Indeteminancy of the inflorescence is normally maintained by an antagonistic feedback loop between TERMINAL FLOWER (TFL) and LFY and AP1 (Shannon and Meeks-Wagner, 1993) whereby LFY initially upregulates TFL in the tip of the IM which  19  then acts to restrict LFY and AP1 to the developing floral primordia (Bradley et al., 1997; Ratcliffe et al., 1999) where, in turn, they repress TFL expression (Liljegren et al., 1999).  1.3.2 Flower development Floral meristems develop into a flower bearing stalk comprised of a proximal pedicel which develops distally into a receptacle (flower base) bearing a series of floral organs. Flowers evade concise definition, reflecting the incredible diversity of extant and extinct floral architectures (Theissen and Melzer, 2007); however, all eudicots, including the Brassicaceae, exhibit standard floral organs: the sepals, petals, stamens and carpels, arranged in whorls (Fig. 1.4; Buzgo et al., 2005), In Arabidopsis, four sepals form the outer whorl followed by a second whorl of alternately spaced four petals, reflecting a tetramerous organ arrangement. Sepals are green, laminar protective structures enclosing the floral bud and attach with a broad base to the receptacle. Petals develop as a distal colourless blade that narrows into a proximal filamentous claw at the receptacle. Sepals and petals are both sterile organs and are collectively referred to as the perianth. The inner whorls consist of the reproductive floral organs. Six male reproductive organs or stamens are arranged in two clusters of the third whorl: two lateral stamens and two pairs of medial stamens. Stamens also attach to the receptacle with a filament, similar in size to the petal claw, and form four distal pollen-bearing locules collectively called the anther. Floral nectaries, small nectar-secreting glands, subtend the bases of stamens. Stamens alternate with the petals such that they develop opposite to the sepals. The central gynoecium, the female reproductive structure, sits in the fourth whorl and is formed by the fusion of two carpels attached via specialised valve margin tissue. Following fertilisation, the gynoecium elongates and matures into a seed-bearing silique, while the remaining floral organs senesce and abscise.  20  After a period of maturation, the siliques dry and shatter along valve margins, releasing seeds (Smyth et al., 1990). Floral organs, like leaves and floral meristems, begin as lateral primordial bulges; however, the floral organs initiate in a whorled rather than spiral phyllotaxy. The sepals develop first, followed by the stamens and petals, and finally the carpels; stamens and carpels mature before the petals which mature last (Smyth et al., 1990). The genetic networks controlling organ identity are relatively well understood and reflect two decades of intense research on floral homeotic mutants (Krizek and Fletcher, 2005). The actions of the floral homeotic genes, as dictated by the ABC model, describe how overlapping expression patterns of three classes of homeotic genes, the A, B and C genes, specify floral organ identity (Fig. 1.4C; Haughn and Sommerville, 1988; Bowman et al., 1991; Lohmann and Weigel, 2002). Class A genes, APETALA1 (AP1) and APETALA2 (AP2) are expressed in whorl 1 to specify sepal identity. Class A along with the Class B genes, APETALA3 (Schultz et al., 1991) and PISTALLATA (PI) are expressed in whorl 2 to pattern petals. In whorl 3, Class B genes and the Class C gene, AGAMOUS (AG), act to promote stamen identity. The fourth whorl is characterised by AG expression to form the carpels. Other than AP2, which encodes a member of the EREBP transcription factor family, all other floral homeotic genes are members of the MADS-box transcription factor family. Loss of function and over-expression studies confirmed the model and showed that floral homeotic genes influence each other’s expression. For example, AP2 is important to repress AG expression in the perianth whereas AG is important to downregulate AP1 expression in whorls 3 and 4, suggesting that class A and class C genes mutually restrict each other’s expression into the appropriate whorls (Bowman et al., 1991; Drews et al., 1991). Another set of MADS-box genes, the SEPALATTAs (SEP1-4) or Class E genes, function with the ABC genes to determine floral organ identity (Pelaz et al., 2000; Honma and Goto, 2001; Ditta et al., 2004). To explain the 21  mechanism of floral homeosis, the floral quartet model suggests that two dimers of ABCE floral homeotic proteins, bound to separate regions of DNA target sequences, interact to form a protein quartet looping the intervening DNA (Fig. 1.4F; Theissen and Saedler, 2001; Theissen and Melzer, 2007; Melzer and Theissen, 2009). Differential regulation of floral homeotic gene expression is crucial for proper floral organ development. Earlier studies and recent high-resolution expression analyses have determined the sequence and spatial patterns of their activation (Krizek and Fletcher, 2005; Wellmer et al., 2006; Urbanus et al., 2009). Besides AP1 which is expressed at stage 1 to mediate floral meristem identity, the first homeotic genes expressed are SEP1, SEP2 and SEP3 activated in late stage 2. SEP3 is a major control point for flower differentiation by promoting the initiation of AP3 and PI expression in stage 3 primordia in conjunction with LFY and UNUSUAL FLORAL ORGANS (UFO), an F-box protein that is itself up-regulated by AP1 (Drews et al., 1991; Flanagan and Ma, 1994; Goto and Meyerowitz, 1994; Jack et al., 1992; Savidge et al., 1995; Wilkinson and Haughn, 1995; Ingram et al., 1995; Samach et al., 1999; Parcy et al., 1998; Ng and Yanovsky, 2000; Kaufmann et al., 2009; Liu et al., 2009). AP3 positively regulates itself and PI, while negatively regulating AP1 (Sundstrom et al., 2006). In the inner whorls of stage 3 flowers, LFY and WUS turn on AG expression (Lohlmann et al., 2001) to promote stamen and carpel formation whereas AP2, AP1 and SEP3, in conjunction with the LEUNIG/SEUSS (LEU/SEU) repressor complex, downregulate AG in the perianth (Drews et al., 1991; Sidhar et al., 2006). AG, in turn, is necessary to restrict AP1 expression, initially activated throughout the primordium, to the first two whorls (Gustafson-Brown et al., 1994). In addition, AG represses and finally extinguishes WUS expression in the central whorl during floral development, thus ensuring floral determinacy through the complete consumption of floral meristem cells (Lenhard et al., 2001). 22  1.3.3 Execution of floral homeotic programs 1.3.3.1 Polarity in flowers and floral organs Flowers develop asymmetries with respect to the apical meristem while the floral organs develop polarity with respect to the floral meristem. Many players previously described (section 1.2.3.1) are also involved in flower development; however, the role of polarity throughout the floral meristem is not well understood. Prior to floral organ development, the abaxial factors, FIL and YAB3 are expressed in the abaxial domain of the floral primordia (Siegfried et al., 1999; Sawa et al., 1999) while the HD-ZIPIII, REV, is expressed in the adaxial domain (Otsuga et al., 2001). The adaxial/abaxial factor juxtaposition in flowers is necessary for flower development as loss of function in either polarity leads to a loss of receptacle formation and the growth of filamentous structures rather than flowers (Talbert et al., 1995; Siegfried et al., 1999; Sawa et al., 1999; Chen et al., 1999; Otsuga et al., 2001). Perhaps due to this, few floral mutants have been described which show aberrations in whole flower polarity. One such mutant is pressed flower (prs) which specifically lacks lateral sepals (Matsumoto and Okada, 2001). Following initiation of the floral organs in stage 3, adaxial and abaxial factors are expressed in their respectve domains of the floral organ primordia. Given the alternating whorled architecture of Arabidopsis flowers, this means that adaxial and abaxial regions of floral organ primordia are often adjacent yet offset from one another. 1.3.3.2 Boundary specification in flowers Boundary specification in flowers involves separation between floral organs and between whorls. The CUC genes, in particular CUC2, play important roles in floral organ separation as well a role in petal identity (Aida et al., 1997; Sakamoto et al., 2001; Baker et al., 2005). The aforementioned RBE and PTL are important for separation between sepals (Brewer et al., 2004; 23  Takeda et al., 2004). PTL likely functions in parallel with the CUCs and is negatively regulated by auxin, AS2, JAG and NUB (Brewer et al., 2004; Xu et al., 2008). RBE acts downstream of the CUCs and involves suppression of AG in the perianth (Nole-Wilson and Krizek, 2006). UFO and SUPERMAN (SUP), encoding a bZIP protein, are essential to maintain third to fourth whorl boundaries (Sakai et al., 1995; Wilkinson and Haughn, 1995; Lee et al., 1997; Sakai et al., 2000). SUP expression is promoted by AP3, PI and AG (Sakai et al., 2000). Many other floral mutants lead to fusion between organs and whorls, suggesting that boundary formation and maintenance is a key event in flower development subject to tight genetic regulation involving multiple players.  1.4 BLADE-ON-PETIOLE1 (BOP1) AND BOP2 GENES 1.4.1 Experimental rationale Our research group, concomitantly with others, described the isolation and characterisation of BLADE ON PETIOLE1 (BOP1) and BLADE-ON-PETIOLE2 (BOP2) a pair of homologous genes that regulate leaf and flower patterning (Ha et al., 2004; Norberg et al., 2005; Hepworth et al., 2005). Initial interest in BOP genes arose due to their homology with NPR1 (NONEXPRESSOR OF PR GENES1) a key mediator of a plant defense response known as Systemic Acquired Resistance (SAR). Neither bop1-3 nor bop2-1 T-DNA insertion lines showed obvious phenotypes. Double bop1 bop2 mutants, however, displayed altered floral and leaf patterning. Regardless of homology to NPR1, bop1 bop2 mutants showed no alteration in disease resistance or suseptibility compared to wild type when challenged with Pseudomonas syringae maculicola AS4326 (Hepworth et al., 2005).  24  1.4.2 blade-on-petiole1 (bop1) and bop2 mutants 1.4.2.1 BOP1 and BOP2 repress growth in the proximal leaf Arabidopsis leaves show dramatic proximal-distal asymmetry (Section 1.2.3; Fig. 1.5A). bop1 bop2 leaves grow much longer than wild type and exhibit expansion of blade-like projections from the petiole of rosette and cauline leaves (Fig. 1.5B-D, F) suggesting that BOP1 and BOP2 function includes repression of leaf elongation and repression of blade expansion in proximal regions. Ha et al (2007) demonstrated that excess blade tissue originated from ectopic meristems formed on the adaxial sides of the petiole. Excess leaf length may occur due to a delayed floral transition, as bop1 bop2 mutants bolt later than wild type (Norberg et al., 2005). It is unknown whether increased leaf length occurs due to extended proliferative growth and/or enhanced cellular expansion. Plants without AS1 or AS2 function also develop leafy petioles and lobed leaves which is correlated with ectopic expression of KNOX genes, BP, KNAT2 and KNAT6 (Byrne et al., 2000; Ori et al., 2000; Semiarti et al., 2001). These KNOX genes are also ectopically expressed in bop1 bop2 leaves (Ha et al., 2004; Norberg et al., 2005; Ha et al., 2007). Double mutants between BOP1 and AS1/ AS2 show a synergistic relationship (Ha et al., 2004), suggesting that BOPmediated repression of KNOX genes may involve an independent, parallel pathway to that employed by AS1 and AS2. Ha et al. (2007) showed no change of BOP expression in as1 or as2 mutants; however, AS2 expression was reduced in bop1 bop2 implying a role for BOP in upregulation of AS2. Ectopic expression of either BOP gene leads to dwarfed, bushy plants with downward pointing siliques and smaller petioles. Many of these phenotypes may be due to repression of KNOX genes and increased expression of AS2 (Ha et al., 2007). In addition to repressing KNOX, AS1 and AS2 are key mediators of adaxial identity (Section 1.2.3.1). Ha et al., (2007) showed that BOP1 and BOP2 similarly promote adaxial identity as bop1 bop2 mutant 25  Figure 1.5 Leaf phenotypes of bop1 bop2 and wild type (Col-0) Twenty-eight-day-old plants are shown for the wild type (A, Col-0) and bop1 bop2 (B). Scale bar, A-B 2 cm. (C) Leaf series from 3-week-old wild-type and bop1 bop2 (bottom row) plants. (D) Cauline leaves from wild-type (left) and bop1 bop2 (middle and right) plants. Hepworth et al. (2005).  www.plantcell.org. Copyright American Society of Plant Biologists.  26  petioles showed features of abaxialisation in the vasculature including phloem adaxially surrounding the xylem. This was correlated with broadened FIL and KAN expression encroaching into the adaxial side of older (but not younger) petioles corresponding to regions of ectopic blade outgrowth. 1.4.2.2 BOP1 and BOP2 repress floral bract and abaxial flower growth Often, bop1 bop2 flowers are subtended by a floral bract or reduced bract-like organs (Fig. 1.6Ai-j; Fig. 1.6Ba-c), suggesting that BOP1 and BOP2 are part of the suite of genes involved in repressing cryptic bract outgrowth. Interestingly, ectopic expression of JAG from the dominant Jag-5D allele also results in bract growth as well as additional blade formation along the petiole, similar to bop1 bop2 (Dinneny et al., 2004; Ohno et al., 2004). Triple mutant analyses showed that functional JAG alleles are required for bract outgrowth observed in leafy (lfy), apetala1 (Hong and Tucker, 1998) and apetela2 (ap2) mutants (Dinneny et al., 2004; Ohno et al., 2004). However, Norberg et al. (2005) showed that, although JAG is expressed ectopically in the bracts of bop1 bop2 flowers, introduction of the jag allele did not suppress bract formation in a bop1 bop2 jag triple mutant. The authors suggest that in the absence of JAG, its close homolog JAGGED-LIKE (now called NUBBIN) is able to replace JAG function in bract promotion, although clearly then, NUBBIN (Dinneny et al., 2006) does not perform this function in jag lfy, jag ap2 and jag ap1 mutants. This hypothesis awaits examination of a bop1 bop2 jab nub quadruple mutant. bop1 bop2 flowers also develop extra floral organs compared to wild-type. Replacing the abaxial sepal in the first whorl are two organs which initially develop as sepals but develop more petal identity and grow outward like wings (Fig. 1.6Ad-h). These organs are slower to develop than the normal sepals (Fig. 1.6Bd-f). The first whorl in bop1 bop2 has a  27  Figure 1.6 Comparison of wild-type and bop1 bop2 floral phenotypes. (A) Morphology of mature flowers: (a) wild-type (Col-0); (b) bop1; (c) bop2; and bop1 bop2 (d) abaxial side; (e) adaxial side; (f) abaxial view of flower with abaxial first-whorl organs removed; (g) abaxial view of young flower showing retarded growth of abaxial organs in the first-whorl; (h) example of fused abaxial first-whorl organs with sepal-petal characteristics. Scale bar, a-e and f-h, 1 mm; (i-k) Bracts and reduced bract-like structures that subtend flowers. Scale bar, 0.5 mm. (B) SEMs of wild-type and bop1 bop2 inflorescence apices depicting inflorescence meristems, floral meristems, and flowers: (a) wild-type inflorescence apex, asterisk marks abaxial sepal; (b) bop1 bop2 inflorescence apex, arrows indicate floral bracts; (c) stage 2 floral primordia with subtending bract (arrow), FM, floral meristem; (d) stage 5 flower with subtending bract and five organs in the sepal whorl such that the adaxial sepal is in the same position as in wild-type. An extra stamen primordium occurs between the two abaxial sepal-whorl organs (asterisks); (e) stage 7 flower showing retarded development of sepals. Smaller abaxial firstwhorl organs are marked by asterisks; (f) example of flower with abaxial first-whorl organs that are half-sepal (bumpy cells) and half-petal (smooth inner cells). Scale bars a-d, 50 µm; e, 100 µm; f, 500 µm. Hepworth et al. (2005).  www.plantcell.org. Copyright American Society of Plant Biologists. 28  pentamerous arrangement; however, the three normal sepals tend to cluster together adaxially due to excessive growth of the abaxial petalloid sepals. Also, stamen number is often increased from six to seven with the additional stamen formed in the middle of the abaxial side between the petalloid sepals (Fig. 1.6B). These aberrations in bop1 bop2 flowers suggest that BOP proteins act to ensure proper organ number and identity in flowers and these functions appear to be restricted to repressing growth on the abaxial face of the flower. It is unknown whether abaxial promoting factors, FIL and YAB3 are also ectopically expressed in the adaxial domain of flowers as in leaves. BOP1 and BOP2 function with PERIANTHIA (PAN), which encodes a bZIP transcription factor (Chuang et al., 1999) to control certain aspects of floral patterning (Hepworth et al., 2005). pan mutants form five organs in the first three whorls reflecting a pentamerous floral symmetry (Running et al., 1995). Triple bop1 bop2 pan mutant flowers show a fusion of bop1 bop2 and pan phenotypes and develop a pentamerous sepal whorl, tetramerous petal whorl and a decreased number of stamens in the third whorl (Hepworth et al., 2005). Importantly, the floral phenotypes of bop1 bop2 do not resemble those of as1 or as2, which are characterised by shorter and narrower sepals and petals (Ori et al., 2000; Byrne et al., 2002; Xu et al., 2008) suggesting that the roles of AS1/AS2 and BOP1 and BOP2 may differ in the flower. 1.4.2.3 BOP1 and BOP2 promote abscission A striking phenotype observed in bop1 bop2 plants is a complete loss of floral organ abscission (Fig. 1.6G). Normally in Arabidopsis flowers, sepals, petals and stamens senesce and are shed (abscised) from the plant shortly following fertilization. However, in bop1 bop2 plants, the floral organs remain firmly attached to the base of the elongating silique indefinitely, suggesting that the BOP1 and BOP2 gene products promote floral organ abscission.  29  1.4.3 Thesis objectives 1.4.3.1 Contributions of BOP1 and BOP2 to abscission Following initial characterisation of bop1 bop2, I chose to investigate the contribution of the BOP proteins to floral organ abscission. I began with a thorough phenotypic characterisation of the lack of floral organ abscission in bop1 bop2 mutant plants. The genetic relationship of BOP1 and BOP2 with other abscission related genes was also examined. Results from this project were published in Hepworth et al. (2005) and McKim et al. (2008) and are presented in Chapter 2. 1.4.3.2 Contributions of BOP1 and BOP2 to nectary development During investigation of the loss of floral organ abscission in bop1 bop2, I discovered that bop1 bop2 plants also lacked full elaboration of nectary glands. This phenotype was fully characterised and genetic interactions with other regulators of nectary development. Much of this work was published in McKim et al. (2008) and is presented in Chapter 3. 1.4.3.3 Suppressors of bop1 bop2 I isolated two recovered abscission lines while conducting a suppressor screen in the bop1 bop2 background. Preliminary genetic and phenotypic analyses are described in Chapter 4. 1.4.3.4 Roles of BOP1 and BOP2 in floral meristem identity Certain features of the bop1 bop2 phenotype, including the development of the floral bract, suggested that floral meristem identity was compromised. In collaboration with Dr. Shelley Hepworth, I also examined the contribution of BOP1 and BOP2 to floral meristem identity. These data will be submitted in Xu et al (2009) and my contribution is discussed in Chapter 5.  30  1.4.3.5 Biochemical mechanism and phylogeny of BOP1 and BOP2 To begin to assess the conservation of BOP roles across land plants, I conducted a phylogenetic analysis of the NPR1 protein family in land plants. Furthermore, the biochemical interactions of BOP1 and BOP2 were investigated using bimolecular fluorescent complementation. These data are presented in Chapter 6.  The following sections (1.5-1.6) will summarise the current literature pertinent to these topics:  1.5 ABSCISSION BIOLOGY 1.5.1 Biological overview of abscission Abscission is a type of cell separation leading to organ shed from the plant body and is an active, developmentally-regulated process (Patterson, 2001; Roberts et al., 2002; Lewis et al., 2006). Once Arabidopsis flowers reach anthesis, defined as the ability to self-fertilise, floral organs, excluding the gynoecium, senesce and abscise (Patterson, 2001). Fertilisation is not a prerequisite for abscission as sterile lines such as bel1 abscise floral organs (Modrusan et al., 1994). Abscission is not an unavoidable consequence of senescence: for example, Arabidopsis rosette leaves senesce but do not abscise. Abscission of floral organs may increase pollinator access to the carpel, eliminate dead weight and remove dead and decaying tissue that would otherwise serve as ideal substrate for pathogen establishment. Given the role of petals in pollinator attraction, petal abscission may also discourage pollinators from visiting already fertilised flowers thereby ensuring that they continually visit flowers at the appropriate stage for cross-pollination. The key step in abscission is a loss of adhesion between the leaving organ and the main plant body. Targeted degradation of the middle lamellae between cell walls is crucial to weaken 31  intercellular attachment (Sexton and Roberts, 1982). Separation occurs within a specialised region known as the abscission zone (AZ) which generally forms at the boundary between plant body and organ. In addition to secretion of middle lamellae-degrading enzymes, AZ cells in the separation layer enlarge and round-up during cell separation (Sexton et al., 1985). Following organ shed, the exposed surface on the plant body differentiates a protective surface layer. Investigation of abscission phenomena dates back at least to Darwin (Darwin, 1877) and has involved study of numerous plant species, notably Phaseolus vulgaris (bean) and Lycopersicon esculentum (tomato). Physiological abscission experiments are rarely undertaken in Arabidopsis given the small size of the AZs; however, a recent laser capture microdissection of Arabidopsis stamen AZs by Cai and Lashbrook (2008) has investigated trends in gene expression leading up to the abscission event which will undoubtedly shed light on the abscission process.  1.5.2 Anatomy and ontogeny of abscission zones Separation of organs from the plant body occurs at anatomically distinct cell files called abscission zones (AZs). Cells in the AZ resemble meristematic cells in being small, isodiametric and cytoplasmically dense in comparison to surrounding tissues (Fig. 1.7). Although adventitious AZs are known to form in response to pathogen attack and other adverse conditions, developmentally prescribed AZs occur in canalised positions often demarcated as morphologically distinct prior to abscission and are commonly formed at the boundary between the lateral organ and main plant body (Addicott, 1982; Taylor and Whitelaw, 2001; Roberts et al., 2002). In Arabidopsis, floral organ AZs develop as a four to six cell layer interface between the floral organs and the receptacle (Bleecker and Patterson, 1997). Vascular traces serving lateral organ are obliged to travel through the AZ. Commonly, these strands anastomose near and 32  Figure 1.7 Floral organ abscission zones (A) Colour-coded model of an Arabidopsis flower with a petal and sepal removed to show the inner organs: blue, sepals; green, petals; orange, stamens; red, carpels. (B) Base of an Arabidopsis silique following floral organ abscission. The abscission zone (AZ) scars left by each organ are as indicated. (C) Toluidine-blue stained longitundinal section of an Arabidopsis flower just after anthesis. The AZ cells are small, densely cytoplasmic whereas surrounding lateral organ and receptacle tissues are highly enlarged and vacuolated. (A-B) Reprinted with permision from The Company of Biologists: Development, Lilijegren et al. (2009).  dev.biologists.org  33  within the AZ and are considered to have a reduced, juvenile character within the zone (Addicott, 1982). In Arabidopsis, highly reduced vascular strands serving the floral organs go through the centre of the AZs (Douglas and Riggs, 2005; Patterson, 2001). Much work on the genetics controlling AZ formation has been done in tomato which forms an AZ in the middle of the flower pedicel, creating a knuckle or joint, which when activated leads to whole flower drop. Formation of tomato pedicel AZs is dependent on the genotype of the L3 cell layer suggesting that communication from inner to outer cell layers is important to signal for AZ differentiation (Symkowiak and Irish, 1999) and agrees with studies in bean that show AZ formation occurs in response to a signal from the stele (Thompson and Osborne, 1994). Several tomato mutants which lack pedicle AZs have been described and the responsible loci cloned. Once such gene is JOINTLESS (JTL), a MADS-box gene essential for the development of pedicel AZs in addition to promotion of inflorescence meristem identity (Mao et al., 2000; Szymkowiack and Irish, 1999, 2006). JTL is not expressed in the developing AZ and is presumed to control AZ development indirectly through its effects on meristem identity (Szymkowiak and Irish, 1999, 2006). The closest homologue to JNT in Arabidopsis is SVP, an inflorescence identity gene (see Section 1.4.1); svp plants do not display abscission deficits but rather precociously enter reproductive development (Hartmann et al., 2000). Another gene implicated in tomato pedicel AZ development, is LATERAL SUPPRESSOR (LS) which encodes a GRAS-like transcription factor (Malayer and Guard, 1964; Schumacher et al., 1999). Interestingly, ls mutants, in addition to lacking petiole AZs, also fail to form axillary buds due to a loss of meristematic character in the leaf axils (Malayer and Guard, 1964). The Arabidopsis homologue, also called LATERAL SUPPRESSOR (LAS), has a similar function in patterning axillary meristems and las mutants were reported to have delayed petal abscission (Greb et al., 2003). LAS shows boundary-specific expression and accumulates very early in a strip between 34  the SAM and the leaf and floral meristem primordia (Greb et al., 2003). However, in both ls and jnt tomato mutants, leaf AZs are normal, suggesting that JNT and LS mediate pedicel-specific effects (Roberts et al., 2002). Two major possibilities exist for the origin of AZ cells. Given the anatomical similarity with meristematic cells and reduced vasculature, AZs cells are traditionally considered to reflect arrested development of meristematic cells that fail to enlarge and vacuolate along with the rest of the organ (Addicott, 1982; Bleecker and Patterson, 1997). However, this notion is not consistent with examples of AZ differentiation in the literature. Significantly, McManus et al (1998) showed that AZ cells in bean petioles developed from mature cortical cells suggesting that AZ formation results from a transdifferentiation event rather than from meristematic arrest. Furthermore, AZs, unlike peripheral meristem cells, are primed for a massive secretion event and must establish a specific middle lamellae composition in the separation layer that will appropriately respond to enzymatic degradation. And lastly, studies indicate that both developmental and adventitious AZs develop in regions flanked on both sides by relatively mature tissue (McManus, 1994). That said, many features of AZ behaviour are shared by peripheral meristematic cells, including proliferative capacity and secretion of enzymes which aid in cell wall loosening. These two views can be reconciled by the notion that AZ cells reflect a reversion to a meristematic-like state during and/or following organ differentiation. Consistent with a proliferative capacity, cells in the AZ commonly divide to give rise to several tiers of cells as abscission approaches (Gawadi and Avery, Jr., 1950; Addicott, 1982) although cell division is not always a precursor to AZ formation (McManus et al., 1998). Until recently, it was unclear exactly when floral organ AZs appear in Arabidopsis; however, new data from Gomez-Mena and Sablowski (2008) suggests that AZ cells start to develop as early as stage 8, just prior to petal outgrowth, and are recognisable as cell files 35  around stage 13, corresponding to anthesis (competency for fertilisation; stages according to Smyth et al., 1990). These cells are flanked by fully vacuolated differentiated tissue from the floral organ distally and the receptacle proximally, again substantiating that they are not simply PZ meristematic cells ‘left behind.’ Furthermore, their development corresponds with expression of cell cycle genes showing activation in the boundary regions during these floral stages (Martinez et al., 1992; Gaudin et al., 2000; Imai et al., 2006). Many mutants exhibiting delayed floral organ abscission also show a prolonged juvenile phase of the flower (Fernandez et al., 2000; Bleecker and Patterson, 2004; Kandasamy et al., 2005a,b; Adamcyzk et al., 2007; Cai and Lashbrook, 2008) hinting that AZ formation may occur due to a developmentally responsive signal; however, it is unclear whether these mutants are delayed in AZ formation or AZ activation (see below). Cai and Lashbrook (2008) suggest a relationship between developmental transitions and AZ development which is attractive given the association between JNT, LS and transition to inflorescence identity. AZs occur at the boundary regions between floral organs and the receptacle but it is unclear whether the CUCs or other boundary-specific genes contribute to AZ differentiation. Mutations in HAWAIIAN SKIRT, encoding an F-box protein, leads to fusion between sepal margins and a delay in abscission but the AZ anatomy appears intact (Gonzalez-Carranza et al., 2007).  1.5.3 Activation of abscission Besides their anatomical distinctiveness, AZ cells are also physiologically primed to respond to signals promoting abscission. During organ shed, at least three major events occur: the release of middle lamellae and cell wall degrading enzymes; the secretion of signal peptides to promote  36  final cell separation; and the enlargement of the abscission zone cells. Abscission occurs along two distinct cell files within the AZ cell tiers known as the separation layer. Mechanisms ensuring that separation takes place within the proscribed separation layer, and not between other cell files of the AZ, are not well understood 1.5.3.1 Enzymes Abscission involves massive secretion of cell wall- and middle lamellae- degrading enzymes. During the lead up to abscission, AZ cells often accumulate starch granules and exhibit enhanced trans-golgi network vesicle formation which may reflect preparation for the secretion event (McManus et al., 1994; Lilijegren et al., 2009). Importantly, breakstrength profiles, which measure the force required to shed an organ, commonly show a gradual weakening of intercellular adhesion prior to organ detachment (del Campillo and Bennett, 1996; Bleecker and Patterson, 1997) showing that abscission occurs in progressive stages. Temporal gene profiling in stamen AZs also supports this notion (Cai and Lashbrook, 2008). Intercellular adhesion occurs from crosslinking between polymers of the middle lamellae, a structure shared between adjacent cell walls (Jarvis et al., 1993). Composition of the middle lamellae is mostly pectin, also a major constituent of the primary cell wall (Carpita and Gibeaut, 1993). Pectin of the middle lamellae is enriched in homogalacturonan (HG; Waldron and Brett, 2007) although epitope studies show that middle lamella composition is considerably heterogenous (Knox et al., 1990). In fact, most adhesion strength is provided by crosslinks within the middle lamella found at tricellular junctions between daughter cells rather than the directly appressed sides (Waldron and Brett, 2007). A diverse suite of enzymes which act on the middle lamellae and the primary cell wall are expressed before, during and after abscission, including pectin modification enzymes, 37  expansins and cellulases (Roberts et al., 2002). Individual contributions to reduced cellular adhesion are unclear as plants lacking function in a single enzyme generally do not show aberrant cell separation, indicating probable redundancy (Lewis et al., 2006). However, polygalacturonidases (PGs), which catalyze hydrolysis of homogalacturonan, in particular, may play a crucial role in several cell separation processes (Tucker et al., 1984; Taylor et al., 1993; Biely et al., 1996; Kalaitzis et al., 1997; Kim et al., 2006; Gonzalez-Carranza et al., 2002; 2007; Jiang et al., 2008; Ogawa et al., 2009). Although delayed, floral organ abscission does occur in Arabidopsis mutants with reduced PG expression, suggesting that PG itself is not essential for eventual abscission (Gonzalez-Carranza et al., 2007; Ogawa et al., 2009). Histological, microscopical and gene expression evidence suggests that the primary cell wall is also profoundly modified during abscission (Clements and Atkins, 2001; Roberts et al., 2002; Lee et al., 2008). This may reflect a combination of functions to weaken cross-links between the cell wall and middle lamellae and to promote cell wall expansion in the distal cell separation layer during organ shed. 1.5.3.2 Separation signals Classic exposure experiments suggested that ethylene promotes while auxin inhibits abscission (Abeles and Rubenstein, 1964; Abeles, 1967; Addicott, 1982; Osborne et al, 1989; Roberts et al., 2002). Several Arabidopsis mutants aberrant in ethylene perception or signalling show delays in floral organ abscission (Patterson and Bleecker, 2004). Although considerably delayed, floral organ abscission occurs in ethylene-insensitive mutants of Arabidopsis, suggesting that ethylene signalling is important for timing rather than AZ formation (Patterson and Bleecker, 2004). The relationship between auxin and abscission is less clear but a dual role has been proposed. Much literature supports the notion that auxin delays abscission by antagonising ethylene. However, 38  as abscission approaches, auxin potentiates, rather than inhibits, ethylene’s promotive effect (Addicott, 1982). Auxin Response Factors (ARFs), transcription factors activated by auxin signalling, are involved in mediating this latter activity (Ellis et al., 2005; Okushima et al., 2005). Both auxin and ethylene signalling mutants eventually abscise, suggesting that these hormones are abscission rate regulators rather than essential signals for organ shed; however, this does not preclude a role for ethylene and auxin in modulating the timing of AZ formation in addition to AZ activation. In the past few years, numerous floral organ abscission mutants have been reported that appear independent of defects in the ethylene pathway. The delayed abscission (dab) mutants all show different degrees of delayed abscission (Patterson and Bleecker, 2004) but to date, the genetic identity of these loci is unknown. Plants carrying mutations in genes encoding the actinrelated proteins ARP4 and ARP7 (Kandasamy et al., 2005a; Kandasamy et al., 2005b) also show delayed abscission which may indicate a role for chromatin remodelling in abscission activation. Investigations of a fascinating mutant, inflorescence deficient in abscission (ida) has greatly contributed to our understanding of signals absolutely required for abscission. Similar to bop1 bop2, ida mutants retain floral organs indefinitely (Butenko et al., 2003). AZs form properly and the middle lamellae begins to dissolve in ida; however, just prior to organ shed, cellular attachments reform, suggesting that the final cell separation event is blocked (Butenko et al., 2003). Cloning of IDA revealed that it encodes a signal peptide (Butenko et al., 2003). When ectopically overexpressed, IDA leads to dramatic premature floral organ abscission as well as pronounced abscission of cauline leaves, an event not observed in Arabidopsis (Stenvik et al., 2006) suggesting that IDA acts as a positive signal at a critical check point during the abscission program. IDA belongs to a family of IDA-LIKE (IDL) signal peptides (Butenko et al., 2003). Stenvik et al., (2008) showed that a conserved small peptide sequence on the C-terminus, called 39  the ‘EPIP’, is functionally redundant between family members suggesting that the IDA and IDLs are mainly controlled through differential expression. IDA may act as a ligand for a pair of receptor-like kinases, HAESA (HAE) and HAESA-LIKE2 (HSL2) which activate a conserved MAPK signalling cascade essential for abscission to occur (Stenvik et al., 2008; Cho et al., 2008). 1.5.3.3 Cellular enlargement Very little is known about the regulation of cellular enlargement during abscission. The cell wall degrading enzymes expressed in AZ may contribute to separation layer expansion (see section 1.6.3.1). AZs cells are considered traditional type II cells that expand isotropically in response to ethylene treatment (Osbourne, 1989). It is tempting to speculate that ethylene accelerates abscission through this mechanism; however, it is unclear what contribution cellular expansion makes to the cell separation event, although a suggested function is severance of the vascular strands (Roberts et al., 2002). One of the dab mutants, dab2-1, shows aberrant cellular expansion but it is unknown if this contributes to the delayed abscission phenotype (Patterson and Bleecker, 2004).  1.5.4 Abscission and protective layers The departing organ leaves an AZ scar on the main plant body - a newly exposed surface which must differentiate a protective layer to avoid opportunistic pathogen invasion. In fact, defenserelated genes are expressed throughout the abscission process presumably to guard against such infection (del Campillo and Lewis, 1992). In herbaceous species, such as Arabidopsis, the protective surface laid down on the newly exposed plant body surface is reported to involve the deposition of suberin, cutin and lignin (Addicott, 1982; Pollard et al., 2008). Direct evidence for suberin involvement in Arabidopsis abscission scars was recently described by Franke et al 40  (2009) who showed that DAISY, an enzyme involved in the production of suberin is specifically expressed in floral organ AZs. It is unclear whether protective surfaces are laid down before or concomitant with abscission; however, based on enzyme expression and epitope analysis, lignin and suberin production and deposition may commence in the distal AZ before organ shed (Lee et al., 2008; Franke et al., 2009). AZ cells also continue isotropic expansion following organ shed such that nascent AZ scars on the receptacle form obvious enlarged clusters of bulbous cells.  1.5.5 Abscission and other cell separation processes Cell separation has other roles throughout the plant life cycle including seed germination, lateral root emergence, formation of stoma, and pollen tube growth (Roberts and Gonzalez-Carranza, 2007). The mechanics underlying the variety of cell separation events may be analogous to one another although the pattern of genetic regulation of separation zone patterning and activation are likely divergent (Roberts et al., 2002). Floral organ abscission does not involve any programmed cell death of abscission zone layers but an activity similar to sloughing of cell layers. In this manner, abscission is similar to other cell separation processes such as lateral root emergence and pollen tube growth. Dehiscence is involved in opening of a structure and may occur due to abscission between cell layers and so is often considered as an abscission event. However, numerous instances of dehiscence may also involve cell death and cell rupture, not characteristics associated with abscission. In Arabidopsis, the silique shatters, or dehisces, to release mature seeds. Separation occurs along the valve margins where the carpels fuse. Significant research has lead to an improved understanding of the genetic players involved in valve margin specification (Girin et al., 2009). Many important parallels may exist with the floral organ  41  abscission zones, including the involvement of PGs and MADS-box proteins (Lilijegren et al., 1999; Ogawa et al., 2009).  1.6 BOP1 AND BOP2 SIGNALLING BOP1 and BOP2 are part of the NPR1 (NON-EXPRESSOR OF PR1) protein family, a sixmember group in Arabidopsis which is characterised by two protein-protein interaction domains, the N-terminal associated BTB/POZ domain and the more C-terminal localised ANKYRIN (ANK) repeat domain (Cao et al., 1997; Aravind and Koonin, 1999). The BTB/POZ domain was initially described in Drosophila melanogaster as bric-à-brac, tramtrack and broad complex (BTB) transcription regulators (Zollman et al., 1994) and in many poxvirus zinc finger proteins (POZ; Bardwell and Triesman, 1994). A variety of roles has been postulated for this domain but an emerging model suggests that it is important for dimerisation with other BTB and non-BTB proteins and for interactions with CUL3-SKP1-CULLIN-F-Box (SCF)-like E3 ubiquitin ligase complex (Krek, 2003). ANK domains are one of the most common eukaryotic protein motifs and are involved in protein-protein interactions to mediate diverse activities (Bork, 1993; Mosavi et al., 2004). In their comprehensive analysis of BTB/POZ domain-containing proteins, Stogios et al (2005) found examples of BTB/POZ –ANK domain proteins in all species examined although the relative orientation of the domains varied.  1.6.1 NPR1 signalling pathway NPR1 is a positive regulator of systemic acquired resistance (SAR), a plant immune response induced following a local infection (Cao et al., 1994; Delaney et al. 1994; Shah et al. 1999; Glazebrook et al. 1996). Both BTB/POZ and ANK protein-protein interaction domains are essential for NPR1 promotion of SAR (Ryals et al., 1997; Cao et al., 1997; Rochon et al., 2006). 42  By enhancing the transcription of a suite of PATHOGENESIS-RELATED (PR) genes throughout the plant, SAR enables distal tissues of the plant to resist subsequent pathogen attack. Although the function of most PR genes is unknown, they are believed to confer some measure of resistance to subsequent pathogen attack through antimicrobial activity (van Loon and VAN STRIEN, 1999). Accumulation of salicylic acid (SA) during SAR at the site of infection as well as in distal tissues (Malamy et al., 1990; Metraux et al., 1990; Rasmussen et al., 1991) is necessary and sufficient for the induction of PR genes (Gaffney et al., 1993; Delaney et al., 1994; Lawton et al., 1994). SA treatment or pathogen attack prompts the nuclear localisation of NPR1 (Kinkema et al., 2000), an event essential for elevated PR gene expression (Mou et al., 2003). Like other members in the NPR1-like protein family, NPR1 is characterised by two aforementioned protein-protein interaction domains but lacks a recognisable DNA binding domain, suggesting that it promotes PR transcription through interaction with another transcription factor. Consistent with this notion, NPR1 is associated with TGACG sequencespecific binding transcription factors (TGAs) known to bind the as-1 (activation sequence-1)type cis-element in PR genes promoters (Zhang and Singh, 1994; Zhang et al., 1999; Depres et al., 2000; Zhou et al., 2000; Kim and Delaney, 2002). Loss of function triple tga2 tga5 tga6, but not single mutants, shows phenotypes similar to npr1 (Zhang et al., 2003) highlighting the convergence of NPR1 action on a set of redundant TGA transcription factors. Extensive research into the biochemical behaviour of NPR1 during SAR has led to the development of the oligomer-monomer switch model (Fig. 1.8). The current working model for NPR1 posits that accumulation of salicylic acid (SA) during SAR causes a reductive shift in cellular redox balance prompting NPR1 to preferentially localise to the nucleus where it interacts with TGAs to activate transcription of PR genes (Dong, 2004). A key tenet of this model is that under noninductive conditions, interprotein disulphide bridges form amongst NPR1 polypeptides 43  Figure 1.8 NPR1 signalling pathway Under non-inductive conditions, NPR1 forms an oligomer in the cytosol stabilized through interprotein disulphide bonds. Following pathogen attack, accumulation of SA promotes a more reducing environment leading to the reduction of the disulphide bonds to thiols. NPR1 monomers are now free to translocate to the nucleus where they interact and activate TGA transcription factors bound to PR gene promoters. Some TGA factors, such as TGA1 must also themselves be reduced to bind with NPR1. Reprinted with permission from Elsevier: Current Opinion in Plant Biology, Pieterse and Van Loon, (2004).  www.sciencedirect.com/science/journal/13695266  44  acting to sequester them as oligomers in the cytoplasm. SA-derived reductive shift reduces these bonds to thiol groups, thus freeing NPR1 monomers for nuclear translocation. This oligomermonomer switch is dependent on a series of conserved cysteine residues. In particular, reduction of Cys82 and Cys216 are essential for SA-mediated nuclear localisation of NPR1 (Mou et al., 2003). Arabidopsis contains eight TGAs which make up the D group of Arabidopsis basic region/ leucine-zipper (bZIP) – type transcription factors (Schultze et al., 2008). Yeast twohybrid results suggest that NPR1 specifically interacts with TGA 2, 3, 6 and 7 (Zhang et al., 1999; Despres et al., 2000; Zhou et al., 2000; Hepworth et al., 2005). Early studies demonstrated that the ANK domain was essential and sufficient for NPR1-TGA interaction (Cao et al., 1997). Further investigations on the association between NPR1 and TGA2 suggest that the complex functions as a SA-dependent enhancesome on the PR promoter (Fan and Dong, 2002; Johnson et al., 2003). In contrast to SA-mediated reduction of Cys82 and Cys216, enhancesome transactivation of TGA2 by NPR1 in vivo hinges on the SA-dependent oxidation of two Cterminal cysteine residues on NPR1, Cys521 and Cys529 (Rochon et al., 2006). Furthermore, NPR1 oligomerisation is itself promoted by SA via GSNO-mediated N-nitrosylation of Cys156 (Tada et al., 2008). Oligomerisation under SAR appears essential for NPR1 protein pool stability in both induced and uninduced conditions (Tada et al., 2008). Differential modulation of specific cysteine residues hints that specific redox-sensitive enzymatic players regulate different regions of the NPR1 protein. In support of this notion, Tada et al (2008) showed that one of eight cytoplasmic thioredoxins (TRXs), TRX-h8, is specifically upregulated during SAR and directly interacts with NPR1; loss of function trx-h8 mutants are defective in SA-induced monomerisation of NPR1, suggestive of a direct role for TRX-h8 in reduction of NPR1 oligomers. In addition, TGA interaction with NPR1 is also modulated by via redox-sensitive disulphide bridges amongst the TGA proteins themselves (Depres et al., 2003). 45  Like other bZIPs, TGAs preferentially dimerise when attaching to DNA (Vinson et al., 1989; Schutze et al., 2008). Interaction with other TGA proteins modifies TGA2 function such that it represses PR gene expression while interaction with NPR1 converts TGA2 into a transcriptional enhancer (Kersawani et al., 2007). Although NPR3 and NPR4 interact with a similar suite of TGAs as NPR1, they function to repress rather than activate PR gene expression as npr3 np4 double mutants show increased basal PR levels and enhanced disease resistance (Zhang et al., 2006). Zhang et al (2006) speculate that NPR3 and NPR4 may act in competition with NPR1 for TGA interaction and the increase in NPR1 availability during SAR leads to a displacement in favour of TGA transactivation. Taken together, these data suggest that competition among TGA-interactors may determine the repressor or activator TGA functions.  1.6.2 Conserved signalling mechanisms between BOP1, BOP2 and NPR1 Several aspects of the BOP1/2 function are similar to that of NPR1. Arabidopsis lines expressing 35S-BOP2-GFP showed localisation of GFP to cytoplasm and the nucleus in root cells (Hepworth et al., 2005). In yeast two-hybrid assays, BOP1 and BOP2 were found to interact with a TGA transcription factor from the same family as those shown to interact with NPR1, PERIANTHIA (PAN, TGA7; Hepworth et al., 2005). Moreover, genetic analyses suggest that BOP1, BOP2 and PAN act in the same genetic pathway to control perianth patterning in flowers (Hepworth et al., 2005). It is unknown whether BOP1/2 activity is controlled posttranscriptionally by redox-contol of nuclear localization and TGA interaction as is the case for NPR1.  46  2 ROLES OF BOP1 AND BOP2 IN ABSCISSION ZONE DEVELOPMENT 2.1 INTRODUCTION Arabidopsis petals, stamens and sepals develop an abscission zone four to six cell layers thick where the bases of the organs meet the receptacle (Bleecker and Patterson, 1997). Following fertilization, these floral organs senesce and abscise. Various genes are specifically expressed at floral organ AZs in Arabidopsis. For instance, the promoter from an Arabidopsis abscissionrelated PG gene was able to drive floral organ AZ-specific expression of β-glucuronidase (GUS) during abscission (Gonzalez-Carranza et al., 2002). Furthermore, GUS constructs driven by the soybean (Glycine max) chitinase (CHIT) promoter or the bean abscission cellulase (BAC) promoter are also upregulated in abscission zones during floral organ abscission (Thompson and Osbourne, 1994; Chen and Bleecker, 1995; Bleecker and Patterson, 1997; Patterson and Bleecker, 2004; Butenko et al., 2006). Floral organ abscission is absent in bop1 bop2 mutants. The first stage of my dissertation investigated the loss of abscission phenotype to determine how BOP1 and BOP2 contribute to the abscission program in Arabidopsis. I undertook a thorough microscopic examination of the abscission zones (AZs) of bop1 bop2 and wild-type plants, expression profiling of several other abscission-related genes in bop1 bop2 and genetic analyses between BOP1, BOP2 and other loci involved in abscission. My results indicate that BOP activity is necessary for the differentiation of the abscission zones in flowers as well as leaves and appears to be the earliest known regulator of abscission zone development characterised to date. This work has important implications with respect to understanding both the process of abscission and the function of the BOP proteins. Data presented in Figures 2.1, 2.4 – 2.6, 2.8-2.14 were published as a research article in Development (McKim et al., 2008) and granted a cover image. Figure 2.7 was 47  published in Hepworth et al., (2005). Data presented in Figures 2.10 and 2.11 was contributed by G.E. Stenvik, M.A. Butenko, W. Kristiansen and R.B. Aalen. Data used to construct graphs in Figures 2.1D and 2.9C were collected by S.K. Cho. Furthermore, the BOP1::GUS transgenic line was constructed by Shelly Hepworth while in working in the Haughn laboratory. All other data were solely my own contribution.  2.2 METHODS 2.2.1 Plant materials and growth conditions Seedlings were germinated onto Arabidopsis thaliana minimal medium agar plates (AT, Haughn and Somerville, 1986), transplanted to soil mix (Sunshine Mix 5; SunGro) and grown in continuous light (~125 microEinsteins) at 20ºC. Wild type was Columbia-0 (Col-0) ecotype, unless otherwise noted. Plants for 35S::IDA experiments were grown at 22°C for 8 hours dark /16 hours light and wild type was C24 ecotype. All alleles used in this thesis are listed in the Appendix. T-DNA mutants bop1-3 and bop2-1 alleles are in the Col-0 background and were obtained from the Arabidopsis Biological Resource Center (ABRC). Both alleles contain 5’UTR T-DNA insertions and were described in Hepworth et al. (2005). The ein3-1 seeds were provided by the Arabidopsis Biological Resource Center (ABRC, Columbus, Ohio). T-DNA ida mutant and IDA::GUS lines are in the C24 ecotype and were described in Butenko et al. (2003). bop1 bop2 ida (Col-0 x C24) triple mutants were generated by crossing and confirmed by genotyping. The 35S::IDA construct described in Stenvik et al. (2006) was transformed into bop1 bop2 plants by floral dip (Clough and Bent, 1998). Analysis was conducted on the T2 generation. BOP1::GUS plants (Col-0) were constructed by Shelley Hepworth and contain transcriptional fusions generated by fusing ~4 kb upstream of the putative BOP1 start codon to the β48  GLUCURONIDASE (GUS) reporter gene in the binary vector pBl101 (Jefferson et al., 1987). Transgenic plants harbouring the BOP1::GUS construct were selected on AT plates containing 50 µg/ ml [w/v] kanamycin. CHIT::GUS (Chen and Bleecker, 1995) and BAC::GUS (Koehler et al., 1996) marker lines have been introgressed into Col-0 and were kindly provided by Dr. Sara Patterson (University of Wisconsin). Primers used for cloning and genotyping are presented in the Appendix. Stages for abscission characterisation were assigned a position. The youngest flower showing visible white petals and was defined as position 1, roughly corresponding to just before anthesis (competency of self-fertilisation). Developmentally older flowers and siliques were numbered sequentially down the stem (basipetally).  2.2.2 BOP2 in situ hybridisation In situ probes were amplified by PCR with Pwo polymerase (Roche Diagnostics) from pCR2BOP2 using primers incorporating the T7 RNA polymerase binding site and spanning the entire BOP2 coding region (see the Appendix for primer sequences). Amplification conditions were 94°C for 2 min, 10 cycles of denaturation at 94°C for 15 s, annealing at 55°C for 30 s, and extension at 72°C for 90 s; 20 cycles of the same conditions but increasing the extension time by 20 s/cycle; followed by a final extension at 72°C for 7 min. Single-stranded RNA probes were transcribed in vitro from the PCR-generated DNA templates using T7 RNA polymerase and digoxygenin-11-UTP-labeled nucleotide mix (Roche Diagnostics) in separate sense (negative control) and antisense orientation with respect to the BOP2 coding region. The probes were then cleaved to approximately 150 bp in length by alkaline hydrolysis for 55 min at 60°C in 0.2 M sodium carbonate buffer (pH 10.4). An antisense CER6 probe was used as a positive control and was described previously (Hooker et al., 2002). 49  Inflorescences were vacuum-infiltrated with FAA (3.7% paraformaldehyde, 5% acetic acid, and 50% ethanol) plus 0.1% Triton X-100 fixative and incubated overnight at 4 °C. Fixative was removed with several washes of phosphate buffer and embedded in paraffin (Paraplast Plus; Sigma). Sections (8 µm) were prepared using a microtome and transferred to positively-charged slides (Fisher Biotech), heat-fixed at 42°C overnight and stored at 4 °C until probing. Paraffin was removed by immersing slides twice in 100% xylene, 50% xylene-50% ethanol, and 100% ethanol for 5 min each. Sections were hydrated by immersion in 95, 85, 70, 50, and 25% ethanol for 5 min each, treated with 2 X SSPE (300 mM NaCI, 20 mM NaH2PO4, 2 mM EDTA, pH 7) at 70°C for 20 min, and incubated for 20 min at 37°C with 1 µg/mL proteinase K in 100 mM Tris-HCI, pH 8, and 50 mM EDTA. Slides were then dehydrated in 25, 50, 75, 85, 95, and 100% ethanol for 5 min each and air dried at 52°C for 10 min. Hybridization was done overnight at 52°C with a digoxigenin-labeled RNA probe (Despres et al., 2000) in 100 µL of hybridization buffer [10 mM Tris-HCI, pH 7.5, 1 mM EDTA, 300 mM NaCI, 50% formamide, 7% dextran sulfate, 1 x Denhardt's solution (1 x Denhardt's solution is 0.02% Ficoll type 400, 0.02% polyvinylpyrrolidone, 0.02% BSA), 500 pg/mL tRNA, and 250 pg/mL poly(A) RNA]. Slides were washed in 2 X SSC (1 X SSC is 0.15 M NaCI, 0.015 M sodium citrate) for 5 min and twice in 0.2 x SSC at 52°C for 30 min. lmmunological detection of the hybridised probe was performed according to (Coen et al., 1990), with a few modifications. Slides were covered for 20 min with 1 mL of 1% blocking reagent (Roche) in 100 mM maleic acid, pH 7, and 150 mM NaCI. Slides were then covered for 30 min in 1 mL of buffer A (1 % BSA [Sigma], 0.3% Triton X-100 [Sigma], 100 mM Tris-HCI, pH 7.5, and 150 mM NaCI). The slides were incubated for 4 hr with 1 mL of dilute (1 :1200) antibody conjugate (Boehringer Mannheim) in buffer A, followed by two washes in buffer A (each for 20 min). For the colour reaction, slides were immersed twice for 5 min in substrate buffer (100 mM Tris-HCI, pH 9.5, 100 mM NaCI, 50  and 50 mM MgC12) and incubated overnight with 0.5 mL of 0.34 mg/mL nitro blue tetrazolium salt and 0.1 75 mg/mL 5-bromo-4-chloro-3-indolylphosphatep-toluidine salt in substrate buffer in the dark. The colour reaction was stopped with 10 mM Tris-HCI, pH 8, and 5 mM EDTA, and slides were viewed before (brown color) or after (blue color) ethanol dehydration, 100% xylene immersion, and coverslip mounting with Entellen (Merck). Sections were photographed using bright field optics.  2.2.3 Senescence and ethylene responses To estimate differences in senescence between floral organs of wild type and bop1 bop2, visual cues such as sepal yellowing, withering and drying of floral organs were assessed in 72 plants of each genotype grown in continuous light and 36 plants of each genotype for long day. Mature silique length was measured to determine if lack of abscission in bop1 bop2 affected silique elongation. The triple response assay used was from Vandenbussche et al (2003). Wild-type, bop1 bop2 and ein3-1 seeds were germinated on AT agar plates supplemented with 100 µM 1aminocyclopropane-1-carboxylic acid (ACC) in the dark at 20 ºC for seven days. Subsequently, hypocotyl length was measured and apical hook formation was scored.  2.2.4 Petal breakstrength Petal breakstrength is the force in gram equivalents required to removal a petal from the receptacle. Petal breakstrength was measured by the apparatus described in (Lease et al., 2006).  2.2.5 β-Glucuronidase activity staining Immediately following harvest, tissue was fixed in 90% acetone for three min. To aid substrate penetration, acetone was replaced with recycled heptane for 5 minutes. Heptane was removed 51  with two washes with β-glucuronidase (GUS) buffer ((100 mM NaH2PO4 pH 7.0, 10 mM EDTA, 0.1% triton-X 100, 0.5 mM potassium ferricyanide, 0.5 mM potassium ferrocyanide). Samples were then incubated with GUS buffer supplemented with 0.05% X-gluc (5-bromo-4-chloro-3indolyl β-D-glucuronide cyclohexylamine) salt (Rose Scientific, Alberta, Canada) in 37 ºC for 2 hours for IDA::GUS plants or overnight (~16 hours) for BOP1::GUS, CHIT::GUS, GLUC::GUS plants. GUS solution was removed by several washes in 70% ethanol. Tissues were cleared overnight in 70% ethanol and further cleared and mounted in 8 choral hydrate: 1 water: 2 glycerol. Photographs were taken using Spot 2 digital camera on a Wild Dissecting Microscope. BOP1::GUS and IDA::GUS samples were stained for GUS activity before embedding in standard Spurr’s resin (see below).  2.2.6 Microscopy Samples for scanning electron microscopy (SEM) were prepared as in Modrusan et al. (Modrusan et al., 1994). Sepals and petals were removed from flowers with forceps prior to fixation in 5% glutaraldehyde in 0.04 M phosphate buffer pH 7.0 overnight at 4 ºC. Fixative was washed out with phosphate buffer and then samples fixed in 0.5% osium tetraoxide for 2 hours. Samples were dehydrated in an ethanol series to absolute 100% ethanol and subsequently critically point dried using a Balzers CPD 020 Critical Point Dryer. The flowers were mounted onto steel stubs which were then coated with gold-palladium in an SEMPrep2 sputter coater (Nanotech, Manchester, UK) and observed using a Hitachi VP-4600 or S-2600 scanning electron microscope (Hitachi Inc., Japan) with an accelerating voltage of 5 kV or 10 kV. For histological AZ and nectary analysis, wild-type and bop1 bop2 flowers at position 2 (approximately stage 15) were fixed overnight at 4 ºC in 2.5% glutaraldehyde in 0.1M potassium buffer, pH 7.0, and then dehydrated in an ethanol series before embedding in standard Spurr’s 52  resin (recipe in Appendix). BOP1::GUS and IDA::GUS samples were first stained for GUS activity, cleared overnight in 70% ethanol and subsequently washed in acetone for 2 hours before embedding in standard Spurr’s resin. Spurr’s embedding followed a microwave infiltration protocol and overnight resin hardening at 65 ºC. For AZ and nectary analyses, sections were cut at 1 µm thickness with a Leica Utracut E Ultramicrotome (Leica Inc., Germany), heat-fixed onto positively-charged glass slides and stained in Toluidine blue. Sections for GUS analysis were cut to 8 µm thickness with a Reichert OM3 Ultramicrotome (Reichert, Austria) and heat-fixed onto slides. Sections were photographed under bright-field and/ or dark-field using a mounted camera (Qimaging, Burnaby, Canada) and images acquired using Openlab 4.0 (Improvision, PerkinElmer, Waltham, MA). Whole plant photographs were taken using a digital camera (Coolpix, Nikon, Japan).  2.3 RESULTS 2.3.1 bop1 bop2 mutants completely lack floral organ abscission Loss of floral organ abscission is a striking phenotype of bop1 bop2 plants (Fig. 2.1A). In wildtype Arabidopsis, sepals, petals and stamens normally abscise shortly after anthesis when pollen is released from stamen locules permitting fertilisation of the carpel roughly corresponding to flower unfurling under the growing conditions described here. The convention used to stage abscission is to label the youngest flower with visible white petals as position 1 (Bleecker and Patterson, 1997). Flowers further down the inflorescence are numbered basipetally down the stem. Anthesis generally occurs between position 2 and 3. Wild-type flowers shed sepals, petals and stamens at position 6.55 ± 1.09 (Fig. 2.1B, Table 2.1) while bop1 bop2 plants floral organs  53  Figure 2.1 Abscission phenotype and petal breakstrength of bop1 bop2 (A) Inflorescences of wild-type plants (Col-0, left) abscise floral organs while bop1 bop2 plants (right) retain their floral organs (arrow). (B) Abscission in wild-type flowers occurs between positions 5 and 7. (C) Floral organs of bop1 bop2 never abscise. (D) Petal breakstrength measurements of wild- type flowers (black bars) decrease with floral age while bop1 bop2 flowers (grey bars) show no change in force over time. Error bars represent standard deviation. n=30 per genotype. McKim et al. (2008)  dev.biologists.org  54  Table 2.1. Characterisation of floral organ abscission in wild-type and bop1bop2 mutants Col-0 n = 72; bop1bop2 n = 68; position = on the inflorescence. McKim et al. (2008) Floral Organs Dry (position)  Abscission (position)  Condition  Genotype  Sepal Yellowing (position)  Continuous Light  Col-0  2.56 ± 0.79  5.4 ± 0.50  6.55 ± 1.09  bop1bop2  2.59 ± 0.82  6.76 ± 0.83  na  Col-0  3.53 ± 0.51  4.25 ± 0.46  5.38 ±0.72  bop1bop2  5.09 ± 0.83  6.46 ± 0.97  na  Long Days dev.biologists.org  55  remain attached to the base of the elongating silique and following silique dehiscence (Fig. 2.1C).  2.3.2 Floral organ senescence in bop1 bop2 Senescence is a distinct developmental response which can occur without obligatory abscission (Taylor and Whitelaw, 2001): for example, in Arabidopsis, both floral organs and leaves senesce but only floral organs abscise. Furthermore, Stenvik et al (2006) described a transgenic line of Arabidopsis showing precocious abscission prior to senescence, demonstrating that abscission can occur in the absence of senescence. Nonetheless, some delayed abscission mutants, such as etr1-1, are also delayed in senescence (Bleecker and Patterson, 1997) suggesting that abscission may be delayed through retarded senescence. To address whether the abscission deficiency in bop1 bop2 results from an independent effect on abscission or is caused indirectly through some defect in the senescence program, I examined senescence kinetics in bop1 bop2 using visible cues such as organ wrinkling (due to organ drying) and sepal yellowing. Although Patterson and Bleecker (1997) reported that turgid organs are shed prior to abscission, under the growth conditions used here, floral organs began to senesce prior to organ shed. Table 2.1 shows that floral senescence is relatively unperturbed in bop1 bop2 apart from a slight delay in floral organ drying. Norberg et al (2005) showed that flowering and whole plant senescence was significantly delayed in the bop1 bop2 mutants grown under long-day conditions. Under our growing conditions of continuous light, bop1 bop2 plants showed only a slight delay in flowering time and plant senescence. bop1 bop2 plants were grown under long-day conditions to determine whether a significant delay in floral organ senescence is detectable under these conditions. A more pronounced retardation in sepal yellowing and floral organ drying was observed in bop1 bop2 compared with wild type under long day conditions (Table 2.1); however, 56  both wild-type and bop1 bop2 floral organs completely senesced by position 6-7 which corresponds to the position of abscission event. These results suggest that the abscission defect in bop1 bop2 is not associated with an inability to senesce.  2.3.3 Ethylene responses in bop1 bop2 Since ethylene perception is essential for proper timing of floral organ senescence and abscission, ethylene responses were characterised in bop1 bop2 using the classic triple response assay. In response to exposure to ethylene in the dark, germinating seedlings display three obvious morphological changes (hence the ‘triple’ response): shortened and thickened hypocotyls, exaggerated apical hook formation and inhibition of root elongation (Neljubov, 1901). Ethylene sensitivity mutants have been isolated using a screen for mutants altered in their triple response following exposure to ethylene (Guzman and Ecker, 1990) or exposure to the ethylene precursor, 1-aminocyclopropane-1-carboxylic acid (ACC). The ACC oxidase enzyme converts ACC to ethylene, carbon dioxide and cyanide. The reaction is not rate-limiting and exposure to ACC is considered equivalent to that of gaseous ethylene exposure (Smalle et al., 1997). Both wild-type and bop1 bop2 (Fig. 2.2A-C) seedlings showed a complete triple response with no significant difference in hypocotyl length or apical hook formation when germinated on ACC-containing plates suggesting that ethylene signalling is unaffected in bop1 bop2 seedlings and that any delay in senescence observed in bop1 bop2 plants is unlikely to be due to altered ethylene responses.  2.3.4 Characterisation of the bop1 bop2 abscission defect I speculated that absence of abscission in bop1 bop2 could result from a block of the final separation event, incomplete or uninitiated abscission zone (AZ) weakening or a lack of AZ development (Fig. 2.3). To distinguish amongst these possibilities, I conducted a series of 57  Figure 2.2 Triple response of wild-type and bop1 bop2 seedlings (A, B) Wild-type (Col-0) and bop1 bop2 seedlings germinated on 100 µM 1-aminocyclopropane1-carboxylic acid (ACC) in the dark showed the triple response consisting of enlarged, thickened hypocotyls, exaggerated apical hook formation and inhibition of lateral root initiation. (C) Histogram showing similar hypocotyl length of wild-type and bop1 bop2 seedlings germinated with and without 100 µM ACC. Scale bars, A,B 1mm. Error bars represent standard error.  58  intercellular adhesion strength measurements, scanning electron microscopy and histological analyses of the bop1 bop2 flowers. 2.3.4.1 Petal AZ breakstrength analysis To narrow down potential causes of the floral organ abscission defect in bop1 bop2, I evaluated the adhesion within bop1bop2 floral organ AZs using a petal removal technique. Due to the progressive degradation of the AZ middle lamellae, the amount of force required to remove a petal from the receptacle decreases as the organs approach abscission (Craker and Abeles, 1969). To assay to what extent, if at all, middle lamellar weakening occurs in bop1 bop2, petal breakstrength was determined in wild-type and bop1 bop2 petals using the apparatus described in Lease et al. (2006). As shown in Figure 2.1D, wild type (black bars) demonstrates the characteristic decline in petal breakstrength until abscission. It is important to emphasise here that weakening of the middle lamellae is progressive, taking several days to complete in wild type under our growing conditions. Dramatically, bop1 bop2 shows no decrease in breakstrength as flowers age (Fig. 2.1D, grey bars) suggesting that cell wall adhesion at the petal-receptacle boundary does not weaken in bop1 bop2. 2.3.4.2 Scanning electron microscopy of floral organ AZs To further examine the nature of AZ intercellular adhesion in bop1 bop2 versus wild-type floral organ AZs, I used scanning electron microscopy (SEM) to observe the floral organ-receptacle boundary of bop1 bop2 and wild-type flowers. Examination of the fracture surface resulting from organ removal indicates the progress of the abscission program (Bleecker and Patterson, 1997). Petals not yet abscised from wild-type and bop1 bop2 flowers at positions 2, 4, 6 and 12 were removed at the receptacle. Prior to declines in petal breakstrength, petal removal from position 2 wild-type and bop1 bop2 flowers reveals a surface of ruptured cells (Fig. 2.4A, B 59  Figure 2.3 Hypotheses for lack of floral organ abscission in bop1 bop2 Lack of abscission in bop1 bop2 could reflect an inability to promote the final cell separation event (1), inadequate or absent degradation of the middle lamellae between AZ cell files (2) or a lack of AZ development and/ or differentiation (3).  3  2  1  60  Figure 2.4 Morphology of floral abscission zones in wild type and bop1 bop2 (A,B) Scanning electron micrographs of petal AZs. The fracture plane on the receptacle (arrow) was observed following removal or natural abscission of petals in wild-type (A; Col-0) and bop1 bop2 (B) flowers at position 2, 4, 6 and 12. (A) Wild-type fracture planes demonstrate progression from broken cells (position 2) to rounded AZ cells (position 6) to protective surface cells (position 12). (B) Fracture planes of bop1 bop2 petals from all positions show broken cells. (C-J) Toluidine blue-stained sections from flowers at anthesis. (C-E) AZ cells at the stamen filament-receptacle (C), petal-receptacle (D) and sepal-receptacle (E) junctions in wild type (arrows). (G-I) Receptacle-organ boundaries of stamen filaments (G), petals (H) and sepals (I) of bop1 bop2 lack cells displaying AZ anatomy (arrows). (F) Longitudinal section along the adaxial and abaxial faces of wild-type flower. (J) Abaxial petalloid sepal of bop1 bop2 lacks an organreceptacle boundary. f, stamen filament; p, petal; s, sepal; ad, adaxial; ab, abaxial Scale bars, A,B 400 µm; C-E, G-I, 35 µm; F,J 70µm. McKim et al. (2008).  dev.biologists.org  61  arrows). However, petal removal from position 4 wild-type flowers does not cause cell rupture but reveals a smooth fracture surface due to the weakening of the middle lamellae typical of wild-type AZ cells (Fig. 2.4A). By position 6, petals from wild-type flowers readily abscise to reveal a mature AZ characterised by spherical, elongated cells (arrow). The exposed plant body then differentiated into a protective surface as seen in position 12 (Fig. 2.4A). In contrast, the fracture surface of bop1 bop2 shows no evidence of AZ middle lamellae degradation; cells consistently rupture following petal removal from flowers of all stages (Fig. 2.4B). Taken together, petal breakstrength and fracture surface SEM results show that weakening of the bop1 bop2 floral organ AZ middle lamellae does not occur at any stage examined, suggesting that the middle lamellae degrading enzymes are either unexpressed or unable to function in bop1 bop2 petal AZs. 2.3.4.3 Histological examination of floral organ AZs To determine more precisely the differences in cellular morphology between bop1 bop2 and wild type, toluidine-blue stained longitudinal sections of bop1 bop2 and wild-type flowers at approximately position 2 were examined. Cytoplasmically-dense AZ cells are visible in the stamen filament-receptacle, petal-receptacle and sepal-receptacle boundaries of wild type (Fig. 2.4C-E, arrows). Such a layer is never observed in bop1 bop2 (Fig. 2.4G-J, arrows). Interestingly, the abaxial petalloid sepals which develop in bop1 bop2 flowers show fusion with the flower receptacle and lack the characteristic pursing of the sepal-receptacle boundary (ab, Fig. 2.4J); however, the remaining sepals do not exhibit this fusion defect but also do not show AZ cell files. In summary, analyses via petal breakstrength (Section 2.3.4.1), SEM (Section 2.3.4.2) and light microscopy presented here strongly suggest that anatomically distinct and active floral 62  AZs do not form in bop1 bop2, indicating that BOP activity is necessary for the development of an AZ.  2.3.5 Vestigial AZ formation in bop1 bop2 Vestigial AZs develop in Arabidopsis at branching points and at the base of pedicels and cauline leaves (Stenvik et al., 2006). These AZs are not normally activated in Arabidopsis but show similar darkened cell-files as floral AZs (Stenvik et al., 2006). Given the absence of floral AZs in bop1 bop2, I examined whether bop1 bop2 also lacks vestigial AZs. 2.3.5.1 SEM of vestigial AZs Although vestigial AZs were postulated to exist at the base of wild-type cauline leaves (Stenvik et al., 2006), it was unknown whether these zones exhibit any developmental changes in intercellular adhesion similar to those observed for floral organs. To address this, I examined the attachment point of cauline leaves with the inflorescence stem by SEM following removal of the leaf, akin to that analysis for floral organ AZs presented in section 2.2.4.2. Cauline leaves were removed from the stem at three maturation phases based on leaf colour: green, yellowing (50%), and yellow. The exposed fracture plane on wild-type stems following removal of green cauline leaves shows breakage of cells along the surface (Fig. 2.5A). Removal of yellowing cauline leaves from wild-type stems reveals enlarged, rounded cells at the edges of the attachment point indicative of an AZ (Fig. 2.5B) and when fully yellow, reveals rounded cells although the vasculature cells remain broken (Fig. 2.5C). This analysis suggests that middle lamellae degradation does occur in cauline leaves of wild type and this can be tracked by SEM. Similar to wild-type, bop1 bop2 stems exhibit ruptured cells on the stem surface following removal of green cauline leaves (Fig. 2.5D). However, unlike wild type, stem surfaces observed after removal of yellowing or complete yellow cauline leaves consistently lacked any 63  Figure 2.5 Morphology of vestigial abscission zones in wild type and bop1 bop2 (A-F) Scanning electron micrographs of stem fracture planes stems following removal of cauline leaves. (A) Fracture plane in wild type (Col-0) shows ruptured cells following removal of green leaves. (B) Removal of 50% yellow leaves reveals a few rounded AZ cells. (C) Fracture planes from yellow leaves often show many rounded AZ cells. Stipules always flank the fracture plane (A, arrow). Corresponding fracture planes of bop1 bop2 from green (D), 50% yellow (E) and yellow (F) cauline leaves show no development of rounded AZ cells or evidence of flanking stipules. (G,H) A furrow of narrowed cells (arrows) flanked by stipules (H, arrowhead) is observed between the abaxial surface of the cauline leaf and the stem in wild type. (J,K) bop1 bop2 cauline leaves lack stipules and leaf-stem boundary furrow. (I) Toluidine blue-stained longitudinal sections show darkened files of cells that mark the vestigial AZ (arrow) at the adaxial boundary between the cauline leaf and stem in wild type and at the branching point with the primary inflorescence stem (arrowhead). The leaf-stem boundary furrow is also apparent (asterisk). (L) No vestigial AZs are visible in bop1 bop2 sections. as, axillary shoot; cl, cauline leaf; st, primary inflorescence stem. Scale bars, A-H,J,K 600 µm; I,L 35 µm. McKim et al. (2008)  dev.biologists.org 64  signs of smooth, enlarged, AZ cells (Fig. 2.5E, F) suggesting that corresponding middle lamellae weakening does not occur in the vestigial AZs of bop1 bop2 cauline leaves. The bases of attached, green cauline leaves were also examined. Abaxial surfaces of wild-type leaves at the stem show a furrow of narrowed cells (Fig. 2.5G, H, arrows) flanked by stipules (Fig. 2.5H, arrowhead). This boundary furrow corresponds to the lower cleavage site formed when wild-type leaves are removed. Abaxial surfaces of bop1 bop2 cauline leaves lack an obvious boundary furrow (Fig. 2.5J, K). Interestingly, stipules which flank the wild-type leaf fracture plane (Fig. 2.5A-C, arrow) and intact wild-type cauline leaves (Fig. 2.5H, arrowhead) were never observed flanking bop1 bop2 fracture planes (Fig. 2.5D-F) or intact leaves (Fig. 2.5K). 2.3.5.2 Histological examination of vestigial AZs To further examine cauline leaf-stem boundaries in bop1 bop2 versus wild type, sections through fully expanded, green cauline leaves were stained with toluidine blue and examined. Sections from wild-type reveal the narrowed cell layers of the leaf-stem boundary furrow (Fig. 2.5I, asterisk). Wild-type sections also show vestigial AZ cells at the adaxial leaf-stem boundary (Fig. 2.5I, arrow) and between the primary and axillary stem branching point (Fig. 2.5I, arrowhead). Wild-type vestigial AZs are readily apparent in serial sections through the sides of the leaf-stem boundary but are less obvious through the vasculature (Fig. 2.6A-C). However, neither cauline leaf nor branching point vestigial AZs are present in any position for expanded, green bop1 bop2 cauline leaves; cells at the junction remain large and vacuolated (Fig. 2.6D-F). In summary, these data indicate that bop1 bop2 show aberrant cauline leaf attachment morphology, lack the progressive middle lamellae weakening observed in wild type and show no obvious vestigial AZ anatomy, suggesting that in addition to floral organ AZs, BOP1 and BOP2 are essential for vestigial AZ formation and activation. 65  Figure 2.6 Serial sections through vestigial abscission zones of wild-type and bop1 bop2 Longitudinal series from a wild-type (Col-0) cauline leaf. (A,B) Vesitigial AZs are manifested as darkly stained cell files between the stem and the adaxial surface of the leaf (arrows). (C) Once in the vasculature the vestigial leaf AZ is not obvious. (D-F) Series through a bop1 bop2 cauline leaf does not show vestigial AZs. Scale bars, A-F, 35 µm. McKim et al. (2008).  dev.biologists.org  66  2.3.6 BOP1 and BOP2 expression analyses To determine whether the observed phenotypes correlate with expression of the BOP genes, the developmental expression pattern of BOP1 and BOP2 was assessed through a combination of in situ hybridisation and reporter-gene analyses. 2.3.6.1 BOP2 in situ hybridisation BOP2 mRNA localization was determined in reproductive tissues by in situ hybridisation. The BOP2 transcript is not expressed above background in the central zone of the inflorescence meristem (IM) but is expressed strongly in floral anlagen (P0) developing on the IM flanks (Fig. 2.7B). Expression persisted throughout stage 1 and stage 2 floral primordia with the strongest expression often seen near the base (Fig. 2.7, panels B-D, arrows). At stage 3 of flower development, expression of BOP2 subsided from the central part of the floral meristem but was detected throughout very young organ primordia in all four whorls as these structures developed (Fig. 2.7, panels E-F, and data not shown). As flowers matured, a strong band of BOP2 transcript was detected at the base of floral organs in the region corresponding to the AZ (e.g. as seen for mature flower shown in Fig. 2.7F). All features of the BOP2 expression pattern were distinct from that of ECERIFERUM 6 (Hooker et al., 2002), encoding a 3-ketoacyl-CoA synthase specific to epidermal cells (Hooker et al., 2002; Fig. 2.7A). These results were published in Hepworth et al. (2005) and are consistent with published reports from Norberg et al. (2005) for the BOP2 gene and with BOP1 in situ hybridisation published by Ha et al. (2004). 2.3.6.2 BOP1::GUS expression patterns I examined the expression pattern in more detail using a GUS reporter gene driven by the BOP1 promoter. (A caveat about GUS staining described here and in later sections is that the enzyme is relatively stable making it difficult to be conclusive with regards to when reporter transcription 67  Figure 2.7 In situ hybridisation of BOP2 mRNA (A) Central section of an inflorescence meristem (im) and young floral buds hybridised with an L1-specific CER6 control probe. (B-F) Longitudinal sections through inflorescence meristems and young flower buds hybridised with an antisense BOP2 probe. BOP2 mRNA is first detected in the floral anlagen (P0) on the flanks of the inflorescence meristem. Arrows in (B) and (D) indicate a strong band of expression at the base of stage 1 (P1) and stage 2 (P2) floral primordia. (E) Cross section of an inflorescence meristem and young floral buds hybridised with an antisense BOP2 probe. Arrows indicate expression in the developing sepal primordia of a stage 3 floral primordium. (F) Cross section of a mature flower showing strong bands of expression at the bases of anthers and petals (arrows) corresponding to developing abscission zones (AZs). The inset shows AZ detail at the base of the anther filament. Scale bars, A-E, 100 µm; F, 50 µm. Hepworth et al. (2008).  www.plantcell.org. Copyright American Society of Plant Biologists.  68  actually ceases). BOP1::GUS is expressed at the base of developing floral organs in stage 9/10 (Fig. 2.8A), stage 12 (Fig. 2.8B) and stage 14 (Fig. 2.8C) flowers, agreeing with previous results (Norberg et al., 2005). In addition, bases of recently shed organs have GUS activity (Fig. 2.8D). BOP1::GUS receptacle expression is detected for a prolonged period following abscission (Fig. 2.8E). The adaxial regions of young pedicel-stem junctions show GUS activity (Fig. 2.8F) that spreads around the entire boundary in older pedicels (Fig. 2.8G). As described above, BOP expression becomes restricted to the proximal segment of organs during development. Since the BOP homologue, NPR1, is activated by external signals it is possible that the BOP proximal expression pattern could be generated by BOP autoregulation following activation by a tissue specific signal. For example, the restriction of BOP expression to the base of developing organs could result from positive feedback mediated by activated BOP proteins. To test this hypothesis, expression of BOP1::GUS in a bop1 bop2 mutant background was examined. BOP1::GUS expression exhibits similar patterns in bop1 bop2 as in wild type (data not shown), showing that BOP1 expression does not depend on functional BOP proteins. Interestingly, BOP1 is ectopically expressed in the bop1 bop2 pedicel where the ectopic bract develops (Fig. 2.8H, compare to 2.8G). In addition, expression of BOP1::GUS is detected at the bases of cauline leaves (Fig. 2.8I) as previously reported (Norberg et al., 2005). Given ethylene’s role in timing of abscission, BOP expression in the mature AZ may be driven in an ethylene responsive fashion. To evaluate this possibility, BOP1::GUS expression was examined in the ethylene response 1 (etr1) mutant background which displays an ethylene insensitive phenotype. BOP1::GUS was expressed in the mature AZ at anthesis in etr1-1 similar to wild type (data not shown) suggesting that BOP1 expression is not regulated by pathways mediated through ETR1 although the possibility remains that BOP1 may be regulated by the activity of other ethylene receptors. 69  Figure 2.8 Expression of BOP1::GUS Whole mounts of BOP1::GUS (Col-0) transgenic plants stained for GUS activity. (A) Expression is restricted to the base of developing organs in stage 9 and 10 flowers. (B) Expression at the base of floral organs and in lateral nectaries of a stage 12 flower (arrow). (C) GUS activity at the base of floral organs, sepal vasculature, style and pollen of stage 14 flowers. (D) Newly-abscised petal and stamen show expression in their AZs and filaments. (E) Position 9 siliques show AZ expression in the receptacle. (F) Young pedicel-stem junctions have an abaxial GUS signal which spreads around the attachment point in older pedicels (G). (H) BOP1::GUS bop1bop2 plant with GUS activity at the base of the ectopic bract (arrow). (I) Cauline leaves show GUS activity at their base. (J-N) Sections of stained BOP1::GUS flowers. (J) Stage 5/6 flower showing GUS activity in stamen primordia (asterisk) and sepals. (K) Staining of a stage 7/8 flower is restricted to the base of stalked stamens (asterisk) and outgrowing sepals. (L) Section through the same stage 7/8 flower showing staining throughout the petal primordia (asterisk). Mature flowers with GUS activity through the AZ (M) and nectaries (N). Scale bars, A, B, D-F 250 µm; C 0.5 mm; G-I 1mm; J-N 35 µm. McKim et al. (2008).  dev.biologists.org 70  To further characterise BOP1 expression, BOP1::GUS stained tissue was sectioned. Corresponding to in situ hybridisation results of BOP2 (Hepworth et al., 2005), BOP1::GUS is expressed in early floral organ primordia (Fig. 2.8J). As the stamens become stalked, GUS activity is basally restricted (Fig. 2.8K, L) while the petal primordia, which develop later, display diffuse staining (Fig. 2.8L). The petal expression also becomes restricted to the petal base as petals grow out (data not shown). Mature flowers show strong staining through the proximal organ and AZ (Fig. 2.8M) in agreement with patterns observed in BOP2 in situ hybridisation (Hepworth et al., 2005). In summary, patterns of BOP1::GUS expression correlate with a putative role in abscission zone development.  2.3.7 Genetic analysis of BOP loci with abscission-related genes Although research into the genetic regulation of abscission in Arabidopsis is relatively recent compared to that of flower and leaf morphology, several loci involved in the regulation of abscission in Arabidopsis have been described. I characterised the genetic interaction between BOP1 and BOP2 and known abscission-related genes, including putative signalling components and enzymatic mediators of abscission, to determine the relative contribution of BOP1 and BOP2 to the abscission program. 2.3.7.1 Suppression of ida and 35S::IDA phenotypes by bop1 bop2 INFLORESCENCE DEFICIENT IN ABSCISSION (IDA) encodes a small protein presumed to act as a signal peptide to promote the final cell separation event in abscission (Butenko et al., 2003; Stenvik et al., 2006). Loss of function ida mutants, as in bop1 bop2, show a complete lack of abscission; however, unlike bop1 bop2, ida flowers differentiate floral organ AZs and show reduced petal breakstrength (Butenko et al., 2003). BOP1 and BOP2 are essential for formation of proper anatomy within the AZ suggesting that these roles are fulfilled prior to IDA function. 71  To test this hypothesis, triple mutants were constructed and examined. The triple mutant bop1 bop2 ida, does not differentiate AZs or show decreases in petal breakstrength (Fig. 2.9) demonstrating that bop1 bop2 is epistatic to ida. My results suggest that absence of abscission in bop1 bop2 occurs due to a lack of proper AZ anatomy. I tested whether overexpression of IDA by the Cauliflower Mosaic Virus (CaMV) 35S promoter in bop1 bop2 plants can circumvent this requirement. The CaMV 35S promoter leads to enhanced expression in most tissue types in Arabidopsis. Compared to wild type (C24) (Fig. 2.10A), precocious floral organ abscission is a striking phenotype of 35S::IDA plants (Fig. 2.10B) as previously described (Stenvik et al., 2006). These plants also exhibit activation of vestigial cauline AZs (Fig. 2.10F) and ectopic abscission at pedicels and branching points (Stenvik et al., 2006). Dramatically, premature floral organ abscission is completely masked in 35S::IDA bop1 bop2 plants which show no floral organ abscission (Fig. 2.10D) and resemble bop1 bop2 (Fig. 2.10C), indicating that bop1 bop2 is completely epistatic to 35S::IDA. Furthermore, ectopic abscission at vestigial AZs in pedicels, branching points (data not shown) and cauline leaves (Fig. 2.10F) is also lost in 35S::IDA bop1 bop2 plants which resemble bop1 bop2 (Fig. 2.10E). Both C24 and Col-0 backgrounds demonstrate these phenotypes when transformed with 35S::IDA, suggesting that the lack of 35S::IDA phenotype in 35S::IDA bop1 bop2 is not due to the Col-0 ecotype. In addition, the 35S::IDA construct is not transcriptionally silenced in bop1 bop2 (Fig. 2.10H). Thus, earlier and ectopic expression of IDA is not sufficient to promote abscission in bop1 bop2 due to an absence of proper AZ architecture, suggesting that, in order to properly function, IDA must be expressed in an AZ with correct anatomy  72  Figure 2.9 Genetic interaction between bop1 bop2 and ida (A, B) Floral organ abscission zones (AZs) in ida (Col-0 x C24) and bop1 bop2 ida (Col-0 x C24) documented by SEM of the petal AZ fracture plane. The fracture plane on the receptacle was observed following removal or natural abscission of petals in ida (A) and bop1 bop2 ida (B) from position 2, 4, 6 and 12. (A) Petal AZs in ida show broken cells at position 2 and rounded AZ cells by positions 4 and 6. However, petal fracture planes revert to show broken cells by position 12. (B) Petal AZs from bop1 bop2 ida always show broken cells. (C) Petal breakstrength of bop1 bop2 ida is similar to bop1 bop2. Error bars represent standard deviation. Scale bars, A,B 400 µm. McKim et al. (2008).  dev.biologists.org  .  73  Figure 2.10 Suppression of premature and ectopic abscission from 35S::IDA in bop1 bop2 (A-D) Flowers from positions 3, 7, and 10. (A) Wild type (C24) shows floral organ abscission by position 6. (B) 35S::IDA (C24) with premature abscission of floral organs by position 3. (C) bop1 bop2 lacks floral organ abscission. (D) 35S::IDA bop1 bop2 (Col-0) showing no floral organ abscission. (E) Cauline leaves do not abscise in bop1 bop2. (F) Cauline leaves of 35S::IDA show ectopic abscission. (G) 35S::IDA bop1 bop2 cauline leaves resemble bop1 bop2. (H) RTPCR from cDNA derived from rossette leaves, buds and flowers from bop1 bop2 and 35S::IDA bop1 bop2. Upper panel shows RT-PCR with IDA primers. IDA is expressed in bop1 bop2 flowers only. 35S::IDA bop1 bop2 plants show expression in rosette leaves, buds and flowers. lower panel, RT-PCR of ACTIN2-7 (ACT) as a positive control. Lane 1 is a genomic PCR control (g). l, leaves; b, buds; f, flowers. McKim et al. (2008).  dev.biologists.org  74  Figure 2.11 Real-time quantitative RT-PCR of IDA and HAESA in wild-type (Col-0) and bop1 bop2 abscission zones. Transcript levels of IDA and HAESA were determined using two biological replicates and were normalised using ACTIN2. Amplification efficiency was assumed to be 2. IDA was expressed at 0.94 and HAESA at 0.95 in bop1 bop2 relative to wild type which was set at 1.00. Error bars indicate range. McKim et al. (2008).  dev.biologists.org  75  2.3.7.2 Abscission-related gene expression in bop1 bop2: IDA::GUS, CHIT::GUS and BAC::GUS reporter gene analyses I was interested in determining whether BOP1 and BOP2 regulate expression of genes encoding putative signalling components, such as IDA, and/or enzymes normally transcribed in AZs. RTPCR results indicated that IDA, HAESA (HAE) and HAWAIIAN SKIRT (HWS) have the same expression levels in bop1 bop2 and wild-type flowers (results not shown). Whereas mutation in HWS leads to delayed abscission and fusion of sepal margins that precedes shedding (GonzalezCarranza et al., 2007), down-regulation of IDA or HAE results in abscission defects only (Jinn et al., 2000). The RT-PCR results were confirmed by quantitative RT-PCR for IDA and HAE; their expression levels in wild-type (Col-0) and bop1 bop2 floral AZs do not differ (the relative expression was very close to 1: 0.94 for IDA and 0.95 for HAE, Fig. 2.11). To further examine IDA regulation in bop1 bop2, IDA::GUS plants (C24) were crossed into bop1 bop2. Wild-type IDA::GUS expression was examined in Col-0 x C24 background to compare with bop1 bop2 IDA::GUS plants generating by crossing. IDA::GUS is expressed shortly following anthesis in bop1 bop2 flowers and persists until after the stage when abscission would normally occur (Fig. 2.12B), a temporal profile akin to that observed in wild type (Fig. 2.12A). Spatially, IDA::GUS expression is present at the base of the floral organs in bop1 bop2, similar to wild type (Fig. 2.12 C, D). Two reporter constructs driven by promoters from genes encoding enzymes which are specifically upregulated in AZs, BAC::GUS and CHIT::GUS, were also examined. As observed previously in wild type (Butenko et al., 2006), BAC::GUS expression appears early (before anthesis) and throughout the AZ by position 1 (Fig. 2.12E). BAC::GUS is expressed also in the proximal petal and stamen filaments. While this temporal pattern is retained in bop1 bop2 (Fig.  76  Figure 2.12 Expression of IDA::GUS, BAC::GUS and CHIT::GUS in wild type and bop1 bop2 genetic backgrounds Floral positions are indicated in the upper right of panels. (A) Wild-type (C24 x Col-0) flowers at position 3 show very weak IDA::GUS activity in the base of floral organs that intensifies throughout position 4 and 5. (B) IDA::GUS expression in bop1 bop2 (Col-0 x C24) is detected at position 4 and more strongly in position 5. (C,D) Darkfield microscopy of stained IDA::GUS flowers in wild type (C) and bop1 bop2 show staining throughout the proximal sepal. (E,F) BAC::GUS in wild type (Col-0) or bop1 bop2 at positions 1, 3-6 stained for GUS activity. (E) BAC::GUS plants show strong staining at the base of all floral organs especially concentrated in the vasculature of position 1 and 3 flowers. Activity persists in position 4 and 5 flowers during abscission and is visible in single AZ cells (arrow). (F) BAC::GUS bop1 bop2 flowers at position 1 show weak staining at the bases of petals and stamens which strengthens and enlarges by position 3. Very slight expression is detected on the adaxial side of sepals in position 4. (G,H) CHIT::GUS expression from flowers in wild type (Col-0) or bop1 bop2 at positions 3-7. (G) CHIT::GUS position 3 and 4 flowers demonstrate weak GUS staining while position 5 and 6 exhibit very strong AZ staining. (H) CHIT::GUS bop1 bop2 position 3 and 4 floral organs show weak GUS activity at their base. GUS activity is stronger in position 5. Staining is maintained in position 6 flowers extending further into the base of these floral organs. Arrowheads indicate expression. f, stamen filament; p, petal; s, sepal. Scale bars, A,B,E-H, 0.5 mm; C,D 35 µm. McKim et al. (2008).  77  dev.biologists.org  78  2.12F), BAC::GUS expression is reduced in intensity, restricted to the bases of the petals and stamens and absent from single sepal AZ cells as seen in the wild type (Fig. 2.12E, arrow). Consistent with published results (Patterson and Bleecker, 2004; Butenko et al., 2006), CHIT::GUS is expressed in the floral organ AZ cells starting at position 4 and increases to maximal intensity coincident with abscission at position 6 after which expression weakens (Fig. 2.12G). Despite the lack of anatomical AZ structure in bop1 bop2, temporal expression of CHIT::GUS is maintained, albeit with lower intensity (Fig. 2.12H). Often CHIT::GUS activity was detected earlier in bop1 bop2 but this was not consistent. CHIT::GUS expression also occurs in the proximal floral organs in the bop1 bop2 background (Fig. 2.12H). These data suggest that activation signals for BAC::GUS and CHIT::GUS are operating in bop1 bop2 regardless of a lack of AZ morphology. Interestingly, ida also shows normal temporal and spatial expression of both these reporters in AZ cells (Butenko et al., 2006) suggesting that signals activating their expression function independently from IDA as well.  2.4 DISCUSSION 2.4.1 BOP1 and BOP2 act early to specify AZ anatomy Abscission occurs in the AZ positioned at the junction between a lateral organ and the main plant body. The AZ is characterized by a unique anatomy and although thought to be essential for abscission, this has not been explicitly demonstrated in Arabidopsis since previously described mutants retain the development of an AZ. I show that two redundant NPR1-like homologues, BOP1 and BOP2 are expressed in the AZ and that lack of BOP1/2 activity results in a complete absence of abscission. Moreover, the absolute loss of floral organ abscission in bop1 bop2 is uniquely correlated with an absence of cellular anatomy typical of the AZ, suggesting that the 79  AZ anatomy is necessary for abscission and that BOP1 and BOP2 initiate differentiation of the AZ. However, I cannot rule out the possibility that BOP1 and BOP2 control other downstream events also necessary for abscission. Several lines of evidence implicate BOP1 and BOP2 in the specification of abscission zone cells as the earliest known step necessary for abscission. First, BOP1 and BOP2 genes are transcribed in early floral organ primordia and resolve to a region corresponding to the future AZ, prior to other known abscission-related genes. Second, appearance of AZ anatomy, absent in bop1 bop2, is the earliest identified event associated with AZ development. Third, the bop1 bop2 phenotype is epistatic to that of both ida and 35S::IDA suggesting that BOP1 and BOP2 act upstream of the only other gene known to be absolutely required for abscission. BOP1 and BOP2 are expressed earlier than IDA, raising the possibility that BOP1 and BOP2 are positive regulators of IDA expression. However, we have shown that IDA is expressed similarly to wild type in bop1 bop2. It is possible that BOP proteins regulate IDA activity posttranscriptionally but given the requirement of BOP1 and BOP2 for AZ-specific anatomy, we favour a model where BOP1/2 function early to specify AZ cell morphology and IDA acts relatively independently to finalize the cell separation process. Significantly, as is the case for IDA (Butenko et al., 2006), BOP1 and BOP2 are not required for the correct temporal transcription of known abscission-related genes tested here, including one encoding a cell wall hydrolytic enzyme. A model of the known essential players in abscission is presented in Figure 2.13. I propose that BOP1 and BOP2 act early to specify AZ-unique anatomy. The characteristics of this anatomy which make it crucial for abscission remain to be identified but could include cell shape and/or cell wall structure amenable to middle lamellae digestion. Upstream factors that regulate initiation of abscission act through both ethylene-dependent and ethylene-independent pathways 80  Figure 2.13 Model of abscission BOP1 and BOP2 act early to promote the development of AZ specific anatomy of small, cytoplasmically dense cell files. Later in flower development, a suite of enzymes involved in middle lamella degradation are expressed specifically in the AZ although with differing temporal patterns. Transcription of these enzymes is independent of BOP-driven formation of AZ anatomy. Abscission occurs following the expression of IDA in the AZ which promotes cell separation. Expression of IDA is also driven independently from BOP1/2 activity.  81  which converge to activate the expression of abscission-related genes including middle lamellae- degrading enzymes. These enzymes are expressed in the AZ, presumably with specific spatial and temporal profiles, to progressively weaken the middle lamellae. This expression is driven independently from BOP1/2-mediated differentiation of the AZ. As yet, none of these enzymes have individually been shown to be essential for abscission. Finally, IDA is necessary for abscission and is expressed in the AZ just prior to abscission, in response to an unknown signal. Given that partial enzymatic dissolution of the middle lamellae occurs in ida mutants, IDA must act downstream from the initiation of abscission. Expression of IDA alone is insufficient for abscission since premature abscission of 35S::IDA plants only occurs at flower positions after anthesis where organs have presumably differentiated AZ that are competent to respond to IDA (Stenvik et al., 2006). New data indicates that IDA function is dependent on the expression of HAE and HSL2 (Cho et al., 2008; Stenvik et al., 2008) which occurs following anthesis (Jinn et al., 2000; Cho et al., 2008). Given the putative role of IDA as a ligand for receptor-like kinases encoded by HAE and HAESA-LIKE2 (HSL2), HAE and HSL2 signalling pathways may also be dependent on BOP1 and BOP2-mediated differentiation of the AZ. The targets of IDA, HAE and HSL signalling are unknown; however, a recent report indicates suggests that the enrichment and activity of the trans-golgi network in AZ cells prior to abscission is essential for cell separation and thus may be a target for IDA/HAE signalling (Liljegren, 2009).  2.4.2 BOP1 and BOP2 also specify vestigial AZ anatomy Although Arabidopsis leaves do not abscise, the observation that ectopic abscission can be induced by 35S::IDA ectopic expression has suggested that vestigial AZs develop at the bases of cauline leaves, branching points, and at the base of pedicels (Stenvik et al., 2006). Several lines 82  of evidence presented here strongly support this hypothesis. First, cauline leaf AZs have characteristic AZ anatomy, a boundary furrow and show progressive degradation of middle lamellae with age similar to floral AZs. Second, the characteristic anatomy, boundary furrow and 35S::IDA-induced abscission at these putative vestigial AZs are dependent on BOP activity. Finally, vestigial AZs develop on the adaxial side of the leaf base corresponding well to BOP1 and BOP2 expression as shown here and in other studies (Ha et al., 2004; Norberg et al., 2005).Taken together, these data suggest that vestigial AZs do develop and that their AZ anatomy is regulated by BOP1 and BOP2.  2.4.3 The role of BOP1 and BOP2 in proximal lateral organ development Previous work on BOP1 and BOP2 has suggested a role for these genes in defining the identity of the proximal regions of leaves (Ha et al., 2003; Hepworth et al., 2005; Norberg et al., 2005). The receptacle may be thought of as a proximal feature of a flower, just as the petiole is a proximal area of a leaf. Therefore, absence of AZs in bop1 bop2 may suggest that BOP1 and BOP2 regulate this aspect of proximal flower and floral organ differentiation. BOP1 and BOP2 promote specific aspects of boundary morphology, such as stipules and AZs, but likely do not contribute to boundary identity per se. Although AZs can be interpreted as defining the organ-plant body junction, bop1 bop2 floral organs do not display gross fusions with the receptacle or themselves as seen in the cuc mutants (Aida et al., 1997; Ishida et al., 2000; Hibara et al., 2006) or in other abscission-defective mutants such as hws (GonzalezCarranza et al., 2007). Furthermore, cauline leaves in bop1 bop2 do not display gross malformations where they meet the stem but rather lack particular features such as the boundary furrow and stipules. Moreover, normal patterns of expression of floral organ AZ markers, such as IDA and HAESA, persist in bop1 bop2 plants suggesting that the positional information is 83  intact. In addition, the boundary-specific expression of a reporter for the LATERAL ORGAN BOUNDARIES gene (Ha et al., 2007) was also detected, albeit at reduced levels, in bop1 bop2 (Ha et al., 2007) which further substantiates the idea that the boundary specific positioning cues are operational. Thus, bop1 bop2 plants lack some but not all of the features of the organ-plant body junction, suggesting that other factors are responsible for overall boundary patterning, such as CUP-SHAPED COTYLEDONS1-3 (Aida and Tasaka, 2006). The BOP1 and BOP2 genes would then act as later effectors of specific aspects of the organ-plant body interface development. How BOP1 and BOP2 promote the formation of AZ anatomy rather than the normal highly vacuolated tissue in the floral organ AZs is unknown. Previous work suggests that BOP1 and BOP2 are important for downregulation of certain KNOX genes and distal identity factors in the proximal regions of leafy organs. BOP1 and BOP2 repress the expression of the class I KNOX, BREVIPEDICELLUS (Colombo et al., 1995), KNOTTED-LIKE2 (KNAT2) and KNOTTED-LIKE6 (KNAT6) genes in leaf petioles (Ha et al., 2003; Ha et al., 2007) and JAG in petioles and bracts (Norberg et al., 2005). Class I KNOX genes are important for maintenance and establishment of shoot-meristem identity, and are normally downregulated in incipient lateral organ primordia (Scofield and Murray, 2006). BP, in particular, has been implicated in boundary and receptacle development (Venglat et al., 2002; Douglas et al., 2002; Douglas and Riggs, 2005; Wang et al., 2006). Reduced BP function leads to failure to develop the characteristic cup-shape bulge of the receptacle, particularly on the abaxial side (Douglas and Riggs, 2005) and to an overproliferation of AZ cells, reminiscent of 35S::IDA plants (Douglas et al., 2002; Wang et al., 2006). This suggests on the one hand that BP promotes receptacle tissue proliferation and on the other that BP represses AZ cell formation. However, ectopic BP expression led to enhanced receptacle bulging accompanied by early abscission of petals (Chuck 84  et al., 1996), suggesting a more complex role for BP in abscission. Excess abaxial growth shown in bop1 bop2 receptacles may result from temporal and/ or spatial misregulation of BP which may then contribute to defects in AZ differentiation. In addition to BP, KNAT2 and KNAT6, other genes involved in promotion of distal organ identity, such as JAGGED (JAG) and NUBBIN, are also misexpressed in bop1 bop2 plants (Norberg et al., 2005). JAG is normally expressed in the distal regions of lateral organs but is misexpressed in bop1 bop2 petioles and floral bracts suggesting that BOP1 and BOP2 activity in proximal regions is important for repression of expression of distal factors (Norberg et al., 2005). JAG, along with AS1/AS2, downregulates CUC and PTL boundary genes during flower development (Xu et al., 2008). Thus, misexpression of JAG in bop1 bop2 flowers could lead to downregulation of boundary-specific genes during AZ development. However, bop1 bop2 jag flowers do not show rescue of floral patterning defects or abscission suggesting that misexpression of JAG does not contribute to these phenotypes or that redundancy with NUB obscures these roles (Norberg et al., 2005).  2.4.4 New candidates for shared function with BOP1 and BOP2 Several new genes involved in abscission zone patterning were recently described including ARABIDOPSIS HOMEOBOX PROTEIN1 (ATH1), which suppresses cellular proliferation in boundaries (Gomez-Mena and Sablowski, 2008). Similar to bop1 bop2, ath1 mutants display loss of certain boundary features in both leaves and flowers, including a loss of boundary furrow and stipules at the base of cauline leaves, and a loss of stamen AZ differentiation (Gomez-Mena and Sablowski, 2008). ATH1 encodes a member of the BELL-LIKE HOMEODOMAIN (Cole et al., 2006) protein family which are known to physically interact with STM and BP (Cole et al., 2006) suggesting a link between regulation of KNOX activity and AZ development. 85  Interestingly, ath1 plants showed a decrease in BP::GUS activity specifically at the organ boundaries in young flowers, indicating that regulation of BP expression may be important for boundary differentiation. Another recent addition to the suite of abscission-related genes is ZINC FINGER PROTEIN2 (AtZFP2) which encodes a bZIP protein in the same family as JAG (Cai and Lashbrook, 2008). AtZFP2 is highly upregulated in stamen AZs yet overexpression leads to extremely delayed abscission and in the case of sepals, complete loss of abscission. It is unknown whether this phenotype reflects a loss of AZ differentiation; however, SEM fracture place analyses suggest that middle lamellae degradation does not occur. Similar to other abscission mutants, overexpressing AtZFP2 lines showed extreme delays in senescence (Cai and Lashbrook, 2008), highlighting a potential regulatory association between flower maturation and AZ development. KLUH (KLU), a cytochrome P450 protein, is not newly described, but its role in abscission has been previously overlooked. Zondlo and Irish (1999) showed that KLU expression was adaxially localised to the base of floral organ boundaries and that ectopic overexpression of KLU leads to a loss of floral organ abscission. KLU has been proposed to mediate a mobile signal distinct from known phytohormones to regulate organ proliferation in later stages of organ growth (Anastasiou et al., 2007). These results offer a tantalising possibility that KLU-induced signalling coordinates the formation of AZs with the growth arrest of the lateral organ and suggest possible upstream regulators of BOP function.  86  3 ROLES OF BOP1 AND BOP2 IN NECTARY DEVELOPMENT 3.1 INTRODUCTION In addition to other floral patterning phenotypes described for bop1 bop2, I discovered that bop1 bop2 flowers are defective in certain aspects of nectary gland formation. The Arabidopsis nectary develops as a continuous ring of tissue along the receptacle between the perianth and the third whorl stamens. Nectary tissue interconnects a series of secretory glands that abaxially subtend the base of stamen filaments and secrete nectar. Lateral nectary glands form a lobed pair which encircles the base of lateral stamens while a single, smaller nectary gland forms at the base of each medial stamen (Davis et al., 1994). Although clearly associated with stamens, the nectary gland is closely appressed by laterally bordering petal claws and basally cupped by sepals. Nectary development is first detected around floral stage 9 as small protrusions at the base of lateral stamens driven by periclinal divisions in the L2 layer of the receptacle (Symth et al., 1990; Bowman and Symth, 1999; Baum et al., 2001). Sustained periods of cellular proliferation between stages 10 through 13 give rise to the nectary glands and interconnecting nectary ridges. During morphogenesis, nectary glands develop clear histological differentiation between an epidermal layer and a cluster of parenchymal layers specialised for nectar secretion, collectively referred to as nectariferous tissue (Fahn, 1988). Acquisition of wavy cuticular ridges and abaxial secretory stomata (modified to be permanently open) are characteristics of the nectary epidermis while swelling of the gland and vascularlisation by phloem sieve tube members reflect the nectar pooling role of the nectary parenchyma (Davis et al., 1994). Secretory stomata are visible as early as stage 10, accompanied by gland outgrowth due to localised cell divisions (Baum et al., 2001). Parenchymal tissue continues to proliferate, producing the swollen  87  protrubences of mature nectary glands filled with nectar which secrete at around stage 14 (Smyth et al., 1990; Bowman and Smyth, 1999; Baum et al., 2001). Unlike other lateral organs which grow outward via a combination of cell division and cell expansion, nectary outgrowth relies primarily on cell division although the secretion phase may involve an increase in vacuole size (Nepi et al., 2007). Nectary tissue proliferation may require reactivation of parenchymal meristematic character in receptacle cells. Nectary tissue has often been likened to meristematic tissue (Gaffal et al., 1998) and is characterised by small cytoplasmically dense cells with high ploidy (Nicolson et al., 2007). KNAT2 and BP, meristem associated KNOX proteins, are expressed in proliferating nectaries, in addition to the receptacle (Baum et al., 2001) also suggesting that nectary proliferation involves meristematic reactivation. However, given the determinate nature of nectary gland development, nectiferous tissue is more comparable with the highly mitotic PZ lateral organ primordia than a true meristem. Nectaries resemble other lateral organs in having two distinct phases: early primodium development followed by morphogenesis. To date, a single gene has been identified as essential for nectary initiation: CRABS CLAW (CRC), which is a member of the YABBY transcription factor family involved in carpel identity and polarity (Bowman and Smyth, 1999; Eshed et al., 1999; Seigfried et al., 1999). The role of CRC in specification of nectaries appears conserved within examined eudicots (Lee et al., 2005a). CRC is expressed in a ring in the third whorl between developing stamens and sepals around stage 6, before any sign of nectary gland anlagens, and is strongly maintained in the nectary tissue throughout past stage 17 (Bowman and Smyth, 1999), hinting that CRC may have roles in the maturation of nectaries following their initiation. However, all crc mutants lack any trace of nectary development, thereby obscuring putative later functions. Nonetheless, there is evidence suggesting that CRC may influence later nectary elaboration. First, nectary-like growths on pedicels formed due to CRC misexpression in 88  the ap1 mutants, developed more nectary-like characteristics when CRC expression was augmented with the 35S::CRC transgene (Baum et al., 2001). Second, Lee et al (2005b), using dissected promoter elements driving genomic CRC, showed correct spatial and temporal CRC expression that rescued lateral nectary development in the crc background; however, medial nectaries were commonly absent and lateral nectaries often lacked stomata. Expression from the promoter elements was assayed by monitoring CRC::GUS activity and thus it is unclear whether the intensity of CRC expression was recovered to wild-type levels. These data imply that CRC may be limiting for nectary differentiation in addition to initiation. A thorough analysis of multiple floral organ mutants showed that nectaries developed in the third whorl regardless of stamen presence, suggesting a dependence of nectary development on third whorl formation rather than stamen identity (Baum et al., 2001). This hypothesis is also supported by the fact that ectopic stamens forming outside the third whorl did not develop nectaries (Baum et al., 2001). Furthermore, superman-1 (sup-1) and 35S::UFO develop multiple reiterations of the third whorl inside the centre of the flower and in this case, ectopic stamens are all associated with nectary tissue. Nectary development is abolished in BC double mutants, implicating B and C genes in nectary initiation and this is interpreted as a loss of third whorl development (Baum et al., 2001). Interestingly, nectaries are also lost in the sep1 sep2 sep3 mutants even though third whorl identity and BC class gene expression is maintained (Pelaz et al., 2000). This suggests that SEP proteins act downstream of third whorl identity to activate CRC (Lee et al., 2005b), similar to the proposed role of SEP in conjunction with AP3/PI and AG to specify stamen identity in the floral quartet model (Section 1.4). Strikingly, nectary glands develop in ABC (ap2 ap3-2 ag-1) triple mutants (Baum et al., 2001). Recovery of nectary development in ABC triple mutants involves contribution from the SHATTERPROOF1/2 genes as ap2 ap3-2 ag-1 shp1 shp2 quintuple mutants lacks nectaries (Lee et al., 2005b). Lee et al 89  (2005b) proposed that SHP1/SHP2 replace lost AP3/PI to promote nectary initiation but only in the ABC mutant background. SHP1/SHP2 are normally expressed in nectaries at the time of nectary emergence (Flanagan et al., 1996; Savidge et al., 2005; Baum et al., 2001) suggesting that they may function redundantly with AP3/PI to specify nectaries. Interestingly, AP3 is expressed early in the nectary primordium and later restricted to the nectary base while PI is not expressed (Baum et al., 2001). AP3 forms obligate heterodimers with PI (Reichmann et al., 1996) to specify floral organ identity (see Introduction, Section 1.4.1). It is unclear if AP3 interacts with a partner other than PI in nectary tissue. I characterised nectary development in bop1 bop2 and investigated the contribution of CRC and cellular proliferation to nectaries in bop1 bop2. Data presented in Sections 4.3.1 and 4.3.2 was published in McKim et al. (2008). All results presented in this chapter are my sole effort.  3.2 METHODS 3.2.1 Plant materials and growth conditions The bop1 bop2 alleles refer to the T-DNA mutants bop1-3 and bop2-1 alleles are in the Col-0 background and were obtained from the Arabidopsis Biological Resource Center (ABRC). Both alleles contain 5’UTR T-DNA insertions and were described in Hepworth et al. (2005). The bop1-5 bop2-2 alleles were described in Norberg et al (2005). CRABS CLAW (CRC)::GUS transgenic plants (Baum et al., 2001) and crc-1 mutants (Bowman and Smyth, 1999) were kindly provided by Dr. John Bowman (Monash University) and are both in the Landsberg erecta (Ler) ecotype background. Floral developmental stages were determined according to Smyth et al. (1990). The sup-1 allele was described in Bowman et al. (1992) and is in the Col background. 90  Triple bop1 bop2 sup-1 mutants were recovered in the F2 population of a bop1 bop2 x sup-1 cross. To control for ecotype effects, bop1 bop2 and sup-1 specimens were taken from the F2 segregating population.  3.2.2 β-Glucuronidase activity staining and microscopy Samples were prepared as described in Section 2.2.5 for β-glucuronidase (GUS) staining and 2.2.6 for scanning electron microscopy (SEM), histological sectioning and light microscopy. CRC::GUS samples were stained 37 ºC for 2 hours.  3.3 RESULTS 3.3.1 BOP1 and BOP2 are necessary for nectary development During my AZ analyses, I noticed that bop1 bop2 flowers did not show obvious nectary glands. Analysis of the F2 progeny of a bop1 bop2 x wild type cross demonstrated that this phenotype co-segregated with other bop1 bop2 phenotypes (data not shown). Wild-type position 4 flowers, with sepals and petals removed, show paired lateral nectary glands protruding at the base of the lateral stamens (Fig. 3.1A). In contrast, lateral stamens of bop1 bop2 lack large nectary outgrowths (Fig. 3.1B). Wild-type lateral nectaries have characteristic deeply reticulated cuticle and associated secretory stomata (Fig. 3.1C), while in bop1 bop2, the area where lateral nectaries normally develop has slightly bulging areas of weak striation and absence of secretory stomata (Fig. 3.1D); medial nectary bulges were not obvious either in scanning electron micrographs or histological section suggesting that these may be absent or strongly reduced. From the onset of nectary development, two distinct nectary cell types exist: an outer epidermal layer and an inner starch granule-containing parenchymal tissue (Baum et al., 2001) as seen in transverse and longitundinal sections (Fig. 3.1E, G); a ridge of connecting nectary tissue  91  Figure 3.1 Nectary morphology and CRC::GUS activity in wild type (Col-0) and bop1 bop2 (A, B) Sepals and petals were removed from wild-type (A) and bop1 bop2 (B) position 4 flowers. Lateral nectary protrusions are obvious in wild type (arrow) but not in bop1 bop2. (C) Pair of lateral nectary glands with secretory stomata (arrow) in wild type. (D) Lateral nectary glands absent in bop1 bop2. (E-H) Toluidine blue-stained sections from position 3 flowers. (E) Wild-type transverse section shows paired lateral nectary glands with distinct epidermal and starch-containing parenchymal regions, a ridge of connecting nectary tissue (arrow) and a medial nectary gland. (F) Corresponding bop1 bop2 cross section shows two primordia (arrows) in the lateral nectary position. (G) A lateral nectary gland visible in longitudinal section of wild type. (H) Corresponding longitudinal section of bop1 bop2 shows a small outgrowth (arrow) in the lateral nectary position. (I-N) CRC::GUS plants stained for GUS activity. (I-K) CRC::GUS in wild type (Col-0 x Ler). (I) Expression in the nectary analgens of a stage 7/8 flower (arrow). (J) Strong nectary gland expression in a mature flower. (K) Position 7 flower with GUS staining nectaries and the ring of connecting nectary tissue. (L-N) CRC::GUS activity in bop1 bop2 (Col0 x Ler). (L) Expression in stage 7/8 flowers in the nectary analgens (arrow). (M) Strong expression in the nectary gland region of a mature flower. (N) GUS staining in position 7 flowers shows strong staining in the lateral nectary gland region and weaker staining in the medial and connecting nectary tissue regions. (O) bop1-5 bop2-2 flowers (null alleles) also show loss of nectary characteristics ln, lateral nectary; ls, lateral stamen; mn, medial nectary; p, petal; s, sepal. Scale bars, C,D,O 500 µm; E-H,J,K,M,N 70 µm; I,M 35 µm. McKim et al. (2008).  92  dev.biologists.org  93  also is present (Fig. 3.1E, arrow). The corresponding section of bop1 bop2 lacks distinct epidermal and parenchymal cells but shows slight bulges where the paired lateral glands would normally arise (Fig. 3.1F, H arrows). Thus, although some residual cell division may occur, bop1 bop2 lacks differentiation of most nectary tissue characteristics. This phenotype correlates with BOP expression patterns as seen for BOP2 expression by in situ hybridisation (data not shown) and BOP1::GUS staining in nectariferous tissue throughout development (Fig. 2.8). BOP2 is expressed at very low levels in bop1 bop2 plants (Hepworth et al., 2005), raising the possibility that the small bulges may result from leaky BOP2 expression. To address this concern, the bop1-5 bop2-2 null allele double mutant (Norberg et al., 2005) was also examined for nectary defects. As shown in Figure 3O, this double null also shows underdeveloped bulges where nectaries would normally develop suggesting that residual nectary bulges are a true loss of BOP function phenotype.  3.3.2 Genetic interaction between CRABS CLAW and BOP1 and BOP2 CRABS CLAW (CRC) is a key gene regulating nectary development in Arabidopsis and encodes a putative zinc finger transcription factor containing a YABBY domain (Smyth et al., 1990; Davis et al., 1994); Siegfried et al., 1999). While bop1 bop2 flowers retain residual bulging reminiscent of nectary glands, crc mutants show a complete loss of nectary development (Bowman and Smyth, 1999). To determine whether loss of nectary growth observed in bop1 bop2 was due to misregulation of CRC expression, the CRC::GUS expression pattern was examined in bop1 bop2. The wild-type CRC::GUS expression pattern was determined in Col-0 x Ler background to compare with bop1 bop2 CRC::GUS plants generated by crossing. As described in Baum et al. (2001), CRC::GUS is expressed in stage 7/8 flowers in the nectary gland anlagen (Fig. 3.1I); expression then expands throughout the connecting nectary tissue 94  between the glands and is maintained after abscission of floral organs (Fig. 3.1J, K). CRC::GUS is also expressed in stage 7/8 bop1 bop2 flowers (Fig. 3.1L), suggesting that BOP1 and BOP2 are not necessary for CRC expression in the nectary anlagen. As bop1 bop2 flowers mature, receptacle regions expressing CRC::GUS expand into presumptive lateral nectary regions and in connecting areas although at reduced levels compared to wild type (Fig. 3.1M, N). Thus, in bop1 bop2, CRC::GUS expression in nectary primordia appears similar both temporally and spatially compared to wild type, suggesting that altered spatial and temporal regulation of CRC transcription is not responsible for the loss of nectary elaboration in bop1 bop2. CRC::GUS is normally restricted to the flower but is observed at the tip of developing floral bracts in bop1 bop2 (Fig. 3.1O). Moreover, expression of BOP1::GUS reporter gene appears unchanged in a crc-1 background (data not shown), suggesting that CRC does not regulate the transcription of BOP1. Nectary development was completely abolished in bop1 bop2 crc-1 triple mutants (data not shown) showing that the severe nectary phenotype of crc-1 is epistatic to that of bop1 bop2.  3.3.3 Restoration of nectary elaboration in bop1 bop2 by superman-1 In bop1 bop2, the nectaries appear arrested at primordia stage and do not develop diagnostic mature nectary surface characteristics, such as wavy epidermal cuticular ridges and modified secretory stomata, referred to here as a loss of nectary elaboration (Fig 3.1). Loss of function in the SUPERMAN gene causes a proliferation of stamens at the expense of carpels, interpreted as an expansion of third whorl identity into the central fourth whorl (Schultz et al., 1991; Bowman et al., 1992; Sakai et al., 1995; 2000). All stamens in sup-1, including ectopic ones, are subtended by nectaries, confirming that nectaries are associated with the third whorl domain (Baum et al., 2001). As previously reported, the outermost nectaries in sup-1 are greatly enlarged (Fig. 3.2C; Baum et al., 2001). 95  Figure 3.2 Restoration of mature nectary surface characteristics in bop1 bop2 sup-1 (A-G) Nectaries in position 4 flowers (A) lateral nectaries in a bop1 bop2 (Col-0) flower are present but lack outgrowth, heavily reticulated cuticle and secretory stomata. (B) Enlarged mature nectaries subtend all stamens in sup-1. (C) Mature paired lateral nectary in Col-0. (D-F) bop1 bop2 sup-1 (D) single protruding lateral nectary shows reticulated cuticle and a secretory stomate (arrow). (E) Medial nectary outgrowth tipped with secretory stomata (arrow). (F) high magnification image of medial nectary showing stamen-like cuticle, * indicates cuticle with smooth surface. Scale bars A-D, 200 µm.  96  Although initiation of nectaries requires third whorl identity only and is independent from floral organ identity, complete realisation of nectary characteristics appears to require activity of floral organ homeotic genes as class B, apetala3-1 (ap3-1) and pistallata-1 (pi-1), and class C, agamous (ag-1), mutants lack stamens and, although not to the degree observed in bop1 bop2, also exhibit stalled nectary development and a loss of nectary elaboration (Baum et al., 2001; Sakai et al., 2000). Nectary morphology was restored in ap3-3 sup-1 and pi-1 sup-1 mutants (Sakai et al., 2000) suggesting that sup-1 is able to promote developed nectaries regardless of compromised B class activity. I reasoned that this enhancement from sup-1 allele may rescue the bop1 bop2 nectary defect as well. To test this prediction, nectary development was assessed in bop1 bop2 sup-1 triple mutants. Nectaries in position 4 flowers of bop1 bop2 sup-1 triple mutants indicate that sup-1 was able to compensate in great measure for loss of nectary elaboration in bop1 bop2. Lateral nectary glands of the triple mutant were larger, fuller in body and developed heavily striated cuticle and secretory stomata (Fig. 3.2D, E) resembling wild type (Fig. 3.2C). The lateral nectary, normally consisting in wild type of two paired glands which encircle the lateral stamen (Fig. 3.2C), often formed as a single, large bar-shaped gland at the base of the lateral stamens in bop1 bop2 sup-1 flowers (Fig. 3.2D) suggesting that sup-1 was not able to recover all aspects of wild-type nectary morphology. Medial nectaries also show rescue, growing outward/upward extensively and exhibiting mature nectary surfaces with reticulated cuticles and secretory stomata (Fig. 3.2E). Strips of smooth-surfaced cells resembling the bases of stamen filaments were often detected at the base of the medial and lateral glands (Fig. 3.2F, asterisk)  97  3.4 DISCUSSION 3.4.1 Loss of nectary elaboration and CRABS CLAW expression in bop1 bop2 During the course of nectary development, two events occur - initiation of the primordia and subsequent sustained cell proliferation and further differentiation of nectaries (Davis et al., 1986; Baum et al., 2001). Nectaries are not entirely absent in bop1 bop2 but greatly reduced in size and lack mature features such as parenchymal secretory tissue and modified stomata suggesting that nectaries are stalled after the initiation event. Elaboration of mature nectary characteristics, such as reticulated cuticle and modified stomata, may depend on attaining a critical cell number during gland development, as small nectaries which display mature surface characteristics and secrete nectar have not been reported. Precise timing of nectary tissue proliferation and nectar exudation is crucial to attract pollinators at appropriate stages of flower maturation; i.e. when pollen is produced and/or the carpels become receptive to fertilisation. Despite this, little is known about the underlying genetic regulation of nectary gland morphogenesis following primordia initiation. Formation of nectary primordia is dependent upon third whorl formation and CRC expression (Baum et al., 2001). Similar to CRC, BOP is expressed very early in nectary development. My analysis suggests that bop1 bop2 mutants retain wild-type spatial and temporal CRC::GUS activity in both the presumptive nectary and the bulges that later develop, indicating that the lack of nectary outgrowth is not due to a loss in CRC activation and that positional cues are still directing nectary specific expression; however, this does not preclude a role for BOP1 and BOP2 in enhancing CRC expression as CRC::GUS activity appeared weaker in bop1 bop2. Similarly, LFY may also contribute to the intensity but not the spatial regulation of CRC expression in nectaries (Lee et al., 2005b). Other evidence suggests that BOP1 and BOP2 share a role with LFY as well as AP1 in regulating CRC expression. Ectopic CRC::GUS expression in 98  floral bracts of bop1 bop2 is similar to that observed on the abaxial side of the pedicel of in lfy-6 and ap1-1 flowers (Baum et al., 2001). In ap1-1, CRC misexpression is associated with ectopic nectary glands and filamentous organs (Baum et al., 2001). These filamentous projections are markedly similar in position and morphology to the highly reduced bracts observed in some bop1 bop2 flowers (Hepworth et al., 2005). In chapter 5, I propose that LFY, AP1 and BOP1 and BOP2 have shared functions in floral meristem identity and bract development and hence may also share a role in limiting CRC::GUS expression to the flower and promoting high levels of expression in presumptive nectary tissue.  3.4.2 Organ identity, boundary position and nectary elaboration Although nectary initiation is independent from floral organ identity, elaboration of nectaries is often reduced and/ or lost in the B class (ap3-1 and pi-1) and C class (ag-1) mutants (Baum et al., 2001) suggesting that differentiation of nectariferous tissue is influenced by floral organ gene expression. As shown in Table 1, elaboration of nectaries as gauged by nectar secretion was most robust in mutants when first and third whorls were occupied by sepals and stamens, respectively, as occurs in wild type, suggesting that the juxtaposition of these two organs may promote nectary development. This potentially explains why in sup-1 and 35S::UFO mutants, those nectaries subtending stamens of the canonical third whorl bordered by sepals are exaggerated (Table 3.1; Baum et al., 2001). Nectaries develop in between the first whorl organs of ap2-2 mutants (Baum et al., 2001), which lack second whorls (Bowman et al., 1991), suggesting that the second whorl may normally restrict nectary growth. Given that petals and nectary gland outgrowth occurs along the same plane, lateral and medial glands are effectively constrained by adjacent petal claws. This relationship is diagrammed in Figure 3.3A and proposes that the  99  Table 3.1 Association between first and third whorl organs and nectary growth Table based on results presented in Baum et al (2001). Se, sepal; Pe, petal; St, stamen; Ca, carpel  Mutant  Floral Plan  Floral  Floral organs at  organ in  1:3 whorl  nd  2 whorl  Nectar Secretion  boundary 1carpel:3stamen  A class (ap2-2)  CaN?, - , St, Ca  None  A class (ap1-1)  Se, - , NSt, Ca  None  B class (ap3-1)  Se, Se, NCa, Ca  Sepal  B class (pi-1)  Se, Se, NCa, Ca  Sepal  C class (ag-1)  (Se, Pet, NPet)n  Petal  lfy-6  Se, Se, NCa, Ca  Sepal  ufo-2  Se, Se, NCa, Ca  Sepal  35S::PI  Se/Pe Pe NSt Ca  Petal  35S::AP3  Se Pe NSt St/Ca  Petal  A, C doubles  (Ca/Lv, St/Pe, NSt/Pe)n  St/Pe  sep1 sep2 sep3  Lv, Lv, Lv, Lv  Lv  Lv:Lv  No  B, C doubles  Se, Se, Se, Se  Sep  -  No  A, B, C  Ca/lv,-, N-  None  sup-1  Se, Pe, N*St, NSt  Pet  35S:UFO  Se, Pe, N*St, NSt  Pet  A:BC 1sepal:3stamen A:BC 1Sepal:3carpel A:C 1Sepal:3carpel A:C 1Sepal:3petal A:AB 1sepal:3carpel A:C 1sepal:3carpel A:C Sepal/petal:stamen AB:BC Sepal:stamen A:BC (2/3):(2/3) ~BC:~ABC  Leaf:carpel C:C Sepal:stamen A:BC Sepal:stamen A:BC  Sometimes  Yes  Infrequent  Infrequent  No  No  No  No  Yes  No  No  Yes  Yes  *enhanced nectariferous tissue formation 100  Figure 3.3 Whorl models and nectary formation (A) wild type flowers are characterised by alternating floral organs in the first, second and third whorls. Nectary development is enhanced due to appression between the third whorl stamens and first whorl sepals. Second whorl petal organs act to repress nectary formation from the first whorl as well as promote sepal boundaries in the first whorl. (B) In bop1 bop2 flowers, the alternation of organs of the first two whorls is lost such that petals often overlie sepals rather than forming between them. This may weaken the ability of sepals to promote nectary gland formation. Furthermore, the abaxial sepals have petal identity and may repress nectary tissue on the opposing stamens. se, sepal; pe, petal claw; ls, lateral stamen filament; ms, medial stamen filament; ln, lateral nectary; mn, medial nectary; ca, carpel; ps, petalloid sepal; br, bract.  B  A se ls  mn pe  ms  ln ca  ps  br wild type bop1 bop2  101  boundary between juxtaposed sepals and stamens enhances nectary proliferation and that second whorl organs restrict nectary gland outgrowth. In bop1 bop2, the pentamerous first whorl results in a different spatial arrangement between the organs such that the sepals and stamens are not directly opposed. In addition, petals overlap the sepals and encroach into the boundary region along the sepal where glands would normally subtend (Fig. 3.3B). Proposed promotive relationship between sepals and stamens may be weakened in bop1 bop2 due to this positioning. Furthermore, petals undergo extensive cell proliferation between stages 7 and 8 (Irish, 2007) while nectary proliferation does not begin in earnest until stage 9 (Davis et al., 1990; Smyth et al., 2001; Baum et al., 2001). Thus, receptacle cells between stamens and sepals normally available for nectary proliferation may be depleted by the intervening petals in bop1 bop2.  3.4.3 Enhanced proliferation permits elaboration of nectary glands in bop1 bop2 Rescue of nectary elaboration by sup-1 suggests that increased proliferation in the third whorl compensates for the nectary defects observed in bop1 bop2. Sakai et al. (1995; 2000) proposed that expansion of third whorl identity into the fourth whorl in sup-1 mutants reflects increased cellular proliferation specifically along the third/fourth whorl boundary region suggesting that SUP’s primary role is to actively repress cellular proliferation. Prunet et al. (2008) recently showed increased proliferation in sup-1 results from prolonged WUS expression and reduction of AG activity (Prunet et al., 2008). Ectopic expression of SUP throughout all four whorls caused reductions in organ number, particularly in the second and third whorls (Yun et al., 2002) indicating that SUP specifically represses B class cellular proliferation. Restoration of nectary growth in bop1 bop2 sup-1 plants may indicate that sup-1 leads to increased cellular proliferation along the region of the third whorl where the ring of nectary tissue normally develops. Rescue of 102  medial and lateral nectaries in bop1 bop2 sup-1 may indicate that a loss of proliferative capacity of initiated nectaries underlies nectary defects in bop1 bop2 flowers.  3.4.4 BOP1 and BOP2 pattern multiple tissues at the receptacle BOP1 and BOP2 promote abscission zone (AZ) anatomy, a feature of floral organ-receptacle boundaries (Chapter 2). AZs develop adjacently to nectaries and both are characterised by small, cytoplasmically dense cells. In this regard, nectaries and AZs resemble meristematic tissue anatomically; however, functionally both tissue types are highly specialised for secretory events: the AZ, of cell wall degrading enzymes and the nectary, of nectar. Thus, BOP1 and BOP2 mediate patterning of two types of secretory tissue in the receptacle. Interestingly, although not mentioned by Cai and Lashbook (2008), micrographs of overexpressing AtZFP2 lines which lack floral organ abscission, clearly show a loss of nectary development, which hints at an association between the development of AZs and nectary formation.  103  4 SUPPRESSORS OF bop1 bop2 4.1 INTRODUCTION Secondary screens are used to discover additional genetic components in regulatory pathways identified by the original mutation (Page and Grossniklaus, 2002). A particularly powerful approach involves screening for additional second-site modifiers which either exacerbate the original defect (enhancers) or weaken the mutant to approximate a wild type phenotype (suppressors). I screened for suppressors of bop1 bop2 rather than enhancers given that an enhancement of a complete loss of abscission would be difficult or impossible to detect. The reversion to wild type observed in suppressor mutants presumably reflects either an intragenic mutation that results in a functional gene product from the original gene or an extragenic mutation in a second gene that results in an altered gene product which compensates for the original mutation. The extragenic suppressors could encode proteins which enable an alternative pathway (Forsburg, 2001) or restore orthodox regulation to a target normally controlled by the primary mutated protein. To uncover other players in developmental programs influenced by BOP function, a mutagenised population of bop1 bop2 seed was grown and screened for plants reverting to wildtype phenotypes including morphology of leaves and flowers as well as plants showing recovered floral organ abscission. Several outcomes from the suppressor screen were possible. I was interested as to whether suppressors which demonstrated complete reversion of all bop1 bop2 phenotypes to wild-type could be recovered. These mutations would presumably affect factors central to the BOP regulatory mechanism. Recovery of mutants that manifest suppression for only certain aspects of the bop1 bop2 phenotype would suggest that the mutation affect genes encoding products with a function unique to that process.  104  4.2 METHODS 4.2.1 Plant materials, mutagenesis and growth conditions Approximately 15 000 bop1 bop2 seeds were mutagenised by incubating in 0.25% ethylmethyl sulphonate (EMS) in water for 13 hours. Seeds were subsequently washed with water and sown to a density of 100 seeds per six inch pot. These M1 bop1 bop2 plants were allowed to selffertilize and M2 seeds harvested as several EMS M2 bulk populations. M2 seedlings were planted in 72 plant dimple trays and grown as described in Section 2.2.1.  4.2.2 Genetic screen, suppressor isolation and segregation analysis A total of 1800 plants were screened by a directed-studies student Nicole Strümpfer, under my supervision, for suppression of leaf, floral patterning and abscission phenotypes of bop1 bop2. Putative suppressors were confirmed to be homozygous for bop1 bop2 by genotyping (primers listed the Appendix). These were re-grown and confirmed to have the suppressor phenotype. Plants were scored for abscission at various flower positions either due to natural organ fall or following mild mechanical stress where the receptacle was lightly squeezed between the thumb and index finger. Suppressor lines were back crossed to bop1 bop2 and resultant F2 generations were scored for the suppression phenotype to determine inheritance patterns (rva1 bop1 bop2 x bop1 bop2, n = 144; rva2 bop1 bop2 x bop1 bop2, n = 101; n = total F2 population scored). Data was analysed with a 2 test (df = 2, p > 0.05).  4.2.3 Microscopy Samples were prepared as described in Section 2.2.6.  105  4.3 RESULTS 4.3.1 Isolation of lines showing recovered abscission in sepals and petals From 1800 EMS mutagenised bop1-3 bop2-1 plants screened, 17 putative suppressors were isolated; however, only three were confirmed to be homozygous for bop1-3 bop2-1 mutations by genotyping. The bop1-3 bop2-1 alleles (hereafter referred to as bop1 bop2) are from the SALK T-DNA collection (http://signal.salk.edu/cgi-bin/tdnaexpress) and thus sequence reversion is unlikely to be a possible suppression mechanism. One plant line, 3.10, showed weak suppression of bop1 bop2 leaf and floral patterning phenotypes which was lost upon secondary screening (data not shown) while two other lines, 24.51 and 26.71 showed recovery of floral organ abscission (Fig 4.1A). These latter lines were renamed recovered abscissor1 (rva1) and recovered abscissor2 (rva2). Both rva1 and rva2 showed restored floral organ abscission of petals and sepals but not stamens in bop1 bop2 plants following mechanical roughing with the fingertips. Such abscission was not observed in bop1 bop2 following mechanical roughing. The suppressor phenotypes for rva1 and rva2 were confirmed and characterised in the next generation.  4.3.2 Penetrance and expressitivity of recovered abscission phenotypes Both supressors exhibited incomplete penetrance with 85% of rva1 bop1 bop2 and 80% of rva2 bop1 bop2 exhibiting the recovered abscission phenotype (Table 4.1). Stronger suppression was observed in rva1 bop1 bop2 plants which manifested a range of expressivities (Table 4.1) from petal and sepal loss by position 6-7, often without mechanical roughing, to almost all sepals and petals shed by position 9-10 (moderate suppression) and finally abscission of variable number of  106  Table 4.1. Phenotype characteristics and segregation of restored floral organ abscission mutants rva1 bop1 bop2 and rva2 bop1 bop2  Suppressor  rva1 bop1 bop2 n = 54  rva2 bop1 bop2 n = 20  Penetrance  85%  80%  Trait severity  Expressivity  3:1 2 crit = 5.99 p > 0.05  strong = petals and sepals abscise at position 6-7  moderate = petals and sepals abscise at position 9-10  mild = some petals and sepals abscise at position >10  strong = > 1 petals or sepals abscise at position 7-8  moderate = > 1 petals or sepals abscise at positions 911  mild = ≥ 1 petals and sepals abscise at positions >11  37%  44%  19%  38%  19%  38%  2.37  2.78  107  Figure 4.1 Recovered abscission phenotypes of rva1 bop1 bop2 and rva2 bop1 bop2 suppressors (A) Position 7 siliques from Col-0 and bop1 bop2 and rva1 bop1 bop2 plants and position 9 silique from rva2 bop1 bop2. All floral organs have abscised from Col-0 siliques while all remain attached in bop1 bop2. Sepals and most petals have abscised from the suppressor rva1 bop1 bop2 and rva2 bop1 bop2 but stamens are retained (arrows). Siliques from rva1 bop1 bop2 and rva2 bop1 bop2 represent the strong expressivity phenotype of each suppressor. Note the shorter silique length in the rva2 bop1 bop2 specimen. (B) Rosette leaves 4-7 and (C) mature flowers from Col-0, bop1 bop2, rva1 bop1 bop2 and rva2 bop1 bop2 indicate that the leaf and floral patterning phenotypes of bop1 bop2 are not suppressed by rva1 or rva2.  108  petals and sepals in position 10 or older siliques (mild suppression). Weaker suppression was observed in rva2 bop1 bop2 which usually required mechanical roughing for floral organ abscission. In this context, the rva2 bop1 bop2 phenotype ranged in expressivity from strong where more than one petal or sepal abscises by position 7-8, moderate where at least one petal or sepal abscises by positions 9-11 and mild as observed by petal or sepal abscission in siliques older than positions 11 (Table 4.1). Interestingly, rva1 bop1 bop2 and rva2 bop1 bop2 retained bop1 bop2 leaf and flower patterning phenotypes (Fig 4.1B, C). Occasionally, an entire cohort of rva2 bop1 bop2 developed dramatically shorter siliques, 50% the length of bop1 bop2 siliques, which were often sterile (Fig 4.1A, Table 4.2). However, rva2 bop1 bop2 populations grown under similar conditions did not exhibit any reduction in silique length. This may also reflect a background mutation segregating in the suppressor background. Single seed descent analysis is ongoing to evaluate this possibility.  4.3.3 Segregation patterns of recovered abscission phenotype of rva1 and rva2 in bop1 bop2 background As shown in Table 4.1, rva1 and rva2 segregate as single, recessive mendelian loci. No novel phenotypes were detected when rva1 bop1 bop2 or rva2 bop1 bop2 were crossed to wild type. Complementation crosses between rva1 bop1 bop2 and rva2 bop1 bop2 are in process.  4.3.4 Abscission zone development in rva1 bop1 bop2 and rva2 bop1 bop2 Restoration of abscission in sepals and petals by rva1 and rva2 may result from several mechanisms. On the one hand, AZ anatomy may be partially restored either in every AZ cell or only in certain files/ regions of the zone leading to the partially recovered abscission phenotype  109  Table 4.2 Mature silique length of rva2 bop1bop2 is shorter than bop1 bop2 and Col-0  Line  Mature silique length ± stdev  % sterile siliques  rva2 bop1 bop2 bop1 bop2 Col-0  6.6 ± 2.8 13.4 ± 3.9 12.2 ± 2.5  21.4 4.8 2.9  110  of the suppressors. Alternatively, enhanced activity of an abscission-promoting signal may compensate for the lack of specialised AZ anatomy. An altered signal is unlikely to be excess IDA because, as described in Section 2.3.7.1, expression of 35S::IDA in bop1 bop2 fails to rescue the loss of abscission defect. To evaluate these possibilities, I examined changes occurring in the AZ at different stages of flower maturation in rva1 bop1 bop2 and rva2 bop1 bop2 via scanning electron microscopy (SEM). 4.3.4.1 Sepal and petal floral organ AZs in rva1 bop1 bop2 show partial development and disorganised cellular expansion Sepals and petals often abscise without mechanical stimulation in rva1 bop1 bop2 plants showing strong suppression. As displayed in Fig 4.2A, a dissected position 6 rva1 bop1 bop2 flower receptacle sports a mature sepal AZ which shed a sepal unaided, a partially developed petal AZ which abscised with mechanical roughing and an inactivated stamen AZ (stamen was dissected away). An equivalently staged Col-0 receptacle shows nascently exposed AZ scars (Fig 4.2B, Fig 2.4A) while a partially dissected bop1 bop2 flower has ruptured cells following dissection of petals and sepals (Fig 4.2C, Fig2.4B). Similar to wild type and bop1 bop2, petal AZs in position 3 and 4 rva1 bop1 bop2 flowers (just after anthesis) showed broken cells following organ removal (Fig 4.2D, E). Petal AZ cell fracture was also common in position 5 rva1 bop1 bop2 flowers (Fig. 4F) although some peripheral cells were intact (Fig. 4F, asterisks) suggestive of middle lamellae weakening (compare to Col-0 Fig. 2A). Intact cells in the sepal AZs of position 5 flowers (Fig 4.2F, G) and petal and sepal AZs in position 6 flowers (Fig 4.2H, I) suggests that the separation layer middle lamellae is degraded; however, this weakening is not  111  Figure 4.2 Scanning electron microscopy of rva1 bop1 bop2 floral organ abscission zones Scanning electron micrographs of position 6 flower receptacles. The fracture plane on the receptacle (arrows) was observed following removal or natural abscission of floral organs. (A) rva1 bop1 bop2 receptacle exhibits a smooth fracture surface from which a sepal has abscised (s); however, the petal AZs revealed on the surface of the receptacle show partial cell breakage and the stamen filament AZ displays mainly broken cells. (B) Wild-type (Col-0) receptacles show differentiation of AZ cells into protective surfaces layers. (C) bop1 bop2 receptacles show broken fracture plane where sepals and petals were attached and have been dissected away. (DK) rva1 bop1 bop2 floral organ AZs at position 3, 4, 5, 6, 8, 12 and 18. (D, E) Position 3 and 4 petal AZs show broken cells following petal removal. (F) Petal removal from position 5 flowers reveals ruptured cells laterally bordered with a few smooth cells (G) Sepal AZs from position 5 flowers often display smooth fracture cavities. (H-I) Petal AZs from position 6 flowers show a range of surface characteristics. (H) fracture plane of the petal AZ is mostly smooth with an obvious central fractured hole. (I) another position 6 petal AZ showing enlarging cells and central fractured cells. (J) broken cells surround a central column jutting out from the receptacle surface in a position 6 flower. (K) position 8 petal AZ shows a ruptured central cell and smooth expanding cells. (L) Expanding surface cells in position 12 AZs extend in irregular tip-like growth pattern (arrowheads). (M) AZs on position 18 siliques reveal intact cells except for broken vascular bundles. Arrow (a) points to the single petal vascular bundle and arrow (b) to the left most one of three sepal bundles. (N, O) Receptacle of position 18 siliques with the sepal AZ and petal AZ demarcation outlined. (N) Receptacle of rva1 bop1 bop2 shows unabscised anther filaments and post-abscission differentiating petal and sepals AZs composed of ruptured cells, irregular surfaces of small, expanding cells and less obvious distinctions between zones. (O) Receptacle of position 18 wild-type siliques show large, relatively uniform cells in clearly delineated AZs. Scale bars A, N, O 600 µm; B, C, 750 µm; D-M, 400 µm.  112  113  uniform as petal AZs showed fracture of the central core, potentially of vascular tissue, (Fig. 4.2H, I, asterisks) which sometimes jutted out from the receptacle plane (Fig. 4.2J, arrow). Petal AZs develop an increasing number of diffusely expanding cells along the fracture plane as depicted in positions 8, 12 and 18 (Fig 4.2K-M). Unlike the roughly spherical, swollen cells of wild type (Fig. 4.2B; Fig. 2.4A), AZ scars in position 12 and 18 rva1 bop1 bop2 siliques exhibited irregularly protruding tubular and bent cells expanding anisotropically outward (arrowheads, Fig. 4.1L, M). New epidermal surfaces on stage 18 siliques are characterised by relatively few, enlarged and rounded cells clustered into distinct scars (Fig. 4.2O). Cells exposed on rva1 bop1 bop2 AZs, however, consist of numerous, irregular, small cells which form a disorganised surface (Fig. 4.2N). Interestingly, centrally located AZ cells remain ruptured on position 6 siliques and may indicate severed vascular bundles (Fig 4.2E, asterisk). Strikingly, several position 18 siliques examined displayed obvious holes on the petal and sepal AZs scars (Fig. 3M, arrows) which may represent sheared vascular strands. Arabidopsis floral organs exhibit highly reduced vasculature compared to leaves (Nelson and Dengler, 2001). Sepals and carpels are vascularized with three main strands and petals and stamens with one main vascular strand each (Bossinger and Smyth, 1995). Thus, the top hole (Fig. 4.2M, arrow a) may correspond to the petal vascular trace while the other (Fig. 4.2M, arrow b) may be a lateral sepal trace. 4.3.4.2 Sepal and petal floral organ AZs in rva2 bop1 bop2 are partially activated Although abscission recovery is less strong in rva2 bop1 bop2 (Table 4.1), rva2 bop1 bop2 receptacles at position 6 show activated petal AZs with intact, rounded AZ cells (Fig. 34.3A, N) more similar to wild type AZs (Fig.3.3 B) than bop1 bop2 (Fig. 4.2C). Petal AZs displayed aberrant clusters of small, proliferating cells (Fig. 4.3A, N, asterisks). Although the middle 114  lamellae of the petal AZ on the rva2 bop1 bop2 flower appears to have weakened, the adjacent sepal AZ has not, as sepal removal resulted in broken, ruptured cells (Fig. 4.3A). As in wild type and bop1 bop2, petal removal from position 3 rva2 bop1 bop2 reveals a surface composed of ruptured cells (Fig. 4.3J). Position 4 petal removal often reveals surfaces with smooth, non-ruptured cells indicating degradation of AZ middle lamellae (Fig. 4.3K, L). However, in other flowers, petal fracture planes at position 5 (Fig. 4.3M) still display ruptured cells as in bop1 bop2 (Fig. 2,4B). Several position 6 AZs show a range of development from expanded cells as previously described (Fig. 4.3N) to smooth, flattened fracture planes in which individual cells appear covered by a layer of residual middle lamellar material (Fig. 4.3O, P). Position 8 petal AZs reveal a combination of broken and intact, spherical cells (Fig. 4.3Q) although the broken cells are not as centrally located as in rva1 bop1 bop2. Position 12 and 18 petal AZs often showed partial differentiation of protective surface yet most exposed AZ cells were deflated (Fig. 4.3R, S, asterisks). Position 12 and 18 receptacles of rva2 bop1 bop2 (Fig. 4.3D, G) depict petal and sepal AZ scars characterised by disordered, deflated, actively proliferating cells (compare with bop1 bop2, Fig. 4.3F, I). The fracture surface does not show much cell breakage but it is difficult to discern a truly broken cell from a deflated or torn edge. The distinction between different floral organ attachment regions, obvious in wild type (Fig. 4.3H) is lost in rva2 bop1 bop2 AZ scars (Fig. 4.3G), similar to the rva1 bop1 bop2 phenotype. However, unlike rva1 bop1 bop2, obvious holes were not commonly observed.  115  Figure 4.3 Scanning electron microscopy of rva2 bop1 bop2 floral organ abscission zones Position 6 receptacles. (A) rva2 bop1 bop2 shows an activated petal AZ (arrow) with enlarged rounded cells and a small cluster of tiny, oblong cells (*). (B) Col-0 position 6 with all floral organs abscised and intact AZ cells swelling into a protective layer. (C) dissected bop1 bop2 flower at position 6 showing cell rupture in the petal and sepal fracture planes. (D-F) Position 12 siliques. (D) rva2 bop1 bop2 (E) Col-0 receptacle showing post-abscission differentiating AZ scars (F) dissected bop1 bop2 receptacle revealing ruptured cells due to organ removal. (G-I) Position 18 siliques. p, petal AZ; s, sepal AZ; f, stamen filament AZ; ln, lateral nectary gland.  116  117  4.3.5 Nectary development in rva1 bop1 bop2 and rva2 bop1 bop2 Although abscission is recovered to varying degrees in rva1 bop1 bop2 and rva2 bop1 bop2, nectary development appears stalled in both lines (Fig. 4.4A, B) although not to the same degree as in bop1 bop2 (see section 2.3.8, Fig 2.4). Nectary-like bulges grow outward from the bases of stamens in post-anthesis rva1 bop1 bop2 and rva2 bop1 bop2 rva2 flowers and often show a reticulated cuticle (Fig.4.4A, B). However, these growths do not form active secretory stomata and thus never become functional nectaries as in wild type (Fig. 4.3C; Fig. 2.13A, E, G).  118  Figure 4.4 Nectary development in rva1 bop1 bop2 and rva2 bop1 bop2 (A-D) Lateral nectary glands. (A, B) Presumptive lateral nectarines form in rva1 bop1 bop2 and rva2 bop1 bop2 but they are smaller and do not develop heavily reticulated cuticle or secretory stomata as compared with wild type (C), similar to bop1 bop2 (D). Scale bars A-C, 600 µm.  119  4.4 DISCUSSION 4.4.1 rva1 and rva2 restore abscission to sepals and petals in bop1 bop2 Floral organ boundaries are characterised by small, cytoplasmically dense cell files of the AZ. The major defect of bop1 bop2 floral organ boundaries is a loss of this anatomy and instead, cells in the AZ position are expanded and vacuolated similar to mature lateral organ tissue. Secondary mutations in rva1 and rva2 rescue the ability of cells at the AZ to respond appropropriately to positional cues which are likely still operating in bop1 bop2 based on reporter gene analysis (see Section 2. 3.7.2). This would suggest that normally RVA1 and RVA2 promote lateral organ fate and/or suppress AZ development in opposition to BOP’s role in restricting proximal lateral organ growth and promotion of AZ formation. Alternatively, RVA1 and RVA2 may have a more general role to enhance intercellular adhesion; however, lack of other phenotypes in rva and rva bop1 bop2 would suggest this is the case. Understanding the molecular roles of RVA1 and RVA2 awaits molecular cloning of the corresponding genes. The morphological changes underlying recovery of sepal and petal abscission could involve restored AZ anatomy. Preliminary results suggest that AZ cell files are not evident in stage 13 flowers of either suppressor in histological section (data not shown). Given that abscission is often delayed in theses lines, development of AZ anatomy may also be delayed. Fine histological examination of the AZ in the suppressor lines at later stages should indicate the extent to which of anatomical recovery mirrors restored abscission behaviour. Alternatively, rva1 and rva2 may induce a change in the abscission program which bypasses the requirement of AZ anatomy. Certainly, features of the AZ scars in both lines suggest that the cells within the AZ are not completely restored to wild type.  120  4.4.2 Recovery of sepal and petal abscission can be separated genetically from stamen abscission Stamens are never seen to abscise in either rva1 bop1 bop2 or rva2 bop1 bop2 suggesting that stamen abscission may be under separate regulatory control, also regulated by BOP1 and BOP2, compared with sepals and petals. Unlike the sterile perianth, stamens are reproductive organs; thus, their retention depends on the fertilisation strategy of the plant which may lead to differential genetic control of abscission in these organs. Furthermore, although Arabidopsis sheds its floral organs in quick succession, this is the not always the case in other species where floral organs within a single flower can be shed along substantially different time frames (Addicott, 1982). Furthermore, the recently described ath1 mutant shows a specific block in stamen abscission while sepals and petals abscise (Gomez-Mena and Sablowski, 2008) also suggesting that perianth and stamen abscission may be regulated differently in Arabidopsis. Interestingly, loss of mature nectary characteristics is not significantly rescued in either rva1 bop1 bop2 or rva2 bop1 bop2. Nectary development is not necessary for AZ formation as crc-1 mutants are devoid of nectaries yet normally abscise floral organs (Bowman and Symth, 1999). These data could suggest that the converse also holds - that capacity to abscise is independent from formation of mature nectary glands. Nonetheless, abscission recovery is observed for sepals and petals only, while nectary development occurs off the stamens in the third whorl. Thus, lack of nectary recovery is still associated with a block of abscission in the third whorl organs in bop1 bop2 background. However, the ath1 mutant which lacks stamen abscission is not reported to show defects in nectary development, suggesting that stamen abscission is not a prerequisite for nectary development.  121  4.4.3 Organ specificity of abscission recovery The rva1 bop1 bop2 suppressor shows more complete temporal reversion to wild-type abscission behaviour for sepals compared with petals; in strong lines, sepals often abscise in a time frame similar to wild type without experimenter mechanical stimulation (compare with Fig. 2.4A) whereas petal abscission and AZ activation was comparatively delayed. Earlier sepal abscission could simply reflect the outer position of the sepal whorl making it more vulnerable to mechanical stress and thus abscission of this may indicate that loss of RVA1 function affects sepal tissue preferentially. Patterson and Bleecker (2004) reported that sepals and stamens abscised before petals in Col and Ws ecotypes. Intriguingly, the rva2 bop1 bop2 samples examined here often showed petal AZ activation prior to or in absence of sepal AZ activation; an opposite trend to that observed in rva1 bop1 bop2. It will be interesting to finitely stage whether petal abscission is always suppressed first in rva2 bop1 bop2.  4.4.4 Vascular abscission is not recovered in rva1 bop1 bop2 A defining aspect of rva1 bop1 bop2 abscission zone behaviour is a consistent rupture of central cells which may represent severed vascular tissue. Organ-plant body vasculature separation during abscission has not received detailed study in Arabidopsis but a variety of behaviours have been documented in other plants (Roberts et al., 2002; Andre et al., 1999; Lee, 1989; Addicott, 1982). Separation of the organ leaving a protruding strand of central vasculature like that observed in some rva1 bop1 bop2 petal AZs (Fig. 4.2J), is presumed to reflect vascular traces broken by purely mechanical forces (Addicott, 1982). Lack of vascular sealing in rva1 bop1 bop2 may reflect insufficient swelling and/or reduced tylose infiltration due to the aberrant expansion behaviour of nascently exposed rva1 bop1 bop2 separation layer cells. Alternatively, 122  non-ruptured cell separation in parenchyma cells and not the vasculature of rva1 bop1 bop2 suggests that the plant may regulate abscission in these two tissues independently. This condition is remarkably similar to vestigial AZs of cauline leaves which always show vascular cleavage (Section 2.3.5, Fig. 2.5) strengthening the notion that vascular severance can be developmentally isolated from enlargement of parenchymal cells in AZ regions. Furthermore, different isoformas of abscission-related enzymes are expressed in the parenchymal versus vascular abscission zone cells (Hong et al., 2000). In addition, bean dissection experiments suggest that the AZ parenchymal tissue is capable of abscising independently from the vascular tissue (Thompson and Osborne, 1994). Therefore, the rva1 bop1 bop2 suppressor may restore abscission behaviour to the parenchymal regions only, approximating a vestigial AZ. IDA is expressed normally in floral organ AZs in bop1 bop2 (Section 2.3.7.2, Fig. 2.12) and would be capable of activating this zone as demonstrated for vestigial cauline AZs when ectopically expressed (Fig. 2.10F; Stenvik et al., 2006).  4.4.5 rva1 bop1 bop2 and rva2 bop1 bop2 shows aberrant cell expansion An unresolved problem in Arabidopsis abscission biology is the extent to which separation layer expansion contributes to abscission; however, in other species, such as Lupinus augustifolius, cellular expansion is considered an active force in cell separation during abscission (Clements and Atkins, 2001). Exposed cells on AZ scars of rva1 bop1 bop2 and rva2 bop1 bop2 display a variety of disorganised and deflated cell shapes raising the possibility that loss of uniform, isotropically expanding cells may contribute then to lack of complete abscission recovery in these lines. Disorganised, reduced cellular expansion and lack of AZ scar features in rva1 bop2 bop2 may also indicate that the cells of the separation layer remaining on the plant body are misidentified. Cell deflation in rva2 bop1 bop2 may indicate inadequate vacuolar expansion and/ 123  or reduction in vacuolar volume following initial expansion or incomplete differentiation of the newly expanded cell wall. Nonetheless, floral organ abscission does occur in these lines suggesting that the ability to abscise is not mutually dependent on cellular enlargement and/or later morphogenesis of the protective scar.  4.4.6 Suppressors for other bop1 bop2 phenotypes were not recovered Both rva1 bop1 bop2 and rva2 bop1 bop2 showed recovery of floral organ abscission for sepals and petals in the bop1 bop2 background but no strong suppression of either the floral patterning or leaf phenotypes suggesting that floral and leaf patterning roles are genetically separable from the AZ differentiation function of BOP1 and BOP2 genes. Absence of leaf and floral patterning suppressors in the screened populations may indicate that suppressors for these phenotypes do not exist either due to the nature of the pathways involved or a redundancy of potential suppressors. Alternatively, isolation of subtle suppression of leaf and flower phenotypes may require more careful analysis of the M2 population.  4.4.7 Conclusions The isolation and characterisation of these two suppressors has advanced our understanding of floral organ abscission in Arabidopsis in several ways. Firstly, recovery of floral organ abscission suggests that other players besides BOP may be involved in AZ anatomy development. Secondly, stamen AZ activation may be differentially regulated from petals and stamens given that stamen abscission is not recovered in these suppressors. Thirdly, floral organ abscission can occur without proper vascular sealing indicating that the parenchymal cells can abscise independently from corresponding responses in the vasculature. Moreover, the correlative lack of normal AZ cell expansion in rva1 bop1 bop2 implies that proper vascular severance may depend on correct expansion of the parenchymal cells in the AZ. 124  5 ROLES OF BOP1 AND BOP2 IN FLORAL MERISTEM IDENTITY 5.1 INTRODUCTION In response to endogenous and environmental cues, the SAM shifts from vegetative to reproductive or inflorescence identity, a tightly-regulated event known as the floral transition. The first phase of the floral transition, sometimes referred to as the early inflorescence, involves a promotion of inflorescence identity in the SAM by the flowering-time genes FLOWERING TIME (FT). SUPPRESSOR OF CONSTANS1 (Liu et al., 2008)and LEAFY (LFY). During the floral transition, key mediators of inflorescence identity, including SOC1, AGAMOUS-LIKE24 (AGL24), FRUITFULL (FUL) and SHORT VEGETATIVE PHASE (SVP) are strongly expressed in the shoot apex and promote inflorescence shoot behaviour. This is characterised by the formation of cauline leaves, rather than rosette leaves, and these develop secondary axillary inflorescence meristems in their axils. The early inflorescence also exhibits substantial internode elongation between lateral organs, resulting in a primary bolting stem that extends from the compressed rosette (Mandel and Yanofsky, 1995; Hempel et al., 1997; Ferrandiz et al., 2000; Yu et al., 2002; Michaels et al., 2003; Gregis et al., 2006). The second phase or late inflorescence involves another switch in activity by the SAM, this time towards the exclusive production of floral meristems. This shift that requires the downregulation of inflorescence identity genes in the the lateral meristems to promote floral meristem fate rather than that of an inflorescence shoot (Yu et al., 2004; Lin et al., 2004; Liu et al., 2007; 2008; Gregis et al., 2008). The floral meristems themselves give rise to determinate floral organs arranged in concentric whorls of outer sterile organs (sepals and petals) and inner reproductive organs (stamens and carpels). After promotion of the initial floral transition, LFY plays an additional central role in specification of floral-meristem identity, a function shared by APETALA1 (AP1; Blazquez et al., 125  2006). Floral meristem initiation begins with the strong up-regulation of LFY in stage 0 floral primordia in the PZ, prior to the initiation of the floral primordium (Weigel et al., 1992; Blazquez et al., 1997; Hempel et al., 1997). As a consequence of direct activation by LFY, as well as by the FT/FD photoperiod induction pathway, AP1 and its homologue CAULIFLOWER (CAL) are expressed in stage 1 floral primordia which are now slight bulges off the side of the SAM (Bowman et al., 1993; Kempin et al., 1995; Parcy et al., 1998; Wagner et al., 1999; Wigge et al., 2005). AP1 and CAL, acting with related MADS-BOX proteins FUL and LATEMERISTEM IDENTITY1 (Saddic et al., 2006), feed-forward to positively enhance LFY expression in the developing floral meristem (Ferrandiz et al., 2000b; Saddic et al., 2006), ensuring that the conversion from shoot to floral meristem fate in the lateral meristems is sharp and irreversible. Mutants impaired in LFY or AP1 function show defects in floral meristem identity featuring modifications to the inflorescence such as secondary flowers, internode elongation and production of leafy organs. LFY and AP1 have further roles in floral architecture, including the specification of floral organ identity and whorl patterning (Chapter 1, Section 1.4). LFY is expressed throughout the floral primordium and, along with specific cofactors, activates the expression of the floral homeotic genes in spatially restricted patterns as dictated by the ABCE model (Krizek and Fletcher, 2005). LFY promotes the expression of the B class genes, APETALLA3 (Schultz et al., 1991) and PISTALLATA (PI) in whorls 2 and 3 (Jack et al., 1992; Weigel and Meyerowitz, 1993) with the assistance of a co-factor, the F-box protein UNUSUAL FLORAL ORGANS (UFO, Wilkinson and Haughn, 1995; Levin and Meyerowitz, 1995; Lee et al., 1997; Lamb et al., 2002; Chae et al., 2008) while LFY activation of AGAMOUS (AG) expression in the inner two whorls (Busch et al., 1999) is achieved through interaction between LFY and the homeodomain protein, WUSCHEL (WUS; Lenhard et al., 2001; Lohlmann et al., 2001). AP1 functions as an A class 126  gene itself and promotes correct organ identity in the first two whorls of the flower along with the other A class gene, APETALA2 (Irish and Sussex, 1990; Mandel et al., 1992; Bowman et al., 1993; Schultz and Haughn, 1993) and may have a role in activation of AP3 (Ng and Yanofsky, 2002). LFY causes AP1 to be expressed throughout the floral primordium (Parcy et al., 1998; Wagner et al., 1999); however, AG expression, also promoted by LFY, leads to repression of AP1 from the inner reproductive whorls (Gustafson-Brown et al., 1994). In turn, AP1 as well as AP2 are important to repress AG expression in the first two whorls (Bowman et al., 1993) likely via the SEUSS/LEUNIG repressor complex (Drews et al., 1991; Liu and Meyerowitz, 1995; Sidhar et al., 2006). LFY and AP1 are also implicated in repression of floral bract growth in Arabidopsis. In most angiosperms, flowers are subtended by floral bracts, which resemble cauline leaves and are defined as pointed, leafy, stipule-flanked organs which subtend flowers and inflorescences and lack petioles (Irish and Sussex, 1990; Bowman et al., 1993). Arabidopsis, similar to other Brassicaceae, is unusual in developing ebracteate flowers, that is, lacking development of the floral bract (Arber, 1931; Weberling, 1981). However, a localised abaxial region in early stage 3 flowers does express a series of lateral organ specific genes, such as FILAMENTOUS FLOWER (FIL), while down-regulating apical and floral meristem identity genes, such as STM and AP1, and thus this region is interpreted as a so-called ‘cryptic bract’ (Long and Barton, 2000). The floral meristem, then, is derived adaxially from the axil of the cryptic bact in a manner similar to secondary inflorescence origins in the axil of the cauline leaf. Laser ablation studies demonstrated that a floral meristem-derived signal is necessary to actively repress cryptic bract growth during floral development (Nilsson et al., 1998). Mutants impaired in floral meristem identity develop floral bracts, which indicates that specification of floral fate is necessary for the bract suppression signal (Schultz and Haughn, 1991; 1993; Huala and Sussex, 1992; Bowman et 127  al. 1993; Weigel et al., 1992; Long and Barton, 2000). Flowers in the bop1 bop2 mutant frequently form floral bracts, hinting that floral meristem identity may be compromised. In collaboration with Dr. Shelley Hepworth, I explored a possible shared function of BOP1 and BOP2 with LFY and AP1 by analysing floral meristem identity and bract suppression in triple bop1 bop2 ap1 and bop1 bop2 lfy plants. Of the data presented in this chapter, Dr. Hepworth contributed Tables 1 and 2 as well as the following plant photographs: Fig. 5.1C-E, Fig. 5.2 A-L, Fig. 5.3 A-F, Fig. 5.4 A-F. All other data were solely my contribution.  5.2 MATERIALS AND METHODS 5.2.1 Plant materials and growth conditions Plants were grown as described in Section 2.2.1. Triple mutants were generated by crossing. Plants were grown in continuous light at 20°C. The number of cauline leaves on the primary inflorescence was scored as described by (Putterill et al., 1995) and the data are expressed as means ± s.e.m. The ap1-1 allele (Irish and Sussex, 1990) was introgressed into Col-0 through backcrossing. Mutant alleles ap1-12 and ap1-10 (Schultz and Haughn, 1993), lfy-1 and lfy-2 (Schultz and Haughn, 1991) were obtained from the Arabidopsis Biological Resource Center (ABRC). Double and triple mutants were constructed by crossing and confirmed by PCR-based genotyping. Primers are listed in the Appendix. Floral stages were determined according to Smyth et al. (1990). Bracts were defined according to Dinneny et al. (2004).  5.2.2 Scanning electron microscopy Samples were prepared as described in Section 2.2.6. To examine the meristem apex, floral primordia were removed with a dissecting needle after stub mounting.  128  5.3 RESULTS 5.3.1 BOP1 and BOP2 expression in developing floral primordia As shown in Fig 2.7 and 2.8 (Chapter 2), BOP1 and BOP2 are expressed early in floral meristem development. In situ hybridisation indicates that BOP2 mRNA is expressed in stage 0 primordia, temporally coincident with time as LFY, and by stage 2 is strongly expressed in the base of the floral primordia. In later stages of floral development, BOP1 and BOP2 localise to the base of floral organs.  5.3.2 bop1 bop2 mutants show slight defects in floral meristem identity Several phenotypes of bop1 bop2 mutants suggest floral meristem identity is weakly impaired. As previously described, bop1 bop2 flowers frequently develop floral bracts (Table 5.1, Fig 5.1B; Hepworth et al., 2005; Norberg et al., 2005), a phenotype also observed in lfy mutants. All bop1 bop2 floral primordia around late stages 1 and 2 show initiating bracts (Hepworth et al., 2005), 20% of which elaborate into visible bracts (Fig 5.1, Table 5.1). Similar to lfy mutants (Schultz and Haughn, 1991; Weigel et al., 1992), bop1 bop2 plants show more secondary inflorescence branches before the first floral node (Table 5.1) and greater internode elongation (S. Hepworth, pers comm) compared to wild type suggesting that the transition to floral production is delayed. A subtle phenotype of bop1 bop2 plants is the development of an ectopic floral meristem in the axil of the sepal leading to a singly branched flower (Fig 5.1D) similar to the branched flowers seen ap1-1 mutants (Fig 5.1E) and infrequently in lfy mutants (Fig 5.2A; Irish and Sussex, 1990; Schultz and Haughn 1991, 1993; Weigel et al., 1992). These data suggest that BOP genes are involved in specifying floral fate and may function together with LFY and/or AP1.  129  Table 5.1 Quantitative analysis of floral-meristem identity phenotypes in wild type and mutants Genotype No. secondary Flowers with Plants with % branched inflorescences bracts branched flowers on flowers plant Col-0  1.94±0.32 (n=36) 2.83±0.14 (n=36) 5.74±0.23 (n=35) 5.51±0.28 (n=29)  0.0% (n=360)  0.0% (n=24)  0.0%  22.3% (n=235) 4.2% (n=407)  14.9% (n=22)  0.3%  54.5% (n=22)  5.6%  100% (n=281)  100% (n=32)  60.1%  Col-0 bop1 bop2  n.d. n.d.  0.0% (n=12) 25.0% (n=12)  0.0% 1.0%  lfy-1 bop1 bop2 lfy-1 Col-0  n.d. n.d. 1.65±0.11 (n=35) 2.67±0.14 (n=36) 2.04±0.16 (n=26) 2.7±0.11 (n=27) 2.94±0.11 (n=32) 5.4±0.34 (n=17)  0.0% (n=333) 28.6% (n=297) 51.5% (n=94) 100% (n=89) 0.0% (n=525)  100% (n=12) 100% (n=6) 0.0% (n=35)  18.1% 29.2% 0%  21.6% (n=540) 0.0% (n=390)  0.0% (n=36)  0%  100% (n=26)  27.7%  100% (n=35) 100% (n=32)  35.8% 81.0%  100% (n=19)  89.5%  bop1 bop2 lfy-2 bop1 bop2 lfy-2  bop1 bop2 ap1-12 ap1-1 bop1 bop2 ap112 bop1 bop2 ap1-1  0.0% (n=525) 31.7% (n=480) 69.2% (n=240)  *branching could not be accurately scored due to high frequency of nodes without flowers (bract only). Table provided by Dr. Shelley Hepworth  130  Figure 5.1 Floral meristem defects in bop1 bop2 (A, B) Inflorescences of Col-0 (A) and bop1 bop2 plants (B). (A) Col-0. (B) bop1 bop2 plants often show a visible floral bract subtending the flower, arrow. (C-D) siliques of Col-0, bop1 bop2 and ap1-1 plants. (D) bop1 bop2 mutants show flower branching rarely in early flowers (right). (E) ap1-1 flower with two branches. Scale bars, D-E 100 µm. Photos C-E contributed by Dr. Shelley Hepworth.  131  5.3.3 bop1 bop2 lfy-2 and bop1 bop2 lfy-1 triple mutants show enhanced loss of floral meristem identity To examine the relationship between BOP1, BOP2 and LFY regulation of floral meristem identity, triple mutants were generated from crossing the bop1 bop2 double mutants with either the weak lfy-2 or strong lfy-1 mutants. Weak lfy-2 mutants rarely develop bracts (5.74%) and show simple, first-order flower branching on 54.5% of plants (Fig 5.2A; Table 5.1). Triple bop1 bop2 lfy-2 mutant plants show greatly enhanced aberrations of floral architecture and floral meristem identity compared either lfy-2 or bop1 bop2 parent. First, 100% of bop1 bop2 lfy-2 plants developed branched flowers and each plant developed more of them compared to lfy-2 alone (Table 5.1). Second, bop1 bop2 lfy-2 branched flowers formed higher order patterns (Fig 5.2B) due to formation of axillary meristems in the axils of multiple leafy-like organs resembling bracts in the primary flower (Fig 5.2P). Third, increased internode elongation between sepal/ bract like structures (Fig 5.2B,C arrows) also suggested a more severe transformation from floral meristem to inflorescence shoot identity in the triple mutant. In addition, all bop1 bop2 lfy-2 triple flowers developed floral bracts which themselves were greatly enlarged compared to floral  132  Figure 5.2 Loss of floral meristem identity and aberrant floral organ identity of lfy mutants are enhanced by bop1 bop2 (A) Flower from weak lfy-2 mutant has simple branched form. (B) bop1 bop2 lfy-2 flower exhibiting highly branched architecture and internode elongation (arrow). (C) Scanning electron micrograph of bop1 bop2 lfy-2 flower showing internode elongation (arrow) between sepalloid bracts. (D) floral bracts from bop1 bop2 lfy-2 (right) show dramatic enlargement compared with bop1 bop2 (left) and lfy-2 (lfy-2) mutants. (E) Strong lfy-1 mutants inconsistently develop small bracts. (F) bop1 bop2 lfy-1 mutant inflorescence with large bracts subtending every flower (arrows) and with floral organs of increased leaf-like character (G) Terminal bract structures of bop1 bop2 lfy-1 lacking associated floral meristems and with ectopic carpelloid tissue on organ margins. (H-I) flower architecture in wild type and mutants. (H) wild-type flower with typical whorled structure (I) bop1 bop2 flower with abaxial petalloid sepals and extra abaxial stamen. (J) lfy-2 flower with mild reduction in petal and stamen number. (K) triple bop1 bop2 lfy-2 flower exhibiting a loss of petals and formation of grossly deformed stamens (L) lfy-1 flowers develop sepals and carpels instead of petals and stamens. (M-Q) SEMs of wild-type and mutant stage 12 flowers. (M) Col-0 control. (N) bop1 bop2 flower with abaxial petalloid sepals slightly fused (right). (O) lfy-2 flower with reduced numbers of petals and stamens and a leaf-like stellate trichome on the right sepal (arrow). (P) bop1 bop2 lfy-2 flower with ectopic floral meristem production (fm’, fm’’) in the axils of multiple sepal/bracts. Ectopic stigmatic papillae are visible on the tips of all developing sepal/bracts. (Q) Ectopic ovule production (*) on the margins of bop1 bop2 lfy-2 sepal/bracts. (R-W) Inflorescence apices of wild type and mutants plants. (R) bop1 bop2 lfy-2 inflorescence apex with most of floral bracts dissected away reveals early sepal/bract fusion on the floral meristem. (S) side view of a bop1 bop2 lfy-2 inflorescence apex with large bract (b) enfolding a young floral meristem. (T-U) inflorescence meristems (IM) and young floral primordia. Numbers indicate the stage of floral primordia (Symth et al., 1990). (T) Col-0 control inflorescence meristem and young floral buds (U) bop1 bop2 inflorescence meristem and young floral buds showing development of small floral bract (arrows, b). (V) lfy-2 inflorescence meristem and young floral buds showing no floral bract and early development of lateral sepals. (W) bop1 bop2 lfy-2 inflorescence meristem and young floral buds. Young floral meristems are mostly occluded by encasing large floral bracts (arrows, b). Scale bars, IM, inflorescence meristem; b, bract; fm’, fm’’ ectopic floral meristems. Photos A-L contributed by Dr. Shelley Hepworth.  133  134  bracts in bop1 bop2 or lfy-2 mutants (Fig 5D). Moreover, bract outgrowth advanced faster in bop1 bop2 lfy-2 than in bop1 bop2 such that initiating floral primordia in the triple were rapidly covered by the growing bract (Fig5.2S,W compare with U; numbers indicate the stage of floral primordia). The lfy-1 mutants show a much stronger loss of meristem identity than lfy-2: lfy-1 plants more frequently develop small floral bracts and always develop a proportion of branched flowers (Fig. 5.2E, L, Table 5.1). When combined with bop1 bop2, these lfy-1 phenotypes are further enhanced agreeing with published results (Norberg et al., 2005). The triple bop1 bop2 lfy-1 plants show greater development of floral bracts (Fig 5.2F, arrows) and display more branched flowers per plant (Table 5.1). Furthermore, the inflorescence of bop1 bop2 lfy-1 flowers often terminated with a cluster of leaf-like organs (Fig 5.2G) interpreted to be bracts lacking formation of an associated lateral meristem. The more severe loss of floral meristem identity in the bop1 bop2 lfy-1 and bop1 bop2 lfy-2 compared to either set of parents suggests that LFY and both BOP1 and BOP2 make independent contributions to this process.  5.3.4 bop1 bop2 lfy-2 and bop1 bop2 lfy-1 triple mutants enhance lfy floral organ defects As described in Chapter 1 and the introduction to this chapter, LFY contributes to floral organ identity via direct activation of the ABC floral homeotic genes. LFY promotion of B-class gene expression is essential to pattern petals and stamens. Strong loss of function lfy-1 flowers show severely aberrant floral organ identity such as sepalloid conversions of petals and dramatic reductions in stamen number as expected from a loss of B-class function (Fig 5.2L, Table 5.2). In lfy-1 mutants, first whorl organs often show bract-like qualities including stellate (branched) trichomes and first and second whorls demonstrate carpelloid features (Fig 5.2L, Table 5.2), 135  Table 5.2. Quantitative analysis of floral-organ identity phenotypes in wild-type and mutants  *Flowers from nodes 1-15 were scored Data provided by Dr. Shelley Hepworth  136  suggested to reflect ectopic expression of AG (Weigel and Meyerowitz, 1993). In contrast, the weak lfy-2 mutant displays mild floral defects in stamen identity and reduced numbers of petals, although petals which do develop appear normal (Fig. 5.2 J,O, Table 5.2). However, in combination with bop1 bop2, floral patterning of bop1 bop2 lfy-2 was severely aberrant and resembled lfy-1. Floral organ development in the second and third whorls was greatly compromised in bop1 bop2 lfy-2 as indicated by a virtual absence of petals and a large drop in stamen number (Table 5.2). The few organs in the second and third whorls often showed a mosaic of sepalloid and/ or carpelloid features (Table 5.2, Fig 5.2K). First whorl sepal/bract organs of bop1 bop2 lfy-2 displayed enhanced transformation to carpel identity (Table 5.2) as indicated by stigmatic papillae (Fig 5.2P, arrows) and occasional ovules initiating along the organ margins (Fig 5.2Q, arrow). Furthermore, bop1 bop2 lfy-2 first whorl organs showed fusion early in development forming an enclosing tube (Fig 5.2R, asterisks); the organs eventually separate during ougrowth as such fused structures are not observed in mature flowers. In combination with the strong lfy-1 allele, triple bop1 bop2 lfy-1 floral patterning phenotypes resembled lfy-1 in nature and showed an enhancement of carpelloid features in the first two whorl organs compared to the lfy-1 parent (Table 5.2). Overall, triple bop1 bop2 lfy-2 and bop1 bop2 lfy-1 mutant plants developed enhanced lfy-like floral organ phenotypes and no sign of bop1 bop2 floral defects. Given that lfy-1 is a null allele, augmented carpelloid tissue development in the first two whorls of bop1 bop2 lfy-1 triple mutants versus either parent suggests that BOP1 and BOP2 may have a redundant role with LFY in the repression of carpelloid identity in the perianth.  137  Figure 5.3 Loss of floral meristem identity in ap1 mutants is enhanced by bop1 bop2 (A-D) Wild type and mutant flowers. (A) Col-0 flower with concentric whorls of floral organs. (B) bop1 bop2 flowers showing petalloid conversion of sepals. (C) strong ap1-1 allele develops ectopic flowers from the sepal axils. (D) bop1 bop2 ap1-1 flower shows internode elongation between sepal/ bracts in addition to ectopic flowers. (E) older bop1 bop2 ap1-1 flower with indeterminate branching architecture. (F) younger bop1 bop2 ap1-1 shows extensive branching with large bracts subtending each node (arrows). Bracts are often branched. (G-L) Scanning electron micrographs of wild-type and mutant inflorescence apices. (G) Col-0 apex. (H) ap1-1 apex (I) bop1 bop2 apex showing development of the floral bract off a stage 2 primordium (*). (J) bop1 bop1 ap1-1 apex with primordia all developing bracts (*). An ectopic floral meristem (fm’) is first detected in late stage 3 primordia. (K) close up of a bop1 bop2 ap1-1 late stage 3 flower with large, branched leafy bracts with an initiating a secondary floral meristem (fm’) within a bract axil. (L) bop1 bop2 ap1-1 stage 3 flower with spiral phyllotaxic pattern of first whorl sepal/ bract organs. Numbers in G to K indicate floral stages. (M) bop1 bop2 ap1-1 stage 11 flower showing multiple ectopic flowers with basal sepal/bracts. Scale bars, A-F, 1cm; G-J, L 100 µm; K, 50 µm; M, 2mm. IM, inflorescence meristem. FM, floral meristem. s, sepal. Photos A-F provided by Dr. Shelley Hepworth.  138  139  5.3.5 bop1 bop2 ap1-1 and bop1 bop2 ap1-12 show enhanced loss of floral meristem identity Another key mediator of floral meristem identity is AP1, which is directly activated by LFY in stage 1 floral primordia (Mandel et al., 1992; Mandel and Yanofsky, 1995; Wagner et al., 1999). The strong loss of function ap1-1 mutant is characterised by a loss of petals and the development of secondary flowers from ectopic floral meristems arising the axil of first whorl bract-like sepals (Irish and Sussex, 1990; Fig. 5.3C). Results in section 5.3.3 and 5.3.4 suggest that BOP1 and BOP2 work in parallel with LFY to ensure determinate floral fate of the floral meristem as well as proper floral organ identity. To determine if BOP1, BOP2 and LFY activities converge to regulate AP1, genetic interactions between bop1 bop2 and ap1 mutants were assessed. Triple bop1 bop2 ap1-1 mutant flowers exhibit severely impaired floral meristem identity compared to either parent. Firstly, the triple flowers produce several ectopic flowers which themselves iterate ectopic flowers resulting in extensive multi-order branches (Fig 5.3D-F,M). The production of ectopic flowers is accompanied by significant internode elongation between developing sepals leading to large branched structures (Fig 5.3D, arrow). First whorl organs of bop1 bop2 ap1-1 develop more like bracts than sepals to form large, leafy, deeply lobed/branched structures (Fig 5.3F,K). As the pedicel of the ectopic flower elongates, the large bract remains at its base and often becomes branched itself (Fig. 5.3F, arrows). Secondly, the bracts/sepals do not arise in a whorled pattern but rather develop in a spiral phyllotaxy off the floral meristem (Fig 5.3K,L,M), similar to the pattern seen for leaves and floral meristem primordia off the SAM. Thirdly, bracts arise from early stage 2 primordia in bop1 bop2 ap1-1 (Fig 5.3L) rather than late stage 2/3 as seen for bop1 bop2 (Fig 5.3I) suggesting that bract formation is initiated earlier in the triple mutant. Sepals develop to look very much like bracts and develop ectopic floral merstems in 140  their axils (Fig 5.3J,K,M). The branched flower structure normally does not terminate in a carpelloid organ except for some late-forming flowers. Triples between bop1 bop2 and the weak ap1-12 allele show similar defects in branching (Table 5.1). Taken together, these data suggest that the bop1 bop2 ap1 triple mutant is compromised in floral meristem identity to greater extent than in either parent, suggesting that BOP1 and BOP2 are involved along with AP1 in promoting floral meristem identity in lateral meristems. In particular, BOP1 and BOP2 and AP1 genes play important roles in suppressing architectures normally associated with inflorescences shoots: bract outgrowth, internode elongation and spiral phyllotaxy.  5.3.6 bop1 bop2 ap1-1 and bop1 bop2 ap1-12 triple mutants enhance ap1 floral organ defects In addition to roles in floral meristem identity, AP1 has class A gene function and thus directs proper morphogeneis of sepals and petals in the first and second floral whorls, respectively. Strong ap1-1 mutants develop secondary flowers in the first whorl and show a loss of sepal and petal identity (Fig 5.4E, Table 5.2) while weak ap1-12 show moderate loss of petals only (Fig 5.4B, Table 5.2). Floral organ phenotypes of triple bop1 bop2 ap1-1 plants reflect enhanced ap11 features characterised by an increased carpelloid character on sepal/bracts (Fig. 5.3F,J,K). This is manifested, in particular, by development of ectopic ovules along sepal margins (Fig 5.3F,K) suggesting that BOP1 and BOP2 have a parallel role with AP1 in suppression of carpel identity in first whorl organs. Similar to combinations with the lfy-2 weak allele, triple bop1 bop2 ap1-12 show phenotypes similar to the strong ap1-1 allele (Fig. 5.3C) showing secondary flower development and a loss of petals. Overall, as is the case for lfy, bop1 bop2 ap1 flowers exhibit 141  Figure 5.4 bop1 bop2 exacerbates floral organ defects in ap1 mutants (A-F) wild-type and mutant flowers. (A) wild-type (Col-0) flower. (B) weak ap1-12 flower develops some petals and is determinate (C) bop1 bop2 ap1-12 flower with secondary flowers in the axil of first whorl sepal/bracts and a loss of most petals (D) bop1 bop2 flower with abaxial petalloid sepals. (E) strong ap1-1 flower forms secondary flowers in the axil of first-whorl sepal/bracts and lacks petals (F) bop1 bop2 ap1-1 flower with rare determinate architecture. (GK) Scanning electrion micrographs of young wild type and mutant flowers. (G) Col-0 flower stage 10/11 (H) bop1 bop2 flower stage 9/10 (I) ap1-1 flower stage 11; note lateral sepal/bracts with stipules and loss of petals. (J) older bop1 bop2 ap1-1 mutant flower with ectopic floral meristems and carpelloid features on first whorl organs. (K) Young stage 8 secondary flower of bop1 bop2 ap1-1 showing ectopic ovules (ov) and papillae (p) along sepal/bract margins. Photos A- F provided by Dr. Shelley Hepworth.  142  143  enhancement of ap1 phenotypes and do not show features associated with bop1 bop2 floral patterning defects.  5.3.7 AGL24, SOC1 and FUL mis-expression in bop1 bop2 ap1 and bop1 bop2 lfy Misexpression of the inflorescence identity factors, AGL24, SOC1 and FUL has been shown to affect the transition to floral fate (Yu et al., 2004; Liu et al., 2007; Gregis et al., 2008). Given the loss of floral meristem identity and exhibition of inflorescence identity characteristics observed in bop1 bop2 lfy and bop1 bop2 ap1, expression of the inflorescence regulators, AGL24, SOC1 and FUL was assessed. These results are contributed by Dr. Shelley Hepworth’s laboratory. AGL24 and SOC1 are normally expressed in inflorescence apical meristems and in young floral primordia; however, they are all repressed in the central dome of late stage 2 flowers and are restricted to the cryptic bract (Samach et al., 2000; Michaels et al., 2002). In bop1 bop2 lfy-2 and bop1 bop2 ap1-1, this expression persists in floral primordia through stage 2 and 3, a greatly expanded expression domain compared with either parent (data not shown). Another regulator of inflorescence identity is FUL, also expressed in the inflorescence apex until late stage 2 where it becomes repressed to the cryptic bract by stage 3; however, FUL plays an important role in valve development and thus its expression reappears in the central zone of the floral primordia (Mandel and Yanovsky, 1995; Hempel et al., 1997). In bop1 bop2, ap1-1 and lfy-2 parents, FUL was misexpressed in stage 2 primordia and showed an enlarged expression domain in stage 3 flowers; these effects were greatly enhanced in the bop1 bop2 lfy-2 and bop1 bop2 ap1-1 triple mutant flowers (data not shown). Thus, BOP1 and BOP2 genes play important roles along with LFY and AP1 in repressing the expression of inflorescence identity genes in emerging floral primordia.  144  5.4 DISCUSSION In Arabidopsis, the floral transition is marked by bolting of a primary inflorescence stem which first initiates several cauline leaves with associated axillary inflorescence shoots then switches to produce single, ebracteate flowers. Multiple regulators, including LFY and AP1, ensure that the transition from vegetative to inflorescence to floral meristem production is sharp and robust. LFY and AP1 are central regulators of floral meristem identity: loss of function in either leads to defects in floral meristem identity suggestive of shoots. lfy mutants show delayed flowering, increased secondary inflorescence production and floral bract development whereas ap1 mutants are characterised instead by single ectopic flower meristem development in the axil of the first whorl organs (Bowman et al., 1993). Outgrowth of floral bracts, increased secondary branching and appearance of branched flowers in bop1 bop2 suggested that BOP1 and BOP2 may also be involved in regulating the transition from shoot identity to floral meristem identity. Genetic analyses suggests that BOP1 and BOP2 share roles with LFY and AP1 in promotion of floral organ identity. Triple mutant bop1 bop2 lfy-1 and bop1 bop2 ap1-1 showed enhanced loss of floral meristem identity compared with either parent suggesting that BOP1 and BOP2 work in parallel with both AP1 and LFY in controlling the transition to flowering.  5.4.1 BOP1 and BOP2 act in parallel with LFY to specifically activate AP1 Synergism between the BOP loci and LFY in repression of bract formation may indicate that they share targets involved in floral patterning. Like LFY, BOP1 and BOP2 are expressed in stage 0 floral primordia and may act with LFY to promote expression of its early target genes in stage 1 flowers. AP1, LMI and CAL are known targets of LFY (Wagner et al., 1999; Williams et al., 2004; Ferrandiz et al., 2000; Saddic et al., 2006). AP1 is normally expressed in stage 1 floral 145  primordia (indicating commitment to floral fate) but in lfy-2 mutants its expression is delayed although it reaches wild type levels by stage 3 (Bowman et al., 1993; Mandel and Yanovsky, 1995; Ruiz-Garcia et al., 1997; Liljegren et al., 1999). In situ hybridisation of wild type and mutant plants has shown that the expression of AP1 is relatively normal in bop1 bop2, appearing around stage 1, but greatly reduced and delayed in bop1 bop2 lfy-2 triple mutants relative to bop1 bop2 or lfy-2 parents (S. Hepworth pers comm) suggesting that under conditions of weak LFY function, BOP1 and BOP2 are critical for AP1 expression. Exacerbation of the loss of floral meristem identity in bop1 bop2 lfy-2 mutants compared to lfy-2 likely reflects this loss of AP1 expression. Genetic and in situ analyses show that BOP1 and BOP2 do not contribute to LMI or CAL activation (S. Hepworth, pers comm.); thus, the interaction between the BOP genes and early LFY targets is specific to AP1. Although LFY expression appears normal in bop1 bop2 mutants inflorescence meristems (S. Hepworth, pers. comm), LFY expression in bop1 bop2 has not been examined in the SAM during the first transition phase to ascertain whether BOP contributes to initial LFY activation. LFY activation is mediated by FT/FD through the activity of the BELL-LIKE HOMEODOMAIN proteins PENNYWISE (PNY) and POUNDFOOLISH (PNF) (Kanrar et al., 2006; 2008) as well as by other long-day induced transcription factors, such as SOC1 together with AGL24, and by giberellin-controlled pathways (Mouradov et al., 2002; Blazquez et al., 2007; Lee et al., 2007; Kobayashi and Weigel, 2007; Liu et al., 2008). It is unknown whether BOP1 and BOP2 contribute to LFY activation via any one of these mechanisms.  146  5.4.2 Regulation of inflorescence identity gene expression by BOP1 and BOP2 In addition to acting along with LFY to activate AP1 transcription, BOP1 and BOP2 also contribute in parallel with AP1 and LFY to repress the development of ectopic floral meristems in the axils of sepals as bop1 bop2 ap1-12 and bop1 bop2 lfy-2 mutants demonstrate greatly enhanced formation of highly branched flowers. ap1-1 cal double mutants are characterised by the proliferation of higher order inflorescence meristems instead of floral meristems off the SAM (Bowman et al., 1993; Ferrandiz et al., 2000) similar to the highly branching phenotype described here for bop1 bop2 ap1-1. Thus, bop1 bop2 enhances this aspect of the ap1-1 phenotype similar to ap1 combinations with cal, suggesting that BOP functions redundantly with AP1 and CAL following specification of floral fate. In ap1-1 mutants, the floral inflorescence identity factors AGL24, SOC1 and SVP are increased in the sepal axils correlating with secondary flower production; introduction of AGL24, SOC1 and SVP loss of function alleles can each rescue this defect (Liu et al., 2007). The expression domain of AGL24 and SOC1 was enlarged even further in bop1 bop2 ap1-1 as well as bop1 bop2 lfy -2 mutants suggesting that the highly branched flowers typical of the triple mutants reflect this misexpression. In fact, flower branching was recovered by introduction of the agl24 allele into bop1 bop2 ap1-1 and bop1 bop2 lfy-2 (S. Hepworth pers comm). Furthermore, these results show that BOP1 and BOP2 must also contribute to the repression of AGL24 and SOC1 in a pathway redundant with AP1 and LFY. AP1 directly represses the expression of AGL24 and SOC1 (Liu et al., 2007). LFY is presumed to partake in suppression of AGL24 indirectly, a role that may be mediated through AP1 (Yu et al., 2004; Liu et al, 2007). Expression of FUL is also enhanced in bop1 bop2 ap1-1 and bop1 bop2 lfy-2 mutants. Similar to SOC1, FUL expression in the rosette and cauline leaves is promoted by FT/FD (Teper147  Bamnolker and Samach, 2005) and was recently shown to be critical to prevent vegetative reversion following transition to flowering (Melzer et al., 2008) supporting the notion that FUL is an inflorescence identity gene. Similar to AGL24 and SOC1, FUL is downregulated by AP1 following during floral meristem specification (Mandel and Yanovsky, 1995). However, unlike AGL24 and SOC1, FUL loss of function mutants do not rescue ap1 branching phenotypes and in fact exacerbate floral meristem defects of ap1 cal mutants by increasing the vegetative characteristics of the lateral organs (Ferrandiz et al., 2000b) suggesting that it plays a secondary role in promotion of floral meristem development. LFY and AP1 are both proposed to antagonise TFL1 expression in the developing floral primordia and restrict TFL1 expression to the SAM where it promotes indeterminancy (Liljegren et al., 1999). TFL1 was shown to be ectopically expressed in lateral meristems of ap1 cal double and ap1 cal ful mutants and significantly, introduction of the tfl1 allele into ap1 cal completely repressed the cauliflower phenotype. This suggests that the conversion of inflorescence to floral meristems regulated by AP1 and CAL may be mediated in large part through TFL1 repression (Ferrandiz et al., 2000b). It is unknown whether ectopic TFL1 may play a similar role in the enhanced branching phenotypes observed in bop1 bop2 lfy and bop1 bop2 ap1 triple mutants.  5.4.3 Other genetic players in bract formation and phyllotaxy of floral organs In Arabidopsis, spiral phyllotaxy of initiated lateral structures, such as leaves and axillary shoots, is a feature of inflorescence shoot identity while whorled phyllotaxy of the floral organs is typical of the determinant floral meristem. Impaired floral meristem identity often is reflected in altered phyllotactic patterns of floral lateral structures. A shift from whorled to spiral phyllotaxy is observed in ap1-1 ap2-1 (Irish and Sussex, 1990) and ap1-1 lfy (Huala and Sussex, 1992) 148  double mutants while ap1 and lfy single mutants show intermediate, uneven arrangements (Irish and Sussex, 1990; Weigel et al., 1992). Spiral phyllotaxy is also seen in triple bop1 bop2 ap1-1 flowers in association with an increased development of sepal/bracts. This suggests that BOP1 and BOP2 contribute to floral meristem identity in part by promotion of a whorled pattern of floral organs in parallel with AP1. Similar to that postulated for BOP1 and BOP2, FILAMENTOUS FLOWER (FIL), a YABBY gene involved in lateral organ polarity also has a role in floral meristem identity, floral phyllotaxis and bract suppression in parallel with LFY and AP1 (Sawa et al., 1999; Goldschmidt et al., 2008). These data highlight a link between the repression of bract development and a shift towards whorled phyllotaxy. It is unknown if BOP1 and BOP2 regulation of bract development and/ or phyllotaxis involves FIL, although Ha et al (2007) showed that FIL is misexpressed in bop1 bop2 leaves. Given that FIL becomes restricted to the cryptic bract when floral organ development begins at stage 3, it may regulate floral organ phyllotaxy in a non-cell autonomous fashion from the bract to the meristem. In fact, Goldschmidt et al (2008) have proposed that LATERAL SUPPRESSOR (LAS), a GRAS transcription factor, contributes to signalling from the bract to the SAM and the FM. LAS is essential for axillary meristem development (Greb et al., 2003) underscoring a relationship between bract development, meristem identity and phyllotaxy. Noncell autonomous signalling likely occurs in both directions as AP1, UFO (see below), LFY, and BOP genes are not expressed in the cryptic bract yet mutations in these genes affect bract production. Similar to bop1 bop2 and fil, mutations in UNUSUAL FLORAL ORGANS (UFO) lead to loss of second and third whorl organ identity and infrequent production of floral bracts (Levin and Meyerowitz, 1995; Wilkinson and Haughn, 1995). Similar to bop1 bop2, combination of ufo 149  with lfy or ap1 alleles lead to an enhancements of the latter's floral meristem identity defects. UFO encodes an F-box protein known to participate in the SCF ubquitin ligase protein degradation pathway (Samach et al., 1999) and acts as a cofactor with LFY to directly activate AP3 expression in the second and third whorls (Ingram et al. 1995; Jack et al. 1992; Lee et al. 1997; Parcy et al. 1998). Chae et al (2008) demonstrated that UFO association with LFY lead to increased proteasome-dependent turnover of the LFY protein postulated to be important to fully potentiate LFY activity (Chae et al., 2008). BOP1 and BOP2 proteins contain a BTB/POZ domain which has been shown to interact with Cullin-3 type proteosome complexes (Pintard et al. 2004; van den Heuvel, 2004; Wang et al. 2004; Dieterle et al. 2005; Figueroa et al. 2005; Gingerich et al. 2005; Weber et al. 2005). Thus, BOP1 and BOP2 may mediate their potentiation of LFY functions through regulated protein degradation similar to UFO.  5.4.4 Floral organ identity and floral organ number Overall bop1 bop2 ap1 mutants display enhanced ap1 phenotypes including enhanced carpelloidy in first whorl organs. Similar enhancement of carpelloid features was observed in bop1 bop2 lfy-1 compared to parents which also suggests that BOP1 and BOP2 work in parallel to LFY to suppress reproductive identity in the first whorl organs. Expression of carpelloid features does not necessarily require AG expression but it remains possible that ectopic AG expression in the bop1 bop2 ap1-1 promotes carpelloid identity. LFY acts to suppress AG expression in the first whorls via positive regulation of B-class genes AP3 and PI. Liu et al (2009) recently reported that expression of SEP3, encoding a MADS-Box partner participating in the AP3/PI quartet and important for initiating and maintaining AP3 and PI expression, is directly repressed by AGL24 and SOC1. Thus, overexpression of AGL24 and SOC1 in bop1 150  bop2 ap1 and bop1 bop2 lfy floral meristems is predicted to cause downregulation of SEP3. Reduced and/or delayed SEP3 levels may impair class B function in the triple mutants leading to reduced petal and stamen identity thus expression of carpelloid features in floral organs. Organ number is drastically reduced in whorl 2 of bop1 bop2 lfy-2 and whorl 3 of bop1 bop2 lfy-2/lfy-1. Furthermore, the reduction the number of organs developing in whorl 2 of bop1 bop2 ap1-12 flowers approximates reductions seen in the strong ap1-1 mutants and ap1-1 cal mutants (this study, Alvarez-Buylla et al., 2006). These phenotypes suggest that bop1 bop2 specifically enhances reductions in floral stem cells available for recruitment into floral organs in whorls 2 and 3, regions patterned by the B-class genes. One potential explanation for the reduction in floral whorl number could be the extensive growth of floral bracts seen in these mutants which may deplete the floral meristem stem cell pool available for the later developing second and third whorl organs.  5.4.5 Roles of BOP1 and BOP2 in floral meristem identity Triple mutant phenotype analysis supports a strong role for BOP1 and BOP2 in AP1 and LFYregulated pathways to suppress inflorescence identity in lateral meristems, to promote their development into whorled, determinate floral meristems, and to further promote of proper floral organ identity in lateral meristems. LFY, AP1 and BOP1 and BOP2 promotion of floral meristem identity is likely mediated in large part by the down-regulation of inflorescence identity genes such as AGL24, SOC1 and FUL in stage 2/3 floral primordia. Although BOP1 and BOP2 function in parallel with LFY and AP1 in determination of floral meristem identity, their role based on bop1 bop2 mutants is primarily one of patterning floral architecture rather through promotion of the floral transition. Ectopic overexpression of 151  either LFY or AP1 in the SAM results in early flowering and the production of solitary flowers instead of an inflorescence due to repression of TERMINAL FLOWER1 (TFL1) expression (Mandel and Yanovsky, 1995; Weigel and Nilsson, 1995; Lijegren et al., 1999) while overexpression of BOP1 and BOP2 leads to dwarfed, epinastic architectures (Norberg et al., 2005; Ha et al., 2007) suggesting that they are not sufficient to repress shoot identity and promote floral identity in Arabidopsis as is the case for LFY and AP1. Thus BOP1 and BOP2 can be construed more as potentiators or enhancers of LFY and AP1 function whose contribution becomes critical under conditions of weak loss of function alleles in LFY or AP1.  152  6 PHYLOGENETIC ANALYSIS OF THE NPR1-LIKE PROTEIN FAMILY AND BIOCHEMICAL MECHANISM OF BOP1/BOP2 6.1 INTRODUCTION Chapters 2 through 5 describe a variety of roles for the BOP1 and BOP2 genes in lateral organ development. The biochemical mechanisms employed by BOP1 and BOP2 proteins to mediate floral and leaf architecture are not yet understood; however, it is likely that they share modes of action with the NON-EXPRESSOR OF PR-1 (NPR1) protein, a key mediator of plant defense responses. BOP1 and BOP2 form a distinct pair within the six member NPR1-like protein family in Arabidopsis (Hepworth et al., 1995). Sequence similarity suggests that NPR1 and its closest homologue, NPR2, also form a separate subgroup as do NPR3 and NPR4 (Hepworth et al., 2005; Figure 6.1). NPR1 was characterized as a crucial mediator of systemic acquired resistance (SAR), a plant immune response induced following a local pathogen infection (Cao et al. 1994, 1997; Delaney et al., 1994; Ryals et al., 1997; Glazebrook et al., 1999; reviewed in Dong, 2004). To ward off subsequent pathogen attack, plants upregulate a suite of PATHOGENESIS-RELATED (PR) genes throughout the plant. Loss of function npr1 mutants are characterised by a lack of elevated PR gene expression following pathogen attack, which leads to enhanced disease susceptibility. NPR1 mediates activation of PR gene expression through interaction with TGACG sequence-specific binding (TGA) transcription factors. This interaction and associated upregulation of PR genes is dependent upon the nuclear localisation of NPR1 (Kinkema et al., 2000; Mou et al., 2003). Seminal work from the Dong (Mou et al., 2003) and Folbert (Depres et al., 2005) laboratories has shed considerable light on the protein activation mechanisms of NPR1. The 153  current working model for NPR1 activation posits that under non-inductive conditions (no pathogen attack), NPR1 forms oligomeric complexes in the cytoplasm with other NPR1 proteins. The oligomeric complex is stabilised through via disulphide bridges formed between conserved cysteine residues (Cys82 and Cys216) amongst adjacent NPR1 polypeptides. In response to pathogen attack, a localised hypersensitive response ensues, prompting the systemic accumulation of salicylic acid (SA). Accumulation of SA shifts the redox balance of the cell such that the disulphide bridges sequestering NPR1 are reduced to thiol groups, thus releasing NPR1 proteins as monomers which are now free to translocate to the nucleus. Once in the nucleus, the reduced form of NPR1 positively interacts with the TGACG sequence-specific binding transcription factors (TGAs) to activate transcription of PR genes (Dong, 2004). Recent data suggest that the redox-regulation of NPR1 and its role in the cytoplasm is more complex. For example, the oxidation rather than reduction of C-terminal Cys521 and Cys529 are required for NPR1 interaction with and transactivation of TGA2 (Rochon et al., 2006). Although NPR1 promotes TGA2 to elicit PR gene expression, NPR1 associates more strongly with non-DNA bound TGA2 (Johnson et al., 2008). These authors propose that NPR1 may act to assemble TGA2 dimers at their cognate DNA binding site (Johnson et al., 2008). The transcriptional activation ability of TGA2 itself also appears to be modulated by SAR, since under non-induced conditions, TGA2 negatively regulates PR gene expression (Kerwasami et al., 2007). In addition to promoting NPR1 monomer release, Tada et al. (2008) reported that SA treatment also enhanced oligomerisation of NPR1 through N-nitrosylation of Cys156. This dual role of SA functions to maintain NPR1 levels as oligomerisation appears essential to protect nascent NPR1 from degradation, thus allowing build up of more polypeptide available for monomer realease during SAR (Tada et al., 2008). These data are consistent with the vastly 154  decreased levels of NPR1 observed in plants expressing a mutated C156Y version of NPR1 (Mou et al., 2003). In addition, the role of oligomerisation in NPR1 protein homeostasis may explain why cytoplasmic localisation of NPR1 is essential for SAR following mycorrhizal infection (Stein et al., 2008). Several aspects of the BOP1/2 signalling mechanism are conserved with NPR1, including interaction with TGA7, PERIANTHIA, which acts in the same genetic pathway to control perianth patterning in flowers (Hepworth et al., 2005). It is unknown whether BOP1/2 activity is controlled by redox-sensitive nuclear localization and TGA interaction as is the case for NPR1. To begin to address the possible shared signalling mechanism between the BOP proteins and NPR1, I employed bimolecular fluorescence complementation (Citovsky et al., 2006) to track subcellular localisation and interaction of BOP proteins and partners in planta. BiFC detects subcellular localisation of protein-protein interactions based on reconstituted fluorescence. The N-terminal and C-terminal halves of a fluorescent protein are fused to putatively interacting proteins and when these proteins interact, the halves of the fluorescent protein are brought into close enough proximity to fluoresce when irradiated under the appropriate absorbed wavelength (Hu et al., 2005; Citovsky et al., 2006). Here, I used split Enhanced Yellow Fluorescent Protein (EYFP) fused to BOP2 and PAN proteins to detect interaction in planta.  6.2 MATERIALS AND METHODS 6.2.1 Plant materials and growth conditions Plants were grown as described in Section 2.2.1.  155  6.2.2 Phylogenetic Analysis Amino acid sequence alignments for Arabidopsis NPR1-like protein family and the BPLs were aligned using the MAFFT (Multiple Alignment using Fast Fourier Transform) tool available online (http://www.ebi.ac.uk/Tools/mafft/index.html) with the BOLSUM62 matrix. The alignment residues were colour-coded based on identity and conservation using the AMAS server (http://www.compbio.dundee.ac.uk/www-amas/). For phylogenetic analysis, the complete BOP1 amino acid sequence was used as a query sequence to assemble homologous proteins from available sequences of land plants. The databases mined include BLASTp (NCBI), the JGI Populus trichocarpa database (http://genome.jgi-psf.org/Poptr1_1/Poptr1_1.info.html), the Medicago trunculata blast genome (http://medicago.org/genome/blast.php), the Maize Genome Database (http://www.maizegdb.org/), and the Physcomitrella patens resource (http://www.cosmoss.org/bm/BLAST?type=1). Proteins were considered members of the NPR1like protein family based on the presence of a N-terminal BTB/POZ domain and a C-terminal ANK repeat domain. Only sequences with a start codon and those supported by expression data were used in the analysis. A multiple protein alignment was constructed using the MAFFT tool with the BLOSUM80 matrix. Based on this alignment, a Maximum Likelihood (ML) analysis using RaXML web-server was executed (Stamatakis et al., 2008). The ML output was fed into TreeView X (http://darwin.zoology.gla.ac.uk/%7Erpage/treeviewx/) to construct an unrooted radial tree which was assembled in TreeIllustrator (http://nexus.ugent.be/geert/).  6.2.3 Generation of pBOP1::BOP2:EYFP transgenic plants The 2X35S promoter of the pSAT6A vector (provided by Stanton Gelvin, Purdue University) was excised by AgeI/ BglII digestion and replaced with the BOP1 4kb promoter fragment 156  described above forming the pBOP1:ENHANCED YELLOW FLUORESCENT PROTIEN (EYFP, Invitrogen, CA) plasmid. Using directional KpnI/SmaI digestion, BOP2 cDNA was cloned into the pBOP1:EYFP plasmid to generate the pBOP1::BOP2-EYFP plasmid. BOP2 cDNA was also cloned into the original pSAT6A vector using KpnI non-directional cloning. Both 35S::BOP2EYFP and pBOP1::BOP2-EYFP plasmids were sequenced to confirm that the fusion was inframe and free from PCR-induced mistakes. The pBOP1:BOP2::EYFP and 35S::BOP2-EYFP constructs were excised and ligated using the homing endonuclease PI-PspI into the 3519 binary vector and transformed into the Agrobacterium strain C58C1 pGV3101 pMP90 (Koncz and Schell, 1986). This construct was transformed into bop1 bop2 by floral dip (Clough and Bent, 1998) and Basta®-resistant transformants selected on soil by spray treatment of seedlings with the herbicide Final ev 150 (AgrEvo). Fluorescence was visualised using epiflourescent microscope (Leica Imagaing).  6.2.4 Bimolecular Florescence Complementation  Bimolecular florescence complementation constructs were cloned using the pSAT modular vectors modified for BiFC, gifts from Dr. Stanton Gelvin at Purdue University (Citovsky et al., 2006). The pSAT6A vector contained full-length EYFP (Invitrogen, CA), pSAT1A contained the C-terminal half of EYFP (residues 1-174) while pSAT4A contained the N-terminal end of EYFP (175-end). Complete BOP2 and PAN cDNAs were cloned into each of these three vectors using KpnI-digestion of cDNA PCR products generated with PwoI proof-reading polymerase (Roche, MD). Primers used for cloning are listed in the Appendix. All constructs were sequenced to confirm translational fusion and that they were error-free. Protoplasts were transformed according to Sang-Dong et al. (2007) using 20 µg of plasmid DNA for single vector controls and 157  10 µg of each plasmid for bimolecular fluorescent complementation cotransformation. Protoplasts were allowed to rest for 12-20 hours before visualisation by epifluorescent microscopy.  6.3 RESULTS 6.3.1 Phylogenetic analysis of the NPR1-like protein family Using a combination of BLASTp database searches, a collection of homologous proteins were assembled. Arabidopsis, Populus trichocarpa (poplar), Zea mays (maize), Oryza japonica (rice) and Physcomitrella patens (moss) sequences retrieved from GenBank were cross-referenced to their available database which was also searched de novo. Unfortunately, insufficient gymnosperm data was available to include in this analysis. Only full-length cDNAs encoding polypeptides containing both N-terminal BTB/POZ and C-terminal ANK oriented domains which also had expression support were selected for alignment. The BLOSUM80 protein alignment matrix, considered the best option for gene family analysis (Wheeler, 2003) was used to construct a multiple protein alignment with MAFFT (Multiple Alignment using Fast Fourier Transform). Maximum Likelihood phylogenetic analysis (Stamatakis, 2006) was performed on the alignment and the data assembled into an unrooted, radial tree presented in Figure 6.1. Four NPR1-like clusters were resolved. Cluster I separates out with maximal support (Bootstrap 100) and consists of three Physcomitrella NPR1-like homologues, presumably representing the most ancestral form of the family. Clusters II – IV contain only angiosperm sequences. Each cluster has dicot and monocot representatives. BOP1 and BOP2 are part of Cluster II, a group distinction strongly supported between Clusters I and Clusters III/IV. Interestingly, Cluster II is 158  Figure 6.1 Phylogenetic Analysis of the NPR1-like protein family Land plants sequences are labelled as moss, monocot, dicot, dicot, Brassicaceae or dicot, Solanaceae. Sequences were aligned by MAFFT using the BLOSUM80 substitution matrix and Maximum Likelihood inferred by RAxML. Four clusters were resolved. Cluster I shows homologues from Physcomitrella patens. Cluster II contains the Arabidopsis BOP1 and BOP2 proteins, while Cluster III resolves NPR1 and NPR2 Arabidopsis homologues and Cluster IV, Arabidopsis NPR3 and NPR4. Monocots and various dicot families form distinct subgroups in Clusters II, III and IV. Species names: Bn, Brassica napus; Bj, Brassica juncea; Bv, Beta vulgaris; Ca, Capsicum annuum; Gh, Gossypium hirsutum; Ha, Helianthus annuus; Hv, Hordeum vulgare subsp. Vulgare; Le, Lycopersicon esculentum; Mt, Medicago trunculata; Mu, Musa acuminate; Nt, Nicotania tabacum; Os, Oryza sativa; Php, Pyscomitrella patens; Pt, Populus trichocarpa; Pyp, Pyrus pyrifolia; Vv, Vitis vinifera; Zn, Zea mays. Arabidopsis family members do not have species identifiers but are shaded in pink.  159  more similar to Cluster I than either Cluster III or IV, suggesting that the BOP proteins members may represent the initial form of the NPR1-like proteins in angiosperms. Support for the distinction between Clusters III and IV is rather weak (Bootstrap 58). BOP homologues exist in both monocots and dicots, which I call BOP-Like (BPL) proteins, in Medicago trunculata, Vitis vinifera, Populus trichocarpa, Oryza satica and Zea mays (Figure 6.1; Table 6.1). Expression data from O. sativa BPLs indicate they are expressed in lateral organs and meristems. The six known NPR1-like proteins from Arabidopsis were examined for amino acid conservation (Fig. 6.2). The alignment presented highlights conserved cysteine residues conserved amongst all family members: Cys82, Cys151, Cys156, Cys233, Cys394 (residue numbers refer to the position in NPR1, Fig 6.2, diamonds) Notably, of the two cysteine residues, each known to be critical for cytoplasmic retention of NPR1, Cys82 and Cys216 (Mou et al., 2003), only C82 is conserved among BOP1 and BOP2 or NPR3 and NPR4 subgroups. In addition, Cys521 and Cys529, whose oxidation is essential for the transactivation of TGA2 by NPR1, are absent in other family members. In contrast, all NPR1-like proteins retain Cys156, a target of Nnitrosylation in NPR1 (Tada et al., 2008). Furthermore, the C-terminal bipartite nuclear localisation sequence crucial for NPR1 nuclear localisation (Nigg et al., 1997; Kinkema et al., 2000) is present in NPR1-4 proteins but is lacking in BOP1 and BOP2. Rather, the C-terminus of the BOP proteins is characterised by a repeated string of histidines at the C-terminus. This feature is absent in the other Arabidopsis NPR1-like family members but is present in the dicot BPL proteins (data not shown) identified in the phylogenetic analysis described above.  160  Table 6.1 BOP protein homologues in land plants Species Medicago truncatula Vitis vinifera Populus trichocarpa Populus trichocarpa Oryza sativa Oryza sativa Oryza sativa Zea mays  Locus ID/ BPL ABD28327.1 MtBPL1 XM_002275944 VvBPL1 eugene3.00160344 PtBPL1 eugene3.00060387 PtBPL2 Os11g04600.1 OsBPL3 Os12g04410 OsBPL2 Os01g0948900 OsBPL1 GRMZM2G096819_P01 ZmBPL1  Expression data EST EST EST EST panicle, shoot, panicle vegetative meristem, leaf, panicle, root EST  161  Figure 6.2 Amino acid alignment of NPR1-like proteins in Arabidopsis Complete polypeptide sequences from all six members of the Arabidopsis NPR1-like protein family were aligned by MAFFT using the BLOSUM62 substitution matrix. Identical residues among all family members are highlighted in red, residues identical in either NPR1-4 or BOP1/BOP2 are shaded in blue while green shading denotes highly conserved residues based on the substitution matrix. Black diamonds show cysteine residues conserved across all family members. Gold dots and gold-shaded boxes label Cys216,Cys519 and Cys529 of NPR1 that are not conserved in other family members. Black underlining bars show the BTB/POZ and ANK repeat domains as labelled. Purple-shading indicates the bipartite nuclear localisation signal (NLS) of NPR1-4 proteins. Peachshaded sequences highlight the histidine–rich (HIS) C-termini of BOP1 and BOP2.  162  163  6.3.2 Complementation of pBOP1:BOP2::EYFP plants In order to determine whether the BOP2-EYFP fusion protein was functional in Arabidopsis, T1 transformants transformed with pBOP1:BOP2::EYFP were assayed by eye for complementation of the bop1 bop2 phenotype (Table 6.2). For leaves, lack of blade on petiole growth was scored as ‘3’ and extreme blade projections off the petiole were scored as ‘0.’ Similarly, bop1 bop2 aberrant floral patterning of extra abaxial petalloid sepals and bracts were scored as 0 and wildtype reversion as ‘3.’ Recovery of floral organ abscission was scored as ‘3’ while complete loss of abscission was scored as ‘0.’ Numbers between 0-3 indicate intermediate phenotypes. All transgenic lines with florescent signals showed complementation; highly expressing pBOP1:BOP2::EYFP transgenic plants showed more complete complementation suggesting that a threshold level of BOP2-EYFP protein is important for recovering BOP function. In addition, 35S::BOP2-EYFP transgenic lines were also generated and demonstrated dwarf, epinastic phenotypes (data not shown) agreeing with published reports (Norberg et al., 2005; Ha et al., 2007).  6.3.3 Subcellular localisation of pBOP1:BOP2::EYFP during lateral organ development Given the important role of subcellular localisation in NPR1 function, BOP2-EYFP protein subcellular localisation was examined in various aerial tissues of highly expressing transgenic  164  Table 6.2 pBOP1::BOP2-EYFP complementation of bop1bop2 Score of 3 (wild type) to 0 (bop1 bop2) for phenotypes in T1 plants  Transgenic lines  fluorescent signal intensity 3=high, 0 = low  entire T1 transformants selected on BASTA n=35 moderate expressors of BOP2-EYFP n=15 high expressors of BOP2-EYFP n=5  floral organ abscission blocked  blade-onpetiole phenotype  first whorl petalloid sepals  1.61±0.92  2.27±1.07  2.07±0.92  2.05±0.79  2.10±0.21  2.77±0.21  2.00±0.94  1.88±0.88  3.00±0.00  2.60±0.55  2.67±0.52  2.38±0.25  165  lines. Consistent with BOP1::GUS and BOP2 in situ hybridisation data presented in Chapter 2 (Figs. 2.7, 2.8), BOP2-EYFP was expressed in developing abscission zones, at the extreme base of lateral organs and at the base of cauline leaves (Fig. 6.3A-C). Cauline leaf expression was particularly intense at the junction between stipule and the base of the leaf (Fig. 6.3A, arrow) which correlated with a function in patterning stipule outgrowth (bop1 bop2 mutants do not show stipules flanking the base of cauline leaves, Fig. 2.5J,K). In addition to strong florescence in this region, the nuclear localisation of BOP2-EYFP was strikingly intense although signal was observed in the cytoplasm as well. Nuclear and cytoplasmic localisation was also observed in petal and sepal AZ cells on the receptacle revealed following organ dissection (Fig. 6.3D,E) and in cells from the basal regions of a petal blade (Fig. 6.3). BOP2-EYFP fluorescence was particularly strong on the adaxial sides of the proximal cotyledons which also showed intense nuclear localisation (Fig. 6.3G). Cytoplasmic and nuclear localisation was observed in the tissue of young rosette petioles; punctate dots of florescence were observed in the cytoplasm of some of these cells (Fig. 6.3I). BOP2-EYFP expression was also detected throughout the stele of the primary root and exhibited dramatic nuclear localisation in vascular tissue directed underlying the emerging lateral root (Fig. 6.3H). 35S::BOP2-EYFP transgenic plants displayed cytoplasmic and nuclear localisation in all tissues examined (data not shown) confirming results from 35S::BOP2-YFP in roots (Hepworth et al., 2005).  166  Figure 6.3 BOP2-EYFP localisation in pBOP1::BOP2-EYFP bop1 bop2 plants BOP2-EYFP localisation in various plant tissues (A) BOP2-EYFP was detected in the stipule junction at the base of cauline leaves. (B) expression in the AZ layer of a detached petal from stage 12 flower. (C) AZ expression in stage 9/10 floral bud. (D-E) nuclear and cytoplasmic expression in AZ cells on the receptacle of stage 12 flowers (D) petal AZ. (E) sepal AZ. (F) nuclear and cytoplasmic localisation in the proximal petal blade. (F) strong nuclear localisation on the adaxial side of the cotyledons. (H) nuclear localisation in the primary root stele. (I) strong nuclear expression with punctuate dots in the cytoplasm of petiole tissue.  167  168  6.3.4 BOP2 interacts with PAN and as well as itself in planta 6.3.4.1 Positive and negative controls BOP2 and PAN were cloned into 35S::EYFP vectors to generate translational fusions. Initially, I attempted biolistic delivery to express these vectors in onion epidermal cells and tobacco leaves. Expression of EYFP alone and BOP2-EFYP yielded both cytoplasmic and nuclear florescent signals (data not shown) while no signal was detected for BOP2-nEYFP and PAN-cEYFP cotransformation or for BOP2-cEYFP and PAN-nEYFP transformation. However, the biolistic method gave highly variable transformation effiencies. Therefore, I switched to an Arabidopsis leaf mesophyll-derived protoplast expression system (Sang-Dong et al., 2007) which provided high and consistent transformation efficiencies. This is especially critical for BiFC applications since a cell must be transfected with both constructs which much express at high enough levels to visualize protein-protein interactions, if these exist. Protoplasts expressing free EFYP demonstrated both strong nuclear and cytoplasmic fluorescence (Fig 6.4B). PAN-EYFP fusion proteins localised exclusively to the nucleus of transformed protoplasts (Fig. 6.4C), agreeing with its role as a transcription factor and with previous studies (Li et al., 2009). Usually, protoplasts transformed with BOP2-EYFP emitted both nuclear and cytoplasmic fluorescence (Fig. 6.4D,E); however, often speckled fluorescence both within and out of the nucleus was observed (Fig. 6.4F). Protoplasts transformed with BOP2-nEYFP or PAN-cEYFP did not show a flourescence signal (Fig. 6.4G,H). Similarly, protoplasts expressing solely BOP2-cEYFP or PAN-nEYFP did not show florescence (data not shown).  169  Figure 6.4 In planta interaction and localisation of BOP2 and PAN fluorescently-tagged fusion proteins Mesophyll protoplasts 12-20 hours following transfection and monitored for YFP signal. Inset in panels show the corresponding DIC image under visible light (A) No vector background florescence control. (B) free EYFP localizes to the nucleus and the cytoplasm. (C) PAN-EYFP fusion protein localises exclusively to the nucleus. (D-E) BOP2-EYFP fusion protein localises to the cytoplasm and the nucleus. (F) infrequently, the fluorescent signal occurs in punctuate spots throughout the protoplast. (G) BOP2-nEYFP fusion protein shows no signal. (H) PAN-cEYFP fusion proteins shows no signal. (I) Protoplasts expressing both BOP2-nEYFP and PAN-cEYFP show nuclear localised florescence. (J) Protoplasts expressing both BOP2-nEYFP and BOPcEYFP often exhibit multiple florescent signals within the nucleus and cytoplasm. (K) BOP2nEYFP and BOP2-cEYFP signal localises to the nucleus as to small dots surrounding the nucleus. (L) Exclusive nuclear signal from BOP2-nEYFP and BOP2-cEYFP transformed protoplasts.  170  171  6.3.4.2 In planta confirmation of BOP and PAN nuclear interaction As described in Hepworth et al (2005), PAN and BOP1 as well as PAN and BOP2 interaction was detected in yeast two-hybrid experiments. To determine whether PAN and BOP2 interact in planta and define the subcellular localisation of this putative interaction, BOP2-nEYFP + PANcEYFP as well as PAN-nEYFP + BOP2-cEYFP vectors were transformed into protoplasts. Both combinations gave an exclusively nuclear signal of reconstituted fluorescence, indicating that BOP and PAN interact in the protoplast system (Fig 6.4D, BOP2-nEYFP + PAN-cEYFP shown). This is the first in planta evidence of BOP and PAN protein interaction. 6.3.4.3 In planta detection of BOP - BOP interaction in cytoplasm and nucleus Given that NPR1 is presumed to form homomultimers in the cytoplasm, I tested the possibility that BOP proteins may also form higher-order protein complexes with themselves. Protoplasts transformed with BOP2-nEYFP and BOP2-cEYFP encoding plasmids showed reconstituted florescence indicating that BOP2 interacts with other BOP2 polypeptides in planta. The subcellular localisation was variable. All fluorescing protoplasts showed nuclear localisation of BOP2-EYFP +BOP2-EYFP interaction (Fig. 64.J-K). Often, the signal was cytoplasmic as well and appeared to be broken into large and small blebs in about 30% of the fluorescing protoplasts (Fig. 6.4J). The nuclear signal was sometimes accompanied by small, punctuate surrounding signals (Fig. 6.4K). Exclusive nuclear localisation was seen in approximately 30% of fluorescing protoplasts (Fig. 6.4L).  172  6.4 DISCUSSION 6.4.1 Distribution of BOP-like proteins across in land plants Phylogenetic analysis presented here suggests that the NPR1-like protein family is split into four distinct subgroups consistently across land plants. NPR1 function and TGA-dependent signalling is conserved in its dicot and monocot homologues (Chern et al., 2005; Malnoy et al., 2007; Zhang et al., 2009; Le Henanff et al., 2009). BOP proteins form a distinct subgroup in the NPR1like protein family across land plants; it will be interesting to determine if the roles of BOP1 and BOP2 in patterning the basal regions of both vegetative and reproductive lateral organs are similarly conserved. Expression data is lacking for most BPLs with the exception OsBPL1, OSBPL2 and OsBPL3, the rice homologues, which show tissue-specific expression in both lateral organs and meristems implying that they may play roles in the development of these tissues. BOP1 and BOP2 are crucial for differentiation of leaf and floral organ abscission zones (Chapter 2), the latter being an almost ubiquitous anatomical feature of plants, first detected in a lycophyte fossil from the Devonian period (Chaloner, 1968). Non-vascular plants exhibit abscission in the form of filament and segment shedding (Addicott, 1982) suggesting that a similar role exists for patterning separation zones in these organisms. Interestingly, three homologues detected in Physcomitrella show stronger homology with the BOP proteins compared to other NPR1-like proteins, suggesting that the patterning functions of BOP, if conserved, originated before the defense-role played by NPR1-4 family members. Although abscission is an extremely common life feature of plants, Endress’ (2008) comprehensive study of perianth biology in the lower angiosperms suggests that persistence of the tepals (sepals) is characteristic in this group. Moreover, histological analyses on the base of these floral organs 173  display features similar to bop1 bop2 in that the small, cytoplasmically-dense cells and the indentation of tissue normally associated with AZs are absent (Endress, 2008). Interestingly, floral nectaries are absent in most of the basal angiosperms (Endress, 2001) again highlighting a relationship between the presence of floral organ boundaries and nectary formation. As more sequence data becomes available, it will be important to examine these non-abscising lower angiosperms for tissue-specific expression and residue conservation of BOP homologues.  6.4.2 Conservation of protein-protein interactions of BOP2 and NPR1 My results suggest similarities exist between NPR1 protein interactions and those of BOP1 and BOP2. Firstly, in support of yeast two-hybrid data (Hepworth et al., 2005), BOP2 proteins interact in planta with the TGA protein, PAN, and this interaction was nuclear localised as shown for NPR1 and TGA interaction. Interaction between NPR1 and TGAs is dependent on the redox-regulation of NPR1 nuclear localisation. It is unknown if BOP2 protein localisation is similarly sensitive to cellular redox state. Exogenous application of dithiothreitol, hydrogen peroxide and SA did not noticeably affect BOP2-EYFP localisation (data not shown). Interstingly, when expressed in bombarded leaf and onion skin, NPR1-GFP showed preferential nuclear fluorescence which was presumed to reflect a SAR-induced state in response to bombardment (Kinkema et al., 2000). Corresponding bombardment experiments showed both strong nuclear and cytoplasmic localisation of BOP2-EYFP, suggesting that its localisation may not be affected in a SAR-sensitive manner. Trangenic pBOP1::BOP2-EYFP plants showed intense nuclear localisation of BOP2EYFP in regions where BOP2 is expected to function suggesting that its subcellular localisation is developmentally regulated. Striking nuclear localisation of BOP2-EYFP was observed in the 174  stele underlying the emerging lateral root primordium (Fig. 6.3H). This expression pattern implies that BOP proteins may be involved in lateral root development; however, no root phenotypes have been described for bop1 bop2. Bimolecular fluorescence complementation indicated that BOP2 + BOP2 proteins interact, a behaviour consistent with that known for NPR1. Punctate cytoplasmic localisation of florescence in pBOP1::BOP2-EYFP plants (Fig. 6.3I) as well as BOP2-EYFP expressing protoplasts (Fig. 6.4F) suggests that the BOP2-EYFP proteins may form cytoplasmic clusters. NPR1 is proposed to form oligomers in the cytoplasm through interprotein disulphide bonds between specific cysteine residues (Mou et al., 2003). However, unlike the putative cytoplasmic localisation of NPR1 oligomers, localisation of the BOP2+BOP2 complexes was predominantly nuclear suggesting that the regulation of subcellular localisation of the BOP proteins may be different than that of NPR1, a notion further supported by the lack of bipartite nuclear localisation signal in BOP1 and BOP2 versus NPR1. The BOP proteins may localise to the nucleus through the interaction with another protein which has a nuclear localisation signal or the BOP proteins may contain an as yet unrecognised nuclear localisation sequence. BOP1 and BOP2 proteins share one (Cys82) of two cysteines important for oligomerisation in NPR1, yet interact in planta, suggesting that Cys261 is not critical to promote interactions between BOP proteins and raising the possibility that other factors promote BOP+BOP interaction. Mutant versions of NPR1 which lack Cys216, like BOP1 and BOP2, showed enhanced nuclear localisation (Mou et al., 2003) which hints at the potential importance of this residue in cytoplasmic retention.  175  6.4.3 Biological significance of BOP and PAN interaction The expression domains of BOP1, BOP2 and PAN overlap and triple mutant analysis suggests a shared role in floral patterning (Chuang et al., 1999; Hepworth et al., 2005; this thesis). Although targets of BOP-PAN regulation are unknown, PAN was recently shown to interact directly with the promoter of the AGAMOUS (AG) to activate its expression in the inner whorls of the flower (Das et al., 2009; Maier et al., 2009). It is unknown whether BOP may participate in PAN regulation of AG; however, results presented Chapter 5 suggest that BOP1 and BOP2 may be involved in repressing AG expression. If this is the case, then BOP proteins may act to inhibit PAN-promotion of AG expression in a manner reminiscent of NPR3 and NPR4 negative regulation of PR gene expression (Zhang et al., 2006). Other potential partners of the BOP proteins are the NIMINS (NIM1-interacting), known interact with NPR1 to hinder the ability of NPR1 to transactivate TGA promotion of PR gene expression during SAR (Weigel et al., 2001; 2005). Recent evidence shows that NIMIN2 interaction with NPR1 modifies NPR1 into a transcriptional repressor (Chern et al., 2008). This suggests that the NPR1-TGA interaction can promote or inhibit PR transcription, depending upon tertiary interactors. The BOP-PAN interaction may be similarly modulated, although a developmental role for NIMINs has not been described. Exciting recent work from the Zachgo laboratory suggest that PAN activity may be redox-regulated (Li et al., 2009). Li et al (2009) showed that PAN, TGA2, TGA3 and TGA7 interact in yeast two-hybrid with ROXY, a nuclear-localised glutaredoxin involved in promotion of petal development (Xing et al., 2008). PAN transactivation of target genes was shown to be dependent on a conserved cysteine residue, suggested to be a target of reduction by ROXY (Li et al., 2009). Based on opposing loss of function phenotypes, ROXY is proposed to inhibit PAN 176  function (Li et al., 2009) which suggests that the reduced thiol form of PAN is inactive. This contrasts with the activation of TGA1 by cysteine residue reduction (Despres et al., 2003). It will be interestingly to determine if BOP and PAN interaction is sensitive to ROXY activity. Since loss of function pan-1 phenotypes appear to be limited to floral symmetry and floral determinancy (Chuang et al., 1999; Das et al., 2009; Maier et al., 2009), it is unclear whether BOP-PAN interactions are important in mediating other roles of the BOP genes, namely, in abscission zone and leaf patterning. In yeast two-hybrid assays, BOP1 and BOP2 interacted most strongly with PAN but were also shown to interact with other TGAs. BOP1 showed interaction with TGA1, TGA4, TGA3 and TGA2 while BOP2 displayed preference for TGA5, TGA6, TGA7. Interestingly NPR1 did not interact with TGA1 or TGA4 in yeast twohybrid (Zhang et al., 1999; Depres et al., 2000; Zhou et al., 2000; Hepworth et al., 2005); however, Depres et al (2003) suggest that this is due to the oxidised state of these proteins in yeast since they do interact in planta following SAR induction. Therefore, interaction between BOP1 and TGA1 and TGA4 may occur under different redox conditions to that of NPR1. Although TGA6 and TGA2 are implicated in regulating PR gene expression (Zhang et al., 2003) this does not preclude possible role(s) in developmental patterning. In fact, TGA2 homologues in tobacco appear to promote petal identity (Thurow et al., 2005). Furthermore, TGA4 interacts with CONSTANS, a key regulator of photoperiod flowering-time as well as with the promoter of FT (Tada et al., 2008). Quantitative RT-PCR suggests that FT transcript levels are ~40% lower in bop1 bop2 plants (S. Hepworth, pers. comm.) hinting at a role for BOP1 and BOP2 proteins in FT expression, which could potentially involve interaction with TGA4. Thus, BOP1 and BOP2 may mediate their variety of functions through interaction with a multitude of TGA partners.  177  Bimolecular flourescence complementation assays will be a quick method to ascertain in planta whether these interactions occur.  178  7 CONCLUSIONS This dissertation examined the roles of the BLADE-ON-PETIOLE1 (BOP1) and BLADE-ONPETIOLE2 (BOP2) genes during lateral organ development with a focus on their function in floral receptacle patterning and roles in floral meristem identity. Section 7.1 will highlight my contributions to the field while Section 7.2 will synthesise my findings to postulate general roles of BOP function during lateral organ development in Arabidopsis.  7.1 THESIS CONTRIBUTIONS 7.1.1 Roles of BOP1 and BOP2 in abscission I determined that BOP1 and BOP2 are required for aspects of AZ-related anatomy in both floral organ and leaf boundaries and that this anatomy was essential for abscission to occur in wildtype as well as 35S::IDA plants. These are the first genes in Arabidopsis shown to be critical for AZ formation in all abscising floral organs and for vestigial AZ formation in cauline leaves. Interestingly, expression of abscission-related genes was relatively unperturbed in bop1 bop2 mutants, indicating that activation of AZ-specific gene expression is independent of AZ anatomy. This project greatly contributed to our understanding of AZ patterning and underscored the genetic separation between the formation of the AZ anatomy and the later activation of the abscission program.  7.1.2 Contributions of BOP1 and BOP2 to nectary development As a manner of course for experiments in objective 7.1, I uncovered an undescribed role for BOP1 and BOP2 in promotion of nectary gland elaboration. Genetic analysis suggests that BOP1 179  and BOP2 mediate this effect downstream of CRABS-CLAW, a key mediator of nectary identity. Furthermore, partial recovery of nectary development by a loss of function in the SUPERMAN gene suggests that lack of nectary elaboration may result from a loss of proliferative capacity in the nectary primordia of bop1 bop2. Thus, BOP1 and BOP2 promote the proliferation of nectariferous tissue in Arabidopsis.  7.1.3 Analysis of recovered abscising lines I conducted a screen for secondary suppressors of bop1 bop2 and recovered two lines showing restoration of floral organ abscission in sepals and petals only, named recovered abscissor1 (rva1) and rva2. Both lines represent recessive and single mutant loci. Preliminary phenotypic analyses suggest that RVA1 and RVA2 may normally repress AZ activation.  7.1.4 Roles of BOP1 and BOP2 in floral meristem identity I evaluated the contribution of BOP1 and BOP2 to floral meristem identity and the extent to which these roles are shared with LEAFY (Lin et al., 2003) and APETALA (Hong and Tucker, 1998). Triple mutant analysis suggests that BOP1 and BOP2 act as potentiators of LFY and AP1 promotion of floral meristem identity and that they function in parallel to repress the expression of inflorescence identity genes.  7.1.5 Biochemical mechanism and phylogenetic analysis of BOP1 and BOP2 A phylogenetic analysis of the NPR1 protein family in land plants suggested that BOP1 and BOP2 homologues, named BOP-LIKEs (BPLs) exist in dicots and monocots and that they cluster independently from NPR1/NPR2 and NPR3/NPR3 protein pairs. I also examined the subcellular localisation of BOP2 during lateral organ development. Furthermore, the 180  biochemical interaction between BOP2 and PERIANTHIA (PAN) and between BOP2 and BOP2 proteins was confirmed in planta using bimolecular fluorescent complementation.  7.2 SYNTHESIS 7.2.1 BOP1 and BOP2 are downstream executors of positional programs Although cell lineage may contribute to cell specification, clonal and molecular studies suggest that the cell fate of any given meristem cell is determined by position-dependent mechanisms (Scheres, 2001). How positional information is established is an area of intensive research although the importance of morphogenic auxin gradients is undisputed (Bowman and Floyd, 2008; Benkova et al., 2009; Wolters and Jurgens, 2009). Cohesive plant architecture requires that positional information be communicated from the meristem to the lateral organs and likely involves non cell autonomous pathways operating between cell layers and between tissue types (Ori et al., 2000). In response to positional cues originating from the meristem, master regulatory factors enlist a suite of effectors whose coordinated activities differentiate the appropriate cell type (Benkova et al., 2009). Based on loss of function phenotypes, BOP1 and BOP2 are involved in the specification of correct cell identity in lateral primordia often by repressing the development of inappropriate cell fates; in particular, BOP1 and BOP2 play early potentiating roles in the floral transition in repression of secondary inflorescence shoot identity and later roles in proximal organ and boundary morphogenesis. Firstly, by and large, floral meristem identity is established in bop1 bop2 mutants excepting the occasional development of the floral bract. Other contributions of BOP1 and 181  BOP2 to floral meristem identity do not manifest unless LFY or AP1 function is attenuated, suggesting that BOP1 and BOP2 function as cofactors or potentiators in this process rather than floral meristem identity genes. Once lateral organs start to develop, positional cues promote differentiation of appropriate cell fate. BOP1 and BOP2 are recruited to repress the formation of inappropriate tissues in proximal lateral organ regions. The type of tissue repressed depends on the developmental context. In leaves, BOP1 and BOP2 are recruited to repress ectopic blade formation from proximal regions in rosette and cauline leaves (Ha et al., 2004, 2007; Hepworth et al., 2005; Norbeg et al., 2005). It is important to note that the ectopic blades of bop1 bop2 are resemble normal blade tissues, suggesting that BOP1 and BOP2 do not impact morphogenesis pathways of leaf cell identity but ensure that positionally appropriate programmes are activated. Distal genes such as JAG and NUB are expressed proximally in bop1 bop2 leaves (Norberg et al., 2005) while at the same time, expression of KNOX genes, such as BP, normally restricted to the leaf base, expand into the petiole (Ha et al., 2003). Furthermore, characteristics associated with leaf–stem boundaries such as the growth of flanking lateral stipules and the vestigial AZs are absent. Although these data suggest that the boundaries between meristem, petiole and blade are fuzzy in bop1 bop2, blades do not encompass the petiole, as seen in as1-1 mutants (Ori et al., 2000; Byrne et al., 2000), and gross fusions between the petiole and the stem, as seen in cuc2 cuc3 mutants (Hibara et al., 2006), are not observed. Thus, positional cues for overall leaf architecture are still present in bop1 bop2, suggesting that BOP1 and BOP2 are important downstream potentiators of this patterning mainly acting to repress differentiation of distal cell types, such as blade tissue, in proximal regions and a second role to recruit cells to differentitate certain features of boundary structure at the bases of these lateral organs. 182  BOP1 and BOP2 roles are modified in the floral developmental context. BOP1 and BOP2 repress development of cryptic bract primordia subtending the abaxial flower pedicel. This function may also be mediated through the repression of lateral organ genes such as JAG. Similarly, extra growth on the abaxial side of the floral meristem can explain much of the floral phenotype of bop1bop2 mutants. Boundary characteristics of the floral organ-receptacle interface, in particular, the development of the AZ, are also regulated by BOP1 and BOP2. In bop1 bop2, the abscission zones cells are replaced by cells which are large and vacuolated, appearing similar to differentiated tissue of the flanking floral organs and receptacle. In addition, nectaries are also unelaborated and show large vacuolated tissue similar to the differentiated receptacle tissue. Similar to leaves, positional information is largely intact in bop1 bop2 as suggested by reporter gene analysis and lack of gross fusion between lateral organs and the receptacle. Thus, in this case, BOP1 and BOP2 act in a variety of floral patterning pathways to repress inappropriate lateral organ differentiation.  7.2.2 BOP1 and BOP2 as negative regulators of growth? BOP1 and BOP2 repression of blade development off the petioles and bract formation from the pedicels imply that these proteins are negative regulators of growth. However, AZs and nectaries develop due to localised proliferative activities, so the loss of their development is inconsistent with a general role of BOP is growth repression. Rather, I propose that BOP1 and BOP2 roles in abscission and leaf and bract formation may reflect that co-opting of BOP1 and BOP2 to different downstream processes that respond to positional signalling in the proximal organ.  183  7.2.3 Relationship between boundaries and proliferation AZs develop at the lateral organ-stem/receptacle boundary while nectaries develop between the stamen and sepal boundaries. Localised proliferation of receptacle tissue leads to nectaries and likely a combination of lateral organ and receptacle tissue contributes to floral organ abscission zones. Before the onset of proliferation, these tissues appear differentiated and are typified by large, vacuolated cells. Thus, formation of AZs and nectaries involves a reversion to quasimeristematic fate which likely involves dynamic regulation of KNOX genes. BP, KNAT2 and KNAT6 are all expressed in developing AZs and nectary tissue (Douglas and Riggs, 2005; Baum et al., 2001; Belles-Boix et al., 2006) and recent results suggest that they dynamically regulate each other’s expression (Ragni et al., 2008). Establishment of boundaries is tightly linked with meristem formation, maintenance and proliferation (Rast and Simon, 2008). CUCs are essential for activation and maintenance of KNOX gene expression in a range of meristematic tissue including SAM, axillary meristems and for serrations and leaflet formation (Aida et al., 1997; Takada et al., 2001; Belles-Boix et al., 2006; Blein et al., 2008; Raman et al., 2008) but it is unknown if they contribute to AZ or nectary gland development. An attractive possibility is that CUC-mediated positional signalling pathways may use BOP1 and BOP2 as downstream effectors to repress differentiated tissue identity in the boundaries. It will be interesting to determine the regulatory hierarchies among KNOX, CUCs and the BOP genes.  184  7.2.4 Abscission zones and nectaries proliferate from cells derived from differentiated tissues - the role of the receptacle Abscission zone differentiation and nectary growth likely both derive, in part, from receptacle tissue. The development of the receptacle has received very little attention from Arabidopsis developmental biologists yet flower architecture depends upon the integrity of this structure and thus, receptacle patterning warrants focused attention. Adaxial factors such as REV and abaxial facors such as FIL are required for receptacle formation as pedicels in rev-1 and fil-1 mutants terminated in filamentous projections lacking a receptacle base (Talbert et al., 1995; Sawa et al., 1999; Chen et al., 1999), hinting that the juxtaposition of adaxial and abaxial identities are required for receptacle formation similar to that observed in leaves. The receptacle develops around stage 9, manifested as swollen bulges directly below the sepals following the development of the pedicel and the floral organs (Douglas and Riggs, 2005; Smyth et al., 1990). This timing is coincidental with the start of nectary gland outgrowth and AZ differentiation (Baum et al., 2001; Gomez-Mena and Sablowski, 2008). Thus, BOP1 and BOP2 may be involved in recruiting and/or sequestering founder cell populations for AZ and nectary formation from differentiated receptacle tissue. The relative contribution of cell division versus cell expansion to receptacle growth is unclear. Although beyond the scope of this thesis, BOP1 and BOP2 are likely involved in receptacle growth since bop1 bop2 receptacles are shorter, thicker and develop on a slightly splayed axis compared with wild type (S. McKim, pers. obs.). Intriguingly, BP is involved in regulation of lignin biosynthesis: overexpression and loss of BP leads to mislocalisation of lignification causing localised areas to fully differentiate into large vacuolated cells which would not normally due so (Mele et al., 2003). Lignin deposition is a characteristic of secondary wall 185  biosynthesis and is considered an irreversible step in cellular differentiation (Groover and Jones, 1999; Evert, 2007). Thus, in bop1 bop2, temporal or spatial misregulation of BP could lead to the irreversible lignification in certain cell types, preventing further recruitment for receptacle and/ or nectary tissue. Loss of function genes in BP, LFY and AP1 result in the loss of the receptacle bulge (Douglas and Riggs, 2005). Given the shared roles between LFY and AP1 and BOP1 and BOP2 and the role of BOP1 and BOP2 in regulating BP expression in the leaf, possible networks regulating receptacle architecture may involve interactions between these players to coordinate the development of AZs, nectaries and receptacle bulges.  7.2.5 Do BOP1 and BOP2 respond to developmentally informative redox cues? Homology of BOP1 and BOP2 with NPR1 suggests an engaging hypothesis that BOP’s developmental roles may be redox modulated as is the case for NPR1. Redox regulation by reactive oxygen species (ROS) has long been associated with proteins involved in responding to oxidative stress due to pathogen or abiotic elicitors (Foyer et al., 1997). However, recent evidence suggests that redox regulation is also involved in developmental patterning. The postulated redox modification of PAN by the thioredoxin ROXY hints that BOP and PAN interactions may be similarly sensitive (Xu et al., 2008; Li et al., 2009). Furthermore, the DNA – binding activity of the HD-ZIPIIIs proteins, which like BOP1 and BOP2, are important mediators of adaxial cell fate in leaves, are controlled by redox-modification of cysteine residues (Comelli and Gonzalez, 2007). Blade expansion in maize leaves and pollen tube growth and root hair development in Arabidopsis are both induced by localised ROS production (Rodriguez et al., 2002; Foreman et al., 2003; Potocky et al., 2007) also highlighting the roles of redox control 186  of developmental patterning. Meristem integrity is also sensitive to cellular redox state (Jiang et al., 2006; Benitez-Alfonso et al., 2009). Finally, the sensitivity of the floral transition to environmental stress may also be mediated through ROS (Martinez et al., 2004; Archard et al., 2008) which highlights the cross-talk which must occur between environmental inputs and developmental programs in plants. A key question to answer is whether cellular redox status changes over the course of lateral organ development. A new redox-sensitive GFP transgenic Arabidopsis line may be a good method to address this problem (Meyer et al., 2007). Furthermore, Fahnenstich et al (2008) recently described an inducible ROS system in Arabidopsis These authors examined the effects following constitutive ROS production; however, the system could be easily modified to drive tissue-specific ROS induction at various stages of development.  187  BIBLIOGRAPHY Addicott,F.T. (1982). Abscission. Berkeley: University of California Press. Aida, M., Ishida, T., Fukaki, H., Fujisawa, H., and Tasaka, M. (1997). 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Plant Journal 19, 259-268.  206  APPENDIX 1)  Alleles used in thesis T-DNA = transposable DNA; EMS = ethylmethane sulfonate Allele name  Method  Ecotype  Type  Reference  bop1-3  T-DNA  Col-0  null  Hepworth et al (2005)  bop2-1  T-DNA  Col-0  slightly leaky  Hepworth et al (2005)  bop2-2  T-DNA  Col-0  null  Norberg et al (2005)  ida  EMS  C24  null  Butenko et al (2003)  etr1-1  EMS  Col-0  dominant  Vandenbussche, et al. (2007)  crc-1  EMS  Ler  null  Bowman et al (1999)  sup-1  EMS  Col-0  strong  Bowman et al (1992)  ap1-1  EMS  introgressed into Col-0  null  Irish and Sussex (1990)  ap1-12  EMS  Col-0  weak  Schultz and Haughn (1993)  lfy-2  EMS  Col-0  null  Schultz and Haughn (1993)  lfy-1  EMS  Col-0  weak  Schultz and Haughn (1993)  207  2) Primer Sequences 2.1) Genotyping Primers Primer  Sequence 5’ – 3’  lfy-2 dCAPs F  GTTTGGGGACAGAGAGACAGAGGGAGGATC  lfy-2 dCAPs R  CGCCACGGTCTTTAGCAATTGTCTGG  bop1-3 LP  GCACAATCTTTCGACTTCATCACC  bop1-3 RP  CGTACCCTTTGATTTTAGTATGCTG  bop2-1 LP  TCGACGCCGAAGTAACGAGAG  bop2-1 LP  CCCTTTTTATAATCAGCATCAAGA  ida 49 (RP)  GGTGTTTCTACTATGCGTGTG  +5074 (ida LP) ATTTGTCGTTTTATCAAAATGTAC 1.2) Cloning Primers Primer BOP1-cDNA-Kpn1F1 BOP2-cDNA-Kpn1F1  Sequence 5’ – 3’ CGCGGTACCATGAGCAATACTTTCGAAG GCGGGTACCGTTTCAGAGAGGAGGAGC  BOP1-cDNA-Kpn1-R  GCGGGTACCGAAGTGATGTTGATGATG  PAN-cDNA-Kpn1-F2 long  TGCAGTATCCGTATCCGGGGTACCATGCAGAGCAGCTTCAAA  PAN-cDNA-Kpn1 R  GCCGGTACCGTCTCTAGGTCTGGCTAA  BOP1-4kb-Age1-F1  GCTACCGGTAGGAGGGCATGTCCATAT  BOP1-4kb-Sac1-R1  CGAAGTACGAGCTCAGCTCCTTTGTTGATTTCTTT  208  1.3) In situ Primers Primer  Sequence 5’ – 3’  BOP2 LP CTCATATGAATGAGGAGCAC BOP2 RP GATAATACGACTCACTATAGGGACTCATACCTTCCCTCTGA CER6 LP ATGCCTCAGGCACCG CER6 RP GATAATACGACTCACTATAGGGTTATTTGAGTACACC 1.4) Quantitative Real-Time PCR Primers Primer  Sequence 5’ – 3’  IDA 68 Right  TCAATGAGGAAGAGAGTTAACAAAAG  IDA 68 Left  CTAAAGGCGTTCCCATTCCT  HAESA 82 Right GAGAGAGGGAATGGAGAG-AAGG HAESA 82 Left  CATGCTCGTCGGACCTTT  ACT2 102 Right  CCGGTACCATTGTCACACAC  ACT2 102 Left  CGCTCTTTCTTT-CCAAGCTC  2) Recipes Spurr’s Standard Resin Chemical  Amount  ERL 4206  10.0g  DER 736  6.0g  NSA  26.0g  DMAE  0.4g  209  AT Minimal Media (Haughn and Somerville, 1986) 5 mM KNO3 2.5 mM KH2PO4 (adjusted to pH 5.5) 2 mM MgSO4 2 mM Ca(NO3) 2, 51 µM Fe-EDTA 70 µM HaBO3 14 µM MnC12 0.5 µM CuSO4 1 µM ZnSO4 0.2 µM NaMoO~, 10 µM NaC1 0.01 µM COC12  210  

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