Starch Function in Ornamental Tobacco Floral Nectary Development Gang Ren Iowa State University

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2007 Starch function in ornamental tobacco floral nectary development Gang Ren Iowa State University

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This Dissertation is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Retrospective Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. Starch function in ornamental tobacco floral nectary development

Gang Ren

A dissertation submitted to the graduate faculty

in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

Major: Genetics

Program of Study Committee: Robert Thornburg, Major professor Martha James Basil Nikolau Drena Dobbs Kang Wang

Iowa State University

Ames, Iowa

2007

UMI Number: 3259503

UMI Microform 3259503 Copyright 2007 by ProQuest Information and Learning Company. All rights reserved. This microform edition is protected against unauthorized copying under Title 17, United States Code.

ProQuest Information and Learning Company 300 North Zeeb Road P.O. Box 1346 Ann Arbor, MI 48106-1346 ii

Table of contents

List of figures xii List of tables x Acknowledgements xi Abstract xiii

Chapter 1 General introduction 1 1.1 Hypothesis 3 1.1.1 Hypothesis 1 5 1.1.2 Hypothesis 2 5 1.2 Plant floral nectary and nectar 6 1.2.1 Plant nectar 7 1.2.2 Nectar secretion models 8 1.2.3 Starch in floral nectaries 10 1.3 Starch metabolism in higher plants 11 1.3.1 Starch biosynthesis 13 1.3.2 Starch degradation 20 1.4 Outline of this dissertation 21

Chapter 2 Starch accumulation in ornamental tobacco floral nectary 23 Abstract 25 2.1 Introduction 26 2.2 Materials and methods 27 2.2.1 Materials 27 2.2.2 Plants 27 2.2.3 Transport of radiolabeled sugars 27 2.2.4 Carbohydrates analysis 28 Thin layer chromatography of radiolabeled sugars 28 Nectar carbohydrate composition 28

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2.2.5 Starch analysis 29 Isolation of starch from nectaries 29 Starch quantitation 29 Starch structural analysis 29 Amylose content analysis 30 2.2.6 Light Microscopy 31 2.2.7 Scanning Electron Microscopy 32 2.3 Results 32 2.3.1 Nectary development 37 2.3.2 Starch in nectaries 40 2.3.3 Starch is dynamically processed during nectary development 45 2.3.4 Role of transported photosynthate in nectar production 51 2.3.5 Is tobacco a model for other plant species? 59 2.4 Discussion 59 2.4.1 Transient nature of the nectary starch 62 2.4.2 Role of transported photosynthate in nectar production 63

Chapter 3 Regulation of starch metabolism in ornamental tobacco floral nectary 69 Abstract 71 3.1 Introduction 72 3.2 Materials and methods 74 3.2.1 Plant Materials 74 3.2.2 rDNA methods 74 3.2.3 PCR methods 75 Primer design 75 General PCR 75 RT-PCR detection 75 Quantitative RT-PCR detection 76 3.2.4 Bioinformatics tools 77 3.2.5 Microscopy methods 77

Transmission Electron Microscopy 77 Quantitation of plastid volume 78 3.2.6 RNA isolation 78 3.2.7 protein methods 79 3.2.8 Real time RT-PCR detection 82 Protein isolation and enzyme activity assay 79 Protein isolation and immunodetection 79 3.3 Results 80 3.3.1 Starch metabolic genes 84 3.3.2 Regulation of starch biosynthesis 97 Sucrose synthase in developing ornamental tobacco floral nectary 97 ADP-glucose pyrophosphorylase in developing floral nectaries 99 3.3.3 Additional starch metabolic genes in developing floral nectaries 102 Starch synthases 102 Starch branching enzymes 105 Starch debranching enzymes 105 3.3.4 Quantitative RT-PCR (qRT-PCR expression) 108 3.4 Discussion 113

Chapter 4 β-carotene meatabolism in ornamental tobacco floral nectary 125 Abstract 127 4.1 Introduction 128 4.2 Materials and methods 129 4.2.1 Plant material 129 4.2.2 Light microscopy 129 4.2.3 Vibratome sections 130 4.2.4 Electron microscopy 130 4.2.5 Biochemical analyses 131 Carotenoids analysis 131 Ascorbic acid analysis 131

4.2.6 Labeling of carotenoids with [14C]-sucrose 132 4.2.7 Oligonucleotides for RT-PCR analysis 132 4.3 Results 133 4.3.1 Light Microscopy: S9 through S12 133 4.3.2 Electron microscopy: S9 through S12 138 S9 plastids 138 The S10 and S11 plastids 141 The S12 plastids 141 4.3.3 Biochemical and Molecular Genetic Analyses 144 4.4 Discussion 165

Chapter 5 Future work 170 5.1 Introduce of transgenic plants based future work 170 5.2 Materials and methods 179 5.2.1 Plante lines 179 5.2.2 RNAi constructs 179 5.2.3 Overexpression constructs 183 5.2.4 Plant transformation and plant selection 183 RNAi expression transgenic plants 183 Overexpression transgenic plants 183 Seeds collection 183 5.2.5 Genomic DNA verification of transgenic plants 184 Plant genomic DNA extraction 184 Genomic DNA PCR 184 5.2.6 Protein isolation 184 5.2.7 Western blot assay 185 5.2.8 Starch staining analysis 185 5.2.9 nectar carbohydrate analysis 185 5.3 Results 186 5.3.1 Transgenic plants verification 186

5.3.2 Starch in ornamental flowers 186

Chapter 6 Summarization and discussion 198 6.1 Starch and floral development 200 6.2 Starch and nectar production 200 6.3 Starch biochemistry in developing tobacco floral nectary 202

Appendices 205 Appendix A Index of EST clones 206 Appendix B Index of oligonucleotides 211 Appendix C Index of constructs 220 Appendix D Index of transgenic plants 225

References 234

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List of figures

Figure 1.1 Structure of tobacco flowers 2 Figure 1.2 The core pathway of starch biosynthesis in higher plants 14 Figure 1.3 The core pathway of starch biodegradation in higher plants 15 Figure 1.4 Biochemical reaction of starch metabolic enzymes 16 Figure 2.1 Light and scanning electron microscopy (SEM) graphs of Arabidopsis thaliana and ornamental tobacco floral nectaries 34 Figure 2.2 Events occurring during development of tobacco nectaries 38 Figure 2.3 Light microscope images of 1-μm-thick LR White resin sections of tobacco nectaries at five different developmental stages stained for differential contrast enhancement with toluidine blue O or PAS technique for cell walls and starch 41 Figure 2.4 Accumulation of starch in nectary 43 Figure 2.5 FACE analysis of nectary starch 46 Figure 2.6 Amylose/amylopectin content of starch isolated from ornamental tobacco floral nectaries at various stages of development 48 Figure 2.7 TLC analysis of radiolabeled nectar 53 Figure 2.8 Labeling of C14-sucrose in various floral stages 55 Figure 2.9 Light microscopic images of 1-μm-thick LR White resin sections of Arabidopsis thaliana floral nectaries stained with PAS technique 60 Figure 2.10 Comparison of biochemistry with nectary development 66 Figure 3.1 Starch granule changes in developing floral nectary parenchyma cells 81 Figure 3.2 Neighbor-joining trees for ornamental tobacco starch metabolism genes 91 Figure 3.3 Sucrose synthases (SuSy) expression in ornamental tobacco 100 Figure 3.4 AGPase expressions in ornamental tobacco floral nectary 103 Figure 3.5 RT-PCR detection of starch metabolism related gene expression profile in ornamental tobacco 106 Figure 3.6 Real-time RT-PCR detection of starch metabolism related gene expression in ornamental tobacco floral nectaries 110

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Figure 3.7 Model of starch mass accumulation and starch metabolic enzyme activity in ornamental tobacco floral nectary 119 Figure S3.1 Comparison of splice-variant of α–amylase 121 Figures 4.1-4.18 Images of isolated whole, and 30-µm-thick cross and longitudinal vibratome sections of tobacco gynoecia and basal nectaries from Stage 2 through Stage 12 observed with normal light or between crossed polarizers 136 Figures 4.19-4.24 Light and transmission electron microscope sections through nectary at Stages 9-12 139 Figures 4.25-4.31 Transmission electron microscope sections showing nectary cell plastids from S9 through S12 142 Figure 4.32 Identification of ornamental tobacco nectary carotenoid as β-carotene 147

Figure 4.33 Analysis of nectary expressed carotenoids extracted from LxS8 149 Figure 4.34.RT-PCR of gene encoding β–carotene biosynthetic enzymes 153 Figure 4.35 Interconnection of biochemical pathways in the nectary 156 Figure 4.36 Thin layer chromatogram (TLC) visualization of nectary expressed carotenoids 159 Figure 4.37 Quantity of ascorbate extracted from six nectaries of ornamental tobacco flowers at different developmental stages 161 Figure 4.38 Ascorbate metabolism 163 Figure 5.1 Scheme of transgenic plants 171 Figure 5.2 3D structure of the DNA-binding domain of sucrose responsive element binding proteins 175 Figure 5.3 Gene expression pattern of SREBP detected by RT-PCR 177 Figure 5.4 Gene expression of mevalonate pathway and MEP pathway genes detected by RTPCR 181

Figure 5.5 Genomic DNA PCR verification of transgenic plants 187 Figure 5.6 Immunodetection of transgenic plants 189 Figure 5.7 Starch accumulation detected by iodine staining in LxS8 flowers 191

Figure 5.8 Starch staining of various transgenic plants line in ornamental tobacco 194 Figure 5.9 Nectar carbohydrates measurement of various transgenic plants line in ornamental tobacco 196 Figure 6.1 Speculated biochemical process in ornamental tobacco floral nectary development 203 Figure A1 Overview of the metabolism pathways in ornamental tobacco floral Nectary 207 Figure A2 Maps of subcloning vectors 208 Figure C1 Map of constructs for transgenic plants 222

List of tables

Table 2.1 Composition of ornamental tobacco LxS8 nectar 36 Table 2.2 Composition of ornamental tobacco LxS8 nectar 52 Table 2.3 Composition of labeled nectar in ornamental tobacco LxS8 58 Table 3.1 Volumes of cell, plastid within cell, and starch granule within plastid in developing ornamental tobacco floral nectary cells 83 Table 3.2 Starch metabolic genes isolated from ornamental tobacco 85 Table 3.3 Oligonucloedites primer for gene isolation 86 Table 3.4 Comparison of Solanum tuberosum starch metabolic genes 98 Table 3.5 RT-PCR primers for ornamental tobacco starch metabolism genes expression detection 93 Table S3.1 Target genes involved in starch metabolism genes in Solanum tuberosum and ornamental tobacco LxS8 123 Table S3.2 1 Volumes of cell, plastid within cell, and starch granule within plastid in developing ornamental tobacco floral nectary cells 124 Table 4.1 Oligonucleotides for RT-PCR analysis 134 Table 4.2 β-carotene biosynthesis genes used in this analysis 145 Table 4.3 Ascorbate biosynthesis genes used in this analysis 152 Table 5.1 Transgenic plants summarization 173 Table 5.2 Primers for transgenic plant verification 180 Table A1 Index of ornamental tobacco LxS8 clones 209 Table B1 Index of oligonucleotides 212 Table B2 Oligonucleotides for transgenic plant constructs 218 Table C1 Index of constructs for tobacco transgenic plants 224 Table D1 Index of detailed tobacco transgenic plants 226

Acknowledgements

This dissertation is dedicated to my beloved wife, Yan Zhou, and my adorable little son, Daniel Ren. I want to thank them for backing me up through my entire PhD study. For almost four years, I have been working in Dr. Robert Thornburg’ lab, a very scientific and energetic group. It has been a great experience to me in my whole life. I want to thank the co-worker, Ms. Rosanne Healy, who performed all of the Microscopy work. I want to thank Dr. Clay Carter in University of Minnesota. Dr. Carter started the pioneer work of ornamental tobacco floral nectar and nectary when he was working under the supervision of Dr. Robert Thornburg for his PhD study. I want to thank three visiting scholars who had worked in Dr. Robert Thornburg’s lab: Dr. Saqlan Naqvi in University of Arid Agriculture, Pakistan, Dr. Adel Guirgis in Menofiya University, Egypt, and Dr. Sanggyu Park in Daegu University, South Korea. We spend a lots of valuable and wonderful time to work together. These three professors had been very helpful to consult me the rightful and efficient way of doing scientific research. They also introduced their own projects in their home countries to me and let me understand the research works out of my study field. I also want to thank Dr. Guangxi Bai, Ms. Anna Klyne, and Mr. Guangyu Liu for discussion and cooperation of the laboratory works. I want to thank Dr. Harry Horner in Department of Genetics, Development and Cell Biology & Microscopy and NanoImaging Facility, Iowa State University, Dr. Horner consulted on the methods of Microscopic work. I want to Dr. Martha James in Department of Biochemistry, Biophysics, and Molecular Biology, Iowa State University. Dr. James consulted on the starch project. She is the Co-PI of the starch project (NSF grant (IBN-0235645) to Dr. Robert Thornburg and Dr. Martha James). She also served as my POS committee. I want to thank all of my committee members: Dr. Martha James, Dr. Basil Nicokau, Dr. Drena Dobbs, Dr. Kan Wang. Thank you all for your assistances during my study.

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And finally, I especially thank my major professor, Dr. Robert Thornburg. Without his advice, guidance, encouragement, and support, it is impossible for me to finish my research project and my PhD study. I want to thank him for bringing up a wonderful research group that has so many professors and postdoctoral fellows working in one lab. I want to thank him for creating a nice working and training environment so that I spent four years of good quality time in this lab. I want to thank him for consulting on the overall creative instructions and supporting for the research project. I want to thank him for contributing the writing, reviewing and editing of this dissertation. And more importantly, I want to thank him for helping me out of those obstacles as that an international student might encounter.

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Abstract

Morphological changes of floral nectary gland and subcellular transition of floral nectary cells of ornamental tobacco LxS8 were exclusively investigated in this study. Enlargement of the floral nectary gland of ornamental tobacco LxS8 that occurs during development is accompanied by a major accumulation of Periodic Acid-Schiff’s staining (PAS-staining) starch grains in nectary amyloplasts. Quantification of starch purified from the nectary at various stages of development showed little starch at an early developmental stage, soon thereafter the amount of starch increased dramatically, reaching a peak approximately 24 hours prior to anthesis. After this time, the amount of starch declined dramatically until the flower reached anthesis, suggesting that the accumulated starch was converted to sugars (sucrose, glucose, and fructose) for nectar production that occurs prior to anthesis. A Transmission Electron Microscopy (TEM) study of plastid development further verfied that ornamental tobacco nectaries accumulate large amounts of starch during floral development and hydrolyze that starch prior to anthesis. Compositional and structural analyses of nectary starch showed that amylose content and degree of amylopectin branching varied during nectary development. The average chain length in amylopectin was relatively short at early developmental stages, reached maximal length towards the end of nectary development, and was reduced again at anthesis, consistent with the synthesis of an increasingly complex form of starch up to middle stages of nectary development, followed by decreases in starch complexity and amount due to starch degradation in the mature nectary. Tobacco floral nectaries undergo changes in form and function. As nectaries change from green to orange, a new pigment is expressed. Analysis demonstrated that it is β-carotene. Plastids undergo dramatic changes. Early in nectary development, they divide and by stage 9 (S9) they are engorged with starch. About S9, nectaries shift from quiescent anabolism to active catabolism resulting in starch breakdown and production of nectar sugars. Starch is replaced by osmiophilic bodies, which contain needle-like carotenoid crystals. Between S9 and S12, amyloplasts are converted to chromoplasts. Changes in carotenoids and ascorbate

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were assayed and are expressed at low levels early in development; however, following S9 metabolic shift, syntheses of β-carotene and ascorbate greatly increase in advance of expression of nectar redox cycle. We propose that biosynthesis of these antioxidants is governed by availability of substrate molecules that arise from starch breakdown. These processes and events may be amenable to molecular manipulation to provide a better system for insect attraction, cross pollination, and hybridization. Thirty-nine different nectary-expressed cDNAs from the nectaries of ornamental tobacco plants that encode 14 different starch metabolic enzymes, 7 different mevalonate metabolic enzymes, 7 different MEP metabolic enzymes, 8 different β-carotene metabolic enzymes, and 3 different ascorbate metabolic enzymes were identified from ornamental tobacco LxS8. The translated protein sequences were used to identify gene functionality by comparison with well characterized protein sequences from Arabidopsis, maize and solanaceous species. Subsequently RT-PCR or real-time RT-PCR were used to evaluate gene expression throughout nectary development. This analysis revealed that starch metabolism was highly regulated at transcriptional level in ornamental tobacco floral nectary. Three different classes of starch metabolic gene expression pattern appeared. Most starch anabolic genes, including starch synthase 1 (SS1), starch synthase 3 (SS3) and granule-bound starch synthase (GBSS), were expressed early in nectary development (Stage 2 and Stage 6), and the expression was significantly down- regulated after Stage 9. In contrast, the starch catabolic genes, including branching enzyme 1 (BE1), isoamylase 1 (ISA1), α-amylase (AMY), β-amylase (BMY), were not highly expressed at early stages, but were significantly induced by Stage 9 of nectary development. A third class of gene expression that included R1, phosphorylase (PHO), and starch branching enzyme 2 (SBE2) were expressed through nectary development. Immnunodetection shows that ADP-glucose pyrophosphorylase (AGPase) small subunit and sucrose synthase (SuSy) were highly expressed at early stages. Enzyme activity analysis of AGPase also shows that enzyme activity was higher at early stages. Transcript analysis for carotenoid

and ascorbate biosynthetic pathways showed that these genes are significantly expressed at S6, prior to the S9 metabolic shift. Thus, formation of antioxidants β- carotene and ascorbate after the metabolic shift is independent of transcriptional regulation. We also evaluated the transport of radiolabeled sugars nectar. Consistency in the amounts and ratios of radiolabeled sugars in nectar indicates that the transported photosynthate sugars were subject to metabolic constraints in the nectary tissue prior to their incorporation into nectar. The velocity of radiolabeled sugars accumulated in nectar illustrates that transported photosynthate contributes to the later nectar production. A series of transgenic plants had been generated. The genetic evidence verified that starch regulates the timing and amount of floral nectar.

Chapter 1 General introduction

Flowers are the reproductive organ systems utilized by angiosperms to produce seeds to complete the plant’s life cycles (Darwin, 1890; Friis and Endress, 1990). Flowers are generally composed of a number of specific organs that function in a coordinated manner to attract pollinators (Darwin, 1890; Endress, 2001; Levy, 1998; McDade, 2004; Nicolson, 2002; Zik and Irish, 2003). These various floral organs generally include sepal, petal, filament and anther, stigma and style, and ovary (Figure 1.1). The female zygotes are located in the ovary, while the male zygotes are produced in the anthers. To achieve successful pollination, the plant must find a way to transfer the male zygotes to the stigma that connects through the transmitting tract to the ovary and the ovules. Higher plants have developed many pollination strategies to achieve the pollen transfer (Aizen, 1998; Darwin, 1859; Darwin, 1890). The transfer of pollen grains from the anther (male organ) to the stigma (female organ) in flowering plants normally relies on wind, gravity, or animals (Creco et al., 1996; Darwin, 1890; Kevan et al.; 1989; Kevan, 1993; Liu et al., 2006). However, perhaps the most commonly and reliably used strategy is the pollinatormediated pollination (Darwin, 1862). Pollinator-mediated pollination relies on other species to mechanically transfer pollen from the anther of one plant to the stigmas of other plants (Darwin, 1862; Feinsinger, 1983; Penny, 1983). For the pollinatormediated strategies to be successful, pollinators must receive a reward to maintain the interests in that process (Penny, 1983). Although floral scent and color play a role in insect attraction that includes sex deception to the insects (Aronne et al., 1993; Schiestl et al., 2003), the typical rewards are pollen and nectar (Cresswell, 1999; Feinsinger 1983; Klinkhamer et al., 1990; Neiland and Wilcock, 1998). In angiosperms it is thought that pollen was the initial reward that was offered to visiting pollinators and still today many species of pollinators continue to collect pollen during their floral visiting (Feinsinger, 1983; Percival, 1961; Temeles and Kress, 2003). During the process of evolution, flowers began to offer rich floral nectar to attract visiting pollinators (Feinsinger, 1983; Temeles and Kress, 2003), 2

Petal

Stigma

Anther

Sepal

Ovary

Nectary

Petiole

Figure 1.1 Structure of tobacco flowers A Nicotiana tobaccum; B Ornamental tobacco. Black bar represents 1 cm. 3

and that process is thought to have begun by the interaction of plants with animal pollinators for the rewards (Temeles and Kress, 2003). This pollinator-plant coevolution strategy has mutual benefits to the plant and the pollinator: highly concentrated sugars in nectar can provide the energy source for the pollinator and the recipients of this reward can help plants for pollen transfers (Feinsinger, 1983; Cresswell 1999; Baker, 1983). The pollinator-plant co-evolution is intensively studied (Cresswell, 1999; Darwin, 1862; Davis, 1997; Faegri et al., 1979; Feinsinger, 1983; Galetto and Bernardello, 2004; Penny, 1983), nonetheless, neither the evolutionary origin of nectar secretion nor the nectar secretion mechanism in various flowering species is clear. The origin of nectar secretion has been thought that it could have originated as a leakage of the phloem solution resulting from the structural weakness of developing tissue exposed to the elevated hydrostatic pressure in the phloem (leaky phloem model), or the nectar production also might have originated as a mechanism to remove the excess sugar solutes supplied by the phloem (sugar secretion model) (de la Barrera and Noble, 2004; Carlsbecker and Carlsbecker, 2005). In both speculations, the sugar in the phloem and polysaccharides (particularly starch) in nectary might be involved in the origin of nectar carbohydrates.

1.1 Hypotheses A voluminous research emerges to explore nectar secretion mechanisms. However, trying to make a conclusive statement about the source of nectar carbohydrates seems like not quite feasible. Nectary structure diversities due to coevolution with various types of pollinators and amount of nectar secreted during floral development (Arumugasamy et al., 1990; Baker, 1978; de la Barrera and Noble, 2004; Davis et al., 1986; Davis et al., 1996; Davis et al., 1997; Fahn, 1979; Freitas et al., 2001; Gillespie and Henwood, 1994) constrain the investigations of the source of nectar carbohydrates. Therefore, due to the complexity of nectaries in different species, interpretation of those observations must be fit for a specific and well-defined system. Two types of floral nectaries are defined based on the plastid 4

developed in nectary cells: chloroplast nectary and amyloplast nectary (Daivs et al., 1986; Figueiredo and Pais, 1992; Nepi et al., 2001). In chloroplast nectary, starch deposited in the chloroplasts accumulates during the day but degrades at night, and the amount of nectar is limited and not always available at any time during pollination (Davis et al., 1986; Figueiredo and Pais, 1992; Vogel, 1997; Vogel, 1998a; Vogel, 1998b; Vogel, 1998c). In amyloplast nectary, starch accumulates in the nectary. The starch accumulated at early floral stages degrades at maturation stages of floral nectaries but before the excessive nectar secretion (Nepi et al., 2001). To study the function of starch deposited in amyloplast nectary, an ornamental tobacco variety LxS8, which has distinguishable developmental floral stages with excessive nectar secreted at late stages of floral development (Koltunow et al., 1990; Kornaga et al., 1997; Thornburg et al., 2004), has been chosen to explore the relationship between nectar and starch towards the understanding of the nectar secretion mechanism. The ornamental tobacco LxS8 is an interspecific plant strain that has been identified as the amyloplast type nectary and secretes large amount nectar during floral development (Koltunow et al., 1990; Kornaga et al., 1997). The floral development in ornamental tobacco LxS8 can be easily identified into 12 different stages, and can be divided into four major developmental stages: pre-floral stages (stage -7 to stage 0), pre-maturation stages (stage 1 to stage 6), maturation stages (stage 7-stage 9), and post maturation stage (stage 10 to stage 12). The pre-floral stages take about one week, the pre-maturation stages and maturation take about two weeks, while the post-maturation stages only takes a few days. During the floral development, flower size, nectary size and color are also changing. At stage 1, the flower bud is about 1 cm. At stage 9, the floral tube reaches the maximum length about 7 cm. At stage 12, the flower is starting to open. 12 hours after stage 12, the flower is fully open. 48 hours after stage 12, the flower is starting to abscise. At stage 12, nectar starts to be secreted. After pollination, nectar starts to be reabsorbed. 5

Hypothesis 1: Starch deposited in ornamental tobacco floral nectary cells triggers floral nectar secretion and contributes to nectar production. The process of starch synthesis and degradation in the ornamental tobacco nectary is unique and represents a novel system that differs from other systems such as leaf transitory starch metabolism (Baroja-Fernandez, 2004; Cobesier et al., 1998; Smith et al., 2004) or the starch metabolism found in long term storage tissues such as maize endosperm (Cao et al., 2000; Dinges et al., 2001; Emes et al., 2003) and rice endosperm (Nakamura, 2002; Piperno, 2004), or in potato tubers (Bustos et al., 2004; Edwards et al., 1999; Geigenberger, 2003; Kubo et al., 1998) and tomato fruit (Park and Chung, 1998). Many research works have suggested that nectar carbohydrates from both of extrafloral nectaries and floral nectaries might be derived from phloem sap (de la Barrera and Noble, 2004; Pacini et al., 2003). However, those models cannot explain well about the various ratios of nectar carbohydrates (Nepi et al., 2001) (the ratio of sucrose: glucose: fructose) in different species found by the intensive studies of nectar carbohydrate components. Therefore, it is a reasonable to suspect that the nectar carbohydrates are not only derived from sugars in flux, but also from the carbohydrate storage materials in nectary cell, to regulate the ratio of nectar carbohydrates, and to attract different types of pollinators. Furthermore, the differences of amyloplast nectary and chloroplast nectary (Daivs et al., 1986; Figueiredo and Pais, 1992; Nepi et al., 2001) suggest that the less starch deposited in nectary, the less nectar is produced. In addition, the availability of starch in different nectaries suggests that starch degradation product regulates the timing of nectar secretion.

Hypothesis 2: Starch in nectary cells not only provides the nectar carbohydrates, but also supply the precursors of other molecules accumulated in nectary for further uses. In both ways, the removal of osmotic pressure introduced by starch degradation in a short time period is involved. Starch accumulation is a massive energy consuming process (Nepi et al., 2001; Southwick, 1984). Why do plants adapt such a huge energy consuming 6

process to produce nectar carbohydrate just to be secreted outside of plants? One feasible answer is starch can be used to regulate the timing of nectar secretion, although there is no experimental evidence to support this speculation. Another possibility is that the nectar will be reabsorbed after pollination to support ovary maturation and further seeds development (Nicolson, 1995; Nepi et al., 2001; Nepi et al., 2003). The reabsorption is observed in plant species (Nepi et al., 2001). Another likely possibility is that the product of starch degradation can be converted to another kind of molecule which can lead to plant defense (Horner et al., 2003; Horner et al., 2007; Carter and Thornburg, 2004). Calcium oxalate in soybean and carotene in tobacco are the chemicals accumulated in the floral nectary after starch degradation (Horner et al., 2003). Although there is no direct evidence to link the starch metabolism with carotene or calcium oxalate metabolism. One observation of nectary cell morphological changes is that mitochondria number increases in the late stages (Horner et al., 2003) suggests the glucose from starch degradation will enter glycolysis and enter TCA cycle to be further converted into the calcium oxalate or carotene deposited in late nectary cells. The molecular conversion process could be stipulated to remove the osmotic pressure introduced by high concentration of endogenous glucose from starch breakdown.

1.2 Plant floral nectary and nectar Nectarferious tissues in plants are the parts responsible for secreting nectar to the outside of nectaries. These tissues are categorized as two major classes, floral nectary and extrafloral nectary (Durkee et al., 1981; Faha, 1987; Vogel, 1987; Vogel, 1988a; Vogel, 1988b; Vogel, 1988c; Schmid, 1988). The extrafloral nectaries are found on outer floral parts or on vegetative parts, and stipulated as a source to attract predatory insects to protect against herbivores (Heads and Lawton, 1985). Extrafloral nectaries generally photosynthesize, and their activity may last from weeks to months. Extrafloral nectaries of angiosperms and nectaries of gymnosperms and ferns often lack direct vascular input but they tend to occur near 7

vascular bundles and frequently are associated with developing organs (Durkee et al., 1981; Nepi et al., 1996a). Floral nectaries are typically embedded in the floral tube, and serve as a source of attracting insect pollinators. Several major nectary structure components are found in floral nectaries: epidermal cells, parenchyma cells, and vascular bundles in some cases (Vogel, 1997; Vogel, 1998a; Vogel, 1998b; Vogel, 1998c). The epidermal cells are thought to be responsible for secretion of carbohydrate and amino acids that are contributing to nectar, while the parenchyma cells store polysaccharides. Nonetheless, floral nectaries of angiosperms have many different anatomy structures that vary from species to species, such as position in flowers, nectary structures, and types of plastids (Vogel, 1997; Vogel, 1998a; Vogel, 1998b; Vogel, 1998c). The structural variation makes the exploration of nectary function more difficult. Perhaps the most critical of the nectary functions that requires study is the origin of the carbohydrates that accumulates in nectar.

1.2.1 Plant nectar Floral nectaries are mainly thought to secret nectar within a flower to attract pollinators. Nectar contains sugars, proteins, amino acids, vitamins, lipids, organic and inorganic acids, glycosides, ions, aromatic compounds, and pigments (Davis et al., 1998; Galetto and Bernardello, 2004; Inouye, 1983; Pleasants and Chaplin, 1983; Wunnachit et al., 1992). A protein component with antimicrobial and antifungal properties was recently recognized in nectar of tobacco (Thornburg et al., 2003). The major sugar components of floral nectar are composed of sucrose and its two monosaccharide components, glucose and fructose (Davis et al., 1998; Pacini et al., 2003; Vogel, 1998a; Vogel, 1998b; Vogel, 1998c). Although the components nectar carbohydrates from all of the species are very similar, the nectar production, quantity, composition, and presentation of nectar vary widely from species to species (Pacini et al., 2003). This variability is thought to respond to environmental changes (Fahn, 1979; Cruden et al., 1983; Marden, 1984; Freeman and Head, 1990; Wyatt et al., 1992; Smets et al., 2000) and physiological 8

changes (Galetto and Bernardello, 1992; Gottsberger et al., 1990; Petanidou et al., 1996). The amount of dry mass or energy allocated to nectar production also varies considerably, from 3% for Pontederia cordata (Harder and Barrett, 1992) to 20% for alfalfa (Medicago sativa) (Teuber et al., 1980) and 35% for milkweed (Asclepias syriaca) (Southwick, 1983). The production of nectar often peaks when pollen is most available, as is the case for Cucurbita pepo (Nepi and Pacini, 1993) and tobacco (chapter 2), while in another case for Stenocereus stellatus, nectar secretion is maximal when the stigma is most receptive (Ibarra-Cerdena et al., 2005). Plants that flower at length or during seasonal transitions undergo large variations in environmental parameters that may affect nectar production and composition as well as nectary cytophysiological characteristics. In floral nectaries, the ratio of disaccharide/monosaccharide varies to attract different types of pollinators (Nepi et al., 1996a; Pacini et al., 2003). In general, honeybee and other insects can be attracted by highly concentrated sucrose-dominated nectar, while hummingbird can be attracted by more diluted monosaccharide-dominated nectar. Since nectar production rate and nectar composition are related to the types of insect pollinators (Baker and Baker 1983; Pellmyr, 1992), changes in these parameters during long flowering periods may be associated with changes in the spectrum of pollinators (Faegri and van der Pijl, 1979).

1.2.2 Nectar secretion models Many factors can affect the manner of nectar release to outside of the nectaries. Three major physiological factors are suggested: water potency, osmotic pressure and modified nectary cell structures (Pacini et al., 2003; de la Barrera and Noble, 2004). The nectar release is though to be the consequence of cooperation of those three major factors. The initiation of nectar secretion in most of the angiosperms is at same time as flower opening, a phenomenon that requires large amount of water to support and facilitate flower opening. The removal of osmotic pressures introduced by highly accumulated sugar contents in nectary parenchyma cells is required to decrease the cytosolic pressure. 9

Nectar is secreted from the floral nectaries, which are specialized superficial glands found in a few ferns (Rashbrook et al., 1992), a few gymnosperms (Wetschig and Depisch, 1999), and most angiosperms (Fahn, 1979; Friis and Endress, 1990). The floral nectaries of angiosperms can be found in many locations within flowers, but most frequently they are located near the inside of flowers (Fahn, 1952; Fahn, 1979). Nectaries have one or multiple layers of specialized nectar-secreting parenchyma cells that underlie an epidermis (Nepi et al., 2003; Horner et al., 2003). The epidermis often has modified stomata that lack subsidiary cells and become permanently opened as the nectary matures (Carter et al., 2007; Horner et al., 2007) or it can have nectar-secreting trachoma (Horner et al., 2003; Kronestedt et al., 1986). Floral nectar is generally rich in carbohydrates (Percival, 1961; Witt et al., 1999). These substances serve as an important energy source for the visiting pollinators (Fahn, 2000; Pacini et al., 2003; Southuick, 1984; Pyke, 1991). Several sources for these substances have been suggested in the literature (Fahn, 2000; de la Barrera and Noble, 2004). However there has been no conclusive biochemical study to demonstrate the source of these substances. Nonetheless, the mechanisms that result in release of nectar from the nectary gland have been investigated. At least five mechanisms have been proposed that mediate release of nectar from the nectary. These mechanisms include modified stomata (Davis and Gunning, 1992; Nepi et al., 1996a; Nepi et al., 2003), cuticle disruption (Wunnachit et al., 1992), cell exocytosis (Fahn, 1979; Kronestedt-robards, 1991), modified external periphery model (Figuereido and Pais, 1992), and programmed cell death (Vesprini et al., 1999; Horner et al., 2003). In the modified stomata model (Davis and Gunning, 1992; Nepi et al., 1996a; Nepi et al., 2003), there are stomata that are expressed on the surface of the nectary. These stomata have lost the capacity to open and close in response to normal physiological stimuli, such as ABA (Carter et al., 2006; Carter et al., 2007). Instead, these stomata are thought to be regulated developmentally during nectary maturation. In this model, nectar secretion from the nectary special parenchyma cells causes a hydrostatic pressure that results in nectar exudation 10

through the opening of modified stomata. Nectaries of this type occur in Vicia (Davis and Gunning 1992), Cucurbita (Nepi et al., 1996a), Linaria (Nepi et al., 2003) and tobacco (Carter et al., 2007). This is a common nectar secretion pattern in angiosperms, and is perhaps the most commonly modified nectary structure, enabling all parenchyma and epidermal cells produce and secrete nectar at the same time. In cuticle disruption, cell exocytosis and modified external periphery models, parenchyma cells operate simultaneously and secretory epidermal cells are synchronous in the releasing of nectar. In the cuticle disruption model, the nectar accumulates in a pocket beneath the cuticle. As nectar secretion increases, the integrity of the cuticle breaks down under the increasing pressure. Species known to secret nectar via cuticle disruption include Anacardium occidentale (Wunnachit et al. 1992). In the cell exocytosis model, the plasma membrane produces invaginations that appear to be similar to those of transfer cells. The increased membrane surface areas facilitate and speed secretion. This model is known to occur in Lonicera japonica (Fahn, 1979). In the modified external periphery model, nectar flows out through the external periphery of the epidermis which has a cuticle, as in Limodorum abortivum spur (Figuereido and Pais, 1992). In the programmed cell death model, the nectary cells gradually die, producing nectar for a given period as in Helleborus (Vesprini et al., 1999). The programmed cell death model is rare, in which the secretory epidermal cells are asynchronous.

1.2.3 Starch in floral nectaries Angiosperm nectaries have been divided based on plastid type (Nepi, 1996a; Pacini et al., 2003). In one type, photosynthetically competent chloroplasts are found throughout the nectary (Vogel, 1997; Vogel, 1998a; Vogel, 1998b; Vogel, 1998c). In these nectaries, nectar sugars produced by photosynthesis make up the majority of nectar sugars. Nectaries of this type often produce a relatively modest amount of nectar but can continue to produce nectar for long periods of time, often several weeks. In the second type of nectary, plastids develop as amyloplasts or as amylochromoplasts (Horner et al., 2007). In nectaries of this type, starch 11

accumulates to high levels in the nectary parenchyma cells during floral development. Then prior to anthesis, the starch in parenchyma is rapidly broken down to provide sugar for nectar. Advantages of this type of mechanism include secreting a large amount of nectar over a short period time (Durkee et al., 1981; Nepi et al., 1996a; Pacini et al., 2003). Amyloplast type nectaries have been observed in Cucurbita (Nepi et al., 1996a; Nepi et al., 1996b), Passiflora sp. (Durkee et al., 1981), Hibiscus rosa-sinensis (Sawidis, 1998), Rosmarinus officinalis (Zer and Fahn 1992), Pisum sativum (Razem and Davis 1999) and the orchid Limodorum abortivum (Figueiredo and Pais 1992). Amyloplasts in the nectar-producing parenchyma are generally almost devoid of stroma because they are fully packed with starch (Nepi et al., 1996b). Because the loading of starch into the starch-storing parenchyma consumes energy (Nepi et al., 2001), the starch storage and degradation that precede nectar secretion are probably evolutionarily derived steps that allow better control for the timing of nectar secretion. Moreover, there are many starch granules per amyloplast; this increases starch surface area, facilitating and speeding hydrolysis at the time of nectar production and secretion. Nectar production may vary in quantity. When it is very abundant amyloplasts may even accumulate in epidermal cells (Nepi et al., 1996b). The abundance of starch in parenchyma cells of Cucurbita pepo and Passiflora biflora (Nepi et al., 1996b; Durkee, 1982; Durkee et al., 1981) has been linked to the high rate of production of nectar with a high sugar concentration. In both types of nectaries, the amount and activity of invertases determine the relative nectar concentration of sucrose versus its hexose components, fructose and glucose (Nepi et al., 2001; Pacini et al., 2003).

1.3 Starch metabolism in higher plants In higher plants carbohydrates are frequently stored in the form of polysaccharides. These polymers reduce osmotic pressure in cells by reducing the concentration of soluble glucose and serve as a repository for these glucose units for future use. Several different types of polysaccharides (glycogen, phytoglycogen, 12

and starch) are known from in various life forms (Preiss, 1982; Landschutz, 2004; Zeeman et al., 2004). All of the storage polysaccharides are composed of glucose monomers linked by α-1,4-glycosidic bonds to form larger linear chain structures, as well as α-1,6-glycosidic linkages to form branches. In contrast, beta-linked glucans such as cellulose and beta-1,3-glucan serve a structural role in plant cell. Among the storage glucans, glycogen is the main storage polysaccharides in animals, while starch is the main storage material in higher plants. Phytoglycogen is a water- soluble polyglucan. It has a similar degree of branching that is to glycogen, but it is significantly smaller. Starch is a storage polysaccharide accumulating as a complex granular structure and is usually defined as a mixture of two distinct components: amylopectin and amylose (Ball et al., 1996; Myers et al., 2000; Smith et al., 1997; Morell et al., 2005). Amylopectin is the major component that is found in the starch granule. It has a very high molecular weight (107~109Da) with about 5% of α-1,6- glycosidic bond at the branching point. While amylose is a smaller linear molecule (105~106Da) with less than 1% of α-1, 6-glycosidic bonds. Starch is synthesized in plant plastids. In leaves, transient forms of starch are synthesized in chloroplasts (Smith et al., 2003). The turnover of transient starch is generally coordinated with circadian cycles. The transient starch synthesized in photosynthetic (source) tissues during the day is broken down at night to provide the carbohydrate metabolites to require for plant growth. In addition to the transient forms, some plant hetertropical (sink) organs also synthesize a form of starch that is termed storage starch (Emes and Neuhaus, 2000). These isoforms differ little in structure from the transient forms but rather accumulate to high levels in the sink tissues where they from a reserve source of metabolites and energy. Although in both source and sink tissues, starch is used for storage and provision of carbohydrate metabolites, starch metabolism in different tissues varies. In photosynthetic tissues, starch is synthesized to capture glucose made during the day, and then degraded to supply other biological processes at night (Smith et al., 2003). In some sink tissues, such as potato tuber and maize kernel, starch accumulates without obvious degradation during development (Emes and Neuhaus, 13

2000). These storage forms are then utilized in other later processes that require energy. For example, the starch formed in maize kernels and in potato tubers is utilized to provide energy for young shoots during the germinations process. In an unusual case of starch storage in a particular sink tissue, floral nectary, starch is accumulated during early stages of nectary development over a 7 to 8 day timeframe (Ren et al. submitted). This storage starch is degraded just prior to anthesis to provide glucose for a variety of processes.

1.3.1 Starch biosynthesis Starch metabolism in higher plants occurs via specific biochemical pathways of starch synthesis and degradation that utilize similar enzymes in a core pathway. The enzymes responsible for the starch synthesis and degradation pathways are highly conserved in all of plant species. Starch synthesis requires the cooperation of at least four different enzymes (Figure 1.2, 1.3, and 1.4). These include ADP- glucose pyrophosphorylase (AGPase, EC2.7.7.27) (Baroja-Fernadez et al., 2003, Chen and Janes, 1997; Geigenberger et al., 2005; Jin et al., 2005; Sanwal et al., 1968), starch synthase (SS, E 2.4.1.21) (Delvalle et al., 2005; Edwards et al., 1999; Hirose and Terao, 2004; Tenorio et al., 2003; Zhang et al., 2005), starch branching enzymes (SBE, EC 2.4.1.18) (Devillers et al., 2003; Myers et al., 2000), and starch debranching enzymes (DBE, EC3.2.1.41, and EC 3.2.1.68) (Dinges et al., 2001; Dinges et al., 2003; Hussain et al., 2003; Wu et al., 2002). Each group contains multiple isoforms. These isoforms are often expressed in different tissues, and they can partially compensate for each other when one of the isoforms is missing (Hussain et al., 2003; Myers et al., 2000; Smith et al., 2005; Zhang et al., 2005). The process of starch synthesis is initiated by the enzyme ADP-glucose pyrophosphorylase (AGPase), and regulation of AGPase activity is critical to starch synthesis (Chen and Janes, 1997; Crevillen et al., 2005). ADP-glucose (ADP-Glc) pyrophosphorylase is the first committed step of starch synthesis. It is responsible for producing ADP-Glc, the soluble substrate for starch synthases. The holoenzyme of AGPase in higher plants is a hetero-tetramer with two large subunits and two 14

Figure 1.4 Biochemical reaction of starch metabolic enzymes

To be continued 15

Figure 1.3 The core pathway of starch biodegradation in higher plants 16

Figure 1.2 The core pathway of starch biosynthesis in higher plants 17

Figure 1.4 Biochemical reaction of starch metabolic enzymes (continued) 18

small subunits. In Arabidopsis, there are four large subunit genes (AGPL1, AGPL2, AGPL3, and AGPL4) and two small subunit genes (AGPS1, and AGPS2) (Crevillen et al., 2005). The large subunits function as regulators of the enzyme activity, while the small subunits function as catalytic subunits (Crevillen et al., 2005). Although AGPase activity is actually regulated by the AGPLs, the activity is also regulated by the tissue redox potential (Geigenberger et al., 2005). The redox potential fluctuates with the diurnal cycles in photosynthetic tissues. In this process, light and sugar signaling activate thrioredoxin to break the disulphide bonds that exist between two catalytic AGPS subunits. Breaking this disulfide bond increases the AGPase holoenzyme activity by increasing substrate affinity and by increasing sensitivity to the activator 3’-phosphoglycerate (3PGA). Once the activated substrate ADP-Glc is synthesized by the action of AGPase, the glucose moiety is transferred onto a glucan primer by the action of one of several different starch synthases (Edwards et al., 1999; Myers et al., 2000; Smith et al., 2005). In starch polyglucan chain elongation, multiple starch synthases (SS) cooperate to form a linear α-1, 4 glycosidic linked starch structures. These enzymes synthesize polysaccharides by transferring the glycosyl group of ADP-Glc to the non-reducing end of a pre-existing glucan primer. Multiple starch synthase isoforms have been identified in plants (Delvalle et al., 2005; Edwards et al., 1999; Hirose and Terao, 2004; Li et al., 2000; Tenorio et al., 2003; Zhang et al., 2005). These isoforms can be categorized into two major groups: the granule-bound starch synthases (GBSS) and the soluble starch synthases (Myers et al., 2000; Smith et al., 2005; Tetlow et al., 2004; van de Wal et al., 1998; Vriten and Nakamura, 2000). GBSS functions in the production of the linear molecular amylose and the soluble starch synthases function in the synthesis of amylopectin. Two different isoforms of GBSS (GBSS1 and GBSS2) have been found in potato. The GBSS 1 is localized in the amyloplasts granule matrix of storage tissues and functions specifically to elongate amylose (van de Wal et al., 1998). GBSS2 is thought to be responsible for transient amylose synthesis in leaf plastids and the plastids of other non-storage tissues (Vriten and Nakamura, 2000). 19

At least four different isoforms of soluble starch synthases have been found in Arabidopsis and maize. These have been named SS1, SS2, SS3, and SS4. The enzymes of the soluble starch synthase group are localized between stroma and starch granules in plastids. These enzymes are involved in amylopectin synthesis. However, the different isoforms appear to have different functions. SS1 is thought to be mainly responsible for the synthesis of the shortest chains (<10 DP, degree of polymerization) (Myers et al., 2000; Smith et al., 2005). SS2 catalyzes the synthesis of intermediate-length glucan chain (10-25 DP) (Craig et al., 1998, Myers et al., 2000). In contrast, the SS3 isozyme is responsible for formation of the longest polyglucan chains (>30 DP) extension (Delvalle et al., 2005; Edwards et al., 1999; Hirose and Terao, 2004; Morell et al., 2003; Zhang et al., 2005, Tenorio et al., 2003). The structure of amylopectin is significantly more complex than that of amylose (Myers et al., 2000; Smith et al., 2005). Amylopectin differs from amylose by the presence of branches in the glucan structure. These branches are introduced into amylopectin by the action of two additional classes of enzymes, starch branching enzymes and starch debranching enzymes. Starch branching enzyme (SBE) introduces an α-1,6 glycosidic bond into the amylopectin structure by cleaving an internal α-1,4 glycosidic bond in one glucan chain and transferring the released reduced end to C6 hydroxyl group of another glucose unit (Devillers et al., 2003; Myers et al., 2000; Tetlow et al., 2004). Two major isoforms of starch branching enzymes, SBE1 (SBE b) and SBE2 (SBE a) are found in maize, rice and Arabidopsis (Tetlow et al., 2004). SBE2 transfers shorter glucan chains and displays a higher affinity towards amylopectin than SBE1, which has higher affinity towards amylose (Myers et al., 2000; Tetlow et al., 2004). The removal of branches within the amylopectin molecule is also important in the formation of the amylopectin structure. This restructuring of the branch points is catalyzed by the action of debranching enzymes (DBE). This activity reorganizes the semicrystalization structure of starch, thereby permitting increased packing of the glucan units. DBE have a mechanism that is essentially the reverse of the SBE mechanism. These enzymes cleave an α-1, 6-linkage in a branched polyglucan and 20

add that removed branch to the nonreducing end of the trimmed polyglucan branch. Debranching enzymes in higher plants can be categorized into two classes: isoamylases (ISA) (Dinges et al., 2001; Hussain et al., 2003; Wu et al., 2002), and pullulanase (PU, also known as limit-dextrinases) (Dinges et al., 2003). Three isoamylases (ISA-1, ISA-2, and ISA-3) and one pullulanase (PU1) genes have been identified in Arabidopsis genome. It is well accepted that debranching enzymes play roles in both of starch synthesis and degradation (Myers et al., 2000). However, the exact roles of debranching enzymes in starch biogenesis have still not been completely elucidated. Two models for the function of debranching enzymes in starch synthesis: glucan-trimming model and glucan-cleaning model proposed (Myers et al., 2000). In the glucan-trimming model, debranching enzymes trim inappropriately positioned glucan branches generated at the growing surface of a starch granule to prevent phytoglycogen accumulation, thereby encouraging starch crystallization and more efficient glucan packing. In the glucan-cleaning model, debranching enzymes are responsible for removing any soluble substrates of amylopectin synthesizing enzymes, such as starch synthases and starch branching enzymes, thereby preventing random futile synthesis of phytoglycogen.

1.3.2 Starch degradation Starch degradation is also the result of multiple enzymes acting in concert. α- Amylase (AMY, EC 3.2.1.1), and β-amylase (BMY, EC 3.2.1.2) are the major enzymes responsible for starch degradation (Smith et al., 2005; Zeeman et al., 2004; Tetlow et al., 2004). However, recent work also shows that glucan water dikinase (GWD or R1, EC 2.7.9.4) and starch phosphorylase (PHO, EC 2.4.1.1) are also involved in starch degradation (Blennow et al., 2002; Ritte et al., 2002; Smith et al., 2005). α-Amylase, β-amylase, disproportionating enzyme (DPE) initiate the process of starch degradation in plants. α-Amylase is an endoamylase, which cuts the α-1,4 linkage inside of polyglycan chain to produce smaller pieces with two or three 21

glucose units. β-Amylase is an exoamylase that hydrolyzes α-1,4 glycosidic linkages of polyglycan chains at the nonreducing end to produce maltose (4-O-α-D- glucopyranosyl-β-d-glucose) during starch breakdown. The DPE breaks the inside α- 1,4 linkages and transfer the resulting glucan moiety to an acceptor molecular through creation of a new α-1,4 linkage (Critchley et al., 2001). The temporal and spatial pattern of starch degradation in plants varies. Most studies about starch degradation regulation have been conduced on one of several biological systems, including barley seed germination, and in Arabidopsis leaves to understand the diurnal fluctuations of starch in leaves (Preiss, 1982; Smith et al., 2005). However, there are very few studies on starch degradation pathways in nonphotosynthetic tissues so far, partially due to little detectable starch degradation in most storage organs (Emes and Neuhaus, 2000). Starch degradation initiates on the surface of intact starch granules (Smith et al., 2005). It has been thought that α-amylase is the key enzyme that initiates the process. However, recent progress has shown that the initiation of starch degradation is a correlation of different enzymes, including AMY3, BAM, DPE, and even ISA3 (Smith et al., 2005). Other enzymes, such as PHO (Blennow et al., 2002), GWD (Ritte et al., 2002) and phosphorylate water diknase (PWD), are also thought to be involved in starch degradation (Smith et al., 2005).

1.4 Outline of the dissertation Taking the advantages of the large amount nectar secretion of ornamental tobacco and the known starch metabolic pathway in other higher plants, we have intensively investigated the function of starch metabolism in the developing ornamental tobacco floral nectaries to understand the floral nectar secretion mechanism. We first have identified the origin of nectar carbohydrate in the nectaries by studying the physiology changes of starch (Chapter 2). The nectar carbohydrate appears to be derived from starch breakdown products prior to anthesis, and is from transported photosynthate after anthesis. The latter process appears to be regulated by the starch breakdown products prior to anthesis. We 22

further investigated the starch metabolic gene expression profile to understand the regulation of starch metabolism in the developing tobacco floral nectary (Chapter 3). Three classes of starch metabolic genes in the nectary were identified: catabolic genes (i.e., starch synthases and AGPase), anabolic genes (i.e., α-amylase, β- amylase, and isoamylase 1), and constitutive expressed genes (i.e., starch branching enzymes, starch phosphorylase, and R1 proteins). Besides the investigation of starch metabolic gene expression pattern, we also have investigated the biomass conversion from starch to β-carotene biosynthesis at later stages of floral nectaries (Chapter 4). This biomass conversion appears to be that the starch breakdown products provide large amount of substrates for β-carotene biosynthesis. That process appears to remove the high osmolarity introduced by the starch breakdown products in the late stages of nectaries. And finally, we have generated a series of transgenic plants using RNAi and/or overexpress approaches to further the investigation of the regulation of nectar secretion (Chapter 5). One set of the transgenic plants included in the investigation is designed to further explore the function of the starch biosynthesis genes in the developing ornamental tobacco floral nectary. These starch biosynthesis genes include starch synthase 1, starch synthase 2, starch synthase 3, AGPase. Another set of the transgenic plants is designed to further explore the function of beta-carotene biosynthesis genes, including ipi, ispH and mdc. Further analyses of those genetic modified plants may provide the genetics evidence for the regulatory role of starch in developing ornamental tobacco floral nectaries, and further pave the way to explore the insect- mediated pollination strategy.

Chapter 2 Starch accumulation in ornamental tobacco floral nectary development: Physiological evidences of starch regulating floral nectar secretion

This chapter is based upon a manuscript that has been submitted to “the Plant Science”

Authors: Gang Ren1, Rosanne A. Healy2, Anna Klyne1, Harry T. Horner2, Martha G. James1, & Robert W. Thornburg1*

Address: 1Department of Biochemistry, Biophysics and Molecular Biology, Iowa State University, Ames, Iowa 50011-3260 USA 2Department of Genetics, Developmental and Cell Biology & Bessey Microscopy Facility, Iowa State University, Ames, Iowa 50011-1020 USA

Keywords: nectar, nectary, Nicotiana, starch, nectar secretion

Contribution of this work:

Unless otherwise noted, the experiments in this manuscript were performed by Mr. Gang Ren. In addition, Mr. Ren was responsible for carrying this project forward throughout the course of the project. Ms. Rosanne Healy performed the Microscopy for Figure 2.1, 2.3, and 2.9. Ms. Anna Klyne performed preliminary experiments on labeling of nectaries with [14C]-sugar that were not included in the final version of this work, and she did the TLC analysis of floral nectar following labels for Figure 2.7. Dr. Harry Horner consulted on the methods of Microscopic sections of this manuscript. Dr. Martha James consulted on starch quantitation and FACE analysis in the characterization of the nectary starch. Dr. Robert Thornburg consulted on the overall creative instructions and support for this work. In addition, he contributed to the writing, reviewing and editing of this work. This work was supported by an NSF grant (IBN-0235645) to Dr. Robert Thornburg and Dr. Martha James.

Abstract Enlargement of the floral nectary gland of ornamental tobacco that occurs during development is accompanied by a major accumulation of Periodic Acid-Schiff’s staining (PAS-staining) starch grains in nectary amyloplasts. Quantification of starch purified from the nectary at various stages of development showed little starch at an early developmental stage, soon thereafter the amount of starch increased dramatically, reaching a peak approximately 24 hours prior to anthesis. After this time, the amount of starch declined dramatically until the flower reached anthesis, suggesting that the accumulated starch was converted to sugars (sucrose, glucose, and fructose) for nectar production that occurs prior to anthesis. Compositional and structural analyses of nectary starch showed that amylose content and degree of amylopectin branching varied during nectary development. The average chain length in amylopectin was relatively short at early developmental stages, reached maximal length towards the end of nectary development, and was reduced again at anthesis, consistent with the synthesis of an increasingly complex form of starch up to middle stages of nectary development, followed by decreases in starch complexity and amount due to starch degradation in the mature nectary. We also evaluated the transport of radiolabeled sugars nectar. Consistency in the amounts and ratios of radiolabeled sugars in nectar indicates that the transported photosynthate sugars were subject to metabolic constraints in the nectary tissue prior to their incorporation into nectar. The velocity of radiolabeled sugars accumulated in nectar illustrates that transported photosynthate contributes to the later nectar production. 26

2.1 Introduction Co-evolution of flowering plants and insects is aptly demonstrated by the fundamentally conserved process of pollinator-mediated hybridization that occurs in plants in spite of the widespread diversity of these organisms. The flower is the basic plant structure that embodies the organs and processes necessary for hybridization. Many floral features are involved in this process. However, rich floral nectar that serves both as an attractant and as a reward for visiting pollinators is arguably the chief component that drives this process. Our research efforts are focusing on the developmental and molecular aspects of nectar production to create strategies that will enhance insect-mediated plant hybridization. We contend that as world populations rapidly approach 10 billion, such enhanced hybridization methods will be required to produce new, cultivated plant lines to feed these burgeoning populations. In most angiosperms, nectar production involves a distinct floral gland called the nectary that occurs in various locations within the flower but most commonly in a cup-like environment at the base of the gynoecium or ovary. The secretion of floral nectar in most angiosperms seems to be under very specific developmental control. Secretion of nectar begins just prior to anthesis and continues while the flower remains receptive to pollination. Nectar flow is often increased by pollinator visitation. However, after pollination, nectar secretion ceases and the nectar remaining in the flower may be reabsorbed (Burquez and Corbet, 1991). Nectar is composed largely of sugars (Baker and Baker, 1981), but also contains amino acids (Baker and Baker, 1973), Krebs cycle intermediates (Baker and Baker, 1975), lipids (Vogel, 1969), vitamins (Griebel and Hess, 1940), and proteins (Peumans et al., 1997, Carter and Thornburg, 2004a, Naqvi et al, 2005). The developmental and molecular mechanisms of floral development that give rise to the nectary secreting tissues and thence to the floral nectar are for the most part unknown. In this study we have focused on starch, a major metabolite, which accumulates in the nectary during floral development, and becomes a source of carbohydrate in the nectar. In 27

addition, we have found that the carbohydrate found in nectar arises not only from starch degradation, but also from transported photosynthates.

2.2 Materials and methods 2.2.1 Materials Radiolabeled sugars (U-[14C]-sucrose and U-[14C]-glucose) were purchased from Amersham (Piscataway, NJ). All other materials were from Sigma Chemical Corp (St. Louis, MO) or Fisher Chemical Company (Pittsburgh, PA) and were of the highest purity available.

2.2.2 Plants The lxs8 line of ornamental tobacco plants used in this study was derived from an interspecific cross between Nicotiana langsdorffii and Nicotiana sanderae (Kornaga et al., 1997). Both of these parents are diploid and belong to the Alatae section of Nicotiana. These plants were previously used to study a genetic instability as well as the tobacco nectar proteins (Carter et al., 1999, Carter and Thornburg, 2000). Nectar was collected as previously described (Carter et al., 1999). Floral stages were determined according to Koltunow et al. (Koltunow et al., 1990).

2.2.3 Transport of radiolabeled sugars Flowers were collected in the greenhouse by cutting through the pedicel close with a razor blade. Cut inflorescences were immediately taken to the laboratory and specifically staged flowers were separated and maintained on ice. The cut end of each flower was immediately placed into 200 μL of feeding solution that was based the composition of on average phloem components, 17% sucrose, 0.01% glucose, and 0.01% fructose (De la Barrera and Nobel, 2004) supplemented with 2µCi of either [14C]-sucrose or [14C]-glucose in 1 mL solution. Six replicates of each identified floral stage were used for labeling. Nectar sample were collected by pipeting 1 mL water into the floral tube, mixing, and decanting the fluid to a fresh microfuge tube. These samples were centrifuged (17,000 x g) for 10 min to remove 28

any particulate material, then 300 uL of each collected sample was used to detect radioactivity by scintillation counting in a Packard 1900 TR liquid scintillation counter. The total carbohydrate in nectar was analyzed by the sensitive anthrone method (Morris, 1948).

2.2.4 Carbohydrate analysis Thin layer chromatography of radiolabeled sugars To characterize the composition of the radiolabeled products in nectar, 300 µL aliquots were analyzed by thin layer chromatography on Whatman PE SIL G silica gel plates in 90% acetonitrile. The solvent was allowed to ascend 3 x with complete drying of the plate between ascents. The plate was then stained with a solution of 5% p-anisaldehyde in ethanol:sulphuric acid (95:5). Sugars were visualized by heating the plate at 110ºC for 3 min and photographed. To quantify the levels of radioactivity, each plate was exposed to a PhosphorImager plate for 92 h then scanned on a Molecular Dynamics Typhoon 8600 PhosphorImager. The amount of radioactivity in each spot was quantified using ImageQuant software. Unmodified labeled sugars were also run on TLC plates as controls.

Nectar carbohydrate composition Nectar was collected from flowers as previously described (Carter et al., 1999) with one exception. To insure that we were collecting all of the nectar, the floral tubes, nectaries, and receptacles were washed with 1 mL of water and this wash was combined with the isolated nectar. The samples were used immediately for carbohydrate determination. To quantify free carbohydrates in nectaries, isolated nectaries were separated from the gynoecia, and approximately 10 to 15 nectaries were homogenized by grinding in a ConTorque tissue homogenizer (Eberbach Corp, Ann Arbor, MI) for 5 min, then the homogenate was brought to a final volume of 1 mL with water. Following centrifugation (17,000xg) for 10 min, the clarified homogenates were used immediately for carbohydrate analysis. The levels of sucrose, glucose, and fructose were evaluated using The Sucrose/D-Glucose/D- 29

Fructose Determination Kit (Boehringer Mannheim/r-Biopharm Cat. No. 10-716-260- 035), according to the manufacturer’s directions.

2.2.5 Starch analysis Isolation of starch from nectaries To isolate starch, we used a published method (Vasanthan, 2001). Nectaries were separated from 50 gynoecia at each flower stage as previously described (Carter and Thornburg, 2000), then the nectaries were frozen in liquid nitrogen and homogenized in a mortar and pestle. To isolate starch, 5 mL of 80% ethanol was added to each nectary homogenate, then incubated for 5 min in 80ºC water bath, followed by centrifugation for 10 min at 1,000 x g, room temperature and the ethanol was gently decanted so as to not disturb the pellet. The remaining pellet was dissolved in 500 μL of 90% aqueous DMSO and heated for 5 min in a boiling water bath. Subsequently the solubilized samples were diluted into 10 volumes of water for enzymatic quantitation. Three independent isolations of starch from nectaries were performed for each stage analysis. All samples were processed at the same time. Starch quantitation Starch was quantitified using the Starch Determination Kit from Boehringer Mannheim/r-Biopharm (Cat. No. 10-207-748-035) according to the manufacturer’s directions, but with one modification. To more completely digest the starch, 1 unit of amyloglucosidase per reaction (Sigma, Cat. No.10130) was incubated with the starch for 17 h at 65ºC (Zhang et al., 2005) prior to enzymatic analysis of hydrolyzed glucose. Starch structural analysis The distribution of linear chains in nectary starch was determined by fluorophore assisted carbohydrate electrophoresis (FACE) according to previously published procedures (O’Shea et al., 1998) with slight modifications (Dinges et al., 2003). Depending on the developmental stage, 300 to 1,500 flowers were required to obtain enough starch for analysis. Nectaries were carefully dissected as 30

described (Carter and Thornburg, 2000), and were stored frozen at -70ºC until used. Because of the large number of nectaries required, the nectaries were separated into smaller groups of 20 to 50 nectaries per purification. As described above, the nectaries were homogenized, DMSO was added to 90%, and the nectaries were boiled for 1 hr with periodic shaking. Following a 15-min centrifugation (17,000 x g), the supernatant was transferred into a new 1.5 mL tube, and 1 mL of 100% ethanol was added to each tube. After mixing, the supernatants were incubated at room temperature overnight and centrifuged at 5000 x g for 5 min to precipitate the extracted starch. The pellets were washed with 80% ethanol and then dissolved in 50 μL 90% DMSO. The final starch concentration was about 5 μg/μL. For FACE analysis, the nectary starch samples were debranched using a commercial Pseudomonas amyloderamosa isoamylase (Megazyme, International, Bray, Ireland). The debranching reaction contained 200 μg starch (dissolved in 30% DMSO), 500 mM sodium acetate (pH 4.4), and 0.6 Units Isoamylase in 50 μL. After overnight incubation at 42ºC, the sample was heated in a boiling water bath for 5 min then centrifuged for 2 min to collect the supernatant. A 10 μL aliquot of the digested starch was dried in a SpeedVac. When dry, 2 μL of 0.1 mg/mL 8- aminopyrene-1, 3, 6-trisulfonic acid (APTS) in 15% acetic acid, and 2 μL of 1M sodium cyanoborohydride (NaCNBH4) were added to label the non-reducing ends of each linear glucose multimer. Following overnight labeling in the dark at 42ºC, 46 μL of Milli-Q water was added to each reaction. The samples were mixed by vortexing and after a subsequent 2 min centrifugation (17,000 x g), a 5 μL of sample was removed and mixed with 195 μL Milli-Q water in FACE vials. These samples were immediately run on a Beckman P/ACE capillary electrophoresis instrument as described (Dinges et al., 2003). Amylose content analysis To evaluate amylose content, a standard curve containing varying ratios of amylose:amylopectin was prepared from (0 to 100%) potato amylose (type III; Sigma Cat. No. A0512) and (100 to 0%) potato amylopectin (Sigma Cat. No. 10118). The total carbohydrate content for the standard curve was maintained constant at 1 31

μg/μL dissolved in 30% DMSO. Then, 100 µL of these amylose:amylopectin

solutions was mixed with 400 μL of I2/KI (0.67% [w/v] I2 and 3.3% [w/v] KI) solution. Starches with unknown amylose:amylopectin ratios at (1 μg/μL starch dissolved in 30% DMSO) were similarly prepared in triplicate. The absorbance at 540 nm for each of the starch solutions was recorded. The composition of unknown starch solutions was established by comparison with the amylose:amylopectin standard curve.

2.2.6 Light Microscopy Gynoecia were physically removed from flowers at floral Stages 2 to 12, and processed in two different ways. For light microscopy, the gynoecia were sliced above the nectaries, and the nectaries were immediately placed in vials containing a fixative consisting of 2% paraformaldehyde and 2% glutaraldehyde in a 0.1 M cacodylate buffer at pH 7.2 and 4ºC. The vials with nectaries were placed in a low vacuum (0.1 Torr) for 30 min on ice, the original fixative was replaced once, and the samples were then stored at 4ºC overnight before they were transferred through an ethanol dehydration series (10%, 25%, 50%, 70%, 95%; 1/2-h steps) which concluded with two changes of pure ethanol (1/2-h steps). The pure ethanol was replaced with increasing concentrations (3:1, 1:1, 1:3, ethanol:resin) of LRW resin, and two changes (several hours each) of pure resin before casting in gelatin capsules and polymerizing them at 60ºC for 1 d. One-μm-thick sections of nectaries at each floral stage were cut with a glass knife on a Reichert Ultracut S ultramicrotome and placed on deionized water droplets on slides and allowed to dry. Sections were either contrast enhanced with toluidine blue O or stained for water-insoluble polysaccharides (particularly starch) using the periodic acid-Schiff (PAS) technique (Horner et al., 2004). Treated and thoroughly-deionized water- washed slides were dried on a warming tray, and then xylol, Permount, and coverslips were added. Sections were viewed with a compound microscope in the bright-field mode. Images were digitized and captured using a Zeiss MRc digital 32

camera with associated software. Stored images were processed in Adobe PhotoShop, Adobe Illustrator, and Macromedia FreeHand.

2.2.7 Scanning Electron Microscopy Gynoecia with nectaries were isolated from tobacco flowers at Stage 12 (see previous section) and mature flowers of Arabidopsis thaliana were fixed in 2% paraformaldehyde and 2% glutaraldehyde in a 0.1 M cacodylate buffer at pH 7.2 and 4ºC. The vials with nectaries were placed in a low vacuum (0.1 Torr) for 30 min on ice, and then the original fixative was changed before they were stored at 4ºC overnight. The samples were washed 3x (20 min per step) with deionized water, followed by a 1 hr exposure to 1% osmium tetroxide in the same buffer and temperature. The samples were briefly washed with deionized water, and carried through the same ethanol dehydration series to pure ethanol (2x). The nectaries and flowers were critical point dried using pure ethanol and carbon dioxide. Dried samples were stored in a desiccator or were adhered to aluminum stubs using double-stick tape and silver paint. The samples were sputter coated with 120 nm AuPd, and then viewed in a JEOL 5800 SEM at 10 kV. Images were digitally captured, stored, and processed.

2.3 Results Studies of the biochemistry of the nectary have lagged behind almost all other plant organs, primarily because the nectary is typically a very small organ that is generally hidden among the internal appendages of the flower. It is difficult to isolate nectary tissue in quantity and because of these factors. Thus, it does not lend itself well to biochemical and molecular studies. This is particularly true in model systems such as Arabidopsis and other species, where light microscopy or even scanning electron microscopy are required to visualize them. Instead of using the nectaries of these model systems, we have chosen to work with a diploid ornamental tobacco line (Kornaga, 1993, Kornaga et al., 1997). While ornamental tobacco is not as desirable as Arabidopsis as a genetic tool, we have chosen tobacco over 33

Arabidopsis as our system for the following reasons: First, Arabidopsis has multiple relatively very small, nectaries (Figure 2.1A-D) of several types (Davis et al., 1998). Tobacco has a single, large nectary embedded in the base of the floral gynoecium and has two zones of stomates 180º opposite each other (Figure 2.1E-H), that are the source of nectar secretion (Carter et al, in press). We calculate that the tobacco nectary is about 1,000-fold larger than the nectaries of Arabidopsis. Second, a single Arabidopsis bolt typically produces fewer than 20 flowers at a time. Ornamental tobacco is indeterminant, continuously producing large numbers of flowers (up to 150) at any given time, permitting the isolation of 100s of mL of nectar and 10s of grams of nectary tissue at any stage for cloning and biochemical analyses. Although tobacco is generally considered to be a self-pollinated species, a significant increase in seed set occurs when insect pollinators visit (McMurtrey et al., 1960). Thus, performing nectary/nectar studies in ornamental tobacco, rather than in Arabidopsis, permits greater latitude in the types of experiments that can be performed and has allowed a much greater understanding of the biochemistry of nectar than can be obtained from Arabidopsis alone (Carter et al., 1999, Carter and Thornburg, 2000, Thornburg et al., 2003, Carter and Thornburg, 2004a, Carter and Thornburg, 2004b, Carter and Thornburg, 2004c, Naqvi et al., 2005, Horner et al., in press, Carter et al, in press). The composition of floral nectar in ornamental tobacco is presented in Table 2.1. The total carbohydrate load is 35%. Approximately half of the sugar present in nectar is sucrose, and the other half is almost evenly divided between glucose and fructose. Quantitation of other ornamental tobacco nectar components have been previously published: protein (Carter et al., 1999); ascorbate (Carter and Thornburg, 2004a); and H2O2 (Carter and Thornburg, 2000). The source of the carbohydrate in ornamental tobacco nectar is not known. Typically two sources of sugars in nectar have been suggested, phloem (Esau, 1997) and nectary-stored starch (Fahn, 1979; Nepi et al., 1996, Peng et al., 2004). However, light and electron microscopic studies of developing nectaries reveal that neither phloem nor xylem invade ornamental tobacco nectaries (Horner, et al., in press). Therefore, we began a study 34

Figure 2.1 Light and scanning electron microscopy (SEM) graphs of Arabidopsis thaliana and ornamental tobacco floral nectaries.

Panel A. SEM of Arabidopsis dissected flower showing two types of nectaries (arrows) sandwiched between petals (removed) and stamens. Bar = 100 µm; Panel B. SEM enlargement of medial nectary (right arrow of A.). Bar = 25 µm; Panel C. SEM enlargement of lateral nectary (left arrow of A.). Bar = 25 µm; Panel D. SEM enlarged portion of nectary in B., showing stomates with relatively small pores. Bar = 10 µm; Panel E. Freshly dissected tobacco gynoecium showing light green ovary and orange basal nectary at Stage 12. The carpel suture line is visible above. The boxed region shows one of the two nectary, stomatal zones. Dark region in box represents depressed region in F. Bar = 1 mm; Panel F. SEM of enlarged region shown in box of E. that represents one of two stomatal zones. Bar = 200 µm; Panel G. SEM of a single stomate from stomatal zone showing large open pore. Bar = 4 µm; Panel H. SEM of a second stomate from stomata zone in F. Bar = 4 µm.

35 36

Table 2.1 Composition of ornamental tobacco LxS8 nectar

Component Value Volume (µL) 24 [protein] (µg/mL) 240 [sugar] (brix) 35 Sucrose (M) 0.55 Glucose (M) 0.47 Fructose (M) 0.43 Ascorbate (µM) 920

H2O2 (mM) 4

to determine whether nectary-stored starch could serve as the source of sugar in the nectar of ornamental tobacco. In tobacco, the nectary consists of a single torus-shaped gland embedded in the base of the gynoecium. Near maturity, it is bright orange due to the accumulation of high levels of β-carotene (Horner et al, in press). Nectar is secreted only from opposing regions on the lateral nectary faces (Figure 2.1E). These regions have an abundance of stomata that all appear to be open in mature flowers (Thornburg et al., 2003, Carter et al., in press). Outside of these opposing lateral regions, stomata are absent. Stomates in nectary function appear to be a conserved mechanism for nectar exudation in many plant species. Indeed, stomates are also visible on Arabidopsis nectaries (Figures 2.1B-D), and are thought to be the source of nectar exudation in this species as well.

2.3.1 Nectary Development Flower development in tobacco is divided into 12 stages, from Stage 1, where the petals and sepals are of equal length to Stage 12 (anthesis) when the flower is fully open and the anthers have dehisced (Koltunow et al., 1990). Development of the nectary gland in ornamental tobacco also consists of several of discrete stages that correlate with the stages of flower development. A profile demonstrating the development of the nectary gland in relation to these floral stages is shown in Figure 2.2. Organ initiation of the nectary occurs at a primordial floral stage while the flower bud is still quite small. By Stage 1 (not shown) of flower development, the nectary is already a well-differentiated ring of cells within the wall of the primordial gynoecium. The initiation of the nectary gland may be controlled by the Crab’s Claw (CRC) transcription factor because Arabidopsis mutants in the CRC gene lack nectaries (Bowman and Smyth, 1999). However, how this transcription factor functions in the development of the nectary remains unknown. The first major phase of nectary development is collectively identified as the filling period during which the nectary characteristically increase in size of several-fold. This stage begins prior to 38

Figure 2.2 Events occurring during development of tobacco nectaries.

Three images at the top of figure show gynoecia (ovary plus nectary) from flowers at three stages of development: Stage 2, nectary from small, developing flowers; Stage 6, presecretory nectary; and Stage 12, fully mature nectary at anthesis.

39 40

Stage 2, continues through Stage 9, and lasts approximately 4 to 5 days. Nectary maturation occurs from Stage 9 through Stage 12. During the maturation period, nectar begins to be secreted from the nectary. Nectar proteins begin to be synthesized at late Stage 10, approximately one day prior to floral opening. The post-maturation period of nectary development begins at anthesis and continues as long as the flower is receptive to pollination. During this time, nectar flow continues although it decreases the longer the flower remains open. This period can last 6 to 8 days if the flower remains unpollinated.

2.3.2 Starch in nectaries To examine the presence of starch in nectaries, we analyzed nectary tissue from ornamental tobacco for the presence of starch using PAS-staining of 1-µm-thick LR White resin sections at five floral Stages (2, 4, 6, 9, and 12) of development. At each stage, nectary sections were first stained with toluidine blue O, a general stain that differentially enhances tissue contrast (Figures 2.3A, D, G, J and M). Figures 2.3B, C, E, F, H, I, K, L, N, and O show similar sections of flowers Stages 2 to 12 stained with PAS at both low magnifications (Figures. 2.3B, E, H, K and N), and higher magnifications to show the starch grains (Figures. 2.3C, F, I, L and O). From Stage 2 through Stage 9 there is an increase in starch in all nectary cells, and at Stage 12 most of the starch is depleted. From the high magnification images it is apparent that starch accumulates both in the outer epidermal layer as well as in the underlying special parenchyma cell layers that make up the majority of the nectary. There appear to be multiple starch grains per plastid at all stages observed. At Stage 12 the starch remaining in the nectary appears to be localized in the outermost cell layers of the nectary (Panel O). To confirm that these images were indeed indicating the metabolism of starch in nectaries, we isolated and quantified starch from similar nectary stages. As shown in Figure 2.4A, starch levels in the nectary increased rapidly during the middle of the filling period. Isolable starch peaked about Stage 9 and thereafter, the level of starch declined. In addition, we also measured the mass of the nectary during its 41

Figure 2.3 Light microscope images of 1-µm-thick LR White resin sections of tobacco nectaries at five different developmental stages (Stages 2, 4, 6, 9, 12) stained for differential contrast enhancement with toluidine blue O (A, D, G, J, M) or PAS technique for cell walls and starch (B, E, H, K, N and C, F, I, L, O).

All sections are longitudinal slices through nectary. B, E, H, K, N are low magnification sequential images by Stage (2 to 12) of nectaries stained with PAS, and C, F, I, L, O are their higher magnification counterparts. All bars = 50 µm.

42 43

Figure 2.4 Accumulation of starch in nectary.

Panel A. Nectaries from flowers at several stages of development were collected and starch was isolated from them. Black bars show quantified level of starch present in nectaries of the indicated stages. The open circles represent the total mass of the nectary at the indicated stages. Panel B. Carbohydrate composition of S9 nectaries.

800 8

600 6

400 4

200 2

0 0 2 4 6 8 9 10 11 12 Floral Stage

B WSG Fructan Fructose Glucose

Sucrose Starch 45

development. As can be seen in Figure 2.4A, the nectary increases in size from about 2 mg/nectary to greater than 6 mg/nectary at anthesis. By comparing isolable starch with total necary mass, we find that at Stage 9, starch represent more than 20% of the total nectary mass.

2.3.3 Starch is dynamically processed during nectary development To determine whether starch composition changed during nectary development, we analyzed starch composition and structure at different floral stages. Initially, we determined the amylose/amylopectin ratio of starch from both early and late stages. As shown in Figure 2.5, at Stage 2, the starch is about 35% amylose and 65% amylopectin. This ratio slowly changes during development to 22% amylose and 78% amylopectin by Stage 12, and remains unchanged 48 h after fertilization, although little total starch remain at this time. The amylose/ amylopectin content of potato starch is shown for comparison. We have also evaluated the structure of starch from several developmental stages using FACE (fluorescence-assisted carbohydrate electrophoresis). These analyses permitted us to identify structural changes that occur in the starch over a time course of synthesis and degradation. FACE analysis utilizes a commercial, bacterial starch debranching enzyme to hydrolyze the α-1,6 linkages in amylopectin. Following debranching, the reducing sugar of each hydrolyzed side chain was labeled with a fluorescent tag. These fluorescent-tagged linear carbohydrate chains were separated by capillary electrophoresis to obtain profiles representative of the relative abundance of specific lengths of linear starch chains at the analyzed stages. As shown in Figure 2.6 (Stages 2, 6, 9, 12; Panels A to D, respectively), FACE analysis for starch from each of the stages examined shows the relative distribution of linear chains extending from a degree of polymerization (DP) of ~4 to >70 glucose units in length. A careful examination of these profiles reveals that starch isolated from developing nectaries (Stage 2, Panel A and Stage 6, Panel B) is composed mostly of short chains with a mean DP of ~12. Note in Panel A the chain length is highly enriched in shorter lengths while the longest chains are present in 46

Figure 2.5 FACE analysis of nectary starch.

Panel A. Stage 2; Panel B. Stage 6; Panel C. Stage 9; Panel D. Stage 12 (anthesis); Panel E. 48 h post fertilization; Panel F. potato starch. control; Panel G. Stage 9 minus Stage 2; Panel H. Stage 9 minus Stage 6; Panel I. Stage 9 minus Stage 12; Panel J. Stage 9 minus 48h PF.

6 A 5 1.0 G 4 0 3 4 7 10 13 16 19 22 25 28 31 34 37 40 43 46 49 52 55 58 61 64 67 70 73

2 - 1. 0 1

0 4 7 1013 16 19 22 25 28 31 34 37 4043 46 4952 55 58 61 64 67 70 73 - 2. 0

5 - 3. 0 1.0 4 B H 0 4 7 10 13 16 19 22 25 28 31 34 37 40 43 46 49 52 55 58 61 64 67 70 73 3

- 1. 0 2

- 2. 0 1

0 - 3. 0 4 7 1013 16 19 22 25 28 31 34 37 4043 46 4952 55 58 61 64 67 70 73

4 C 1.0 I 3 0 4 7 10 13 16 19 22 25 28 31 34 37 40 43 46 49 52 55 58 61 64 67 70 73

2 - 1. 0

1 - 2. 0

0 4 7 1013 16 19 22 25 28 31 34 37 4043 46 4952 55 58 61 64 67 70 73 - 3. 0

6 1.0 X X XXXX X XX X X X X X XXXXX J XX XXX XX X 5 X XX XX XX X X XX X 0 XXXX 4X 7 10 13 16 19 22 25 28X 31X X 34 37 40 43 46 49 52 55 58 61 64 67 70 73 X X X XX X 4 XX X D X X X X - 1. 0 X 3 X X

X X X 2 - 2. 0 X

1 X - 3. 0

0 4 7 1013 16 19 22 25 28 31 34 37 4043 46 4952 55 58 61 64 67 70 73

7 6

6 5

5 E 4 F 4 3 3

2 2

1 1

0 0 4 7 1013 16 19 22 25 28 31 34 37 4043 46 4952 55 58 61 64 67 70 73 4 7 1013 16 19 22 25 28 31 34 37 4043 46 4952 55 58 61 64 67 70 73 chain length chain length 48

Figure 2.6 Amylose/amylopectin content of starch isolated from ornamental tobacco floral nectaries at various stages of development.

2, 6, 9, and 12 are nectary stages from which starch was isolated. 48 h PF are nectaries from flowers 48 h after fertilization. Potato starch is included as a reference.

Amylopectin 80 Amylose

0 2 6 9 12 48h PF Potato Starch 50

only very low levels. In Panel B the intermediate length chains (DP ~15 to 25) are present in higher levels compared with Stage 2. In addition, the medium-long chains (DP ~35 to 50) are also significantly increased in Stage 6. By Stage 9 (Panel C), the chain lengths peak for all of the intermediate and long chain lengths. This finding fits well with the proposed buildup of a more highly crystalline form of starch during the filling stage of nectary development. By Stage 12, starch has undergone degradation to produce sugar for nectar secretion. As shown in Panel D, starch from Stage 12 contains fewer long chains and more short chains when compared with Stage 9. This indicates that at Stage 9, the longer chains make up larger mass of the total starch. This pattern continues to post fertilization stages (Panel E). Panel F represents the side chain pattern for potato starch as a control for both calibration and comparison with isolated starch from tobacco. These patterns are better illustrated in subtraction plots. Panel G represents the starch chain lengths at Stage 9, minus chain lengths at Stage 2. This plot illustrates that Stage 2 is enriched in the short side chains (from DP = 4 to ~20) and is deficient in longer length chains (DP > 20). Panel H shows Stage 9 minus Stage 6. At Stage 6 nectaries have begun to increase in size even though starch still contains more short chains (DP = 4 to 20) and fewer long chains (DP > 25), than Stage 9 starch. We interpret these data to mean that there is an increase in the overall lengths of all chains as the nectary develops and that by Stage 9, the maximum chain lengths are attained. Between Stages 9 and 12, starch appears to be degraded. By Stage 12, all chain lengths are shorter, and the chains with DP ~ 8 to 20 are the most abundant. Finally, 48 h after fertilization, starch is difficult to isolate, but it contains significantly more short and intermediate length chains (DP ~13 to 35) than the Stage 9 starch (Panel I). As interpreted, these data indicate that starch from later stages is significantly degraded when compared to starch at Stage 9. To evaluate how the accumulation of carbohydrate in nectaries compared with nectar sugar output at anthesis, we ran a complete carbohydrate analysis on Stage 9 nectaries to compare with total nectar sugars. In Stage 9 nectaries, most of 51

the carbohydrate is present in starch (67.6%), sucrose (17.7%), glucose (8.2%), and fructose (6.4%). There was very little detectable water soluble glucan (<1%) or fructan (<0.5%) present in the nectary tissues (Figure 2.4B). The total amount of carbohydrate detected in Stage 9 nectaries was 5.36 µmoles of glucose equivalents per flower (Table 2.2). For a comparison, nectar was isolated from flowers immediately upon anthesis (Stage 12) and the amount of sucrose, glucose, and fructose were quantitated. In Stage 12 nectaries, we detected 6.05 µmoles of glucose equivalents per flower. Thus, the total amount of starch and free sugars present in Stage 9 nectaries is able to account for the majority of the total carbohydrate present in the nectar and in the nectary indicating that other sources of carbohydrates besides nectary starch are required for a robust nectar production at anthesis. However, these studies also demonstrated that nectar volume and nectar carbohydrates continued to increase as the flower remained open after anthesis.

2.3.4 Role of transported photosynthate in nectar production To evaluate whether transported photosynthate participates in the flow of nectar, we initiated a series of experiments in which [14C]-labeled sucrose and [14C]- labeled glucose was fed to flowers through the pedical of cut flowers. These studies demonstrated that [14C]-labeled sugars accumulated to the highest level in nectaries over all other floral organs (data not shown). In addition, to determine whether the photosynthetically transported sugars accumulating in nectar had been metabolized, we evaluated the chemical composition of the labeled nectar sugars using thin layer chromatography (TLC). Figure 2.7 shows a TLC analysis of nectar from five individual flowers labeled with [14C]-glucose and five individual flowers labeled with [14C]-sucrose. Panel A is stained with p-anisaldehyde to show total nectar carbohydrates, while Panel B is a PhosphorImager scan to identify labeled compounds. As can be seen in Figure 2.7, the labeled sugars appearing in the nectar all contain similar forms. No other forms of labeled compounds were identified in nectar (Figure 2.8B) nor were any other unlabeled sugars identified at measurable levels in the nectar of these plants (Figure 2.7A). Regardless of which labeled sugar 52

Table 2.2 Composition of ornamental tobacco LxS8 nectar

Stage 9 nectaries Sample µg/flower µmoles/flowera Percentage (%) Starch 658 + 92 4.06 75.8 Sucrose 172 + 40 0.50 9.4 Glucose 80 + 19 0.44 8.3 Fructose 63 + 10 0.35 6.5 TOTAL 5.36 1.00

Stage 12 nectariesb Sample µg/flower µmoles/flowera Percentage (%) Starch 67 + 27 0.42 6.9 Sucrose 785 + 38 2.30 38.0 Glucose 300 + 21 1.67 27.6 Fructose 300 + 31 1.67 27.6 TOTAL 6.05 1.00

a molar equivalents were calculated as follows: µMoles Starch = µg Starch/(180- 18); µMoles Sucrose = µg Sucrose/((2*180)-18); µMoles Glucose = µg Glucose/180; µMoles Fructose = µg Fructose/180. b Stage 12 here is the flowers just after anthesis. 53

Figure 2.7 TLC analysis of radiolabeled nectar.

Positions of labeled sugar standards were run at each edge of nectar samples. Nectar samples 1 through 5 were from individual flowers and were labeled with [14C]-glucose. Nectar samples 6 through 10 were from other individual flowers and were labeled with [14C]-sucrose.

Panel A. TLC plate was stained with p-anisaldehyde as described in Materials and Methods. Positions of migration for nectar sugars are shown; Panel B. TLC plate was exposed to a PhosphorImager plate for 92 h and visualized on a Molecular dynamics Typhoon 8600 PhosphorImager.

1 2 3 4 5 6 7 8 910 1 2 3 4 5 6 7 8 910

A B 55

Figure 2.8 Labeling of C14-sucrose in various floral stages

Identify floral stage as stage 9, stage 10, stage 11, stage 12s (stage 12 + 0 h, stage 12 + 4 h, stage 12 + 8 h, stage 12 + 12 h, stage 12 + 24 h, stage 12 + 36 h, stage 12 + 48 h) were used in this experiment. Six replicates were used in this experiment for each floral stage. 2 µCi of [14C]-sucrose were put into 1 mL of feeding solution (17% sucrose, 0.01% glucose and 0.01% fructose). Each flower with identified stages was put into 200 uL of feeding solution for 2 h.

Panel A. Nectar volume in various floral stages; Panel B. Total carbohydrate accumulation of nectar in various floral stages detected by anthrone; Panel C. Initial accumulation rate of label in various floral stage nectar following the feeding of [14C]-sucrose.

(sucrose or glucose) was fed to flowers, the labeled sugars present in nectar were sucrose + glucose + fructose in the same ratio as these same compounds found in wildtype Lxs8 nectar (Table 2.3). To determine whether cleavage of labeled sucrose occurred prior to or post-secretion, we incubated [14C]-sucrose with whole soluble nectar. Even after a 24 h incubation, no cleavage of labeled sucrose into labeled glucose and fructose was observed (data not shown). There we conclude that invertase activity is not present in the nectar of ornamental tobacco and that processing of transported sucrose is most likely to occur within the nectary. To more finely dissect the process of photosynthate accumulation in nectar we evaluated the photosynthetic flow into nectar at different floral stages. Initially we examined the total flow of carbohydrate that accumulated in extracellular nectar. This was done by washing the inside of the floral tube with 1 mL of water and quantitating the total sugar composition with the anthrone assay (Morris, 1948). As shown in Figure 2.8A, extracellular carbohydrate (secreted nectar) begins to accumulate by stage 10, however the total accumulation of extracellular carbohydrate remains low anthesis at Stage 12. Subsequently we evaluated the high-level flow of nectar that was produced by these flowers. This parameter was evaluated by physically measuring the total volume of nectar that was produced by these flowers. These measurement showed that higher volume nectar flow was not significant until anthesis (Stage 12), when nectar began to accumulates at a rapid rate and continued throughout the remainder of the experiment (Figure 2.8B). Finally to evaluate transport of photosynthate we incubated the pedicels of stage flowers in artificial phloem plus [14C]-radiolabled sucrose and quantitated the amount of sugar transported, during a 2 h timeframe. As can be seen (Figure 2.8C), little sugar was transported into nectar prior to anthesis (Stage 12). However, in the hours after anthesis these was a burst of photosynthate transported into nectar. This process peaked 12 h after anthesis and the transport declined thereafter. Based on these observations, we conclude the following: A) that nectar begins to be secreted at low levels by floral stage 10; B) that a high-level flow of 58

Table 2.3 Composition of labeled nectar in ornamental tobacco LxS8

% compositionn Sample Reference Sucrose Glucose Fructose Nicotiana LxS8 wildtype 53.7% 24.2% 22.1% Carter et al., in press nectar [14C]-sucrose nectar 54.4% 23.5% 22.2% This work - scans of [14C]-glucose nectar 44.4% 31.3% 24.3% Figure 8 n = 5 replicates 59

nectar begins following anthesis and continues for at least 48 hours thereafter; and C) that the high-level flow of nectar is coordinated with the transport of photosynthate into nectar.

2.3.5 Is tobacco a model for other plant species? To evaluate whether the nectaries of other plant species also accumulate starch during nectary development, we have performed PAS-staining of the floral nectaries of Arabidopsis thaliana plants, prior to and at anthesis. Arabidopsis has six individual floral nectaries sandwiched between the stamen filaments and the gynoecium (Davis et al., 1998). Figure 2.9A shows a PAS-stained Arabidopsis nectary prior to anthesis (approximately Stage 8). It is clear that starch grains are present in this nectary. Figure 2.9B, shows a nectary from a just opened flower (Stage 12). Nectaries at this latter stage contain no visible PAS-staining starch grains. These results indicate that Arabidopsis nectaries do indeed accumulate some starch, and that this starch is hydrolyzed prior to anthesis. Thus, Arabidopsis appears to be analogous to ornamental tobacco with regard to the timing of the accumulation and degradation of starch and ornamental tobacco may indeed serve as a model for other plant species.

2.4 Discussion Plants attract visiting pollinators by offering metabolically-rich floral nectar as a reward. Chief among the nectar ingredients are various forms of carbohydrates (sucrose, glucose and fructose), although many other components also are present (Carter et al, 2006). Because the levels of sugars that accumulate in nectar are much higher than the levels that occur in phloem (35% in nectar versus about 17% in phloem), the nectary requires a sugar concentration strategy to provide a ready storehouse for nectar secretion. Initial observations in the development of the floral nectary demonstrate that the nectary undergoes two very obvious changes. First is a change in color that we have shown to be due to the accumulation of β-carotene (Horner et al., in press). 60 61

Figure 2.10 Comparison of biochemistry with nectary development.

Appearance of gynoecium with nectary at base is shown at three stages: Stage 2 (emerging bud), Stage 6 (middle stage), and Stage 12 (anthesis). Nectary stages have been defined (organ initiation, filling, ripening, maturation, and post maturation). Biochemical events that we have characterized include: Starch biosynthesis during the filling stage of nectary development forming amyloplasts; starch hydrolysis during the maturation stage of nectary development releasing sucrose, glucose and fructose into nectar; synthesis and secretion of nectarins, particularly NEC1 and NEC5 that function to generate high levels of hydrogen peroxide that drives the Carter-Thornburg Nectary Redox Cycle.

Second is a change in size that is due to the accumulation and storage of starch within the nectary during the filling period. After accumulation of starch through floral stages 9, the starch is hydrolyzed during the maturation period (Stage 9 to Stage 12). We have demonstrated this accumulation and degradation of starch in the nectary using two independent methods. First, we used light microscopy of PAS- stained nectaries. This method stains starch very efficiently and reveals the massive accumulation and subsequent degradation of starch. The granular accumulation of the PAS-positive starch observed in Figure 2.3, Panels F to O implies that the starch is localized in amyloplasts. The second method used was the physical isolation and characterization of starch from nectaries at the different floral stages (Figure 2.4). Both of these methods demonstrated that the nectary accumulates a large starch reserve during development, and that the starch was mostly degraded immediately prior to anthesis.

2.4.1 Transient nature of the nectary starch Starch is the most efficient form of sugar storage in cells. Starch granules consist almost entirely of the glucose polymers amylose and amylopectin. Amylose molecules are essentially linear, containing 102 to 104 of α-1,4 linked glucosyl residues. Amylopectin molecules are larger (104 to 105 monomers) and also contain α-1,6 branch points within their (Robin et al., 1974, French, 1984, Kainuma, 1988, Imberty et al., 1991). We characterized starch at different stages of nectary development with respect to both composition and structure. Together, these analyses indicate that a highly crystalline form of starch accumulates at Stage 9, the developmental stage at which nectary starch is at its peak. This was shown first by evaluation of the amylose/amylopectin ratio, which revealed a general trend towards a higher amylopectin content in the later stages of nectary development. Because the more highly branched, complex structure of the amylopectin molecule is known to account for the semicrystalline nature of starch granules (French, 1984), starch granule 63

crystallinity and maximum stored carbohydrate are predicted to be highest at Stage 9. This is supported by the evaluations of starch structures during nectary development to compare the relative abundance of specific lengths of linear chains in the amylopectin molecules. These structural analyses demonstrated that there were increases in the proportions of both short and long chains up to Stage 9, when the maximum chain lengths were attained. In particular, intermediate length chains with DP ~12 to 25 were shown to increase during this period of starch production. Recent analyses of starches from a number of plant species indicates that chains of these intermediate lengths are involved in the double helical formation of linear chains that forms the basis of starch granule crystallinity (Umemoto et al., 2002, Morell et al., 2003, Zhang et al., 2004). Following Stage 9, nectary starch appears to be significantly reduced in complexity as would be expected during a period of starch degradation.

2.4.2 Role of transported photosynthate in nectar production We have also evaluated the role of transported photosynthate in the production of nectar by feeding radiolabeled sugars to flowers, and following the movement and accumulation of the radiolabels in nectar. Despite the fact that the nectary is not innervated by either phloem or xylem (Horner et al., in press), it accumulates very high level of all floral organs when flower are fed with [14C]-labeled sugar through the pedical. TLC analysis of the radioactivity accumulating in nectar showed that regardless of whether glucose or sucrose was fed, the labeled sugars secreted into nectar were the same (sucrose + glucose + fructose, [molar ratio 1:1:1]). This implies that during the nectary transit, the sugars were metabolized into the same secreted products that are found in the wildtype Lxs8 nectar. These studies also demonstrate that nectar is produced from at least two sources. First it appears that hydrolysis of nectary starch in the late phase of nectary development releases a bolus of sugar that can account for the sugars present in nectar at anthesis. However, within hours following anthesis, there is a second phase of nectar production that is 4 to 5 times larger than that the amount of sugar derived 64

from nectary starch. This second phase of sugar production appears to arise from transported photosynthate that transits the nectary. Thus, in ornamental tobacco, nectar production is due to two process, sugar from starch hydrolysis and sugar from transported photosynthate. Since the hydrolysis of starch proceeds the transport of photosynthate, the question arises as to whether the hydrolysis of starch is related to the increase in photosynthate transport. Degradation of the large reserve of nectary starch (20% by mass) in the short timeframe between Stage 9 and Stage 12 (anthesis), produces a huge concentration of glucose in a relatively sort time frame. Based on the amount of starch present at Stage 9 nectaries and the total nectary volume, we estimate that their hydrolysis produces a sugar concentration of 0.5 M in the nectary. This would be expected to raise the osmolarity of the nectary to 1 MPa or even higher. The result of this rapid and dramatic increase in intracellular osmolarity will lead to the water potential, ψ, of the nectary dramatically decrease. Because the water potential of phloem in leaves and stems is thought to smaller than flower (Kaufmann and Kramer, 1967, Chapotin et al., 2003, Nerd and Neumann, 2004), this would result in a large influx of water from the surrounding phloem to floral nectary. However, the accumulation of fluid within the nectary also increases the hydrostatic pressure of fluid within the nectary. This pressure is alleviated by the escape of this fluid from the nectary pores that are located on the opposite faces of the nectary. In this mechanism, hydrolysis of the stored nectary starch serves both as an initial source of carbohydrate for nectar as well as a mechanism to “prime the pump” and drive high level nectar secretion due to phloem influx into the nectary. A similar starch hydrolysis hypothesis has been advanced to explain how flower petal open precisely at anthesis (Hammond, 1982, Ho and Nichols, 1977, Bieleski et al., 2000). In this petal opening hypothesis, starch accumulating during floral development is hydrolyzed just prior to anthesis releasing high levels of glucose that results in a huge influx of water into the petals causing cells to enlarge thereby driving the petal opening. Thus, a similar mechanism may underlie opening of the flower at anthesis and the generation of nectar flow. 65

Thus rapid production of glucose from the breakdown of nectary starch has at least three consequences. First, sugar sensing mechanism are know that mediate gene expression in response to sugar concentration (Rolland et al, 2002). These mechanism may also function in regulating the maturation phase of nectary development during which time the nectary develops from a nonsecretory anabolic organ into the plant’s premier secretory organ. Second, the hydrolysis of starch results in the high level production of carbon substrates that can be utilized by other metabolic pathways. We have recently demonstrated that these high levels of glucose provides a substrate for the large scale of synthesis of β-carotene that occurs in the nectary during the maturation phase of development (Horner et al, in press). Third, the rapid production of glucose from starch hydrolysis following Stage 9 results in a rapid increase in intracellular osmolarity. Thus, these results have permitted us to define a crucial step in the biochemistry of nectary development. Based on them, we propose the model shown in Figure 2.10 to explain all known nectary biochemistry. At early stages of nectary development, plastids are present, giving the nectary a lime-green color (see Figure 2.2). During the filling period (Stages 2 through 9), starch accumulation in the nectary, reaches a very high level. This, along with increase in number and size of cells (data not shown), causes the nectary to increase in size. Beginning about Stage 9, the maturation period of nectary development begins and starch is hydrolyzed to provide sugars (sucrose, glucose, and fructose) for nectar production. At the same time, photosynthates arising from other tissues will be transported into nectary cells to be released as nectar. Deciphering the molecular events underlying this mechanism will provide new insight into floral biology and into the interaction between plant and their pollinator to be released as nectar. About midway through the maturation period, nectar proteins are synthesized and secreted into the apoplast, where they are carried along with the soluble nectar. These proteins (particularly Nectarin I and Nectarin V) are responsible for the high levels of hydrogen peroxide that accumulate in the nectar 66

Figure 2.9 Light microscopic images of 1-µm-thick LR White resin sections of Arabidopsis thaliana floral nectaries stained with PAS technique.

Panel A. Nectary from flower prior to anthesis (Stage 8) shows small, scattered starch grains in nectary tissues (arrows) and portion of a stomate (*) are shown. Panel B. Nectary from a Stage 12. There are almost no starch grains in nectary tissues. Two stomates are clearly visible (*). Bars = 25 µm.

67 68

and drive antimicrobial activity of the recently described Carter-Thornburg Nectar Redox Cycle (Carter and Thornburg, 2004a). Because of their size and demonstrated research potential, ornamental tobacco nectaries have become a model system for study of nectar/nectary biology. The nectaries are sufficiently large that enough tissue can be isolated for enzymatic and other biochemical assays. This is not possible with nectaries of other model plants, such as Arabidopsis. However, although Arabidposis nectaries exist as separate conical cellular protuberances they appear to be structurally similar with tobacco nectaries, particularly with regard to having a stomatal zone (compare Figures 2.1 and 2.9) and to starch formation and degradation (see Figures 2.3 and 2.9). While differences between the two species probably do exist, we anticipate that the use of Arabidopsis as a genetic model, coupled with the use of ornamental tobacco as a biochemical and developmental model, will allow us to better focus on nectary biology in general. This fusion of these complementary models should permit us to make rapid progress in understanding the mechanisms that underlie nectar production, and should permit us to design novel molecular strategies to enhance the attraction of insect pollinators for more efficient pollination, and ultimately greater yield.

Chapter 3 Expression of starch metabolic genes in the developing nectaries of ornamental tobacco plants

This chapter is based upon a manuscript that has been submitted to “The Plant Sciences”

Authors: Gang Ren1, Rosanne A. Healy2, Harry T. Horner2, Martha G. James1, & Robert W. Thornburg1

Abbreviations: TIGR, The Institute for Genomic Research

Acknowledgements: The authors would like to thank NSF #IBN-0235645 (RWT), the Carver Trust (RWT), the Hatch Act and State of Iowa Funds (RWT), and a USDA Service Center Agency Current Research Information System project 3625- 21000-029-01S (HTH). The authors also would like to graciously thank Dr. Yutaka Miyazawa for providing us with several Nicotiana tabaccum starch biosynthetic genes. And Dr. Prem Chourey, Florida State University and Dr. Thomas Okita Washington State University, who supplied the antisera for these studies.

Keywords: nectar, nectary, Nicotiana, starch metabolic gene, gene regulation 70

Contribution of this work: Unless otherwise noted, the experiments in this manuscript were performed by Mr. Gang Ren. In addition, Mr. Ren was responsible for carrying this project forward throughout the course of the project. Ms. Rosanne Healy performed the Microscopic experiment for Figure 3.1. Dr. Harry Horner consulted on the methods of Microscopic sections of this manuscript. Dr. Martha G. James consulted on starch gene isolation and Real-time RT-PCR in this manuscript. Dr. Robert Thornburg consulted on the overall creative instructions and support for this work. In addition he contributed to the writing, reviewing and editing of this work. This work was supported by an NSF grant (IBN-0235645) to Dr. Robert Thornburg and Dr. Martha James. This work was supported by an NSF grant (IBN-0235645) to Dr. Robert Thornburg and Dr. Martha James. 71

Abstract The accumulation of starch within the ornamental tobacco floral nectary gland was studied by TEM analysis of plastid structure throughout flower development. Amyloplasts accumulated in the nectary from early stages of nectary development until Stage 9, and then rapidly disappeared prior to anthesis (Stage 12). We examined expression of starch metabolic genes in the nectary to identify the underlying molecular mechanisms responsible for the transient accumulation of starch in the floral nectary. Initially, we isolated 18 nectary-expressed cDNAs that encoded different starch metabolic enzymes. The encoded protein sequences were compared with well-characterized protein sequences to identify gene functionality. We examined several steps of starch metabolism. Experiments showed that sucrose synthase were strongly expressed in nectaries at both the mRNA level and protein level. We also examined ADP-glucose pyrophosphorylase (AGPase), AGPase mRNA, protein and enzyme activity was strongly expressed at early stages of nectary development, but expression declined as the nectary matured. Starch synthase genes were differentially expressed in ornamental tobacco nectaries. SS3 was the most strongly expressed isoform and was expressed at early stages but declined at later stages. Starch branching enzymes and starch debranching enzymes were also differentially expressed in nectaries. Quantitative RT-PCR revealed three different regulatory pattern of gene expression in the nectary. Most starch anabolic genes, including AGPS, SS3 and SS1, were expressed early in nectary development (Stage 2 and Stage 6), and were down regulated after Stage 9. In contrast, the starch catabolic genes, including ISA1, AMY, BMY, were not detected at early stages, but were induced by Stage 9 of nectary development. A third class of gene expression that included R1 protein, Pho, and SBE1 were expressed throughout nectary development. The switch from starch anabolism to catabolism may be an important key to the normal development and function of nectary. 72

3.1 Introduction The floral nectary, first described by Carl Linnaeus, is a remarkable organ that serves to provide the carbohydrate rich nectar to visiting pollinators in return for gamete transfer from flower to flower (Linnaeus, 1751). In addition, in ornamental tobacco, the nectary recently has been shown to play a defensive role in protecting the rich nectar against the infection of microorganisms vectored by microbe-carrying pollinators (Carter et al., 2006; Carter et al., 2007; Thornburg et al., 2003). The floral nectar of ornamental tobacco contains highly concentrated sugars, amino acids (Carter et al., 2006), vitamins (Carter and Thornburg, 2004c), and proteins (Carter and Thornburg, 2004a; Carter and Thornburg, 2004b; Carter and Thornburg, 2004c). This defensive strategy is based on the accumulation in nectar of a high level of hydrogen peroxide (Carter and Thornburg, 1999). The hydrogen peroxide is produced by the action of Nectar Redox Cycle, a novel biochemical pathway in gynoecium which allows the cells to survive in the presence of high levels of hydrogen peroxide. Both extracellular and intracellular antioxidants are also produced by the nectary (Carter and Thornburg, 2004a; Horner et al., 2007). Because carbohydrate metabolism is a major aspect of nectary function, we have undertaken studies to evaluate this process in the nectary throughout floral development. These studies have shown that starch transiently accumulates in the nectary during floral development, and becomes one source of carbohydrate in the nectar (Ren et al., submitted). An additional source of nectar carbohydrate arises from transported photosynthate (Ren et al., submitted). Starch, a mixture of two distinct components: amylopectin and amylase, forms a highly efficient carbohydrate storage molecule that is deposited in specific plastids termed amyloplasts (Ball et al., 1996; Morell et al., 2005; Myers et al., 2000; Smith et al., 1997). Starch is synthesized by four major groups of genes: ADP- glucose pyrophosphorylase (AGPase, EC 2.7.7.27), the rate-limiting step of starch biogenesis provides activated glucose moieties for starch production (Crevillen et al., 2005; Geigenberger et al., 2005; Jin et al., 2005; Sanwal et al., 1968). Starch synthase (SS, EC 2.4.1.21), catalyzes the polymerization of glucose units into 73

polysaccharides (Delvalle et al., 2005; Edwards et al., 1999; Hirose and Terao, 2004; Tenorio et al., 2003; Zhang et al., 2005). Two additional groups of enzymes, the starch branching enzyme (SBE, EC 2.4.1.18) and starch debranching enzymes (DBE, EC 3.2.1.41, and EC 3.2.1.68), function to modulate the architecture of starch structure to permit the most efficient packing of sugar into the starch granule (Myer et al., 2000). Starch degradation is also the result of enzymes cooperation. Alpha- amylase (AMY, EC 3.2.1.1) and β-amylase (BMY, EC. 3.2.1.2) are the major enzymes responsible for starch degradation (Smith et al., 2005; Tetlow et al., 2004; Zeeman et al., 2004). Alpha-amylase is responsible for endogenous digestion of polyglucan chains, while β-amylase is mainly responsible for exogenous digestion of polyglucan chains. Recent progress shows that glucan water dikinase (GWD, EC 2.7.9.4, also named as R1 protein) and starch phosphorylase (Pho, EC 2.4.1.1) are also involved in starch synthesis and starch degradation (Blennow et al., 2002; Ritte et al., 2002; Reimann et al., 2002; Smith et al., 2005). The R1 enzyme functions in starch phosphorylation and may predispose starch for degradation (Blennow et al., 2002; Ritte et al., 2002; Reindeer et al., 2002). Despite the well investigated core pathways of starch metabolic genes, the molecular and developmental mechanisms of starch metabolism in developing floral nectary are for the most part unknown. In this study we have focused on the transcriptional regulations of starch metabolism in ornamental tobacco. While starch metabolism has been widely studied in a variety of organisms, including maize (Hannah, 1997; James et al., 2003), Arabidopsis (Zeeman et al., 2002), Potato (Geigenberger, 2003), and other organisms, this work and our earlier contributions (Ren et al., submitted), represent the first attempt to define the transient metabolism of starch in plant nectaries. Starch is known to accumulate in the floral nectaries of a number of plant species including Cucurbita pepo (Nepi et al., 1996), Passiflora sp. (Durkee et al., 1981), Hibiscus rosa-sinensis (Sawidis, 1998), Rosmarinus officinalis (Zer and Fahn, 1992), Pisum sativum (Razem and Davis, 1999) and Limodorum abortivum (Figueiredo and Pais, 1992). Whether these other species used sugar capture and metabolism that found in ornamental tobacco 74

is not clear; however, the mechanisms that we are describing in ornamental tobacco provides a molecular model against which they can be compared.

3.2 Materials and methods All materials were from Sigma Chemical Corp (St. Louis, MO) or Fisher Chemical Company (Pittsburgh, PA) and were of the highest purity available. An aliquot of antibody raised against the maize sucrose synthase were kindly provided by Dr. Prem Chourey, Florida State University. An aliquot of antiserum raised against the potato ADP-glucose pyrophosphorylase small subunit were kindly provided by Dr. Thomas Okita, Washington State University.

3.2.1 Plant materials The LxS8 line of ornamental tobacco plants used for these nectary studies have been previously described (Carter et al., 2007). Nectary stages were identified and isolation of nectar and floral tissues was as previously described (Carter et al., 1999). If not used immediately, tissues were maintained at -20˚C until use. Plants were grown under 16h day/8h night conditions in the greenhouse. The 12 different floral stages of tobacco were determined according to the method of Koltunow (Koltunow et al., 1990). Materials were collected on ice, and nectaries from flower Stages 2, 6, 9, and 12 were dissected away from ovaries. Samples of young leaves (approximately 25 mm long) and for dark samples Stage 12 nectaries were also collected under green illumination two hours before local sunrise (prior to nautical twilight).

3.2.2 rDNA methods Methods describing the construction and use of the several nectary cDNA libraries as well as the isolation and characterization of 13,696 EST sequences will be forthcoming in a separate manuscript (Taylor et al., in preparation). Primers used for isolation of specific starch metabolic genes by PCR were based upon conserved 75

regions of those genes determined by sequence comparisons of nucleotide sequences from other solanaceous species and are presented in Table 3.3.

3.2.3 PCR methods primer design Comparisons of gene structures with cDNA sequences from available Arabidopsis and solanaceous sequences were used to predict putative exon-intron junctions in the target ornamental tobacco gene. A webpage-based program (http://seq.yeastgenome.org/cgi-bin/web-primer) was used to design primers for RT- PCR and quantitative (real-time) RT-PCR. Both pairs of primers were crossed the predicted exon-intron junctions to cross out genomic DNA contamination during RNA isolation and cDNA synthesis. The annealing temperatures for both pairs of primers were 58˚C to 60˚C, and the amplicon sizes ranged from 88 to 317 bp (Table 3.5).

General PCR The general PCR reaction included: 50 µM of each primer, 1 X PCR master mix (Eppendorf, cat # 200 250 2250), and 200 ng of DNAs prepared from various nectary cDNA libraries (Navqi et al., 2005). PCR products that had the expected size were subcloned into pGEM-T easy vector (Invitrogen, cat # A3600) and following ligation were transformed into E. coli DH5a cells. Plasmids from positive transformants were purified and both DNA strands were completely sequenced to insure a correct sequence.

RT-PCR detection A reverse transcription system (cMaster™ RTplusPCR system, Eppendorf, cat # 003 003 358) was used to synthesize first strand cDNA from each of the RNA isolations according to manufacture’s instruction. A 2.5 X PCR master mix (Eppendorf, cat # 200 250 2250) was used for PCR amplification. The PCR system includes: 300 nM primers, 500 ng of first strand cDNA, 1 X PCR master mix (Eppendorf, cat. 200 250 2250). PCR reactions were performed with the following 76

program (94ºC 30 sec, 55ºC 30 sec, 72ºC 1 min) for 25 to 28 cycles. PCR products were run on 1.5% agrose gel containing ethidium bromide, visualized under UV lights, and photographed. The ornamental tobacco 18S rRNA and 26S rRNA were used as internal controls to monitor and normalize gene expression level.

Quantitative RT-PCR detection A StrataScript® First Strand cDNA Synthesis System (Stratagene, cat# 200420) was used to synthesize first strand cDNA from RNA isolated above. A Brilliant® SYBR™ Green QPCR master mix (Stratagene, cat# 600548) was used to visualize the RT-PCR reaction. Each reaction contained 0.5 µL first strand cDNA and 300 nM of each primer. The quantitative RT-PCR reaction was hot-started at 55ºC for 10 min, then performed for 35 cycles (95ºC for 45 sec, annualized at 55ºC for 45 sec and elongated at 72ºC for 1 min). The florescence was detected by QPCR machine (Stratagene M4400). Three independent RNA isolations were performed; with two replications for each of the RNA isolations (6 independent replicates for each) was assayed for each specific tissue or stage. A glyceraldehyde-3-phosphate dehydrogenase (GAPDH) clone (001G06) from the LxS8 EST project (Taylor et al., in preparation) was used as an internal control to monitor and normalize mRNA expression. Fluorescence of each assay was monitored throughout the 40-cycle PCR experiment. A fluorescence threshold (T) was set during the early linear phase of amplification at about 100 dR fluorescence units. The cycle number that gives this

threshold value was then calculated for all samples. This cycle threshold value (CT)

was used for all calculations. CT values were calculated for both GAPDH and for the

individual genes for each assay in the experiment. The CT for GAPDH was compared between replicates, between different samples, and between different experiments to insure that results were comparable. To evaluate gene expression

(-∆CT) - for any gene relative to GAPDH, we used the formula (Ngene/NGAPDH=2 ; where

∆CT=CTgene-CTGAPDH). These analysis, therefore provide a level of gene expression for any gene relative to that of GAPDH at each time point.

3.2.4 Bioinformatics tools Completely sequenced DNA sequences were used to blast against the National Center for Biotechnology Information (NCBI)’s GenBank, and The Institute for Genomic Research (TIGR)’s gene index by using either tblastn or blastn (http://www.ncbi.nlm.nih.gov/blast and http://compbio.dfci.harvard.edu/tgi/plant.html) algorithms. Homologous gene sequences identified from these searches were used for ClastalW (http://align.genome.jp) analysis to produce the phylogenetic (neighbor- joining) trees used in these studies.

3.2.5 Microscopy methods Transmission Electron Microscopy For general ultrastructure, small portions of fresh, isolated nectaries were initially fixed in 2% glutaraldehyde and 2% paraformaldehyde in a 0.1M sodium cacodylate buffer, pH 7.2, at 4ºC overnight, washed in cold buffer, postfixed in 1% osmium tetroxide in same buffer at 4ºC for 1.5 h, washed with buffer and deionized water (deiwater), and en bloc stained with 2% aqueous uranyl acetate in the dark. The specimens were washed in deiwater. Then the specimens were divided into two batches for further processing into either Quetol or Spurr’s resin mixtures. For Quetol resin embedding (manufacturer’s instructions; www.polysciences.com), following deiwater washes, specimens were processed through increasing concentrations of Quetol and decreasing concentrations of deiwater to pure Quetol monomer, and then into Quetol resin mixture [Quetol 651 (15 g), NMA (10 g), NSA (20 g), and DMP-30 (1 g)], followed by casting in aluminum trays, and polymerized at 75ºC for 1 d. For Spurr resin embedding (Spurr, 1969) following deiwater washes, specimens were dehydrated in an ethanol series, transferred to pure acetone, and infiltrated in acetone and Spurr’s hard resin mixture. Specimens were then transferred into pure resin mixture, cast in aluminum trays and polymerized at 70ºC for 2 d. Specimens embedded in both resin mixtures were sectioned using either glass or diamond knives. Thin 60-nm-thick sections were picked up with copper slot grids and deposited on Formvar-coated racks, and stained with 5% aqueous uranyl 78

acetate and lead citrate (Reynold, 1963). Observations were made on a JEOL 1200EX TEM (www.jeol.com), and images were captured photographically on Kodak SO-163 film (www.kodak.com), which was later digitized for further processing using an Epson Perfection 3200 PhotoScanner (www.epson.com).

Quantitation of plastid volume For quantitation of starch, five cells located three cells inward from the epidermis from each stage were photographed in cross section at 3K. All measurements were done using a Soft Imaging System Analysis program (http://www.soft-imaging.net). Each cell area was measured and the average determined. Within each cell, the numbers of plastids and starch grains were quantified, and the area of each plastid and each starch grain was measured. Area measurements were converted to volumes to provide realistic numbers per cell volume per nectary stage.

3.2.6 RNA isolation RNA was isolated from 16 different ornamental tobacco organs, including leaf, stem, root, sepal, petiole, petal, floral tube, ovary, stigma, anther, and floral nectaries from Stage 2, 6, 9, 11, 12, and 48 hours after fertilization, were independently collected on ice, dissected from the plants, and immediately frozen in liquid nitrogen. Three independent tissue collections and RNA isolations were performed to represent a specific tissue, or a specific stage of nectary. Approximately 300 to 400 mg of each tissue (except for nectary tissue) was used for each of the RNA isolations. For nectaries, 100 to 200 mg of tissues was dissected from about 50 flowers. Frozen tissues were homogenized with a mortar and pestle. A Promega SV RNA extraction kit (Promega, cat # Z3500) was used to isolate total RNA from the homogenized tissues according to the manufacture’s direction. 5 Units DNase I was added for 15 min to completely digest genomic DNA. Phenol:chloroform extraction and isopropanol precipitation was subsequently used to further purify and concentrate the RNA. 79

3.2.7 Protein methods Protein isolation and enzyme activity assay To isolate proteins from each floral stage, 50 flowers were collected, floral tissues were dissected and the tissues were homogenized with liquid nitrogen. To each sample, we added 250 µL of 62.5 mM Tris-HCl (pH 6.5), 50 mM DTT solution, containing 1% protease inhibitor cocktail (Sigma, cat # P9599). Samples were then centrifuged at 4ºC, 5000 rpm for 30 min and the supernatant was collected as a crude protein extract. Protein in each sample was quantified using Bradford method [54] [Bradford, 1976] with bovine serum albumin as a standard. Each sample was then diluted to 1 µg/µL for enzyme activity assay. AGPase enzyme activities were detected by using the method from Appledoorn (Appledoorn et al., 1999) with one modification: NADH formation instead of NBT formation was used to record the absorbance changes. 100 µg of extracted protein in a 1 mL reaction medium that contained 100 mM HEPES buffer

(pH 8.0), 0.44 mM EDTA, 5 mM MgCl2, 0.1% BSA, 1 mM NAD, 2 Units phosphoglucose mutase (PGM) rabbit muscle, 6 Units glucose-6-phosphate dehydrogenase (G6PDH), 20 µM glucose-1,6-bisphosphate, 2 mM 3- phosphoglyerate, 10 mM NaF, 1.4 mM sodium pyrophosphate (PPi). Each reaction was started by addition of 10 µL of 2mM ADP-glucose and the absorbance changes

at 340 nm were recorded corresponding to the initial velocity (V0).

Protein isolation and immunodetection 30 µg of each sample was loaded on a 12% SDS-PAGE gel (Laemmli, 1970) and separated under 200 Voltages for 90 min. After transfer of protein to a nitro- cellulose membrane (Osmonic Inc., cat # WP4HY00010), the membrane was incubated with a rabbit antibody raised against potato AGPase small subunit (Nakata et al., 1994) or maize sucrose synthase (Koch et al., 1992). A rabbit secondary antibody conjugated with peroxidase was used and the peroxidase activity was detected on a Kodak x-ray film using an ECL™ Western Blotting Detection Reagents (Amersham, cat # RPN2109). 80

3.3 Results From previous work (Ren et al., submitted), it is clear that ornamental tobacco floral nectary accumulates starch at about 20% of floral nectary mass during its development and it degrades that starch just prior to anthesis. To confirm and extend these results we conducted an examination of the nectary using transmission electron microscopy (TEM). As shown in Figure 3.1, at the earliest stages of floral development (Stage 2) when the flower is just emerging from the bud stage (Figure 3.1A and 1B), there are already well-developed amyloplasts present in the perinuclear space surrounding the nucleus. Both the number and size of these amyloplasts increase as the nectary develops through Stage 4 (Figure 3.1C) and Stage 6 (Figure 3.1D), culminating at Stage 9 (Figure 3.1E) when starch accumulation in the nectary reaches a peak. Then between Stage 9 and Stage 12 (Figure 3.1F) the amyloplasts disappear from the nectary and are replaced by amyloplast ghosts and electron dense bodies. We have previously identified these electron dense bodies as β–carotene (Horner et al., 2007). These observations are consistent with our earlier findings that nectary starch increases in complexity during these early stages to reach a maximal complexity at Stage 9 before the starch structure becomes more simplified due to degradation between Stage 9 and Stage 12 (Ren et al., submitted). In addition, we also quantitated the volume of starch accumulation in the nectary at different stages of its development by measuring the areas of parenchyma cells, plastids, and starch granules when observed by TEM. Based upon the assumption that those entities were spheres, we then calculated the radius of each entity, and further calculated the estimated volume of each entity. This quantitation verified that starch began to accumulate at early stages at lower rate (Stage 2 and Stage 6), but had higher accumulation rate at later stages (Stage 9). In addition, cell volume was increased 2 times from Stage 2 to Stage 9 (Table 3.1). The volume of plastid per cell was increased from 5.6% at Stage 2 to 21.8% at Stage 9. The total starch volume per cell was increased from 1.7% at Stage 2 to 17% at Stage 9. The total starch volume per plastid was increased from 29.6% at Stage 2 to 81

Figure 3.1 Starch granule changes in developing floral nectary parenchyma cells Starch granule accumulation and morphology changes in floral nectary parenchyma cells during floral nectary development are observed under TEM. Red arrow indicates starch granule; Green arrow indicates chromoplast; Blue arrow indicates vacuole; Dark arrow indicates nucleus. A flowers from Stage 2, Stage 6, Stage 9 and Stage 12; B Stage 2; C Stage 4; D Stage 6; E Stage 9; F Stage 12. Bar in 1A represents 1 cm. Bar in 1B-F represents 10 µm.

A B

2 4 6 9 12 CD

E F 83

Table 3.1 Volumes of cell, plastid within cell, and starch granule within plastid in developing ornamental tobacco floral nectary cells

Stage S2 S4 S6 S9 (Plastid Volume/Cell Volume) per Cell (%) 5.6 6.4 8 21.8 (Starch Volume/Cell Volume) per Cell (%) 1.7 2.6 4.9 17 (Starch Volume/ Plastid Volume) per Cell (%) 29.6 40.8 58.8 77.7 Number of plastids per Cell 11.8 9.7 8.6 8.5

77.7% at Stage 9. These observations were consistent with our previously finding that starch made up about 20% of the total nectary mass at stage 9 (Ren et al., submitted). Based upon these and earlier observations, we conclude that starch biosynthetic apparatus is active early in nectary development, but that starch degradation machinery is activated about floral Stage 9.

3.3.1 Starch metabolic genes To further our understanding of starch metabolism in the nectary during floral development, we attempted to identify the genes involved in this process. We began by evaluating starch metabolism in Arabidopsis, maize, and potato. From these studies, we identified a list of 22 target genes (Supplemental Table S3.1) that participate in starch metabolism in these various species. Using this target list as a guide, we began to isolate cDNAs from these genes that might function in the metabolism of starch in the ornamental tobacco nectaries. Taking advantage of a nectary EST project (Taylor et al., in preparation), we searched 13,696 nectary- expressed ESTs for these starch metabolic genes; however, these studies identified only five of these 22 genes (Table 3.2; R1 protein (DQ021469), starch phosphorylase (DQ021468), isoamylase 3 (DQ021471), alpha-glucosidase (not submitted), and β-amylase (DQ021457)). We used an alternative strategy to isolate the other genes. We compared the homologous sequences of starch metabolic genes from various Solanaceous species to identify any regions of conserved identity. We relied heavily on the TIGR potato, tomato, Capsicum and Nicotiana EST databases for these studies (now located at the Dana Farber Institute, http://compbio.dfci.harvard.edu). We then designed oligonucleotide primers from these regions of high identity and PCR amplification. For template DNA we used one of three different ornamental tobacco nectary cDNA libraries (Stage 6 library, Stage 12 library, and Stage PF library), from an ornamental tobacco leaf cDNA library or from fresh first strand cDNA prepared from ornamental tobacco nectaries. The oligonucleotide primers used for these experiments are presented in Table 3.3. Fragments that were of the expected size Table 3.2 Starch metabolic genes isolated from ornamental tobacco Reference Clone (TIGR) a Ornamental Tobacco Clone b GenBank Clone Identity Gene Name Accession Clone Name a Species GenBank# Length ATG Stop 5’end 3’end Vector length % % nuc AA Sucrose synthase 1 Susy1-2 DQ021467 Potato M18745 2711 76 2491 1250 946 2113 90 94 pGEM-T Sucrose synthase 2 SuSy2-4 DQ021470 Potato M18745 2711 76 2491 1250 946 2113 90 93 pGEM-T Alpha-amylase (large isoform) AAML-3 DQ021455 Potato M79328 1962 541 1764 1083 558 1640 91 90 pGEM-T c 558 689 Alpha-amylase (small isoform) AAMS-2 DQ021456 Potato M79328 1962 541 1764 696 91 84 pGEM-T 1077 1640 Starch branching enzyme 1 SBE-2 DQ021459 Potato X69805 3114 121 2706 552 1420 1971 95 97 pGEM-T Starch branching enzyme 2 SBE2-3 DQ021460 Potato AJ011888 2982 149 2776 363 2396 2726 92 90 pGEM-T Isoamylase 1 ISO-5 DQ021461 Potato AY132996 2706 70 2451 793 199 937 90 93 pGEM-T Isoamylase 2 ISO2-1 DQ021462 Potato AY132997 2831 64 2700 1671 481 2140 90 86 pGEM-T Isoamylase 3 330G04 DQ021471 Potato AY132998 2634 32 2332 291 2045 2335 90 87 pDNR-LIB Starch synthase 1 SS1-2 DQ021463 Potato Y10416 2360 68 1993 1009 518 1521 93 91 pGEM-T Starch synthase 2 SS2-1 DQ021466 Potato X87988 2631 64 2367 640 1743 2380 94 94 pDNR-LIB Starch synthase 3 SS3-1 DQ021464 Potato X94400 4167 207 3899 797 1982 2772 94 95 pGEM-T d Granule-bound starch synthase GBSS-1 DQ021465 Tomato TC125827 2450 277 2099 793 989 1780 92 93 pGEM-T AGPase (small subunit) AGPS-1 DQ021458 Potato AY186620 1845 64 1629 539 0976 1509 94 97 pGEM-T Beta-amylase 051E09 f DQ021457 Potato TC118938d d 2058 117 1855 1904 117 1855 90 90 pTripl-Ex Starch phosphorylase 204C12 DQ021468 Potato X52385 3101 44 2944 259 2184 2442 93 97 pTripl-Ex

R1 protein 224D07 DQ021469 Potato Y09533 4851 105 4499 618 3680 4297 94 96 pDNR-LIB 85 Alpha-glucosidase 003C02 N.S. e Potato AJ001374 2992 62 2821 233 2142 2369 91 89 pTripl-Ex Nicotiana Sucrose phosphatase 342F09 N.S. AY729656 1278 1 1278 545 1061 1278 95 94 pDNR-LIB tobaccum Sucrose-phosphate synthase 1 044E08 N.S. Potato AF194022 3165 1 3165 364 3102 3165 98 100 pTripl-Ex Nicotiana Sucrose-phosphate synthase 2 352E08 N.S. DQ213015 3195 1 3195 455 2323 3195 90 92 pDNR-LIB tobaccum a: The reference clone is the potato or Nicotiana tobaccum cDNA from TIGR or GenBank that permits us to map our clones. The accession number, the length, and the positions of the ATG start and stop codons are indicated for each reference clone. b: The length of the isolated gene fragment, its 5’ and 3’ ends relative to the reference clone, the percent identities to the reference clone at both the nucleotide and amino acid level and the vector that the fragment is cloned in are indicated. c: The small isoforms of the alpha-Amylase clone appear to be an alternatively spliced variant of the large isoform. d: The TC number is from TIGR’s EST database. e : Not submitted to GenBank. Table 3.3 Oligonucleotides primers for genes isolation

Potato Expected Gene name Primer name Primer sequence homolog Size (bp) ADP-glucose pyrophosphorylase NL-AGPS-F 5’-AGCAAAGACGTGATGTTAAACC-3’ AY186620 539 small subunit NL-AGPS-R 5’-TCTTCACATTGTCCCCTATACG-3’ NL-AGPSF-F1 5’-GCAATCACACTCTACCACACA-3’ ADP-glucose pyrophosphorylase NL-AGPS-F2 5’-CAAGCTTTGTTAACAATGGCG-3’ AY186620 1648 small subunit a NL-AGPSF-R 5’-TACCTCATGGCTGTTTAGCC-3’ NL-SU-F 5’-TATTTCGCCCAGGAAAATGTC-3’ Sucrose synthase M18754 1250 NL-SU-R 5’-TGAACGATGATCTCAGCTGGA-3’ NL-SUSYF-F1 5’-TTGTCTGAGGATTTCCCATCTG-3’ Sucrose synthase a NL-SUSYF-F2 5’-TTCCCATCTGCTGAATCAACTG-3’ M18754 2730

NL-SUSYF-R 5’-TTTTTTTTTGGACAAATTGAAA-3’ 86 NL-GB-F 5’-TCCCTGCTACCTAAAGTCGAT-3’ Granule-bound starch synthase TC125827 792 NL-GB-R 5’-CACAGATTGGCACTGTTCCAT-3’ NL-SS1-F 5’-TTTGTGTGTTACAAGTAGTGTTTC-3’ Starch synthase 1 Y10416 1009 NL-SS1-R 5’-CGACGGGTTCCAATCATTAA-3’ NL-SS2-F 5’-TGGCAAGCCTCAATGTAAAG-3’ Starch synthase 2 X87988 640 NL-SS2-R 5’-CACCACTGATACTTAGCAGCG-3’ NL-SS3-F 5’-CAAGCATGCCATTGCTTATG-3’ Starch synthase 3 X94400 797 NL-SS3-R 5’-CTTCCACCTTTCCAAACCAT-3’ NL-BE1-R 5’-TTCGATGCTGTATGTTCACCA-3’ Starch branching enzyme 1 X69805 556 NL-BE2-F 5’-ATGGCCAAACTCATTATCCAT-3’ NL-BE2-R 5’-ATGAGTTTATGACTTCAGAACA-3’ Starch branching enzyme 2 AJ011888 363 NL-BE-F 5’-TACTAGTGCATAGACCACTGCTG-3’ a : Full size cDNA To be continued Table 3.3 Oligonucleotides primers for genes isolation (continued)

Potato Expected Gene name Primer name Primer sequence homolog Size (bp) NL-ISO1-F 5’-GTGTAGATTTTGGGAGTATT-3’ Starch isoamylase 1 AY132996 793 NL-ISO1-R 5’-ACCCAATACAGAGTTATAACT-3’ NL-ISO2-F 5’-GCTTCATTTGCTTTATGCCTC-3’ Starch isoamylase 2 AY132997 1665 NL-ISO2-R 5’-CACACTCGTCTCCCATGTTAA-3’ NL-AAM-F 5’-TCAGCAGTCTGATCCATTGGT-3’ α-Amylase M79328 1083 NL-AAM-R 5’-TTTGGCTGTGCCTCAAGAAT-3’ a : Full size cDNA

87 Table 3.5 RT-PCR primers for ornamental tobacco starch metabolism genes expression detection

Tm Gene name Primer name Primer sequence Size (bp) (°C) ADP-glucose pyrophosphorylase RT-agps-F 5’-GATTTTAGCTTCTACGACCGATCAGC-3’ 58 125 small subunit RT-agps-R 5’-CAGTTCTTGATCACACAACCTTCACC-3’ 58 RT-susy1-F 5’-TACAGAGTTGTTCACGGCATTGAC-3’ 58 Sucrose synthase 1 189 RT-susy1-R 5’-CCTGTCCTTTAGCACACACAGATGTT-3’ 59 RT-susy2-F 5’-GTGTTCGATCCCAAATTCAACG-3’ 59 Sucrose synthase 2 156 RT-susy2-R 5’-TAGCACACACAGATGCTCCTCGT-3’ 59 RT-be1-F 5’-ATGGATCATCTTGTGAAGCGCA-3’ 59 Starch branching enzyme 1 135 RT-be1-R 5’-CATGACCAGTCTATTGTGGGTGACA-3’ 58 RT-be2-F 5’-CGAAGGTGAAATATTCGGCATTG-3’ 58 Starch branching enzyme 2 91 RT-be2-R 5’-CCTGGAAAATACAAGATTGTCTTGGAC-3’ 59 RT-gbss-F 5’-ATCTGAATGCCAAGGTCGCTTT-3’ 58 88 Granule-bound starch synthase 135 RT-gbss-R 5’-GCTTTTCATATCCATCAATGAAATCG-3’ 59 RT-ss1-F 5’-TGACCTCTCTGGAAAGGTTAAGTGC-3’ 59 Starch synthase 1 88 RT-ss1-R 5’-AAATCCAATCAGCGGACAATCG-3’ 59 RT-ss2-F 5’-ATGATGATCGGATGAGGCTTTACAG-3’ 58 Starch synthase 2 115 RT-ss2-R 5’-TCTTCTTTCTGCAATCTGTTGAAGTAGG-3’ 58 RT-ss3-F 5’-ATGGCACCTATTGCAAAGGTGG-3’ 58 Starch synthase 3 134 RT-ss3-R 5’-GTCCTTCACCTGATTCATCTTCAAAC-3’ 59 RT-iso1-F 5’-TTGGATCCTTATGCCAAGGTCA-3’ 58 Starch isoamylase 1 135 RT-iso1-R 5’-TAGGTCTCCTTCCCAGTCAAACTGAT-3’ 59 RT-iso2-F 5’-TCTTTGCCGTTTAAGACCTACGGT-3’ 59 Starch isoamylase 2 317 RT-iso2-R 5’-GCTCAGTTTTTCGGTAATGCCA-3’ 58

To be continued Table 3.5 RT-PCR primers for ornamental tobacco starch metabolism genes expression detection (continued)

Tm Gene name Primer name Primer sequence Size (bp) (°C) RT-iso3-F 5’-AACGAGATTACCTGGCTTGAGGA-3’ 58 Starch isoamylase 3 279 RT-iso3-R 5’-TGTTTTGCTTCCAGAAGAATAGCAG-3’ 59 RT-aaml-F 5’-ATATTGCAGGCCTTCAACTGGG-3’ 58 α-Amylase 147 RT-aaml-R 5’-TGGAAGGTAACCTTCAGGAGACAA-3’ 59 RT-bam-F 5’-TGAGCTATTGGAAATGGCGAAGA-3’ 59 β-Amylase 103 RT-bam-R 5’-AAGAGGGATCGTGCAGGAATCA-3’ 58 RT-R1-F 5’-AGAAATATATGTCGAGGTGGTCAGGG-3’ 59 Starch R1 enzyme 115 RT-R1-R 5’-TTGGGTAACCTAACACTTGAGGAGAGT-3’ 59 RT-pho-F 5’-AATCATGATGTGCAAGCGAAGA-3’ 58

Starch phosphorylase 142 89 RT-pho-R 5’-TGTAATCTGGCACAAATATTACCTTCAA-3’ 59 RT-18S-F 5’-TGGTCATGGCCGTTCTTAGT-3’ 59 18S rRNA 239 RT-18S-R 5’-TCGGCCAAGGCTATAAACTCGT-3’ 58 RT-26S-F 5’-TAAACGGCGGGAGTAACTATGACTCT-3’ 59 26S rRNA 235 RT-26S-R 5’-TTTCGGCTCCCACTTATACTACACCT-3’ 59 RT-gap2-F 5’-TTGAAGGGTGGTGCCAAGAAA-3’ 58 GAPDH 161 RT-gap2-R 5’-CATTTATGACCTTTGCCAAGGGAG-3’ 58

were cloned into plasmids and sequenced to identify partial cDNAs encoding the various starch metabolic genes. Using this approach between 2 and 6 independent copies of 13 additional starch metabolic genes were isolated (Table 3.2). In all cases except for alpha-amylase the independent copies of each gene had identical sequences with each other and a single copy was deposited in the GenBank. Two different isoforms of alpha-amylase were isolated, including AMY-L (DQ021455) and AMY-S (DQ021456). The AMY-S appears to be an alternative splicing product of the AMY-L (Supplemental Figure S3.1). The GenBank accession numbers for each gene are presented in Table 3.2. Sucrose synthase 1 (SuSy1, DQ021467) and sucrose synthase 2 (SuSy2, DQ021470), were identified in ornamental tobacco LxS8 (Table 3.3). One cDNA fragment encoding the AGPase small subunit (AGPS, DQ021458) was isolated. Despite repeated attempts, we were unable to isolate the AGPase large subunit genes. Each of these cDNAs showed very high identity with the potato homolog at both the nucleotide and amino acid level. In addition, cDNAs encoding a number of starch synthases, starch branching enzymes and starch debranching enzymes were identified. Because these starch metabolism genes exist as extensive gene families with conserved isoforms, we confirmed the identity of each isolated genes using the clustal W algorithm (http://align.genome.jp) to generate neighbor-joining trees that contained the isolated gene sequences with Arabidopsis, barley, rice, bean, pea, mung bean, potato, sorghum, wheat, and maize. Figure 3.2A shows a comparison of our four isolated clones with various starch synthase isoforms from the SWISS. This phylogenetic analysis shows that each of the previously identified starch synthase isoforms (SS1, SS2, SS3, and GBSS) clearly falls into independent clades and identifies each isoform. Clones encoding SS4 or SS5 were no identified. For each of these ornamental tobacco starch synthases clones, the most closely related species was Solanum tuberosum. Table 3.2 shows that the ornamental tobacco starch synthase isozymes share 92 to 94% nucleotide sequence identity and 91 to 95% amino acid identity with the corresponding potato reference genes. This high identity confirms the isoform specificity for each of the tobacco genes because this cross-species 91

Figure 3.2(A) Neighbor-joining trees for ornamental tobacco starch metabolism genes Protein sequences obtained from either GenBank or SWISS-PORT were used to generate neighbor-joining tree using a webpage-based Clustal W program (http://align.genome.jp). A Starch synthases. All protein sequences were from SWISS. SS1: Arabidopsis thaliana SS1: Q9FNF2; LxS8 SS1: DQ012453; Oryza sativa SS1: Q40739; Phaseolus vulgaris SS1: Q75T80; Solanum tuberosum SS1: P93568; Triticum aestivum SS1: Q43654; Zea mays SS1: O49064. SS2: LxS8 SS2: DQ012456; O. sativa SS2-1: Q7XE48; O. sativa SS2-2: Q6Z2T8; O. sativa SS2-3: Q5DWW9; P. vulgaris SS2: Q75T79; P. vulgaris SS2b: Q75T41; S. tuberosum SS2: Q43847; T. aestivum SS2-3: Q9LEE2; T. aestivum SS2-2: Q9LEE3; Z. mays SS2a: O48899; Z. mays SS2b: O48900. SS3: A. thaliana SS3: Q9SAA5; LxS8 SS3: DQ012455; O. sativa SS3: Q8W1P1; S. tuberosum SS3: Q43846; T. aestivum SS3: Q9LKW6. GBSS: A. thaliana GBSS: Q9MAQ0; LxS8 GBSS: DQ012457; O. sativa GBSS1: P19395; O. sativa GBSS2: Q8VYU1; P. vulgaris GBSS1: Q9XIS6; P. vulgaris GBSS1b: Q84LK2; S. tuberosum GBSS: Q43176; T. aestivum GBSS1: P27736; Z. mays GBSS: P04713.

A SS2 SS2 * SS2 SS2b

SS2-3 SS2 SS2-2 SS2a SS2-3 SS2b SS2-2 SS2-1 SS1 SS1 * SS1 SS1 SS1 SS1 SS1 SS1 GBSS1 GBSS GBSS1 GBSS GBSS GBSS * GBSS GBSS1b GBSS2 GBSS1 SS3 SS3 SS3 SS3 SS3 * SS3 93

Figure 3.2(B) Neighbor-joining trees for ornamental tobacco starch metabolism genes Protein sequences obtained from either GenBank or SWISS-PORT were used to generate neighbor-joining tree using a webpage-based Clustal W program (http://align.genome.jp). B Starch branching enzymes. All protein sequences were from GenBank. SBE1: Aegilops tauschii SBE1: AAD30187; LxS8 SBE1: DQ012459; Hordeum vulgare SBE1: AAP72268; O. sativa SBE1: BAA016161; Pisum sativum SBE1: CAA56320; P. vulgaris SBE1: BAA82349; Sorghum bicolor SBE: AAD50279; S. tuberosum SBE1: CAA70038; T. aestivum SBE1: AAG27622; Vigna radiata SBE1: AAT76445; Z. mays SBE1: AAC36471. SBE 2: A. tauschii SBE2b: AAW80632; A. thaliana SBE2: AAB03100; LxS8 SBE2: DQ012460; H. vulgare SBE2a: AAC69753; H. vulgare SBE2b: AAC69754; O. sativa SBE2: XP_466059; P. vulgaris SBE2: BAA82348; P. sativum SBE2: CAA56319; S. bicolor SBE2: AAP72267; S. tuberosum SBE2: CAB40747; T. aestivum SBE2a: AAK26822; T. aestivum SBE2b: AAW80631; V. radiata SBE2: AAT76444; Z. mays SBE2: AAC33764.

B SBE2b

SBE2a

SBE2b

SBE2*

SBE2

SBE2 SBE2 SBE2

SBE2a

SBE2b

SBE2

SBE1 *

SBE1

SBE1 SBE1 SBE1

SBE1

SBE1 95

Figure 3.2(C) Neighbor-joining trees for ornamental tobacco starch metabolism genes Protein sequences obtained from either GenBank or SWISS-PORT were used to generate neighbor-joining tree using a webpage-based Clustal W program (http://align.genome.jp). C Starch debranching enzymes. All protein sequences were from GenBank. ISA1: A. thaliana ISA1: NP_181522; H. vulgare ISA1: BAB72000; LxS8 ISA1: DQ021461; O. sativa ISA1: BAC75533; S. tuberosum ISA1: AAN15317; T. aestivum ISA1: AAL31015; Z. mays ISA1: AAB97167. ISA2: A. thaliana ISA2: NP_973751; H. vulgare ISA2: BAD08581; LxS8 ISA2: DQ021462; O. sativa ISA2: AAT93894; S. tuberosum ISA2: AAN15318; Z. mays ISA2: AAO17048. ISA3: A. thaliana ISA3: NP_192641; LxS8 ISA3: DQ021471; H. vulgare ISA3: BAD89532; O. sativa ISA3: NP_001063433; S. tuberosum ISA3: AAN15319; Z. mays ISA3: AAO17049. PU: A. thaliana PU: AAO00771; H. vulgare PU: AAF98802; O. sativa PU: BAA28632; T. aestivum PU: AAK52485; Z. mays PU: AAD11599.

C ISA3 ISA3

ISA3 ISA3 ISA3

ISA3 *

ISA3

ISA1 *

ISA1

ISA1 ISA1 ISA1

ISA1

ISA2 *

ISA2 ISA2 ISA2

ISA2

ISA2 ISA2

PU PU PU PU

PU 97

identity is significantly higher than the identity between potato isoforms. For example the SS1 gene show 91% amino acid identity with the ornamental tobacco SS1 cDNA, while the potato SS1 gene shows only 49%, 23% and 36% amino acid identity with the potato SS2, SS3 and GBSS gene (Table 3.4). Similar observation (Figure 3.2B) showed that the starch branching enzymes, SBE1 and SBE2 formed independent clades. Again, the high identity of the potato and tobacco clones confirmed the identity because the different potato isoforms showed only 48% protein identity (Table 3.4). Likewise, Figure 3.2C showed that each isoform of starch debranching enzyme (ISA1, ISA2, ISA3) also can be classified to the isoforms from other higher plant species, and again the identity among the potato isoforms (Table 3.4) is much lower than the potato/tobacco isoforms.

3.3.2 Regulation of starch biosynthesis There are three very important steps in starch biosynthesis. The first of these is catalyzed by sucrose synthase that serves to trap sugar in sink tissues by hydrolyzing sucrose into UDP-glucose and fructose (Martin et al., 1993; Munoz et al., 2005; Nolte and Koch, 1993; Zrenner et al., 1995). The second important step is catalyzed by ADP-glucose pyrophosphorylase that forms the committed substrate (AGP-glucose) for starch biosynthesis (Emes et al., 2003). The third important step is the actual structural formation of starch. This last step is catalyzed by starch synthases (SS), which polymerize linear glucan chains (James et al., 2003) and by starch branching enzymes (SBE), which introduce branched linkages into the starch structure and trim polyglucan chain into the actual starch structure (James et al., 2003). We investigated each of these steps independently. To investigate the expression of these starch metabolic genes, we designed primer pairs for each of the identified starch metabolic genes to perform RT-PCR (Table 3.5).

Sucrose synthase in developing ornamental tobacco floral nectary Since the floral nectary is a heterotrophic organ it is unable to photosynthesize, therefore, sugar for starch synthesis must be captured from the 98

Table 3.4 Comparison of Solanum tuberosum starch metabolic genes

Genes DNA Sequence Protein Sequence Similarity (%) a, b Similarity (%) b SS1 : SS2 46 40 SS1 : SS3 5 23 SS1 : GBSS 41 36 SS2 : SS3 13 23 SS2 : GBSS 41 34 SS3 : GBSS 5 23 SBE1 : SBE2 50 47 ISA1 : ISA2 7 28 ISA1 : ISA3 51 44 ISA2 : ISA3 20 29 a: Only the cDNA sequence in coding region was only for DNA sequence similarity calculation. b: The sequence similarity was calculated by using the webpage based (http://www.ebi.ac.uk/clustalw/) program. 99

phloem. While phloem does not penetrate the ornamental tobacco nectary (Horner et al., 2007), there are 23 phloem bundles that lie between the nectary and the gynoecium. Because of its importance in the capture of sugar, we evaluated the expression of sucrose synthase in the developing floral nectary using RT-PCR. Two different primer sets (RT-susy1-F/RT-susy1-R and RT-susy2-F/RT-susy2-R, Table 3.5), which can distinguish between SuSy1 and SuSy2 (data not shown), were used for the RT-PCR detection of these two genes. As shown in Figure 3.3A, both SuSy genes have similar patterns of gene expression. In all cases, our RT-PCR analyses used two standardization controls, 26S and 18S rRNAs. Oligonucloetides specific for the 26S and 18S rRNAs showed constant levels of expression in RNA preparations from all organs and confirmed the quality of our RNA and the loading of RNA in each reaction. Both sucrose synthase genes are expressed in all floral organs (Figure 3.3A and B). Analysis of both of these genes showed that the sucrose synthases were induced to a high level of expression at early stages but their expressions were decreased at later stages (Figure 3.3A). The mRNA expression level of SuSy1 appeared to be higher than SuSy2 during the floral nectary development; however the mRNA expression level of SuSy2 appeared to be higher than SuSy1 in leaves. We also examined the level of sucrose synthase protein present in the nectaries Western Blot analysis. Figure 3.3C shows that the sucrose synthase protein accumulated to very high levels at early stages of nectary development. This protein remained at moderate levels at later stages (Stage 6, Stage9, Stage 12, Stage PF). This level appeared to be same level as found in leaves. We therefore conclude that sucrose synthase is expressed most highly in early stage nectaries, but its expression persists throughout nectary development.

ADP-glucose pyrophosphorylase in developing floral nectaries A second important step in starch biosynthesis is AGPase because it catalyzes the first committed step in the synthesis of starch. Therefore, we examined AGPase small subunit (AGPS) in detail. To investigate AGPS gene expression in developing floral nectary, a pair of oligonucleotide primers (RT-AGPS-F and RT- 100

A SuSy1 SuSy2 18S 26S PF1211962 Nect ar y stag es

B SuSy1 SuSy2 18S 26S RSLPeOvStSePtFt Fl oral Organ s

C 2 6 911 12 PF Nectary stages 101

Figure 3.4 AGPase expressions in ornamental tobacco floral nectary A RT-PCR detection of AGPase small subunit. Five different stages (Stage 2, Stage 6, Stage 9, Stage 11, Stage 12, and Stage PF) were used to represent the whole floral nectary development. 26S and 18S rRNA were used as internal controls. B Immunoblotting of AGPase small subunit. Five different stages (Stage 3, Stage 6, Stage 9, Stage 12 and Stage post-fertilization) are used to represent the whole floral nectary development. Polyclonal antibody against potato AGPase small subunit was used for the immuno-blotting. C AGPase holoenzyme activity assay at different stages (Stage 2, Stage 6, Stage 9, and Stage 12) of nectary development. Three independent protein isolations were used to represent a specific stage. 200 µg of isolated total protein were used in each assay. Closed square: Nectaries, open square: ovaries.

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AGPS-R) was designed from the sequence of DQ021458. Like SuSy expression, the AGPS mRNA accumulates at early stages (Stage 2 and Stage 6), and decreases at later stages (Stage 9, Stage 11 and Stage 12) (Figure 3.4A). To test whether AGPS protein level is consistent with the AGPS mRNA level, western blotting using polyclonal antibody against potato AGPase small subunit was used. Western Blots of nectary proteins showed a pattern of expression of the AGPS protein that mirrored the AGPS mRNA expression pattern (Figure 3.4B), (i.e., the protein was most highly expressed at early developmental times and the level dropped at later times). We next examined the AGPase enzyme activity throughout floral development. These studies showed that nectary AGPase enzyme activity increased from Stage 2 to Stage 6 when the maximum level was obtained. Thereafter the level declined with 60% of AGPase enzyme activity remaining at Stage 9, but only 30% enzyme activity remaining at Stage 12 (Figure 3.4D). The level of AGPase enzyme activity in ovary throughout the floral development remains stable at the level of nectaries of Stage 12. This is further verified that nectary is a strong sink tissue requiring more AGPase enzyme activity to metabolize captured sugar through sucrose synthases.

3.3.3 Additional starch metabolic genes in developing floral nectaries The third major step in starch biosynthesis is the assembly of glucose moieties into amylase and amylopectin. Especially for amylopectin, this is a complex process requiring the action of many enzymes (Ball et al., 1996; Myers et al., 2000; Smith et al., 1997; Tetlow et al. 2004; James et al., 2003; Zeeman et al., 2002).

Starch synthases To investigate the gene expression pattern of starch synthases enzymes, we designed the RT-PCR primer pairs (Table 3.5) based on the four major starch synthases isoforms that were found in this study, SS1 (DQ021463), SS2 (DQ021466), SS3 (DQ021464), and GBSS (DQ021465). While all four isoforms 103

Figure 3.3 Sucrose synthases expression in ornamental tobacco Five different floral nectary stages (Stage 2, Stage 6, Stage 9, Stage 12, and Stage PF (post-fertilization)) were represented alive of inflorescence development. A RT- PCR detection of sucrose synthases gene expression in ornamental tobacco floral nectary. Primer pairs of RT-susy1-F and RT-susy1-R, and RT-susy2-F and RT- susy2-R were used for RT-PCR detection as of SuSy1 and SuSy2, respectively. Primer pairs used for internal controls as of 26S and 18S rRNA listed in Table 3.5. B RT-PCR detection of sucrose synthases gene expression in other ornamental tobacco organs. C Immunodetection of SuSy at different floral nectary stages (Stage 2, Stage 6, Stage 9, Stage 12 and Stage PF). Polyclonal antibody against maize sucrose synthase was used for the immunoblotting.

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A AGPs 18S 26S PF1211962 Nectary stages B AGPs 18S 26S RSLPeOvStSePtFt Floral Organs C 2 6 9 12 Nectary stages D 1.4 1.2 1.0 0.8 0.6 0.4 0.2 0 2 6 9 12 Nectary stages 105

were well expressed in leaves, these starch synthase isoforms were differentially expressed in the developing nectary (Figure 3.5A). SS1 and GBSS were expressed in all plant tissues at low levels throughout nectary development. Although well- expressed in leaves, SS2 was not expressed in the nectary, nor was it expressed in other floral organs (Figure 3.5A). In contrast, the most interesting of the starch synthases was SS3 that was highly regulated in developing nectary. It was expressed at early nectary stages but its expression dramatically declined as the nectary matured (Figure 3.5A), suggesting that SS3 may be important in the buildup of starch during nectary development.

Starch branching enzymes We isolated two starch branching enzymes (SBE1 and SBE2) in the course of this study. To investigate the expression in nectaries, primer sets derived from DQ021459 and DQ021460 were used for RT-PCR (Table 3.5). Although both starch branching enzymes were expressed throughout the developmental program (Figure 3.5B), their expression appears to differ. SBE2 is expressed at earlier time points and is down-regulated at later time points, while SBE1 is appears to be more highly expressed at later stages of nectary development. In the floral tube, petals and sepals the SBE2 appears to be more highly expressed than SBE1, while other organs showed similar levels of expression for both genes.

Starch debranching enzymes Our studies identified three debranching enzymes, ISA1, ISA2 and ISA3 in the nectaries. Despite repeated attempts, a pullulanase clone was not identified. RT- PCR showed that these three isoamylases were expressed differentially in developing floral nectaries (Figure 3.5C). ISA1 was universally expressed in plant, but it is highly expressed in late stages (Figure 3.5C). ISA2 and ISA3 showed similar expression patterns. They were highly expressed in middle stages of floral nectary development, while their expression could not be demonstrated in most of the other floral parts. 106

Figure 3.5 RT-PCR detection of starch metabolism related gene expression profile in ornamental tobacco Six different nectary stages (Stage 2, Stage 6, Stage 9, Stage 11, Stage 12 and Stage PF) and six different inflorescent tissues (floral tube, petal, sepal, stigma, ovary, and petiole) plus leaf, stem and root) were assayed for gene expression. Primer pairs used for each specific gene were listed in Table 3.5. 26S rRNA and 18S rRNA were used as internal controls. A RT-PCR detection of starch metabolic gene expression in ornamental tobacco floral nectary. Lane 1: Stage 2; lane 2: Stage 6; lane 3: Stage 9; lane 4: Stage 11; lane 5: Stage 12; lane 6: Stage postfertilization. B RT-PCR detection of starch metabolic gene expression in other organs. Lane 1: floral tube; lane 2: petal; lane 3: sepal; lane 4: stigma; lane 5: ovary; lane 6: petiole; lane 7: leaf; lane 8: stem; lane 9: root.

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A GBSS SS1 SS2 SS3 BE1 BE2 ISA1 ISA2 ISA3 18S 26S GAPDH PF1211962 Nectary stages B GBSS SS1 SS2 SS3 BE1 BE2 ISA1 ISA2 ISA3 18S 26S GAPDH RSLPeOvStSePtFt Floral Organs 108

3.3.4 Quantitative RT-PCR (qRT-PCR) expression Based upon these RT-PCR analyses, it appears that there are some starch metabolic genes that are expressed during early development (SuSy, AGPS, SS3, and SBE2) and others that are expressed later in nectary development (SBE1, ISA1), we decided to expand these analyses by examining these and other starch metabolism genes using a quantitative RT-PCR (qRT-PCR) method. In some cases, analysis was also performed to analyze the enzyme glyceraldehyde-3-phosphate dehydrogenase (GAPDH). As can been seen in Figure 3.5, GAPDH is invariant in the nectary throughout its development and is well expressed in other plant organs, although the petal, stigma, and root appear to show lower levels of expression. Also note that the level of expression of GAPDH is similar in foliage and nectaries. Because its expression was invariant through nectar development, we utilized glyceraldehydes 3-phosphate dehydrogenase (GAPDH) as a control for the temporal patterns of expression. To avoid contamination from nuclear DNA in qRT-PCR analyses, it is preferable that one or both of the qRT-PCR oligonucleotides should cross an intron junctions, therefore, oligonucleotides for the qRT-PCR procedure were designed by comparisons of our cDNA sequence with genomic sequences either from Arabidopsis, Rice, or other genomic sequences in the GenBank to identify the putative intron junction and in most cases two putative intron-exon junctions. The use of such intron-spanning oligonucleotides permits the specific amplification of signals from mRNAs even in the presence of genomic DNA. For each gene, the amplicon size was between 88 and 317bp. For all qRT-PCR assays, our oligonucleotide pairs amplified a single species, as evidenced by fluorescent spectra with a single melting temperature throughout the experiment (data not shown). All assays were performed in 96-well plates in duplicate on three independent RNA isolations as indicated in Materials and Methods. The cycle

threshold value (CT) was determined for each to provide a level of expression of each gene relative to the expression of GAPDH. These gene expression levels are shown in Figure 3.6 for 12 of the 18 starch metabolic genes. 109

In Figure 3.6A, the two isoforms of sucrose synthases are each very highly expressed at Stage 2 in nectary development. The level of expression for these genes, particularly SuSy1 is remarkable. By Stage 6, the level of expression of SuSy1 is 25 times the level of expression of GAPDH control. This level of expression is much higher than for any other nectary-expressed gene. Even at Stage 12, when the level of expression has significantly declined there is still 2 times the level of GAPDH expression. As a comparison examine the expression of SuSy1 and SuSy2 in leaves (Figure 3.6A) or the nectary-expression of starch branching enzyme 1 (Figure 3.6G) or starch phosphorylase (Figure 3.6H) which are expressed at a much lower level. In leaves, a source organ, SuSy1 and SuSy2 are expressed at 4% and 3% of leaf GAPDH, respectively, significantly lower than the 2500% found in the nectary. This very high level of SuSy1 and SuSy2 expression in nectaries should make this organ an especially strong sink in these ornamental tobacco flowers. The first committed step of starch biosynthesis (ADP-glucose pyrophosphorylase small subunit; AGPS) is also quite strongly expressed in the nectaries (2 to 4 times of GAPDH at Stage 2 and Stage 6); however, it is even more strongly expressed in leaves (3.5 times of GADPH at leaves) (Figure 3.6B). Because leaves are especially strong source tissue, this higher level of expression in leaves may be expected. In the nectary, AGPS expression decline during development and by Stage 9, when starch accumulation reaches its peak, AGPS mRNA is expressed at low levels (40% of GADPH at Stage 9). It remains low through anthesis (Stage 12; 20% of GADPH at Stage 12), and then increases following fertilization (3.6 times of GADPH). As can be seen in Figure 3.6B, AGPS mRNA level is 1.8 to 2.2 folds of GAPDH mRNA at early stages, and only 0.2~0.4 folds of GAPDH mRNA at late stages. While AGPS mRNA level in leaf was higher than nectary tissues, with 3.6 folds of GAPDH mRNA. Whether this level of expression indicates additional biological function is unclear. However, it is interesting to note that in some plants, following pollination, any residual nectar remaining in the nectary cup is reabsorbed. Presumably, this eliminates a rich source of sugars that could serve as a potential growth medium for infectious agents. Whether increased expression of AGPS 110

Figure 3.6 Quantitative RT-PCR detection of starch metabolism related gene expression in ornamental tobacco Three independent RNA isolations were performed to represent a specific stage. First strand cDNA were performed twice to represent an independent RNA isolation. Five different floral nectary stages (Stage 2, Stage 6, Stage 9, Stage 12, and Stage PF) were used to represent the whole inflorescence development. Expression level was normalized according to GAPDH expression level. A sucrose synthases (SuSy1 and SuSy2); B: AGPase small subunit; C starch synthases (SS1, SS3 and GBSS); D α-amylase (AMY); E β-amylase (BMY); F isoamylase 1 (ISA1); G starch branching enzyme 1 (SBE1); H starch phosphorylase (Pho); I R1 protein.

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40 AMY BE1 SuSy1 3.0 0.3 35 A SuSy2 D G 2.5 30 0.2 25 2.0 20 1.5 15 1.0 0.1 10 0.5 5 0 0 0 2 96 PF12 Leaf 2 96 PF12 Leaf 2 96 PF12 Leaf

PHO B AGPase small subunit 1.2 E H 4.0 2.0 1.0 BMY 3.0 0.8 1.6

2.0 0.6 1.2 0.4 0.8 1.0 0.2 0.4

0 0 0 2 96 PF12 Leaf 2 96 PF12 Leaf 2 96 PF12 Leaf

1.0 SS1 ISA1 C SS3 0.5 F 2.5 I R1 protein 0.8 GBSS 0.4 2.0 0.6 0.3 1.5 0.4 0.2 1.0 0.2 0.1 0.5 0 0 2 96 PF12 Leaf 0 2 96 PF12 Leaf 2 96 PF12 Leaf 112

following fertilization participates in this process is not clear, but is suggestive of that possibility. Figure 3.6C shows the patterns of expression of the three nectary-expressed starch synthases (SS1, SS3 and GBSS). SS2 expression was not detected in the nectary. As can be seen, the expression of SS1 is less than 10% of GAPDH throughout nectary development. At early stages of nectary development, its expression level (5% of GAPDH) is comparable to its expression level in foliage (5% of GAPDH). Subsequently, its expression declines as the nectary develops. In contrast, SS3 is 10 times more strongly expressed than SS1 at early developmental stages (Stage 2 and 6, 70% and 36% of GAPDH respectively) and is 5 times more strongly expressed in nectaries than in foliage (15% of GAPDH). This suggests that SS3 may be the starch synthases isoform that is most important in nectary starch biosynthesis. The level of expression of SS3 drops dramatically in later stages of nectary development (Stages 9 and 12, less than 0.3% of GAPDH both), indicating that expression of these starch biosynthetic enzymes is down regulated prior to the peak of starch accumulation. Together, the AGPS and SS3 and perhaps SuSy enzymes define a unique pattern of gene expression within the nectary that we term the anabolic pattern. These enzymes are strongly expressed at early stages of development and their expression is turned off at later stages of development. The switch between on and off occurs about or just prior to Stage 9 of floral development. The correlation of the expression of these genes with the previously observed filling of the nectary with starch (Ren et al., submitted) strongly suggests that the enzymes encoded by these genes are responsible for filling the nectary with starch. An alternative pattern of expression was observed with the genes that encode predicted starch catabolic enzymes. Figure 3.6D-F show the expression patterns of alpha-amylase, β-amylase and isoamylase 1 genes, respectively. Each of these genes encoding starch degradative enzymes is expressed either not at all or at low levels in early nectary development, but once starch catabolism begins at about Stage 9 (Ren et al., submitted), the expression of these genes is induced to quite 113

high levels (alpha-amylase increases 20 times (Figure 3.6D), β-amylase increases 25 times (Figure 3.6E), and isoamylase 1 (Figure 3.6F) increases 22 times when compared from Stage 12 to Stage 2). We term this latter pattern of nectary gene expression the catabolic pattern of expression. Again, the switch from the “off” stage to the “on” state is just prior to Stage 9. Finally, another set of genes were expressed throughout nectary development. Starch branching enzyme 1 (SBE1, Figure 3.6G) and starch phosphorylase (Pho, Figure 3.6H) are examples of genes that are not strongly regulated during nectary development and may be necessary for both starch synthesis and degradation. In addition, the R1 protein (glucan water dikinase) also shows the anabolic pattern of expression. The function of this enzyme is not well understood, however, it is generally thought to catalyze the addition of phosphate groups to starch and is consistent with its anabolic pattern of expression (Ritte et al., 2002; Reimann et al., 2002).

3.4 Discussion The ornamental tobacco floral nectaries accumulate high levels of starch from very early stages of development until approximately floral Stage 9 when nectary starch attains peak levels (Ren et al., submitted). Thereafter, the nectary starch is rapidly degraded to provide sugars for nectar production (Ren et al., submitted) as well as substrates for the biosynthesis of important nectary metabolites (Horner et al., 2007). The evidence for the accumulation of starch in nectaries is three-fold. We have previously used PAS staining of ornamental tobacco nectaries to identify the presence of carbohydrates storage forms inside nectary cells (Ren et al., submitted). We have also physically isolated and characterized the starch from ornamental tobacco nectaries (Ren et al., submitted). These studies showed that the nectary accumulates about 20% of its total mass as starch, that the nectary starch most complex at its Stage 9 peak and that the starch is rapidly degraded between floral Stage 9 and 12 (Horner et al., 2007; Ren et al., submitted). Third, in this study, we have examined the buildup and breakdown of amyloplasts in ornamental tobacco 114

nectaries using transmission electron microscopy (TEM). Those studies showed that amyloplasts accumulated in nectaries from Stage 2 flowers, through Stage 9 when the peak levels of amyloplasts were observed. Thereafter, the amyloplasts were emptied of starch but remained as ghosts within the nectary parenchyma cells. In these studies we also estimated the amount of starch present inside the cells. These studies (Table 3.1) estimated that little starch (1.7% of nectary cell volume) was present as starch at early (Stage 2) developmental stages. This number increases until peak accumulation occurred at Stage 9 when starch was estimated to represent 17% of nectary cell volume. This number is in excellent agreement with our earlier estimate that nectary starch represented 20% of total nectary mass (Ren et al., submitted). Based upon these three independent lines of investigation, it is clear that the ornamental tobacco floral nectary accumulates massive amounts of starch during its development and degrades this starch prior to anthesis. Because it is clear that starch metabolism is transient during nectary development, we have investigated starch metabolic gene expression in the nectary during this time frame. To accomplish this we first targeted genes that were known to be involved in starch metabolism in maize, potato, and Arabidopsis. This provided us with a list of 22 target genes (Table 3.1). We next isolated and characterized cDNAs encoding 18 of these 22 targeted genes. Bioinformatic analyses permitted us to identify each sub-family member for the starch synthases, the starch branching enzymes, and the starch debranching enzymes, with the exception that a pullulanase clone was not identified. We were also unable to identify pullulanase enzyme activity expressed in the ornamental tobacco floral nectaries (Klyne & Thornburg, unpublished observation). After identifying these starch metabolic genes, we used several methods to evaluate gene expression. We used RT-PCR to determine the pattern of expression of the starch biosynthetic genes. Some of these starch biosynthetic genes were expressed throughout nectary development; however, others were expressed only at early stages of development and not at maturity. 115

Because the levels of several genes appeared to vary significantly at different stages of nectary development, we developed quantitation (real-time) RT-PCR assays for 12 of the 18 starch metabolic genes, and we quantitated expression of these genes. These studies identified at least three different patterns of gene expression. The first of these patterns showed relatively high levels of gene expression at early developmental stages, but were turned off at later stages of development. We term this pattern the anabolic pattern of expression because several starch biosynthetic genes (AGPS, SS3, SS1) clearly showed this pattern of expression. The second of these patterns, the catabolic pattern of expression showed little or no expression in nectaries at early developmental stages, but showed high levels of expression at late nectary developmental stages (ISA1, alpha–amylase, β-amylase). The final distinct pattern of expression (constitutive expression), showed good expression levels at both early and late stages of development (Pho, SBE1). The expression of these starch anabolic genes and the starch catabolic genes not only defines unique patterns of gene expression, and require different regulatory schemes, but these patterns of expression also correlated with previously observed synthesis and degradation of starch in the developing floral nectary (Ren et al., submitted). Based on these observations, we conclude that starch biosynthesis and degradation in the nectary is most likely to be controlled at the transcriptional level. However, we have not excluded the possibility that fine-tuning of this process may be regulated by other mechanism known to regulate starch metabolism [redox potential (Geigenberger et al., 2005), or protein phosphorylation (Blennow et al., 2002)]. The regulation of these processes is clearly of great interest for the process of nectary function. The ornamental tobacco nectary has two primary functions in the plant. First it secretes nectar to serve as an attractant for pollinators [10] [Ren et al., submitted] and secondly, it serves to protect the gynoecium from microbes vectored to the flower by visiting pollinators (Horner et al., 2007, Thornburg et al., 2003). This second process is mediated by the Nectar Redox Cycle (Thornburg et al., 2003), a novel biochemical pathway that function to generate high 116

levels of hydrogen peroxide in soluble nectar via at least two different mechanism (Carter and Thornburg, 2003; Carter et al., 2007). In order to deal with these very high levels of hydrogen peroxide, the nectary produces at least two different antioxidants, ascorbate and β-carotene. Both of these compounds are produced from sugars in the nectary liberated following starch breakdown (Horner et al., 2007). Starch breakdown also provides sugars for nectar secretion (Ren et al., submitted). Indeed, the rapid breakdown of starch into free sugars appears to trigger a secondary flow of photosynthate into nectar that results in the accumulation of very high levels of nectar production in these plants (Ren et al., submitted). The processes that regulate starch metabolism in the nectary appear to be relatively simple. Starch biosynthetic genes are induced very early in floral development, while the flower is still at the bud stages. The most highly expressed genes are the sucrose synthase genes. Following transport of sucrose into cells, sucrose synthase cleaves the intracellular sucrose into fructose and UDP-glucose, thereby capturing sucrose from the phloem. The two sucrose synthase genes were found to be expressed at 25 and 10 fold the level of GAPDH, respectively. This is much higher than has been observed from any other nectary-expressed gene that we have investigated. This very high level of expression suggests that the nectary should be a strong sink tissue within the flower. This was confirmed in earlier studies that explored photosynthate capture by floral organs. Those studies demonstrated that the nectary was the strongest sink organ within the flower (Ren et al., submitted). Sucrose synthase western blots showed a similar pattern of expression with highest levels of activity found in the mature (Stage 2 and Stage 6) nectary. Another gene that was highly expressed was the ADP-Glucose pyrophosphorylase (AGPase) small subunit (AGPS). AGPase is the first committed step in starch metabolism, is a heterotetramer protein made up of two regulatory large subunits (AGPL) and two catalytic small subunits (AGPS) (Crevillen et al., 2005; Jin et al., 2005). Our analysis was limited to the small subunit of this enzyme because despite repeated attempts we were unable to isolate an AGPL clone. The inability to isolate an AGPL clone may suggest that the conserved sequences used 117

for potato preparation within the tobacco AGPL gene maybe more variable than in other Solanaceous genes. The AGPS gene showed the anabolic pattern of expression that is on at early developmental stages and is down regulated at later developmental stages. Western blotting analysis demonstrated that a similar pattern of expression was detected at the protein level for the AGPase small submit and likewise the activity analysis showed that AGPase activity was also highly expressed at the early stages of nectary development but was down-regulated at late stages of development. Together, the RNA, protein and activity analyses of the sucrose synthase and AGPase biosynthetic steps suggest that biosynthesis of starch is transcriptionally regulated in the ornamental tobacco floral nectary. We were able to draw a similar conclusion from the qRT-PCR analyses, that starch metabolism is largely controlled at the transcriptional level. These studies defined several clear pattern of gene regulation: an anabolic pattern, a catabolic pattern and a constitutive pattern. It is very interesting that the anabolic genes are down-regulated at the same time that the catabolic genes are up-regulated. Whether there is a molecular connection between these two processes is unknown, but the basis of this transcriptionally controlled metabolic switch is of particular interest. We are focusing on transcription factors that are expressed in the nectary during this timeframe. To date we have identified several MYB transcription factors whose expression is regulated in this timeframe. The involvement of these transcription factors in this process is still uncertain. Understanding the mechanism that control starch metabolism in the nectary manipulation of nectary metabolism can be used to achieve both higher accumulation of starch within the nectary as well as to increase the quantity and quality of nectar production in these flowers. It is well known that increased levels of sugar in nectar results in greater pollination or visitation to those flowers (Silva et al., 2000), and it is also known that high levels of pollinator visitation results in higher levels of seed set (Wallace et al., 1996; Roubik, 2002). The possibility of engineering floral gene expression for the purpose of attracting greater numbers of pollinating insects to increase seed set and ultimately yield of important fruit, nut, 118

and field crops is an intriguing outcome of this work. We are currently investigating the role of starch metabolic genes using RNAi and overexpression strategies to evaluate the effects of modifying nectary gene expression on nectar composition and quality. Such novel strategies of production will be required as the world’s population approaches 10 billion in the coming decades (Yan and Kerr, 2002). 119

Figure 3.7 Model of starch mass accumulation and starch metabolic enzyme activity in ornamental tobacco floral nectary A Simplified starch accumulation pattern and starch metabolic gene expression pattern in developing floral nectary. B Model of nectary starch anabolism. C Model of nectary starch catabolism.

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anabolic catabolic phase phase A D anabolic free sug ars gene expression

nect ary B st arch catabolic gene expression C

4 6 9 12 Stage of Floral Development

Sucrose Nectary Starch Anabolism B extracellular intracellular Sucrose UD P SuSy UD PG PPi ATP PPi Pi Fructose UG PS G1P AD PG AD P AG PS UT P HK PGM AD PG AD P UD P GBSS SSI/II/III F6P G6P SB E DBE PGI GWK Pho Starch (Gn) Starch (Gn+1) plastid Glycolysis

C Nectary Starch Catabolism -Amyl ase HK PGM UG Pase GG G + G Glucose G6P G1P UD PG DPE UT P 2 Pi ATP AD P PGI Maltose (GG) UD P (mtx) M altose F6P transporter S6P synthase Maltose S6P GWD /Pho Sucrose S6P DBE, ISA1,2,3 phosphatase -Amyl ase Starch (Gn) Sucrose plastid Invertase Nectar Fructose Glucose 121

Supplemental Figure S3.1 Comparison of splice-variant of α–amylase A Diagram of potato and ornamental tobacco α–amylase cDNA. PotAmy23: potato α–amylase 23. LxS8 AAM-L: α–amylase large fragment. LxS8 AAM-S: α–amylase small fragment. B Comparison of α–amylase cDNA sequence between potato and ornamental tobacco AAM-L and AAM-S. C Left border of α–amylase RNA-splicing site. D Right border of α–amylase RNA alternative splicing site.

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A PotAMY23

LxS 8 AAM -L

LxS 8 AAM -S

B LxS8 AAM-L ------QQSDPLVVIRNGKEIILQAFNWESHKHDWWRNLDRKVPDIAKSGFTTVWLPPAS LxS8 AAM-s ------QQSDPLVVIRNGKKVILQAFNWESHKHDWWRNLDRKVPDIAKS------Pot AMY23 MALDESQQSDPLVVIRNGKEIILQAFDWESHKHDWWLNLDTKVPDIAKSGFTTAWLPPVC *************::*****:********* *** ******** LxS8 AAM-L QSLSPEGYLPQNLYSLNSSYGSEHLLKALLNKMKQYKVRAMADIVINHRVGTTQGHGGMY LxS8 AAM-s ------Pot AMY23 QSLAPEGYLPQNLYSLNSKYGSEDLLKALLNKMKQYKVRAMADIVINHRVGTTQGHGGMY

LxS8 AAM-L NRYDGIPLSWDEHAVTSCTGGRGNKSTGDNFNGVPNIDHTQSFVRRDLTDWMRWLRSSVG LxS8 AAM-s ------G Pot AMY23 NRYDGIPMSWDEHAITSCTGGRGNKSTGDNFNGVPNIDHTQSFVRKDLIDWMRWLRSSVG *

LxS8 AAM-L FQDFRFDFAKGYSPKYVKEYIEGAKPIFSVGEYWDTCNYKGSYLDCNQDSHRQRIINWID LxS8 AAM-s FQDFRFDFAKGYSPKYVKEYIEGAKPIFSVGEYWDTCNYKGSYLDCNQDSHRQRIINWID Pot AMY23 FQDFRFDFAKGYASKYVKEYIEGAEPIFAVGEYWDTCNYKGSNLDYNQDSHRQRIINWID ************:.**********:***:************* ** ************** LxS8 AAM-L QTGQLSSAFDFTTKAILQEAVKGEFWRLRDSKGKPPGVLGWWPSRAVTFIDNHDTGSTQA LxS8 AAM-s QTGQLSSAFDFTTKAILQEAVKGEFWRLRDSKGKPPGVLGWWPSRAVTFIDNHDTGSTQA Pot AMY23 GAGQLSTAFDFTTKAVLQEAVKGEFWRLRDSKGKPPGVLGLWPSRAVTFIDNHDTGSTQA :****:********:************************ ******************* LxS8 AAM-L HWPFPSNHIMEGYAYILTHPGIPSVFYDHFYDWGNSTHDQIVKLIDIRRHQGIHSRSSVQ LxS8 AAM-s HWPFPSNHIMEGYAYILTHPGIPSVFYDHFYDWGNSTHDQIVKLIDIRRHQGIHSRSSVQ Pot AMY23 HWPFPSRHVMEGYAYILTHPGIPSVFFDHFYEWDNSMHDQIVKLIAIRRNQGIHSRSSIR ******.*:*****************:****:*.** ******** ***:********::

LxS8 AAM-L ILEAQP------LxS8 AAM-s ILEAQP------Pot AMY23 ILEAQPNLYAATIDEKVSVKIGDGSWSPAGKEWTLATSGHRYAVWQK ******

C 38ProAspIleAlaLysSerG lyPheThrThrValTrp 49 LxS8 AAM-s GATAGAAAAGTTCCTGATATTGCAAAATCTG ------LxS8 AAM-L GATAGAAAAGTTCCTGATATTGCAAAATCTG^GCTTCACAACAGTATGGTT 150 *******************************

D 168TrpLeuArgSerSerValG lyPheGlnAspPheArgPheAspPhe 182 LxS8 AAM-s ------GCTTCCAAGATTTTCGTTTTGATTTTGCT LxS8 AAM-L GGTGGCTAAGATCCTCTGTCG^GCTTCCAAGATTTTCGTTTCGATTTTGCT 550 ******************* ********* 123

Supplemental Table S3.1 Target genes involved in starch metabolism genes in Solanum tuberosum and ornamental tobacco LxS8

Potato Homolog in Number Gene Name GenBank 1 ADP-glucose pyrophosphorylase small subunit AY186620 2 ADP-glucose pyrophosphorylase large subunit 1 X61187 3 ADP-glucose pyrophosphorylase large subunit 2 X74982 4 Sucrose synthase 1 M18745 5 Granule-bound starch synthase TC125827 a 6 Starch synthase 1 Y10416 7 Starch synthase 2 X87988 8 Starch synthase 3 X94400 9 Starch synthase 4 TC159092 a 10 Starch synthase 5 TC153222 a 11 Starch branching enzyme 1 X89605 12 Starch branching enzyme 2 AJ011888 13 Starch isoamylase 1 AY132996 14 Starch isoamylase 2 AY132997 15 Starch isoamylase 3 AY132998 16 Pullulanase TC137237 a 17 Disproportionating enzyme X68664 18 Alpha-amylase M79328 19 Beta-amylase TC118938 a 20 Alpha-glucosidase AJ001374 21 Starch R1 enzyme AY027522 22 Starch phosphorylase X52385

a: Sequence information is not found in GenBank. The TC number is from TIGR. 124

Table S3.2 Volumes of cell, plastid within cell, and starch granule within plastid in developing ornamental tobacco floral nectary cells

Average Cell Average Total Average Total Stage/ Area in µm2 Plastid Area in Starch Area in Cell or structure Area Ave Cell µm2 per Cell µm2 per Cell Volume Measured S (µm2) 193 27.9 13.0 S2 Calculated r (µm) 7.84 3.0 2.0 Calculated V (µm3) 2017.5 113.0 33.5 Measured S (µm2) 188 30.8 16.9 S4 Calculated r (µm) 7.74 3.1 2.3 Calculated V (µm3) 1942.3 124.7 50.9 Measured S (µm2) 225 42.9 31.0 S6 Calculated r (µm) 8.46 3.7 3.1 Calculated V (µm3) 2536.3 212.1 124.7 Measured S (µm2) 334 121.5 100.7 S9 Calculated r (µm) 10.3 6.2 5.7 Calculated V (µm3) 4572.2 997.8 775.3

Volume of sphere (V) =4/3πr3; Area of Circle (S) = πr2; r= radius; π=3.14.

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Chapter 4 β-carotene metabolism in ornamental tobacco floral nectary

Amyloplast to chromoplast conversion in developing ornamental tobacco floral nectaries provides sugar for nectar and antioxidants for protection

This section is based on a published paper in American Journal of Botany (Horner HT, Healy RA, Ren G, Fritz D, Klyne A, Seames C, and Thornburg RW. Amyloplast to chromoplast conversion in developing ornamental tobacco floral nectaries provides sugar for nectar and antioxidants for protection. Am. J. Bot. 2007 Jan; 94(1):1224).

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Contribution of this work: The authors thank the USDA SCA #58-3625-3-104 for partial support of this project to H.T.H., the National Science Foundation Grant #IBN-0235645 to R.W.T., and the Microscopy and NanoImaging Facility and its staff for use of its facilities under the directorship of H.T.H. Mr. Gang Ren performed all the molecular biological experiments, including Table 4.1, 4.2, 4.3 and Figure 4.34 and 4.38, Ms. Rosanne Healy performed all of the Microscopic experiment, including Figure 4.1-4.18, Figure 4.19-4.24, Figure 4.25-4.31. Ms. Anna Klyne performed TLC experiment (Figure 4.33). Dr. Harry Horner consulted on the methods of Microscopic sections of this manuscript. Dr. Robert Thornburg consulted on the overall creative instructions and support for this work. In addition he contributed to the writing, reviewing and editing of this work. 127

Abstract Tobacco floral nectaries undergo changes in form and function. As nectaries change from green to orange, a new pigment is expressed. Analysis demonstrated that it is β-carotene. Plastids undergo dramatic changes. Early in nectary development, they divide and by stage 9 (S9) they are engorged with starch. About S9, nectaries shift from quiescent anabolism to active catabolism resulting in starch breakdown and production of nectar sugars. Starch is replaced by osmiophilic bodies, which contain needle-like carotenoid crystals. Between S9 and S12, amyloplasts are converted to chromoplasts. Changes in carotenoids and ascorbate were assayed and are expressed at low levels early in development; however, following S9 metabolic shift, syntheses of β-carotene and ascorbate greatly increase in advance of expression of nectar redox cycle. Transcript analysis for carotenoid and ascorbate biosynthetic pathways showed that these genes are significantly expressed at S6, prior to the S9 metabolic shift. Thus, formation of antioxidants β- carotene and ascorbate after the metabolic shift is independent of transcriptional regulation. We propose that biosynthesis of these antioxidants is governed by availability of substrate molecules that arise from starch breakdown. These processes and events may be amenable to molecular manipulation to provide a better system for insect attraction, cross pollination, and hybridization.

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4.1 Introduction The changes in plastid ultrastructure and function during development have been studied in a variety of organs and plants. These changes typically follow various pathways, which may lead to formation of amyloplasts (Bechtel and Wilson, 2003), and in the cases of petal and fruit development, into chromoplasts (Pyke, 1999; Camara et al., 1995; Pyke and Page, 1998; Weston and Pyke, 1999; Pyke and Howells, 2002). These changes may also occur in certain roots and underground stems without passing through the chloroplast stage. The switchover from plastids containing chlorophylls to those containing carotenoids (Cheung et al., 1993; Fraser et al., 1994; Ronen et al., 2000; Busch et al., 2002; Castillo et al., 2005) is related to ripening and senescence. Even though this process has been documented many times, some systems may have unique aspects that warrant further study. This is the case for tobacco floral nectaries where large, engorged amyloplasts are initially converted into intermediate plastids—amylochromoplasts (plastids containing both starch and β-carotene crystals)—and finally to chromoplasts. In addition, the catabolic and anabolic processes associated with these conversions define a complex and well-coordinated series of biochemical reactions that involve nectar production (Pacini et al., 2003; Carter et al., 2006) and nectar, nectary, and gynoecium protection. Plastids are dynamic and versatile double-membrane-bounded organelles that are classified into at least six distinct categories—chloroplasts, leucoplasts, elaioplasts, amyloplasts, chromoplasts, and undifferentiated plastids or proplastids (Waters and Pyke, 2004). They have a range of sizes, shapes, and numbers per type of cell. They are related to the age of a cell and its developmental state (Osteryoung and McAndrew, 2001). Plastids seem to be well suited to adapting to the complexity and function of the changing cell in which they exist. Even though both extrafloral and floral nectaries have been variously studied (O'Brien et al., 1996; Horner et al., 2003; Pacini et al., 2003; Stpiczynska et al., 2004, 2005), only cursory attention has been directed to the changes in the plastids. Therefore, the developing tobacco floral nectary provides an interesting system in which to study these 129

changes as part of a gland whose dynamics are associated with starch synthesis and catabolism, nectar and hydrogen peroxide production, and antioxidant (β- carotene and ascorbate) synthesis—a series of integrated processes reported together in this study for the first time. The purpose of this study is to follow the developmental changes in the plastids up to maturity and beyond, to relate both associated biochemical and molecular processes with these changes, and to understand the underlying mechanisms that control them. This combined information will serve as the basis for developing strategies to manipulate the components comprising the nectar, which in turn will have the potential for increasing insect visitation and ultimately cross-pollination for hybridization and crop improvement.

4.2 Material and methods 4.2.1 Plant lines Ornamental tobacco plants that overproduce nectar [from interspecific cross of Nicotiana langsdorffii x N. sanderae Hort. Var Sutton's Scarlet line LxS8 (Kornaga et al., 1997)] were grown in a temperature-controlled glasshouse on the Iowa State University campus under a photoperiod regimen that allowed for continuous flowering. Most plants provided an array of flowers at different stages of development that can be identified by stage (stages 2, 4, 6, 9, 10, 11, and 12; Figure 4.1--7) per Koltunow et al. (1990). The flowers were processed for both light and electron microscopy, and biochemically analyzed for starch, carotenoids, and ascorbic acid according to the following methods.

4.2.2 Light microscopy For general observations requiring 1-µm-thick sections, whole gynoecia with nectaries or isolated nectaries were fixed in 4% paraformaldehyde in a 0.1 M sodium cacodylate buffer, pH 6.8, at 4°C for 3 h, washed in cold buffer and deionized water, dehydrated in an ethanol series to 100% ethanol, and infiltrated in a 3 : 1, 1 : 1, and 1 : 3 mixture of pure ethanol : LR White Resin (http://www.emsdiasum.com). Infiltration was followed by pure resin, specimens were embedded in resin in 130

covered aluminum trays to eliminate oxygen, and they were polymerized at 55°C for 24 h. Sections were cut on a Leica-Reichert Ultracut S ultramicrotome (http://www.leica-microsystems.com) using glass knives and mounted on Probe-On Plus slides (http://www.fisherscientific.com). Some sections were treated with the periodic acid Schiff (PAS) Technique to localize non-water soluble polysaccharides (Ruzin, 1999).

4.2.3 Vibratome sections Fresh floral gynoecia at S9 through S12 were glued in two orientations (trans- and longi-planes through gynoecia and nectaries) to the base reservoir with a fast- drying epoxy glue. Deionized water was added just above the top of each specimen, and 30-µm-thick sections were cut and placed in drops of deionized water on slides. Cover slips were added and sections were viewed either in bright-field (white light [WL]) mode or between crossed polarizers (CP), and digitally captured using a Zeiss MRc Axiocam camera (http://www.zeiss.de) mounted on an Olympus BH10 microscope (http://www.olympus-global.com).

4.2.4 Electron microscopy For general ultrastructure, small portions of fresh, isolated nectaries were initially fixed in 2% glutaraldehyde and 2% paraformaldehyde in a 0.1 M sodium cacodylate buffer, pH 7.2, at 4°C overnight, washed in cold buffer, postfixed in 1% osmium tetroxide in same buffer at 4°C for 1.5 h, washed with buffer and deionized water, and en bloc stained with 2% aqueous uranyl acetate in the dark. The specimens were washed in deionized water. Then the specimens were divided into two batches for further processing into either Quetol or Spurr's resin mixtures. For Quetol resin embedding (manufacturer's instructions; http://www.polysciences.com) after deionized water washes, specimens were processed through increasing concentrations of Quetol and decreasing ratios of deionized water to pure Quetol monomer, and then into Quetol resin mixture (Quetol 651, 15 g; methyl nadic anhydride [MNA], 10 g; nonyl succinic anhydride [NSA], 20 g; and 131

tridimethylaminomethyl phenol [DMP-30], 1 g), followed by casting in aluminum trays. Polymerization was done at 75°C for 1 d. For Spurr resin embedding (Spurr, 1969) following deionized water washes, specimens were dehydrated in an ethanol series, transferred to pure acetone, and infiltrated in acetone and Spurr's hard resin mixture. Specimens were then transferred into pure resin mixture, cast in aluminum trays, and polymerized at 70°C. Specimens embedded in both resin mixtures were sectioned using either glass or diamond knives. Thin 60-nm-thick sections were mounted on Formvar-coated slotted grids and stained with 2% aqueous uranyl acetate and lead citrate (Reynolds, 1963). Observations were made on a JEOL 1200EX TEM (www.jeol.com, and images were captured photographically on Kodak SO-163 film (www.kodak.com), which was later digitized for further processing using an Epson Perfection 3200 PhotoScanner (http://www.epson.com).

4.2.5 Biochemical analyses Carotenoids analysis Nectaries were harvested from flowers at different stages (Koltunow et al., 1990) as previously described (Carter and Thornburg, 2000). Approximately 10 nectaries were combined in a glass tissue homogenizer with a total volume of 300 µl acetone. After homogenization, the organic layer was removed, and the process was repeated, first with 300 µl acetone, then with 300 µl hexane. The three organic layers were combined, dehydrated with sodium sulfate, and evaporated to dryness, taken up in 50 µl hexane and used for analysis. Thin layer chromatography (TLC) was performed on silica gel plates using a 9 : 1 hexane : acetone solvent. HPLC was performed exactly as described (Barua and Olson, 1998) on a Waters HPLC system (http://www.waters.com) using a 3-mm Microsorb-MV column (http://www.rainin- global.com). Carotenoids were estimated at 450 nm.

Ascorbic acid analysis Nectaries were harvested from staged flowers as previously described. Dissected nectaries from approximately 30 flowers were homogenized in an equal 132

volume of 1% oxalic acid in a Con-Torque tissue homogenizer (http://www.eberbachlabtools.com). Following homogenization, the samples were centrifuged (13 000 x g, 5 min) to remove debris. The pellet was re-extracted with an additional volume of 1% oxalic acid and reprocessed. The supernatants were combined and a 50 µl sample in a total volume of 2 ml of 1% oxalic acid was titrated to a pink endpoint with 0.05% 2,6-dichlorophenolindophenol (DCIP) in 0.1 M phosphate buffer, pH 7.0. A standard curve of the sample containing up to 20 µg of ascorbate was prepared to quantitate levels of ascorbate.

4.2.6 Labeling of carotenoids with [14C]-sucrose Flower buds at S9 were harvested and placed with pedicels in water at room temperature. Subsequently 0.8 µCi [14C]-sucrose (per flower) was added to the water. Incubation continued for 48 h with additional water added when necessary. Nectaries (three per sample, duplicate samples) were extracted 2x with acetone and once with hexane. The combined organic phases were dried in a rotary evaporator, resuspended in 2 : 1 : 1 acetone : hexane : acetonitrile, and a 10 µL sample was counted by scintillation. The thin layer chromatograms with 10 µL per sample (Kodak 13181 Silica Gel; http://www.kodak.com) were developed in 9 : 1 hexane : acetone. Autoradiograms were prepared by exposing each chromatogram to a phosphorimager plate overnight.

4.2.7 Oligonucleotides for RT-PCR analysis For synthesis of the oligonucleotides for RT-PCR analysis, the ornamental tobacco cDNA clones were mapped onto the Arabidopsis thaliana genomic clones encoding the same genes. This permitted the localization of the Arabidopsis introns in the tobacco cDNA sequences. Oligonucleotides were then synthesized to cDNA regions that spanned the hypothetical intron junctions. All oligonucleotides were first tested with genomic DNA to insure that the oligonucleotides did not amplify a fragment with genomic DNA. The oligonucleotides used in this analysis are listed in 133

Table 4.1. Control oligonucleotides were also prepared for the 18S and 26S rRNA genes.

4.3 Results In ornamental tobacco, the nectary is embedded in the base of the gynoecium. It encircles the base and has a continuous cuticle over its entire surface (Figure 4.1--7). It has two stomatal zones 180° in line with the internal septum (Figure 4.8; arrowheads; Thornburg et al., 2003). All the vascular tissues (veins) emanating from the pedicel below the nectary pass through the gynoecial walls next to the locules and interior to the nectary but do not penetrate the nectary special parenchyma anywhere (Figure 4.8; asterisks). Beginning at S2, the special parenchyma accumulates starch. We have begun a series of studies to evaluate development of the floral nectary. We have used macromicroscopy of gynoecia and nectaries throughout nectary development to identify developmental changes in this floral organ. Between S2 and S9 both the gynoecium and nectary increase in size (G. Ren et al., unpublished data). In addition, the nectary color changes from lime green (S2 and S4; Figure 4.1, 2) to yellow (S6; Figure 4.3) to a light orange (S9; Figure 4.4). Between stages S10 and S12, the nectary matures and becomes a deep, bright orange (Figure 4.5--7). Nectar secretion is initiated at late S10 or early S11 and continues beyond S12.

4.3.1 Light microscopy: S9 through S12 Previous studies indicate that approximately 20% of the nectary mass is composed of starch (G. Ren et al., unpublished data) by S9, when it is engorged with starch (Figure 4.9, 10, 15, 19). Beginning at S2, the special parenchyma accumulates starch. Starch continues to accumulate through S9 when the nectary changes greatly and the accumulated starch is degraded to produce sugars for nectar production. To better evaluate these changes during this maturation phase of nectary development, we prepared vibratome longitudinal and cross sections Table 4.1 Oligonucleotides for RT-PCR analysis

Gene Oligo name Primer sequence Tm Amplicon size (bp) RT-IPI-F 5’-TCTTCAGCAACGATCAGCAACA-3’ 58 ipi 130 RT-IPI-R 5’-TTGTGCAGCATTTCTTACCCCA-3’ 59 RT-FPS-F 5’-TGGTACAAACTACCTAAGGTTGGCA-3’ 58 fps 152 RT-FPS-R 5’-GCAGTCTGGAACTCAACCTCATTAAA-3’ 58 RT-GGPS-F 5’-GTTGCTGATAAGGTAACTTATCCCAAAC-3’ 58 gps 225 RT-GGPS-R 5’-CAGATCACTCATGTTTTCACCTTCTATT-3’ 57 RT-PYS-F 5’-CTGTGTCATCAGAGAAAAAGGTGTATG-3’ 57 pys 252

RT-PYS-R 5’-TGCACCACACATATATTGCCCA-3’ 58 134 RT-CISO-F 5’-AGCATAGAAGATTGGGAGGGACTCT-3’ 58 ciso 305 RT-CISO-R 5’-TTGGTGTCCCCACCTCCTTAAA-3’ 59 RT-PDS-F 5’-ATTACGAGTTACTTCTTGGCCGGA-3’ 59 pds 83 RT-PDS-R 5’-GAAGCAACATTTTAGTTCACCATGC-3’ 58 RT-ZDS-F 5’-AGTGTCAAAGCAGGTTTTGGCA-3’ 58 zds 188 RT-ZDS-R 5’-CGATGTAGTCCTGTTTTGTATATGAGCC-3’ 59

To be continued

Table 4.1 Oligonucleotides for RT-PCR analysis (continued)

Gene Oligo name Primer sequence Tm Amplicon size (bp) RT-LYC-F 5’-CATCCCTCAACAGGTTATATGGTAGC-3’ 58 lyc 225 RT-LYC-R 5’-AAACCTTCTCGTAGCGGGTAAATC-3’ 58 RT-CCD-F 5’-ACCCGAAGGTTGACCCTGTAACT-3’ 58 ccd 224 RT-CCD-R 5’-TTCACCATTTCCTTTGGTTTGAAG-3’ 58 RT-NEO-F 5’-GACCATTTGAATGTGAAAGGTTGG-3’ 58 neo 158 RT-NEO-R 5’-CTTCCATTTCTTCTTTTACCTTTTCACT-3’ 57 RT-GPI-F 5’-TTGGACAGTTACTGGCCATCTATGA-3’ 58 gpi 235

RT-GPI-R 5’-GAAGGATCAGAAGGAACATCAGCA-3’ 58 135 RT-GDH-F 5’-TATCGGAAAAGGTACTAGGGAAGGCT-3’ 59 gdh 228 RT-GDH-R 5’-GCGTCTCATTCACAATCTGATCAAG-3’ 58 RT-GLDH-F 5’-TGTCGGGTCAGGTTTGTCTCC-3’ 58 gldh 235 RT-GLDH-R 5’-ATGAGCACCAACCTGGACAATG-3’ 58 26S rRNA RT-26S-F 5’-TAAACGGCGGGAGTAACTATGACTCT-3’ 59 235 control RT-26S-R 5’-TTTCGGCTCCCACTTATACTACACCT-3’ 59 18S rRNA RT-18S-F 5’-TGGTGCATGGCCGTTCTTAGT-3’ 59 239 control RT-18S-R 5’-TCGGCCAAGGCTATAAACTCGT-3’ 58

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Figs. 4.1–18. Images of isolated whole (Figs. 4.1--7) and 30-µm-thick cross (Figs. 4.8, 4.15--18) and longitudinal (Figs. 4.9--14) vibratome sections of gynoecia (G) and basal nectaries (N) from stage 2 (S2) through stage 12 (S12) observed with white light (WL) or between crossed polarizers (CP)

1. S2 with basal green nectary. 2. S4 with basal lime-green nectary. 3. S6 with yellow-green nectary. 4. S9 with basal orange nectary. 5. S10 with basal orange nectary. 6. S11 with basal orange nectary. 7. S12 with bright orange nectary. 8. S12 vibratome cross section through nectary showing relationship of vascular bundles (asterisks) interior to nectary and locule. Arrows indicate stomatal pore regions where nectar is secreted. 9. S9 longitudinal WL view through gynoecium, one locule and whitish basal nectary. 10. Same as Fig. 4.8, CP view of starch-filled nectary and calcium oxalate crystal layer (arrows) in gynoecium wall and in septum. 11. S11 longitudinal WL view through gynoecium, one locule and orange basal nectary. 12. Same as Fig. 4.11, CP view of nectary with both starch and carotenoids; oxalate crystals are prominent. 13. S12 longitudinal WL view through gynoecium, one locule, and very orange nectary. 14. Same as Fig. 4.13, CP view showing differentially colored, bright orange nectary. 15. S9 vibratome section through nectary showing starch-engorged cells, CP. 16. S10 vibratome section through nectary and its epidermis showing mixture of starch-filled and carotenoid-containing cells, CP. 17. S11 vibrarome section through nectary and epidermis showing fewer cells with starch grains and more cells with carotenoids, CP. 18. S12 vibratome section through nectary showing cells engorged with multicolored ß-carotene crystals, CP. Bar: 1 mm, Figs. 4.1--14; 50 µm, Fig. 4.15; 200 µm, Fig. 4.16; 100 µm, Figs. 4.17, 18.

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through the gynoecium and nectary. These showed that the nectary color changes from the inner part of the nectary toward the single-layered epidermis during S10 and S12 (Figure 4.11--14). At S9, the end of the nectary starch filling period, each special parenchyma cell contains plastids that in turn contain multiple starch granules. The 30-µm-thick cross and longitudinal sections through the nectary at S9 through S12 showed a progression of completely starch-filled special parenchyma, as shown between crossed polarizers, to special parenchyma that progresses from whitish to an orange color from the inside of the nectary near the vascular bundles to the outside of the nectary near the epidermis (Figure 4.15–18). At high magnification of these regions S9 plastids display multiple starch grains (Figure 4.15) that transition during S10 (Figure 4.16) and S11 (Figure 4.17) to plastids with multicolored β-carotene crystals that give an overall orange appearance to the entire nectary at S12 (Figure 4.7, 13, 18). The transition of starch-filled to β-carotene-filled plastids occurs over a period of 36 h. At S12, there are still a few plastids that contain small amounts of starch scattered throughout the special parenchyma (Figure 4.20; see next section). The nectary remains orange, and the special parenchyma cells stay intact for up to about 48 h beyond S12, if the flower has not been pollinated.

4.3.2 Electron microscopy: S9 through S12 S9 plastids The enlarged S9 plastids are uniformly filled with multiple starch grains and have a minimum of lamellae and stroma (Figure 4.21, 25). There are very few, small osmiophilic bodies visible in the stroma. There is an average of 8.5 plastids per special parenchyma cell area with each plastid volume averaging about 1000 µm3. Each plastid volume consists of about 78% starch, so that at S9 the ratio of cell volume to starch volume is 5.9 : 1. This figure corresponds well with the previously estimated quantification of starch in nectaries of 20% of total nectary mass, which represents starch (G. Ren et al., unpublished data). 139

Figs. 4.19–24. Light (Figs. 4.19, 20) and transmission electron microscope (Figs. 4.21--24) sections through nectary at stages 9–12

19. S9 periodic acid-Schiff (PAS)-stained LR White section through nectary showing magenta cell walls and multiple starch grains per cell. Clear cell centers represent unstained nuclei. 20. S12 PAS-stained LR White section through nectary showing magenta-colored cell walls with scattered cells showing few starch grains. 21. S9 nectary cell engorged with plastids containing multiple starch grains and dense lamellae. Note small, relatively few cytoplasmic vacuoles and large central nucleus. 22. S10 nectary cell with engorged plastids containing multiple starch grains, more osmiophilic bodies, and more small cytoplasmic vacuoles. 23. S11 nectary cell with enlarged plastids, some of which appear to be partially degenerated, more osmiophilic bodies, and fewer large cytoplasmic vacuoles. 24. S12 nectary cell with plastids with no starch grains but with many osmiophilic bodies and dense, wavy-appearing crystals and crystal profiles. Vacuoles are large and peripheral. Bar: 50 µm, Figs. 4.19, 20; 5 µm, Figs. 4.21--24.

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The S10 and S11 plastids With further development, some of the S10 plastids in the inner special parenchyma have some loss of starch accompanied by an increase in stroma. Osmiophilic bodies are more numerous, and some of these bodies contain small acicular crystals (Figure 4.26). These crystals are confined to these bodies and appear to penetrate them, and sometimes they extend beyond the borders of the osmiophilic bodies (Figure 4.26 insert). These changes continue from the inner most part of the nectary near the locule wall toward the outer epidermis. The transition between S10 and S11 (Figure 4.27) shows an increase in the number of plastids containing less starch and more osmiophilic bodies with crystals. We use the term amylochromoplasts to designate the plastids at these two stages and into S12. In some cells the plastids appear partially degenerated (Figure 4.23, 27), which suggests they give rise to some of the vacuoles seen in Figure 4.23, as well as to vacuoles seen at S12 (Figure 4.24). These observations suggest that some of the cells lose their plastids via degeneration, while others retain intact plastids.

The S12 plastids By S12 (anthesis), many but not all of the plastids in the special parenchyma have lost their starch (Figure 4.24), while plastids in other cells have a few starch grains as well as an increased number of crystals (Figure 4.28--31). These latter plastids contain many osmiopilic bodies, and there are many crystals that have enlarged and extended the boundary membranes of the plastids (Figure 4.31). In addition, stroma lamellae in some plastids have proliferated into what appear to be sheets or reticulate networks in some plastids (not shown). The crystal-containing plastids are consistent with the plastids seen in the Vibratome sections (Figure 4.17, 18). If the flowers are not pollinated, these plastids with their crystals continue for at least 48 h before they degenerate. The special parenchyma cells at S12 are highly vacuolated (Figure 4.24), but retain their cellular identity with intact nuclei, mitochondria, ER, and vacuoles.

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Figs. 4.25–31. Transmission electron microscope sections showing nectary cell plastids from S9 through S12

25. S9 plastid engorged with multiple starch grains. 26. S10 plastid with fewer starch grains and more osmiophilic bodies with small dense acicular crystals (inset). 27. S11 plastid with one starch grain, large osmiophilic bodies with associated acicular crystals. 28. S12 plastids with several small starch grains, osmiophilic bodies, and acicular crystals, some wavy. 29. S12 plastids with several starch grains, large osmiophilic bodies and larger acicular crystals, some curved. 30. S12 plastid with little starch, large osmiophilic bodies, and ghosts of acicular crystals (possibly extracted during processing). 31. S12 plastid with starch grains, many osmiophilic bodies, and a portion of a long tail associated with long acicular crystals that extend from plastid stroma. Bar: 2 µm, Figs. 4.25--28, 30, 31; 1 µm, Fig. 4.29; 0.5 µm, Fig. 4.26 inset.

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4.3.3 Biochemical and molecular genetic analyses To accompany these anatomical and developmental results, a series of biochemical and molecular genetic analyses designed to identify and clarify the roles of these compounds in nectary function was conducted. Initially, the yellow pigment that accumulates in the earlier stages and appears at its maximum in the mature S12 nectaries was isolated and characterized. To identify the yellow pigment, extracts of the nectary for analysis were prepared. As shown in Figure 4.32B, the nectary extract gives a single peak by HPLC analysis. On the same HPLC column, three common carotenoids (lutein, lycopene, and β-carotene) were run (Figure 4.32A). The nectary extracted material co-chromatographed with β-carotene. Further analysis indicated that this yellow pigment had an identical visible absorption spectrum to β-carotene (Figure 4.32C). To evaluate the production of β-carotene in the nectary, we used TLC at different developmental stages. A yellow pigment was present in the different stages of the developing nectary (Figure 4.33A). The absorbance spectrum of this pigment at each stage of development was characteristic of β-carotene (data not shown) indicating that the only pigment extractable from nectaries was β-carotene. The level of β-carotene in nectary extracts also was quantified. As shown in Figure 4.33B, early stages of nectary development contained little β-carotene and even by S9, only moderate levels were produced. However, between S9 and S12, the level of β- carotene increased 10-fold. To further evaluate expression of the β-carotene biosynthetic pathway, we isolated cDNAs encoding portions of each of the β-carotene biosynthetic enzymes, and the latter were assayed. Most of these genes were identified from a study of expressed sequence tags (EST) (K. Taylor et al., unpublished data). These clones range in size from small gene fragments to full-length cDNA clones. These β- carotene biosynthetic genes are shown in Table 4.2 by gene name; GenBank accession number; length of each of the LxS8 nectary-expressed genes along with the most closely related reference gene (from other solanaceous species) and its length; the percentage identity of the LxS8 gene vs. the reference gene; and finally Table 4.2 β-carotene biosynthesis genes used in this analysis

Gene name Clone GenBank references Position of EST size GenBank Size GenBank Species in identity to Symbol Identity in accession (bp) accession GenBank (%) reference GenBank Ipi ipp isomerase DQ212769 1157 AB049815 L. esculentum 1107 96 Full length farnesyl fps pyrophosphate DQ212770 700 X84695 C. annuum 1228 92 280-990 synthase

geranylgeranyl 145 gps pyrophosphate DQ212775 458 X92893 C. roseus 1271 83 860–poly A synthase phytoene psy DQ212779 2102 L23424a L. esculentum 11191 92 Full length synthase carotenoid 1020–poly crtiso DQ212772 1018 AF416727 L. esculentum 2387 94 isomerase A

a: here is no full-length phytoene synthase clone in the GenBank. To be continued Table 4.2 β-carotene biosynthesis genes used in this analysis (continued)

Gene name Clone GenBank references Position of clone EST size GenBank Size GenBank Species in identity relative Symbol Identity in accession (bp) accession GenBank (%) to GenBank reference phytoene 1686–poly pds DQ212776 91 X68058 C. annuum 1929 95 desaturase A ζ-carotene zds DQ212773 853 AF195507 S. tuberosum 1931 92 1289-poly A desaturase 146 lycopene β- lyc DQ212774 797 X81787 N. tabacum 1614 96 981–poly A cyclase carotene ccd celavage DQ212781 1279 AY576002 L. esculentum 1638 94 652-poly A dioxygenase neoxanthin 1430–poly nxs DQ212780 307 AF254793 L. esculentum 1666 90 synthase A a: here is no full-length phytoene synthase clone in the GenBank. 147

Fig. 4.32. (A–C). Identification of nectary carotenoid as ß-carotene

HPLC elution detector response at 450 nm. (A) Carotenoid standards, a = lutein; b = lycopene; c = ß-carotene. (B) Carotenoids isolated from ornamental tobacco nectaries. (C, inset) Detector response for purified nectary carotenoid. Solid line is standard ß-carotene, and dashed line is carotenoid isolated from ornamental tobacco nectaries.

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Fig. 4.33. (A–C). Analysis of nectary-expressed carotenoids extracted from LxS8

(A) Thin-layer chromatogram of carotenoids isolated from various stages of nectary development (stages 2, 6, 9, and 12). Std, ß-carotene standard. The origin (ori), solvent front, and migration of ß-carotene are indicated. (B) Absorbance (450 nm) of ß-carotene in nectary extracts at various developmental stages.

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the location of the cloned LxS8 gene relative to the reference gene. All these genes are closely related to their reference gene. With the exception of the geranylgeranyl pyrophosphate synthase gene, which is 83% identical to its reference gene, the remainder share >90% identity. Together this group of genes represents a complete set of unique tools for the analysis of β-carotene biosynthesis. Based on the nucleotide sequence of each of these clones, RT-PCR oligonucleotides pairs (Table 4.3) were developed to permit analysis of this biosynthesis pathway in nectaries. To evaluate the patterns of expression for each of these genes, we performed RT-PCR using mRNA isolated from various tissues and at different developmental stages of the nectary. This analysis is shown in Figure 4.34. As can be seen, these genes were widely expressed throughout the plant. Various levels were found in all of the organs examined. The nonfloral organs (leaf, stem and root—lanes 13–15) expressed each of the carotene biosynthetic genes. The non-nectary floral organs (lanes 7–12) also expressed varying levels of these genes. Among the non-nectary floral organs, the ovary (lane 11) appeared to express each gene at higher levels than any of the other tissues. Phytoene synthase appeared to be constitutively expressed in all organs and tissues studied. The nectary also expressed each of these genes and, with the possible exception of phytoene synthase ( pys), each gene was transcriptionally regulated during nectary development (lanes 1–6). The farnesyl pyrophosphate synthase ( fps), carotenoid isomerase (crtiso), and lycopene B-cyclase (lyc) genes had the greatest changes in transcription as the nectary developed. The expression of each gene increased to maximal levels at S6 and S9 (lanes 2 and 3). Following S9, the expression levels of the genes appeared to decline slightly or to disappear by S12 (lane 5). Then after pollination (lane 6), most of the genes were greatly down-regulated. The only gene that did not undergo such a great down-regulation was phytoene synthase (Cookson et al., 2003). Thus, during floral development the genes encoding these β-carotene biosynthetic enzymes were regulated (both up and down) in the nectary. A surprising result was that β-carotene biosynthetic gene expression reached maximal levels by S6, yet the levels of β- carotene remained low until S9. Because of this observation, the regulation of β-

Table 4.3 Ascorbate biosynthesis genes used in this analysis

gpi phosphate DQ202348 361 AY690423 S. tuberosum 1992 93 152 A isomerase L-galactose Complete gdh DQ202349 965 TC126912 S. tuberosum 1316 93 dehydrogenase ORF L-galactono- mgldh 1,4-lactone DQ202350 962 AB048530 N. tabacum 1951 95 121 – 986 dehydrogenase

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Fig. 4.34. RT-PCR of genes encoding ß-carotene biosynthetic enzymes.

Genes: ipi, isopentenyl pyrophosphate isomerase; fps, farnesyl pyrophosphate synthase; gps, geranylgeranyl pyrophosphate synthase; pys, phytoene synthase; crtiso, carotenoid isomerase; pds, phytoene desaturase; zds, -carotene desaturase; lyc, lycopene ß-cyclase; ccd1, carotenoid cleavage dioxygenase; nxs, neoxanthin synthase; 26S rRNA; 18S rRNA. GenBank accession numbers for genes are in Table 4.1. First strand cDNAs were prepared and analyzed as described in Materials and Methods.

Tissues: lane 1, nectary S2; lane 2, nectary S6; lane 3, nectary S9; lane 4, nectary S11; lane 5, nectary S12; lane 6, nectary postfertilization; lane 7, floral tube; lane 8, petal; lane 9, sepal; lane 10, stigma; lane 11, ovary; lane 12, pedicel; lane 13, leaf; lane 14, stem; lane 15, root.

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carotene production must be at a level other than the transcriptional level. While posttranscriptional regulation of those genes (either at the translational or posttranslational level [protein regulatory]) could account for this observation, we believe that a simpler and more plausible explanation is simply the availability of substrates for β-carotene production. In addition, we evaluated the expression of the carotene dioxygenase, ccd1, which uses carotene as a substrate for the formation of apocarotenoids. This gene was also strongly expressed in S6 and S9 nectaries and its reduced expression continued through S12, but it was turned off after fertilization. Figure 4.35 is a summary of the biochemistry in the maturing nectaries. We previously showed that starch accumulates to high levels during the filling stage of nectary development (S2 through S9) (G. Ren et al., unpublished data). From that study, we found that during this timeframe, the biosynthetic enzymes for β-carotene production are also expressed. At about S9, the nectary undergoes a shift in metabolism that we refer to as the "anabolic/catabolic shift," and the accumulated starch that is stored in the nectary begins to be degraded to glucose. The resulting glucose has several fates within the cell. Foremost, it is converted into sucrose, the predominant sugar secreted into nectar. Somewhere during this secretory process, the action of several invertases (extracellular, vacuolar, and acidic forms) hydrolyze approximately 50% of the sucrose (S) to glucose (G) and fructose (F). These three sugars, (S/G/F) in a 1 : 1 : 1 molar ratio, make up the sugar constituents of nectar (G. Ren et al., unpublished data). Second, glucose feeds into the ascorbate biosynthetic pathway to produce another antioxidant, ascorbate, and eventually oxalic acid/oxalate. Finally, glucose also enters the plastidal 2-C-methyl-d-erythritol- 4-phospahte (MEP) pathway leading to the production of isopentenyl pyrophosphate (IPP), the precursor of β-carotene. Because the β-carotene biosynthetic genes are expressed at high levels by S6, it is likely that the biosynthetic machinery for β-carotene synthesis is in place early in nectary development. This is confirmed by the low levels of β-carotene accumulation up to S9 (see Figure 4.33B). However it is not until starch breakdown began after S9 that β-carotene accumulated to high levels. This suggests that the 156

Fig. 4.35. Interconnection of biochemical pathways in the nectary

Oxidative stress is generated by the nectar redox cycle. ß-Carotene and ascorbate function as antioxidants inside the nectary and in soluble nectar, respectively. IPP = isopentenyl pyrophosphate; Glc = glucose; Fruc = fructose.

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substrates leading into β-carotene biosynthesis are the limiting features of the accumulation of β-carotene and that the sugars from starch degradation might feed into the β-carotene biosynthetic pathway. To evaluate whether sugars can serve as precursors for β-carotene in the nectary, [14C]-sucrose was fed through the pedicel of immature (S9) flowers and after incubation for 2 d, the nectaries were dissected from the flowers, and the carotenoids were isolated and evaluated by TLC. As can be seen in Figure 4.36, Lanes 1 and 2, β-carotene is the predominant pigment isolated from the nectaries of these flowers. When the thin layer plate was exposed to reveal the location of the isotope, the β-carotene was highly labeled confirming the biochemistry shown in Figure 4.35, indicating that sugars can feed through the MEP pathway into β- carotene biosynthesis. Because the antioxidant β-carotene is mostly produced after S9, we hypothesized that ascorbate, the other major antioxidant involved in the nectar redox cycle, might also be most significantly produced late in development. Therefore, nectaries from different developmental stages were harvested, ground, and assayed for the presence of ascorbate. As shown in Figure 4.37, ascorbate is present in nectaries at quite low levels throughout the filling stage (S2 to S9) of floral development. However, between S9 and S12, the amount of ascorbate increased approximately 10-fold. This is highly reminiscent of the pattern observed with β- carotene production. The question of whether the ascorbate biosynthetic pathway was regulated similarly to the β-carotene biosynthetic pathway was pursued. To evaluate expression of the ascorbate biosynthetic pathway, a subset of the genes encoding the entire pathway was utilized. The ascorbate biosynthetic pathway consists of eight enzymatic steps, starting with glucose-6-phosphate (Figure 4.38A). Many of these steps produce intermediates that are involved in biochemistry unrelated to ascorbate metabolism (e.g., cell wall and glycoprotein biosynthesis). Because of these overlaps with other biochemical pathways, we focused on the two enzymatic steps that are directly related to ascorbate biosynthesis, l-galactose dehydrogenase and l-galactono-1,4-lactone dehydrogenase. These enzymes are the 159

Fig. 4.36. Thin-layer chromatogram (TLC) visualization of nectary- expressed carotenoids.

Pedicels of S9 flowers were placed in [14C]-sucrose for 48 h. Subsequently, nectaries were harvested and carotenoids were isolated. A TLC of duplicate flowers was run and photographed (lanes 1 and 2). The TLC plate was then exposed to a phosphorimager plate (lane 3 corresponds to lane 1, lane 4 to lane 2). The origin is indicated with the position of ß-carotene and an unidentified compound.

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Fig. 4.37. Quantity of ascorbate extracted from six nectaries of ornamental tobacco flowers at different developmental stages

Six nectaries were isolated and ascorbate quantified as outlined in Materials and Methods.

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Fig. 4.38. (A, B). Ascorbate metabolism.

(A) Biochemical pathway for ascorbate. (B) RT-PCR of genes encoding ascorbate biosynthetic enzymes. Genes used: gpi, glucose phosphate isomerase; galdh, l-galactose dehydrogenase; Mt_glodh, mitochondrial l- galactono-1,4-lactone dehydrogenase; 26S rRNA; 18S rRNA. First strand cDNAs were prepared and analyzed as described in Materials and Methods.

Tissues used: lane 1, nectary S2; lane 2, nectary S6; lane 3, nectary S9; lane 4, nectary S11; lane 5, nectary S12; lane 6, nectary postfertilization; lane 7, floral tube; lane 8, petal; lane 9, sepal; lane 10, stigma; lane 11, ovary; lane 12, pedicel; lane 13, leaf; lane 14, stem; lane 15, root.

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final two steps in the biosynthesis of ascorbate. Therefore, cDNAs were cloned from LxS8 nectary mRNA encoding each of these enzymes along with glucose-6- phosphate isomerase, the first step in the pathway. As shown in Table 4.3, cloning cDNA fragments encoding each of these cDNAs was successful. These fragments ranged in size from 360 to 960 nucleotides in length and share high identity with their reference clone. Together these clones represent the first and final two steps in ascorbate biosynthesis, and together they make a unique set of tools for ascorbate gene analysis in the nectaries of ornamental tobacco. To evaluate the patterns of expression for each of these genes, we performed RT-PCR using mRNA isolated from various tissues and at different developmental stages of the nectary. This analysis is shown in Figure 4.38B. As was observed for the β-carotene biosynthetic pathway, the ascorbate biosynthesis genes were also widely expressed throughout the plant. Various levels were found in all of the organs examined. In the nectary, each of these three genes was expressed maximally by S6, and then expression gradually declined. Thus, it seems that as observed for β- carotene biosynthesis, ascorbate biosynthesis likely takes place in the nectary at early developmental times; however, major levels of ascorbate did not accumulate until after S9, when starch breakdown began. It appears that the biochemical pathways for both β-carotene and ascorbate are present early in nectary development, yet accumulation of these antioxidants did not occur at high levels until starch breakdown began at about S10. These data indicate that the production of these antioxidants appears to be regulated in late nectary development by substrate availability.

4.4 Discussion The tobacco floral nectary is a highly active gland whose early and intermediate developmental stages (S2 through S9) are characterized by the synthesis of starch in plastids and, to a lesser degree, carotenoids in the plastid thylakoids. These events are marked externally by the increase in sizes of the 166

gynoecium and the basal nectary, and the change in color of the nectary from a lime green (S2) to bright orange (S9). Based on an ongoing study (G. Ren et al., unpublished data), we believe the plastids in the young nectaries are not chloroplasts, based on the lack of distinct grana and their capacity to divide, but are plastids containing photosynthetic pigments with the signals to focus on starch synthesis. At S9, the special parenchyma has become engorged with starch-filled amyloplasts containing a few dense thylakoids. Between S9 and S10, the flower opens, and the secretion of nectar begins and lasts through at least S12. The events that follow S9 are remarkable. The starch in the amyloplasts is rapidly catabolized to form nectar sugars (G. Ren et al., unpublished data) and a substrate source for β-carotene and ascorbate biosynthesis. The biosynthesis of the carotenoids is so great that the β-carotene forms multiple crystals in the majority of plastids. We identify these dual-function plastids that contain both starch and β- carotene during the later developmental stages as amylochromoplasts. In addition, other metabolic events occur late in nectary development. They are the expression of a novel MYB transcription factor that regulates NEC1 expression prior to anthesis (G-Y. Liu et al., unpublished data), and the resulting NEC1-mediated accumulation

of H2O2 in the secreted nectar (Thornburg et al., 2003). Durkee (1983) has characterized several types of floral nectaries developmentally based on whether their internal cells degenerate during secretion, such as holonecrine in soybean (Horner et al., 2003) or as eccrine when the internal secretory cells retain their integrity for some time following the major secretory phase, as is the case for tobacco. The tobacco nectaries have been shown to continue to secrete beyond S12 if they have not been pollinated. Even chromoplast development in a variety of fruits and other plant structures has been followed in other studies (discussed in the introduction), the present study clearly documents that the plastids from the early stages of nectary development through S12 may be considered to have a dual function. We thus give them the name amylochromoplast. Biochemical analysis of carotenoids during the entire period of nectary development (S2 through S12) showed that in the earlier stages 167

carotenoids were formed most likely in the plastid lamellae, and at S9 this process greatly increased with the spectacular formation of β-carotene crystals from the osmiophilic bodies. We believe these osmiophilic bodies are the later sites of the carotenoids and serve as the centers for β-carotene crystal development. Their production occurs in all of the nectary cells to a greater or lesser extent, giving the nectaries their distinct orange appearance from S10 through S12. Different methods of preservation and embedments were used to better maintain them. However, they were only undistorted in the fresh Vibratome sections. In that our primary goal was not to better characterize these crystals microscopically, but only to show their presence, we did not pursue cryopreservation and cryosectioning methods. Thornburg et al. (2003) concluded that the production of the NEC1 protein was involved in H2O2 production in the nectar. This oxidative compound provides the nectar with an oxidizing medium to serve as an antimicrobial environment to protect the nectar sugars and other diverse components present (Davis et al., 1998; Pacini et al., 2003; C. Carter et al., in press). The expression of this protein and its transcription factor beginning at S9, using in situ hybridization and immunocytochemistry, supports this contention (C. Carter et al., in press). Because the S9 nectary cells that produce NEC1 remain viable through S12 and beyond, it seems that the cells of the nectary, and possibly the rest of the gynoecium, need to be protected. Our results strongly suggest that the production of β-carotene and ascorbic acid provide the counterbalancing antioxidants needed to accomplish this protection. Furthermore, and in concert with a number of other recent studies (Smirnoff and Wheeler, 2000; Green and Fry, 2005), some of the ascorbic acid may be converted into calcium oxalate crystals (Horner et al., 2000), which come to line the gynoecial wall and are interspersed in the septum between the two locules during the later stages of gyneocium development. These crystal-filled idioblasts, containing prismatic and crystal sand crystals are evident in Figure 4.10, 12, and 14. These crystals may serve a secondary function as a physical and chemical barrier to 168

protect the gynoecium and the enclosed developing seeds from insects and other organisms that can use them as a source of food (Korth et al., 2006). Nectar begins to be secreted by late S10 and early S11. One of the primary characteristics of nectar secretion in ornamental tobacco, in addition to the sugar secretion is the production of H2O2 via the Carter–Thornburg nectar redox cycle. This recently discovered biochemical pathway (Carter et al., 1999; Carter and

Thornburg, 2000, 2004a--c) produces high levels of H2O2 (up to 4 mM) that are antimicrobial and protect the gynoecium and the developing ovules from microbes transferred to the flower by visiting pollinators or by wind/weather. Thus one notable feature of the nectar of ornamental tobacco flowers is the highly oxidative environment found in nectar. As a consequence, the nectary likely produces antioxidants to help alleviate this oxidative stress. We have previously identified ascorbate as the major extracellular antioxidant in soluble nectar (Carter and Thornburg, 2004a, b), and we propose that β-carotene serves as the major intracellular antioxidant in the nectary. Its production peaks at anthesis (S12) exactly when the oxidative stress is greatest. To evaluate gene expression affecting carotenoid and ascorbate biosynthesis, we isolated 12 cDNAs in these two biosynthetic pathways. Based on the nucleotide sequence of each of these clones, we developed RT-PCR oligonucleotides pairs (Table 4.1) to permit analysis of this biosynthetic pathway in nectaries. As shown in Figure 4.36, the entire pathway of β-carotene biosynthesis is well expressed in nectaries, and maximal levels of these cDNAs are expressed by developmental S6 to S9. Thus, the levels of mRNAs encoding the β-carotene biosynthetic enzymes are highly expressed before the maturation phase (S9 to S12), when most of the β-carotene is produced. Clearly another factor, other than transcriptional regulation, is responsible for regulating the production of carotenoids in the nectary. There are two possible regulatory paradigms that could function in this regard. First, posttranscriptional regulation of those genes, either at the translational level or posttranslational (protein regulatory) level, could account for this observation. However, these phenomena generally do not function to regulate 169

an entire pathway. Further, because low levels of β-carotene are being produced at these early stages (see Figure 4.1--4), we postulate that wholesale down-regulation of the entire pathway is unlikely. A simpler and more elegant explanation is the availability of substrates for β-carotene production. In summary, the tobacco nectary represents a very complex factory that simultaneously involves a wide variety of anabolic and catabolic processes that together contribute to nectar production and protection. This study has provided fundamental information for future studies that may help to alter nectar production and composition for better insect visitation, cross pollination, and hybridization of important crop and horticultural plants.

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Chapter 5 Future work

5.1 Introduction of transgenic plants based future work Based upon the work described in Chapter 2, 3, and 4, a series of experiments based on transgenic plant methods have been initiated. These experiments are designed to answer the following questions: 1) Does the rate of the transient starch accumulated in floral nectary affect the nectar production in the developing nectary? 2) Does the alteration of the structures of transient starch in tobacco nectary regulate the timing of nectar production? 3) Does sucrose responsive element binding proteins (SREBP) regulate starch metabolism? 4) Does the β-carotene accumulated in later stages of floral nectary development remove starch degradation products? To address these questions, we have generated a series of transgenic plants that either over express or knockout express of specific genes involved in starch metabolism in the nectary based upon the previously tested metabolic pathways (Figure 5.1). Overexpression studies utilize the vector, pEarlyGate 101 (Keith et al., 2006), which contains the strong CaMV 35S promoter upstream of a fusion protein. The fusion protein contains a gene of interest with an YFP-HA tag at C-terminal. The pEarlyGate 101 vector contains a Kanr gene for subcloning and a Barr gene inside of left bounder and right bounder for plant selection (Figure C1). Gene knock-out studies utilize the vector, pHellsGate 2 (Chris et al, 2002), that contains the strong CaMV 35S promoter upstream of two inverted repeat cassettes of a specific gene (Figure C1) to generate the RNAi. The pHellsGate 2 contains a Specr gene for subcloning selection and a Kanr resistent gene for plant selection. To address the question 1, whether the rate of the transient starch accumulation affect the nectar production, we have generated sucrose synthase (SuSy) expressed Nicotiana tobaccum transgenic plants. In addition, we also have 171

Sucrose ADPGlucose 1 S S 2 SuSy S S S P 3 G S S S S A B G

Fructose Glucose Starch

GPA+Pyruvate AcetylCoA

IS P DC H M IPP

IPI

DMAPP

GGPP

βcarotene

Figure 5.1 Scheme for transgenic plants 172

generated a series of AGPase over expressed and RNAi expressed Nicotiana tobaccum and LxS8 transgenic plants (Table 5.1). Since the floral nectary is a heterotrophic organ and it is unable to photosynthesize, it is necessary to define the metabolism of carbohydrate used for starch synthesis in nectary. Sucrose synthase, which catalyzes sucrose into fructose-1-P and UDP-glucose, has been suggested for the cleavage of sucrose into utilizable sugar. Previously data had shown that sucrose synthase was highly expressed in nectary, especially in early stages (Ren et al., submitted). Thus, sucrose synthase knock-down would regulate the transported photosynthates supply to nectary cells, and further alter the starch accumulation rate. AGPase is the first committed step in the synthesis of starch. This enzyme is responsible for producing ADP-glucose, the soluble substrate required for starch synthases. AGPase holoenzyme in higher plants is a hetero-tetramer with two large and two small subunits. In Arabidposis, AGPL is represent by four large subunits (AGPL1, AGPL2, AGPL3, and AGPL4) and AGPS is represented by two small subunits (AGPS1 and AGPS2). AGPase large subunits functions as regulators of the catalytic AGPase small subunits (Crevillen et al., 2005). Hence, regulation of AGPase would also lead to alteration of starch accumulation rate in nectary cells. Previous data had also shown that osmorality in developing floral cell introduced by starch degradation products were used to tigger the large scale amount of nectar at late stage (Stage 12). Therefore, SuSy and AGPase genetic modified transgenic would alter the starch accumulation rate in developing floral nectary, thereby to investigate the relationship of the rate of starch accumulation and the amount of nectar carbohydrate. To address the question 2, whether the alteration of the structures of transient starch in tobacco nectary regulates the timing of nectar production, we have made a series of starch synthase RNAi Nicotiana tobaccum transgenic plants, including transgenic plants for SS1, SS2, SS3, GBSS (Table 5.1). In addition, the SS2 and SS3 RNAi transgenic plants for ornamental tobacco LxS8 line had been generated. Previous data showed that the mRNA level of SS1, SS3 and GBSS were highly 173

Table 5.1 Transgenic plants summarization

Nicotiana Ornamental Gene of Interest Construct tobaccum tobacco LxS8 pHG-AGPS 14 4 AGPase pEG-AGPS 12 0 SuSy pHG-SuSy 8 0 SREBP1 pHG-SRE1 13 3 SREBP2 pHG-SRE2 14 0 GBSS pHG-GBSS 13 0 SS1 pHG-SS1 20 0 SS2 pHG-SS2 35 1 SS3 pHG-SS3 20 3 pHG-IPI 13 0 IPI pEG-IPI 0 0 pHG-MDC 11 0 MDC pEG-MDC 5 0 pHG-ISPH 27 0 ISPH pEG-ISPH 16 0

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expressed at early stages in LxS8 floral nectary gland. The expression levels of those three genes were lower at late stages (Chapter 3). The mRNA level of SS3 was highly expressed in nectaries, while the mRNA level of SSII was not detectable in nectaries. Hence, regulation of different starch synthase isoforms would alter starch accumulation patterns and starch structures in developing floral nectaries. These alterations would cause nectar production to be released at different time period during floral nectary development. Therefore, regulation of starch accumulation rate and starch structure would give us a hint whether starch was used to regulate the timing of nectar secretion. To address the question 3, whether sucrose responsive element binding proteins (SREBP) regulates starch metabolism, the RNAi expressed Nicotiana tobaccum transgenic plants for SREBP1 and SREBP2 were generated. In addition, the RNAi expressed ornamental tobacco LxS8 transgenic plants for SREBP1 were generated (Table 5.1). SREBP1 and SREBP2 are two transcription factors categorized to the R2R3 myb family (Figure 5.2). DNA sequence analysis shows that SREBP1 and SREBP2 have different regulatory domains. Those two transcription factors were highly inducible by higher concentration of sucrose in potato leaf (Kim et al., 1993, Kim et al., 1997). RT-PCR data shows that SREBP1 is near floral nectary specifically expressed. The SREBP1 mRNA is highly accumulated in Stage 9, but neither expressed in early stages and nor in later stages (Figure 5.3). While the SREBP2 is constitutively expressed in all of the plant organs (Figure 5.3). Investigation of the starch accumulation patterns in SREBP mutants and starch metabolic gene expression patterns would give us the evidence that whether SREBP1 is one of the transcription factors regulating starch metabolism. To address the question 4, whether the β-carotene accumulated in later stages of floral nectary development remove starch degradation products, we have generated a series of over expressed and RNAi expressed Nicotiana tobaccum transgenic plants, including mevalonate diphosphate decarboxylase (MDC), 1- hydroxy-2-methyl-2(e)-butenyl 4-diphosphate (HM-2B4PP) reductase (ispH), and 175

Figure 5.2 3D structure of the DNA-binding domain of sucrose responsive element binding proteins. A SREBP1; B SREBP2; C lmsfC; D lmseC; E lh89C; F lh88C. (C,D,E,F were from PDB).

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A D

B E

C F

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Figure 5.3 Gene expression pattern of SREBP detected by RT-PCR S2: Stage 2 nectary; S6: Stage 6 nectary; S9: Stage 9 nectary; S11: Stage 11 nectary; S12: Stage 12 nectary; PF: Post-fertilized nectary; Ft: floral tube; Pe: Petal; Se: Sepal; St: Stigma and filament; Pt: Petiol; Ov: Ovary; L: leaf; S: Stem; R: Root.

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SREBP1

SREBP2

26S rRNA

18S rRNA S2 S6 S9 S11 S12 PF Ft Pe Se St Pt Ov L S R

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isopentenyl diphosphate (IPP) isomerase (IPI) (Table 5.1). β-carotene biosynthesis is thought to start with mevolonate pathway (Goldstein and Brown, 1990), however, MEP pathway has been found to be the major metabolic pathway to start β-carotene biosynthesis in plastids (Lichtenthaler 1999, Rohmer 1999, Eisenreich et al. 2001, Dubey et al., 2003). Therefore, those two parallel pathways (Figure 5.1) may crossed talk with each other to produce isopentenyl diphosphate to initiate β- carotene biosynthesis in nectary cells. Gene expression detection of those two pathway was illustrated in Figure 5.4. As can been seen in Figure 5.4, the gene expression of the mevalonic pathway and MEP pathway were highly regulated during floral nectary development. Therefore, the down-regulation of MDC, ispH and IPI might decrease β-carotene accumulation in nectary cells, and the up-regulation of MDC, ispH and IPI might increase β-carotene accumulation in nectary cells. The relationship of β-carotene accumulation in nectaries and nectar carbohydrate would tell us whether β-carotene were one of the strategies that nectaries had been adapted to release the increased intracellular osmorality in nectaries.

5.2 Materials and methods 5.2.1 Plant lines Wild type of Nicotiana LxS8 and wild type of Nicotiana tobaccum were used in this study. The ornamental tobacco LxS8 line of ornamental tobacco plants used in this study was derived from an interspecific cross between Nicotiana langsdorffii and N. sanderae (Kornaga et al., 1997).

5.2.2 RNAi constructs A PCR based method was used to generate the RNAi constructs. Primer pairs used for RNAi constructs were listed in Table 5.2. Amplified PCR products were subcloned into pHellgate 2 vector (Chris et al, 2002) using a Gateway® BP clonase™ II kit (Invitrogen, cat# 11789-020) and the constructs were verified by sequencing using primers (H8-L-F, H8-L-R, H8-R-F, H8-R-R) listed in Table B.1.

180

Table 5.2 Primers for transgenic plant verification

Primer Gene Construct Primer Sequences Size (bp) Name AGPS-IR2 TGGGAAGATATTGGTACC pHG-AGPS 800 HG8-L-R TATCATGCGATCATAGGCGTCT AGPase P-AGPS TGGGAAGATATTGGTACC pEG-AGPS 800 EG-R ACGCTGAACTTGTGGCCGTTTA FSY-F1 ATTCAGATGGATTTCTTC SuSy pHG-SuSy 450 HG8-L-R TATCATGCGATCATAGGCGTCT SER1-IF2 TGAACGGGCTTCGTCACT SREBP1 pHG-SRE1 550 HG8-L-R TATCATGCGATCATAGGCGTCT SRE-IF1 ACCAGGAAGCTACAGGAG SREBP2 pHG-SRE2 600 HG8-L-R TATCATGCGATCATAGGCGTCT P-GBSS TTCAATGTCCCTTTGGCTCACA GBSS pHG-GBSS 232 HG2-L-R GGGGTACCGAATTCCTCGA P-SS1 TGTTGGCATAATCCTGTCAGCA SS1 pHG-SS1 228 HG2-L-R GGGGTACCGAATTCCTCGA P- SS2 TGCTTACTGACTTATCGCGAGTACA SS3 pHG-SS2 236 HG2-L-R GGGGTACCGAATTCCTCGA P-SS3 CCCGTGCAGTTCAAGATTTAAACC SS3 pHG-SS3 271 HG2-L-R GGGTACCGAATTCCTCGA IPI-IF2 GAACAGCTGAAAGAGCTT pHG-IPI 500 HG8-L-R TATCATGCGATCATAGGCGTCT IPI IPI-IF2 GAACAGCTGAAAGAGCTT pEG-IPI 500 EG-R ACGCTGAACTTGTGGCCGTTTA MDC-IF2 ATTGGCTGTGTTGAGAA pHG-MDC 700 HG8-L-R TATCATGCGATCATAGGCGTCT MDC MDC-IF2 ATTGGCTGTGTTGAGAA pEG-MDC 700 EG-R ACGCTGAACTTGTGGCCGTTTA ISPH-IF4 GGTGACATCTGGTGCATCCA pHG-ISPH 550 HG8-L-R TATCATGCGATCATAGGCGTCT ISPH ISPH-IF4 GGTGACATCTGGTGCATCCA pEG-ISPH 550 EG-R ACGCTGAACTTGTGGCCGTTTA

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Figure 5.4 Gene expression of mevalonate pathway and MEP pathway genes detected by RTPCR A mevalonic pathway gene expression; B MEP pathway gene expression. S2: Stage 2 nectary; S6: Stage 6 nectary; S9: Stage 9 nectary; S11: Stage 11 nectary; S12: Stage 12 nectary; PF: Postfertilized nectary; Ft: floral tube; Pe: Petal; Se: Sepal; St: Stigma and filament; Pt: Petiol; Ov: Ovary; L: leaf; S: Stem; R: Root

182

A ACCT1 ACCT2 HMGS HMGR

MVK PMK MDC

rRNA rRNA

S2 S6 S9 S11 S12 PF Ft Pe Se St Pt Ov L S R

B DXS2 DXR

ISPD ISPE

ISPF ISPG ISPH

26S rRNA 18S rRNA S2 S6 S9 S11 S12 PF Ft Pe Se St Pt Ov L S R 183

5.2.3 Overexpression constructs A pEarleygate 101 vector with 35S promoter (Keith et al., 2006) was used to make C-terminal YFP-HA fusion protein construct. pHellsGate constructs from above were subcloned into pEarleygate 101 using a Gateway® LR clonase™ II kit (Invitrogen, cat 11791-020) according to manufactory instruction. The constructs were verified by sequencing using primers (H8-L-F, GFP-R) listed in Table B.1.

5.2.4 Plant transformation and plant selection RNAi expression transgenic plants An Agrobacterium tumefaciens (LAB4404) introduced tobacco leaf disc transformation method (Curtis et al., 1995) was used to generate both ornamental tobacco LxS8 and Nicotiana tobaccum transgenic plants. Co-cultivation medium (CC, pH 5.7) contains 8% phytagel (Sigma, cat. 2751175), 4.4 g/L Murashige & Skoog (RPI, cat. M10400-25.0), 30 g/L sucrose, 1 mg/L BAP and 0.1 mg/L NAA). Callus induction medium (CI, pH 5.8) contains 8% phytagel, 4.4g/L Murashige & Skoog, 30 g sucrose, 1mg/L BAP, 0.1 mg/L NAA, 50 mg/L carbonicillin and and 100 mg/L Kanamycin. Root induction medium (RI, pH 5.8) contained 5% phytagel, 4.4 g/L Murashige & Skoog, 30 g sucrose, 50 mg/L carbonicillin, and 50 mg/L Kanamycin.

Overexpression transgenic plants All of the medium used in overexpression constructs were the same as above, except the antibiotics for plant selection. 25 mg/L glufosinate ammonium was used for callus selection of pEarlyGate 101 derivatives. 10 mg/L glufosinate ammonium was added in root induction medium for pEarlyGate 101 derivatives.

Seeds collection Brown paper bags were used to collect the self-pollinated seeds for each transgenic plant. About 10 pods of mature seeds from each transgenic plant were collected, recorded, and stored (Table D2). 184

5.2.5 Genomic DNA verification of transgenic plants Plant genomic DNA extraction 250 mg of leaves from each plant were collected and grinded up in 1.5 mL eppendorf tube with 500 μL Shorty buffer (10 μg/L glycogen, 0.2 M Tris-HCl (pH9.0), 0.4 M LiCl, 25 mM EDTA, 1% SDS). Followed the supplementation of 500 µL PCI (Phenol:Cloroform:Isoamyl Alcohol at a ratio of 25:24:1), the samples were vigorously vortexed for 30 seconds and then centrifuged at maximum speed to remove any particulates. The genomic DNA in the supernatant were precipitated by equal volume of isopropanol, then washed by 70% ethanol. The extracted genomic DNA were dissolved in 50 μL TE buffer (pH 8.0) supplemented with 1mg/mL RNase.

Genomic DNA PCR A PCR based method was used to verify whether the target constructs were integrated into the genome of host plants. One primer in the PCR reaction was designed from specific gene sequences and another primer was designed from vector sequences (Table 5.2). Primer HG2-L-R was designed according to the pHellsGate 2 vector sequences, and primer EG101-R was designed according to the pEearleyGate 101 vector sequences. The size of amplicons were from 220 bp to 280 bp in order to optimize the consistency of PCR efficiency (Table 5.2). PCR was performed in the condition as: 94 °C for 30 sec, 50 °C for 45 sec, 72 °C for 45 sec, with 35 cycles. The PCR products were separated on 1.5% agrose gels with ethidium bromide, and then visualized under UV lights.

5.2.6 Protein isolation To isolate proteins from each floral stage, 400 mg leaves from each plant lines were homogenized with liquid nitrogen. To each sample, we added 250 µL of 62.5 mM Tris-HCl (pH 6.5), 50 mM DTT solution, containing 1% protease inhibitor cocktail (Sigma, cat # P9599). Samples were then centrifuged at 4ºC, 5000 rpm for 30 min and the supernatant was collected as a crude protein extract. Protein in 185

each sample was quantified using Bradford method (Bradford, 1976) using bovine serum albumin as a standard.

5.2.7 Western blot assay 30 µg protein of each sample prepared above was loaded on a 12% SDS- PAGE gel (Laemmli, 1970) and separated under 200 Voltages for 90 min. After transfer of protein to a nitro-cellulose membrane (Osmonic Inc., cat # WP4HY00010), the membrane was incubated with a rabbit antibody raised against potato AGPase small subunit. A rabbit secondary antibody conjugated with peroxidase was used and the peroxidase activity was detected (Amersham, cat # RPN2109).

5.2.8 Starch staining analysis To compare starch content in developing ornamental tobacco flowers and in various transgenic plants, the starch was detected by an iodine staining method according to Zhang et al. (Zhang et al., 2005). We collected flowers with identified stages at the same time. Then the harvested flowers were boiled in 50 mL 80% ethanol for 10 min to be decolorized. The decolorized flowers were stained with a freshly prepared solution of I2/KI (5g KI and 0.5g I2 in 500 mL water) for 5 min, destained in water for 10 min, and photographed immediately. In all instances, all of the flowers in any given experiment were collected, stained, destained and photographed simultaneously.

5.2.9 Nectar carbohydrate analysis Nectar was collected from flowers as previously described (Carter et al., 1999) with one exception. To insure that we were collecting all of the nectar, the floral tubes, nectaries, and receptacles were washed with 1 mL of water and this wash was combined with the isolated nectar. The samples were used immediately for carbohydrate determination. The total carbohydrate in nectar was analyzed by the sensitive anthrone method (Morris, 1948). 186

5.3 Results 5.3.1 Transgenic plants verification The positive transgenic plants were verified by PCR using one gene specific primer and one vector specific primer (Table 5.2, Figure 5.5). Figure 5.5 showed that approximately 85% of the transgenic plants were have the target insertion in the genomic DNA. To test whether that the RNAi expressed plants have an effect on the protein level. Antibody from potato AGPase small subunit was used to for immunodetection. Twenty micrograms of total protein from various RNAi expressed ornamental tobacco LxS8 were used in this test. These LxS8 transgenic plants include pHGAGPS1, pHGAGPS2, pHGSS-a, pHGSS3-b, pHGSREBP1-a, pHGSREBP1-b, pHGMYB305-1, pHGMYB305-2. Immunodetection of AGPase showed that the RNAi expressed AGPS LxS8 transgenic plants were have 30-60% knockdown at the protein level (Fig. 5.6 A). To test whether the RNAi expressed Nicotaina tobaccum plants have the effect at the protein level, antibody from maize sucrose synthase was used in this test. Figure 5.6B showed the pHGSuSy1 and pHGSuSy2 of Nicotiana tobaccum transgenic plants were a candidate of sucrose synthase knockout lines.

5.3.2 Starch in flowers from various transgenic plants To test that floral nectary was the major organ responsible for starch storages, we used iodine based staining method to monitor starch accumulation in the whole inflorescence. Figure 5.7 showed that starch in the developing nectary were accumulated at early stages, but degraded at late stages. At the early stages (Stage 2, and Stage 6), starch were not highly deposited at nectary, but were accumulated at those early stages of floral tube and petals (Figure 5.7A, B). At Stage 9, iodine staining showed more intensity in nectary than other floral components (Figure 5.7C). At the later stages (Stage 12s) starch deposited in nectary were starting to be degraded, so were starch accumulated in floral tube and 187

Figure 5.5 Genomic DNA PCR verification of transgenic plants PCR was used to identify the construct insertion in genomic DNA. A pHGSRE1 Nicotiana tobaccum transgenic plant lines; B pHGSRE2 Nicotiana tobaccum transgenic plant lines; C pHGAGPS Nicotiana tobaccum transgenic plant lines; D pEGMDC Nicotiana tobaccum transgenic plant lines; E pEGISPH Nicotiana tobaccum transgenic plant lines; F pEGAGPS Nicotiana tobaccum transgenic plant lines; G pHGAGPS ornamental tobacco transgenic plant lines.

188

A pHG-SRE1 (N. tobaccum) pRT011 pRT001 pRT002 pRT003 pRT004 pRT005 pRT006 pRT007 pRT008 pRT009 pRT010 pRT012 pRT013 pRT014 pRT025 pRT015 pRT016 pRT017 pRT018 pRT019 pRT020 pRT021 pRT022 pRT023 pRT024 pRT026 pRT027 B pHG-SRE2 (N. tobaccum) pRT011 pRT001 pRT002 pRT003 pRT004 pRT005 pRT006 pRT007 pRT008 pRT009 pRT010 pRT012 pRT013 pRT014 pRT025 pRT015 pRT016 pRT017 pRT018 pRT019 pRT020 pRT021 pRT022 pRT023 pRT024 pRT026 pRT027

C pHG-AGPS (N. tobaccum) pRT135 pRT136 pRT134 pRT137 pRT133 pRT214 pRT125 pRT126 pRT127 pRT128 pRT129 pRT130 pRT131 pRT132 D pEGM D C (N. tobaccum) pRT155 pRT153 pRT151 pRT152 pRT154

E pEGI SPH (N. tobaccum) pRT201 pRT194 pRT195 pRT196 pRT197 pRT198 pRT199 pRT190 pRT192 pRT202 pRT203 pRT200 pRT191 pRT193 pRT189 pRT204

F pEGAGPS (N. tobaccum) pRT219 pRT216 pRT217 pRT220 pRT210 pRT218 pRT221 pRT211 pRT213 pRT212 pRT214 pRT215 G pHGAGPS (LxS8) pRT221 pRT220 pRT222 189

Figure 5.6 Immunodetection of transgenic plants

A Antibody against potato AGPase small subunit was used to test the protein level of RNAi expressed transgenic plants.

B Antibody against maize sucrose synthase was used to test the protein level of RNAi expressed transgenic plants. 190

A pRT224 pRT222 pRT223 xxs8 wt pRT226 pRT227 myb305-1 myb305-2

B pRT121 pRT123 pRT122 pRT116 pRT117 pRT118 pRT119 pRT120 N. tobaccum wt tobaccum N. 191

Figure 5.7 Starch accumulation detected by iodine staining in LxS8 flowers Two flowers represent each floral stage were used to be dissected for iodine staining. A Stage 2; B Stage 6; C Stage 9; D Stage 11; E Stage 12 (anthesis); F: Stage 12 (12 hours after anthesis); G: Stage 12 (24 hours after anthesis); H: Stage 12 (36 hours after anthesis).

192

A B C D E

F G H I 193

petals (Figure 5.7D-H). However, the starch in floral tube was depleted earlier than the starch in petals (Figure 5.7F-H). To further to verify that the ornamental tobacco LxS8 transgenic plants have the effect on starch synthesis, an iodine-based method (Zhang et al. 2005) was used to visualize the starch in various floral parts. And to further test whether the starch synthesis have an impact on nectar carbohydrate secretion, an anthrone based method were used to measure the total sugar in various ornamental transgenic plants (Morris, 1948). Figure 5.8 showed that AGPase knockdown transgenic plants had similar starch deposition pattern as the wild type, the SS3 transgenic plants had more starch deposited in developing floral parts, and the myb305 transgenic plants had less starch deposited in developing. Figure 5.9 showed that SS3 transgenic plants had more nectar sugar than the wild type, while the myb305 had less nectar sugar than the wild type. 194

Figure 5.8 Starch staining of various transgenic plants line in ornamental tobacco All transgenic plant lines were RNAi expressed transgenic ornamental tobacco lines. Left column of each panel was the flowers 24 hours before anthesis, the right column of each panel was the flowers 24 hours after anthesis. A wild type ornamental tobacco; B: wild type ornamental tobacco; C myb3051; D myb3052; E srebp1a (pRT-230); F agpsa (pRT-222); G ss3a (pRT-226); H ss3b (pRT-227).

195

A B

D C

E F

H G

196

Figure 5.9 Nectar carbohydrates measurement of various transgenic plants line in ornamental tobacco

All transgenic plant lines were RNAi expressed transgenic ornamental tobacco lines. Left column of each panel was the flowers 24 hours before anthesis, the right column of each panel was the flowers 24 hours after anthesis.

197 198

Chapter 6 Summarization and discussion

We had shown that ornamental tobacco was a plant suitable for studying floral nectary development (Chapter 2, 3, 4, 5). The LxS8 line of ornamental tobacco plants used in this study was derived from an interspecific cross between Nicotiana langsdorffii and N. sanderae (Kornaga et al., 1997). Both of these parents are diploid and belong to the Alatae section of Nicotiana. These plants were previously used to study a genetic instability as well as the tobacco nectar proteins (Carter et al., 1999, Carter and Thornburg, 2000). Flower development in tobacco is divided into 12 stages, from Stage 1, where the petals and sepals are of equal length to Stage 12 (anthesis) when the flower is fully open and the anthers have dehisced (Koltunow et al., 1990). Development of the nectary gland in ornamental tobacco also consists of several of discrete stages that correlate with the stages of flower development (Figure 2.2). Large amount of nectar carbohydrates is secreted outside of developing floral nectaries at later stages (Table 2.1). To study the functions of starch in the ornamental tobacco developing floral nectaries, we first demonstrated that starch in developing floral nectaries illustrated transit nature (Chapter 2). The starch in the developing floral nectaries was accumulated up to Stage 9, and then degraded after Stage 9 (Figure 2.3, 2.4). The starch polyglycan distributions in the developing floral nectaries were dynamically changed indicating that starch metabolism were highly regulated in floral nectaries (Figure 2.5). The gene expression patterns of starch metabolic genes indicated that the starch metabolism in the developing floral nectaries were highly regulated at transcriptional level (Chapter 3). The mRNA levels of starch anabolic genes, including AGPase, sucrose synthase, and starch synthases, were highly expressed at early stages of the developing floral nectaries, and then declined at later stages (Figure 3.4, 3.5, 3.6, 3.7). While the mRNA levels of starch catabolic genes, including α-amylase and β-amylase, were not highly expressed at early stages of the developing floral nectaries, but increased at later stages (Figure 3.7). The protein 199

levels of AGPase and sucrose synthase displayed similar pattern as the mRNA levels (Figure 3.4, 3.5). Second, we also had illustrated that nectar carbohydrate secretion in the developing floral nectaries was characterized into two phases, the slow phase and the rapid phase (Chapter 2). Nectar carbohydrate quantification illustrated that a small scale of nectar carbohydrates in the slow phase was secreted after Stage 10 followed by a large scale of nectar carbohydrates in the rapid phase was flushed out after anthesis (Figure 2.8, Table 2.1). 14C-sugar chasing experiments verified that the large scale of nectar carbohydrates were from transported photosynthates (Figure 2.8). In addition, the sugar chasing experiments also indicated that the transported sugars entered the floral nectary cells to be further processed as the nectar products (Figure 2.7). Third, we had illustrated that nectary cells were determined to autonomous cellular changes (Chapter 2, 3, 4). The number of nectary cells was not changed significantly, instead the size of nectary cells increased about 3 fold during floral nectary development (Figure 2.3), indicating that the development of nectary gland was autonomous. Microscopy works also indicated that vacuole was enlarged and chromoplast was emerged at Stage 12 (Figure 3.2, 4.19-31). Transition of chloroplast to chromoplast was also illustrated by Microscopy works (Figure 4.19- 31). In addition, large amount of β-carotene, a presumable pre-antioxidant material, was deposited as a crystal-like form in late stages of the developing floral nectary cells (Figure 4.1-12, Figure 4.32). Gene expression pattern indicated that the β- carotene metabolic pathway (Figure 4.8) was not highly regulated during the development of floral nectary, indicated that the accumulation of β-carotene was regulated by the supplication of substrates of the β-carotene metabolic pathway. And fourth, we have also generated a series of transgenic plants, either using Nicotiana tobaccum or ornamental tobacco, to investigate the molecular mechanisms of starch regulations and biomass conversions in developing floral nectary (Chapter 5). Phenotypical observations transgenic plants indicated that starch metabolic genes were not only responsible for plant growth, but also 200

responsible for nectar carbohydrates production. The AGPase down-regulated transgenic plant in LxS8 suggested that starch deposited in plants regulated not only the growth of plants, but also the development of flowers (Chapter 5). The down- regulated starch synthase transgenic plants suggested that starch synthase 2 was the one controlling plant growth, starch synthase 1 and starch synthese 3 were responsible for starch accumulation in nectary gland (Chapter 5). These observation were consistent with the starch synthase gene expression pattern in ornamental tobacco (Figure 3.6, 3.7).

6.1 Starch and floral development Here, we are demonstrating that starch might be the element chose to adapt to nectar-pollinator co-evolution. We had shown that starch deposited were regulating plant growth including flower development (Chapter 5). The AGPase down-regulated transgenic plant in LxS 8 clearly indicated that the shortened floral tubes were embedded in sepal during the flower development although the structures of flowers remained intact (Ren, unpublished data). A similar shorten floral tube was observed in Myb 305 down-regulated LxS8 transgenic plants (Liu et al., unpublished data). The Myb 305 was thought to be a transcription factor responsible for regulating Nectarin genes, as well as for regulating starch metabolism (Liu et al., unpublished data). In addition, the petal size of Lily was shortened by the use of starch degradation inhibitors (Bieleski et al., 2000). These observations suggested that starch was not only used for floral tube elongation but also used for flower opening. The floral tube elongation and flower opening allowed pollinators to drink the nectar.

6.2 Starch and nectar production It had been arguably suggested that either starch provided the source of nectary carbohydrates or phloem solutes supplied the source of nectary carbohydrates. However, no conclusive statements of the source of floral nectaries 201

have been reached, partially due to the complexity of nectary types and the availability of collectable nectar. We had shown that floral nectar secretion was divided by two phases, the slow phase and the rapid phase (Figure 2.8). The observations indicated that starch deposited in developing floral nectary cells was not only contributed as a source of nectar production in the slow phase, but also as a regulator for nectar production in the rapid phase (Chapter 2). We also had shown that utilizing starch as the regulator would be relatively an energy saving process, although the starch deposition in developing floral nectary cells was an energy consuming process. The ornamental tobacco could use a small amount of starch to build up the endogenous osmolarity in nectary cell for rapid nectar secretion. Only 4 µL of phloem sap were needed to build up 700 µg of starch to build the osmolarity to attract 70 µL of phloem soluts at the rapid phase of nectar secretion (Calculated from Figure 2.4 and 2.8). The build-up osmolarity in floral nectary cells could change the water potential in nectary gland. The water potential changes could be used to pump the phloem solutes into nectary cells to be further secreted as the nectar. Hence, we conclude that no matter what the evolution origin of nectar production was, the starch was the carrier and regulator for nectar production secretion. We also had shown that depletion of starch in floral tube impaired nectar secretion (Chapter 5, Figure 5.5 and 5.6) by preventing flower opening. Further evidences of that conclusion could be verified by transgenic plants designed to introduce various starch accumulation rate and different starch structures in nectary cells. We have demonstrated that starch can regulate nectar production secretion. However, the timing of nectar secretion is another important factor used for nectar- pollinator co-evolution. The rate of starch accumulation and the structure of starch granules might be the major factors used for the timing controls. The genetic modified transgenic plants would give us the hint about this question. Starch synthase 2 down-regulated transgenic plants were growing slower and smaller than the starch synthase 1 and starch synthase 3 down-regulated transgenic plants. These observation indicated that starch synthase 1 and starch synthase 3 were the 202

major starch synthase isoforms for regulating the starch structure and the starch accumulation rate, while the starch synthase 2 was the one for regulating plant growth (data not shown). Further evidences were needed to test the relationship of the timing of nectar secretion with the starch structure and the starch accumulation rate.

6.3 Starch biochemistry in developing tobacco floral nectary Based on the observation we have made previously, we were able to generate a working model for the functions of starch in ornamental tobacco floral nectary. Starch was starting to build up at the early stages of the development of floral nectary (Figure 6.2A) and then degraded at later stages (Figure 6.2B, C). The products from starch degradation would face several destinations: A) contribute as a source of nectar production in slow phase; B) build up the osmolarity in nectary cells to attract photosynthates in phloem; C)provide the substrates for β-carotene synthesis.

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Figure 6.1 Speculated biochemical process in ornamental tobacco nectary development Panel A: Starch accumulation phase (starting from Stage 1 to Stage 9 of floral nectary development); Panel B: Nectar secretion phase, slow phase (starting from Stage 10 to early Stage 12); Panel C: Nectar secretion phase, rapid phase (starting from late Stage 12).

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Appendices

I am hereby providing the information of materials had been used and will be used in this project. Four appendices were provided, including Appendix A, Index of ornamental tobacco LxS8 cDNA clones; Appendix B, Index of oligonucleotides; Appendix C, Index of constructs for tobacco transgenic plants; Appendix D, Index of tobacco transgenic plants.

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Appendix A Index of ornamental tobacco LxS8 cDNA clones

In this appendix, one table and three figures were provided: Table A1, Index of ornamental tobacco LxS8 cDNA clones; Figure A1, Overview of the metabolism pathways in ornamental tobacco floral nectary; Figure A2, Maps of subcloning vectors. Thirty-nine genes involved in starch metabolic pathway (14 genes), mevalonate pathway (7 genes), MEP pathway (7 genes), β-carotene pathway (8 genes), and ascorbate pathway (3 genes) were identified from ornamental tobacco LxS8 (Figure A1). Fifty-three LxS8 clones from these metabolic pathways had been isolated and identified from LxS nectary cDNA libraries either using gene specific primer sets (Appendix B) or searching ESTs from the EST project (Taylor et al., unpublished data). These clones were subcloned to four different vectors, pCR- Topo, pDNR-LIB, pGEM-T easy, and pTriplEx (Figure A2, Table A1). GenBank accession numbers and detailed information were provided in Table A1. MEP pathway Mevolonate pathway GAP + pyruvate Acetyl-CoA DXS 2, 1-deoxy-d-xylulose-5-phosphate (DXP) synthase 2 ACCT, acetoacetyl-CoA thiolase DXR, 1-deoxy-d-xylulose-5-phosphate (DXP) reductoisomerase HMGR, HMG-CoA reductase ispD, 2-C-methyl-D-erythritol 4-phosphate (MEP) cytidyltransferase HMGS, HMG-CoA synthase ispE, 4-diphosphoytidyl-2C-methyl-D-erythritol (CDP-M) kinase MVK, mevalonate kinase ispF, 2-C-methyl-D-erythritol 2,4- cyclodiphosphate synthase PMK, phosphomevalonate kinase ispG, 1-hydroxy-2-methyl-2(e)-butenyl 4-diphosphate (HM-2B4PP) synthase MDC, mevalonate diphosphate decarboxylase ispH, 1-hydroxy-2-methyl-2(e)-butenyl 4-diphosphate (HM-2B4PP) reductase IPP DMAPP IPI, IPP isomerase IPP DMAPP IPP isomerase

Starch metabolic pathway 207 glucose Ascorbate pathway β-carotene pathway AGPase, ADP-glucose pyrophosphorylase GGPP D-galacturonate

SS1, starch synthase 1 CRTISO, carotenoid isomerase GPI, glucose-6-phoshate SS2, starch synthase 2 SS3, starch synthase 3 FPS, farnesyl pyrophosphate synthase isomerase GPS, geranylgeranyl pyrophosphate synthetase GBSS, granule-bound starch synthase GLDH, L-galactose LYC, lycopene β-cyclase dehydrogenase SBE1, starch branch enzyme 1 PDS, phytoene desaturase SBE2, starch branch enzyme 2 ZDS, ζ -phytoene desaturase MGLDH, Mitochondrial PYS, phytoene synthase ISA1, isoamylase 1 L-galactono-1,4-lactone ISA2, isoamylase 2 dehydrogenase ISA3, isoamylase 3 β-carotene L-Ascorbate R1, PHO, starch phosphorlase AMY, α-amylase BMY, β-amylase glucose

Figure A.1 Overview of the metabolism pathways in ornamental tobacco floral nectary

208

A B

D C

Figure A2 Maps of subcloning vectors

A: pGEM-T easy; B: pDNR-LIB; C: pTriplEx; D: pCR2.1 TOPO 209

Table A1 Index of ornamental tobacco LxS8 cDNA clones

GENBANK CONSTRUCT SYMBOL GENE NAME VECTOR NO. pRT700 SS1 starch synthase 1 pGEM-T DQ021463 pRT701 SS2 starch synthase 2 pGEM-T DQ021466 pRT702 SS3 starch synthase 3 pGEM-T DQ021464 pRT703 GBSS granule boundary starch synthase pGEM-T DQ021465 pRT704 AMYL α-amylase (large) pGEM-T DQ021455 pRT705 AMYS α-amylase (small) pGEM-T DQ021456 pRT706 BAM β-amylase pTRIPLEX DQ021457 pRT707 ISA1 isoamylase 1 pGEM-T DQ021461 pRT708 ISA2 Isoamylase 2 pGEM-T DQ021462 pRT709 ISA3 isoamylase 3 pDNR-LIB DQ021471 pRT710 SBE1 starch branching enzyme 1 pGEM-T DQ021459 pRT711 SBE2 starch branching enzyme 2 pGEM-T DQ021460 pRT712 PHO starch phosphorylase pTRIPLEX DQ021468 pRT713 R1 starch R1 enzyme pTRIPLEX DQ021469 pRT714 AGPS ADP-glucose pyrophosphorylase pGEM-T DQ021458 pRT715 SuSy1 sucrose synthase 1 pGEM-T DQ021467 pRT716 SuSy1 sucrose synthase 1 pGEM-T nsa pRT717 SuSy2 sucrose synthase 2 pGEM-T DQ021470 pRT718 SREBP1 sucrose responsive element binding factor 1 pTRIPLEX nsa pRT719 SREBP2 sucrose responsive element binding factor 2 pDNR-LIB nsa pRT720 SuP sucrose phosphatase pDNR-LIB nsa

1-deoxy-d-xylulose-5-phosphate (DXP) a pRT721 DXS2 pDNR-LIB ns synthase 2 1-deoxy-d-xylulose-5-phosphate (DXP) pRT722 DXR pGEM-T nsa reductoisomerase 2-C-methyl-D-erythritol 4-phosphate (MEP) pRT723 ISPD pGEM-T nsa cytidyltransferase

To be continued

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Table A1 Index of ornamental tobacco LxS8 clones (continued)

GENE GENBANK CONSTRUCT GENE NAME VECTOR SYMBOL NO.

2-C-methyl-D-erythritol 2,4- a pRT724 ISPF pDNR-LIB ns cyclodiphosphate synthase ISPG, 1-hydroxy-2-methyl-2(e)-butenyl 4- pRT725 pDNR-LIB nsa GcpE diphosphate (HM-2B4PP) synthase 1-hydroxy-2-methyl-2(e)-butenyl 4- pRT726 ISPH pGEM-T nsa diphosphate (HM-2B4PP) reductase pRT727 ACCT1 cytosolic acetoacetyl-CoA thiolase pDNR-LIB nsa

LXS8 peroxisomal acetoacetyl-CoA a pRT728 ACCT2 pTRIPLEX ns thiolase a pRT729 HMGR HMG-CoA reductase pDNR-LIB ns a pRT730 HMGS HMG-CoA synthase pDNR-LIB ns pRT731 MVAK mevalonate kinase pGEM-T nsa pRT732 PMK phosphomevalonate kinase pGEM-T nsa pRT733 MDC mevalonate diphosphate decarboxylase pGEM-T nsa pRT734 CCD carotene cleavage dioxygenase pTRIPLEX DQ212781 pRT735 CTISO carotenoid isomerase, pCR2.1- DQ212772 TOPO pRT736 FPS farnesyl pyrophosphate synthase pGEM-T DQ212770 pRT737 GPS geranylgeranyl pyrophosphate synthetase pDNR-LIB DQ212775 pRT738 IPI IPP isomerase pTRIPLEX DQ212769 LCY, pRT739 Lycopene β-cyclase pTRIPLEX DQ212774 CRTL pRT740 NXS Neoxanthia synthase pDNR-LIB DQ212780 pRT741 PDS phytoene desaturase pCR2.1- DQ212776 TOPO pRT742 PYS Phytoene synthase pTRIPLEX DQ212779 pRT743 ZDS ζ-carotene desaturase pCR2.1- DQ212773 TOPO pRT744 GPI glucose-6-phoshate isomerase pDNR-LIB DQ202348 pRT745 GLDH L-galactose dehydrogenase pGEM-T DQ202349 Mitochondrial L-galactono-1,4-lactone pRT746 MGLDH pGEM-T DQ202350 dehydrogenase nsa :not submitted to GenBank

211

Appendix B Index of oligonucleotides

I had designed oligonucleotides for gene isolation, RT-PCR (or Realtime RT- PCR) testing, and constructs verification in this study (Table B1). In addition, I had also designed primer sets to make transgenic plant constructs (Table B2) using BP cloning (Invitrogen, cat# 11789020) or LR cloning methods (Invitrogen, cat# 11791020). The forward primers for transgenic plant constructs contained the attB1, one of the BP recombination sites that is located at the upstream of the 5’ of the specific gene sequences. While the reverse primers for transgenic plant constructs contained the attB2, one of the BP recombination sites that is located at the upstream of the 3’ of these specific gene sequences. 212

Table B1 Index of oligonucleotides

OLIGONUCLEOTIDE NAME SEQUENCE PURPOSE

NL-AGPS-F AGCAAAGACGTGATGTTAAACC Gene isolation

NL-AGPS-R TCTTCACATTGTCCCCTATACG Gene isolation

NL-AGPSF-F1 GCAATCACACTCTACCACACA Gene isolation

NL-AGPSF-F2 CAAGCTTTGTTAACAATGGCG Gene isolation

NL-AGPSF-R TACCTCATGGCTGTTTAGCC Gene isolation

NL-SU-F TATTTCGCCCAGGAAAATGTC Gene isolation

NL-SU-R TGAACGATGATCTCAGCTGGA Gene isolation

NL-SUSYF-F1 TTGTCTGAGGATTTCCCATCTG Gene isolation

NL-SUSYF-F2 TTCCCATCTGCTGAATCAACTG Gene isolation

NL-SUSYF-R TTTTTTTTTGGACAAATTGAAA Gene isolation

NL-GB-F TCCCTGCTACCTAAAGTCGAT Gene isolation

NL-GB-R CACAGATTGGCACTGTTCCAT Gene isolation

NL-SS1-F TTTGTGTGTTACAAGTAGTGTTTC Gene isolation

NL-SS1-R CGACGGGTTCCAATCATTAA Gene isolation

NL-SS2-F TGGCAAGCCTCAATGTAAAG Gene isolation

NL-SS2-R CACCACTGATACTTAGCAGCG Gene isolation

NL-SS3-F CAAGCATGCCATTGCTTATG Gene isolation

NL-SS3-R CTTCCACCTTTCCAAACCAT Gene isolation

NL-BE1-F TACTAGTGCATAGACCACTGCTG Gene isolation

NL-BE1-R TTCGATGCTGTATGTTCACCA Gene isolation

NL-BE2-F ATGGCCAAACTCATTATCCAT Gene isolation

NL-BE2-R ATGAGTTTATGACTTCAGAACA Gene isolation

NL-ISO1-F GTGTAGATTTTGGGAGTATT Gene isolation

NL-ISO1-R ACCCAATACAGAGTTATAACT Gene isolation

NL-ISO2-F GCTTCATTTGCTTTATGCCTC Gene isolation

NL-ISO2-R CACACTCGTCTCCCATGTTAA Gene isolation

NL-AAM-F TCAGCAGTCTGATCCATTGGT Gene isolation

NL-AAM-R TTTGGCTGTGCCTCAAGAAT Gene isolation

NL-GLDH-F ATGGCAGCTCAGACATTGCA Gene isolation

To be continued

213

Table B1 Index of oligonucleotides (continued)

OLIGONUCLEOTIDE NAME SEQUENCE PURPOSE

NL-GLDH-R TCAGCTTTGTTGGATACCGCTA Gene isolation

NL-FPS-F TTAAAGCATCTGCTCTTGGC Gene isolation

NL-FPS-R CTCGTATTCCAGAAATACTTCCTCA Gene isolation

NL-MGLDH-F TCAAATGCTCCGTTCCTTCA Gene isolation

NL-MGLDH-R ATAGGATTGCATGTCACGACCA Gene isolation

NL-DXR-F CTGGAAGGGCTGTTGCTGA Gene isolation

NL-DXR-R TGGGATGCTTCAAAGCATCA Gene isolation

NL-ISPD-F ACAACATGTCTACTCTTCAATTGGG Gene isolation

NL-ISPD-R CCCTAAGATTCTTCAATGGTTCAA Gene isolation

NL-ISPF-F GTCAAACCCCAAATTGATTCTTT Gene isolation

NL-ISPF-R TTTCCTATAAAGATTGAGGGCCTT Gene isolation

NL-ISPH-F CATGGCTATTCCTCTCCAATTCT Gene isolation

NL-ISPH-R AGGCCAATTGTAAGGCTTCTTC Gene isolation

NL-MVK-F GCTTTTCTTTGGCTGTTTCACCTC Gene isolation

NL-MVK-R CATTTCCACCAATTCCAGCA Gene isolation

NL-PMK-F GTTAAATAATGGCAGTAGTGGCCTC Gene isolation

NL-PMK-R AAGGGAAGCCAATGACTTAGGT Gene isolation

NL-MDC-F AACACCCACAAACATAGCAGTGA Gene isolation

NL-MDC-R TTTCGTCAGGTAGCAAAACTGG Gene isolation RT-18S-F TGGTGCATGGCCGTTCTTAGT RT-PCR RT-18S-R TCGGCCAAGGCTATAAACTCGT RT-PCR RT-26S-F TAAACGGCGGGAGTAACTATGACTCT RT-PCR RT-26S-R TTTCGGCTCCCACTTATACTACACCT RT-PCR

RT-GAP2-F TTGAAGGGTGGTGCCAAGAAA RT-PCR

RT-GAP2-R CATTTATGACCTTTGCCAAGGGAG RT-PCR

RT-AGPS-F GATTTTAGCTTCTACGACCGATCAGC RT-PCR

RT-AGPS-R CAGTTCTTGATCACACAACCTTCACC RT-PCR

RT-SUSY1-F TACAGAGTTGTTCACGGCATTGAC RT-PCR

RT-SUSY1-R CCTGTCCTTTAGCACACACAGATGTT RT-PCR

RT-SUSY2-F GTGTTCGATCCCAAATTCAACG RT-PCR

To be continued

214

Table B1 Index of oligonucleotides (continued)

OLIGONUCLEOTIDE NAME SEQUENCE PURPOSE

RT-SUSY2-R TAGCACACACAGATGCTCCTCGT RT-PCR

RT-GBSS-F ATCTGAATGCCAAGGTCGCTTT RT-PCR

RT-GBSS-R GCTTTTCATATCCATCAATGAAATCG RT-PCR

RT-SS1-F TGACCTCTCTGGAAAGTGTTAAGTGC RT-PCR

RT-SS1-R AAATCCAATACAGCGGACAATCG RT-PCR

RT-SS2-F ATGATGATCGGATGAGGCTTTACAG RT-PCR

RT-SS2-R TCTTCTTTCTGCAATCTGTTGAAGTAGG RT-PCR

RT-SS3-F ATGGCACCTATTGCAAAGGTGG RT-PCR

RT-SS3-R GTCCTTCACCTGATTCATCTTCAAAC RT-PCR

RT-BE1-F ATGGATCATCTTGTGAAGCGCA RT-PCR

RT-BE1-R CCTGGAAAATACAAGATTGTCTTGGAC RT-PCR

RT-BE2-F CGAAGGTGAAATATTCGGCATTG RT-PCR

RT-BE2-R CATGACCAGTCTATTGTGGGTGACA RT-PCR

RT-ISO1-F TTGGATCCTTATGCCAAGGTCA RT-PCR

RT-ISO1-R TAGGTCTCCTTCCCAGTCAAACTGAT RT-PCR

RT-ISO2-F TCTTTGCCGTTTAAGACCTACGGT RT-PCR

RT-ISO2-R GCTCAGTTTTTCGGTAATGCCA RT-PCR

RT-ISO3-F AACGAGATTACCTGGCTTGAGGA RT-PCR

RT-ISO3-R TGTTTTGCTTCCAGAAGAATAGCAG RT-PCR

RT-AAML-F ATATTGCAGGCCTTCAACTGGG RT-PCR

RT-AAML-R TGGAAGGTAACCTTCAGGAGACAA RT-PCR

RT-AAMS-F ATTGCAAATCTGGCTTCCAAGATT RT-PCR

RT-AAMS-R TCTTTGCCTGTGACTATCTTGGTTG RT-PCR

RT-BAM-F TGAGCTATTGGAAATGGCGAAGA RT-PCR

RT-BAM-R AAGAGGATCGTGCAGGAATCA RT-PCR

RT-GLU-F GAATTTATACGGAGCGAGCGAAAC RT-PCR

RT-GLU-R TCGTACTTGTGTGTTTGACCTCCTT RT-PCR RT-R1-F AGAAATATATGTCGAGGTGGTCAGGG RT-PCR RT-R1-R TTGGGTAACCTAACACTTGAGGAGAGT RT-PCR

RT-PHO-F AACCATGATGTGCAAGCGAAGA RT-PCR

To be continued

215

Table B1 Index of oligonucleotides (continued)

OLIGONUCLEOTIDE NAME SEQUENCE PURPOSE

RT-PHO-R TGTAATCTGGCACAAATATTACCTTCAA RT-PCR

RT-SRE1-F ACCGCCCTTTTACTGCTGAAGA RT-PCR

RT-SRE2-R GACTACCCGGATTCATTGACATTG RT-PCR

RT-SRE2-F ACAGCAAGATAAGGTGTTTGTGCC RT-PCR

RT-SRE2-R CTTGTTGCATACACAAACCTAACCCT RT-PCR

RT-CCD-F ACCCGAAGGTTGACCCTGTAACT RT-PCR

RT-CCD-R TTCACCATTTCCTTTGGTTTGAAG RT-PCR

RT-CRTISO-F AGCATAGAAGATTGGGAGGGACTCT RT-PCR

RT-CRTISO-R TTGGTGTCCCCACCTCCTTAAA RT-PCR

RT-FPS-F TGGTACAAACTACCTAAGGTTGGCA RT-PCR

RT-FPS-R GCAGTCTGGAACTCAACCTCATTAAA RT-PCR

RT-GPS-F GTTGCTGATAAGGTAACTTATCCCAAAC RT-PCR

RT-GPS-R CAGATCACTCATGTTTTCACCTTCTATT RT-PCR

RT-IPI-F TCTTCAGCAACGATCAGCAACA RT-PCR

RT-IPI-R TTGTGCAGCATTTCTTACCCCA RT-PCR

RT-LCY-F CATCCCTCAACAGGTTATATGGTAGC RT-PCR

RT-LCY-R AAACCTTCTCGTAGCGGGTAAATC RT-PCR

RT-NXS-F GACCATTTGAATGTGAAAGGTTGG RT-PCR

RT-NXS-R CTTCCATTTCTTCTTTTACCTTTTCACT RT-PCR

RT-PDS-F ATTACGAGTTACTTCTTGGCCGGA RT-PCR

RT-PDS-R GAAGCAACATTTTAGTTCACCATGC RT-PCR

RT-PYS-F CTGTGTCATCAGAGAAAAAGGTGTATG RT-PCR

RT-PYS-R TGCACCACACATATATTGCCCA RT-PCR

RT-ZDS-F AGTGTCAAAGCAGGTTTTGGCA RT-PCR

RT-ZDS-R CGATGTAGTCCTGTTTTGTATATGAGCC RT-PCR

RT-ACCT1-F AGCTGTATCTTTGGGGCACCC RT-PCR

RT-ACCT1-R CCTCCTCCACCATTGCAAACA RT-PCR

RT-ACCT2-F CAGGACTTAAAGCAACAATGATTGC RT-PCR

RT-ACCT2-R GAGGCATCAAACTTTATTAAACCTTCA RT-PCR

RT-HMGR-F AAGATCTACGAAAGGTGTTAGGTGGG RT-PCR

To be continued

216

Table B1 Index of oligonucleotides (continued)

OLIGONUCLEOTIDE NAME SEQUENCE PURPOSE

RT-HMGR-R GGGCAAACATAAAGAGGTAGAGATCC RT-PCR

RT-HMGS-F GGCTTTTGCACTGAGGTTGAAGAT RT-PCR

RT-HMGS-R TGTCGGTATTCCCATGTTTCTCA RT-PCR

RT-MVK-F ATGGAATTCAGCCACCAAGGTT RT-PCR

RT-MVK-R CCTTTGTATTTCTCCCGACCTTTG RT-PCR

RT-PMK-F TGCTCCTGGAAAGGTTCTAATGACT RT-PCR

RT-PMK-R CGCTTCAATCTGATTCCGATACG RT-PCR

RT-MDC-F CTGATCTAAACAGCTATGTCATAGGCG RT-PCR

RT-MDC-R CAAAACTGGACCTCTACCTGGTCTT RT-PCR

RT-DXS2-F AGGGAATACCAATTGAGGTTGGTAA RT-PCR

RT-DXS2-R TCTGGAAGTACCATTGACCTCAACTT RT-PCR

RT-DXR-F GCTGATCAGGTCAAAACATTCAGAC RT-PCR

RT-DXR-R GCGGGCAACCTCGATGATA RT-PCR

RT-ISPD-F TTTGGGAAATGCAGACACCTCA RT-PCR

RT-ISPD-R ATCGTCGGGTGTCGTAACCTTTAT RT-PCR

RT-ISPF-F AAGCTCACTCGGATGGTGATGTT RT-PCR

RT-ISPF-R CTAGGTTTCCCAATTCATAACCTGC RT-PCR

RT-ISPG-F TTGCTTCAAGGTTGCAGAATGC RT-PCR

RT-ISPG-R GTTTTACCACCGTCTTGCCGA RT-PCR

RT-ISPH-F GGTAACAAAGGTCTGGAACACCGT RT-PCR

RT-ISPH-R ATCACAGACATAAGTTGCCTCTGCC RT-PCR

RT-GPI-F TTGGACAGTTACTGGCCATCTATGA RT-PCR

RT-GPI-R GAAGGATCAGAAGGAACATCAGCA RT-PCR

RT-GLDH-F TATCGGAAAAGGTACTAGGGAAGGCT RT-PCR

RT-GLDH-R GCGTCTCATTCACAATCTGATCAAG RT-PCR

RT-MGLDH-F TGTCGGGTCAGGTTTGTCTCC RT-PCR

RT-MGLDH-R ATGAGCACCAACCTGGACAATG RT-PCR P-F TTGCGGTTCTGTCAGTTCCA Constructs verification GFP-F TCGACCAGGATGGGCACC Constructs verification GFP-R ATCACATGGTCCTGCTGGAGTT Constructs verification

To be continuedd

217

Table B1 Index of oligonucleotides (continued)

OLIGONUCLEOTIDE NAME SEQUENCE PURPOSE H8-L-F GCAAGTGGATTGATGTGACATCT Constructs verification H8-L-R CGTCTTACACATCACTTGTCATATT Constructs verification H8-R-F GTGTTATCATTGATCTTACATTTGG Constructs verification H8-R-R TATCATGCGATCATAGGCGTCT Constructs verification

EG-R ACGCTGAACTTGTGGCCGTTTA Transgenic plant verification HG2-L-R GGGGTACCGAATTCCTCGA Transgenic plant verification P-AGPS ATGTTCAAGAAGCGGCTAGGGAAA Transgenic plant verification P-GBSS TTCAATGTCCCTTTGGCTCACA Transgenic plant verification P-IPI GGCGAGGGAGGTCTGAAGCTAT Transgenic plant verification P-MDC AAGACCAGGTAGAGGTCCAGTTTT Transgenic plant verification P-ISPH GGTGACATCTGGTGCATCCA Transgenic plant verification P-SREBP1 CCCAGAATTTTTGGCACTTTTGC Transgenic plant verification P-SREBP2 GGGTTAGGTTTGTGTATGCAACAAG Transgenic plant verification P-SS1 TGTTGGCATAATCCTGTCAGCA Transgenic plant verification P-SS2 TGCTTACTGACTTATCGCGAGTACA Transgenic plant verification P-SS3 CCCGTGCAGTTCAAGATTTAAACC Transgenic plant verification P-SUSY ATTTCAACCGGTGGCCTGAA Transgenic plant verification

218

Table B2 Oligonucleotides for transgenic plant constructs

GENE SIZE FULL PRIMER NAME PRIMER SEQUENCE NAME (BP) SIZE 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPCSYF TCGAAGGAGATAGAACCATGGTCGACATGGCTGA ACGTGTACTGACTC-3’ 2415 YES 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC SUCROSE BPCSYR GCGGCCGCACTCAGCAGCCAATGGAACAGCT-3’ SYNTHASE 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPSYFA TACGCACGGCTAAGGGAGTTAGTTA-3’ 0622 NOb 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC BPSYRA AATGGAACAGCTTCAGCCATCTT-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPCAGPSF TCGAAGGAGATAGAACCATGGTCGACATGGCTTC AGPASE CATTGGAGCC-3’ 1560 YES 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC BPCAGPSR GCGGCCGCAGATGATAATTCCACTGGGAAT-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT a BPCSRE1F TCGAAGGAGATAGAACCATGGTCGACATGGAAGC 0846 YES GATTAAGGAAACGG-3’

SREBP1 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT b BPCSRE1I 0486 NO CGAAACTCCTCAGCAGCCGC-3’ 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC BPCSRE1R GCGGCCGCAACAAAAGCATTTCTTATTGC-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT a BPCSRE2F TCGAAGGAGATAGAACCATGGTCGACATGGATCG 1056 YES GGTTAAAGGTCCATG-3’ SREBP2 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPCSRE2I AACAGCCGCCATTAAAGAGAT-3’ 0682 NOb 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC BPCSRE2R GCGGCCGCAATCGATTTTGCTTACTCCC-3’

a: forward primers (XXXF) and reverse primers (XXXR) can be used to pull out full size cDNAs; b: the intermediate primers (XXXI) and reverse primers (XXXR) are used to make pHellsgate constructs.

To be continued

219

Table B2 Oligonucleotides for transgenic plant constructs (continued)

GENE SIZE FULL PRIMER NAME PRIMER SEQUENCE NAME (BP) SIZE 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPISPHF TCGAAGGAGATAGAACCATGGTCGACATGGCTAT ISPH TCCTCTCCAAT-3’ 1380 YES 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC BPISPHR GCGGCCGCAGGCCAATTGTAAGGCTTC-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPIPIF TCGAAGGAGATAGAACCATGGTCGACATGTCGCT IPI GACAACAACGCC-3’ 875 YES 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC BPIPIR GCGGCCGCAAGTCAATTTGCGGATGGTT-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPMDCF TCGAAGGAGATAGAACCATGGTCGACATGGCAGC MDC AGAACAATCAA-3’ 1269 YES 5’-GGGGACCACTTTGTACAAGAAAGCTGGGTC BPMDCR GCGGCCGCACTTGGGCAGTCCAGTTTCTG-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPGBSSF GAATATATCTGAATGCCAAGGTCG-3’ GBSS 726 NO 5’-GGGGACCACTTTGTACAAGAAAGCTGGGT BPGBSSR GCATGTAACTGAATGAGACCACAAG-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPSS1F TCACCTAGGTATCTGAATGGAGG-3’ SS1 895 NO 5’-GGGGACCACTTTGTACAAGAAAGCTGGGT BPSS1R TTGCTGACAGGATTATGTCAACACC-3’ 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPSS2F TGCAGAAGGAACTTGGTTTACCA-3’ SS2 578 NO 5’-GGGGACCACTTTGTACAAGAAAGCTGGGT BPSS2R CATAGTTATGAGCAGCATTATCC 5’-GGGGACAAGTTTGTACAAAAAAGCAGGCT BPSS3F CAAGACTTCCATACCATTGTCCCC-3’ SS3 745 NO 5’-GGGGACCACTTTGTACAAGAAAGCTGGGT BPSS3R CCATACTTTTATTTCAGTCCCAC-3’ a: forward primers (XXXF) and reverse primers (XXXR) can be used to pull out full size cDNAs; b: the intermediate primers (XXXI) and reverse primers (XXXR) are used to make pHellsgate constructs.

220

Appendix C Index of constructs for tobacco transgenic plants

To investigate the gene functions using transgenic plant methods, fifteen different constructs had been made, including four overexpression and eleven RNAi expression constructs (Table C1). The overexpression vector used in the constructs was pEarleyGate 101 (Keith et al., 2006), which contains the strong CaMV 35S promoter upstream of a fusion protein. The fusion protein contains a gene of interest with an YFP-HA tag at the C terminal of that gene. The pEarlyGate 101 vector contains a Kanr gene for subcloning and a Barr gene inside of the left bounder (LB) and the right bounder (RB) for plant selection (Figure C1). The RNAi expression vector used in the constructs was pHellsGate 2 (Chris et al, 2002), that contains the strong CaMV 35S promoter upstream of two inverted repeat insertions of a specific gene (Figure C1). The pHellsGate 2 contains a Specr gene for subcloning selection and a Kanr gene for plant selection. Long primers conjugated with attB sites (Table B2) were used to first subcloned to pHellsGate 2 to make the RNAi expression constructs using a BP clonase method (Invitrogen, cat# 11789020), then subcloned to pEarleyGate 101 to make the overexpression constructs (Invitrogen, cat# 11791020). The products of BP/LR reactions were transformed into an E. Coli (DH5α) strain. The positive clones of pHellsGate 2 derivatives were selected on LB media containing 30 µg/mL chloramphenicol and 50 µg/mL spectinomycin. The positive clones of pEarleyGate 101 derivatives were selected on LB media supplemented with 200 µg/mL kanamycin. Constructs were verified by sequencing using primers based on the vector sequences (Table B1). The pEarleyGate 101 derivatives were sequenced using the primers as P-F and GFP-R (Table B1), and the pHellsGate 2 derivatives were sequenced using primers as H8-L-F, H8-L-R, H8-R-F and H8-R-R (Table B1). Verified constructs were then transformed into an Agrobacterium tumefaciens LBA 4404 strain. pHellsGate 2 derivatives were selected on LB media containing 221

200 µg/mL spectinomycin, and pEarleyGate 2 derivatives were selected on 200 µg/mL kanamycin. 222

Figure C1 Map of constructs for transgenic plants

A pEarleyGate 101: vector used for overexpression in tobacco. pEarleyGate 101 had a YFP-HA tag at the C-terminal of fusion protein cassette.

B pHellsGate 2: vector used for RNAi expression in tobacco. pHellsGate 2 was used to construct the inverted repeat insertions of a specific gene to express the dsRNA.

Both of pEarleyGate 101 and pHellsGate 2 vectors were utilizing CaMV 35S

(P35S) promoter and nos terminator (T). LB: left bounder of T-DNA. RB: right bounder of T-DNA. H8-L-F, H8-L-R, H8-R-F, H8-R-F were primers used for sequencing verification of insertions of pHellsGate 2 derivatives. H8-L-F and GFP-R were primers used for sequencing verification of insertions of pEarleyGate 2 derivatives. ER2 and HG2-L-R were used for PCR verification using transgenic plant’s genomic DNA as the template. For primers sequences information, check the Appendix B1.

223

A pEarleyGate 101

LB R1 R2 T RB

Kanr Barr PS35 ccd YFP-HA

H8-L-F GFP-R EG-R

B pHellsGate2

LB R1 R2 R2 R1 T RB

spacer Specr Kanr PS35 ccd ccd

H8-L-F H8-L-R H8-R-F H8-R-R HG2-L-R

224

Table C1 Index of constructs for tobacco transgenic plants

CONSTRUCT NAME NOTE

PRT747 PHG2 AGPS KNOCKDOWN

PRT748 PHG2 GBSS KNOCKDOWN

PRT749 PHG2 SS2 KNOCKDOWN

PRT750 PHG2 SS3 KNOCKDOWN

PRT751 PHG2 SS3 KNOCKDOWN

PRT752 PHG2 SRE1 KNOCKDOWN

PRT753 PHG2 SRE2 KNOCKDOWN

PRT754 PHG2 ISPH KNOCKDOWN

PRT755 PHG2 IPI KNOCKDOWN

PRT756 PHG2 MDC KNOCKDOWN

PRT757 PHG2 SUSY KNOCKDOWN

PRT758 PEG AGPS OVEREXPRESSION

PRT759 PEG ISPH OVEREXPRESSION

PRT760 PEG IPI OVEREXPRESSION

PRT761 PEG MDC OVEREXPRESSION

PRT762 PZEO ISPH SUBCLONING

PRT763 PZEO IPI SUBCLONING

PRT764 PZEO MDC SUBCLONING

PRT765 PZEO AGPS SUBCLONING

PRT766 P211 SSI SUBCLONING

PRT767 P211 SUSYS SUBCLONING

225

Appendix D Index of tobacco transgenic plants

Two hundred and thirty-two tobacco transgenic plants, including 221 Nicotiana tobaccum and 11 ornamental tobacco LxS8 plant lines, had been generated (Table D1). The constructs from Appendix C were transformed into either ornamental tobacco Nicotiana tobaccum or ornament tobacco LxS8 plant lines using a modified tobacco leaf disc method (Curtis et al., 1995, Chapter 5). Self-pollinated seeds (Table D1) from the transgenic plants were collected as in the method described in Chapter 5.

226

Table D1 Index of detailed tobacco transgenic plants

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPIEICES

PRT-001 pHG-SREBP1 pHGSRE-01 PRT-T1-001 N. tobaccum PRT-002 pHG-SREBP1 pHGSRE-02 PRT-T1-002 N. tobaccum PRT-003 pHG-SREBP2 pHGSRE-03 PRT-T1-003 N. tobaccum PRT-004 pHG-SREBP2 pHGSRE-04 PRT-T1-004 N. tobaccum PRT-005 pHG-SREBP2 pHGSRE-05 PRT-T1-005 N. tobaccum PRT-006 pHG-SREBP1 pHGSRE-06 PRT-T1-006 N. tobaccum PRT-007 pHG-SREBP2 pHGSRE-07 PRT-T1-007 N. tobaccum PRT-008 pHG-SREBP2 pHGSRE-08 PRT-T1-008 N. tobaccum PRT-009 pHG-SREBP2 pHGSRE-09 PRT-T1-009 N. tobaccum PRT-010 pHG-SREBP1 pHGSRE-10 PRT-T1-010 N. tobaccum PRT-011 pHG-SREBP1 pHGSRE-11 PRT-T1-011 N. tobaccum PRT-012 pHG-SREBP2 pHGSRE-12 PRT-T1-012 N. tobaccum PRT-013 pHG-SREBP2 pHGSRE-13 PRT-T1-013 N. tobaccum PRT-014 pHG-SREBP1 pHGSRE-14 PRT-T1-014 N. tobaccum PRT-015 pHG-SREBP1 pHGSRE-15 PRT-T1-015 N. tobaccum PRT-016 pHG-SREBP2 pHGSRE-16 PRT-T1-016 N. tobaccum PRT-017 pHG-SREBP1 pHGSRE-17 PRT-T1-017 N. tobaccum PRT-018 pHG-SREBP1 pHGSRE-18 PRT-T1-018 N. tobaccum PRT-019 pHG-SREBP2 pHGSRE-19 PRT-T1-019 N. tobaccum PRT-020 pHG-SREBP2 pHGSRE-20 PRT-T1-020 N. tobaccum PRT-021 pHG-SREBP1 pHGSRE-21 PRT-T1-021 N. tobaccum PRT-022 pHG-SREBP1 pHGSRE-22 PRT-T1-022 N. tobaccum PRT-023 pHG-SREBP2 pHGSRE-23 PRT-T1-023 N. tobaccum PRT-024 pHG-SREBP1 pHGSRE-24 PRT-T1-024 N. tobaccum PRT-025 pHG-SREBP2 pHGSRE-25 PRT-T1-025 N. tobaccum PRT-026 pHG-SREBP2 pHGSRE-26 PRT-T1-026 N. tobaccum PRT-027 pHG-SREBP1 pHGSRE-27 PRT-T1-027 N. tobaccum PRT-028 pHG-SS2 pHGSS2-01 PRT-T1-028 N. tobaccum PRT-029 pHG-SS2 pHGSS2-02 PRT-T1-029 N. tobaccum

To be continued

227

Table D1 Index of detailed tobacco transgenic plants (continued)

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPECIE

PRT-030 pHG-SS2 pHGSS2-03 PRT-T1-030 N. tobaccum PRT-031 pHG-SS2 pHGSS2-04 PRT-T1-031 N. tobaccum PRT-032 pHG-SS2 pHGSS2-05 PRT-T1-032 N. tobaccum PRT-033 pHG-SS2 pHGSS2-06 PRT-T1-033 N. tobaccum PRT-034 pHG-SS2 pHGSS2-07 PRT-T1-034 N. tobaccum PRT-035 pHG-SS2 pHGSS2-08 PRT-T1-035 N. tobaccum PRT-036 pHG-SS2 pHGSS2-09 PRT-T1-036 N. tobaccum PRT-037 pHG-SS2 pHGSS2-10 PRT-T1-037 N. tobaccum PRT-038 pHG-SS2 pHGSS2-11 PRT-T1-038 N. tobaccum PRT-039 pHG-SS2 pHGSS2-12 PRT-T1-039 N. tobaccum PRT-040 pHG-SS2 pHGSS2-13 PRT-T1-040 N. tobaccum PRT-041 pHG-SS2 pHGSS2-14 PRT-T1-041 N. tobaccum PRT-042 pHG-SS2 pHGSS2-15 PRT-T1-042 N. tobaccum PRT-043 pHG-SS2 pHGSS2-16 PRT-T1-043 N. tobaccum PRT-044 pHG-SS2 pHGSS2-17 PRT-T1-044 N. tobaccum PRT-045 pHG-SS2 pHGSS2-18 PRT-T1-045 N. tobaccum PRT-046 pHG-SS2 pHGSS2-19 PRT-T1-046 N. tobaccum PRT-047 pHG-SS2 pHGSS2-20 PRT-T1-047 N. tobaccum PRT-048 pHG-SS2 pHGSS2-21 PRT-T1-048 N. tobaccum PRT-049 pHG-SS2 pHGSS2-22 PRT-T1-049 N. tobaccum PRT-050 pHG-SS2 pHGSS2-23 PRT-T1-050 N. tobaccum PRT-051 pHG-SS2 pHGSS2-24 PRT-T1-051 N. tobaccum PRT-052 pHG-SS2 pHGSS2-25 PRT-T1-052 N. tobaccum PRT-053 pHG-SS2 pHGSS2-26 PRT-T1-053 N. tobaccum PRT-054 pHG-SS2 pHGSS2-27 PRT-T1-054 N. tobaccum PRT-055 pHG-SS2 pHGSS2-28 PRT-T1-055 N. tobaccum PRT-056 pHG-SS2 pHGSS2-29 PRT-T1-056 N. tobaccum PRT-057 pHG-SS2 pHGSS2-30 PRT-T1-057 N. tobaccum PRT-058 pHG-SS2 pHGSS2-31 PRT-T1-058 N. tobaccum

To be continued

228

Table D1 Index of detailed tobacco transgenic plants (continued)

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPECIE

PRT-059 pHG-SS2 pHGSS2-32 PRT-T1-059 N. tobaccum PRT-060 pHG-SS2 pHGSS2-33 PRT-T1-060 N. tobaccum PRT-061 pHG-SS2 pHGSS2-34 PRT-T1-061 N. tobaccum PRT-062 pHG-SS2 pHGSS2-35 PRT-T1-062 N. tobaccum PRT-063 pHG-SS3 pHGSS3-01 PRT-T1-063 N. tobaccum PRT-064 pHG-SS3 pHGSS3-02 PRT-T1-064 N. tobaccum PRT-065 pHG-SS3 pHGSS3-03 PRT-T1-065 N. tobaccum PRT-066 pHG-SS3 pHGSS3-04 PRT-T1-066 N. tobaccum PRT-067 pHG-SS3 pHGSS3-05 PRT-T1-067 N. tobaccum PRT-068 pHG-SS3 pHGSS3-06 PRT-T1-068 N. tobaccum PRT-069 pHG-SS3 pHGSS3-07 PRT-T1-069 N. tobaccum PRT-070 pHG-SS3 pHGSS3-08 PRT-T1-070 N. tobaccum PRT-071 pHG-SS3 pHGSS3-09 PRT-T1-071 N. tobaccum PRT-072 pHG-SS3 pHGSS3-10 PRT-T1-072 N. tobaccum PRT-073 pHG-SS3 pHGSS3-11 PRT-T1-073 N. tobaccum PRT-074 pHG-SS3 pHGSS3-12 PRT-T1-074 N. tobaccum PRT-075 pHG-SS3 pHGSS3-13 PRT-T1-075 N. tobaccum PRT-076 pHG-SS3 pHGSS3-14 PRT-T1-076 N. tobaccum PRT-077 pHG-SS3 pHGSS3-15 PRT-T1-077 N. tobaccum PRT-078 pHG-SS3 pHGSS3-16 PRT-T1-078 N. tobaccum PRT-079 pHG-SS3 pHGSS3-17 PRT-T1-079 N. tobaccum PRT-080 pHG-SS3 pHGSS3-18 PRT-T1-080 N. tobaccum PRT-081 pHG-SS3 pHGSS3-19 PRT-T1-081 N. tobaccum PRT-082 pHG-SS3 pHGSS3-20 PRT-T1-082 N. tobaccum PRT-083 pHG-SS1 pHGSS1-01 PRT-T1-083 N. tobaccum PRT-084 pHG-SS1 pHGSS1-02 PRT-T1-084 N. tobaccum PRT-085 pHG-SS1 pHGSS1-03 PRT-T1-085 N. tobaccum PRT-086 pHG-SS1 pHGSS1-04 PRT-T1-086 N. tobaccum PRT-087 pHG-SS1 pHGSS1-05 PRT-T1-087 N. tobaccum

To be continued

229

Table D1 Index of detailed tobacco transgenic plants (continued)

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPECIE

PRT-088 pHG-SS1 pHGSS1-06 PRT-T1-088 N. tobaccum PRT-089 pHG-SS1 pHGSS1-07 PRT-T1-089 N. tobaccum PRT-090 pHG-SS1 pHGSS1-08 PRT-T1-090 N. tobaccum PRT-091 pHG-SS1 pHGSS1-09 PRT-T1-091 N. tobaccum PRT-092 pHG-SS1 pHGSS1-10 PRT-T1-092 N. tobaccum PRT-093 pHG-SS1 pHGSS1-11 PRT-T1-093 N. tobaccum PRT-094 pHG-SS1 pHGSS1-12 PRT-T1-094 N. tobaccum PRT-095 pHG-SS1 pHGSS1-13 PRT-T1-095 N. tobaccum PRT-096 pHG-SS1 pHGSS1-14 PRT-T1-096 N. tobaccum PRT-097 pHG-SS1 pHGSS1-15 PRT-T1-097 N. tobaccum PRT-098 pHG-SS1 pHGSS1-16 PRT-T1-098 N. tobaccum PRT-099 pHG-SS1 pHGSS1-17 PRT-T1-099 N. tobaccum PRT-100 pHG-SS1 pHGSS1-18 PRT-T1-100 N. tobaccum PRT-101 pHG-SS1 pHGSS1-19 PRT-T1-101 N. tobaccum PRT-102 pHG-SS1 pHGSS1-20 PRT-T1-102 N. tobaccum PRT-103 pHG-GBSS pHGGBSS-01 PRT-T1-103 N. tobaccum PRT-104 pHG-GBSS pHGGBSS-01 PRT-T1-104 N. tobaccum PRT-105 pHG-GBSS pHGGBSS-02 PRT-T1-105 N. tobaccum PRT-106 pHG-GBSS pHGGBSS-03 PRT-T1-106 N. tobaccum PRT-107 pHG-GBSS pHGGBSS-04 PRT-T1-107 N. tobaccum PRT-108 pHG-GBSS pHGGBSS-05 PRT-T1-108 N. tobaccum PRT-109 pHG-GBSS pHGGBSS-06 PRT-T1-109 N. tobaccum PRT-110 pHG-GBSS pHGGBSS-07 PRT-T1-110 N. tobaccum PRT-111 pHG-GBSS pHGGBSS-08 PRT-T1-111 N. tobaccum PRT-112 pHG-GBSS pHGGBSS-09 PRT-T1-112 N. tobaccum PRT-113 pHG-GBSS pHGGBSS-10 PRT-T1-113 N. tobaccum PRT-114 pHG-GBSS pHGGBSS-11 PRT-T1-114 N. tobaccum PRT-115 pHG-GBSS pHGGBSS-12 PRT-T1-115 N. tobaccum PRT-116 pHG-SuSy pHGSuSY-01 PRT-T1-116 N. tobaccum

To be continued

230

Table D1 Index of detailed tobacco transgenic plants (continued)

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPECIE

PRT-117 pHG-SuSy pHGSuSY-02 PRT-T1-117 N. tobaccum PRT-118 pHG-SuSy pHGSuSY-03 PRT-T1-118 N. tobaccum PRT-119 pHG-SuSy pHGSuSY-04 PRT-T1-119 N. tobaccum PRT-120 pHG-SuSy pHGSuSY-05 PRT-T1-120 N. tobaccum PRT-121 pHG-SuSy pHGSuSY-06 PRT-T1-121 N. tobaccum PRT-122 pHG-SuSy pHGSuSY-07 PRT-T1-122 N. tobaccum PRT-123 pHG-SuSy pHGSuSY-08 PRT-T1-123 N. tobaccum PRT-124 pHG-AGPS pHGAGPS-01 PRT-T1-124 N. tobaccum PRT-125 pHG-AGPS pHGAGPS-02 PRT-T1-125 N. tobaccum PRT-126 pHG-AGPS pHGAGPS-03 PRT-T1-126 N. tobaccum PRT-127 pHG-AGPS pHGAGPS-04 PRT-T1-127 N. tobaccum PRT-128 pHG-AGPS pHGAGPS-05 PRT-T1-128 N. tobaccum PRT-129 pHG-AGPS pHGAGPS-06 PRT-T1-129 N. tobaccum PRT-130 pHG-AGPS pHGAGPS-07 PRT-T1-130 N. tobaccum PRT-131 pHG-AGPS pHGAGPS-08 PRT-T1-131 N. tobaccum PRT-132 pHG-AGPS pHGAGPS-09 PRT-T1-132 N. tobaccum PRT-133 pHG-AGPS pHGAGPS-10 PRT-T1-133 N. tobaccum PRT-134 pHG-AGPS pHGAGPS-11 PRT-T1-134 N. tobaccum PRT-135 pHG-AGPS pHGAGPS-12 PRT-T1-135 N. tobaccum PRT-136 pHG-AGPS pHGAGPS-13 PRT-T1-136 N. tobaccum PRT-137 pHG-AGPS pHGAGPS-14 PRT-T1-137 N. tobaccum PRT-138 pHG-IPI pHGIPI-01 PRT-T1-138 N. tobaccum PRT-139 pHG-IPI pHGIPI-02 PRT-T1-139 N. tobaccum PRT-140 pHG-IPI pHGIPI-03 PRT-T1-140 N. tobaccum PRT-141 pHG-IPI pHGIPI-04 PRT-T1-141 N. tobaccum PRT-142 pHG-IPI pHGIPI-05 PRT-T1-142 N. tobaccum PRT-143 pHG-IPI pHGIPI-06 PRT-T1-143 N. tobaccum PRT-144 pHG-IPI pHGIPI-07 PRT-T1-144 N. tobaccum PRT-145 pHG-IPI pHGIPI-08 PRT-T1-145 N. tobaccum

To be continued

231

Table D1 Index of detailed tobacco transgenic plants (continued)

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPECIE

PRT-146 pHG-IPI pHGIPI-09 PRT-T1-146 N. tobaccum PRT-147 pHG-IPI pHGIPI-10 PRT-T1-147 N. tobaccum PRT-148 pHG-IPI pHGIPI-11 PRT-T1-148 N. tobaccum PRT-149 pHG-IPI pHGIPI-12 PRT-T1-149 N. tobaccum PRT-150 pHG-IPI pHGIPI-13 PRT-T1-150 N. tobaccum PRT-151 pHG-MDC pHGMDC-01 PRT-T1-151 N. tobaccum PRT-152 pHG-MDC pHGMDC-02 PRT-T1-152 N. tobaccum PRT-153 pHG-MDC pHGMDC-03 PRT-T1-153 N. tobaccum PRT-154 pHG-MDC pHGMDC-04 PRT-T1-154 N. tobaccum PRT-155 pHG-MDC pHGMDC-05 PRT-T1-155 N. tobaccum PRT-156 pHG-MDC pHGMDC-06 PRT-T1-156 N. tobaccum PRT-157 pHG-MDC pHGMDC-07 PRT-T1-157 N. tobaccum PRT-158 pHG-MDC pHGMDC-08 PRT-T1-158 N. tobaccum PRT-159 pHG-MDC pHGMDC-09 PRT-T1-159 N. tobaccum PRT-160 pHG-MDC pHGMDC-10 PRT-T1-160 N. tobaccum PRT-161 pHG-MDC pHGMDC-11 PRT-T1-161 N. tobaccum PRT-162 pHG-ISPH pHGISPH-01 PRT-T1-162 N. tobaccum PRT-163 pHG-ISPH pHGISPH-02 PRT-T1-163 N. tobaccum PRT-164 pHG-ISPH pHGISPH-03 PRT-T1-164 N. tobaccum PRT-165 pHG-ISPH pHGISPH-04 PRT-T1-165 N. tobaccum PRT-166 pHG-ISPH pHGISPH-05 PRT-T1-166 N. tobaccum PRT-167 pHG-ISPH pHGISPH-06 PRT-T1-167 N. tobaccum PRT-168 pHG-ISPH pHGISPH-07 PRT-T1-168 N. tobaccum PRT-169 pHG-ISPH pHGISPH-08 PRT-T1-169 N. tobaccum PRT-170 pHG-ISPH pHGISPH-09 PRT-T1-170 N. tobaccum PRT-171 pHG-ISPH pHGISPH-10 PRT-T1-171 N. tobaccum PRT-172 pHG-ISPH pHGISPH-11 PRT-T1-172 N. tobaccum PRT-173 pHG-ISPH pHGISPH-12 PRT-T1-173 N. tobaccum PRT-174 pHG-ISPH pHGISPH-13 PRT-T1-174 N. tobaccum

To be continued

232

Table D1 Index of detailed Nicotiana transgenic plants (continued)

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPECIE

PRT-175 pHG-ISPH pHGISPH-14 PRT-T1-175 N. tobaccum PRT-176 pHG-ISPH pHGISPH-15 PRT-T1-176 N. tobaccum PRT-177 pHG-ISPH pHGISPH-16 PRT-T1-177 N. tobaccum PRT-178 pHG-ISPH pHGISPH-17 PRT-T1-178 N. tobaccum PRT-179 pHG-ISPH pHGISPH-18 PRT-T1-179 N. tobaccum PRT-180 pHG-ISPH pHGISPH-19 PRT-T1-180 N. tobaccum PRT-181 pHG-ISPH pHGISPH-20 PRT-T1-181 N. tobaccum PRT-182 pHG-ISPH pHGISPH-21 PRT-T1-182 N. tobaccum PRT-183 pHG-ISPH pHGISPH-22 PRT-T1-183 N. tobaccum PRT-184 pHG-ISPH pHGISPH-23 PRT-T1-184 N. tobaccum PRT-185 pHG-ISPH pHGISPH-24 PRT-T1-185 N. tobaccum PRT-186 pHG-ISPH pHGISPH-25 PRT-T1-186 N. tobaccum PRT-187 pHG-ISPH pHGISPH-26 PRT-T1-187 N. tobaccum PRT-188 pHG-ISPH pHGISPH-27 PRT-T1-188 N. tobaccum PRT-189 pEG-ISPH pEGISPH-01 PRT-T1-189 N. tobaccum PRT-190 pEG-ISPH pEGISPH-02 PRT-T1-190 N. tobaccum PRT-191 pEG-ISPH pEGISPH-03 PRT-T1-191 N. tobaccum PRT-192 pEG-ISPH pEGISPH-04 PRT-T1-192 N. tobaccum PRT-193 pEG-ISPH pEGISPH-05 PRT-T1-193 N. tobaccum PRT-194 pEG-ISPH pEGISPH-06 PRT-T1-194 N. tobaccum PRT-195 pEG-ISPH pEGISPH-07 PRT-T1-195 N. tobaccum PRT-196 pEG-ISPH pEGISPH-08 PRT-T1-196 N. tobaccum PRT-197 pEG-ISPH pEGISPH-09 PRT-T1-197 N. tobaccum PRT-198 pEG-ISPH pEGISPH-10 PRT-T1-198 N. tobaccum PRT-199 pEG-ISPH pEGISPH-11 PRT-T1-199 N. tobaccum PRT-200 pEG-ISPH pEGISPH-12 PRT-T1-200 N. tobaccum PRT-201 pEG-ISPH pEGISPH-13 PRT-T1-201 N. tobaccum PRT-202 pEG-ISPH pEGISPH-14 PRT-T1-202 N. tobaccum PRT-203 pEG-ISPH pEGISPH-15 PRT-T1-203 N. tobaccum

To be continued

233

Table D1 Index of detailed Nicotiana transgenic plants (continued)

PLANT LINE CONSTRUCT PLANT NUMBER SEED SPECIE

PRT-204 pEG-ISPH pEGISPH-16 PRT-T1-204 N. tobaccum PRT-205 pEG-MDC pEGMDC-01 PRT-T1-205 N. tobaccum PRT-206 pEG-MDC pEGMDC-02 PRT-T1-206 N. tobaccum PRT-207 pEG-MDC pEGMDC-03 PRT-T1-207 N. tobaccum PRT-208 pEG-MDC pEGMDC-04 PRT-T1-208 N. tobaccum PRT-209 pEG-MDC pEGMDC-05 PRT-T1-209 N. tobaccum PRT-210 pEG-AGPS pEGAGPS-01 PRT-T1-210 N. tobaccum PRT-211 pEG-AGPS pEGAGPS-02 PRT-T1-211 N. tobaccum PRT-212 pEG-AGPS pEGAGPS-03 PRT-T1-212 N. tobaccum PRT-213 pEG-AGPS pEGAGPS-04 PRT-T1-213 N. tobaccum PRT-214 pEG-AGPS pEGAGPS-05 PRT-T1-214 N. tobaccum PRT-215 pEG-AGPS pEGAGPS-06 PRT-T1-215 N. tobaccum PRT-216 pEG-AGPS pEGAGPS-07 PRT-T1-216 N. tobaccum PRT-217 pEG-AGPS pEGAGPS-08 PRT-T1-217 N. tobaccum PRT-218 pEG-AGPS pEGAGPS-09 PRT-T1-218 N. tobaccum PRT-219 pEG-AGPS pEGAGPS-10 PRT-T1-219 N. tobaccum PRT-220 pEG-AGPS pEGAGPS-11 PRT-T1-220 N. tobaccum PRT-221 pEG-AGPS pEGAGPS-12 PRT-T1-221 N. tobaccum PRT-222 pHG-AGPS pHGAGPS-a PRT-T1-222 LxS8 PRT-223 pHG-AGPS pHGAGPS-b PRT-T1-223 LxS8 PRT-224 pHG-AGPS pHGAGPS-c PRT-T1-224 LxS8 PRT-225 pHG-AGPS pHGAGPS-d PRT-T1-225 LxS8 pRT-226 pHG-SS2 pHG-SS2-a PRT-T1-226 LxS8 PRT-227 pHG-SS3 pHG-SS3-a PRT-T1-227 LxS8 PRT-228 pHG-SS3 pHG-SS3-b PRT-T1-229 LxS8 PRT-229 pHG-SS3 pHG-SS3-c PRT-T1-229 LxS8 PRT-230 pHG-SREBP1 pHG-SREBP1-a PRT-T1-230 LxS8 PRT-231 pHG-SREBP1 pHG-SREBP1-b PRT-T1-231 LxS8 PRT-232 pHG-SREBP1 pHG-SREBP1-c PRT-T1-232 LxS8

234

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