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I. .0 . . 0 . 0 00 CI ’ 30. 0.00 20H This is to certify that the dissertation entitled TISSUE- AND ORGAN-SPECIFIC PHYTOCHROME-MEDIATED RESPONSES IN ARABIDOPSIS THALIANA presented by SANKALPI NADEEKA WARNASOORIYA has been accepted towards fulfillment of the requirements for the Doctoral degree in Genetics .\ .:’:‘e.f_".'_..‘.'.'_'.: ’ ' at. "1;--i ........... f‘éfleim" ‘x Major Professorls Signature ILICZ/lfl Date MSU is an Affirmative Action/Equal Opportunity Employer LIBRARY Michigan State University - —A--‘-Ini_-_;Q'-L-.-A- PLACE IN RETURN BOX to remove this checkout from your record. To AVOID FINES return on or before date due. MAY BE RECALLED with earlier due date if requested. DATE DUE DATE DUE DATE DUE 5/08 K:IProj/Aoc&PrelelRC/Date0ue.indd TISSUE- AND ORGAN-SPECIFIC PHYTOCHROME-MEDIATED RESPONSES IN ARABIDOPSIS THALIANA By Sankalpi Nadeeka Wamasooriya A DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Genetics 2010 ABSTRACT TISSUE- AND ORGAN-SPECIFIC PHYTOCHROME-MEDIATED RESPONSES IN ARABIDOPSIS THALIANA By Sankalpi Nadeeka Wamasooriya The red/far-red light-absorbing phytochrome photoreceptor family regulates numerous responses throughout the life cycle of Arabidopsis thaliana. Despite the discovery of individual and redundant phytochrome functions through mutational analyses, conclusive reports on distinct sites of photoperception, and the mechanisms by which localized pools of phytochromes act at the molecular level in mediating tissue- and organ-specific responses are limited. The objectives of this thesis research are to identify sites of phytochrome photoperception in Arabidopsis, correlate them with the regulation of tissue- and organ-specific responses and recognize candidate downstream target genes that define the molecular bases of such responses. To address these objectives, a mammalian gene encoding an enzyme capable of reduction of functional phytochromes was expressed in a tissue-specific manner in transgenic Arabidopsis plants. Transgenic lines were probed for perturbed phytochrome-mediated responses under blue, red and far- red illumination and comparative phenotypic and photobiological analyses of transgenic lines with constitutive, mesophyll- and meristem-specific phytochrome inactivation revealed distinct functions of localized pools of phytochromes. Mesophyll-localized phyA was found to exert a dominant role on hypocotyl inhibition, whereas hypocotyl- localized phyA was implicated in regulation of hypocotyl elongation in the absence of the inhibitory action of mesophle-localized phyA under far-red light. Through comparative microarray-based gene expression profiling of transgenic Arabidopsis lines with constitutive and mesophyll—specific phytochrome inactivation and subsequent phenotypic characterization of mutants, this study also identified two proteins, a GNAT family protein and a caldesmon-related protein, as putative signaling intermediates in regulating phyA-mediated hypocotyl development under far-red light. Furthermore, phytochrome- dependent sucrose-stimulated anthocyanin accumulation was distinctively altered in transgenic lines with mesophyll-specific phytochrome inactivation. The analysis of anthocyanin pigmentation responses confirmed a functional role for mesophyll-localized phyA in regulating anthocyanin accumulation in far-red light and its contribution in blue light. Individual phytochrome isoforrns were recognized to have divergent roles in anthocyanin accumulation under red light; phyA through phyD exhibit inductive roles and phyE functions as a novel suppressor of anthocyanin accumulation. Additionally, metabolic inactivation of the phytochrome chromophore in roots suggested that root- localized phytochrome and/or the phytochrome chromophore is vital for the photoregulation of root development and confers sensitivity to jasmonic acid. Thus, conclusive evidence from this study indicates that the analyses of spatially isolated pools of phytochromes is an effective tool for providing novel insight into the complex signaling pathways controlled by phytochromes. DEDICATION To my mother, for her support from pre-school through graduate school ACKNOWLEDGMENTS I would like to acknowledge the people that have shared their knowledge, advice, experience and time with me during my graduate school years. I am grateful to my advisor, Dr. Beronda Montgomery, for her support, for investing time in discussions and for providing a stimulating work environment. Each member of my guidance committee, Dr. Jianping I-Iu, Dr. Robert Larkin, Dr. Gregg Howe and Dr. Daniel Jones, has given me thoughtful advice and constructive criticism which has been of great help. Dr. Barb Sears, the members of the Genetics Program and the Plant Research Laboratory have been supportive and encouraging throughout my graduate work at Michigan State University. A big thank you goes to past and present members of the Montgomery lab, especially to Melissa Whitaker, Bagmi Pattanaik, Sookyung Oh, Julie Bordowitz, and Stephanie Costigan whose help and support enabled me to complete this work. Lastly, I would like to express my sincere appreciation to my mother, Mrs. Padma Wamasooriya, my brother, Aminda Wamasooriya, and my husband, Ravin Kodikara, for the support and encouragement given during the years of my education. TABLE OF CONTENTS LIST OF TABLES ............................................................................................................. ix LIST OF FIGURES ............................................................................................................. x KEY TO ABBREVIATIONS ........................................................................................... xii CHAPTER 1 Introduction .................................................................................................... 1 1.1 Overview .................................................................................................................... 2 1.2 Substructure and Biosynthesis of Phytochromes ....................................................... 4 1.3 Phytochrome Activity at the Molecular Level ........................................................... 6 1.4 Physiological Responses Regulated by Phytochromes .............................................. 7 1.5 Tissue- and Organ-Specific Phytochrome Responses ................................................ 9 1.6 Probing Spatial-specific Responses via Targeted Chromophore Reduction ............ l 1 1.7 Summary .................................................................................................................. 15 1.8 References ................................................................................................................ 20 CHAPTER 2 Regulation of Far-red Light-Mediated High Irradiance Responses by Spatial-specific Phytochromes ........................................................................................... 29 2.1 Overview .................................................................................................................. 30 2.1.1 Phytochrome-dependent High Irradiance Responses ........................................ 31 2.1.2 Light Perception by Phytochrome A ................................................................. 32 2.1.3 Phytochrome A and Downstream Components in Signaling ............................ 34 2.1.4 Outlook .............................................................................................................. 36 2.2 Materials and Methods ............................................................................................. 37 2.2.1 Photomorphogenesis of Transgenic Lines with Targeted Chromophore Inactivation in PRC ..................................................................................................... 37 2.2.2 Quantification of Anthocyanin Levels in Transgenic Lines with Targeted Chromophore Inactivation .......................................................................................... 38 2.2.3 Gene Expression Analysis ................................................................................. 39 2.2.4 Validation of At] g26220, At4g02290 and At1g52410 Expression in Transgenic Lines with Targeted Chromophore Inactivation by RT-PCR ................................... 41 2.2.5 Confirmation of T-DNA Insertion Mutants ...................................................... 43 2.2.6 Hypocotyl Inhibition Assay ............................................................................... 44 2.3 Results and Discussion ............................................................................................. 45 2.3.1 Mesophyll-specific Phytochromes Have Distinct Regulatory Roles in F ar-red Light ........................................................................................................................... 45 2.3.2 Phytochrome A is Involved in Regulating Anthocyanin Accumulation in FRc and Be ......................................................................................................................... 49 2.3.3 Mesophyll-localized Phytochrome Inactivation Leads to Distinct Gene Expression Patterns .................................................................................................... 52 2.3.4 GCNS- and Caldesmon~related Proteins are Implicated in the Regulation of Hypocotyl Development under F Rc ........................................................................... 57 2.3.5 Summary ............................................................................................................ 60 vi 2.4 Future Perspectives .................................................................................................. 61 2.6 References ................................................................................................................ 79 CHAPTER 3 Phytochrome-mediated Light-dependent Anthocyanin Accumulation in Red Light ................................................................................................................................... 87 3.1 Overview .................................................................................................................. 88 3.1.1 Anthocyanin Biosynthesis and Accumulation ................................................... 89 3.1.2 Spatial-specific Accumulation of Anthocyanins ............................................... 93 3.1.3 Functions of Phytochromes in Spatial-specific Anthocyanin Accumulation....95 3.1.4 Outlook .............................................................................................................. 97 3.2 Materials and Methods ............................................................................................. 98 3.2.1 Quantification of Anthocyanin Levels in Transgenic Lines with Targeted Chromophore Inactivation ......................................... . ................................................. 98 3.2.2 Confirmation of Apophytochrome Mutants ...................................................... 99 3.2.3 Quantification of Anthocyanin Levels in Apophytochrome Mutants ............. 101 3.2.4 Expression Levels of Anthocyanin Marker Genes .......................................... 101 3.2.5 Complementation of phyE Mutant .................................................................. 103 3.2.6 Arabidopsis Seedling Extracts ......................................................................... 105 3.2.7 Expression of PH YE-m6 in Complemented phyE Mutant .............................. 106 3.2.8 Quantification of Anthocyanin Levels in Complemented phyE Mutant ......... 106 3.3 Results and Discussion ........................................................................................... 107 3.3.1 Spatial-specific Phytochromes Regulate Anthocyanin Accumulation in Rc ..107 3.3.2 Phytochrome Family Members Have Differential Roles in Anthocyanin Accumulation in Rc .................................................................................................. 109 3.3.3 Anthocyanin Marker Genes are Differentially Expressed in Rc ..................... 114 3.3.4 Summary .......................................................................................................... 1 18 3.4 Future Perspectives ................................................................................................ 119 3.5 References .............................................................................................................. 132 CHAPTER 4 Regulation of Root Development by Phytochromes and Jasmonic Acid..141 4.1 Overview ................................................................................................................ 142 4.1.1 Role of Phytochromes in Root Growth and Development .............................. 143 4.1.2 Biology of Jasmonic Acid ............................................................................... 146 4.1.3 Phytochrome and Jasmonic Acid Signaling .................................................... 148 4.1.4 Targeted Chromophore Deficiency through Transactivation of Biliverdin Reductase .................................................................................................................. 150 4.1.5 Outlook ............................................................................................................ 151 4.2 Materials and Methods ........................................................................................... 152 4.2.1 Transactivation of BVR .................................................................................... 152 4.2.2 Whole-mount lmmunohistochemistry ............................................................. 153 4.2.3 Arabidopsis Seedling Extracts ......................................................................... 155 4.2.4 Immunoblotting ............................................................................................... 156 4.2.5 Hypocotyl Inhibition Assays ........................................................................... 157 4.2.6 Root Inhibition Assays .................................................................................... 157 4.2.7 Expression Levels of Jasmonic Acid-inducible Marker Genes ....................... 158 4.3 Results and Discussion ........................................................................................... 159 vii 4.3.1 Transactivation of BVR in M0062>>UAS-BVR ............................................. 160 4.3.2 Root-localized Phytochrome Inactivation does not Affect Hypocotyl Inhibition Response ................................................................................................................... 161 4.3.3 Phytochrome or Phytochromobilin Affects Root Elongation in Arabidopsis .162 4.3.4 Root-localized Phytochrome or Phytochromobilin Reduces J asmonic Acid- mediated Root Inhibition .......................................................................................... 165 4.3.5 Root-localized Phytochrome Deficiencies Impact Expression of Jasmonic Acid-inducible Marker Genes .................................................................................. 169 4.3.6 Summary .......................................................................................................... g 173 4.4 Future Perspectives ............................................................................................. 174 4.5 References .............................................................................................................. 191 APPENDIX ...................................................................................................................... 199 A1 Overview ................................................................................................................ 200 A2 Materials and Methods ........................................................................................... 201 A21 Generation of 10571>>UAS-BVR .................................................................. 201 A22 Plant Growth .................................................................................................... 202 A23 Reagent Preparation ......................................................................................... 202 A24 Leaf Protoplast Isolation .................................................................................. 203 A25 Protoplast Sorting by Fluorescence-Activated Cell Sorting (FACS) .............. 205 A3 Results .................................................................................................................... 206 A4 Discussion .............................................................................................................. 207 A5 References .............................................................................................................. 218 viii LIST OF TABLES Table 1 List of genes with unique differential expression in CAB3::pBVR2 vs. 358::pBVR3 ....................................................................................................................... 71 Table 2 F old-difference in root lengths of wild-type, BVR-expressing and mutant seedlings ........................................................................................................................... 186 Table A1 GFP-positive protoplasts before and after Fluorescence Activated Cell Sorting (F ACS) ............................................................................................................................. 213 Table A2 Quantification of RNA ..................................................................................... 217 LIST OF FIGURES "Images in this dissertation are presented in color." Figure 1.1 Conserved domain structure of phytochrome in plants .................................... 17 Figure 1.2 Biosynthesis of holophytochrome in Arabidopsis ............................................ 18 Figure 1.3 Inactivation of holophytochrome through expression of BVR ......................... 19 Figure 2.1 Photomorphogenesis of F Rc-grown wild-type and transgenic BVR plants ..... 66 Figure 2.2 Anthocyanin content of wild-type and transgenic BVR seedlings ................... 67 Figure 2.3 Development of wild-type, transgenic BVR and phy mutant seedlings under continuous blue light .......................................................................................................... 68 Figure 2.4 Anthocyanin content of wild-type and transgenic BVR seedlings ................... 69 Figure 2.5 Differential expression patterns in 35S::pBVR3 and CAB3::pBVR2 ............. 70 Figure 2.6 Validation of microarray analysis for At1g26220, Ar4g02290 and Atlg52410 through analysis of transcript accumulation ...................................................................... 72 Figure 2.7 Expression profiles in response to different light conditions ........................... 73 Figure 2.8 Expression profiles in different tissues of Arabidopsis seedlings .................... 74 Figure 2.9 Site of T-DNA insertion and transcript accumulation in SALK_062388 ........ 75 Figure 2.10 Site of T-DNA insertion and transcript accumulation in SALK_101567 ...... 76 Figure 2.1 1 Site of T-DNA insertion and transcript accumulation in SALK_151393 ...... 77 Figure 2.12 Mean hypocotyl length of T-DNA insertion mutants ..................................... 78 Figure 3.1 Anthocyanin content of wild-type and transgenic BVR seedlings ................. 124 Figure 3.2 Sites of T-DNA insertions in apophytochrome mutants and locations of gene- specific oligonucleotide annealing ................................................................................... 125 Figure 3.3 Analysis of T-DNA alleles in apophytochrome mutants ............................... 126 Figure 3.4 Development and sucrose-dependent anthocyanin accumulation of wild-type and apophytochrome mutants .......................................................................................... 127 Figure 3.5 Sucrose-dependent anthocyanin accumulation in the complemented phyE mutants ............................................................................................................................. 128 Figure 3.6 Phytochrome protein accumulation in wild-type and the complemented phyE mutants ............................................................................................................................. 129 Figure 3.7 Expression of anthocyanin marker genes ....................................................... 130 Figure 3.8 Quantification of expression levels of anthocyanin marker genes ................. 131 Figure 4.1 Whole-mount immunolocalization of BVR protein accumulation in roots of M0062>>UAS-BVR by Confocal laser scanning microscopy ........................................ 178 Figure 4.2 BVR protein accumulation in No-O wild-type and 3SSzchVR1 transgenic seedlings ........................................................................................................................... 179 Figure 4.3 Photomorphogenesis of wild-type and BVR-expressing seedlings ................ 180 Figure 4.4 Mean hypocotyl lengths of wild-type and BVR-expressing seedlings ........... 181 Figure 4.5 Mean root lengths of wild-type, BVR-expressing and mutant seedlings ........ 182 Figure 4.6 Mean root lengths of wild-type, BVR-expressing and mutant seedlings ........ 184 Figure 4.7 Relative expression levels of 0PR3 in wild-type, BVR-expressing and mutant seedlings ........................................................................................................................... 187 Figure 4.8 Relative expression levels of VSPI in wild-type, BVR-expressing and mutant seedlings ........................................................................................................................... 189 Figure A1 GAL4 enhancer-trap-based induction of Biliverdin reductase (B VR) expression in transgenic Arabidopsis thaliana plants ........................................................................ 209 Figure A2 Expression patterns of green fluorescent protein in enhancer trap lines ........ 21 1 Figure A3 Fluorescence Activated Cell Sorting (F AC S) acquisition dot plots ............... 212 Figure A4 Fluorescence Activated Cell Sorting (F ACS) acquisition dot plots ............... 214 Figure A5 Confocal laser scanning microscopy of plant protoplasts used for fluorescence activated cell sorting ........................................................................................................ 215 Figure A6 Confocal laser scanning microscopy of protoplasts of J0571>>UAS-BVR used for fluorescence activated cell sorting ............................................................................. 216 xi KEY TO ABBREVIATIONS avg - average B-bMe Bc - continuous blue bHLH - basic helix-loop-helix BphPs - bacteriophytochromes BR - bilirubin BV - biliverdin BVR - biliverdin reductase BVR - gene encoding biliverdin reductase CAB3 - promoter of gene encoding chlorophyll a/b binding protein CaMV - Cauliflower Mosaic Virus promoter chl - chlorophyll CLSM - confocal laser scanning microscopy Cphs — Cyanobacterial phytochromes cry2 - cryptochrome 2 D - dark DEG -differentially expressed genes F ACS - fluorescence-assisted cell sorting FP - forward primer F phs — fungal phytochromes FR - far-red light F Rc - continuous far-red light xii FR-HIR - F ar-red high irradiance responses GFP - green fluorescent protein GNAT - GCNS-related N-acetyltransferase HCI - hydrochloric acid HIR - high irradiance responses HRP - horseradish peroxidase Ile - isoleucine JA - jasmonic acid JA-Ile -jasmonoy1-L-isoleucine KOH - potassium hydroxide LC-MS - liquid chromatography—mass spectrometry LFR - low fluence responses MeJA - methyl jasmonate MES - 2-(N-morpholino) ethanesulfonic acid NDPKZ - nucleoside diphosphate kinase 2 NIH - National Institute of Health NTE - plant-specific amino-terminal extension PAS - Per—Amt—Sim PBS - phycobilisome PCB - phycocyanobilin PCR - polymerase chain reaction PCDB - phytochromobilin P¢R - phytochromorubin xiii qRT-PCR - quantitative reverse transcription polymerase chain reaction QTL - quantitative trait locus R-md Rc - continuous red R-HIR - red high irradiance responses RP - reverse primer rpm - revolutions per minute RT-PCR - reverse transcription polymerase chain reaction SD - standard deviation SDS - sodium dodecyl sulfate SDS-PAGE - sodium dodecyl sulfate -polyacrylamide gel electrophoresis suc -sucrose TAIR - The Arabidopsis Information Resource T-DNA - transfer-DNA UAS - upstream activating sequence UBC2I — gene encoding ubiquitin-conjugating enzyme 21 VLF R - very low fluence responses W - white Wc - continuous white WT - wild-type YEP - yeast extract-peptone xiv Chapter 1 Introduction Some of the information included in chapter 1 was published in a book chapter: Wamasooriya SN and Montgomery BL, Using Transgenic Modulation of Protein Accumulation to Probe Protein Signaling Networks in Arabidopsis thaliana, Plant Biotechnology and Transgenic Research, Bentham Science Publishers, Oak Park, IL (in press). 1.1 Overview Perception of changes in the environment is important to any living organism. To monitor environmental variations, plants are equipped with multiple sensory systems. Among the environmental factors that determine plant survival and reproduction, light is the most variable and dominating factor. In addition to being the source of energy for photosynthesis, it mediates a myriad of growth, developmental and adaptive processes throughout the life cycle of plants (Chory et a1., 1996; Franklin and Quail, 2010; Franklin and Whitelam, 2004; Neff, F ankhauser, and Chory, 2000; Schepens, Duek, and Fankhauser, 2004). Proper onset and fine-tuning of developmental transitions and adaptive processes not only require detection of the presence or absence of light but also spectral quality, quantity, directionality and periodicity. To perceive fluctuations in the temporal and spatial patterns of light, photoreceptors function at the interface between the organism and the environment. The regulation of plant growth and development by light signals, otherwise known as photomorphogenesis, involves three main classes of photoreceptors, blue (B)/UV-A-absorbing cryptochromes and phototropins and red (R)/far-red (FR)-absorbing phytochromes (Chen, Chory, and Fankhauser, 2004; Ulm and Nagy, 2005). The most studied and best-characterized group among plant photoreceptors is the R/FR light-absorbing phytochromes (Franklin and Quail, 2010; Neff, F ankhauser, and Chory, 2000; Schepens, Duek, and Fankhauser, 2004; Smith, 2000). Phylogenetic and genomic analyses have revealed that phytochromes and/or phytochrome-like chromoproteins are present not only in plants but also in cyanobacteria (Kehoe and Grossman, 1996; Rockwell and Lagarias, 2010; Yeh et a1., 1997), bacteria (Davis, Vener, and Vierstra, 1999; Hughes, 2010; Jiang et a1., 1999) and fungi (Blumenstein et a1., 2005; Hughes, 2010; Idnurm and Heitman, 2005). Existing evidence supports that photosynthetic organelles, chloroplasts, in plant cells evolved from a bilin sensor protein in the bacterial progenitor which in turn gave rise to phytochromes (Montgomery and Lagarias, 2002). Thus, the evolutionary origins of higher plant phytochromes are represented by prokaryotic genes (J iang et a1., 1999). Bacteriophytochromes (BphPs), cyanobacteria] phytochromes (Cphs) and fungal phytochromes (F phs) fall into distinct functional clades (Kamiol et a1., 2005) and phytochrome—like proteins, e.g., regulator of chromatic adaptation E (RcaE) and phytochrome-like protein A (PlpA), are present in some cyanobacteria] systems allowing the organisms to respond to predominant light conditions in the environment (Kehoe and Gutu, 2006; Montgomery, 2007). In Arabidopsis, phytochromes regulate a range of developmental and adaptive responses, such as seed germination, de-etiolation, gravitropic orientation, stomatal development, shade avoidance, entrainment of the circadian clock and flowering (Chen, Chory, and F ankhauser, 2004; Franklin and Quail, 2010; Mathews, 2006). Multiple BphPs could be present in bacteria, allowing them to regulate capacity of photosynthesis, movement towards or away from light and pigmentation process (Vierstra and Davis, 2000). F phA in Aspergillus nidulans regulates asexual sporulation in R light (Blumenstein et a1., 2005) and is potentially a phytochrome with many functional roles in regulating morphological and physiological differentiations, as well as tuning of asexual and sexual reproduction in response to light (Brandt et a1., 2008; Purschwitz et a1., 2008). A histidine kinase protein, Cphl in the cyanobacterium Synechocysris sp. strain PCC 6803, allows the organism to adapt to light-dark transitions (Garcia-Dominguez et a1., 2000). Photoreceptor RcaE in the cyanobacterium, F remyella diplosiphon, regulates the green-red photoreversibility of complementary chromatic adaptation and enables the organism to adapt to changes in ambient light in freshwater (Kehoe and Gutu, 2006). In Synechocystis sp. strain PCC 6803, PlpA regulates growth (Vierstra and Davis, 2000) and potentially has a role for regulating growth in B light (Wilde et a1., 1997). Despite the apparent diversity observed in photosensory and functional roles among the members of the phytochrome-class photosensors, all members appear to share a common photochemical mechanism for light sensing (Rockwell and Lagarias, 2010). Numerous responses/adaptations to light in organisms across kingdoms suggest that, from eubacteria to higher plants, informational light cues are utilized to maintain their growth, metabolism and developmental transitions in accordance with the environment. 1.2 Substructure and Biosynthesis of Phytochromes Phytochromes are soluble chromoproteins and consist of an apoprotein and a chromophore (Montgomery, 2009; Terry, Wahleithner, and Lagarias, 1993). A nuclear gene family encodes the apoprotein and the chromophore biosynthesis is localized in the plastid (Emborg et a1., 2006; Kohchi et a1., 2001). Thus, the synthesis of a holophytochrome involves coordination of two physically separated subcellular biosynthetic pathways (Montgomery, 2008). Multiple phytochrome species are present in plants and three distinct apoprotein encoding genes (PH YA-PH YC) are conserved within angiosperms (Mathews, Lavin, and Sharrock, 1995). The phytochrome family contains only three members (PH YA—PH Y C) in monocotyledonous plants (Takano et a1., 2005), whereas in dicotyledonous plants, additional apophytochrome-encoding genes have arisen due to recent gene duplication events (Mathews, Lavin, and Sharrock, 1995; Mathews and Sharrock, 1997). In the model plant Arabidopsis thaliana, a family of five nuclear genes, PH YA-PH YE encodes the apoproteins (F ankhauser and Staiger, 2002; Quail, 1994). The canonical photosensory core of phytochromes consists of an N- terminal sensory module of 3 distinct domains (Figure 1.1); Period/ARNT/Sim (Garcia- Dominguez et a1., 2000), cGMP phosphodiesterase/adenylate cyclase/FhIA (GAF) and phytochrome-specific (PHY; Hughes, 2010; Rockwell and Lagarias, 2010). Recent data suggest that phytochromes can dimerize to form homodimers and heterodimers (Clack et a1., 2009; Sharrock and Clack, 2004) and site-directed mutagenesis has revealed that the Per—Amt—Sim 2 (PASZ) domain is required for dimerization (Figure 1.]; Kim et a1., 2006). The C-terminal transmitter module consists of a dimerization/phosphoacceptor domain and an ATPase catalytic domain (Figure 1.1; Hughes, 2010). The bilin linear tetrapyrrole chromophore is covalently attached to the GAP domain (Figure 1.1; (Hughes, 2010; Rockwell and Lagarias, 2010) and the chromophore can either be phycocyanobilin (PCB), phytochromobilin (PCDB), or biliverdin lXa (BV) depending on the organism (Rockwell and Lagarias, 2010). All higher plant phytochromes utilize phytochromobilin as the chromophore, which is essential for phytochrome photoperception (Lagarias and Rapoport, 1980; Terry, Wahleithner, and Lagarias, 1993). The first committed step of plastid-localized chromophore biosynthesis is the oxidative cleavage of heme by multiple heme oxygenases to form biliverdin IXa (Figure 1.2; Emborg et a1., 2006). A ferredoxin-dependent phytochromobilin synthase further reduces biliverdin to (3Z)-phytochromobilin (Figure 1.2; Kohchi et a1., 2001). Phytochromobilin attaches to a conserved Cys residue in the GAP domain in each monomer via a thio-ether linkage (Figure 1.1; Franklin et a1., 2003; Kamiol et a1., 2005) through intrinsic autocatalytic activity of the apophytochrome molecule (Kamiol et a1., 2005). The phytochromes are R/FR reversible between two forms, the R-absorbing Pr (7t max~660 nm) and the F R-absorbing Pfr (2. max~730 nm). The biologically inactive form of phytochrome, Pr, is synthesized in the dark, and through photoconversion Pfr, the biologically active form is generated (Franklin and Quail, 2010). Photointerconvertibility between Pr and Pfr forms results in a dynamic photoequilibrium in natural light conditions (Franklin and Quail, 2010). The analysis of the three-dimensional structure of the ~ 20 kDa GAF domain fragment of SyB-Cphl phytochrome from Synechococcus OSB’ suggests that light-induced rotation of the A pyrrole ring determines photoconversion (Ulijasz et a1., 2010). However, a wealth of data still suggest that isomerization of the D ring is likely correct for other phytochromes (F odor, Lagarias, and Mathies, 1990; Kneip et a1., 1999; Rudiger et a1., 1983). 1.3 Phytochrome Activity at the Molecular Level The elucidation of phytochrome function at the molecular level has been the focus of plant photobiology for many years. Studies with PHYA-GFP and PHYB-GFP fusion proteins confirmed that upon conversion to the Pfr form, phytochromes display light- dependent nuclear translocation (Chen, C hory, and Fankhauser, 2004; Kircher et al., 1999; Nagatani, 2004; Yamaguchi et a1., 1999). However, for nuclear translocation of phyA, involvement of additional proteins, FHL and F HY, has been implicated recently (Hiltbrunner et a1., 2006). The best-studied mechanism of phytochrome signalling is the physical interaction of a nuclear-translocated photoreceptor with a class of bHLH transcription factors called PHYTOCHROME INTERACTING FACTORS (PIFs; Bauer et a1., 2004; Kim et a1., 2003; Monte et al., 2007; Shen et a1., 2007). Recent studies suggest that nuclear translocated phyB directly interacts with PIF 3 in a light-dependent manner, to regulate transcription of a number of genes with light responsive elements (Martinez-Garcia, Huq, and Quail, 2000; Monte et a1., 2004; Terzaghi and Cashmore, 1995). Moreover, extensive yeast two-hybrid screens confirmed that phytochrome kinase substrate 1 (PKSl; F ankhauser et a1., 1999) and nucleoside diphosphate kinase 2 (NDPK2; Choi et a1., 1999) are involved in direct physical interactions with some phytochrome isoforms. To date, numerous downstream signal transduction components have been identified (Kim et a1., 2003; Quail, 2000; Quail, 2002; Ryu et a1., 2005; Shen et a1., 2007), suggesting the existence of a transcriptional cascade that follows R/FR light perception by phytochromes. 1.4 Physiological Responses Regulated by Phytochromes Phytochromes perceive changes in R and FR light to modulate multiple physiological responses, from seed germination and seedling establishment through onset of reproductive development. Based on responses of seedlings to R and FR light, several classes of phytochrome responses have been characterized. Regulation of antagonistic and complementary functions by phytochrome family members has resulted through gene duplication and divergence (Mathews, 2010). The light-labile phyA molecule is the most abundant phytochrome in etiolated seedlings (Clough and Vierstra, 1997) and is the sensor for irreversible, very low fluence responses (VLF R) including absorption of FR light (Furuya and Schafer, 1996). Notably, in R light, nuclear-localized phyA has a role in irradiance-dependent photoprotection (Franklin, Allen, and Whitelam, 2007). Despite a wealth of data obtained through biochemical localization assays and injection of signaling intermediates (i.e. cyclicGMP, calmodulin and calcium; Nagy and Schafer, 2002), functions mediated by phytochromes remaining in the cytosol are poorly understood. However, for cytosolic phyA, a number of physiological functions have been reported (Rdsler, Klein, and Zeidler, 2007). phyB through phyE comprise the light—stable phytochrome molecules and phyB is predominantly responsible for the perception of R light and functions as a classic R/FR light reversible molecular switch regulating low fluence responses (LFR; Furuya and Schafer, 1996). Light-stable phytochromes collectively regulate end-of—day FR response and the shade-avoidance response in adult plants (Azari et a1., 2010). Irreversible high irradiance responses (HIR) are also regulated by phytochromes (Smith, 1995) with phyA mediating FR-H1R(Jiao, Lau, and Deng, 2007) and light-stable phytochromes mediating R-HIR (Azari et a1., 2010). The isolation of individual null mutants in all five phytochromes of Arabidopsis and construction of higher-order mutants has enabled thorough examination of responses regulated by phytochromes at seedling stage (Franklin et al., 2003; Franklin and Quail, 2010; Sullivan and Deng, 2003). Through transgenic approaches, molecular mechanisms underlying spatially localized pools of phytochromes in regulating distinct phytochrome- dependent responses have been elucidated (Endo et a1., 2007; Endo and Nagatani, 2008; Endo et a1., 2005; Montgomery, 2009; Wamasooriya and Montgomery, 2009). However, cell autonomous and cell non-autonomous signaling mediated by phytochromes during short-term physiological responses and long-term adaptive responses await further experimentation. 1.5 Tissue- and Organ-Specific Phytochrome Responses As phytochrome-mediated light responses are often localized to specific plant tissues or organs (Goosey, Palecanda, and Sharrock, 1997), the distribution of phytochromes at many developmental stages has been analyzed through multiple techniques, each having varying levels of sensitivities in detecting phytochromes in vivo or in vitro (Nagatani, 1997). Moreover, sites of light perception and sites of light action or sites displaying the physiological response do not always overlap in the plant, highlighting the significance of inter-cellular communication in plant growth and development (Bou-Torrent, Roig-Villanova, and Martinez-Garcia, 2008). Transmission of perceived light signals between tissues or organs that would result in grth and developmental changes at a site, distinct from the cells or tissue that received the light stimulus has been observed in planta and at the physiological level, the roles for spatial- specific phytochrome pools have been determined (Montgomery, 2008). Early physiological studies aimed at studying cell non-autonomous responses have utilized microbeam irradiation of plants or irradiation of detached plant parts to photoactivate phytochromes at a particular site and assay for a measurable phytochrome- dependent phenotypic change at a distant site (De Greef and Caubergs, 1972a; De Greef and Caubergs, 1972b; Mandoli and Briggs, 1982b). For example, in dark-grown bean seedlings, perception of light by leaves and the embryonic axis is required for leaf expansion (De Greef and Caubergs, 1972a). The onset of the transition from dark-grown to light-grown state (also known as de-etiolation) is marked by an increase in respiration. Cell layers in the epidermis and hypodermis of the epicotylar ribs displayed the highest ATP breakdown (De Greef and Caubergs, 1972b) suggesting the presence of epidermal cell layer responses in dark—grown bean seedlings. The perception of light in cotyledons is responsible for the inhibition of hypocotyl elongation and apical hook opening through intercellular signaling (Black and Shuttleworth, 1974; Caubergs and De Greef, 1975), as well as the pattern and level of accumulation of anthocyanins (Nick et a1., 1993). The most well-studied cell non-autonomous phytochrome response is the photoperiodic induction of flowering in photoperiod-sensitive plant species. In these plants, floral initiation involves perception of light signals in the leaves and the subsequent transport of a photo-induced stimulus from the leaves to the shoot apex through the phloem (Parcy, 2005; Zeevaart, 2006). Through photoreceptor mutant analyses, it is known that phyA and cryptochrome 2 (cry2) are involved in perceiving long-day photoperiods and thus control flowering (Franklin and Quail, 2010). Although promoter-fusion studies indicate that these photoreceptors are ubiquitously expressed throughout the seedlings (Toth et a1., 2001), a wealth of prior studies has confirmed that light perception in leaves is associated with the photoperiodic induction of flowering (Zeevaart, 2006). In dark-grown oat seedlings, photoperception within the mesocotyl mediates its own development. In contrast to the coleoptilar node and tip where phytochromes are most abundant, the sites above and below the coleoptilar node regulate promotion of coleoptile elongation through light piping (Mandoli and Briggs, 1982a). Thus, the sites of photoperception may or may not coincide with the sites of highest phytochrome abundance. Studies with microbeam irradiation focused on phytochrome responses at a distant site and cabxsluciferase reporter gene expression experiments have shown luciferase activity in non-irradiated tissues distant from the few irradiated cells in dark-grown IO Arabidopsis cotyledons (Bischoff et a1., 1997). Results from these studies validate the presence of cell- and tissue-specific phytochrome signaling pathways that regulate distinct aspects of light—dependent growth and development through intercellular and/or interorgan coordination. However, conclusive evidence from many of these early physiological studies that utilized excised plant organs and microbeam irradiation, were confounded due to light piping and light scattering (Mandoli and Briggs, 1982b). 1.6 Probing Spatial-specific Responses via Targeted Chromophore Reduction The isolation of various mutants in ArabidOpsis enabled phenomenal progress in light signaling research and the elucidation of the functional roles of phytochromes. Extensive analyses of individual and multiple apophytochrome mutants have uncovered the existence of unique and overlapping photoregulatory roles among the five types of phytochromes (Franklin and Whitelam, 2004). Chromophore biosynthetic mutants, hyl and hy2, exhibit multiple phytochrome-deficient phenotypes throughout their life cycle due to an absence of all five phytochromes (Hudson, 2000); their study has contributed to our understanding of photoregulatory functions at a global level. However, mutations in a number of genes that encode for phytochrome-interacting proteins have resulted in distinct phenotypes in different tissues within the seedling (Neff, F ankhauser, and Chory, 2000). Therefore, each localized pool of phytochromes may control only a subset of light-mediated responses in planta. Definitive information on how these localized pools of phytochromes mediate distinct aspects of plant growth and development is currently limited. ll A molecular tool that can render all five types of phytochromes non-functional is essential for probing tissue- and organ~specific phytochrome responses. One tool currently utilized is the expression of a gene encoding a rat kidney enzyme, biliverdin IXa reductase (BVR, Figure 1.3). BVR converts biliverdin IXa, a precursor of phytochrome-chromophore biosynthesis, into bilirubin and metabolizes the chromophore, phytochromobilin to phytochromorubin (Figure 1.3; Terry, Wahleithner, and Lagarias, 1993). The end products of the BVR enzyme cannot covalently assemble with apophytochromes (Terry, Wahleithner, and Lagarias, 1993). Constitutive expression of BVR in Arabidopsis has led to the perturbation of many light-dependent phytochrome- mediated responses (Lagarias et a1., 1997; Montgomery et a1., 1999). Moreover, depending on the subcellular localization of reduced levels of the phytochrome chromophore through BVR expression, light-dependent, as well as light-independent phenotypes have been observed in Arabidopsis and Tobacco (Nicotiana tabacum cv Maryland Mammoth; Montgomery et a1., 2001; Montgomery et a1., 1999). Such BVR- dependent phenotypes are very similar to responses displayed by an Arabidopsis phytochrome chromophore-deficient mutant, hyI (Montgomery et a1., 1999) and indicate loss of photosensory activities of multiple phytochromes. Thus, expression of B VR is responsible for altered phytochrome-mediated responses due to reduction of holophytochrome levels (Lagarias et a1., 1997). Transgenic approaches such as over-expression, ectopic expression and mis- expression of transgenes, are useful for the analysis of gene function (Rutherford et a1., 2005; Wamasooriya and Montgomery, in press). With tissue- and organ-specific promoters, the expression of BVR can be restricted to a distinct subset of cells, tissues or organs. Spatial-specific regulation of transgene expression has been successfully utilized in many studies that were aimed at further understanding the photoreceptor functions during the life cycle of Arabidopsis (Wamasooriya and Montgomery, in press). One such example is transgenic Arabidopsis lines in which the cry2—GFP fusion was expressed under the control of mesophyll-specific, vascular bundle-specific and epidermal-specific promoters in a cry2-deficient mutant background. Only cry2-GF P accumulation or expression in vascular bundles was able to rescue the late flowering phenotype of the cry2 mutant suggesting that the site of cry2 photoperception that regulates flowering is the vascular bundles (Endo et a1., 2007). Studies with stable transgenic lines displaying mesophyll-specific and meristem- specific phytochrome chromophore deficiencies have revealed that localized pools of phytochromes can regulate distinct physiological responses and established the efficacy of targeted BVR expression as a novel molecular technique to investigate sites of light perception (Wamasooriya and Montgomery, 2009). A current limitation is the availability of cloned and characterized promoters that can direct the gene expression in a targeted manner. The limited number of well-characterized promoters and corresponding expression patterns restricts the cell types and developmental processes that can be targeted (Wamasooriya and Montgomery, in press). A two-component, enhancer-trap mis-expression system based on the yeast GAL4 transcription factor has been used successfully to study regulatory mechanisms during embryonic development in Drosophila (Brand and Perrimon, 1993), root development in Arabidopsis (Laplaze et a1., 2005), and improving salinity tolerance in rice (Plett et al., 2010). The bipartite enhancer-trap strategy results in transactivation of the expression of 13 a gene under control of the Upstream Activation Sequence (UAS) element by a transcriptional activator (Laplaze et a1., 2005). A more stringent regulatory system based on promoter/enhancer trap activation can be achieved with transcription factors with sequence-specific DNA-binding activities that are not normally found in plants (Moore et a1., 1998). Transgenic enhancer trap lines are T-DNA insertion lines with diverse expression patterns of the yeast transcription factor, GAL4, whose expression depends on the presence of native genomic enhancer sequences. The GAL4- responsive mGFP5 gene marks the expression pattern mediated by genomic enhancers in green fluorescence (Haseloff, 1999; Laplaze et a1., 2005). Two-component transactivation systems overcome the limited availability of cloned and characterized promoters by using native genomic enhancers within a host genome (Wu et a1., 2003) and circumvent the necessity to maintain and genotype multiple stable transgenic lines, which is laborious and labor intensive (Wamasooriya and Montgomery, in press). Another advantage of GAL4/U AS two-component enhancer trap system is that distinct expression patterns can be achieved in a localized manner, which makes it an effective tool to determine sites of physiological process such as photoperception that have been shown to have spatial-specific aspects. Use of the Cauliflower mosaic virus (CaMV) 355 minimal promoter-based enhancer-trap system to express PH YB-GFP in a phytochrome B (phyB) mutant of Arabidopsis revealed that PH YB-GF P expression in the mesophyll but not in vascular bundles suppresses the expression of a key flowering regulator, FLOWERING LOC US T(F7) in vascular bundles of cotyledons (Endo et a1., 2005). This finding indicates that a novel inter-tissue signaling mechanism occurs between mesophyll and vascular bundles making it a critical step in the regulation of flowering by phyB (Endo et a1., 2005). I4 Probing spatial-specific phytochrome responses necessitates reduction of holophytochromes in a selective manner. Enhancer trap-based two-component strategy requires two transgenic parents: the enhancer trap and the UAS-BVR. Genetic crosses between the UAS-BVR parent and a range of GAL4-enhancer trap parents will result in progeny with diverse expression patterns of the BVR gene. The sites of mGF P5 expression should be identical to the sites of BVR expression in the progeny. Selective mis-expression of BVR through an enhancer trap system will result in site-specific reduction of chromophore availability, inducing local phytochrome-deficiencies within the plant and the functions of localized phytochrome pools can be probed to characterize the sites of photoperception. Comparative analyses of such progeny will expand our current understanding of phytochrome-regulated tissue- and organ-specific responses and possibly gain insight into molecular bases of novel inter-tissue and/or inter-organ signaling mechanisms mediated by phytochromes in planta. 1.7 Summary Plants are sessile organisms solely dependent on light for photosynthesis and thus need to have photosensory mechanisms to adapt to sub-optimal light conditions in the surrounding environment. Photoreceptors render plants the capacity to detect and respond to frequent fluctuations in many components of light. Among the very well characterized plant photosensory systems, phytochromes regulate a vast number of growth and developmental processes in Arabidopsis. Unparalleled progress has been made in determining the physiological roles and dissecting the phytochrome-signaling cascade. Despite the wealth of experimental findings on tissue- and organ-specific responses at 15 physiological and molecular level, definitive molecular evidence about sites of photoperception, cellular signaling mechanisms of specific pools of phytochromes is yet limited. The identification of the molecular bases of tissue- and organ-specific phytochrome responses and their downstream molecular effectors will broaden our current understanding of the complex signal transduction network in Arabidopsis. Molecular evidence on mechanisms by which phytochromes or different phytochrome isoforms regulate responses at distinct sites in the plant will aid in fine-tuned manipulation of agronomically desirable physiological responses in important crop species. 16 .88 £08.85 Scum ceases we; oSwE .EsEoc >=E£Sazm Beak omniv— oEEHmE .ommmhxximf ”BE—8-59:3 < omens. 2:253 JOE: ”83:3 2:252 mo 0558.830 £85 new 2668 m5 couaeoomosamoca a memo—02 EmEoc cows—8-08:2 3%sz Ravi ”couentofimc com Ems—ow N Swizz/Thom .m38me Ewes—o m SIA— wEmEoc ”20 of £53» 2.2%: Amxov 0:658 @3338 m 8 8:083 .E—EoEoEooSa—Q dug—Q0885 080508.03 of .mem mmR holophytochrome apophytochrome mRNA . nucleus . Figure 1.3 Inactivation of holophytochrome through expression of BVR. Plastid-targeted biliverdin reductase (BVR) converts a precursor of the phytochromobilin chromophore (PCDB), biliverdin IXa (BV), into bilirubin (BR) or PCDB into phytochromorubin (PCDR). BVR expressed in the cytosol metabolizes PCDB into PCDR. 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Detection of spatial-specific phytochrome responses using targeted expression of biliverdin reductase in Arabidopsis. Plant Physiol 149(1), 424-433. Wamasooriya, S. N., and Montgomery, B. L. (in press). Using Transgenic Modulation of Protein Accumulation to Probe Protein Signaling Networks in Arabidopsis thaliana. Plant Biotechnology and Transgenic Research, Bentham Science Publishers, Oak Park, IL. Wilde, A., Churin, Y., Schubert, H., and Bomer, T. (1997). Disruption of a Synechocystis sp. PCC 6803 gene with partial similarity to phytochrome genes alters growth under changing light qualities. F EBS Lett 406(1-2), 89-92. Wu, C., Li, X., Yuan, W., Chen, G., Kilian, A., Li, J., Xu, C., Li, X., Zhou, D. X., Wang. S., and Zhang, Q. (2003). Development of enhancer trap lines for functional analysis of the rice genome. Plant J 35(3), 418-427. Yamaguchi, R., Nakamura, M., Mochizuki, N., Kay, S. A., and Nagatani, A. (1999). Light-dependent translocation of a phytochrome B-GFP fusion protein to the nucleus in transgenic Arabidopsis. J Cell Biol 145(3), 437-445. 27 Yeh, K. C., Wu, S. H., Murphy, J. T., and Lagarias, J. C. (1997). A cyanobacterial phytochrome two-component light sensory system. Science 277(5331), 1505- 1508. Zeevaart, J. A. (2006). Florigen coming of age after 70 years. Plant Cell 18(8), 1783- 1789. 28 Chapter 2 Regulation of Far-red Light-Mediated High Irradiance Responses by Spatial-specific Phytochromes Some of the work included in chapter 2 was published in the journal, Plant Physiology. Wamasooriya SN and Montgomery BL, (2009), Detection of Spatial-Specific Phytochrome Responses Using Targeted Expression of Biliverdin Reductase in Arabidopsis, Plant Physiology, 149(1), 424-433. Some of the work in chapter 2 is from a manuscript submitted to the journal, Plant, Cell and Physiology. Wamasooriya SN, Porter KJ and Montgomery BL, Tissue- and lsoform-Specific Phytochrome Regulation of Light-Dependent Anthocyanin Accumulation in Arabidopsis thaliana (submitted). 29 2.1 Overview Phytochromes, in response to R and FR light, regulate numerous and distinct aspects of light-mediated growth and development (Chen, Chory, and F ankhauser, 2004; Franklin and Quail, 2010; Mathews, 2006). The regulation of these physiological responses encompasses complex intra- as well as, inter-cellular signaling cascades (Montgomery, 2008). A number of studies have confirmed that phytochromes mediate cell- and tissue-specific signaling pathways in controlling discrete aspects of light- dependent growth and development through intercellular and/or inter-organ coordination (Black and Shuttleworth, 1974; Caubergs and De Greef, 1975; De Greef and Caubergs, 1972a; De Greef and Caubergs, 1972b; Mandoli and Briggs, 1982; Nick et a1., 1993). Expression of distinct fractions of the Arabidopsis genome in different organs or tissue types in response to developmental and environmental cues results in spatial- specific responses. Discrete spatial-specific light responses are likely to be impacted by several factors. Light sensitivity of cells or organs is influenced by their physiological differences; i.e. pigmentation due to photosynthetic activity, and metabolic status can affect light sensitivity of a particular tissue (Bou-Torrent, Roig-Villanova, and Martinez- Garcia, 2008). Although spatial expression analyses have concluded that phytochromes are found in all tissues analyzed, even in roots, (Clack, Mathews, and Sharrock, 1994; Nagatani, 1997; Toth et a1., 2001) and thus, are R- and FR-sensitive, in a given tissue or organ, phytochromes and interacting partners downstream of R/FR light perception might not be uniformly distributed (Bou-Torrent, Roig-Villanova, and Martinez-Garcia, 2008). In fact, several concerted efforts related to tissue- and organ-specific expression profiling have confirmed that each organ or tissue type has a well defined genome expression 30 pattern (J iao, Lau, and Deng, 2007; J iao et a1., 2005; Ma et a1., 2005) and spatial-specific involvement of signaling cascades in different organs and cell types in response to divergent light effects. Thus, most spatial-specific expression patterns of early targets of phytochrome signaling likely define the cellular mechanisms of localized pools of phytochromes that eventually result in different, even opposing, intercellular and/or organ-specific phytochrome responses to the same light stimulus (Bou-Torrent, Roig- Villanova, and Martinez-Garcia, 2008). 2.1.1 Phytochrome-dependent High Irradiance Responses Plants are exposed to prolonged periods of high intensity light irradiation in natural environments. HIRs are characterized as physiological responses that require relatively high photon fluence rates over a long duration with strong irradiance dependence and a lack of photoreversibility (Caubergs and De Greef, 1975; Deng and Quail, 1999; Neff, F ankhauser, and Chory, 2000). HIRs that have been examined extensively are light-mediated inhibition of hypocotyl elongation and anthocyanin production in planta (De Greef and Caubergs, 1972a; De Greef and Caubergs, 1972b). B- , R- and F R-mediated HIRs were first identified through the analysis of action spectra for anthocyanin formation under prolonged irradiation in cabbage and turnip (De Greef and Caubergs, 1972a) and Mohr (1972) later speculated that Hle with action spectra having a peak in the FR light region were FR-HIR, and the possible involvement of phytochrome in the regulation of F R-HIR. Recently, Shinomura, Uchida, and F uruya, (2000) reported that a transitory signal generated during photoconversion from Pfr to Pr form of 31 phytochrome is required for F R-HIR in contrast to the more stable phyB-produced signal in R-HIR (Ahmad and Cashmore, 1997; Caubergs and De Greef, 1975). Typical HIRs include inhibition of seed germination, hypocotyl elongation, and opening of the apical hook, expansion of the cotyledons, accumulation of anthocyanin and a FR-light preconditioned block of greening during seedling development (Neff, F ankhauser, and Chory, 2000). Germination and growth of Arabidopsis seedlings in continuous FR light (F Re) can induce partial photomorphogenesis without the accumulation of chlorophyll. Under these conditions, although seedlings are chlorophyll deficient, hypocotyl elongation is inhibited and the cotyledons are expanded (Whitelam et a1., 1993). phyA and phyB have recognized roles in mediating HIR of light-dependent inhibition of hypocotyl elongation in Arabidopsis under FR and R light, respectively (Quail et a1., 1995). 2.1.2 Light Perception by Phytochrome A phyA is unique among the other phytochrome family members due to a number of characteristics in its stability and light perception. In contrast to light-stable phyB-phyE, phyA is light-labile and is responsible for the VLF R and F R—HIR (Nagy and Schafer, 2002; Sharrock and Clack, 2002; Wang and Deng, 2003). phyA is relatively stable in its Pr form and is rapidly degraded upon conversion to the Pfr form (Clough and Vierstra, 1997). Since phyA accumulates to high levels in etiolated seedlings, the primary function of phyA is to promote seed germination during early stages of de-etiolation (Mohr, 1972). However, phyA that exists in de-etiolated tissues is known to regulate numerous responses in the life cycle of light-grown seedlings, namely, inhibition of stem elongation 32 growth during shade avoidance (Yanovsky, Casal, and Whitelam, 1995), light input to the endogenous clock (Johnson et a1., 1994), inhibition of intemode elongation (Devlin, Patel, and Whitelam, 1998) and regulation of leaf expansion (Franklin et a1., 2003). The observation that inhibition of hypocotyl elongation or anthocyanin accumulation can be partially reversed if the B light pulses are followed by saturating F R-light pulses suggested that phytochromes are involved in B light perception (Casal and Boccalandro, I995; Mancinelli, Rossi, and Moroni, 1991). Indeed, in later studies, phyA has been shown to have a role in regulating responses under B illumination, i.e. chloroplast gene expression during leaf development, hypocotyl inhibition and anthocyanin accumulation (Chun, Kawakami, and Christopher, 2001; Duck and Fankhauser, 2003; Neff and Chory, I998; Poppe et a1., 1998; Whitelam et a1., 1993; Yadav et a1., 2005). Analysis of monogenic and high order mutants in apophytochrome genes has revealed distinct functional roles for phyA during de-etiolation of Arabidopsis seedlings in FR light. phyA mutants have long hypocotyls and closed cotyledons as opposed to short hypocotyls and expanded cotyledons in WT seedlings under FR light (Nagatani, Reed, and Chory, 1993; Parks and Quail, 1993; Whitelam et a1., 1993). These studies confirmed that phyA is an FR sensor and regulates seedling de-etiolation. Single mutants of Arabidopsis deficient in phyB-phyE resembled WT plants in F Rc (Aukennan et al., 1997; Reed et a1., 1994) and thus confirm that phyA solely mediates VLFR and F R-HIR (Wang and Deng, 2003) in FR light. Until recently, phyA was thought to be neither necessary nor sufficient for R light perception (Quail et a1., 1995) although several studies with phyAphyB double mutant and phyB mutant in R light (< 50 umol m-2 8") indicated that phyA might potentially have a role in regulating hypocotyl inhibition, hook 33 opening and expansion of cotyledons (Neff and Chory, 1998; Reed et a1., 1994). Franklin, Allen, and Whitelam, (2007) reported that phyA is able to regulate de-etiolation and possesses increased stability, as nuclear phyA is subject to irradiance-dependent photoprotection under high intensity R light (> 160 umol rn-2 5.1). Results from this study revealed the significance of nuclear phyA being an irradiance-dependent R light sensor as plants are exposed to high intensities of light in natural environments. 2.1.3 Phytochrome A and Downstream Components in Signaling phyA-mediated signaling, based on light-dependent nuclear transport and differential stability, involves compartmentalization, transcriptional regulation and differential degradation (Hiltbrunner et a1., 2006; Rdsler, Klein, and Zeidler, 2007; Wang and Deng, 2003; Yang et a1., 2009; Yanovsky et a1., 2002). Since phyA is solely responsible for regulating responses in F R light conditions (Nagatani, Reed, and Chory, 1993; Parks and Quail, 1993; Whitelam et a1., 1993), the approach for identifying the downstream candidates of phyA signaling has been to assess for the disruption of phytochrome-mediated responses in FR in mutants. Based on elongated hypocotyls and unexpanded cotyledons under FR light, more than 15 mutants have been isolated and characterized that encode signal transduction components in the phyA-mediated signaling cascade (Wang and Deng, 2003; Yang et a1., 2009). Many of the signal transduction components appear to be transcription factors with which nuclear-localized phyA interacts to mediate discrete responses (Ballesteros et a1., 2001; Duek and Fankhauser, 2003; F airchild, Schumaker, and Quail, 2000; Ni, Tepperman, and Quail, 1998; Rosler, Klein, and Zeidler, 2007). Moreover, since VLF Rs saturate upon activation of phyA with 34 short pulses of very low fluence R or FR light, while HIRs require sustained activation of phyA with higher fluences of FR light, VLF R- and HIR-mediated signaling can be dissected genetically at certain loci that function downstream of phyA in the signaling pathway (Luccioni et a1., 2002; Yanovsky, Casal, and Luppi, 1997; Yanovsky, Whitelam, and Casal, 2000). This suggests that signaling downstream of light perception by phyA may branch out into two individual cascades, depending on the mode in which phyA is activated. Distinct physiological roles have been attributed to nuclear-localized phyA. F HL- and FHY-regulated nuclear transport of phyA allows interaction of phyA with transcription factors (Hiltbrunner et a1., 2006; Hiltbrunner et a1., 2005). In fact, phyA has been identified to interact with LAFI and HFR in regulating inhibition of hypocotyl elongation under FR light (Yang et a1., 2009). Recently, an analysis of the fhl/fhyl double mutant has shown that cytoplasmic phyA is able to regulate R light-enhanced phototropism, abrogation of gravitropism, and inhibition of hypocotyl elongation in B light (Rbsler, Klein, and Zeidler, 2007). This raises the possibility that nuclear- and cytoplasmic-localized pools of phyA may engage different downstream interacting partners to regulate discrete physiological responses. Moreover, the analysis of gene expression profiles in roots, hypocotyls and cotyledons of Arabidopsis seedlings (Ma et a1., 2005) has indicated that the expression patterns of PH YA, PH YB, several PIF3 and a number of early targets of phytochrome signaling action during both de-etiolation and shade avoidance response (i.e. AT HB2, AT HB4, HA T2, PAR l, PILI, RIP; Roig-Villanova et a1., 2006) vary in response to light in the three organs. Thus, upon light perception by 'nuclear- or cytoplasmic-localized phyA, the transcriptional network might immediately 35 diverge in a cell- and organ-specific manner, thereby increasing the complexity of phyA- mediated signaling. Although, a significant number of downstream molecular effectors have been identified in the phyA signaling pathway, identification of target genes involved in spatial-specific phyA-mediated signaling requires further analysis. 2.1.4 Outlook Although much progress has been made in understanding spatial-specific light perception by phytochromes and the subsequent downstream signaling network, definitive molecular evidence about distinct sites of phytochrome photoperception and cellular mechanisms of pools of phytochromes in regulating tissue- and organ-specific phytochrome responses in Arabidopsis is limited. The importance of regulation of such spatial-specific responses is beginning to be recognized, however, much work remains to be accomplished to identify these responses and elucidate them fully at the molecular level (Bou-Torrent, Roig-Villanova, and Martinez-Garcia, 2008; Montgomery, 2008). Knowledge of spatial-specific phytochrome responses to the changing light environment and their underlying molecular bases will provide groundwork for dissecting the complex intercellular and inter-organ molecular processes underlying R- and FR-dependent plant growth and development. The knowledge base generated at the molecular level will help to modify light responses in specific tissues or organs, reducing the detrimental and pleiotropic effects observed when phytochrome action is manipulated in the whole plant. A molecular tool that is being currently utilized for phytochrome inactivation is the expression of a gene encoding a rat kidney enzyme, biliverdin IXa reductase (B VR). Constitutive expression of BVR in Arabidopsis has led to the perturbation of many light- 36 dependent phytochrome-mediated responses (Lagarias et a1., 1997; Montgomery et a1., 1999). Such BVR-dependent phenotypes are very similar to responses displayed by an Arabidopsis phytochrome chromophore-deficient mutant, hyI (Montgomery et a1., 1999) and indicate a loss of photosensory activities of multiple phytochromes. In this chapter, two tissue-specific promoters (i.e. CAB3 and MERIS) were utilized to drive the expression of the BVR in mesophyll cells and shoot meristamatic tissues, respectively. Studies with stable transgenic lines displaying mesophyll- and meristem-specific phytochrome chromophore deficiencies revealed that localized pools of phytochromes can regulate distinct physiological responses and established the efficacy of targeted BVR expression as a novel molecular technique to investigate sites of light perception (Wamasooriya and Montgomery, 2009). 2.2 Materials and Methods 2.2.] Photomorphogenesis of Transgenic Lines with Targeted Chromophore Inactivation in FRc Seeds of No-O WT, 35S::pBVR3, CAB3::pBVR2, Col-0 WT and phyA (SALK_ 014575; Ruckle, DeMarco, and Larkin, 2007) were sterilized with 35 % (v/v) commercial bleach and 0.025 % (v/v) SDS solution and were rinsed 6 times with ultrapure water (Milli-Q, Millipore, MA) according to the protocol published in (Wamasooriya and Montgomery, 2009). Sterilized seeds were planted in 100- x 25-mm petri dishes on media containing 1X Murashige and Skoog salts (Catalog No. MSP09, Caisson Laboratories, UT), 0.9 % (w/v) Phytablend (Catalog No. PTPOI, Caisson Laboratories, UT) with 1 % (w/v) Sucrose (Catalog No. 4072-05, J .T. Baker, NJ), adjusted to pH 5.7 with KOH. 37 Imbibing seeds were cold-stratified at 4 °C for 3 days in darkness. Plates were transferred to a temperature- and humidity-controlled growth chamber with F Rc . . . -2 -l . . . . illumination of 5 umol m s . Seedling images were scanned at high resolution to show characteristic phenotypes observed in wild-type and transgenic Arabidopsis seedlings during photomorphogenesis in FR light. 2.2.2 Quantification of Anthocyanin Levels in Transgenic Lines with Targeted Chromophore Inactivation To determine whether mesophyll- and meristem-localized phytochrome deficiencies distinctly affected sucrose-stimulated anthocyanin accumulation, anthocyanin levels were quantified in 4-d-old seedlings grown under F Re and Be light. Seeds of No-0 WT, 358::pBVR3, CAB3::pBVR2, Col-0 WT and phyA (SALK_ 014575; Ruckle, DeMarco, and Larkin, 2007) were sterilized and planted as described in section 2.2.1. Sterilized seeds were treated with R pulse (approximately 75 umol m.2 s") for 5 min prior to imbibition to synchronize germination. Imbibing seeds were cold-stratified at 4 °C for 3 days in darkness. Plates were transferred to a temperature- and humidity- controlled growth chamber with Bc illumination of 25 umol m-2 s, FRc illumination of 5 umol m-2 s", or in darkness for 4 days at 22 °C. Anthocyanins were extracted from whole-plant seedlings using I 0/o (v/v) HCI (Catalog No. HX0603-l3, EMD Chemicals Inc., NJ) in methanol (Catalog No. 9070-03, J .T. Baker, NJ) as described previously (Feinbaum and Ausubel, 1988) with the following modifications. The number and weight of fresh whole-plant seedlings were recorded before transferring the harvested seedlings 38 into glass vials. Anthocyanins were extracted in l ”/0 (v/v) HCl in methanol at a ratio of 20 uL/mg of fresh tissue overnight (~ 16 hr) with gentle shaking at 4 °C (adapted from Rabino and Mancinelli, (1986). Chloroform (Catalog No. 9180-01, J .T. Baker, NJ) was added at a ratio of 20 uL/mg of fresh tissue. Chloroform-water partitioning was performed by adding sterile H20 at 0.4 volume of 1 % HCI in methanol and chloroform volumes, followed by centrifugation at 13,000 rpm at room temperature for 5 min (Denville 260D, NJ; adapted from Kerckhoffs et a1., (1997). Anthocyanin content was estimated per seedling by measuring the A535 minus the A650 of the aqueous phase spectrophotometrically (Montgomery et a1., 1999). Two-tailed, unpaired Student’s t-test was performed to compare anthocyanin contents relative cognate WT seedlings. 2.2.3 Gene Expression Analysis To identify candidate downstream target genes in regulating FR-dependent hypocotyl development, gene expression analysis was performed as follows. Seeds of No-O WT, 35S::pBVR3 and CAB3::pBVR2 were sterilized as described in section 2.2.1. Seeds were planted in 150-x 15-mm petri dishes on media prepared according to section 2.2.1. To synchronize germination, plates with seeds were exposed to R light of 75 umol m.2 s.1 for 5 min and imbibing seeds were cold-stratified at 4 °C in darkness for 3 days. 7-d-old whole seedlings were quickly (< 1 min) harvested and immediately frozen in liquid nitrogen inside the FR chamber. Total RNA was isolated using RNeasy® Plant Minikit (Catalog No. 16419, Qiagen, CA) according to manufacturer’s instructions. After assessing the quality of isolated RNA on the Agilent 2100 Bioanalyzer (Agilent 39 Technologies, Inc., CA), 500 ng of total RNA were subjected to the synthesis of TM amplified RNA (aRNA) using the MessageAmp Premier RNA Amplification Kit (Catalog No. 4385821, Applied Biosystems/Ambion, TX) according to manufacturer’s instructions with the following modifications. For in vitro transcription reaction, samples were incubated for 8 hrs at 40 °C. Binding of aRNA to magnetic beads following addition of 100 % ethanol was carried out for 5 min with gentle shaking. To capture the magnetic beads-aRNA complex, the U bottom plate was held on the magnetic stand for ~ 6 min. aRNA was eluted off of magnetic beads through vigorous shaking for ~ 7 min. The quality of biotin-labeled aRNA was determined on the Agilent 2100 Bioanalyzer (Agilent Technologies, Inc., CA). 1 1-12 ug of labeled aRNA were submitted to the Research Technology Support Facility at Michigan State University for hybridization with the GeneChip® Arabidopsis ATHI Genome Array (Catalog No. 900385, Affymetrix, Inc., CA) and acquisition of scanned probe arrays. Each labeled aRNA was hybridized to an individual ATHI Genome Array and the microarray analysis was conducted with three independent RNA extractions per sample. The expression data were subjected to per chip normalization (shift to the 75th percentile) with baseline transformation to the median of all samples using GeneSpring GX 10.0 (Agilent Technologies Inc., CA). The data were then filtered on flags (present or marginal in at least 1 out of the 9 samples). Analysis of variance on log-transformed expression values was carried out by applying the Benjamini and Hochberg multiple testing correction with a p-value cut off of 0.05 across the three groups (No-0 WT, 35S::pBVR3 and CAB3::pBVR2) to control the false discovery rate. An expression filter of fold change 40 greater or equal to 2.0 was applied for 358::pBVR3 and CAB3::pBVR2 samples against the No-O WT sample. Expression data of CAB3::pBVR2 was compared against 358::pBVR3 to identify genes with changes in gene expression that are unique to hypocotyl development observed in CAB3::pBVR2 in FR light. Based on the differential expression pattern observed in CAB3::pBVR2 relative to 358::pBVR3, three genes were selected to prioritize initial analysis: At1g26220, A t4g02290 and At1g52410. The expression of At] g26220, At4g02290 and Atlg52410 genes were analyzed using publicly available gene expression data. Mean-normalized logz values from AtGenExpress and BAR Heatmapper Plus (http://www.bar.utoronto.ca) were used to generate heat maps for Atlg26220, At4g02290 and At1g52410 genes. 2.2.4 Validation of At1g26220, At4g02290 and Atlg52410 Expression in Transgenic Lines with Targeted Chromophore Inactivation by RT-PCR To confirm the respective fold-reduction in the expression levels of At1g26220, A t4g02290 and At1g52410 observed in CAB3::pBVR2 through microarray analysis, total RNA isolated from No-O WT, 358::pBVR3 and CAB3::pBVR2 in section 2.2.3 was used in RT-PC R. The quantity of RNA for each sample was analyzed by spectrometry (NanoDroplOOO, Thermo Scientific, MA). Oligo(dT)15 primed-first-strand cDNA was synthesized from 1 pg of total RNA using a Reverse Transcription System (Catalog No. A3500, Promega, WI) according to manufacturer’s instructions with the following modifications: RT reactions were incubated at 42 °C for 1 hr followed by 95 °C for 5 min and 4 0C for 5 min. cDNA was stored at -20 °C overnight before PC R amplification. First-strand cDNA synthesis reactions were diluted 1:16 with nuclease-free water. PC R 41 was conducted with GoTaqGreen (Catalog No. M7123, Promega, WI) and 4 ul of the diluted cDNA product was used as template in a 25 ul reaction. The gene-specific oligonucleotides for At1g26220 and Atlg52410 were designed based on the full length cDNA sequence: for At1g26220- forward 5 ’- AGGC AC AATCTCCACTCCTCCAGC-3 ’ and reverse 5’- CGGGTCAGAGACGAACCCAAGCG-3 ’, for Atlg52410- forward 5’- GACAATAACGAAGAAGAACACGCTGC -3’ and reverse 5’- AATGAGAATCTGACCGAAATCTTTACG -3 ’. The gene-specific oligonucleotides for At4g02290 were designed using a free online database, AtRT Primer (Han and Kim, 2006); forward 5’- TCTI‘CTTCCTCCTCCTATGCCCTCA -3 ’ and reverse 5’- TGCAAAAACACTGATGGCTCGTTT -3 ’. PCR amplification was carried out with gene-specific oligonucleotides at 10 11M. The following thermal cycling conditions were used for the PCR amplification: for At1g26220, (l) 1 cycle of denaturation at 94 °C for 2 min, (2) 27 cycles of denaturation at 94 °C for 30 s, annealing at 56 0C for 45 5, extension at 72 °C for 38 s and (3) final extension at 72 °C for 5 min with a hold at 4 °C. For At4g02290, (1) 1 cycle of denaturation at 94 °C for 2 min, (2) 28 cycles of denaturation at 94 °C for 30 s, annealing at 56 0C for 45 5, extension at 72 °C for 50 s and (3) final extension at 72 °C for 5 min with a hold at 4 °C. For At1g52410, (l) 1 cycle of denaturation at 94 °C for 2 min, (2) 27 cycles of denaturation at 94 °C for 30 s, annealing at 56 °C for 45 3, extension at 72 °C for 1 min 10 s and (3) final extension at 72 °C for 5 min with a hold at 4 °C. Expression of UBC2] (At5g25 760) was analyzed as an internal control by including UBC21-specific primers, forward 5’- 42 CCTTACGAAGGCGGTGTTTI‘TCAGJ’ and reverse 5’- CGGCGAGGCGTGTATACATTTG-3’ at 10 uM in the same reaction. A 7 uL aliquot of the PCR product was visualized by electrophoresis for 1 hr 30 min at 85V on a 1.5 % agarose gel containing ethidium bromide at 0.02 ug/mL (Catalog No. 15585-01 l, Invitrogen, CA). Ultraviolet images were obtained using the Gel Doc system (Bio-Rad Laboratories, Inc., CA) at subsaturation settings. 2.2.5 Confirmation of T-DNA Insertion Mutants SALK_062388, SALK_101567 and SALK_151393 mutants with a T-DNA insertion in an exon of At1g26220, At4g02290 and At1g52410 respectively, were selected from the Salk T-DNA insertion mutant collection (Alonso ct a1., 2003) and were genotyped by PCR. Seeds of Col-0 WT and homozygous SALK_062388, SALK_101567 and SALK_151393 mutants were sterilized, planted and subjected to cold stratification as described in section 2.2.1. Plates were transferred to a temperature- and humidity- controllcd growth chamber with FRc illumination of 5 umol m.2 s.I for 7 days at 22 °C. Total RNA was isolated from 7-d-old whole seedlings using RNcasy® Plant Minikit (Catalog No. 16419, Qiagen, CA) including on-column DNase treatment (Catalog No. 79254, Qiagen, CA) according to manufacturer’s instructions. The quantification of RNA, synthesis of cDNA and analysis of transcript accumulation in Col-0 WT and homozygous SALK_062388 (for Atlg26220), SALK_101567 (for At4g02290) and SALK_151393 (for At1g52410) mutants were performed as described in section 2.2.4 with the following modifications: for At4g02290 and Atlg52410, annealing temperature 43 was 55 °C. Number of amplification cycles in the PCR step was 31 for At4g02290 and At1g52410. A 7 uL aliquot of the PCR product was visualized by electrophoresis for 1 hr at 90V on a 1.5 % agarose gel containing ethidium bromide at 0.02 ug/mL (Catalog No. 15585-011, Invitrogen, CA). Ultraviolet images were obtained using the Gel Doc system (Bio-Rad Laboratories, Inc., CA) at subsaturation settings. 2.2.6 Hypocotyl Inhibition Assay To determine whether the T-DNA insertion mutants in candidate genes selected based on microarray analysis display impaired hypocotyl development in FRc, seeds of homozygous SALK_0623 88, SALK_101567 and SALK_151393 were sterilized and planted as described in section 2.2.1. Imbibing seeds were cold-stratified at 4 °C for 3 days in darkness. Plates were transferred to a humidity-controlled chamber with FRc illumination of 5 umol m.2 5-] or in darkness for 7 days at 22 oC. Seedlings were scanned and plant images were used to quantify hypocotyl lengths using ImagcJ software (NIH). The hypocotyl inhibition assay was repeated 3 times. Percentage dark length and standard deviations of percentage dark length were calculated according to the following equations. Two-tailed, unpaired Student’s t-tcst was performed to compare the percentage dark length of hypocotyls relative to Col-0 WT seedlings. Percentage dark length, °/o = Length of hypocotyl in BC, Rc or F RC (X) * 100% Length of hypocotyl in dark (Y) Standard Deviations (with ratio: % = X/Y * 100%): 44 SDX 2 + SDY ang avgY SD: Where: SD is standard deviation avg is mean hypocotyl length X is length of hypocotyl in Bc, Rc or FRc Y is length of hypocotyl in dark 2.3 Results and Discussion Previously, in Arabidopsis, constitutive expression of BVR was shown to result in phytochrome inactivation and has led to perturbation of distinct light-mediated growth and developmental processes. For an example, 35 S::pBVR lines displayed elongated hypocotyls under all light conditions tested due to inactivation of phytochromes (Lagarias ct a1., 1997; Montgomery et a1., 1999). Targeted expression of BVR using tissue-specific promoters was utilized in this chapter as a molecular tool to investigate the sites of photoperception responsible for regulating discrete responses, and to identify their candidate molecular effectors in FR-mediated photomorphogenesis in Arabidopsis. 2.3.1 MesophyII-specific Phytochromes Have Distinct Regulatory Roles in F ar-red Light Selective expression of plastid-targeted BVR under the control of CAB3 and MERIS promoters affected leaf morphology and hypocotyl growth in comparison to No-O WT under FRc conditions. Notably, CAB3::pBVR2 transgenic line with mesophyll- 45 specific inactivation of phytochromes exhibited closed cotyledons as opposed to open cotyledons in No-O WT, 35S::pBVR3 and MER15::pBVR1 (Figure 2.1). A similar phenotype with closed cotyledons was observed in the null phyA mutant relative to C 01- OWT (Figure 2.1). The phenotype of MER15::pBVR1 with meristem-spccific phytochrome inactivation was identical to the No-0 WT (Figure 2.1). Furthermore, cotyledons of No-O WT, 35S::pBVR3 and MER15zzpBVR remained yellow even after approximately 1.5 h of exposure to ambient white light during imaging, whereas in CAB3::pBVR2 line, exposure to white light resulted in enhanced greening. Color change in cotyledons of CAB3::pBVR2 line from yellow to green under white light following growth in FR light indicates that this line is defective in the FR block-to-greening response. Defective FR block-to-greening has been observed in phyA and hfrl mutants previously (Barnes et a1., 1996; F airchild, Schumaker, and Quail, 2000; Yanovsky et a1., 2002). The growth of Arabidopsis seedlings in FRc can induce partial photomorphogenesis without the accumulation of chlorophyll and is characterized by inhibition of hypocotyl elongation and open cotyledons (Whitelam et a1., 1993). These responses attribute to classic FR-HIR and thus the CAB3::pBVR2 line is impaired in aspects of FR-HIR. Barnes et al., (1996) reported that FR block-to-grecning is regulated by phyA. Similarity of impaired F R-HIR between the phyA mutant and CABB::pBVR2 line implies a potential role for mesophyll-specific phyA in the regulation of cotyledon opening and FR block-to-grecning under FR light. A distinct hypocotyl phenotype was obvious in the CAB3::pBVR2 line relative to No-O WT, 35S::pBVR3 and MER15::pBVRl lines (Figure 2.1). The CAB3::pBVR2 line displayed elongated hypocotyls relative to No-O WT under FRc. Although an elongated 46 hypocotyl phenotype was observed in the 35S::pBVR3 line relative to No-O WT, this phenotype was not as severe as what was observed for the CAB::pBVR2 line (Figure 2.1). In contrast to the elongated hypocotyls of 35S::pBVR3 and CAB3::pBVR2 lines, the MER15::pBVR] line displayed wild-type hypocotyl growth inhibition (Figure 2.1). The expression of BVR in CAB3::pBVR lines is regulated both spatially and by light, whereas in MER15::pBVR lines, B VR expression is regulated spatially. The analysis of fluence-rate dependence of hypocotyl inhibition response for BVR expressing transgenic lines 3SS::pBVR3, CAB3::pBVR and MER15::pBVR showed that hypocotyl inhibition was affected distinctively in CAB3::pBVR under F Rc illumination. Comparison of hypocotyl lengths of CAB3::pBVR lines and the 358::pBVR3 line under F Rc indicated that CAB3::pBVR lines were as impaired as 35S::pBVR3 line in response to F Rc (Figure 4E, Wamasooriya and Montgomery, 2009). However, MER15::pBVR lines responded similarly to No-0 WT seedlings (Figure 4F, Wamasooriya and Montgomery, 2009). The observation that the hypocotyl inhibition response of CAB3::pBVR lines was as deficient as 35S::pBVR3 line implies that mesophyll-localized phytochromes are involved in regulating hypocotyl inhibition under F Rc illumination. phyA has been recognized as the predominant phytochrome isoforrn responsible for the FR-dependent inhibition of hypocotyl elongation (Nagatani, Reed, and Chory, 1993; Parks and Quail, 1993; Whitelam et a1., 1993). Although (Toth et a1., 2001) showed that both PH YA and PH YC are expressed at high level in Arabidopsis cotyledons under FRc conditions, a clear role for phyC has not been recognized in F R-mediated inhibition of hypocotyl elongation (Balasubramanian et a1., 2006; Monte et a1., 2003). Thus, elongated hypocotyls in CAB3::pBVR lines under F Rc illumination are a result of 47 phytochrome deficiency in mesophyll tissues and mesophyll-localized phyA is likely responsible for initiating a signaling cascade in regulating FR-mcdiated inhibition of hypocotyl elongation. The recognized role for mesophyll-localized phyA in this study corroborates with previous reports indicating that cotyledons are the site of a phytochrome-induced signal that controls hypocotyl growth inhibition (Black and Shuttleworth, 1974) and that cotyledon-localized FR perception is able to impact gene expression in the hypocotyl (Tanaka et a1., 2002). Although CAB3::pBVR lines appeared to be as deficient as the 358::pBVR3 line in hypocotyl inhibition, close comparison of percentage dark lengths of CAB3::pBVR lines indicate that their hypocotyls were longer under increasing fluences of FR light than those of the 358::pBVR3 line (Figure 4E, Wamasooriya and Montgomery, 2009). Percentage dark lengths close to 100 % observed for 3SS::pBVR3 line suggest that hypocotyls of these lines in FRc were nearly identical in length to dark-grown controls (Figure 4E, Wamasooriya and Montgomery, 2009). This observation suggests that 35S::pBVR3 line was blind to F Rc illumination. However, under these conditions, CAB3::pBVR lines are able to perceive FR light and induce hypocotyl elongation. Induction of hypocotyl elongation by FR light leads to an increase in percentage dark lengths (> 100 %) for CAB3::pBVR lines and the hypocotyl elongation response showed further increase in a fluencc rate-dependent manner (Figure 4E, Wamasooriya and Montgomery, 2009). Fluence rate-dependent increase in hypocotyl length in CAB3::pBVR lines suggested that CAB3::pBVR lines are able to perceive FR light and induce hypocotyl elongation. As BVR accumulation occurs only in mesophyll tissues of CAB3::pBVR 48 lines (thus only mesophyll-localized phytochrome inactivation), the photoactive phyA present in the hypocotyls likely mediate F R-dependent hypocotyl growth through hypocotyl-localized phytochrome signaling. The analysis of publicly available gene expression data using the eF P browser (http://bbc.botany.utoronto.ca/efp/developmcnt) indicated that expression of CAB3 is relatively similar in Re, F Rc, Bc, or We light. Thus, the phenotypic differences observed for CAB3::pBVR lines relative to No-O WT, 35S::pBVR3 and MER15::pBVR lines under F Rc illumination indeed reflect distinct regulatory roles of phytochromes, and the possibility that such phenotypic differences resulted from varying levels of BVR expression was ruled out (Wamasooriya and Montgomery, 2009). 2.3.2 Phytochrome A is Involved in Regulating Anthocyanin Accumulation in FRc and Bc A regulatory role of phytochromes in the induction of sucrose-stimulated anthocyanin accumulation has been reported through comparative analyses of constitutive BVR expressing transgenic lines (Montgomery et a1., 2001; Montgomery et al., 1999) and through the analysis of pif3 mutants and PIF 3 over-expresser lines (Kim et a1., 2003; Shin, Park, and Choi, 2007). To determine the effects of localized phytochrome inactivation on sucrose-stimulated anthocyanin accumulation and to gain insight into the roles of localized pools of phytochromes in FRc, sucrose-stimulated anthocyanin accumulation levels were compared in FRc-grown and dark-grown representative CAB3::pBVR2, 358::pBVR3 and No-O WT plants. Anthocyanin accumulation was barely detectable in dark-grown seedlings and no significant differences in the levels 49 were obvious between any of the lines (Figure 2.2). In F Re, the levels of accumulated anthocyanins were lower in both of the BVR expressing transgenic lines relative to No-O WT (Figure 2.2). However, in comparison to No-O WT, the reduction observed for 3SS::pBVR3 was not significant (p=0.0597). Notably, CAB3::pBVR2 line showed a highly significant reduction in anthocyanin levels relative to No-O WT (p>0.0001) and a similar reduction was observed for the phyA mutant, relative to Col-0 WT (p>0.0001). This observation suggested that the CAB3::pBVR2 line and the phyA mutant are completely deficient in the FR-HIR of sucrose-stimulated anthocyanin accumulation and thus, mesophyll-localized phyA is involved in the regulation of F R-HIR of inducing sucrose-stimulated anthocyanin accumulation (Wamasooriya and Montgomery, 2009). Previous reports confirm that phyA has a significant role in the induction of anthocyanin accumulation under B illumination (Duck and Fankhauser, 2003; Neff and Chory, 1998; Poppe et a1., 1998; Weller et a1., 2001 ). Anthocyanin levels were quantified in a representative BVR-expressing transgenic line, CAB3::pBVR2, to determine whether mesophyll-localized phytochrome inactivation leads to a reduction of sucrose-stimulated anthocyanin accumulation under Bc illumination as observed under FRc illumination. In the presence of sucrose, anthocyanins were visible in the No-O WT under B light (Figure 2.3). The CAB3::pBVR2 line showed reduced levels of anthocyanins compared to No-O WT in Be (Figure 2.4). Phytochromes in meSOphyll tissues are responsible primarily for the induction of anthocyanins under Be as the 3SS::pBVR3 and CAB3::pBVR2 lines, which exhibited elongated hypocotyls under Bc (Figure 1A and 1B, Montgomery, 2009) displayed comparable level of reduction in anthocyanin accumulation relative to No-O WT. In comparison to the levels observed for No-O WT, 35 S::pBVR3 and 50 ,3 CAB3::pBVR2 lines accumulated only ~ 41 % and 39 % of No-O WT level of anthocyanins, respectively (Figure 2.4). However, under FRc illumination, CAB3::pBVR2 and 3SS::pBVR3 accumulated 1.8 % and 67 % of the No-0 WT level of anthocyanins, respectively (Figure 2.2). In contrast to the drastic reduction in anthocyanin accumulation observed in the CAB3::pBVR2 line relative to the 3SS::pBVR3 line under F Rc illumination, a similar degree of reduction in anthocyanin levels in these two transgenic lines in B illumination may be indicative of the possible roles of functional cryptochromes in regulating sucrose-stimulated anthocyanin accumulation under Bc light as reported previously (Ahmad, Lin, and Cashmore, 1995; Lin, Ahmad, and Cashmore, 1996; Mancinelli, 1985; Mancinelli, Rossi, and Moroni, 1991). Moreover, the higher amount of BVR accumulation in cotyledons of CAB3::pBVR3 relative to cotyledons of 3SS::pBVR3 is also related to the observed differences in anthocyanin levels in PRC and Be. Although B VR is expressed constitutivcly in the 3SS::pBVR3 line and in mesophyll tissues of the CAB3::pBVR3 line, tissue-specific expression of BVR within the cotyledons of the two transgenic lines could differ slightly. Such differences could have distinct functional importance on the amount of phytochrome inactivated in the cotyledons of CAB3::pBVR2 and 3SS::pBVR3 under F Rc vs. Bc light. Similar to the function of mesophyll-localized phytochromes under F Rc (Wamasooriya and Montgomery, 2009), mesophyll-localized phyA is primarily responsible for the phytochrome-dependent induction of anthocyanin accumulation in Be (Figure 2.4). Under Bc illumination, the levels of anthocyanins in CAB3::pBVR2 and the phyA mutant were reduced to comparable levels relative to their cognate WT parent, i.e., CAB3::pBVR2 accumulates ~ 39 % of the level of anthocyanins accumulated in No-O 51 WT, whereas the phyA mutant accumulates ~ 46% of the Col-O WT level of anthocyanins. Notably, a phyB mutant accumulates at least as much anthocyanin as the Col-0 WT under Bc (Figure 2.4). Light- and/or sucrose-inducible anthocyanins in most species accumulate in vacuolar space of cells in photosynthetic tissues, i.e. palisade and spongy mesophyll (Gould and Quinn, 1998; Lee and Collins, 2001) and in epidermal cells (Kubo et a1., 1999). The comparative analysis of anthocyanin accumulation in the CAB3::pBVR2 line vs. the 35S::pBVR3 and the phyA mutant line indicates that phyA in the mesophyll tissues is responsible for the mesophyll-specific induction of anthocyanin accumulation, and may impact inter-tissue anthocyanin accumulation in the epidermis, under Be, as well as, under FRc illumination. 2.3.3 Mesophle-localized Phytochrome Inactivation Leads to Distinct Gene Expression Patterns Comparison of phenotypes between FRc-grown CAB3::pBVR2 and 35S::pBVR3 lines indicated that the CAB3::pBVR2 line was impaired in a number of F R-HIR, i.e. hypocotyl inhibition, cotyledon expansion, FR block-to-greening and sucrose-induced anthocyanin accumulation (Wamasooriya and Montgomery, 2009). Moreover, a unique fluence-ratc dependent hypocotyl elongation response mediated by hypocotyl-localized phyA was also apparent in the CAB3::pBVR2 line lacking mesophyll-localized phyA (Wamasooriya and Montgomery, 2009). To identify and characterize genes regulating spatial-specific FR-HIR mediated by phyA, a comparative microarray-based gene expression profiling of F Rc-grown No-O WT, 3SS::pBVR3 and CAB3::pBVR2 whole 52 seedlings was performed. As 7-d-old seedlings of 3SS::pBVR3 and CAB3::pBVR2 were phenotypically different in PRC, through comparative gene expression profiling, the aim was to obtain potential insight into the molecular basis of F R-HIR in Arabidopsis. To identify candidate genes regulating F R-HIR based on the differential gene expression between CAB3::pBVR2 line vs. 3SS::pBVR3 line, an expression filter of fold change greater or equal to 2.0 was applied for these two lines against the No-O WT sample. The comparison of differentially expressed genes in CAB3::pBVR2 vs. 35S::pBVR3, 3SS::pBVR3 vs. No-O WT and CAB3::pBVR2 vs. No-O WT revealed that 180 genes are shared among the three groups (Figure 2.5). Relative to No-O WT, 348 genes are differentially expressed commonly in the CAB3::pBVR2 and 358::pBVR3 lines, whereas 17 genes and 1115 genes show unique differential expression in the 35S::pBVR3 and CAB3::pBVR2 lines relative to No-0 WT, respectively (Figure 2.5). The phenotypic differences between No-O WT and CAB3::pBVR2 as evident in Figure 2.1, may be a result of differential expression patterns of the 11 15 genes enriched in CAB3::pBVR2 relative to No-O WT. The comparison of CAB3::pBVR2 vs. 3SS::pBVR3 and 35S::pBVR3 vs. No-O WT showed only 3 genes in common that were differentially expressed. These 3 genes are known to encode a transporter-related protein involved in ion transport, an F-box family protein and a nodulin MtN21 family protein (The I Arabidopsis Information Resource, TAIR). Although 518 genes are differentially expressed in common between CAB3::pBVR2 vs. No-0 WT and CAB3::pBVR2 vs. 35S::pBVR3, only 1 1 genes were shown to have unique differential expression in CAB3::pBVR2 vs. 3SS::pBVR3 (Figure 2.5). Based on the information from functional categorization of gene ontology annotations, the entity list of 11 genes with unique 53 differential expression in CAB3::pBVR2 vs. 3SS::pBVR3 did not include genes with recognized implications in FR-HIR (Table 2.1). Thus, to narrow down the list of candidate genes, the gene entity list obtained from comparison of CAB3::pBVR2 and 3SS::pBVR3 was utilized. From the 712 differentially expressed genes between CAB3::pBVR2 and 3SS::pBVR2, ~ 30 genes were selected based on the fold change observed in CAB3::pBVR2 and on information available from functional categorization of gene ontology annotations, TAIR and tissue- and light-dependent gene expression patterns from publicly available gene expression data (AtGenExpress, http://www.weigelworld.org/resources/microarray/AtGenExpress). To prioritize the initial analysis of candidate genes that could be involved in hypocotyl development under F Rc, 3 genes were selected based on the fold-change observed in CAB3::pBVR2 vs. 35S::pBVR3: At1g26220 (probe ID, 245877_at), At4g02290 (probe ID, 255517_at) and At1g52410 (probe ID, 59609_at). At1g26220, At4g02290 and At1g52410 genes are known to encode a GCNS-related N-acetyltransferase (GNAT) family protein, a glycosyl hydrolase family 9 protein and a caldesmon-related protein with a novel calcium—binding repeat sequence, respectively (TAIR). Fold-change in gene expression for At] g26220, At4g02290 and At1g52410 in CAB3::pBVR3 relative to 35S::pBVR3 was - 2.03, + 5.55 and + 6.27, respectively. The validation of microarray data by RT-PCR analysis of transcript accumulation of candidate genes, At1g26220, At4g02290 and At1g52410 in F Rc-grown No-O WT, 35S::pBVR3 and CAB3::pBVR2 indicated that transcript accumulation of At1g26220 was down-regulated, whereas, transcript accumulation of A t4g02290 and At1g52410 was up-regulated in the CAB3::pBVR2 line relative to No-O WT and 35S::pBVR3 line 54 (Figure 2.6). Thus, the analysis of transcript accumulation by RT-PCR validated the fold- change in gene expression levels in the CAB3::pBVR2 line as evident through microarray analysis for the 3 candidate genes selected for further analysis. Tissue- and light-specific gene expression analysis of At1g26220, At4g02290 and At1g52410 using publicly available gene expression data (AtGenExpress, http://www.weigelworld.org/resources/microarray/AtGenExpress) revealed unique expression patterns in light vs. dark and in cotyledons and/or hypocotyls in young wild- type Arabidopsis seedlings. The expression of At1g26220 was induced under longer duration of exposure to W, B, and R light conditions relative to darkness, whereas exposure to shorter or longer duration of FR light did not change the expression level (Figure 2.7). However, validation of At1g26220 expression in No-0 WT, 35 S::pBVR3 and CAB3::pBVR2 by RT-PCR indicated that accumulation of At1g26220 transcript expression was higher in No-0 WT (Figure 2.6). As FR light is of lower energy wavelength, exposure of No-O WT for a prolonged time period (~ 7d) could result in up-regulation of At] g26220 and down- regulation of this gene in 35S::pBVR3 and CABB::pBVR2 due to phytochrome W inactivation under F Rc. t At4g02290 showed distinct patterns of gene expression in dark vs. short and long duration of exposure to different light conditions. In general, the expression of At4g02290 was up-regulated in dark relative to light and longer duration of darkness, further increased its expression (Figure 2.7). Exposure to shorter duration of B, R and FR illumination led to up-regulation of At4g02290; however, longer duration of B, R and FR illumination was able to down-regulate its expression (Figure 2.7). Distinct expression 55 patterns of At4g02290 dependent on the presence of light and its duration, especially in FR light and the reduction of its transcript in No-0 WT could implicate a possible role in phenotypic difference observed for CAB3::pBVR2 in FRc. Although expression of At1g52410 was constant in darkness and under different light conditions, tissue-specific expression analysis showed distinct patterns. The expression of At1g52410 was clearly up-regulated in the hypocotyl, shoot apex and rosettes of vegetative wild-type seedlings (Figure 2.8). Although transcript accumulation of At1g5241 0 was barely detectable in No-O WT (Figure 2.6), its distinct expression in the hypocotyl makes it a good candidate for further analysis. The expression of At1g26220 was up-regulated in spatially-discrete patterns, especially in cotyledons, leaves, vegetative rosettes and green tissues of wild-type seedlings (Figure 2.8). Microarray results and analysis of transcript accumulation by RT-PCR for this gene in No-O WT also indicated higher expression/transcript accumulation relative to CAB3::pBVR2 with mesophyll-specific phytochrome inactivation. This observation implies that At] g26220 expression is reduced upon mesophyll-specific inactivation of phytochromes and a putative role in mesophyll-localized phytochrome signaling. Despite the very low transcript accumulation in No-O WT (Figure 2.6), clear up-regulation was observed for At4g02290 in the hypocotyls of wild-type seedlings relative to other tissues (Figure 2.8). No-O WT seedlings have functional mesophyll- and hypocotyl-localized phytochromes and the CAB3::pBVR2 line has functional hypocotyl-localized phytochromes, but lacks functional mesophyll-localized phytochromes. Since microarray analysis and validation of At4g02290 transcript accumulation by RT-PCR were performed with RNA extracted from FRc-grown whole seedlings, elongated hypocotyls 56 of the CAB3::pBVR2 line may have contributed a larger proportion of RNA to the total RNA pool (due to more abundant transcripts from hypocotyl-specific At4g02290 expression in the CAB3::pBVR2 line) and thus are indicative of increased expression of At4g02290 in the CAB3::pBVR2 line relative to No-O WT. The hypocotyl-specific increased expression as apparent in the Heatmap (Figure 2.8) and the up-regulation of At4g02290 in the CAB3::pBVR2 line lacking mesophyll-localized phytochromes implicate a putative role of At4g02290 in hypocotyl-localized phytochrome signaling. 2.3.4 GCNS- and Caldesmon-related Proteins are Implicated in the Regulation of Hypocotyl Development under FRc To determine whether candidate genes have functional roles in phytochrome- regulated hypocotyl development in F Rc, T-DNA insertion mutants having T-DNA insertions in an exon of At1g26220, At4g02290 and At1g52410 (Figure 2.9 A, 2.10 A and 2.11 A) were obtained from the Salk T-DNA insertion mutant collection (Alonso et a1., 2003). RT-PCR analysis of transcript accumulation of At1g26220 and At4g02290 in confirmed homozygous T-DNA insertion mutants indicated that respective transcripts were absent in SALK_062388 (Figure 2.9 B) and SALK_101567 (Figure 2.10 B), respectively, and that these lines were null mutants. However, a T-DNA insertion mutant for At1g52410, SALK_151393, displayed a very low level of transcript accumulation and was similar to that of Col-0 WT (Figure 2.11 B). As the T-DNA insertion in SALK_151393 is in the last exon and the primers for RT—PCR analysis were designed to anneal to a region upstream of the T-DNA insertion site, SALK_151393 may accumulate a truncated transcript and lack a fiinctional protein corresponding to At1g52410. 57 Comparative analysis of F Rc-grown CAB3::pBVR2 and 3SS::pBVR3 indicated that CAB3::pBVR2 has elongated hypocotyls relative to No-O WT and 35S::pBVR3 and was indicative of hypocotyl-localized phytochrome signaling resulting in hypocotyl elongation in the absence of mesophyll-localized phytochrome action on hypocotyl inhibition (Figure 2.1; as discussed in section 2.3.1). To assess whether the selected candidate genes were involved in hypocotyl development under F Rc, hypocotyl inhibition response was quantified as a percentage dark length in FRc-grown 7-d-old homozygous T-DNA insertion mutants, SALK_062388, SALK_101567 and SALK_151393 (Figure 2.12). The SALK_0623 88 line displayed elongated hypocotyls relative to hypocotyls of Col-0 WT and the increase in length was significant (p=0.0002). The increase in the percentage dark length for SALK_0623 88 was ~ 1 1 % greater than the percentage dark length of Col-0 WT. A significant increase in the hypocotyl length in SALK_101567 relative to Col-0 WT was not apparent (p=0.2881) under F Rc (Figure 2.12). However, a marginally significant increase in the hypocotyl length for SALK_151393 relative to Col-O WT (p=0.0375) was observed under FRc. The increase in the percentage dark length for SALK_151393 was ~ 2 % of Col-0 WT percentage dark length (Figure 2.12). Based on the comparative microarray-based gene expression profiling and validation by RT-PCR analysis, the expression of At1g26220 was down-regulated in the CAB3::pBVR2 line in PRC. As the T-DNA insertion in an exon of At1g26220 eliminated the accumulation of its transcript (Figure 2.9 B) in the SALK_0623 88, the increase in hypocotyl length observed for SALK_062388 relative to Col-0 WT is a result of the lack of the GNAT family protein encoded by At1g26220 and implicates that in the wild-type 58 _ E "l Arabidopsis plants, the GNAT family protein has a regulatory role in hypocotyl inhibition in PRC. Although a short hypocotyl phenotype in a T-DNA disruption mutant of the Arabidopsis GCN5, gcn5-1 under Wc light has been reported (Vlachonasios, Thomashow, and Triezenberg, 2003), any reports on roles of the GNAT family protein encoded by Atlg26220 under F Rc condition has not emerged. However, the observation that a mutation of GCN5 in the gcn5-I mutant, leads to shorter hypocotyls in We conditions does indicate that a GNAT family protein encoded by a homolog of At1g26220 as having a putative role in hypocotyl development in Arabidopsis. As a number of molecular effectors are involved in the regulation of hypocotyl development, the contribution of each candidate gene to the overall phenotype can be minor and thus a ~ 11 % increase in hypocotyl length observed for SALK_062388 is reflective of At1g26220 being a candidate gene in the regulation of hypocotyl development in Arabidopsis under F Re. The analysis of the hypocotyl inhibition response of SALK_151393 indicated that the increase in the percentage dark length was ~ 2 % of Col-0 WT percentage dark length. As the microarray analysis indicated that At1g52410 expression was up-regulated by 6.27 X in the CAB3::pBVR2 relative to 35S::pBVR3, a decrease in hypocotyl length was anticipated for SALK_151393 in the absence of the At1g52410 transcript. However, RT-PCR analysis of SALK_151393 showed very low At1g52410 transcript accumulation that was similar to the level in Col-0 WT (Figure 2.1 l B). The ~ 2 % increase in the percentage dark length relative to Col-0 WT may be indicative of possible residual activity of the caldesmon-related protein encoded by At1g52410. Previous reports indicate that At1g52410 encodes TSAI (TSK-associating protein 1) that interacts with a 59 protein complex, TSK/MGO3/BRU, a key factor in cell division control and plant morphogenesis (Suzuki et a1., 2005; Yamada et a1., 2008) and could have implications in FR-dependent morphogenesis in Arabidopsis. 2.3.5 Summary Comparative phenotypic analysis of CAB3::pBVR2 and 3SS::pBVR3 under FRc and Be illumination revealed that spatially-distinct pools of phytochromes are able to perceive light and regulate discrete aspects of photomorphogenesis through inter-tissue and inter-organ signaling pathways. The comparison of hypocotyl inhibition responses of 3SS::pBVR3 and CAB3::pBVR2 across increasing fluences of F Rc illumination showed that mesophyll-localized phyA regulates FR-mediated hypocotyl inhibition. However, in the absence of mesophyll-localized phyA, the functional phyA in hypocotyls is able to mediate cell elongation through hypocotyl-localized phytochrome signaling resulting in FR-dependent hypocotyl elongation. The observation that this growth promotive response, i.e. hypocotyl elongation, is only present in the absence of mesophyll-localized phyA suggests that the inhibitory activity by mesophyll-localized phyA on the hypocotyl is more pronounced relative to the elongation response exerted by hypocotyl-localized phyA. The dissection of growth inhibitory and stimulatory effects of spatial-specific pools of phytochromes indicates that de-etiolation involves coordination between distinct pools of phytochromes. Thus, targeted BVR expression has essentially disrupted an inter- tissue signaling between mesophyll tissues and hypocotyl in the CAB3::pBVR2 line allowing the. identification of distinct functions mediated by localized pools of phytochromes. Moreover, additional regulatory roles for mesophyll-localized phyA such 60 _.—7 as cotyledon opening and FR block-to-greening that are classic F R-HIR could be identified. The similar levels of anthocyanin accumulation under F Re in the CAB3::pBVR2 line and the phyA mutant that is completely deficient in the FR-HIR implied that sucrose-stimulated anthocyanin accumulation is regulated by mesophyll- localized phyA specifically and its possible contribution to induction of anthocyanin accumulation under Bc. Comparative microarray-based gene expression profiling of FRc- grown No-O WT, 35 S::pBVR3 and CAB3::pBVR2 whole seedlings allowed initial selection of candidate genes that have significant changes in gene expression levels between CAB3::pBVR2 and 3SS::pBVR3. The initial analysis of hypocotyl inhibition responses of T-DNA insertion mutants in At1g26220 (encodes a GNAT family protein), At4g02290 (encodes a glycosyl hydrolase family 9 protein) and At1g52410 (encodes a caldesmon-related protein with a novel calcium-binding repeat sequence) under FRc identified a GNAT family protein and a caldesmon-related protein as candidate signaling intermediates in regulating F R-mcdiated hypocotyl development by phyA in Arabidopsis. 2.4 Future Perspectives Comparative microarray-based gene expression profiling of F Rc-grown No-0 WT, 35S::pBVR3 and CAB3::pBVR2 whole seedlings revealed that distinct gene expression patterns are observed for CAB3::pBVR2 relative to 35S::pBVR3 and the implications of such gene expression patterns likely represent the molecular and cellular bases of phenotypic differences between the two transgenic lines. Thus, differentially expressed genes in the CAB3::pBVR2 vs. 35S::pBVR3 may have functional roles in hypocotyl inhibition regulated by mesophyll-specific phyA and hypocotyl elongation 61 regulated by hypocotyl-specific phyA under FRc and/or characteristic F R-HIRs, i.e. cotyledon expansion and FR-block-to-greening regulated by phyA. Although a number of signaling intermediates have been identified in the phyA-specific signaling pathway in FR light (as discussed in section 2.1.3), a comprehensive understanding of the sites of photoperception and the molecular and cellular mechanisms of spatial-specific phyA- mediated responses is yet limited. Therefore, the comparative microarray-based gene expression profiling of CAB3::pBVR3 relative to 35S::pBVR3 aids in the initial analyses and selection of candidate signaling intermediates involved in mesophyll- and hypocotyl- specific phyA signaling underlying the fine-tuning of F R-HIRs in Arabidopsis. As evident by microarray analysis and subsequent validation by RT-PCR, the expression of At1g26220 was down-regulated in CAB3::pBVR2 with mesophyll-specific phytochrome inactivation relative to 35S::pBVR3 with constitutive phytochrome inactivation (Figure 2.6). If At1g26220 has a putative role in regulating hypocotyl inhibition under FR light, a mutant in the At1g26220 is expected to display more elongated hypocotyls relative to WT seedlings in FR light. The observation that J SALK_0623 88 (a T-DNA insertion mutant in At1g26220) had an ~ 1 1 % increase in percentage dark length relative to Col-0 WT implicates a role of At1g26220 in regulating rum 3 hypocotyl inhibition mediated by mesophyll-localized phyA. To rule out the possibility of positional effects of T-DNA insertion on the observed hypocotyl elongation in SALK_0623 88, the assessment of additional mutant alleles of At1g26220 for hypocotyl inhibition response under FRc is required. SALK_150736 and SALK_022035 from the Salk T-DNA insertion mutant collection (Alonso et a1., 2003) contain a T-DNA insertion in exons of At1g26220. If SALK_I 50736 and SALK_022035 also display a similar 62 degree of hypocotyl elongation relative to Col-0 WT as SALK_062388, this result would suggest that the At1g26220 gene has a role in inhibition of hypocotyl elongation in FR light. However, the complementation of T-DNA insertion mutants in the At1g26220 gene that display elongated hypocotyls under FR light, with a WT copy of At1g26220 under the regulatory control of the native promoter and subsequent restoration of WT phenotype in the complemented transgenic lines, will confirm that the At1g26220 gene encoding a GNAT family protein as a signaling intermediate of the F R-dependent inhibition of hypocotyl elongation regulated by mesophyll-specific phyA. Using microarray analysis and validation by RT-PCR, the expression of At1g52410 was determined to be up-regulated in CAB3::pBVR2 relative to 35S::pBVR3 (Figure 2.6). Thus, if At1g52410 has a regulatory role in hypocotyl inhibition under FR light, a mutant in the At1g52410 may be expected to display hypocotyls of reduced length relative to WT seedlings. However, a marginally significant increase in the hypocotyl length of SALK_I 51393 relative to Col-0 WT was apparent (p=0.0375) under FRc and the increase in the percentage dark length for SALK_151393 was ~ 2 % of Col-0 WT percentage dark length. Moreover, the analysis of transcript accumulation of At1g5241 0 by RT-PCR revealed similar levels of transcript accumulation in Col-0 WT, as well as in SALK_151393 (Figure 2.1 1 B) indicating that SALK_151393 is not a true null mutant. Thus, the analysis of hypocotyl inhibition response in additional T-DNA insertion mutant alleles of At1g5241 0, such as SALK_001 102 and GK-299G09-015515 from the Salk T- DNA insertion mutant collection (Alonso et a1., 2003) having a T-DNA insertion in the promoter and in an exon, respectively, under FR light can provide additional insight into the involvement of At] g524 10 as a regulator of phyA-mediated hypocotyl inhibition in 63 FR light. The complementation of T-DNA insertion mutants (with a WT copy of At1g52410 under the regulatory control of the native promoter) and restoration of WT hypocotyl inhibition response in the complemented lines under FR light will confirm that the caldesmon-related protein encoded by At1g52410 as a signaling intermediate of inhibition of hypocotyl elongation regulated by mesophyll-specific phyA under FR light. The phenotypic similarity of CAB3::pBVR2 line and phyA null mutant with respect to cotyledon greening induced by white light following growth in FR light, closed cotyledons and elongated hypocotyls indicated that both lines are defective in a number of F R-HIRs (Wamasooriya and Montgomery, 2009). Defective F R block-to- greening has also been observed in hfrl mutants previously (Fairchild, Schumaker, and Quail, 2000). HFR] is known to encode a phyA-specific signaling intermediate (Duck and F ankhauser, 2003; Fairchild, Schumaker, and Quail, 2000; Jang et a1., 2005; Kim et a1., 2002). Although F Re is able to inhibit hypocotyl elongation in hfrl mutants, the inhibition is significantly impaired in the hfrI mutants in moderate (5 umol rn-2 3.1) and strong (42 umol rri'2 3.1) F Re light (Fairchild, Schumaker, and Quail, 2000). This phenotype contrasts with the complete blindness to F Rc of the phyA mutant (Barnes et a1., 1996; Fairchild, Schumaker, and Quail, 2000; Yanovsky et a1., 2002; Yanovsky, “i Whitelam, and Casal, 2000). Moreover, hfrl mutants displayed cotyledons separation in contrast to phyA null mutantsl(Fairchild, Schumaker, and Quail, 2000). Due to the similarity of phenotypes among CAB3::pBVR3, phyA and hfrI mutants with respect to hypocotyl inhibition and FR block-to-greening induced by white light following growth in 5 umol m.2 s"1 FR light, the analysis of expression ofAt1g26220 and At1g52410 in 64 CAB3::pBVR3, phyA and hfrl mutants by qRT-PCR under F Rc would reveal the gene expression patterns of putative candidate genes. The similarity of expression levels for At1g26220 and At1g52410 among CAB3::pBVR3, phyA and hfrl mutants will further substantiate the involvement of a GNAT family protein and a caldesmon-related protein as signaling intermediates in regulating phyA-mediated hypocotyl development in Arabidopsis seedlings under FR light. 65 A. “3‘ \lljkl5: pl))\l)\3 I‘l3\l{l [lllllfl No-(l (TAPS ' \\"l‘ pliVR: Vol-(1 \\"l' Figure 2.1 Photomorphogenesis of F Rc-grown wild-type and transgenic BVR plants. No-O wild-type (No-0 WT), 35S::pBVR3, CAB3::pBVR2, MER15::pBVR], Col-0 wild-type (Col-0 WT) and phyA (SALK_014575) were grown at 20 °C on Phytablend medium containing 1 % Suc for 7 (1 under FRc illumination of 5 umol rri'2 3']. Above each seedling, cotyledons are shown that were separated and arranged to display the full cotyledon surface area. 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AW- odm 2.6 References Ahmad, M., and Cashmore, A. R. (1997). The blue-light receptor cryptochrome 1 shows functional dependence on phytochrome A or phytochrome B in Arabidopsis thaliana. Plant J 11(3), 421-427. Ahmad, M., Lin, C., and Cashmore, A. R. (1995). Mutations throughout an Arabidopsis blue-light photoreceptor impair blue-light-responsive anthocyanin accumulation and inhibition of hypocotyl elongation. Plant J 8(5), 653-658. Alonso, J. M., Stepanova, A. N., Leisse, T. J ., Kim, C. J., Chen, H., Shinn, P., Stevenson, D. K., Zimmerman, J ., Barajas, P., Cheuk, R., Gadrinab, C., Heller, C., Jeske, A., Koesema, E., Meyers, C. C., Parker, H., Prednis, L., Ansari, Y., Choy, N., Deen, H., Geralt, M., Hazari, N., Horn, 13., Kames, M., Mulholland, C., Ndubaku, R., Schmidt, 1., Guzman, P., Aguilar-Henonin, L., Schmid, M., Weigel, D., Carter, D. E., Marchand, T., Risseeuw, E., Brogden, D., Zeko, A., Crosby, W. L., Berry, C. C., and Ecker, J. R. (2003). 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Poppe, C., Sweere, U., Drumm-Herrel, H., and Schafer, E. (1998). The blue light receptor cryptochrome 1 can act independently of phytochrome A and B in Arabidopsis thaliana. Plant J 16(4), 465-471. Quail, P. H., Boylan, M. T., Parks, B. M., Short, T. W., Xu, Y., and Wagner, D. (1995). Phytochromes: photosensory perception and signal transduction. Science 268(5211), 675-680. Rabino, I., and Mancinelli, A. L. (1986). Light, Temperature, and Anthocyanin Production. Plant Physiol 81(3), 922-924. Reed, J. W., Nagatani, A., Elich, T. D., F agan, M., and Chory, J. (1994). Phytochrome A and Phytochrome B Have Overlapping but Distinct Functions in Arabidopsis Development. Plant Physiol 104(4), 1139-1149. Roig-Villanova, 1., Bou, J., Sorin, C., Devlin, P. F., and Martinez-Garcia, J. F. (2006). Identification of primary target genes of phytochrome signaling. Early transcriptional control during shade avoidance responses in Arabidopsis. Plant Physiol 141(1), 85-96. Rosler, J., Klein, 1., and Zeidler, M. (2007). Arabidopsis flil/fhyl double mutant reveals a distinct cytoplasmic action of phytochrome A. Proc Natl Acad Sci U S A 104(25), 10737-10742. Ruckle, M. E., DeMarco, S. M., and Larkin, R. M. (2007). Plastid signals remodel light signaling networks and are essential for efficient chloroplast biogenesis in Arabidopsis. Plant Cell 19(12), 3944-3960. Sharrock, R. A., and Clack, T. (2002). Patterns of expression and normalized levels of the five Arabidopsis phytochromes. Plant Physiol 130(1), 442-456. Shin, J ., Park, E., and Choi, G. (2007). PIF 3 regulates anthocyanin biosynthesis in an HYS-dependent manner with both factors directly binding anthocyanin biosynthetic gene promoters in Arabidopsis. Plant J 49(6), 981-994. Shinomura, T., Uchida, K., and F uruya, M. (2000). Elementary processes of photoperception by phytochrome A for high-irradiance response of hypocotyl elongation in Arabidopsis. Plant Physiol 122(1), 147-156. 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Dissecting the phytochrome A-dependent signaling network in higher plants. Trends Plant Sci 8(4), 172-178. Wamasooriya, S. N., and Montgomery, B. L. (2009). Detection of spatial-specific phytochrome responses using targeted expression of biliverdin reductase in Arabidopsis. Plant Physiol 149(1), 424-433. Weller, J. L., Beauchamp, N., Kerckhoffs, L. H., Platten, J. D., and Reid, J. B. (2001). Interaction of phytochromes A and B in the control of de-etiolation and flowering in pea. PlantJ 26(3), 283-294. Whitelam, G. C., Johnson, E., Peng, J ., Carol, R, Anderson, M. L., Cowl, J. S., and Harberd, N. P. (1993). Phytochrome A null mutants of Arabidopsis display a wild-type phenotype in white light. Plant Cell 5(7), 75 7-768. Yadav, V., Mallappa, C., Gangappa, S. N., Bhatia, S., and Chattopadhyay, S. (2005). A basic helix-loop-helix transcription factor in Arabidopsis, MYC2, acts as a repressor of blue light-mediated photomorphogenic growth. Plant Cell 17(7), 1953-1966. Yamada, K., Nagano, A. J., Nishina, M., Hara-Nishimura, I., and Nishimura, M. (2008). NA12 is an endoplasmic reticulum body component that enables ER body formation in Arabidopsis thaliana. Plant Cell 20(9), 2529-2540. Yang, S. W., Jang, 1. C., Henriques, R., and Chua, N. H. (2009). FAR-RED ELONGATED HYPOCOTYLI and FHYl-LIKE associate with the Arabidopsis transcription factors LAF 1 and HFRI to transmit phytochrome A signals for inhibition of hypocotyl elongation. Plant Cell 21(5), 1341-1359. Yanovsky, M. J ., Casal, J. J ., and Luppi, J. P. (1997). The VLF loci, polymorphic between ecotypes Landsberg erecta and Columbia, dissect two branches of phytochrome A signal transduction that correspond to very-low-fluence and hi gh- irradiance responses. Plant J 12(3), 659-667. 85 Yanovsky, M. J ., Casal, J. J ., and Whitelam, G. C. (1995). Phytochrome A, phytochrome B and HY4 are involved in hypocotyl growth responses to natural radiation in ArabidOpsis: weak de-etiolation of the phyA mutant under dense canopies. Plant, Cell & Environment 18(7), 788-794. Yanovsky, M. J ., Luppi, J. P., Kirchbauer, D., Ogorodnikova, O. B., Sineshchekov, V. A., Adam, B, Kircher, S., Staneloni, R. J ., Schafer, E., Nagy, F ., and Casal, J. J. (2002). Missense mutation in the PAS2 domain of phytochrome A impairs subnuclear localization and a subset of responses. Plant Cell 14(7), 1591-1603. Yanovsky, M. J ., Whitelam, G. C., and Casal, J. J. (2000). fl1y3-1 retains inductive responses of phytochrome A. Plant Physiol 123(1), 23 5-242. 86 Chapter 3 Phytochrome-mediated Light-dependent Anthocyanin Accumulation in Red Light Some of the work included in chapter 3 was published in the journal, Plant Physiology. Wamasooriya SN and Montgomery BL, (2009), Detection of Spatial-Specific Phytochrome Responses Using Targeted Expression of Biliverdin Reductase in Arabidopsis, Plant Physiology, 149(1), 424-433. Most of the work in chapter 3 is from a manuscript submitted to the journal, Plant, Cell and Physiology. Wamasooriya SN, Porter KJ and Montgomery BL, Tissue- and Isoform-Specific Phytochrome Regulation of Li ght-Dependent Anthocyanin Accumulation in Arabidopsis thaliana (submitted). 87 3.1 Overview Exogenous and endogenous stimuli can lead to biosynthesis and accumulation of flavonoid compounds: flavones, flavonols, isoflavonoids and anthocyanins and among them anthocyanins are the most abundant in plants (Shi and Xie, 2010). The accumulation of flavonoid compounds is highly regulated in response to environmental stimuli such as light, temperature, and nutrient availability (Feinbaum and Ausubel, 1988). Among the external stimuli, light is one of the most important environmental stimuli regulating the expression of flavonoid structural genes (Mol et a1., 1996). Metabolites, hormones, and the developmental stage of the tissues are among the endogenous stimuli that regulate anthocyanin biosynthesis and deposition (Mol et a1., 1996) Synthesis and deposition of anthocyanins in different tissues within the plant can exert specific functions. Light-induced anthocyanin biosynthesis and accumulation in the epidermis is thought to have evolved for protection of plants against excess or damaging solar radiation (Drumm-Herrel and Mohr, 1985). Additionally, anthocyanins are important antioxidant molecules and aid in protecting plants from damage by active oxygen species (Nagata et a1., 2003). Biosynthesis and accumulation of pigments is one of main aspects of photomorphogenesis in Arabidopsis seedlings upon emergence into the light environment. Photomorphogenesis is regulated by the action of at least two major classes of photoreceptors: phytochromes in R and FR illumination (Mancinelli, 1985) and cryptochromes in UV-A and B illumination (Ahmad, Lin, and Cashmore, 1995). A small nuclear gene family of five members, PH YA — PH YE, has been identified in Arabidopsis (Fankhauser and Staiger, 2002; Quail, 1994) and each gene encodes a 88 phytochrome apoprotein that covalently attaches to a single linear tetrapyrrole chromophore, phytochromobilin (PCDB; Terry, Wahleithner, and Lagarias, 1993). phyA is the phytochrome primarily responsible for FR-dependent growth responses (Nagatani, Reed, and Chory, 1993; Whitelam et a1., 1993) and has an additional role in regulating photoresponses under B illumination (Chun, Kawakami, and Christopher, 2001; Duck and F ankhauser, 2003; Neff and Chory, 1998; Whitelam et a1., 1993; Yadav et a1., 2005). phyB through phyE contribute to the regulation of growth and development, primarily in response to R illumination (Aukerman et a1., 1997; Franklin et a1., 2003; Monte et a1., 2003; Nagatani, Reed, and Chory, 1993; Reed et a1., 1993). Anthocyanin biosynthesis and accumulation is one of the characteristic aspects of the photomorphogenesis process and differential expression of anthocyanin structural genes, as well as their regulatory genes is important for mediating anthocyanin biosynthesis. Despite major advances in understanding light-regulated anthocyanin synthesis, information on the involvement of phytochromes in complex signaling cascades to mediate anthocyanin'synthesis and deposition is limited. Thus, investigating light-dependent anthocyanin biosynthesis and accumulation is a useful undertaking to gain insight into the photoregulation of pigmentation, as well as to define the discrete and overlapping roles of phytochrome isoforms under different light conditions. 3.1.1 Anthocyanin Biosynthesis and Accumulation Flavonoids are aromatic secondary metabolites with numerous biological functions (Agati and Tattini, 2010; Lepiniec et a1., 2006). However, flavonoid compounds are largely nonessential for plant viability (Nesi et a1., 2000). In plants, the 89 main flavonoid compounds are flavones, flavonols, isoflavonoids and anthocyanins; however, Arabidopsis cannot produce isoflavonoids due to lack of chalcone reductase and isoflavone synthase (Aoki, Akashi, and Ayabe, 2000). The anthocyanin biosynthetic pathway is characterized by two groups ofco-regulated structural genes: early biosynthetic genes. i.e., ('HS, CHI, F3H, and FLS, and the late biosynthetic genes, i.e., BER and LDOX, during seedling development (Kubasek et al., 1992). Many ofthe early and late biosynthetic genes are induced by sucrose and to a lesser degree by other sugars (Gollop et a1., 2002; Martin, Oswald, and Graham, 2002; Solfanelli et a1., 2006). The expression of early and late biosynthetic genes is highly regulated by the products of multiple regulatory genes and tissue-specific expression of biosynthetic genes and anthocyanin deposition are correlated (Procissi et a1., 1997). The mostly nonessential nature of anthocyanins for plant viability has made the generation of mutants in this biosynthetic pathway feasible and has facilitated the genetic and molecular dissection of the pathway. Isolation and characterization of mutants and functional genomics approaches have identified a number of such regulatory gene families. Regulatory proteins can directly or indirectly regulate the biosynthesis and/or accumulation of anthocyanins. A majority of these regulatory proteins belongs to two of the largest families of regulatory proteins in plants: the MYB and bHLH families (Lepiniec et a1., 2006) in addition to WD40, WRKY, WIP, Homeodomain, and bMADS families (Shi and Xie, 2010); The biosynthesis and accumulation of anthocyanins in response to light mainly occur due to transcriptional regulation of genes in the biosynthetic pathway (Martin and Gerats, 1993; Taylor and Briggs, 1990). For example, under high-intensity light conditions, accumulation of anthocyanins is induced in the 90 leaves and stems of Arabidopsis seedlings and coincides with an increase in chalcone synthase (CHS) activity, which is partly related to an increased rate of CHS transcription (Feinbaum and Ausubel, 1988). Differential regulation of transcription factors involved in biosynthesis adds another regulatory level to fine-tune anthocyanin deposition. Regulatory genes that are required for mediating anthocyanin biosynthesis can be expressed constitutively, e.g. TTGI, or can be induced by light (Cominelli et a1., 2008). Two such examples of light-induced genes are PIF 3 (Phytochrome-Interacting Factor 3, a member of the bHLH family) and H Y5 (Long Hypocotyl 5, a member of the bZIP factors). In R and FR light conditions, PIF 3 (Kim et a1., 2003) and HY5 (Ang and Deng, 1994; Somers et a1., 1991) play a positive role in regulating anthocyanin biosynthesis. Sequence analysis indicated that the promoters of all anthocyanin biosynthetic genes, in addition to the G-box elements, have multiple ACGT-containing elements and E-box elements, which are common to light-regulated genes (Shin, Park, and Choi, 2007). PIF3 and HY5 regulate anthocyanin biosynthesis by directly binding only to G-boxes and ACGT-containing sequence elements, respectively, within the promoters of anthocyanin biosynthetic genes and collaboratively regulate phyA-mediated anthocyanin biosynthesis (Shin, Park, and Choi, 2007). Moreover, PIF3-dependent regulation occurs by activation of the transcription of anthocyanin biosynthetic genes in a HY5-dependent manner (Shin, Park, and Choi, 2007). Reports have also shown that anthocyanin biosynthesis in Arabidopsis can be induced by different abiotic and biotic factors and induction in biosynthesis is characterized by notable changes in the transcripts of biosynthetic, as well as regulatory genes. An R2R3-MYB factor, PAP] (also called AtMYB 75; At1g56650) is an essential 91 regulatory protein of anthocyanin biosynthesis in Arabidopsis (Rowan et a1., 2009; Shi and Xie, 2010). In wild-type Arabidopsis seedlings, PAPI expression is induced by light (Cominelli et a1., 2008; Lea et a1., 2007; Lillo, Lea, and Ruoff, 2008; Solfanelli et a1., 2006; Teng et a1., 2005). PAP] stimulates light induction of anthocyanin biosynthesis in seedlings (Cominelli et a1., 2008; Stracke et a1., 2007) by regulating anthocyanin biosynthetic genes (Gonzalez et a1., 2008). Expression of PAP] is also induced by sucrose (Solfanelli et a1., 2006; Teng et a1., 2005). Interestingly, sucrose-induced accumulation of anthocyanin is marked by either induction or increase in the expression of PAPI (Lea et a1., 2007; Rowan et a1., 2009; Teng et al., 2005). Furthermore, PAP] expression and induction of anthocyanin biosynthesis by sucrose levels is positively correlated (Lillo, Lea, and Ruoff, 2008; Solfanelli et a1., 2006; Teng et a1., 2005). Notably, PAPI expression is induced after 6 hrs of exposure to R light (Cominelli et a1., 2008), indicating a possible role of phytochromes in the PAPI-regulated anthocyanin accumulation. While PIF3, HY5 and PAP] have a positive regulatory role, some of the MYB and bHLH regulatory proteins act as repressors of anthocyanin accumulation in Arabidopsis (Dubos et a1., 2008; Matsui, Umemura, and Ohme-Takagi, 2008; Yadav et a1., 2005; Zhu et a1., 2009). Anthocyanin biosynthesis is indeed subjected to environmental and developmental regulation and induction of biosynthesis and/or accumulation by abiotic and biotic stress factors requires proper coordination of synthesis and deposition in a spatial-, as well as a temporal-specific manner. Deposition of anthocyanins correlates with the transcript accumulation of corresponding early and late biosynthetic genes. The regulation of structural genes at the transcriptional level by regulatory proteins with 92 inductive and suppressive roles comprises a complex regulatory network of gene expression with positive and negative roles to fine-tune the biosynthesis, accumulation and deposition of anthocyanins in discrete tissues at a defined developmental stage in the life cycle of Arabidopsis. 3.1.2 Spatial-specific Accumulation of Anthocyanins Accumulation of anthocyanins occurs in specific tissues at discrete developmental stages and is strictly regulated (Tonelli et a1., 1994). Among the environmental factors that regulate this process are light, temperature, nutrients, and stress (Rabino and Mancinelli, 1986). Developmental information from such cues is transduced through complex signaling cascades to numerous genes that affect the synthesis, amount and distribution of anthocyanins within seedlings and adult plants. During de-etiolation, even though anthocyanin biosynthesis is limited to the outer cell layers of young seedlings, its accumulation predominantly occurs in the sub- epidermal layer of the hypocotyl and lower epidermis of the cotyledons (Huub, Kerckhoffs, and Kendrick, 1997). By contrast, in adult plants, anthocyanins are synthesized mainly in the outer cell layers of young developing leaves and the amount of anthocyanins is diluted as the leaf matures (Kerckhoffs et a1., 1992). Differential spatial- specific accumulation in young seedlings and adult plants suggests that anthocyanin deposition is developmentally regulated. In tomato, the anthocyanin responses occurring in the de-etiolation process display strong tissue specificity in the hypocotyls, restricted to a single layer of subepidermal cells (Neuhaus et a1., 1993). 93 Tissue-specific expression of certain regulatory genes is able to fine-tune the expression of structural genes, which in turn mediates anthocyanin accumulation in distinct tissues. In maize, regulatory genes that control transcription of structural genes of the anthocyanin biosynthetic pathway exert spatial and temporal regulation of anthocyanin accumulation in numerous tissues (Procissi et a1., 1997). Light-inducible tissue-specific activation of myb class genes regulates the spatial-specific anthocyanin biosynthesis at different developmental stages in maize seeds. For example, an ACGT- containing sequence element, upstream of the transcription start site of myb genes, C1 and p1, confers light inducibility (Cone et a1., 1993; Kao et a1., 1996). RT-PCR analysis of pl and CI transcripts indicated that expression of these regulatory genes is restricted to the pericarp and aleurone layer, respectively (Procissi et a1., 1997). Light-inducible tissue-specific activation of p1 regulates pigmentation in the pericarp during the early stages of maize seed development, whereas C1 regulates pigmentation in the aleurone layer during the advanced stages (Procissi et a1., 1997). An example of a regulatory protein conferring spatial-specific activation of anthocyanin biosynthetic genes is a bHLH family protein, Transparent TESTA8 (TT8) that modulates the expression of certain genes of the late flavonoid biosynthetic pathway in Arabidopsis siliques (Mehrtens et a1., 2005). Therefore, differential expression of structural genes and regulatory genes in discrete tissues play a major role in regulating spatial-specific anthocyanin biosynthesis and accumulation in planta. Transcriptional regulation of anthocyanin biosynthetic genes is subjected to both spatial and temporal regulation through the activities of regulatory proteins. Previously published data support the existence of spatial-, as well as temporal-specific de novo 94 synthesis and/or accumulation of anthocyanins in Arabidopsis and other plant species (Borevitz et a1., 2000; Kubo et a1., 1999; Nesi et a1., 2000). Despite the extensive amount of work reported on the spatial-specific biosynthesis and accumulation of anthocyanins, the mechanisms that control spatial and temporal regulation of anthocyanin accumulation have not yet been fully elucidated in Arabidopsis. Both phytochromes and cryptochromes regulate anthocyanin biosynthesis and accumulation in plants (Ahmad, Lin, and Cashmore, 1995). Phytochromes and cryptochromes that are localized in specific tissues and/or organs are able to mediate distinct light-dependent responses (Endo et a1., 2007; Montgomery, 2009; Wamasooriya and Montgomery, 2009). Localized pools of photoreceptors may regulate anthocyanin synthesis and disposition in which they perceive light or can mediate at distant sites from the site of photoperception. This phenomenon suggests that complex signaling mechanisms exist between perception of light and anthocyanin accumulation in specific tissues within plants. Whether the spatial- specific accumulation of photoreceptors contributes to the synthesis and deposition of anthocyanins in discrete tissues or organs and the functions of different phytochrome isoforms in regulating anthocyanin accumulation under different fluences of R/FR light are yet to be analyzed. 3.1.3 Functions of Phytochromes in Spatial-specific Anthocyanin Accumulation Light-dependent anthocyanin biosynthesis in seedlings displays characteristics of typical phytochrome-mediated HIR (Mancinelli, 1985). Analysis of photoreceptor mutants has enabled understanding of specific physiological functions mediated by phytochrome isoforms in phytochrome-dependent anthocyanin deposition under different 95 light conditions. In FR light, the HIR of anthocyanin accumulation was completely absent in a phyA mutant, whereas a phyB mutant displayed no significant reduction (Kunkel et a1., 1996). Therefore, in FR light, phyA is solely responsible for anthocyanin accumulation (Kunkel et a1., 1996). It was also reported that phyA has a significant role in the induction of anthocyanin accumulation in B (Duck and Fankhauser, 2003; Neff and Chory, 1998). Under R illumination, 3 phyB mutant had only a slight reduction in anthocyanin levels compared to WT or a phyA mutant. This observation suggests that, phyA or other phytochromes, but not phyB, play a significant role in R illumination (Kunkel et a1., 1996). In contrast to the evidence for the role of phyA in R illumination in regulating anthocyanin synthesis, phyB] of tomato is responsible for a significant proportion of anthocyanin biosynthesis in R (Kerckhoffs et a1., 1997). A reduction of anthocyanin levels reported for a phyD mutant in the Wassilewskija (Ws) ecotype under Wc, was not evident in a phyD mutant in the Landsberg erecta (Ler) ecotype, indicating that anthocyanin accumulation is primarily regulated by phyB in the Ler background (Aukerman et a1., l997). Currently, no reports have been published on the impact of phyC or phyE on anthocyanin accumulation. Spatially localized pools of phytochromes are known to contribute to the photoregulation of distinct phytochrome-dependent responses (Endo et a1., 2007; Endo and Nagatani, 2008; Endo et a1., 2005; Montgomery, 2009; Wamasooriya and Montgomery, 2009). In Arabidopsis, exposure to high-intensity light conditions is known to induce accumulation of anthocyanins in the leaves and stems via an increased rate of CHS transcription and it is possible that this response is mediated by a photoreceptor (Feinbaum and Ausubel, 1988). Distinct patterns of anthocyanin deposition can be 96 attributed to the members of the phytochrome family in Arabidopsis and tomato (Kerckhoffs et a1., 1997; Wamasooriya and Montgomery, 2009). Based on the fact that both phytochromes and anthocyanins are localized in distinct tissues during different developmental stages of plants, localized pools of phytochromes may have specific physiological functions related to anthocyanin synthesis, accumulation, and/or deposition. In fact, in Arabidopsis, mesophyll-localized phyA is known to regulate F R- dependent induction of anthocyanins and additional mesophyll-localized phytochromes, apart from phyB, have a role in R-induced anthocyanin accumulation (Wamasooriya and Montgomery, 2009). Despite the information available on the spatial- and temporal- specific regulation of phytochrome activity in mediating anthocyanin synthesis and deposition, conclusive reports on whether a particular localized pool of phytochromes can regulate intra- and/or inter-tissue accumulation of anthocyanins under R light have not yet been published. 3.1.4 Outlook Prior investigations into phytochrome-dependent regulation of anthocyanin accumulation revealed that mesophyll-specific phyA regulates F R-mediated induction of anthocyanin accumulation (Wamasooriya and Montgomery, 2009). The objective of this chapter is to utilize targeted phytochrome inactivation to further analyze the sites of photoperception and gain insight into the roles of phytochrome isoforms in sucrose- stimulated, light-dependent anthocyanin accumulation under Rc conditions. Transgenic Arabidopsis lines with tissue-specific chromophore deficiencies are probed under Rc for phenotypic perturbations. In contrast to the role of mesophyll-localized phyA in 97 regulating F R-mediated induction of anthocyanin accumulation, under Rc, additional mesophyll-localized phytochromes, in addition to phyB, mediate R-induced anthocyanin accumulation. Given the prior investigations into the functions of phyA and phyB in Arabidopsis and tomato in light-inducible anthocyanin accumulation, the analysis of apophytochrome mutants revealed novel roles for phytochrome isoforms in Re conditions: phyA, B, C and, D essentially regulate the induction of anthocyanin accumulation, whereas phyE imposes suppression on anthocyanin accumulation in Arabidopsis. 3.2 Materials and Methods 3.2.1 Quantification of Anthocyanin Levels in Transgenic Lines with Targeted Chromophore Inactivation Seeds of No-O WT, 3SS::pBVR3, CAB3::pBVR2, Col-0 WT and phyB (SALK_ 022035; Ruckle, DeMarco, and Larkin, 2007) were sterilized and planted as described in section 2.2.], and were subjected to treatment with R pulse (approximately 75 umol m-2 s-l) for 5 min prior to imbibition. Imbibing seeds were cold-stratified at 4 °C for 3 days in darkness. Plates were transferred to a temperature- and humidity-controlled growth chamber with Rc illumination of 50 umol m-2 s-1 or in darkness for 4 days at 20 oC. Anthocyanins were extracted from whole-plant seedlings and quantified as described in section 2.2.2. 98 3.2.2 Confirmation of Apophytochrome Mutants To gain insight into the differential roles of phytochrome isoforms in Rc conditions, apophytochrome mutants, each with a T-DNA insertion in an exon of the respective genes were selected from the Salk T-DNA insertion mutant collection (Alonso et a1., 2003). Seeds of Col-0 WT, phyA (SALK_014575; (Ruckle, DeMarco, and Larkin, 2007), phyB (SALK_ 022035; (Ruckle, DeMarco, and Larkin, 2007), phyC (SALK_007004), phyD (SALK_027336) and phyE (SALK_092529) were sterilized as described in section 2.2.1. Sterilized seeds were treated with R pulse (approximately 75 umol m-2 s!) for 5 min prior to imbibition. Imbibing seeds were cold-stratified at 4 °C for 3 days in darkness. Plates were transferred to a temperature- and humidity-controlled growth chamber with RC illumination of 50 umol m-2 s-1 for 4 days at 22 °C. 4-d-old whole seedlings were quickly (< 1 min) harvested and immediately frozen in liquid nitrogen inside the R light chamber. Using RNeasy® Plant Minikit (Catalog No. 16419, Qiagen, CA) including on-column DNase treatment (Catalog No. 79254, Qiagen, CA), total RNA was isolated according to manufacturer’s instructions. The quantity of RNA was analyzed by spectrometry (NanoDroplOOO, Therrno Scientific, MA). Oligo(dT)15 primed-first-strand cDNA was synthesized from 1 ug of total RNA using a Reverse Transcription System (Catalog No. A3 500, Promega, WI) according to manufacturer’s instructions with the modifications as described in section 2.2.4. cDNA was stored at -20 °C overnight before PCR amplification. First-strand cDNA synthesis reactions were diluted 1:16 with nuclease-free water. PCR was conducted with GoTaqGreen (Catalog No. M7123, Promega, WI) and 4 u] of the diluted cDNA product was used as template in 99 a 25 u] reaction. The gene-specific oligonucleotides for respective genes were designed using a free online database, AtRTPrimer (Han and Kim, 2006); PH YA (At1g095 70)— forward 5’-AAGGAGAATGCACCCAAGGTCATC-3’ and reverse 5’- CACCTTCAATCCGCGTAAACTTGTC-3’, PHYB (At2gl8790)— forward 5’- TCGAGGGAAAGGTTATTGGGGCTTT-3’ and reverse 5’- GGAACATGTCTCGGACTAGCTCTGG-3 ’, PHYD (At4g16250)— forward 5 ’- GATCGCAAAGGGGAATTCATTCAGG-3’ and reverse 5’- TTCCATGGGTGCATAACGGACA-3 ’and PHYE (At4g18130)— forward 5 ’- GCTTACGGGATGGTCAAAACACGA-3’ and reverse 5 ’- GGCGACTTCAACCCTTAGTTGTGAG-3 9. The gene-specific oligonucleotides for PHYC (At5g35840) were designed based on the full length cDNA sequence: PH Y C— forward 5 ’-CCCTCAACAAATTGGCATATCTCCGCC-3 ’ and reverse 5 ’- AGATCCTCAGGCAGTCCTGGTGC-3 ’. PCR amplification was carried out with gene- specific oligonucleotides at 10 uM. The following thermal cycling conditions were used for the PCR amplification: (1) 1 cycle of denaturation at 94 °C for 2 min, (2) 30 cycles of denaturation at 94 °C for 30 s, annealing for 45 s (for PH YA, PH YB and PH YE at 60 °C, for PHYC at 61 °C and PHYD at 59 0C), extension at 72 °C for 38 s and (3) final extension at 72 °C for 5 min with a hold at 4 °C. Expression of UBC21 (At5g25 760) was analyzed as an internal control by including UBC21-specific primers, forward 5 ’- CCTTACGAAGGCGGTGTTTTTCAG-3’ and reverse 5’- 100 CGGCGAGGCGTGTATACATTTG-3’) at 10 uM in the same reaction. To rule out the possibility of dependency of transcript accumulation on the number of PCR cycles, amplification with respective gene-specific oligonucleotides was repeated at 45 cycles. A 10 uL aliquot of the PCR product was visualized by electrophoresis for 2 h at 80V on a 1.5% agarose gel containing ethidium bromide at 0.02 ug/mL (Catalog No. 15585-011, Invitrogen, CA). Ultraviolet images were obtained using the Ge] Doc system (Bio-Rad Laboratories, Inc., CA) at subsaturation settings. 3.2.3 Quantification of Anthocyanin Levels in Apophytochrome Mutants Levels of anthocyanins were quantified in apophytochrome mutants that were confirmed to be null in section 3.2.2 according to the protocol described in section 2.2.2, to gain further insight into the roles of individual phytochrome isoforms in the regulation of light-dependent anthocyanin accumulation in Arabidopsis. 3.2.4 Expression Levels of Anthocyanin Marker Genes Two MYB genes, MYB75/PAP1 and MYB90/PAP2 are known to regulate anthocyanin biosynthesis, but only MYB75/PAP1 is required for sucrose-induced anthocyanin accumulation (Borevitz et a1., 2000). According to (Teng et a1., 2005), MYB75/PAP1 gene is an important quantitative trait locus (QTL) for sugar-induced anthocyanin induction in Arabidopsis. DFR is a late biosynthetic gene and specific for the anthocyanin branch of the flavonoid biosynthetic pathway (Kubasek et a1., 1992). An active DF R enzyme is essential for anthocyanin biosynthesis and MYB75/PAPI is required for sucrose-stimulated DFR expression (Teng et a1., 2005). Among the types of 101 sugars that can induce MYB75/PAP1 and DFR, sucrose appears to be the most effective trigger for both, however, the structural genes upstream of DFR display lower induction by sucrose and can also be induced, to a slight degree, by other sugars (Solfanelli et a1., 2006; Teng et a1., 2005). Due to the requirement of MYB75/PAPI and DFR for sucrose- induced anthocyanin accumulation in Arabidopsis, MYB75/PAP1 and DFR were selected as anthocyanin marker genes for transcript analysis by RT-PCR. The levels of MYB75/PAPI and DFR transcript accumulation in the presence or absence of sucrose was analyzed by RT-PCR to determine if the changes observed in the levels of anthocyanin accumulation in apophytochrome mutants are reflected at the transcript level of selected anthocyanin marker genes. Seeds of Col-0 WT and apophytochrome mutants (phyA, phyB, phyC, phyD and phyE) were sterilized and planted as described in section 2.2. 1. Sterilized seeds were treated with R light pulse (approximately 75 umol rn-2 5.1) for 5 min prior to imbibition. Imbibing seeds were cold- stratified at 4 °C for 3 days in darkness. Plates were transferred to a temperature- and humidity-controlled grth chamber with Rc illumination of 50 umol m-2 s-1 for 4 days at 22 °C. Extraction and quantification of total RNA were performed as described in section 3.2.2. cDNA was synthesized from 1 ug of total RNA using Reverse Transcription System (Catalog No. A3500, Promega, WI) according to manufacturer’s instructions with the modifications as described in section 2.2.4. F irst-strand cDNA synthesis reactions were diluted 1:16 with nuclease-free water. PCR was conducted with GoTaqGreen (Catalog No. M7123, Promega, WI) and 4 u] of the diluted cDNA product was used as template in a 25 u] reaction. PCR amplification was carried out with the following gene-specific oligonucleotides at 10 uM: MYB75/PAP1— forward 5’- 102 GCTCTGATGAAGTCGATCTTCJ’ and reverse 5 ’- CTACCTCTTGGCTTTCCTCT- 3’, DFR] — forward 5 ’- GGTTTCATCGGTTCATGGCT-3’ and reverse 5 ’- GGTTTCATCGGTTCATGGCT-3 ’ based on Teng et a1., (2005) and Cominelli et al., (2008), respectively. The following thermal cycling conditions were used for the PCR amplification. For MYB75/PAP1 (1) 1 cycle of denaturation at 94 °C for 2 min, (2)40 cycles of denaturation at 94 °C for 30 s, annealing at 56.5 0C for 45 5, extension at 72 0C for 3 min and (3) final extension at 72 0C for 10 min with a hold at 4 °C (Teng et a1., 2005). For DFR], (l) 1 cycle of denaturation at 94 °C for 3 min, (2) 30 cycles of denaturation at 94 °C for 30 s, annealing at 57 0C for 35 3, extension at 72 °C for 45 s and (3) final extension at 72 °C for 5 min with a hold at 4 oC. Expression of At5g25 760 (UBC21) was analyzed as an internal control by including UBC21-specific primers as described in section 3.2.2. A 7-uL aliquot of the PCR product was visualized by electrophoresis for 2 h at 80V on a 1.5% agarose gel containing ethidium bromide. Ultraviolet images were obtained using the Ge] Doc system (Bio-Rad Laboratories, Inc., CA) at subsaturation settings. Levels of transcript accumulation were quantified using Quantity One®, Version 4.6.3, Bio-Rad Laboratories, Inc., CA). 3.2.5 Complementation of phyE Mutant To test if the phenotype with respect to increased levels of anthocyanin in the phyE null mutant can be restored to WT levels, the phyE null mutant was complemented with a construct containing a transgene encoding C-terminal myc epitope-tagged phyE driven by the PH YE native promoter. pBI-PpHygtphyE-myc6 (PH YE-mo hereafter) plant 103 transformation construct was obtained from Robert A. Sharrock (Clack et a1., 2009). The PH YE-m6 construct was initially introduced into the GV3101pM90 strain of A grobacterium tumefaciens by electroporation (Weigel and Glazebrook, 2006). Transformants were selected on YEP medium containing kanamycin (50 ug/mL; Catalog No. K-4378, Sigma, MO), gentamycin (50 ug/mL; Catalog No. 61-098-RF, Cellgro®, Mediatech, Inc.,VA) and rifampicin (20 ug/mL; Catalog No. R8883, Sigma, MO). Arabidopsis (Arabidopsis thaliana) phyE mutant (SALK_092529) in the Col-0 WT background was transformed with GV3101 (PH YE-m6) using standard Agrobacterium- mediated floral dip transformation (Clough and Bent, 1998). Kanamycin selection of transfonnants was performed in 150- x lS-mm petri dishes containing 1X Murashige and Skoog salts (Catalog No. MSP09, Caisson Laboratories, UT), 0.6 % (w/v) Phytablend (Catalog No. PTPO], Caisson Laboratories, UT), 1 % (w/v) Sucrose (Catalog No. 4072- 05, J.T. Baker, NJ) with kanamycin (Catalog No. K-43 78, Sigma, MO) at 50 ug/mL. T1 transgenic seedlings were selected through a rapid selection protocol previously described for identifying transformed Arabidopsis seedlings following floral dip transformation (Harrison et a1., 2006). Since the phyE mutant (SALK_092529) turned pale green/yellow on media containing kanamycin under We, indicating that the kanamycin resistance gene is co-suppressed in the phyE mutant background, segregation analysis on complemented phyE mutants was performed as described above for selection of primary transformants. T2 seedlings segregated for kanamycin resistance in a 3:1 ratio, which is consistent with the resistance being determined by a single insertion. Based on kanamycin resistance, T3 seedlings that were homozygous for a single T-DNA insertion 104 were selected to obtain seeds for further analyses on complemented phyE mutants (PHYE-m6/phyE-I and PHYE-m6/phyE-11). 3.2.6 Arabidopsis Seedling Extracts Seeds of Col-0 WT, complemented phyE mutants (PH YE-m6/phyE-1 and PH YE- m6/phyE-11), and phyE mutant were sterilized, planted and subjected to cold stratification as described in section 2.2.1. Plates were kept in a humidity—controlled chamber with Rc illumination of 50 umol m.2 s-1 for 10 days at 22 °C. Whole-seedling tissues were quickly harvested (< 1 min), weighed and transferred to individual 15 mL sterile tubes. After immediately freezing in liquid nitrogen, tissues were crushed to a powder using a microgrinder. Crude Arabidopsis extracts were prepared according to a protocol adapted from Lagarias et a1., (l997); whole-seedling tissues were immediately homogenized in plant extraction buffer: 50 mM Tris-HCI, pH 8.0 (Catalog No. 15568- 025, Invitrogen, CA), 100 mM NaCl (Catalog No. 24740-0] 1, GIBCO®, Invitrogen Co., NY), 1 mM EDTA (Catalog No. 15575-038, GIBCO®, Invitrogen Co., NY), 1 mM EGTA (Catalog No. E3889, Sigma, MO), 143 mM 2-mercaptoethanol (Catalog No. M3148, Sigma, MO), 1 mM phenylmethanesulfonyl fluoride (Catalog No. P7626, Sigma, MO), 1 % dimethylsulfoxide (Catalog No. 9224-01, J. T. Baker, NJ), 1X protease inhibitors (Catalog No. 11-836-170-001, Roche, IN), 5 % glycerol (Catalog No. 15514- 01], Invitrogen, CA), the latter five components were added just before use. For homogenization, plant extraction buffer was added at a ratio of 3 volumes/ fresh weight (mg). The crude homogenates in 15 mL tubes were centrifuged at 4750 rpm for 2 min at 105 4 °C (Allegra® X-15R Centrifuge, Beckman Coulter, CA) to pellet debris and partially clarified supematants were transferred to microcentrifuge tubes on ice followed by centrifugation at 13,000 rpm for 15 min at 4 oC (Sorvall® Fresco, Heraeus, NC). Aliquots of clarified supematants were stored at -80 °C for immunoblot analysis as described in section 3.2.7. 3.2.7 Expression of PH YE-m6 in Complemented phyE Mutant Total soluble proteins extracted from lO-day-old whole-seedlings were quantified as reported (Wamasooriya and Montgomery, 2009). Protein (~ 60 pg) was separated by SDS-PAGE and subsequent immunoblot analyses were performed as described (Montgomery et a1., 1999) using rabbit anti-myc (1:2000; Catalog No. 2278, Cell Signaling, MA). Secondary antibody incubation was performed with goat anti-rabbit IgG (H+L) conjugated to horseradish peroxidase (HRP; 1:4000, Catalog No. 7074, Cell Signaling, MA). Antibody signal (chemiluminescence) was detected using SuperSignal® West Dura Extended Duration substrate (Catalog No. 34075, Therrno Fisher Scientific ® TM . . Inc., IL) on Molecular Imager VersaDoc MP 4000 System (Bio-Rad Laboratories Inc., CA). 3.2.8 Quantification of Anthocyanin Levels in Complemented phyE Mutant Seeds of Col-0 WT, complemented phyE mutants (PH YE-m6/phyE—1 and PH YE- m6/phyE-I I), and phyE mutant were sterilized, planted and subjected to cold stratification as described in section 2.2.1. Plates were kept in a humidity-controlled 106 chamber with Rc illumination of 50 umol m-2 s-1 for 4 days at 22 °C. Levels of anthocyanins were quantified according to the protocol described in section 2.2.2. Two- tailed, unpaired Student’s t-test was performed to compare the anthocyanin content relative to Col-0 WT and phyE mutant seedlings. 3.3 Results and Discussion As previously reported, constitutive expression of BVR inhibits sucrose- stimulated anthocyanin synthesis in transgenic Arabidopsis plants (Montgomery et a1., 2001; Montgomery et al., 1999) and suggests a regulatory role of phytochromes in anthocyanin biosynthesis. Recent analysis of transgenic lines displaying mesophyll- localized phytochrome deficiencies indicated that mesophyll-localized phyA regulates F R-mediated induction of anthocyanin accumulation (Wamasooriya and Montgomery, 2009). Analyses of the phyA mutant and CAB3::pBVR lines suggested that phyA in the mesophyll is solely responsible for the regulation of anthocyanin accumulation, as anthocyanin levels in CAB3::pBVR lines and the phyA mutant were similar (Wamasooriya and Montgomery, 2009). The objective of this chapter was to utilize targeted expression of BVR using tissue-specific promoters to limit the BVR activity to a particular tissue. This novel experimental approach was utilized to investigate the sites of photoperception for phytochrome-dependent anthocyanin accumulation in transgenic Arabidopsis plants. 3.3.1 Spatial-specific Phytochromes Regulate Anthocyanin Accumulation in Re Accumulation of anthocyanins occurs in specific tissues at discrete developmental 107 stages (Tonelli et a1., 1994) and a transient developmental peak in flavonoid accumulation occurs when young seedlings are ~ 4-days-old (Kubasek et a1., 1992). Therefore, in Arabidopsis, to investigate the impact of monochromatic Rc illumination and determine the roles of localized pools of phytochromes on sucrose-induced accumulation of anthocyanins in seedlings, anthocyanin levels were quantified in 4-d-old representative 35S::pBVR3, CAB3::pBVR2 and null phyB T-DNA insertion mutant (Ruckle, DeMarco, and Larkin, 2007) and the cognate wild-type plants. Anthocyanin accumulation in Re light was low for all lines compared to their cognate WT plants in the presence, as well as in the absence of sucrose (Figure 3.1). However, in the 35S::pBVR3 line, the anthocyanin levels were not significantly reduced compared to No-O WT (p=0.6253), while the CAB3::pBVR2 line displayed significant reduction compared to No-O WT (p<0.0001, Figure 3.1). This observation suggests that in Re illumination, inactivation of phytochromes in mesophyll tissues has a more pronounced effect on anthocyanin accumulation at whole-seedling level than the constitutive inactivation of phytochromes. More pronounced effect on anthocyanin accumulation upon mesophyll-localized phytochrome inactivation as compared to constitutive inactivation could also arise due to the differences in the levels of BVR expression in cotyledons and tissue specificity of cotyledon tissues vs. mesophyll tissues between 3SS::pBVR3 and CAB3::pBVR2 lines. Higher level of BVR expression in the cotyledons of CAB3::pBVR2 compared to the level in the cotyledons of 35S::pBVR3 could lead to a drastic reduction of phytochromes within cotyledons causing CAB3::pBVR2 line to have a lesser amount of anthocyanins than all of the other lines, including the phyB mutant compared to its wild-type parent, Col-0 WT (Figure 3.1). The 108 phyB mutant displays a significant reduction in the levels of anthocyanin compared to Col-0 WT in Rc illumination (p=0.0383) indicating that phyB has a role in the induction of anthocyanin accumulation in Re as previously reported (Kunkel et a1., 1996) . The response of CAB3::pBVR2 compared to No-O WT is different from that observed for the phyB null mutant (in comparison to Col-0 WT) and hence suggests that phyA and/or light-stable phytochrome isoforms other than phyB have a role in inducing anthocyanin accumulation under Rc illumination. Anthocyanins are actively sequestered into cell vacuoles for ergastic storage and in most species, they accumulate predominantly in leaf mesophyll (Gould et a1., 2000) and epidermal cells (Kubo et a1., 1999). Analysis of light- and/or sucrose-inducible anthocyanin accumulation in CAB3::pBVR lines suggests that mesophyll-localized phytochromes are responsible for the induction of anthocyanins in mesophyll tissues in Arabidopsis, and may impact inter-tissue anthocyanin accumulation in the epidermis under Rc. 3.3.2 Phytochrome Family Members Have Differential Roles in Anthocyanin Accumulation in Re According to previously published results, phyB is primarily responsive to R wavelengths (Reed et a1., 1993). phyA was also recognized to play a significant regulatory role with respect to anthocyanin accumulation in R based on the observation that a phyA mutant had a notable reduction in anthocyanin levels compared to WT in contrast to a slight reduction in phyB mutant (Kunkel et a1., 1996). However, phyB] of tomato is responsible for a significant proportion of anthocyanin biosynthesis in R 109 (Kerckhoffs et a1., 1997). These observations suggest that phytochrome family members may have differential roles in mediating the anthocyanin accumulation response. To determine which phytochrome isoform(s) may be involved in the regulation of anthocyanin levels in Re, that anthocyanin levels were quantified in 4-d-old Rc-grown seedlings of apophytochrome mutants. Single apophytochrome mutants obtained from the Salk T-DNA insertion mutant collection were assessed for the absence of respective transcript accumulation by RT-PCR analysis. In the selected T-DNA insertion mutants, the T-DNA was inserted in the exon region of each apophytochrome gene (Figure 3.2). No respective transcript was detected by RT-PCR in apophytochrome mutants and this confirmed that T-DNA insertion mutants were null (Figure 3.3). Levels of anthocyanin accumulation were quantified in the confirmed null mutants under Rc illumination following a 4-d growth period and this analysis revealed that the phyA, phyB, phyC and phyD mutants displayed lower accumulation of anthocyanins than Col-0 WT (Figure 3.4 C). As previously published, the total levels of anthocyanins are lower for Col-0 WT in Re than Wc (Neff and Chory, 1998; Wamasooriya and Montgomery, 2009). In the presence of sucrose, anthocyanin levels in the phyA, phyB, phyC and phyD mutants were ~ 76 %, 52 % 66 % and 86 % respectively, relative to Col-0 WT, indicating that the levels measured in these mutants were further reduced under Rc. The reduction in the anthocyanin levels for the phyB and phyC mutants in comparison to Col-0 WT were significant (p=0.0004 and p=0.0079, respectively). Although the levels in the phyA and phyD mutant were lower with respect to Col-0 WT, levels for the phyA mutant were not quite significantly different (p=0.0559) and the phyD mutant was not significantly different (p=0.323 8). However, compared to Col-0 WT, the phyE mutant accumulated 110 significantly more anthocyanins (p=0.0022, Figure 3.4 C). In the phyE mutant, levels were ~ 67 % higher than Col-0 WT in the presence of sucrose and anthocyanins were clearly visible at the junction of hypocotyls and cotyledons (Figure 3.4 A). Elevated levels of anthocyanin accumulation is a characteristic of a hyperphotomorphogenic phenotype as previously observed in a transgenic line overexpressing a mutant form of HY5 (HY5-AN77) in We, Be, Rc and F Rc (Ang et a1., 1998). The analysis of anthocyanin accumulation levels in apophytochrome mutants indicated that phyA, phyB, phyC, and phyD contribute to the light—dependent, sucrose-stimulated accumulation of anthocyanins, whereas phyE has a role in repressing anthocyanin synthesis and/or accumulation under Re. The lowest levels of anthocyanin accumulation in the phyB mutant suggest that, in Re, phyB is having a major quantitative role in the induction of anthocyanins. Under Rc, in the absence of sucrose, anthocyanin levels for the phyC and phyD mutants were less compared to Col-0 WT and were not quite significant (p=0.4468, p=0.2336, respectively, Figure 3.4 C). However, the reduction observed in the phyB mutant was extremely significant (p<0.000]) compared to WT and confirms that phyB indeed has a major role in sucrose-stimulated anthocyanin accumulation in Re. Notably, the phyA and phyE mutants accumulate more anthocyanins (~ 29 %, ~79 %, respectively) in the absence of sucrose compared to WT. This increase in anthocyanin levels was quite significant in the phyA mutant (p=0.0086) and extremely significant in the phyE mutant (p<0.0001) with respect to Col-0 WT. Anthocyanin levels were compared in apophytochrome mutants with respect to l % sucrose vs. 0 % sucrose to determine if a specific member of the phytochrome family could have a role in sucrose-induction of anthocyanin accumulation. Col-0 WT and all 111 apophytochrome mutants accumulated lower levels of anthocyanins in the absence of sucrose. The fold increase with respect to 1 % sucrose vs. 0 % sucrose was 2.0 for Col-0 WT and 1.8 for the phyB, phyD and phyE mutants (Figure 3.4C). Anthocyanins were visible only in the phyE mutant regardless of the presence or absence of sucrose (Figure 3.4 A and B). The anthocyanin levels with respect to 1 % sucrose vs. 0 % sucrose was quite significant in the Col-0 WT and phyE mutant (p=0.0004, p=0.0002, respectively) and was extremely significant in both the phyB and phyD mutants (p<0.0001). The fold increase for the phyA and phyC mutants was 1.2 and 1.4, respectively (Figure 3.4 C) and the difference in anthocyanin levels with respect to l % sucrose vs. 0 % sucrose was significant for the phyC mutant (p=0.0111), but not significant for the phyA mutant (p=0.2474). In the phyA mutant, the accumulation of similar levels of anthocyanins regardless of the presence or absence of sucrose in the medium may imply a lack of sucrose stimulation and thus suggest a possible requirement of phyA for sucrose- induction of anthocyanin accumulation in Re. An apparent difference in the fold increase of anthocyanins was not observed in the phyE mutant compared to Col-0 WT with respect to l % sucrose vs. 0 % sucrose. This observation may indicate that suppression of phyE on anthocyanin accumulation is independent of the presence of sucrose in the medium. To confirm that a lack of phyE is responsible for the altered anthocyanin levels observed in the phyE mutant line and thus phyE is a true suppressor of anthocyanin accumulation in Arabidopsis, the phyE mutant was complemented with a C -terminal myc-tagged PH YE construct, i.e., ProPHygsPHYE-md Based on kanamycin resistance, T3 seedlings that were homozygous for a single transgene insertion were selected to 112 obtain seeds for further analyses. In T3 seedlings for two independent complemented transgenic lines, i.e., PH YE-m6/phyE-1 and PH YE—m6/phyE-1 1, a reduction of anthocyanin levels in seedlings grown under Rc was observed (Figure 3.5). The degree of complementation correlated well with levels of myc-tagged phyE protein accumulating in the complemented transgenic lines (Figure 3.6). The observation of reduction in anthocyanin levels was noticeably true for complemented transgenic lines grown under Rc in the presence of sucrose. Complemented line PH YE-m6/phyE—I I , which had a greater accumulation of myc-tagged phyE protein than line PH YE-m6/phyE-1 , exhibited a greater reduction of anthocyanin accumulation in the presence of sucrose, i.e. ~72.5 % of that of the phyE parent, than did PH YE-m6/phyE-I , which exhibited ~82.5 % of the level of anthocyanins measured for phyE (Figure 3.5). The reductions observed for sucrose-dependent anthocyanin accumulation for the complemented PH YE-m6/phyE-1 and PH YE-m6/phyE-Il lines relative to the phyE mutant were significant, i.e., p=0.0001 and p<0.0001, respectively, as compared to the ~58 % level of anthocyanin accumulation observed for Col-0 WT relative to the phyE mutant (p<0.0001). Based on analyses of transgenic plants with mesophyll-specific phytochrome deficiency and subsequent analysis of single phy mutants in all five of the apophytochrome genes, all phytochrome family members were shown to contribute to the light-dependent, sucrose-stimulated accumulation of anthocyanins, but with divergent regulatory roles. Notably, phyE is unique in its role of suppressing anthocyanin synthesis and/or accumulation under Rc, whereas other members of the phytochrome family stimulate anthocyanin accumulation under these conditions. These results are of particular interest as, to date, few repressors of anthocyanin biosynthesis have been 113 reported. A bHLH transcription factor, AtMYC2/J IN] is a repressor of anthocyanin biosynthesis since the anthocyanin levels in the atmyc2-3 mutant are higher in FR (Yadav et a1., 2005; Zhu et a1., 2009). Zhu et a1., (2009) reported that a single-repeat R3 MYB transcription factor like CPC (CAPRICE) is a negative regulator of anthocyanin biosynthesis and confers suppression by competing with R2R3-MYB proteins- i.e., MYB75/PAP] and MYB90/PAP2. Increased levels of anthocyanin accumulation has been observed in the loss of function of a mutant of AtMYBL2 (myblZ) and the suppression of anthocyanin accumulation by AtMYBL2 occurs through negative regulation of the expression of structural and regulatory genes of anthocyanin biosynthesis in Arabidopsis seedlings (Dubos et a1., 2008; Matsui, Umemura, and Ohme- Takagi, 2008). 3.3.3 Anthocyanin Marker Genes are Differentially Expressed in Rc According to previously published data, MYB75/PAP1 is essential (Borevitz et a1., 2000) and an important QTL for sucrose-induced anthocyanin accumulation in Arabidopsis (Teng et a1., 2005). DFR is specific for the anthocyanin branch of the flavonoid biosynthetic pathway (Kubasek et a1., 1992) and an active DF R enzyme is essential for anthocyanin biosynthesis (Teng et a1., 2005). Sucrose appears to be the most effective trigger in inducing anthocyanin accumulation for both MYB 75/PAP1 and DF R (Solfanelli et a1., 2006; Teng et a1., 2005) and MYB75/PAP1 is required for sucrose- stimulated DFR expression (Teng et a1., 2005). Thus, MYB75/PAP1 and DFR were ideal candidates to determine if the differential accumulation of anthocyanins observed in apophytochrome mutants correlates with steady-state accumulation of MYB 75/PAP1 and 114 DF R transcripts in the presence/absence of sucrose in 4-d-old Rc-grown apophytochrome mutants. The analysis of steady-state levels of transcript accumulation for MYB 75/PAPI in apophytochrome mutants indicated that Col-0 WT, phyA, phyD and phyE mutants accumulate similar levels of MYB75/PAP1 transcript in the absence as well as, in the presence of ] % sucrose (Figure 3.7 A, B and 3.8 A). The phyC mutant accumulates a higher level of transcript on 1 % sucrose compared to 0 % sucrose (Figure 3.7 A and B). Quantification or relative transcript accumulation for the phyC mutant on 0 % vs. 1 % also indicated that in the presence of sucrose, the level of relative transcript accumulation for MYB75/PAP1 was higher. Notably, the level of relative transcript accumulation for MYB75/PAP1 was higher in the phyB mutant compared to rest of the lines (Figure 3.7 A, B and Figure 3.8 A) and the sucrose-induced transcript accumulation was the highest for phyB mutant (Figure 3.7 A and B). The phyB mutant displayed significantly higher levels of transcript accumulation for MYB75/PAP1 compared to Col-0 WT on 1 % sucrose (p=0.0007, Figure 3.7 A, B and Figure 3.8 A). In the phyB mutant, on 0 % vs. 1 % sucrose, the relative transcript accumulation level for MYB 75/PAP1 was significantly higher in the presence of sucrose (p=0.0422, Figure 3.7 A, B and Figure 3.8 A). This observation implies that the phyB mutant is responsive to sucrose in the medium. It was previously observed that MYB75/PAP1 transcript accumulates rapidly within 6 h under R light with an intensity of 125 umol m-2 3-1 indicating that MYB75/PAP1 transcript is light induced (Cominelli et a1., 2008). According to Teng et a1., (2005), the expression of MYB75/PAP1 is induced by a sucrose-induced signaling pathway and as a consequence, the anthocyanin biosynthetic pathway is activated. Despite the expression of 115 MYB75/PAP1 being a QTL for sucrose-induced anthocyanin accumulation in Arabidopsis as previously described (Teng et a1., 2005), compared to Col-0 WT, the levels of anthocyanins and relative transcript accumulation for MYB75/PAP1 in the phyB mutant were inversely correlated. Hence, the anthocyanin levels and MYB 75/PAP1 expression can not be correlated for the phyB mutant. This observation implies that sucrose-stimulated, light-dependent anthocyanin accumulation in Re requires a functional phyB, whereas induction of MYB75/PAP1 expression under Rc does not. In the phyB mutant, the remaining type II phytochromes are able to perceive R light and induce MYB75/PAPI expression. However, after sucrose and R light trigger the MYB75/PAPI expression, the positive regulatory role of MYB75/PAP1 on the late anthocyanin biosynthetic genes leading to an induction of anthocyanin biosynthesis possibly require functional phyB under Rc. Although phyB through phyE are primarily responsive to R light (Aukerman et a1., 1997; Monte et a1., 2003; Reed et a1., 1993) and phyB is the predominant type II phytochrome regulating R-HIR and LF R (Nagy and Schafer, 2002; Quail, 2002), functional phyC, phyD and phyE family members in the phyB mutant possibly perceive R light resulting in an induction of sucrose-stimulated MYB75/PAP1 expression under Rc. phyA may not contribute in this regard as it is only known to be an irradiance-dependent light sensor at very high fluences of R light of > 160 umol m-2 8.1 (Franklin, Allen, and Whitelam, 2007). The significantly higher levels of transcript accumulation for MYB75/PAP1 compared to Col-0 WT on 0 % vs. 1 % sucrose could also reflect a possible feedback regulatory role of anthocyanin levels on MYB 75/PAP1 transcript accumulation where Arabidopsis seedlings are responding to low anthocyanin 116 levels by trying to up-regulate biosynthesis through induction of MYB 75/PAP1 transcript accumulation. Analysis of transcript accumulation for DFR in single apophytochrome mutants indicated that DFR expression is higher in Col-0 WT and all of the single apophytochrome mutants in the presence of sucrose compared to barely detectable levels of DFR transcript accumulation in the respective lines on 0 % sucrose (Figure 3.7 C and D) and confirms that DFR expression is subjected to sucrose-dependent up-regulation as previously published (Solfanelli et a1., 2006). The level of DFR transcript accumulation observed in the phyA mutant was more or less similar to the level observed for Col-0 WT on 1 % sucrose (Figure 3.8 B) and possibly coincides with the similar levels of anthocyanins in the phyA mutant compared to Col-0 WT (p=0.0559, Figure 3.4 C). Quantification based on densitometry indicated that compared to Col-0 WT, the phyB mutant displayed similar levels of DFR transcript accumulation in the presence of sucrose. Compared to Col-0 WT, even though, the phyE mutant accumulated significantly higher level of anthocyanins on 0 % and l % sucrose (p<0.0001 and p=0.0022 respectively, Figure 3.4 C), the level of DFR transcript accumulation was not reflective of this observation (Figure 3.7 C). On 0 % vs. 1 % sucrose, the phyB mutant showed significantly higher levels of DFR transcript accumulation (p= 0.049], Figure 3.7 C, D and Figure 3.8 B). Despite the specificity of DFR gene for anthocyanin biosynthesis within the flavonoid biosynthetic pathway (Kubasek et a1., 1992), the difference in the level of accumulated DFR transcript may not correlate with the amount of anthocyanin levels when multiple signaling pathways, i.e. phytochrome and sucrose, are involved in determining the overall level of anthocyanin accumulation. 117 3.3.4 Summary The quantification of anthocyanin levels in transgenic BVR-expressing lines with mesophyll-specific phytochrome inactivation indicates that mesophyll-localized phytochromes regulate sucrose-stimulated accumulation of anthocyanins under Rc. The analysis of anthocyanin levels in single apophytochrome mutants suggests that all phytochrome isoforms contribute to the regulation of anthocyanin accumulation under R light. In this regard, disparate activities of different phytochrome family members on the accumulation of anthocyanins were identified, with phyE functioning as a suppressor and the remaining phytochromes acting as promoters of anthocyanin accumulation under Rc illumination. However, the suppression of phyE on anthocyanin accumulation response appears to be independent of sucrose. By contrast, in other studies, phyB and phyD were shown to negatively impact phyA-mediated seed germination in FR (Hennig et a1., 2001; Hennig et a1., 2002), however, phyE promotes phyA-mediated germination under FR (Hennig et a1., 2002). Thus, fine-tuning of distinct aspects of photomOrphogenesis by opposing activities of individual phytochrome isoforms is not limited to the observation of the impact of phytochrome family members on anthocyanin accumulation under Rc. Among the phytochrome isoforms, phyB has the greatest quantitative role in the induction of anthocyanins under Rc (Figure 3.4 C). Although phyA has previously been implicated in the R-dependent regulation of anthocyanin (Kunkel et a1., 1996), the levels of anthocyanin accumulation in the phyA mutant in this study additionally suggest that phyA has a distinct role in regulating the responsiveness to sucrose under Rc in Arabidopsis seedlings (Figure 3.4 C). Notably, a prior report for a phyD mutant in the Ler background reported lower levels of anthocyanin under white light (Aukerman et al., 118 1997), although levels were on average lower in the phyD mutant under Rc, a significant reduction in anthocyanin levels relative to the Col-0 WT parent in the absence of phyD was not apparent (Figure 3.4 C). No prior reports have been evident for phyC and phyE in the regulation of anthocyanin, but results obtained through quantification of anthocyanin levels for both mutants and the complementation of the phyE mutant suggest that phyC and phyE proteins have significant, but divergent roles in regulating anthocyanin levels under Rc. phyC is involved in the induction of anthocyanins, whereas phyE exhibits a distinctive, novel role in the suppression of anthocyanin accumulation under Rc (Figure 3.4 C and Figure 3.5). The analysis of transcript accumulation levels of anthocyanin marker genes, MYB 75/PAPI and DFR showed sucrose induction in the phyB mutant and confirmed sucrose-dependent up-regulation of DF R expression as previously published. A correlation between the level of transcript accumulation for MYB 75/PAP1 or DF R and the amount of accumulated anthocyanins was not apparent for the apophytochrome mutants and in-depth analysis of genetic as well as biochemical interactions among phytochrome family members would provide more conclusive evidence for the molecular mechanism of regulating sucrose-stimulated anthocyanin biosynthesis and accumulation by phytochromes under Re. 3.4 Future Perspectives Probing the biological functions of phytochromes has yielded phenomenal progress through the use of mutants harboring mutations in the genes encoding phytochrome apoproteins, as well as chromophore biosynthetic enzymes. Comparative 119 phenotypic and photobiological analyses of apophytochrome mutants has aided in studying discrete photoregulatory functions of individual phytochromes and their roles in light-mediated plant growth and developmental processes (Aukerman et a1., 1997; Franklin, Lamer, and Whitelam, 2005; Franklin and Quail, 2010; Franklin and Whitelam, 2004; Neff, Fankhauser, and Chory, 2000). Chromophore biosynthetic mutants have been used to probe the global effects upon loss of photosensory roles of all phytochromes. For example, hyI and hy2 mutants display multiple phytochrome-deficient phenotypes throughout the life cycle due to the lack of holophytochromes (Hudson, 2000; Terry, 1997). Given the discrete and overlapping functions among the phytochrome family members, the analysis of anthocyanin accumulation response in apophytochrome mutants could overlook the contributions of multiple members on the overall levels of anthocyanins. Additionally, phyA, phyB and phyD form homodimers and, phyB, phyD and phyE are known to exist in all possible heterodimeric combinations (Sharrock and Clack, 2004) whereas phyC and phyE do not homodimerize, but display obligate heterodimerization (Clack et a1., 2009). The lack of one partner could cause an imbalance in the amounts of dimerization partners and affect the overall stability of the existing partners (Sharrock and Clack, 2004). Furthermore, a mutation in one of the apophytochrome genes may influence the functions of one or more of the other isoforms and the accumulation of some phytochrome isoforms in light is coordinately regulated, at least in part by the levels of other members of the phytochrome family (Hirschfeld et a1., 1998). Thus, heterodimerization and functional inter-dependence of phytochrome family members further increases the complexity in the array of phytochrome functions and the analysis of combinatorial interactions of phytochrome isoforms is pivotal in defining the 120 steady-state levels, as well as the array of multiple and differentially competent phytochromes (Sharrock and Clack, 2004). Additive and synergistic interactions among the phytochrome family members can be determined through the comparative analysis of higher order mutants (Reed et a1., 1993) and such studies have enabled the elucidation of overlapping functions among the five isoforms (Franklin and Quail, 2010). For example, gradually lower levels of anthocyanins accumulate in phyB, phyD, and phprhyD mutants in Ws background, suggesting that phyB and phyD have an additive contribution and individual contributions of phyB and phyD isoforms are similar in anthocyanin accumulation response (Aukerman et a1., 1997). However, in the Ler ecotype, phyB is highly dominant over phyD in regulating levels of anthocyanin (Aukerman et a1., 1997). Currently, information on additive and/or synergistic interactions among the phytochrome family members in regulating anthocyanin biosynthesis and/accumulation remains limited. The analysis of anthocyanin accumulation in higher order mutants would reveal the additive roles and/or synergistic relationships of phytochrome isoforms in regulating sucrose-dependent anthocyanin accumulation under R. A limitation of comparative analysis of higher order mutants is the dissection of additive or synergistic roles conferred by homodimers and heterodimers. However, in the case of obligate heterodimerizing partners, i.e. phyC and phyE (Clack et a1., 2009), based on comparative analysis of anthocyanin accumulation response in the phyC, phyE and phprhyE mutants, additive or synergistic roles of phyC-phyE heterodimers can be elucidated. The relative expression levels for MYB75/PAP1 and DFR quantified by real-time RT-PCR would indicate whether the varying levels of accumulated anthocyanins correlate with the 121 expression of anthocyanin marker genes at the molecular level upon loss of multiple phytochrome family members. Through a comprehensive analysis of the anthocyanin accumulation response in high order mutants, the molecular mechanism of phytochrome function can be elucidated. A wealth of data indicates that phytochrome family members are expressed in a spatial- as well as temporal-specific manner and such localized pools of phytochrome can mediate discrete physiological functions (Bischoff et a1., 1997; De Greef and Caubergs, 1972a; De Greef and Caubergs, 1972b; Goosey, Palecanda, and Sharrock, 1997; Montgomery, 2008; Parcy, 2005; Sharrock and Clack, 2002; Zeevaart, 2006). Varying fractions of homodimeric and heterodimeric forms could arise due to differential spatial- and/or temporal-specific expression patterns of PH Y genes (Sharrock and Clack, 2004). Such physiological consequences could confound conclusions based on comparative phenotypic and photobiological analyses. Therefore, an enhancer trap-driven transactivation of BVR expression can be utilized to knock down the five types of phytochromes in a tissue- and/or organ-specific manner to gain insight into phytochrome- mediated inter-tissue signaling cascades underlying spatial-specific anthocyanin accumulation. Numerous regulatory proteins are involved in the regulation of light-dependent sucrose-stimulated anthocyanin accumulation and are spatially and developmentally regulated. Tissue-specific anthocyanin accumulation is commonly seen in vegetative tissues/organs (Borevitz et a1., 2000; Nesi et a1., 2000). Anthocyanins accumulate in the vacuolar space of cells in photosynthetic tissues, i.e. palisade and spongy mesophyll (Gould and Quinn, 1998; Lee and Collins, 2001) and epidermal cells (Kubo et a1., 1999). 122 Enhancer trap lines with GFP expression patterns that correlate with spatial-specific expression patterns of anthocyanin accumulation (i.e. J 1071- vascular/dermal expression throughout the seedling, J 1491- dermal expression in shoots and roots, J2093- root cap, epidermis, hypocotyl, apex and stomates, J2662- epidermis in hypocotyl, some expression is seen in cotyledons, (Haseloff, 1999), http://www.plantsci.cam.ac.uk/Haseloff/geneControl/catalogues) can be crossed with UAS-BVR lines to transactivate BVR and induce localized phytochrome inactivation at sites that are indicated by GFP expression. If cell autonomous signaling is perturbed, localized phytochrome inactivation may lead to a reduction of anthocyanin accumulation at the same site. Previous studies indicate that localized pools of phytochrome can regulate physiological responses at sites away from the site of photoperception (Bischoff et a1., 1997; De Greef and Caubergs, 1972a; De Greef and Caubergs, 1972b; Goosey, Palecanda, and Sharrock, 1997; Montgomery, 2008; Parcy, 2005; Zeevaart, 2006). If inter-tissue signaling is involved in phytochrome-mediated anthocyanin accumulation, a reduction in the levels of accumulated anthocyanins may be seen at distant sites away from the site of phytochrome inactivation indicated by GFP expression. Changes in the patterns of anthocyanin accumulation in the F3 progeny resulting from the cross between selected enhancer trap parents and UAS-BVR parent can be analyzed through bright field microscopy using cross-sections of fresh tissues. Thus, R/FR photoperception can be correlated with localized pools of phytochrome and anthocyanin accumulation response. Furthermore, molecular bases of cell autonomous and cell non-autonomous phytochrome-mediated signaling of spatial-specific anthocyanin accumulation can be elucidated. 123 "‘2 25 E 0.4 DD $1? ’3 8 0.2 ‘13 <3 0.0 . h :1 No-O 35S:: CAB:: Col-0 phyB WT pBVR3 pBVR2 WT Figure 3.] Anthocyanin content of wild-type and transgenic BVR seedlings. No-O wild-type (No—O WT), 35S::pBVR3, CAB3::pBVR2, Columbia-0 wild- type (Col-0 WT), and phyB (SALK_022035) were grown at 20 °C on Phytablend medium containing 1 % Suc for 4d under Rc illumination of 50 umol m2 s'1 or darkness. Black bars (Rc) and white bars (dark) represent the mean (+SD) of three independent measurements. 124 T-DNA phyA (SALK_014575) A t] g095 70 ATG Stop T-DNA .380 0.8030 0800 080808800 :0 00.00 000.8030 0:03 m.0>0. 8600:0080 ..-m N.8 .081 020 50088:... .00 :00:: 2% .x. . :0 0 0880800 88008 0:030:30 :0 0.. mm :0 850:0 008.0000 0.0-0-0 80:0 00:00:80 003 <20 00:00:00: 80.00.05 .0 ..0 am i .0>0. :0.mm0:0x0 :008 0:00.08 0:0m 80.8000 800 wmwnzé Numb 00 00080200 0 00 00:00 0.00.00 26 X . :0 .800 08.30 e\e 0 :0 30003103..me 0.0.: 0%. 638 103.00: 0.0:: 0.880 0.80000: .0808 03%. 0:080:2510550 00.: .95 0-30: 80-28 0-05528 5 00000003 :20 .0: 0:0 0080020: 855 0.3% 55an 2. 00:00 :0:.:08 8:050:80 00 m.0>0. :0.mm0:0x0 00 :0.:00_...::0:O Wm 0:00.“. 85 00.: 90.: 0.0.: 0.0.: as: 0-60 [5090/2150 % 0.0:: 00.: 0.0:: 0.0:: as: 0-60 <:'>. 3 Izoan/(thd)§z 81W % r000 .000 0.00. < 00m-_H_ 131 3.5 References Agati, G., and Tattini, M. (2010). Multiple functional roles of flavonoids in photoprotection. 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Plant Cell 5(7), 757-768. 139 Yadav, V., Mallappa, C., Gangappa, S. N., Bhatia, S., and Chattopadhyay, S. (2005). A basic helix-loop-helix transcription factor in Arabidopsis, MYC2, acts as a repressor of blue light-mediated photomorphogenic growth. Plant Cell 17(7), 1953-1966. Zeevaart, J. A. (2006). Florigen coming of age after 70 years. Plant Cell 18(8), 1783- 1789. Zhu, H. F ., Fitzsimmons, K., Khandelwal, A., and Kranz, R. G. (2009). CPC, a single- repeat R3 MYB, is a negative regulator of anthocyanin biosynthesis in Arabidopsis. Mo] Plant 2(4), 790-802. 140 Chapter 4 Regulation of Root Development by Phytochromes and Jasmonic Acid 141 4.1 Overview Light is the primary energy source for photosynthesis and the most important environmental signal regulating plant growth and development throughout the plant life cycle (Chory et a1., 1996; Franklin and Quail, 2010; Franklin and Whitelam, 2004; Neff, Fankhauser, and Chory, 2000; Schepens, Duck, and F ankhauser, 2004). The regulation of physiological responses depends upon complex intracellular, intercellular and inter-organ signaling cascades (Montgomery, 2008). Through inter-tissue and inter-organ signaling, opposing physiological responses in different plant tissues or organs can be regulated by the same light stimulus (Bou-Torrent, Roig-Villanova, and Martinez-Garcia, 2008). Although definitive information on molecular mechanisms of such inter-tissue and inter- organ communication is limited, tissue-specific gene expression analyses suggest that there are distinct subsets of light-mediated genes in discrete tissues in several plant species. In Arabidopsis, in cotyledons, hypocotyls, and roots, less than 1 % of light- regulated genes are common to all three types of tissues (J iao et a1., 2007). Despite the similarity in the mechanism of photoperception and initial signaling in cotyledons and roots in Arabidopsis (Cashmore et a1., 1999; Quail, 2002), distinct subsets of light- regulated genes have been identified from cotyledons vs. roots (J iao, Lau, and Deng, 2007; Ma et a1., 2005). In rice, roots appear to have more light-regulated genes than shoots (J iao, Lau, and Deng, 2007). Root-specific, light-regulated gene expression suggests that perception of light by root-localized photoreceptors has biological importance. Light penetration has been observed in the upper layers of soil up to several millimeters in natural environments (Mandoli et a1., 1990). Phytochromes, as well as 142 other photoreceptors, are localized in roots and render roots capable of sensing and responding to light (Kiss et a1., 2003; Okada and Shimura, 1992; Somers and Quail, 1995). By 'light piping' through the vascular tissue, the light signals perceived above ground can be extended to roots in deeper layers of soil (Mandoli and Briggs, 1982; Mandoli and Briggs, 1984). In the light spectrum, FR light is conducted most efficiently through internal light piping (Sun, Yoda, and Suzuki, 2005; Sun et a1., 2003). Thus, light that is perceived from aboveground portions of the plant can travel to the root to regulate root photomorphogenesis (Lauter, 1996). Root-localized phytochromes are able to perceive light directly and impact root growth and development in natural environments. Published data confirm that phytochromes are expressed in roots of Arabidopsis (Toth et a1., 2001) and are known have an important role in root development (Correll and Kiss, 2005; Salisbury et a1., 2007). Localization of phytochromes and light-dependent growth responses in roots indicate that light perception by roots is an important component of photomorphogenesis in plants. 4.1.] Role of Phytochromes in Root Growth and Development The ability of light to penetrate into the upper soil layers allows germinating seeds and roots in natural environments to perceive light (Mandoli et a1., 1990; Tester and Morris, 1987). Perception of light by germinating seeds is critical for detecting the location within the soil stratum. Phytochromes play a major role in ensuring that germination is induced at a specific time that a young seedling is able to reach the soil surface post-germination (Seo et a1., 2009). Salisbury et a1., (2007) reported that phyA, 143 phyD and phyE are highly expressed in primary and lateral root tips, whereas phyD is expressed throughout the elongation zone of the primary root. Detection of light by root-localized phytochromes and transmission of light through internal light piping can impact root growth and development in plants and discrete functions of phytochromes have been identified within the root system of Arabidopsis. Root-localized phytochromes regulate gravitropism in Arabidopsis in response to R light (Correll et a1., 2003; Correll and Kiss, 2005). In roots of maize seedlings, both phyA and phyB are involved in regulating light-induced gravitropism (F eldman and Briggs, 1987). In Arabidopsis, phyA and phyB play a key role in R light- induced positive phototropism in roots (Kiss et a1., 2003). Phytochrome action in roots is not limited to the tropic responses. Root-localized phytochromes in Arabidopsis, regulate root elongation under R light (Correll et a1., 2003; Correll and Kiss, 2005) and phyA and phyB have been shown to control R light-mediated elongation of the primary root (Correll and Kiss, 2005). Phytochromes also mediate the orientation of lateral roots (Kiss et a1. 2002) and lateral root production is regulated by stimulatory effects of phyA, phyB and phyE, and inhibitory effect of phyD (Salisbury et a1., 2007). The observation that initiation of lateral root grth is regulated by phytochromes in Arabidopsis plants grown on soil indicates that even in a more natural environment roots display phytochrome- dependent development (Salisbury et a1., 2007). Comparison of total amount of root hairs and root hair densities in light vs. dark- grown Arabidopsis seedlings by De Simone, Oka, and Inoue, (2000) indicated that light enhances root hair formation. Notably, under R light, phyA and phyB contribute to root hair development (De Simone, Oka, and Inoue, 2000; Reed et a1., 1993) and the 144 observation that a hy3 mutant (phytochrome B deficient or phyB mutant) exhibited longer root hairs than the wild type suggests that the perception of light by phyB could inhibit root hair elongation (Reed et a1., 1993). In fact, expression profiling of Arabidopsis roots using microarrays after 1 hr of exposure to R light indicates that expression of some genes involved in photomorphogenesis and root development is regulated by R light (Molas, Kiss, and Correll, 2006). A role for phytochromes in root hair initiation in lettuce seedlings has been reported and a site within the root itself may be a potential site of light perception for this response (De Simone, Oka, and Inoue, 2000). Not only R light, but also continuous exposure to FR light promotes root growth. In a phyA mutant, the lack of stimulation of root grth only under FR light suggests that phyA stimulates root growth in FR light (Kurata and Yamamoto, 1997). However, the observation that white, R or FR light was unable to stimulate root growth in a phyB mutant, suggests that phyB is essential for the roots to respond to light (Kurata and Yamamoto, 1997). According to Reed et a1., (1993), phyA is the only phytochrome mediating root hair formation under F Re. In addition to the recognized roles for phytochromes in Arabidopsis, emerging data indicate that phytochromes regulate root development in rice. Shimizu et a1., (2009) reported that seminal root grth inhibition in FR light was mediated exclusively by phyA, whereas in R light, both phyA and phyB were functional. Despite the detection of phyA, phyB and phyC in roots immunochemically, phyC appeared to have a minor or no role in growth inhibition of seminal roots in rice (Shimizu et a1., 2009). In seminal roots, the photoperceptive site for phytochrome-dependent inhibition appeared to be phytochromes within the roots themselves (Shimizu, Shinomura, and Yamamoto, 2010; 145 Shimizu et a1., 2009). Although reports on functions of phytochromes during regulation of root grth and development are available, biological importance of global loss of phytochromes on root development in Arabidopsis has just begun to be explored.— Interactions between hormone and light signaling have been extensively studied in the plant kingdom (Alabadi and Blazquez, 2009; Jaillais and Chory, 2010; Seo et a1., 2009). The plant hormone, jasmonic acid (JA), in addition to its recognized role in regulating defense responses, is a regulator of plant growth and development (Browse, 2005). J A inhibits root elongation (Staswick, Su, and Howell, 1992) and a number of reports indicate that phytochrome chromophore deficiency affects JA-mediated root inhibition (Muramoto et a1., 1999; Zhai et a1., 2007), but the current understanding of interaction of IA and phytochrome signaling in light-regulated root development is limited. 4.1.2 Biology of Jasmonic Acid JA and related compounds, collectively called jasmonates, are involved in numerous processes related to plant growth, development and survival. Among the physiological processes regulated by jasmonates are defense responses against biotic stresses (i.e. herbivore attack and pathogen infection), reproduction, secondary metabolism and senescence (Avanci et a1., 2010; Browse, 2005; Seo et a1., 2001; Wastemack, 2007; Wastemack and Hause, 2002). The jasmonates-JA, methyl jasmonate, J A conjugated to leucine and isoleucine and octadecanoid precursors-are produced from a-linolenic acid present on chloroplast membranes (Avanci et a1., 2010). Upon biotic or abiotic stress, phospholipases release a-linolenic acid from chloroplast membranes and a- linolenic acid serves as the precursor for JA biosynthesis via the octadecanoid pathway (Mueller et a1., 1993) involving three sub-cellular compartments: chloroplasts, 146 peroxisomes and cytoplasm (Browse, 2009; Mueller et a1., 1993; Seo et a1., 2001). The enzyme encoded by JAR], belonging to the acyl-adenylate enzyme class, catalyzes the conjugation of JA to the amino acid isoleucine (Ile) and generates jasmonoyl-L- isoleucine (JA-Ile; Guranowski et a1., 2007; Staswick and Tiryaki, 2004; Staswick, Tiryaki, and Rowe, 2002; Suza and Staswick, 2008). The observations that in vitro synthesized JAR] protein had the enzymatic activity to conjugate JA to lle and that J A- lle was able to inhibit root growth was confirmed by the jar] mutant and indicated that JA-Ile is the bioactive form of JA as a signaling molecule (Fonseca et a1., 2009; Staswick and Tiryaki, 2004; Thines et a1., 2007). Inhibition of root growth by JA has been widely used to identify mutants impaired in JA-biosynthesis and signaling based on insensitivity to growth inhibition upon exogenous application of JA or JA analogs. A number of key signaling intermediates in the JA signal transduction pathway have been identified in Arabidopsis (Avanci et a1., 2010; Browse, 2009; Wastemack, 2007; Wastemack and Hause, 2002). The C011 gene was identified in such a screen in the presence of a JA-Ile analog, coronatine (Feys et a1., 1994) and the cot] mutant was impaired in all known JA responses indicating that the F- box protein CO] is essential for JA signaling (Xie et a1., 1998). The CO] protein was later identified as a receptor for the bioactive form of JA (F onseca et a1., 2009; Thines et a1., 2007). Several groups identified that CO] 1 binds to S-PHASE KINASE-ASSOCIATED PROTEIN] and CULLIN to form the SCFCO” complex, an E3 ubiquitin ligase that degrades target proteins via the 26S proteosome in response to J A (Devoto et a1., 2002; Xu et a1., 2002). The CO] protein confers target specificity to the SCFCO“ complex . . COIl (Feys et a1., 1994; Xie et a1., 1998). The target proteins of the SCF complex are 147 jasmonate ZIM domain (JAZ) proteins that repress transcription of J A-responsive genes by blocking the activity of transcriptional activators (Chini et a1., 2007; Thines et a1., 2007). Binding of JA-Ile to the SCFCO“ complex induces degradation of J AZ repressors via the 26S proteosome and de-represses transcriptional activators allowing transcriptional activation of early response genes by jasmonates (C hini et a1., 2007; Thines et a1., 2007). 4.1.3 Phytochrome and Jasmonic Acid Signaling The contribution of jasmonates to regulation of cell expansion, cell division, growth orientation, and tissue and organ formation is known to affect plant morphogenesis (Avanci et al., 2010; Koda, 1997). Extensive overlap exists between the growth and developmental processes regulated by light and plant hormones (J aillais and Chory, 2010) and plant hormones function to integrate light responses (Bou-Torrent, Roig-Villanova, and Martinez-Garcia, 2008). Although limited. several breakthroughs in understanding the signaling network mediated by phytochromes and J A have been made in recent years. The initial report on a connection between JA and phytochrome signaling is the identification of a mutant displaying an F R-specific long hypocotyl phenotype in C ol-O WT background and the mutant was found to have a mutation in the FAR-RED- INSENSI T I VE2 1 9 (FIN219) gene, a suppressor of the constitutive photomorphogenesis1 (COPI) mutation (Hsieh et a1., 2000). Later, the finZl 9 mutation was found to be allelic to the jar] mutation (Staswick, Tiryaki, and Rowe, 2002) and JAR] encodes a J A-amino synthetase, catalyzing the conjugation of JA to Ile, required for optimal signaling in 148 jasmonate responses in Arabidopsis (Staswick and Tiryaki, 2004). Both jar] and finZ 1 9 mutants display a long hypocotyl phenotype under FRc compared to WT (Chen et a1., 2007) suggesting that mutants are impaired in phytochrome-mediated signaling in FR. Another example of a link between JA and phytochrome signaling is jasmonate insensitive] (iinI), an allele of MYCZ (Lorenzo et a1., 2004) and ZBF] (Yadav et a1., 2005). In FR light, the patterns of expression of certain gene are altered in the presence of mutations in ZBF] gene (Yadav et a1., 2005). Moreover, published data indicate that phytochrome chromophore biosynthesis and J A signaling are interconnected. H Y], H02, H03 and H04 genes encode multiple heme oxygenases (Emborg et a1., 2006) and HY] is expressed in roots of Arabidopsis (Davis et a1., 2001; Davis, Kurepa, and Vierstra, 1999; Emborg et a1., 2006). Under white light, a hy] mutant (hy][21.84N]) displays longer roots compared to the Ler WT parent (Muramoto et a1., 1999). By contrast, Zhai et a1., (2007) reported that hy] mutants in Col-0 WT background (hy]-100 and hy]-101) have shorter roots and upon JA treatment and HY] expression is down-regulated. This observation indicates that JA impacts phytochrome chromophore biosynthesis in Arabidopsis. Significance of interactions between JA and phytochrome signaling in the natural environment was recently discovered through comparative analyses of Arabidopsis plants subjected to insect herbivory after growth in high density or exposure to high FR light (Moreno et a1., 2009). Plants which are grown in shade or supplemented with FR light display characteristic shade-avoidance responses (Ballare, 2009; Ballare, Scope], and Sanchez, 1990). phyB detects changes in the R/F R ratio of light and is able to suppress shade avoidance responses under light with high R/FR ratio (Franklin et a1., 2003). Work by Moreno et a1., (2009) confirmed that, while phyB induces shade 149 avoidance responses in plants grown in high density or exposed to high FR light, induction of shade-avoidance responses also leads to a reduction in sensitivity to jasmonates accompanied by suppression of JA-mediated gene expression and defense responses upon insect herbivory. Selective insensitivity to jasmonates may have a role in diverting resources away from defense pathways and channeling towards stem elongation to reach light and maximize photosynthesis. Additionally, selective desensitization to jasmonates can also have implications in avoiding the inhibitory effects of jasmonates on cell growth, which could affect stem elongation, a characteristic response of the shade avoidance response (Yan et a1., 2007). Light perceived in leaves can regulate the development of organs at a distant site through manipulation of hormone signaling (Salisbury et a1., 2007). Global organ responses to R and FR light require the integration of cell autonomous responses with intercellular and inter-organ signaling to connect and coordinate them. Phenotypic and photobiological analyses of transgenic lines with targeted phytochrome inactivation will be an attractive tool to expand currently limited understanding of the cellular and molecular nature of signaling cascades involved in regulating phytochrome-dependent root development in Arabidopsis. 4.1.4 Targeted Chromophore Deficiency through Transactivation of Biliverdin Reductase Constitutive expression of BVR in Arabidopsis perturbs many light-dependent phytochrome-mediated responses (Lagarias et a1., 1997; Montgomery et a1., 1999). Studies with stable transgenic lines displaying mesophyll-specific and meristem-specific 150 phytochrome chromophore-deficiencies have revealed that localized pools of phytochromes can regulate distinct physiological responses and established the efficacy of targeted BVR expression as a novel molecular technique to investigate sites of light perception (Wamasooriya and Montgomery, 2009). A current limitation is the availability of cloned and characterized promoters that can direct the gene expression in a targeted manner. A two-component, enhancer-trap mis-expression system overcomes the limited availability of cloned and characterized promoters by using native genomic enhancers within a host genome (Wu et a1., 2003) and circumvents the necessity to maintain and genotype multiple stable transgenic lines (Wamasooriya and Montgomery, in press). Genetic crosses between the UAS-BVR parent and a range of GAL4-enhancer trap parents will result in progeny with diverse expression patterns of the BVR gene and results in distinct BVR expression patterns in a localized manner, which makes it an effective tool to determine sites of physiological processes, such as photoperception, that have been shown to have spatial-specific aspects. 4.1.5 Outlook Numerous studies establish the interactions between JA- and phytochrome- mediated signaling (Ballare, 2009; Ballare, Scope], and Sanchez, 1990; Chen et a1., 2007; Lorenzo et a1., 2004; Moreno et al., 2009; Muramoto et a1., 1999; Robson et a1; 2010; Yadav et a1., 2005; Zhai et a1., 2007). Even though published work has confirmed the presence and emphasized the importance of the link between the two signaling pathways, information on the possible tradeoffs between phytochromes- and JA-mediated signaling is limited. Moreover, collective functional roles of phytochromes and JA, and possible 151 sites of phytochrome photoperception during regulation of root elongation in Arabidopsis require further analysis. Muramoto et a1., (1999) observed longer roots in a chromophore biosynthetic mutant, hy] mutant (hy][21.84N]), compared to the Ler WT, whereas, Zhai et a1., (2007) reported that hy] mutants in Col-0 WT background (hy]-100 and hy]-10]) have shorter roots upon JA treatment. Thus, the objective of this study is to analyze if root-localized chromophore deficiencies alter light-dependent root morphogenesis and/or sensitivity of roots to JA-mediated root inhibition. A UAS-BVR transgenic line was crossed to the M0062 enhancer trap line displaying root-specific GFP expression (Haseloff, 1999). Comparative phenotypic analyses of transgenic lines with constitutive-, mesophyll- and meristem-specific chromophore deficiencies and M0062>>UAS-BVR progeny with root-specific chromophore deficiency indicate that lack of root-localized phytochrome/chromophore affects root development in a light-dependent manner in A. thaliana. 4.2 Materials and Methods 4.2.1 Transactivation of BVR Wild type Arabidopsis ecotype C24 plants were transformed with UAS-BVR construct by floral dip as described by (Clough and Bent, 1998). Kanamycin selection of transformants was performed as described in section 3.2.5. T1 transgenic seedlings were selected through a rapid selection protocol for identifying transformed Arabidopsis seedlings following floral dip transformation (Harrison et a1., 2006). T3 plants of UAS- BVR] were crossed with M0062 enhancer trap line (Haseloff, 1999) and F1 progeny were planted on kanamycin for selection. The genotyping of F1 seedlings was performed 152 by PCR with GoTaqGreen (Catalog No. M7123, Promega, WI). PCR amplification was carried out with the following gene-specific oligonucleotides at 10 uM in a 25 ul reaction: BVR— forward 5’—GCTGAGGGAC'ITGAAGGATCCAC-3’ and reverse 5’— CACTTCTTCTGGTGGCAAAGCTTC—3’, GAL4— forward, 5’— AGTGTCTGAAGAACAACTGGGAG—3 ’and reverse 5 ’— CGAGTTTGAGCAGATGTTTACC—3’. Thermal cycling conditions were (1) 1 cycle of denaturation at 95 °C for 2 min, (2) 40 cycles of denaturation at 95 0C for l min, annealing for 1 min (for BVR at 60 °C and for GAL4 at 58 °C), extension at 72 °C for l min and (3) final extension at 72 °C for 5 min with a hold at 4 0C. A lO-uL aliquot of the PCR product was visualized by electrophoresis for 2 h at 80V on a 0.8% agarose gel containing ethidium bromide (Catalog No. 15585-011, Invitrogen, CA) at 0.02 pg/mL (w/v). Ultraviolet images were obtained using a Gel Doc system (Bio-Rad Laboratories, Inc., CA) at subsaturation settings. F 1 seedlings that were positive with both primers sets were transferred to soil to obtain F2 and F3 seeds. The F3 and F4 seeds of UAS-BVR] x M0062 cross were used for subsequent analyses (M0062>>UAS-BVR). 4.2.2 Whole-mount lmmunohistochemistry Seeds of C24 WT and M0062>>UAS-BVR were sterilized as described in section 2.2.1 and were planted in 100- x 100- x lS-mm square petri dishes on media containing 1X Murashige and Skoog salts (Catalog No. MSP09, Caisson Laboratories, UT), 0.8 % (w/v) Phytablend (Catalog No.PTP01, Caisson Laboratories, UT), 0.05 % (w/v) 4- Morpholineethanesulfonic acid (Catalog No. M3672, Sigma, MO), 1 % (w/v) Sucrose (Catalog No. 4072-05, J.T. Baker, NJ), adjusted to pH 5.7 with KOH. Imbibing seeds 153 were cold-stratified at 4 0C for 3 days in darkness. Plates were kept vertically in a humidity-controlled chamber with Wc illumination of 100 umol m-2 s.1 for 3.5 days at 22 °C. 3.5-d-old seedlings were subjected to whole-mount in situ protein localization to visualize proteins in root tips, lateral roots, and embryos, as previously described with limited modifications (Sauer et a1., 2006). Seedlings were treated with the paraformaldehyde-based fixative solution (4 % paraformaldehyde, Catalog No. P6148, Sigma, MO in 1X PBS, supplemented with 0.1 % Triton X-100, Catalog No. A3352, Research Products International Corp., IL) for 30 min followed by washing with 1X PBS for 2x10 min and with sterile water for 2x5 min. Fixed seedlings were mounted on Poly- Prep slides (Catalog No. P0425, Sigma, MO) in a droplet of water and were air-dried for 2.5 h at room temperature. Cell walls were digested with 2 % Driselase (Catalog No. D9515, Sigma, MO) in 1X PBS for 30 min at 37 °C. Slides were washed with ]X PBS for 3x10 min. Tissues were permeabilized with 3 % IGEPAL CA—630 (Catalog No. 1302], Sigma, MO) containing 10 % dimethylsulfoxide (Catalog No. 9224-01, J. T. Baker, NJ) for l h at room temperature, followed by washing with 1X PBS for 4x10 min. Blocking with 3 % Bovine Serum Albumin Fraction V (Catalog No. 03 116 964 001, Roche Diagnostics, IN) was carried out for l h at room temperature. Fixed and permeated seedlings were incubated with rabbit anti-BVR antibody (Catalog No. 56257-100, QED Biosciences Inc., CA) at 1:2000 dilution in 1X PBS or with 1X PBS alone for control samples overnight at 4 °C. Excess primary antibody was removed by washing slides with 1X PBS for 3x10 min. Following incubation with the primary antibody and washing, seedlings were incubated with goat anti-rabbit IgG (H+L) conjugated to Hilyte PlusTM 555 (Catalog No. 61056-P1us555, AnaSpec, CA) at 0.005 mg/mL dilution in 1X PBS for 154 6 h at 37 °C. To remove excess secondary antibody, slides were washed with 1X PBS for 4x10 min. Drops of antifade mounting medium, Citifluor (Catalog No. 19470, Ted Pella Inc., CA) were placed on treated seedlings and covered with cover slips. Slides were stored overnight in darkness at 4 °C before imaging. Root tips of seedlings were imaged on an inverted Axiovert 200 Zeiss LSM 510 Meta confocal laser scanning microscope (Carl Zeiss Microlmaging, NY) using differential interference contrast (DIC) optics and fluorescence excitation/emission filters. A 20x0.75 Plan Apochromat objective lens was used for imaging. DIC imaging was performed using the 543-nm laser. Fluorescence from the secondary antibody was collected using a 543-nm laser for excitation and a 560 — 615 nm band pass filter for emission. Images were acquired using the LSM FCS Zeiss 510 Meta AIM imaging software (Carl Zeiss Microlmaging, NY). 4.2.3 Arabidopsis Seedling Extracts Seeds of No-O WT and 358::cBVRl were sterilized as described in section 2.2.1. Imbibing seeds were cold-stratified at 4°C for 3 days in darkness in 1.5 mL microcentrifuge tubes. In Phytatrays (Catalog No. P1552, Sigma, MO), imbibed seeds were planted on NITEX®nylon membrane (Catalog No. 03-100/32, Sefar Filtration Inc., NY) placed on a raft in contact with a liquid medium containing 1X Murashige and Skoog salts (Catalog No. MSP09, Caisson Laboratories, UT) with 1 % (w/v) Sucrose (Catalog No. 4072-05, J .T. Baker, NJ), adjusted to pH 5.7 with KOH. Phytatrays were kept in a humidity-controlled chamber with We illumination of 100 umol m-2 s-] for 21 days at 22 oC. Leaf and root tissues were quickly harvested (< 1 min) above and below the nylon membrane, weighed and transferred to separate 15 mL sterile tubes. After 155 immediately freezing in liquid nitrogen, tissues were crushed to a powder using a microgrinder. Crude Arabidopsis extracts were prepared according to a protocol adapted from (Lagarias et a1., 1997), leaf and root tissues were immediately homogenized in plant extraction buffer as described in section 3.2.6, however, for homogenization, plant extraction buffer was added at a ratio of 2 volumes/fresh weight (mg). The crude homogenates were clarified as described in sections 3.2.6. The aliquots of clarified supernatants were stored at -80 0C for immunoblot analysis in section 4.2.4. 4.2.4 Immunoblotting Total soluble proteins extracted from 21-day-old whole-plants were quantified as reported (Wamasooriya and Montgomery, 2009). For immunoblot analysis to confirm BVR accumulation in shoot and root extracts of 3SS::cBVR1 , ~ 25 pg of total protein was separated by SDS-PAGE and subsequent immunoblot analyses were performed as described (Montgomery et a1., 1999) using rabbit anti-BVR antibody (123000; Catalog No. 56257-100, QED Biosciences Inc., CA) and ImmunoPure® goat anti-rabbit IgG (H+L) conjugated to horseradish peroxidase (HRP; 1:5000, Catalog No. 31460, Pierce Biotechnology, Inc., IL) as the secondary antibody. Antibody signal (chemiluminescence) was detected using SuperSignal® West Dura Extended Duration substrate (Catalog No. 34075, Thermo Fisher Scientific Inc., IL) on Molecular Imager® TM VersaDoc MP 4000 System (Bio-Rad Laboratories Inc., CA). 156 4.2.5 Hypocotyl Inhibition Assays Seeds of No-O WT, 35S::cBVR1, C24 WT, and UAS-BVR1>>M0062 were sterilized and planted as described in section 2.2. l. Imbibing seeds were cold-stratified at 4 °C for 3 days in darkness. Plates were kept in a humidity-controlled chamber with Be or Rc illumination of 30 umol m'2 5'] and 50 umol m-2 s“, respectively, or in darkness for 7 days at 22 °C. Seedlings were scanned and plant images were used to quantify hypocotyl lengths using ImageJ software (NIH). The hypocotyl inhibition assay was repeated 3 times. Percentage dark length and standard deviations of percentage dark length were calculated according to the equation described in section 2.2.6. Two-tailed, unpaired Student’s t-test was performed to compare the percentage dark length of hypocotyls of transgenic lines relative to cognate WT seedlings, except for C24 WT, and UAS- BVR1>>M0062 grown in Rc and F Re, where two-tailed, unpaired Mann-Whitney test U- test was performed to compare the percentage dark length of hypocotyls of UAS- BVR1>>M0062 to that of C24 WT . 4.2.6 Root Inhibition Assays Seeds ofNo-O WT, 35S::pBVR3, 35S::cBVRl, CAB3::pBVR2, MER15::pBVR], Col-0 WT, jar] , myc02-05, C24 WT, M0062>>UAS-BVR, C20 WT, hy]-1 and hy2-1, were sterilized and planted on media prepared as described in section 4.2.2 with or without 20 uM Jasmonic acid (Catalog No. 392707, Sigma, MO). Imbibing seeds were cold-stratified at 4 0C for 3 days in darkness. Plates were kept vertically in a humidity- controlled chamber with We illumination of 100umol m'2 s'1 for 10 days at 22 °C. Plates were scanned and plant images were used to quantify root lengths using ImageJ software 157 (NIH). The root elongation assay was repeated 6 times. Two-tailed, unpaired Student’s t- test was performed to compare the means of root lengths. 4.2.7 Expression Levels of Jasmonic Acid-inducible Marker Genes To determine if spatial-specific deficiency of chromophore in vivo affects the expression of JA-inducible marker genes, real-time quantitative RT-PCR (qRT-PCR) was performed to quantify the levels of transcripts of a JA biosynthetic gene, 12-0PDA REDUCTASE3 (0PR3; At2g06050; Zhai et a1., 2007), and a JA-inducible marker gene, vegetative storage protein] (VSPI ; At5g24 780; Zhai et a1., 2007). Seeds of No-O WT, 35S::pBVR3, 3SS::cBVR1, CAB3::pBVR2, MER15::pBVR], Col-0 WT,jar1, myc02- 05, C24 WT, M0062>>UAS-BVR, C20 WT, hy]-I and hy2-1 were planted in 245- x 245- x 18-mm square petri dishes on media prepared as described in section 4.2.2 with or without 20 uM Jasmonic acid (Catalog No. 392707, Sigma, MO). Imbibing seeds were cold-stratified at 4 °C for 3 days in darkness. Plates were kept vertically in a humidity- controlled chamber with We illumination of 100 umol m'2 s-1 for 10 days at 22 °C. IO-d- old whole seedlings were quickly (< l min) harvested and immediately frozen in liquid nitrogen. Using RNeasy® Plant Minikit (Catalog No. 16419, Qiagen, MD) including on- column DNase treatment (Catalog No. 79254, Qiagen, MD), total RNA was isolated according to manufacturer’s instructions. Quantity of the RNA was analyzed by spectrometry (NanoDroplOOO, Therrno Scientific, DA). First-strand cDNA synthesis was performed using the Reverse Transcription System (Catalog No. A3500, Promega, W1) with random primers according to the manufacturer’s instructions using a 20 uL reaction volume. The incubation times of first-strand cDNA synthesis with total RNA of 0.2 pg 158 was (1) 10 min at room temperature, (2) l h (instead of 15 min) at 42 °C, (3) 5 min at 95 °C and (4) 5 min at 4 °C. The cDNA reaction mixture was diluted forty-fold with Nuclease-Free Water, and 4 uL of the diluted cDNA product was used as template in a 10 uL qPCR reaction using the Applied Biosystems FAST 7500 real-time PCR system in FAST mode with Fast SYBR® Green Master Mix (Applied Biosystems, CA) according to manufacturer’s instructions. For transcript analysis, annealing/extension temperature was 60 °C for both the 0PR3 and VSPI primer sets. Reactions were performed in triplicate and products were checked by melting curve analysis. The abundance of transcripts was analyzed using the ddCt method based on relative quantitation, normalizing to the reference transcript UBC21 (At5g25 760). All qRT-PCR experiments were repeated with three independent biological replicates. 4.3 Results and Discussion To regulate light-dependent root elongation and certain tropic responses within the Arabidopsis root system, roots themselves appear to be the site of photoperception (Correll et a1., 2003; Correll and Kiss, 2005). Elongated roots have been observed in a phytochrome chromophore biosynthetic mutant, hy] (hy][21.84N]; having a 13 bp deletion in the H Y I gene) compared to Ler WT by Muramoto et a1., (1999), in contrast to shorter roots in hy] mutants (hy]-100 and hy]-10]) in Col-0 WT background as observed by Zhai et a1., (2007). Thus, definitive evidence on the impact of phytochrome or phytochromobilin on JA-mediated root inhibition in Arabidopsis has been confounding. Furthermore, published reports indicate that JA and related compounds regulate distinct aspects of plant morphogenesis (Avanci et al., 2010; Koda, 1997) and 159 JA- and phytochrome-mediated signaling pathways are linked (Ballare, 2009; Ballare, Scope], and Sanchez, 1990; Chen et a1., 2007; Lorenzo et a1., 2004; Moreno et a1., 2009; Muramoto et a1., 1999; Yadav et a1., 2005; Zhai et a1., 2007). Since FR light is conducted most efficiently through internal light piping (Sun, Yoda, and Suzuki, 2005; Sun et a1., 2003) and phytochrome is the only known photoreceptor that is able to absorb R and FR light, at least in Arabidopsis, two novel molecular approaches were employed for targeting phytochrome inactivation to roots and to study the effects of phytochrome inactivation on root development. The sensitivity of roots to JA and JA-mediated root inhibition response in transgenic lines accumulating BVR in discrete tissues were analyzed in this chapter. Comparative phenotypic analyses of transgenic lines with constitutive, mesophyll-, meristem- and root-specific chromophore deficiencies indicate that lack of root-localized phytochrome or phytochrome chromophore affect root development in a light-dependent manner in A. thaliana. 4.3.1 Transactivation of B VR in M0062>>UAS-BVR A GAL4-based bipartite enhancer trap approach (Laplaze et a1., 2005) was used to generate an Arabidopsis line having phytochrome chromophore deficiency in roots. Through genetic crosses of the enhancer trap line, M0062 with root-specific GF P expression, and the UAS-BVR line, a transgenic BVR line, M0062>>UAS-BVR was isolated. Accumulation of BVR protein was confirmed by whole-mount immunohistochemistry on roots of the M0062>>UAS-BVR line (Figure 4.1). In the M0062>>UAS-BVR line, expression of BVR leads to root-specific holophytochrome 160 deficiencies and is used as a tool to study the impact of root-specific phytochrome inactivation on light-dependent root development in Arabidopsis. 4.3.2 Root-localized Phytochrome Inactivation does not Affect Hypocotyl Inhibition Response The observation that constitutive BVR-expressing seedlings had elongated hypocotyls in Re, FRc and Be light indicated that lack of phytochromes within the seedlings is responsible for the phenotype (Lagarias et a1., 1997). As 'light piping' allows the light signals perceived above ground to affect responses in roots (Mandoli and Briggs, 1982; Mandoli and Briggs, 1984), it is possible that phytochrome inactivation in roots might affect physiological responses in tissues other than roots. To test whether root- localized phytochrome inactivation affects hypocotyl inhibition and rule out residual activity ofthe BVR enzyme in shoots as would be evident by elongated hypocotyls in B, R and FR light, phytochrome-mediated hypocotyl inhibition response was analyzed in M0062>>UAS-BVR and 3SS::cBVRl under Bc, Re and FRc. The 3SS::CBVR1 line accumulates BVR protein in shoots and roots (Figure 4.2). The analysis ofhypocotyl inhibition response in the 3SS::cBVRl line showed a significant increase in hypocotyl length compared to the No-O WT parent under Be. Re and FRc (p>0.0001, Figure 4.3 and Figure 4.4) and as reported by Montgomery et al., (1999). The percentage dark length of M0062>>UAS-BVR compared to C24 WT was not significantly different under the Be light condition tested (p=().7l46, Figure 4.3 and Figure 4.4). However, in Re and FRc light, the frequency distribution analysis of hypocotyl lengths measured for C24 WT and M0062>>UAS-BVR seedlings showed that 161 lengths for C24 WT represented a normal distribution, whereas the lengths for M0062>>UAS-BVR seedlings represented a slightly negatively skewed distribution. Thus, two-tailed, unpaired Mann-Whitney test U-test was performed to compare the percentage dark length of hypocotyls of M0062>>UAS-BVR to that of C24 WT. In both Re and F Re, the percentage dark lengths of M0062>>UAS-BVR compared to C24 WT was not significantly different (Figure 4.4, p=1.0 in Re and FRc). This observation suggests that root-localized phytochromes have no or a minor role in regulating hypocotyl inhibition in Re, FRc and Be and residual BVR activity is absent in shoots of M0062>>UAS-BVR. 4.3.3 Phytochrome or Phytochromobilin Affects Root Elongation in Arabidopsis To more fully understand the regulation of light-dependent root development by phytochromes or phytochromobilin, root elongation responses were analyzed in transgenic Arabidopsis lines with constitutive, mesophyll-, meristem- and root-specific phytochrome inactivation through the expression of BVR. Root lengths of cognate WT plants and BVR-expressing transgenic lines were quantified in We illumination. BVR accumulation in the 3SS::pBVR3 and 35S::cBVRl roots was confirmed by immunoblot analyses (Wamasooriya and Montgomery, 2009 and Figure 4.2). The 35S::pBVR3 and 3SS::cBVR1 lines had significantly longer roots relative to No-O WT (p<0.0001, Figure 4.5). The roots of35S::pBVR3 and 3SS::cBVRl seedlings were ~ 45 % and ~ 90 % longer, respectively, than the roots of No-O WT seedlings. As previously published, the CAB3::pBVR2 seedlings accumulated BVR in mesophyll tissue, but not in roots and thus leads to mesophyll-specific phytochrome inactivation (Wamasooriya and Montgomery, 162 2009). The CAB3::pBVR2 seedlings showed ~ 21 % longer roots compared to No-O WT (p=0.0165, Figure 4.5) and the percentage elongation was not to the same degree as observed in the 3SS::pBVR3 and 35S::cBVRl. The root lengths of MER15::pBVR] with shoot-apex-specific BVR accumulation (Wamasooriya and Montgomery, 2009) were on average ~ 8 % longer, but not significantly different from No-O WT (p=0.3424, Figure 4.5). Zhai et a1., (2007) observed shorter roots in the hy]-10] mutant compared to Co]- 0 WT. In comparative analysis with BVR-expressing transgenic lines, root lengths were quantified in two chromophore biosynthetic mutants, hy]-I and hy2-1. Relative to the C20 WT, roots of hy]-1 were ~ 15 % shorter and roots of hy2-1 were ~ 23 % longer (Figure 4.5). However, compared to the C20 WT, the differences observed in root length for hy]-I and hy2-1 were not statistically significant (p=0.1524 and p=0.0859, respectively). M0062>>UAS-BVR with root-localized phytochrome chromophore deficiencies showed roots ~ 12 % longer in length relative to C24 WT (Figure 4.5), a response that was similar to the root elongation observed for 35S::pBVR3, 35S::cBVR1 and hy2-1. A characteristic phytochrome-deficient phenotype of 35S::pBVR3 and 35 S::cBVR1 transgenic lines with constitutive phytochrome inactivation is elongation of hypocotyls under R, FR and Be (Lagarias et a1., 1997; Montgomery et a1., 1999) and such a phenotype was not observed in the M0062>>UAS-BVR line as described in section 4.3.2 (Figure 4.3 and 4.4). This observation provides additional evidence that M0062>>UAS-BVR lacks root-localized phytochromes and the elongated roots likely represent a specific perturbation of a phytochrome-regulated root inhibition response. 163 On average, statistically significant increases in root lengths under Wc illumination were observed for transgenic lines with BVR accumulation in roots (i.e. 3SS::pBVR3, 35S::cBVR1 and M0062>>UAS-BVR, thus root-localized phytochrome inactivation) with respect to lines lacking BVR accumulation in roots (i.e. No-O WT, CAB3::pBVR2 and MER15::pBVR], Figure 4.5). A similar phenotype was also apparent in the chromophore-deficient mutant, hy2-1 relative to C20 WT (Figure 4.5). As previously reported, subcellular localization of BVR affects distinct subsets of light- mediated and light-independent processes in Arabidopsis (Montgomery et a1., 1999). For example, 35S::pBVR3 with plastid-localized BVR expression is intolerant to higher light fluences and displays a fluence-rate-dependent reduction in chlorophyll levels accompanied by an increase in the chlorophyll a/b ratio (Lagarias et a1., 1997). This phenotype could be related to changes in plastid metabolism (Franklin et a1., 2003) upon BVR activity in plastids and/or due to lack of BV/phytochromobilin that would otherwise play an important regulatory role within the plastid compartment (Montgomery et a1., 1999). A response to higher light fluences, as such, was not obvious in 35S::cBVR] with cytosolic BVR expression (Montgomery et a1., 1999). Since 3SS::cBVR1 and M0062>>UAS-BVR exhibit cytosolic BVR expression, increased root lengths observed in these lines compared to their cognate WT plants are expected to be associated with lack of phytochromobilin or phytochromes and likely reflect disruption of the phytochrome-mediated root inhibition response as reported for R light-dependent inhibition of root elongation (Correll and Kiss, 2005). The presence of longer roots in the CAB3::pBVR2 line relative to No-O WT may be due to the disruption of phytochrome-mediated signaling between mesophyll- and 164 root-localized phytochrome pools and implies that mesophyll-localized phytochromes may be involved in long-distance communication to regulate root elongation. This observation corroborates an already demonstrated phenomenon by Salisbury et a1., (2007) that light-perception by shoot-localized phytochromes can impact root development in Arabidopsis. The impaired phytochrome-mediated root inhibition in transgenic lines with mesophyll- and root-specific phytochrome inactivation with respect to lines with constitutive phytochrome inactivation suggests that both mesophyll- and root-localized phytochromes distinctly contribute to light-dependent regulation of root development in Arabidopsis. 4.3.4 Root-localized Phytochrome or Phytochromobilin Reduces Jasmonic Acid- mediated Root Inhibition Zhai et a1., (2007) confirmed that phytochromobilin deficiency in the hy]-101 mutant enhances JA-mediated root inhibition. Inhibition of root elongation is promoted by JA and derivatives of JA (Staswick, Su, and Howell, 1992). By measuring root lengths of BVR-expressing transgenic lines, root inhibition responses were quantified in the presence of exogenous methyl JA (MeJA) to determine whether the lack of phytochromobilin impacts JA-mediated root inhibition. Comparative analysis of root lengths indicated that lines with phytochrome inactivation in roots had reduced sensitivity to MeJA. Root lengths of3SS::pBVR3, 35S::cBVR1, hy]-1 and hy2-1 were ~ 101 %, ~ 82 %, ~ 73 % and ~ 97 % longer than the root lengths of their cognate WT parents, respectively, and were significantly longer (p 50.0001 for all) in the presence of 20 uM of MeJA (Figure 4.6). Root elongation to a similar degree in the presence of MeJA was 165 also observed in the JA-insensitive mutants, jar] and myc02-5, with ~ 100 % and ~ 6] % increase respectively, compared to root lengths measured for Col-0 WT (Figure 4.6). In the absence of MeJA, the root lengths of 35S::pBVR3 and 3SS::cBVRl were longer than No-0 WT (Figure 4.5), thus, for more accurate quantitative comparison of sensitivity of roots to MeJA, relative MeJA-sensitivity was calculated. For a particular line, the relative MeJA-sensitivity was the ratio of the root length in the absence of exogenous MeJ A to the root length in the presence of 20 uM of MeJA (Table 2). In the absence of MeJA, the 3SS::pBVR3 line with plastid-targeted BVR expression in roots displayed a 3.45-fold difference in root length relative to 4.75-fold difference in No-O WT (Table 2). The difference between No-0 WT and 3SS::pBVR3 reflects a ~ 27 % reduction in sensitivity to MeJA application (Table 2) and of all the lines tested, J A-insensitive mutants, jar] and myc02-5 displayed the most reduction in JA-sensitivity (~ 66 % and ~ 49 %, respectively) compared to Col-0 WT (Table 2). A 4.95-fold increase was apparent for 35S::cBVRl with cytosolic BVR accumulation in the absence of MeJA relative to 20 uM of MeJ A (Table 2) and suggests that 3SS::cBVR1 is as sensitive as No-O WT to JA-mediated inhibition of root elongation. In the 35S::cBVRl line, the roots were significantly longer than the roots of No-O WT on 0 uM as well as 20 11M MeJA. The relative JA-sensitivity in M0062>>UAS-BVR also indicated a reduced response to 20 11M of MeJA. In the absence of MeJA, the fold difference for M0062>>UAS-BVR was 4.46 compared to 4.8 for C24 WT (Table 2). The roots of M0062>>UAS-BVR were significantly longer than the roots of C24 WT (p=0.0266) and showed ~ 7 % reduction in sensitivity to MeJA application (Table 2). As noted for 3SS::pBVR3 and M0062>>UAS-BVR, chromophore biosynthetic mutants, hy]-1 and hy2-1 also exhibited hyposensitivity to MeJA treatment 166 (Figure 4.6). Reduction in relative JA-sensitivity for hy]-1 and hy2-1 was ~ 51 % and ~ 37 % respectively (Table 2) and both lines displayed longer roots compared to C20 WT (Figure 4.6). However, contrasting responses were observed in hy]-1 and hy2-1 in the absence of MeJA; roots were shorter in hy]-1, but longer in hy2-1 relative to C20 WT (Figure 4.5). The presence of longer roots in hy]-I on MeJA treatment was distinctive from the shorter roots reported for hy]-10] allele in Col-0 WT background by Zhai et a1., (2007). However, the relative JA-sensitivity in hy]-101 (Zhai et a1., 2007) and hy]-1 appeared to be similar despite the differences in sensitivities of different Arabidopsis ecotypes to treatment with JA (Matthes, Pickett, and Napier, 2008), duration of growth and MeJA concentration. Transgenic lines with phytochromobilin deficiencies in roots, i.e. 35S::pBVR3 and M0062>>UAS-BVR, relative to other BVR-expressing lines displayed the most reduction in sensitivity to MeJA (~ 27 % and ~ 7 %, respectively) and suggests that lack of phytochrome or phytochromobilin in roots impacts light-dependent, JA-mediated root inhibition. The response of hy]-1 on 0 uM and 20 uM of MeJA was analogous to jar] and myc02-5 having shorter roots than their cognate WT lines in the absence of MeJA and longer roots upon exogenous application of 20 uM of MeJA. The response of the hy2-I mutant resembled closely the responses observed in the 35S::pBVR3 line and M0062>>UAS-BVR, having longer roots relative to cognate WT lines regardless of the absence or the presence of MeJA treatment, but all three lines displayed hyposensitivity to MeJA. The 35S::cBVR1 line was discrete in its relative response to application of MeJA. Even though the 35S::cBVR1 line had longer roots compared to No-O WT regardless of the absence or presence of MeJA, the line was not impaired in relative 167 sensitivity to MeJA compared to No-O WT. Despite their cytosolic BVR expression, the contrasting response to MeJA in 35S::cBVRl and M0062>>UAS-BVR, i.e. 35S::cBVR1 being as sensitive as No-0 WT and M0062>>UAS-BVR being hyposensitive relative to C24 WT, may be due to a perturbation of communication between the shoot and root and/or between phytochrome- and JA-mediated signaling. The differences in the amounts of BVR being accumulated in roots of 3SS::cBVR1 and M0062>>UAS-BVR could also account for the disparate responses to MeJA application. A possible reason for the 3SS::cBVRl line to have similar sensitivity to MeJA as No-O WT and hyposensitivity to MeJA in the 35S::pBVR3 line could be due to indirect effects of changes to plastid metabolism on JA biosynthesis. pB VR expression could affect the stability of phytochromobilin synthase (Montgomery et a1., 1999) and/or alter plastid heme levels (Lagarias et a1., 1997; Montgomery et a1., 1999). As targeting of BVR expression to plastids could alter plastid metabolism (Franklin et a1., 2003; Montgomery et a1., 1999) and JA biosynthesis involves chloroplasts/plastids (Mueller et a1., 1993), changes to plastid metabolism could impact early steps of JA biosynthesis occurring in the chloroplasts/plastids (Wastemack, 2007). Based on the comparative analysis of relative sensitivities to MeJA in the 35S::pBVR3 line with plastid-localized BVR accumulation and the M0062>>UAS-BVR line with cytosolic BVR accumulation, it is evident that the observed hyposensitivity to MeJA is likely related to a lack of phytochromobilin and/or phytochrome, but not due to indirect consequences of accumulated BVR on plastid metabolism. 168 4.3.5 Root-localized Phytochrome Deficiencies Impact Expression of Jasmonic Acid- inducible Marker Genes To determine whether the presence of longer roots in lines with root-localized phytochrome inactivation is correlated with lower endogenous levels of JA and/or whether the hyposensitivity to JA-mediated root inhibition is reflected at gene expression level, the expression of 0PR3 (At2g06050) and VSP] (At5g24 780) was analyzed by qRT- PCR. In the absence of MeJA treatment, the expression of 0PR3, a JA-biosynthetic gene, was low in all the lines tested (Figure 4.7). Expression levels of 0PR3 in 35S::pBVR3 and 3SS::cBVR1 with constitutive phytochrome inactivation was ~ 74 % and ~ 66 %, respectively, relative to No-O WT (Figure 4.7). The expression level of 0PR3 in M0062>>UAS-BVR with root-localized phytochrome inactivation was ~ 90 % compared to its WT, C24. This is only an ~ 10 % reduction in expression as opposed to ~ 26 % and ~ 34 % reduction observed in the 3SS::pBVR3 and 3SS::cBVR1 lines, respectively. Thus, distinct differences in the levels of 0PR3 expression are obvious for lines with constitutive vs. root-specific phytochrome chromophore deficiency. ~ 22 % and ~ 16 % reduction in 0PR3 expression levels relative to No-0 WT was apparent in the CAB3::pBVR2 and MER15::pBVR] lines, respectively, lacking phytochrome inactivation in roots (Figure 4.7). In contrast to what was observed for the 3SS::pBVR3 and 35S::cBVRl lines with constitutive phytochrome inactivation in the absence of MeJA, ~ 10 % and ~ 25 % increase in 0PR3 expression level relative to C20 WT was obvious in hy]-1 and hy2-1, respectively, (Figure 4.7). Previously, an increased level of 0PR3 expression in the absence of MeJA treatment was reported for a different hy mutant, hy]-101, relative to Col-0 WT by Northern blot analysis (Zhai et a1., 2007). 169 Relative to Col-0 WT, the jar] mutant had ~ 13 % increase, whereas myc02-5 had ~ 15 % decrease in 0PR3 expression level (Figure 4.7). Reductions in 0PR3 expression levels have been reported for JA-insensitive mutants as determined previously by Northern blot analysis (Chung et a1., 2008; Koo et a1., 2009). Upon exogenous application of 20 uM of MeJA, the expression of 0PR3 was induced relative to expression levels on 0 uM of MeJA in all the lines, but the degree of induction was clearly variable in comparison to their cognate WT plants. The 35S::pBVR3 and 35S::cBVR1 lines with phytochrome inactivation in roots showed ~ 64 % and ~ 54 % levels of 0PR3 expression, respectively, relative to No-O WT (Figure 4.7). However, the level of 0PR3 expression for M0062>>UAS-BVR with root-localized phytochrome inactivation was more or less identical to the expression level observed for C24 WT (Figure 4.7). The 0PR3 expression was ~ 62 % in the CAB3::pBVR2 line with mesophyll-specific phytochrome inactivation, whereas the level for the MER15::pBVR] line with meristem-specific phytochrome inactivation was ~ 74 % relative to No-O WT. In the hy]-1 mutant, the 0PR3 expression was ~ 88 % and by contrast, in the hy2-I mutant, it was slightly higher (~ 4 %) than that of No-O WT (Figure 4.7). An expression level of ~ 82 % and ~ 75 % was observed in the jar] and the myc02-5 mutants, respectively, relative to Col-0 WT (Figure 4.7). Most lines with hyposensitivity to JA- regulated root inhibition displayed lower levels of 0PR3 expression relative to their cognate WT plants, with M0062>>UAS—BVR and hy2-1 being exceptions. Even though variations in the level of 0PR3 expression were apparent, all phytochrome-deficient lines were responsive to exogenous application of MeJA. 170 The expression level of JA-inducible marker gene VSP] was undetectable or barely detectable in all of the lines in the absence of exogenous MeJA application (Figure 4.8). However, all the lines tested showed robust induction of VSP] expression in the presence of MeJA treatment suggesting that 0PR3 and VSPI have distinct expression patterns. Lower levels of VSP] expression were evident in the 358::pBVR3, 3SS::cBVRl, CAB3::pBVR2, MER15::pBVR], jar] and myc02-5 lines with respect to the expression levels in cognate WT plants. The 3SS::pBVR3 and 3SS::cBVR1 lines showed ~ 58 % and ~ 53 % VSP] expression, respectively, comparative to No-O WT, whereas the CAB3::pBVR2 and MER15::pBVR] lines showed ~ 67 % and ~ 69 % VSP] expression, respectively. These results suggest distinct differences in VSP] expression level for lines with constitutive and tissue-specific phytochrome deficiencies. ~ 50 % increase in VSP] expression relative to C24 WT was observed for M0062>>UAS-BVR with root-localized chromophore deficiency and suggests that M0062>>UAS-BVR is responsive to exogenous application of MeJA. Chromophore deficient hy]-I and hy2-1 mutants showed a similar increase, ~ 59 % and ~ 50 %, in VSP] expression, respectively, with respect to C20 WT. This observation correlates with a substantial increase in VSP] expression in the hy]-10] mutant relative to Col-0 WT on MeJA application as previously reported by (Zhai et a1., 2007). As determined by Northern blot analysis of VSP] expression in JA-insensitive mutants in prior studies (Chung et a1., 2008; Koo et a1., 2009), ~ 48 % and ~ 36 % reductions were evident in jar] and myc02-5, respectively, compared to their WT, Col-0 (Figure 4.8). The analysis of expression of JA-inducible marker genes, 0PR3 and VSP] by qRT-PCR in BVR-expressing transgenic lines indicated that 0PR3 and VSPI expression 17] is generally reduced for lines with constitutive and tissue-specific phytochrome chromophore deficiencies relative to cognate WT plants. 0PR3 expression was detectable on 0 uM, whereas VSP] expression was barely detectable. However, all of the lines tested were responsive to MeJA and VSP] expression showed robust induction on 20 uM. The 3SS::pBVR3 and 3SS::cBVRl lines with constitutive phytochrome chromophore deficiency had reduced levels of 0PR3 and VSP] expression relative to No- 0 WT. Even though such a reduction was apparent in the CAB3::pBVR2 line with mesophyll-specific chromophore deficiency and in the MER15::pBVR] line with meristem-specific chromophore deficiency, the reduction was not as extensive as in the 3SS::pBVR3 and 35 SzchVRl lines. A possible explanation for the reduction of 0PR3 and VSP] expression in the CAB3::pBVR2 and MER15::pBVR] lines is disruption of inter-tissue phytochrome- and JA-mediated signaling upon mesophyll- and meristem- specific phytochrome deficiencies. The M0062>>UAS-BVR line showed similar 0PR3 expression to that of C24 WT on MeJA, but expression was increased by ~ 50 % for VSP] relative to C24 WT on MeJA application. In general, the expression pattern of 0PR3 and VSP] observed for M0062>>UAS-BVR is unique from two other lines (i.e. 3SS::pBVR3 and 3SS::cBVR1) with phytochrome inactivation in roots and may be related to a higher level of BVR accumulation in roots or a distinct phenotype at the molecular level due to root-localized phytochromobilin or phytochrome deficiencies. JA- insensitive mutants, jar] and myc02-5, both showed reductions in 0PR3 and VSPI expression relative to Col-O WT as previously reported (Chung et a1., 2008; Koo et a1., 2009). The increase in 0PR3 expression for hy2-l and increase in VSPI expression for hy]-1 and hy2-1 correlated with the apparent increase in the expression of the two genes 172 in the hy]-10] allele in Col-O WT background as previously published (Zhai et a1., 2007). However, the reduction in 0PR3 expression evident for the hy]-I mutant may be a result of ecotypic differences between C20 WT and Col-0 WT. 4.3.6 Summary Increased root lengths of 35S::cBVR1 and M0062>>UAS-BVR lines with cytosolic BVR expression compared to their cognate WT plants are expected to be associated with a lack of phytochromobilin or phytochromes and likely reflect disruption of the phytochrome-mediated root inhibition response. The presence of longer roots in the CAB3::pBVR2 line relative to No-O WT may be due to the disruption of phytochrome-mediated signaling between mesophyll- and root-localized phytochrome pools and implies that mesophyll-localized phytochromes may be involved in long- distance communication to regulate root elongation. This observation corroborates an already demonstrated phenomenon by Salisbury et a1., (2007) that light-perception by shoot-localized phytochromes can impact root development in Arabidopsis. The impaired phytochrome-mediated root inhibition in transgenic lines with mesophyll- and root- specific phytochrome inactivation with respect to lines with constitutive phytochrome inactivation suggests that both mesophyll- and root-localized active phytochromes contribute to light-dependent regulation of root development in Arabidopsis. Expression analysis of 0PR3 and VSP] indicates that the two genes have distinct expression patterns in the lines with chromophore deficiencies. 0PR3 is expressed to detectable levels in the absence of MeJA and is induced in all of the lines on exogenous MeJA application. By contrast, the level of VSP] expression was barely detectable in the 173 absence of MeJA and shows robust induction in all of the lines on exogenous MeJ A treatment. The PHYTOCHROME AND F LOWERING TIME] (PF T1) fiinctions as a subunit of the Mediator complex and negatively regulates phytochrome signaling (Kidd et a1., 2009). PF T1 is also required for JA-dependent expression of defense genes in Arabidopsis (Kidd et a1., 2009). As JA and phytochrome signaling have antagonistic effects (Zhai et a1., 2007), PF Tl also exhibits divergent fianctions (Kidd et a1., 2009) indicating a definitive molecular link between phytochrome and JA-inducible marker genes. Arabidopsis lines with root-localized BVR accumulation have defects in light- dependent root elongation. Arabidopsis lines with root-localized BVR expression, i.e. 35S::pBVR3 and M0062>>UAS-BVR, exhibit hyposensitivity to exogenous application of the JA-derivative, MeJA. These results provide evidence that the root-specific phytochrome chromophore or root-localized phytochromes are vital for photoregulation of root elongation and impact JA sensitivity. 4.4 Future Perspectives The experimental findings of this chapter and previously published data confirm that JA- and phytochrome mediated signaling is interconnected (Lorenzo et a1., 2004; Moreno et a1., 2009; Robson et a1., 2010; Staswick, Tiryaki, and Rowe, 2002) and the lack of phytochromobilin or phytochromes in roots negatively impacts root inhibition by JA (Zhai et a1., 2007). The elongation of roots observed for BVR-expressing transgenic lines with phytochrome deficiencies in roots (notably, where BVR expression was targeted to the plastids, i.e. 3SS::pBVR3) could be due to already lower levels of 174 endogenous JA in them. The accumulation of BVR in plastids is known to affect plastid metabolism (Franklin et a1., 2003) and the involvement of plastids in JA biosynthesis (Wastemack, 2007) could potentially have indirect effects on overall JA levels. In lines with root-localized phytochrome deficiencies, even though the expression level of a J A- biosynthetic gene, 0PR3, was similar to the level noted for their cognate WT, the lack of phytochromobilin or phytochrome could impact downstream or upstream enzymatic activities relative to OPR3 activity. To determine whether the accumulation of BVR or its end products, i.e. bilirubin and phytochromorubin, has an impact on JA biosynthetic genes leading to changes in JA production, the endogenous JA and JA-Ile levels can be quantified by liquid chromatography-mass spectrometry (LC-MS) before and after mechanical wounding as previously published (Chung et a1., 2008; Koo et a1., 2009; Li et a1., 2005). Furthermore, to develop a mechanistic interpretation of data from LC-MS, the expression levels of several JA-biosynthetic genes, such as LOX2 (At3g45140), AOS (At5g42650) and AOC1 (At3g25 760), as well as JA-inducible marker genes, T hionin (T hi2. 1 , At] g72260), JAZ5 (At1g17380) and JAZ 7 (At2g34600) can be analyzed by qRT- PCR. qRT-PCR analysis of JA-biosynthetic, as well as JA-inducible marker genes, together with LC-MS data will reveal if the endogenous JA levels are markedly different among the lines with chromophore deficiencies relative to WT. Ecological significance of cross talk between phytochrome and JA signaling was revealed by Moreno et a1., (2009). In plants grown in high density or exposed to high FR light, phyB mediates shade avoidance responses accompanied by a reduction in sensitivity to jasmonates. Hyposensitivity to JA is characterized by suppression of JA- mediated gene expression and defense responses upon insect herbivory (Moreno et al., 175 2009). As BVR accumulation leads to reduction of phytochromobilin and thus a reduction of the five types of phytochromes (Lagarias et a1., 1997; Montgomery et a1., 1999), to determine whether the lines with chromophore deficiencies are compromised in resistance to insect herbivory, feeding assays using the generalist Spodoptera exigua followed by the analysis of J A-marker gene expression can be performed. Feeding assays with S. exigua would represent conditions encountered by plants in a more natural environment and information fi'om the feeding assays could expand the understanding of ecological importance of an antagonistic relationship between phytochrome and J A signaling. Perception of light by root-localized phytochromes (Correll et a1., 2003; Correll and Kiss, 2005) and transmission of perceived light information through internal light piping (Mandoli and Briggs, 1982; Mandoli and Briggs, 1984) can impact growth, development and tropic responses in the Arabidopsis root system. Although phytochromes are known to be expressed in roots and root tips of Arabidopsis (Salisbury et a1., 2007; Toth et a1., 2001), information on perception of light by phytochromes localized in different root tissues that regulate root photomorphogenesis is lacking. Enhancer trap lines; J 1 103, KS074 and Q0171, with GFP expression in the root cap and , emerging lateral roots, root tip and root cap, respectively (Haseloff, 1999), http://www.plantsci.cam.ac.uk/Haseloff/geneControl/catalogues) can be crossed with UAS-BVR lines to transactivate BVR and induce localized phytochrome inactivation at sites that are indicated by GFP expression. Comparative phenotypic analysis of F3 progeny would indicate whether phytochromes localized in specific root tissues are involved in light perception. 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No-O wild-type (No-0 WT), 3SS::pBVR3, 3SS::cBVRl,CAB3::pBVR2, MER15::pBVRl, C24 wild-type (C24 WT), M0062>>UAS-BVR, Col-0 wild-type (Col-0 WT), jar], myc02-5, C20 wild-type (C20 WT), hy]-1 and hy2-I lines were grown on Phytablend medium containing 0.8 % Suc for 10 d at 22 °C under Wc light of 100 umol m.2 s". Data points represent means (:9: SD) of 6 or more roots measured from 6 independent experiments. F old-difference values for root lengths determined for seedlings with respect to cognate wild-types are indicated. 182 :3 8 20000 mm... wwd N06 00.0 N... 00.. .N. 00.. 20.. ”0000:0000 00? 29: :DW AWQV %0 none Cc 0v “V O\ ...n» ...nc ..no ....00 00¢ ........00-00.00.10.04004003 ., . , . _ , y . . .0 .J ... h 0 1 1 m _ W L ,, 0 IO.ON N 0 a m m . v m . m- : l—r 130.3. m. Fl . # m... . . -000 m 0.00 183 Figure 4.6 Mean root lengths of wild-type, BVR-expressing and mutant seedlings. No-O wild-type (No-O WT), 3SS::pBVR3, 3SS::cBVRl,CAB3::pBVR2, MER15::pBVR], C24 wild-type (C24 WT), M0062>>UAS-BVR, Col-0 wild- type (Col-O WT), jar] , my602-5, C20 wild-type (C20 WT), hy]-1 and hy2-1 lines were grown on Phytablend medium containing 0.8 % Suc with 20 pM jasmonic acid for 10 d at 22 °C under Wc light of 100 umol m'2 s". Data points represent means of 6 or more roots measured from 6 independent experiments. Fold- difference values for root lengths determined for seedlings with respect to cognate wild-types are indicated. 184 3.3 3 0.3305 Sc m: G._ SN Sir _N._ N: a: .3 sausage $00 av 28 re ow O p. .c a? 9 fig». ago», a» or 9% A? a ow as. aw a A a e n 4 av .v7 «V nvv avv xi: nvv «”7 r o o N tutu tn 1.1131191 1001 meow 0d? 185 Table 2 Fold-difference in root lengths of wild-type, BVR-expressing and mutant seedlings. Fold-difference values for root lengths determined for seedlings on 0 uM JA relative to 20 pM JA are indicated. Fold difference (-JA/+JA) Plant line (relative sensitivity to MeJA) ‘ No-O we 4.75 " ' 35S::pBVR3 i N f 3.45 3SS::cBVRl . it 4.95 CAlB3::pBVR2 4.74 ' Makisépiévm 7 f 4.23 I c24Wr 4.30 .4 M0062>>UAé-BVR t i 4.46 ‘4 Col-O WT * f 4.85 jar] 1.62 mchZ-5 ‘ 2.48 C20 wr 5.76 hAyI-I. f 2.82 i hy2-1 r 3.60 186 Figure 4.7 Relative expression levels of 0PR3 in wild-type, BVR-expressing and mutant seedlings. No-O wild-type (No-0 WT), 3SS::pBVR3, 3SS::CBVR1,CAB3::pBVR2, MER15::pBVRl, C24 wild-type (C24 WT), M0062>>UAS-BVR, Col-O wild- type (Col-0 WT), jar] , my602-5, C20 wild-type (C20 WT), hy]-1 and hy2-1 lines were grown on Phytablend medium containing 0.8 % Suc with 0 or 20 uM jasmonic acid for 10 d at 22 °C under Wc light of 100 umol m'2 s". Expression of UBC21 (At5g25 760) was analyzed as a reference. Bars, black bars, -JA and white bars, + JAzo uM- Quantification by qRT-PCR was performed with 3 independent experiments. 187 we; wwo pd Nwd mwd m—a 2.? E5 8 023.8 no; #50 No.0 vw .o wad “353% 20m 03- E5 8 832v 006 wwd mhd wwd E .o ”monocotmo 0%. 2e o (IEDEUVSHdO) uorssoxdxe GAIIBIQH v '1‘: '— Siom>UAS-BVR, Col-O wild-type (Col-0 WT), jar] , myc02-5, C20 wild-type (C20 WT), hy]-1 and hy2-1 lines were grown on Phytablend medium containing 0.8 % Sue with 0 or 20 pM jasmonic acid for 10 d at 22 °C under We light of 100 umol m2 s". Expression of UBC21 (At5g25 760) was analyzed as a reference. Bars, black bars, -JA and white bars, + JAZO “M. Quantification by qRT-PCR was performed with 3 independent experiments. 189 om; am; :6 mmd mwd mod mud wvd a. 09 see a. a x. .2 94 «e [e A. cw.— nm.m 9%,? hm. ano fif: mush? owd omdo 4.4% an? no.0 mmd w—A mmd O 3% 9D..09nenc 2:. Es 0. 3:28 mm .o ”Ooh—Oh ohm—u 20.4 AS- 55 2 e233 _ 0.0 beep—0&6 2Q 9¢...n.9nonc 04/: 44a 449 one 444 have 449 6% 9% : :: 21 8 fl. 1U 0.0 V“. o o (1508./yum) uorssardxa muep‘d I W. M S.-. ed 190 4.5 References Alabadi, D., and Blazquez, M. A. (2009). 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Wamasooriya SN and Montgomery BL, (2010), Investigating Tissue- and Organ- specific Phytochrome Responses using F ACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana, Journal of Visualized Experiments, 39. https://www.jove.com/index/details.stp?id== l 925, doi: 10.3791/1925 199 A1 Overview Phytochromes are the red/far-red light-absorbing photoreceptor class and they mediate unique and overlapping functions in all higher plant systems in which they have been studied (Franklin and Quail, 2010). In the life cycle of Arabidopsis, an array of light responses is regulated by phytochromes and such responses are often localized to specific plant tissues or organs (Montgomery, 2008) and indicate the existence of spatial-specific phytochrome-dependent responses. Even though much progress on the discovery and elucidation of individual and redundant phytochrome functions have been made through mutational analyses, conclusive evidence on discrete photoperceptive sites and the molecular mechanisms of localized pools of phytochromes that mediate spatial-specific phytochrome responses still remain elusive. Based on the hypotheses that specific sites of phytochrome photoperception regulate tissue- and organ-specific aspects of photomorphogenesis, and that localized phytochrome pools engage distinct subsets of downstream target genes in intracellular and/or in cell-to-cell signaling, a biochemical approach to selectively reduce functional phytochromes in a tissue- and organ-specific manner within transgenic plants was developed. The biochemical approach is based on a bipartite enhancer-trap strategy that results in transactivation of the expression of a gene under control of the Upstream Activation Sequence (UAS) element by the transcriptional activator GAL4 (Laplaze et a1., 2005). In the UAS-BVR parent, the biliverdin reductase (B VR) gene under the control of the UAS is silently maintained in the absence of GAL4- mediated transactivation (Costigan, Wamasooriya and Montgomery, unpublished data). Genetic crosses between a UAS-BVR transgenic line and a GAL4-GP P enhancer trap line (Haseloff, 1999) result in specific expression of the BVR gene in cells marked by 200 GFP expression (Costigan, Wamasooriya and Montgomery, unpublished data). BVR accumulation in Arabidopsis plants results in phytochrome chromophore deficiency at specific locations in vivo (Lagarias et a1., 1997; Montgomery et a1., 1999; Wamasooriya and Montgomery, 2009). Thus, progeny transgenic plants that have been produced through genetic crosses display GAL4-dependent GFP expression, as well as activation of the BVR gene that leads to biochemical inactivation of phytochrome (Figure A1). Through comparative photobiological and molecular genetic analyses of BVR transgenic lines, insight into tissue- and organ-specific phytochrome-mediated responses that are associated with corresponding sites of photoperception can be gained. Putative downstream target genes involved in mediating spatial-specific phytochrome responses can be identified by fluorescence activated cell sorting (FACS) of GFP-positive, enhancer-trap-induced BVR-expressing plant protoplasts followed by cell-type-specific gene expression profiling by microarray analysis. FACS-mediated cell-type-specific expression profiling will expand our understanding of sites of light perception, the mechanisms through which various tissues or organs cooperate in light-regulated plant growth and development, and advance the molecular dissection of complex phytochrome-mediated cell-to-cell signaling cascades. A2 Materials and Methods A2.l Generation of J0571>>UAS-BVR UAS-BVR plants at T3 generation were obtained as described in section 4.2.1. T3 plants of UAS-BVR were crossed with the J057l enhancer trap line (Figure A1), which exhibits GFP expression in cortex/endodermis including initials, with a broad pattern of 201 expression in hypocotyl and cotyledons (Figure A2; Haseloff, 1999). Selection of F1 seedlings based on kanamycin resistance and PCR screening was performed as described in section 4.2.1. Fl seedlings that were positive in PCR with BVR- and GAL4-specific primer sets were transferred to soil to obtain F2 seeds and to progress to the F3 generation. F3 seeds of the UAS-BVRXJ0571 cross were used for subsequent analyses (J0571>>UAS-BVR). A2.2 Plant Growth Wild-type (C24 WT), parental enhancer trap line (J0571) and confirmed J0571>>UAS-BVR progeny isolated as described in section A2.1, were sown on soil, i.e. ~ 2000 sterilized seeds per line. J107l, which exhibits GFP expression in vascular/dermal tissue throughout the seedling (Figure A2), was included as a control to determine the efficiency of F ACS where GFP expression is observed in limited tissues. C24 WT, J0571, J 1071 and J0571>>UAS-BVR plants were grown for 5 weeks on soil under white illumination of 100 pmol m-2 s.1 at 22 °C and 70 % humidity. A2.3 Reagent Preparation TEX buffer was prepared as follows. For 1 liter TEX buffer, the following components: 3.1 g of Gamborg's BS salts (Catalog No. G5768, Sigma, M0), 0.5 g of 2- (N-morpholino) ethanesulfonic acid (MES; at 2.56 mM, Catalog No. M3671, Sigma, MO), 0.75 g of calcium chloride dihydrate (CaC12.2H20; at 6.75 mM, Catalog No. C2536, Sigma, MO), 0.25 g of ammonium nitrate (NH4NO3; at 3.12 mM, Catalog No. , 202 A3795, Sigma, MO), 136.9 g of sucrose (at 0.4 M, Catalog No. 4072-05, J .T. Baker, NJ) were weighed and dissolved completely in ~ 900 mL of deionized, distilled water (ddHZO) and the pH was adjusted to 5 .7 with l M KOH. Final volume was adjusted to 1 liter and was filter sterilized with a 0.2 pm bottle-top filter connected to a vacuum pump. 10X leaf digestion stock solution was prepared by dissolving 2 % (w/v) Macerozyme R- 10 (Catalog No. MSPC 0930, SERVA Electrophoresis GmbH, Crescent Chemical Company, NY), 4 % (w/v) Cellulase “Onozuka” R-lO (Catalog No. PTC 001, SERVA Electrophoresis GmbH, Crescent Chemical Company, NY) in TEX buffer followed by filter sterilization with 0.2 pm filter. 5 mL aliquots were frozen at -80 °C. 1X leaf digestion solution was prepared by adding 45 mL fresh TEX buffer to a 5 mL aliquot of 10X leaf digestion stock solution prior to use. A2.4 Leaf Protoplast Isolation Leaf protoplast isolation protocol was adapted from Denecke and Vitale (1995). Green, healthy leaves were collected from 5-week-old plants (~ 250 mL of leaves loosely packed in a beaker) and rinsed 4x with ~ 40 mL ddeO, followed by rinsing twice with sterile ddeQ. Using a #20 scalpel, leaves were cut into thin strips and leaf tissue strips were divided equally into two 50 mL sterile plastic tubes. The 1X leaf digestion solution was prepared as described in section A2.3 and ~ 25 mL of IX leaf digestion solution per 50 mL tube was added to cover all leaf tissue. Leaf tissue in 1X digestion solution was vacuum infiltrated in open tubes for 1 hr at room temperature using a vacuum desiccator connected to a water pump followed by 3-hr incubation at room temperature on a rocker 203 with gentle shaking. Capped tubes with leaf tissue in 1X leaf digestion solution were kept wrapped in aluminum foil during this incubation to prevent exposure to light. After 3 hrs, the rocker speed was increased for ~ 2 min to release protoplasts. The crude protoplast suspension was filtered through two layers of sterile cheese cloth to remove debris and the filtrate was collected in a sterile glass beaker. The filtrate was filtered through a sterile 100 pm nylon mesh into a sterile Petri dish. The flow through was collected and transferred to a new sterile 50 mL tube. About 15-20 mL of fresh TEX buffer was _used to wash the sterile Petri dish and any protoplasts adhering to the surface were collected. The flow through was centrifitged using a swing bucket rotor at lOOxg at 10 °C for 15 min (Acceleration 6, Deceleration 0; Allegra® X-l 5R Centrifuge, Beckman Coulter, CA). ~ 25 - 30 mL of the liquid (which contains residual and pelleted debris) below the floating protoplast layer, was removed with a sterile 9” glass Pasteur pipette connected to a peristaltic pump (Model 3100, Welch Rietschle Thomas, IL) without disturbing the floating protoplast layer and ~ 1.0 — 15 mL volume was left in the 50 mL tube. Fresh TEX buffer was added to a final volume of 40 mL while gently resuspending the protoplasts. Centrifugation, removal of liquid below the floating protoplast layer and gentle resuspension of the protoplasts by adding fresh TEX buffer were repeated two times as described above to remove as much cellular debris as possible. The centrifiJgation time is reduced to 10 min in the first repetition and to 5 min in the final repetition. Floating protoplasts were aspirated with a cut, sterile 1 mL transfer pipette into a new 15 mL tube wrapped in aluminum foil to prevent exposure to light and were kept on ice until sorting. Sorting was performed directly afier isolation of leaf protoplasts. 204 A2.S Protoplast Sorting by Fluorescence-Activated Cell Sorting (FACS) Before isolated protoplasts were subject to sorting, protoplasts were examined by Confocal Laser Scanning Microscopy (CLSM) using a 488-nm argon laser for excitation to confirm protoplast integrity, minimal amount of debris (to avoid clogging the FACS sorting nozzle) and the presence of GFP fluorescence in the protoplast pool. Isolated protoplasts were sorted in TEX buffer via FACS (BD F ACSVantage SE, BD Biosciences, CA) using a ZOO-pm nozzle on a macro sort head at event rates between 6,000 and 15,000, with a system pressure of around 3-9 p.s.i. following an adapted protocol (Bimbaum et a1., 2005). C24 WT non-GF P protoplasts were used to determine the autofluorescence thresholds. To collect GFP-positive protoplasts in J0571, J1071 and J 0571>>UAS-BVR suspensions, protoplasts were sorted using an air-cooled argon laser (Spectra Physics Model 177, Newport Corporation, CA) operated at 100 mw on a 488- nm argon line. GFP fluorescence was detected using a 530/30 band pass filter. For J0571>>UAS-BVR, while GFP-positive protoplasts were being collected, in a separate channel, GFP-negative protoplasts were collected as a negative control for subsequent microarray analyses. Following the collection of GFP-positive protoplasts, sorted protoplast fractions were examined for presence/absence of GFP fluorescence and protoplast integrity by CLSM as described above. Total RNA from sorted protoplast fractions was extracted using RNeasy® Plant Minikit (Catalog No. 16419, Qiagen, CA) including on-column DNase treatment (Catalog No. 79254, Qiagen, CA) according to manufacturer’s instructions. The quantity of RNA in the samples for C24 WT (GFP negative), J057l (GF P positive), J 1071 (GP P positive) and J0571>>UAS-BVR (GF P positive) was analyzed by spectrometry (NanoDroplOOO, Thermo Scientific, MA). 205 A3 Results A significant number of GFP-positive protoplasts were detected in the GF P channel by FACS for J0571, J 1071 and J0571>>UAS-BVR samples (Figure A3 and A4). In optimization assays using GFP-enhancer trap lines, J0571 with GF P expression in cortex/endodermis including initials, with a broad pattern of expression in hypocotyl and cotyledons, displayed ~ 17 % to 24 % GFP-positive protoplasts, whereas the line with vascular and dermal expression, J 1071, had ~ 1.4 % GFP-positive protoplasts. F3 progeny, J0571>>UAS-BVR, displayed ~ 32 % GFP-positive protoplasts (Table A l ). Sorting of 3 x 500 uL (~ 1.5 mL total of protoplast suspension) of 10571 protoplasts for l h gave ~ 100,000 GFP-positive protoplasts. Sorting of 3 x 500 pL (~ 1.5 mL total protoplast suspension) of J 1071 protoplasts for 1.5 h gave ~ 3,000 GFP-positive protoplasts. Sorting of 3 x 500 pL (~ 1.5 mL total protoplast suspension) of J0571>>UAS-BVR protoplasts for 1.5 h gave ~ 104,000 GFP-positive protoplasts and 109, 000 GFP-negative protoplasts (Table A1). Confocal images indicated a very high yield of protoplasts for both J0571 and J 1071 samples before sorting was carried out (Figure AS-C and E), and the sorted fractions contained only bright GFP-fluorescent protoplasts (Figure AS-G and E). The non-GF P protoplasts can also be sorted and collected in a separate channel. For J0571>>UAS-BVR, the protoplasts in non-GF P protoplast fraction displayed only chlorophyll autofluorescence (Figure A6-G). This observation indicates that intact GFP-positive protoplasts can be sorted via F ACS and the fractions, which were sorted based on the presence of GF P, do not contain GFP-negative protoplasts. RNA extraction from isolated protoplasts (1 mL) yields RNA of sufficient quantity for detection by fluorospectrometry (Table A2). RNA yields from pre-sorted 206 C24 wild-type protoplasts or F ACS-sorted, GFP-positive protoplast fractions exceeded the minimum 20 ng needed for use in RNA-labeling assays for microarray (Table A2). cDNA can be prepared for hybridization as described in Affymetrix GeneChip® Expression Analysis Technical Manual and then hybridized to GeneChip®Arabidopsis ATHl Genome Array (Catalog No. 900385, Affymetrix, Inc., CA) for cell-, tissue- or organ-type-specific gene expression profiling. A4 Discussion Gene expression profiling through microarrays has revealed that more than 30 % of the genes in Arabidopsis seedlings are light regulated (Ma et a1., 2001) and has identified a vast group of genes encoding light signal transduction components involved in the phytochrome signaling cascade (Chen, Chory, and Fankhauser, 2004; Ulm and Nagy, 2005). Such studies suggest that light induces rapid and long-term changes in gene expression. A wealth of data indicate that phytochrome family members are expressed in a spatial- as well as temporal-specific manner and localized pools of phytochrome can mediate discrete physiological functions in planta (Bischoff et a1., 1997; De Greef and Caubergs, 1972; De Greef and Verbelen, 1972; Parcy, 2005; Goosey, Palecanda, and Sharrock, 1997; Montgomery, 2008; Sharrock and Clack, 2002; Zeevaart, 2006). Each localized pool of phytochromes may control only a subset of distinct developmental and adaptive responses. Moreover, it is likely that downstream signaling components interact with phytochromes in a cell- and tissue-specific manner in mediating discrete physiological responses (Ma et al., 2005; Montgomery, 2008; Neff, Fankhauser, and Chory, 2000). Through comparative analysis of gene expression changes in GAL4-GF P 207 X UAS-BVR progeny and parental lines, distinct changes in gene expression that result fi'om localized phytochrome deficiencies can be determined. Based on gene expression profiles, genes that encode negative and positive regulators of phytochrome-mediated cell-to-cell signaling can be elucidated. Recent data suggest that several hundred transcripts are induced by protoplasting of plant tissues (Bimbaum et a1., 2003). Thus, the ideal negative control for microarray analysis is RNA isolated from non-GF P protoplasts collected during sorting. Transcripts that are induced by protoplasting of plant tissue can be excluded from data analysis using these controls, resulting in the identification of target genes specifically involved in tissue— and organ-specific phytochrome signaling and/or responses. To confirm steady-state changes in expression of genes identified via cell-type-specific expression profiling, qRT-PCR can be carried out on poly (A) RNA isolated from the sorted GFP-positive and GFP-negative protoplasts. Genes whose expression is changed significantly in microarray analyses and confirmed by qRT-PCR are identified as candidate genes involved in discrete phytochrome-mediated intercellular signaling. If T-DNA insertion mutants are already available in the identified candidate genes, they can be analyzed for light-impaired phenotypes in R and FR. By comparing the phenotypes of mutants carrying mutations in candidate genes and known phytochrome mutants to wild type, specific responses in which candidate genes have regulatory roles can be identified. If the mutants with mutations in candidate genes display phytochrome-deficient phenotypes in R and/or FR conditions, this will confirm that the identified downstream target genes are potentially involved in mediating distinct phytochrome-regulated responses. 208 Figure Al GAL4 enhancer-trap-based induction of Biliverdin reductase (B VR) expression in transgenic Arabidopsis thaliana plants. (A). An individual selected from a library of GAL4-based enhancer trap lines, which contain a GAL4-responsive green fluorescent protein (GFP) marker gene, is crossed with a line containing a GAL4-responsive target gene (B VR) to induce expression of the BVR gene in GAL4-containing cells marked by GFP fluorescence. Based on a figure from Dr. Jim Haseloff (http://www.p1antsci.cam.ac.uk/Haseloff/geneControl/GAL4Frame.html). (B) Production of phytochrome chromophore, phytochromobilin (P¢B) and holophytochrome in parent lines. 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W., Wang, J. Y., Lambert, G. M., Hirst, J. A., Galbraith, D. W., and Benfey, P. N. (2005). Cell type-specific expression profiling in plants via cell sorting of protoplasts from fluorescent reporter lines. Nat Methods 2(8), 615-619. Bimbaum, K., Shasha, D. B., Wang, J. Y., Jung, J. W., Lambert, G. M., Galbraith, D. W., and Benfey, P. N. (2003). A gene expression map of the Arabidopsis root. Science 302(5652), 1956-1960. Bischoff, F., Millar, A.J., Kay, S.A., and F uruya, M. (1997) Phytochrome-induced intercellular signalling activates cabzzluciferase gene expression, The Plant Journal 12(4), 839-849. Chen, M., Chory, J ., and Fankhauser, C. (2004). Light signal transduction in higher plants. Annu Rev Genet 38, 87-117. De Greef, J .A., and Caubergs, R. (1972). Interorgan correlations and phytochrome: leaf expansion. Arch Int Physiol Biochim 80, 961 -962. De Greef, J .A., and Verbelen, J .P. (1972). Interorgan correlations and phytochrome: changes in plastid ultrastructure during de-etiolation processes. Arch Int Physiol Biochim 80, 962-963. Denecke, J ., and Vitale, A. (1995). The use of protoplasts to study protein synthesis and transport by the plant endomembrane system. Methods Cell Biol 50, 335-48. Franklin, K. A., and Quail, P. H. (2010). Phytochrome functions in Arabidopsis development. J Exp Bot 61(1), 1 1—24. Goosey, L., Palecanda, L., and Sharrock, R. A. (1997). Differential patterns of expression of the Arabidopsis PHYB, PHYD, and PHYE phytochrome genes. Plant Physiol 115(3), 959-969. Haseloff, J. (1999). GP P variants for multispectral imaging of living cells. Methods Cell Biol 58, 139-151. Lagarias, D. M., Crepeau, M. W., Maines, M. D., and Lagarias, J. C. (1997). Regulation of photomorphogenesis by expression of mammalian biliverdin reductase in transgenic Arabidopsis plants. Plant Cell 9(5), 675-688. Laplaze, L., Parizot, B., Baker, A., Ricaud, L., Martiniere, A., Auguy, F., Franche, C ., Nussaume, L., Bogusz, D., and Haseloff, J. (2005). GAL4-OF P enhancer trap lines for genetic manipulation of lateral root development in Arabidopsis thaliana. J Exp Bot 56(419), 2433-2442. 218 Ma, L., Li, J., Qu, L., Hager, J., Chen, Z., Zhao, H., and Deng, X. W. (2001). Light control of Arabidopsis development entails coordinated regulation of genome expression and cellular pathways. Plant Cell 13(12), 2589-2607. Ma, L., Sun, N., Liu, X., Jiao, Y., Zhao, H., and Deng, X. W. (2005). Organ-specific expression of Arabidopsis genome during development. Plant Physiol 138(1), 80- 91. Montgomery, B. L. (2008). Right place, right time: Spatiotemporal light regulation of plant growth and development. Plant Signal Behav 3(12), 1053-1060. Montgomery, B. L., Yeh, K. C., Crepeau, M. W., and Lagarias, J. C. (1999). Modification of distinct aspects of photomorphogenesis via targeted expression of mammalian biliverdin reductase in transgenic Arabidopsis plants. Plant Physiol 121(2), 629-639. Neff, M. M., Fankhauser, C., and Chory, J. (2000). Light: an indicator of time and place. Genes Dev 14(3), 257-271. Parcy, F. (2005) Flowering: a time for integration. Int J Dev Biol 49, 585-593. Sharrock, R. A., and Clack, T. (2002). Patterns of expression and normalized levels of the five Arabidopsis phytochromes. Plant Physiol 130(1), 442-456. Ulm, R., and Nagy, F. (2005). Signalling and gene regulation in response to ultraviolet light. Curr Opin Plant Biol 8(5), 477-482. Wamasooriya, S. N., and Montgomery, B. L. (2009). Detection of spatial-specific phytochrome responses using targeted expression of biliverdin reductase in Arabidopsis. Plant Physiol 149(1), 424-433. Zeevaart, J. A. (2006). F lorigen coming of age after 70 years. Plant Cell 18(8), 1783- 1789. 219 MICHIGAN STATE UNIVE SITY LIB 111 I! II 1111116111111“ 93 322 4 I l 3 . . .... ............... ............ ................ ....... c . n 0 .......... ..... .......... v o ..... ..... ......... .......... ................... ...... ........ ........... ........ ..........