Enhancement of the Free Amino Acid and Protein Content Of

ENHANCEMENT OF THE FREE AMINO ACID AND PROTEIN CONTENT OF

CASSAVA STORAGE ROOTS AND EVALUATION OF ROOT-SPECIFIC

PROMOTERS IN CASSAVA.

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Elisa Leyva-Guerrero

Graduate Program in Plant Cellular and Molecular Biology

The Ohio State University

2011

Dissertation Committee: Dr. Patrice Hamel, Advisor Dr. James Metzger Dr. Richard Sayre, Co-Advisor Dr. Randy Scholl Dr. Dave Somers

Elisa Leyva-Guerrero

2011

ABSTRACT

Cassava is an important staple crop for millions of people around the world in particular in Sub-Saharan Africa. Cassava is preferred given its many agronomical attributes such as ability to grow in poor soils and drought resistance. Cassava storage roots are also a good source of calories (starch), however, they are deficient in protein and other micronutrients such as iron and zinc. Protein malnutrition is widespread in the regions were cassava is widely consumed and consumption of cassava as a staple has been linked to reduced protein intake in the diet. A cassava storage root with higher protein could potentially impact the nutrition and well being of millions people.

There are several proposed strategies to increase the protein and free amino acid content in crops such as modification of the amino acid biosynthetic pathways and over expression of native storage proteins. In this thesis we propose two different approaches for increasing free amino acids and protein. One approach was to increase nitrate assimilation through the expression of a mutated nitrate reductase. Nitrate reduction to nitrite is followed by a further reduction to ammonium which can be readily incorporated into amino acids. A rate limiting step in nitrate metabolism is the reduction of nitrate to nitrite by nitrate reductase. This enzyme is highly regulated, however, it has been observed that through mutation of a key regulatory serine the enzyme remains active and the assimilation of nitrate increases resulting in increased free amino acid levels. In

cassava the root-specific expression of a mutated nitrate reductase resulted in a the doubling of the storage root free amino acid content, however, no root protein increase was observed. A second approach was to utilize the nitrile group present in the cassava cyanogenic glucoside linamarin as a reduced nitrogen source. The hydrolysis of linamarin releases acetone cyanohydrin which in turn can degrade to release cyanide. In all plants there is a cyanide assimilation pathway involving β-cyanoalanine synthase, the end products of this pathway are aspartate and ammonia. Through increased hydrolysis of linamarin, we sought to increase the assimilation of cyanide in to aspartate and ammonia.

Both of these compounds can be utilized for the synthesis of additional amino acids. The linamarase enzyme is responsible for linamarin hydrolysis. In wild-type cassava the interaction between linamarase and linamarin is limited spatially; linamarase is found in the cell wall and linamarin in the vacuole. Through the expression of a vacuolar targeted linamarase, we proposed to increase the hydrolysis of linamarin and as a result provide more reduced nitrogen (nitrile) for free amino acid and protein synthesis. The expression of a vacuolar linamarase in cassava storage roots resulted in a doubling of the free amino acid pool in this organ but no increase in protein. It was only through the dual expression of vacuolar linamarase and the storage protein sporazein that both an increase in free amino acid and protein in the storage roots was observed. Interestingly the expression of a mutated nitrate reductase and the expression of a vacuolar linamarase with or without sporazein in the cassava storage roots resulted in an increase in free amino acid and protein levels in the leaves. This indicates the presence of amino acid transport from the

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storage roots to a strong nitrogen utilization organ in this case the leaves where the amino acids are incorporated in to protein. Importantly even with elevated root expression of sporazein the leaves had increased free amino acids and protein indicating that the expression of a root storage protein is not sufficient to substantially reduce leaf nitrogen sink strength.

Although cassava is a main staple food the molecular biology tools available for its transformation are very limited. In the final chapter of this thesis we evaluated four different Arabidopsis root promoters in cassava to determine their functionality and tissue specificity. Two promoters named A14 and E40 were determined to be functional in cassava roots with minimal leaf expression. The availability of an increased set of root promoters for cassava may increase the development of transgenic cassava with increased nutritional and industrial value.

DEDICATION

Too the most wonderful and dedicated parents in the world

ACKNOWLEDGEMENTS

I thank first and beyond anyone God, for giving me life, for giving me a wonderful family and friends and for being a constant presence and a source of support.

I thank my two wonderful and dedicated parents Roberto and Rosa Maria for it is because of them I have made it this far. I am a regular person who had great parents believing in her, believing that I could make it this far. Thank you for everything.

I thank my advisor Dr. Richard Sayre for taking me in his lab, for helping me grow and being there throughout the years. Thank you for allowing me to pursue this dream and for all your guidance.

I thank my little sister Alejandra for always being a loving sister and someone who helped me keep my spirits high at all times. I thank my very soon to be husband Andy for all his love in this last years and specially in these last months, for taking care of little things and big things and making a big difference in my life.

I thank all my grandmothers Pita and Virginia and my grandfathers Roberto and Jacobo for their presence in life or from heaven, thank you for your love. I thank all my family, my aunts and uncles, all my cousins, thank you for always believing in me.

I thank Adriana for her friendship throughout all these years, for your kindness and for always being there for me. I thank Luis for his friendship in grad school and beyond, for always listening to me no matter what time of day or length of phone call, for making me laugh whether it was intentional or not. I thank Yuriko for being my surrogate little sister when our own sisters were away, thank you for your friendship.

I thank my labmates that are more than that, my friends Vanessa, Zoee, Anil and

Narayanan, I don’t know how I could have made it without you guys, thank you in particular to Narayanan for all his guidance and help with my projects and Vanessa for reading my thesis at the last minute.

I thank all those that have gone through the Sayre Lab, those that have taught me and those that have allowed me to teach, in particular to Uzo for teaching me cassava subculturing. I would like to thank also Dr. Nigel Taylor for teaching me cassava techniques.

I thank the Keppler family for making me feel so welcomed and for all the family outings that have made these last years so much more enjoyable. To all my friends a long long

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list to go through, but I thank my friends in Mexico for their long distance friendship and your encouragement as always on facebook or by email, you don’t know how much it meant to me. I thank Ro for always caring, for being there no matter what, for letting me vent and listening. To Niro for being my first friend in Columbus and for always being so kind. I thank the Jose’s and Sergio for the dancing and partying nights that made life in

Columbus all the much livelier, Roxana for being my super concert friend and for letting me stay with her all the time in Columbus and because no matter what we had to do we always managed to sneak to Chicago to see Julieta. To Shely and her family, the Zakins and the Fans for opening your homes to me. To Nadja and Xuan, for being my first friends in St. Louis. To all my friends in St. Louis and all over the world! I thank also all my friends in dachshund and dog rescue land.

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VITA

1981…………Born- San Luis Potosi, SLP, Mexico

2004………….B.S. Chemistry, Universidad Autonoma de San Luis Potosi

2004-2011……Graduate Research Associate, The Ohio State University

FIELD OF STUDY

Major Field: Plant Cellular and Molecular Biology

TABLE OF CONTENTS

ABSTRACT ...... ii

DEDICATION ...... v

ACKNOWLEDGEMENTS ...... vi

VITA ...... ix

FIELD OF STUDY ...... ix

LIST OF FIGURES……………………………………………………………………..xix

LIST OF TABLES…………………………………………………………………..…xxiv

CHAPTER 1 INTRODUCTION ...... 1

1.1 BIOLOGY OF CASSAVA ...... 1

1.2 CASSAVA IN AFRICA ...... 7

1.3 CYANOGENESIS OF CASSAVA ...... 11

1.4 LINAMARIN TRANSPORT ...... 16

1.5 CYANIDE ASSIMILATION AND DETOXIFICATION PATHWAYS ...... 19

1.6 FUNCTION OF CYANOGENIC GLUCOSIDES ...... 23

1.7 DIETARY CYANIDE CONSUMPTION ...... 24

1.7.1 Konzo...... 25

1.7.2 Tropical Ataxic Neuropathy ...... 25

1.7.3 Goiter ...... 26

1.8 CASSAVA FOOD PROCESSING ...... 26

1.8.1 Peeling ...... 27

1.8.2 Boiling ...... 27

1.8.3 Sun-drying ...... 27

1.8.4 Fermentation ...... 28

1.8.5 Grating and Soaking ...... 28

1.9 PROTEIN CONTENT IN CASSAVA ...... 30

1.10 GENETIC TRANSFORMATION OF CASSAVA ...... 34

1.11 OBJECTIVES ...... 39

CHAPTER 2 EXPRESSION OF A MUTATED NITRATE REDUCTASE TO

INCREASE THE FREE AMINO ACID AND PROTEIN CONTENT IN CASSAVA

STORAGE ROOTS ...... 40

2.1 INTRODUCTION ...... 40

2.2 MATERIALS AND METHODS ...... 44

2.2.1 E.coli and Agrobacterium strains ...... 44

2.2.2 Chemical reagents and supplies ...... 44

2.2.3 Cassava cultivar, in vitro and greenhouse growth ...... 44

2.2.4 Nitrate reductase gene ...... 45

2.2.5 General molecular techniques ...... 45

2.2.6 Codon optimization ...... 46

2.2.7 Cloning of XI-NIA in to the pCAMBIA2300 binary vector ...... 53

2.2.8 Agrobacterium transformation ...... 55

2.2.9 Generation of Friable Embryogenic Callus ...... 56

2.2.10 Transformation of cassava ...... 59

2.2.11 Cassava DNA extraction and transformation validation with PCR ...... 59

2.2.12 Dot blot analysis of putative transgenic plants ...... 60

2.2.13 Cassava root RNA extraction ...... 63

2.2.14 Semiquantitative RT-PCR of the XI-NIA gene ...... 64

2.2.15 Real-time RT-PCR analysis of XI-NIA expression ...... 65

2.2.16 Linamarin determination in storage root and leaf ...... 67

2.2.16.1 Standard curve of linamarin and internal standard ...... 68 2.2.16.2 Tissue sample extraction for linamarin quantification ...... 68 2.2.16.3 Derivatization of linamarin and internal standard ...... 69 2.2.16.4 Selection of ion for single ion monitoring of linamarin content ...... 69 2.2.16.5 Linamarin Standard curve ...... 73 2.2.17 Free amino acid extraction and analysis from storage root and leaf ...... 77

2.2.18 Total protein analysis of storage root pulp and leaf ...... 77

2.2.19 Western blot of nitrate reductase in CNIA and wild-type plants ...... 78

+ 2.2.20 Growth experiment on free NH4 media ...... 81

2.2.21 Analysis of calorie content in cassava storage roots ...... 81

2.2.22 Statistical analysis...... 82

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2.3 RESULTS...... 83

2.3.1 Codon optimization and mutation of Arabidopsis NIA2 ...... 83

2.3.2 Construction of the pCNIA vector ...... 88

2.3.3 Production of transgenic cassava ...... 90

2.3.4 PCR and dot-blot analysis ...... 92

2.3.5 Analysis of cassava storage roots by semiquantitative RT-PCR and real-time

RT-PCR ...... 94

2.3.6 Harvest weight ...... 98

2.3.11 Western Blot ...... 103

2.3.7 Storage Root Free Amino Acid Pool analysis ...... 105

2.3.8 Leaf Total Free Amino Acid Pool ...... 121

2.3.9 Protein analysis ...... 124

2.3.10 Linamarin analysis ...... 128

2.3.12 Growth experiment in ammonia free media ...... 133

2.3.13 Analysis of calorie content in cassava storage roots ...... 137

2.3.14 Summary of CNIA lines ...... 139

2.4 DISCUSSION ...... 141

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CHAPTER 3 EXPRESSION OF A VACUOLAR TARGETED LINAMARASE AND

THE STORAGE PROTEIN SPORAZEIN TO INCREASE THE FREE AMINO ACID

POOL AND PROTEIN CONTENT IN CASSAVA STORAGE ROOTS ...... 150

3.1 INTRODUCTION ...... 150

3.2 MATERIALS AND METHODS ...... 158

3.2.1 E.coli and Agrobacterium strains ...... 158

3.2.2 Chemical Reagents and supplies ...... 158

3.2.3 Cassava cultivar and in vitro and greenhouse growth ...... 158

3.2.4 General molecular techniques ...... 159

3.2.5 Cloning of the linamarase gene ...... 160

3.2.6 Determination of the signal peptide...... 164

3.2.7 Cloning of an ΔN-ter linamarase ...... 164

3.2.8 Cloning of a vacuolar targeted linamarase ...... 167

3.2.9 Cloning of the pCambia2300-VL and pCambia2300-SVL vectors ...... 170

3.2.10 Generation of Friable Embryogenic Callus ...... 171

3.2.11 Transformation of cassava ...... 172

3.2.12 DNA extraction and PCR of putative transgenic ΔN-linamarase cassava lines

...... 173

3.2.13 DNA extraction and PCR of putative transgenic VL and SVL cassava lines

...... 174

3.2.14 Dot blot analysis of putative VL and SVL transgenic cassava plants ...... 175

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3.2.15 Harvest weight ...... 175

3.2.16 RNA extraction and cDNA synthesis from cassava storage roots ...... 175

3.2.17 Semiquantitative RT-PCR analysis of vacuolar linamarase and sporazein

expression ...... 176

3.2.18 Linamarin determination ...... 177

3.2.18.1 Standard curve ...... 177 3.2.18.2 Tissue sample extraction ...... 178 3.2.18.3 Derivatization of linamarin and internal standard ...... 179 3.2.19 Free amino acid extraction and analysis ...... 179

3.2.20 Total protein analysis...... 179

3.2.21 Western blot ...... 179

3.2.22 Total nitrogen content ...... 182

3.3 RESULTS...... 183

3.3.1 Cloning of a ΔN-ter linamarase and a vacuolar linamarase ...... 183

3.3.2 Cloning of the pCambia2300-VL and pCambia2300-SVL vectors ...... 187

3.3.4 Generation of transgenic cassava expressing ΔN-ter linamarase ...... 189

3.3.5 Generation of transgenic cassava VL and SVL lines ...... 191

3.3.6 Greenhouse growth of wild-type, VL and SVL lines ...... 195

3.3.7 Analysis by semiquantitative RT-PCR of vacuolar linamarase and sporazein

expression...... 202

3.3.8 Western blot ...... 204

3.3.9 Linamarin content analysis of storage root and leaf ...... 206 xv

3.3.10 Total free amino acid content in leaf and storage roots ...... 215

3.3.11 Individual free amino acid analysis in the storage root pulp ...... 219

3.3.12 Individual free amino acid analysis in leaves ...... 234

3.3.13 Protein content ...... 246

3.3.14 Total Nitrogen analysis of leaves and storage root ...... 250

3.3.15 Summary of VL and SVL lines ...... 255

3.4 DISCUSSION ...... 258

3.5 Summary table of strategies for the increase of free amino acid and protein content

in cassava storage roots ...... 266

CHAPTER 4 ALTERNATIVE ROOT GENE PROMOTERS FOR EXPRESSION OF

TRANSGENES IN CASSAVA STORAGE ROOTS ...... 268

4.1 INTRODUCTION ...... 268

4.2 MATERIALS AND METHODS ...... 285

4.2.1 E.coli and Agrobacterium strain ...... 285

4.2.2 Chemical Reagents ...... 285

4.2.3 Cassava cultivar and in vitro and greenhouse growth ...... 285

4.2.4 Bioinformatic analysis of the Arabidopsis promoters ...... 286

4.2.5 Arabidopsis promoters and entry vector ...... 286

4.2.6 Codon optimization and gene synthesis ...... 290

4.2.7 Destination vector ...... 290

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4.2.8 Expression vector ...... 298

4.2.9 Generation of Friable Embryogenic Callus ...... 299

4.2.10 Transformation of cassava ...... 299

4.2.11 DNA extraction and PCR ...... 300

4.2.12 Dot blot ...... 301

4.2.13 Microscopy ...... 302

4.2.14 RNA extraction ...... 303

4.2.15 Semiquantitative RT-PCR ...... 305

4.2.16 Real-time RT-PCR ...... 307

4.3 RESULTS...... 309

4.3.1 Arabidopsis promoters ...... 309

4.3.2 Codon optimization ...... 309

4.3.3 Expression system ...... 312

4.3.4 Generation of transgenic cassava ...... 316

4.3.5 Characterization of transgenics by PCR, dot-blot and widefield fluorescence

...... 318

4.3.6 Analysis of young cassava plants by semi-quantitative RT-PCR ...... 324

4.3.7 Analysis of young cassava plants by confocal microscopy ...... 326

4.3.8 Analysis of cassava leaves and storage roots by semi-quantitative RT-PCR . 330

4.3.9 Real-time RT-PCR analysis ...... 333

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4.3.10 Identification of cis-acting elements and transcription start site ...... 340

4.4 DISCUSSION ...... 344

BIBLIOGRAPHY ...... 350

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LIST OF FIGURES

1.1 Images of Manihot esculenta Crantz ...... 2 1.2 Cassava root images ...... 4 1.3 Images of cassava stems ...... 6 1.4 Worldwide production of cassava ...... 8 1.5 Sub-Saharan Africa cultivation of cassava ...... 10 1.6 Synthesis of linamarin in cassava ...... 12

1.7 Cyanogenesis of linamarin ...... 15 1.8 The linustatin degradation pathways ...... 18 1.9 Cyanide assimilation pathway or β-cyanoalanine synthase ...... 20

1.10 Cyanide detoxification or rhodanese pathway ...... 22 1.11 Different cassava processing techniques ...... 29 1.12 Protein and energy consumed per capita ...... 32

1.13 Transformation systems for cassava ...... 35 2.1 Outline of a fusion PCR reaction used for codon optimization ...... 47 2.2 Cassava transformation tissue culture images ...... 58

2.3 GC-MS analysis of linamarin and phenyl-β-glucopyranoside ...... 71 2.4 Mass spectrometer scan of linamarin and phenyl-β-glucopyranoside ...... 72 2.5 Standard curve of linamarin and phenyl-β-glucopyranoside ...... 76

2.6 Fusion PCR reaction ...... 84 2.7 Gene sequence alignment of Arabidopsis NIA2 and XI-NIA ...... 85 2.8 Protein sequence alignment of Arabidopsis NIA2 and XI-NIA ...... 87

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2.9 T-DNA region of pCNIA vector ...... 89 2.10 Schematic of cassava regeneration after transformation ...... 91 2.11 PCR screening of in vitro transgenic CNIA plantlets ...... 93

2.12 RT-PCR analysis of XI-NIA expression in storage roots ...... 95

2.13 Real-time RT-PCR analysis of CNIA lines ...... 97 2.14 Storage root harvest weight of CNIA lines ...... 99 2.15 Shoot harvest weight of CNIA lines ...... 101

2.16 Images of cassava storage roots from CNIA lines and wild type ...... 102 2.17 Western blot analysis of CNIA lines ...... 104 2.18 Total free amino acids in storage root of wild type and CNIA lines ...... 106

2.19 The glutamate family in the free amino acid pool of CNIA and wild type...... 110

2.20 The aspartate family in the free amino acid pool of CNIA and wild type ...... 113 2.21 The alanine/valine/leucine and serine in the free amino acid pool of CNIA and wild type ...... 116 2.22 The aromatic group and histidine in the free amino acid pool of CNIA and wild type ...... 119

2.23 Total free amino acids in leaves of wild type and CNIA lines ...... 123

2.24 Total soluble protein in cassava storage root of CNIA lines and wild type ...... 126 2.25 Total soluble protein in cassava leaves of CNIA lines and wild type ...... 127 2.26 Linamarin content in storage root of CNIA lines and wild type ...... 130

2.27 Linamarin content in leaves of CNIA lines and wild type ...... 132

2.28 In vitro growth experiment of CNIA lines and wild type in media with or without reduced nitrogen ...... 133 2.29 Dry weight of CNIA lines and wild type grown in different media with or without reduced nitrogen ...... 136 2.30 Calorie content of CNIA lines and wild type storage roots ...... 138

2.31 Summary of CNIA lines ...... 140

3.1 Cyanide assimilation and detoxification pathways in plants ...... 153

3.2 T-DNA region of the 3Dfls plasmid ...... 163

3.3 T-DNA region of the 3Dls plasmid ...... 166 3.4 T-DNA region of the 3DVL plasmid ...... 169

3.5 Cloning of a ΔN-ter linamarase and a vacuolar targeted linamarase ...... 185

3.6 Prediction of the signal peptide of linamarase ...... 186 3.7 T-DNA region of the pCambia2300-VL vector ...... 188 3.8 T-DNA region of the pCambia2300-SVL vector ...... 188

3.9 PCR analysis of putative transgenic cassava lines transformed with the 3Dls vector ...... 190

3.10 Dot blot analysis of SVL and VL lines ...... 192 3.11 Images of VL and SVL roots at 4 months of greenhouse growth ...... 196 3.12 Storage root images from select VL and SVL lines ...... 198

3.13 Storage root fresh weight ...... 200

3.14 Shoot fresh weight ...... 201 3.15 Semiquantitative RT-PCR analysis of VL and SVL lines ...... 203 3.16 Western blot analysis of VL and SVL ...... 205

3.17 Single ion chromatogram for linamarin from wild type sample ...... 207

3.18 Single ion chromatogram for PGP from wild type sample ...... 208

3.19 Single ion chromatogram for linamarin from VL-04 line sample ...... 209

3.20 Single ion chromatogram for PGP from VL-04 line sample ...... 210

3.21 Storage root linamarin from wild type and transgenic VL and SVL ...... 212

3.22 Leaf linamarin from wild type and transgenic VL and SVL ...... 214 3.23 Total free amino acid content in storage roots of wild type and transgenic VL and SVL ...... 216 3.24 Total free amino acid in the leaf of wild type and transgenic VL and SVL ...... 218

3.25 Analysis of free amino acids from the glutamate family of wild type and transgenic VL and SVL lines storage root ...... 221 3.26 Analysis of the free amino acids from the aspartate family of wild type and transgenic VL and SVL lines storage root ...... 225

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3.27 Analysis of the free amino acids from the alanine/valine/leucine group and serine/cysteine/glycine group in wild type and transgenic VL and SVL lines roots ...... 228

3.28 Analysis of the free amino acids from the aromatic group and histidine in wild type and transgenic VL and SVL lines roots ...... 231

3.29 Analysis of the free amino acids from the glutamate family in wild type and transgenic VL and SVL leaves ...... 238

3.30 Analysis of the free amino acids from the aspartate family of wild type and transgenic VL and SVL lines leaves ...... 240 3.31 Analysis of the free amino acids from the alanine/valine/leucine group and serine/cysteine/glycine in wild type and transgenic VL and SVL leaves ...... 242

3.32 Analysis of the free amino acids from the aromatic group in wild type and transgenic VL and SVL leaves ...... 244

3.33 Storage root protein analysis in wild type and transgenic VL and SVL ...... 247

3.34 Leaf protein analysis in wild type and transgenic VL and SVL lines ...... 249

3.35 Total moles of reduced nitrogen in wild type and transgenic VL and SVL……251 3.36 Summary of VL and SVL lines ...... 256

4.1 Expression maps of Arabidopsis root promoters ...... 271

4.2 A14 promoter sequence ...... 273

4.3 E40 promoter sequence ...... 275

4.4 E49 promoter sequence ...... 279

4.5 S8 promoter sequence ...... 281 4.6 Codon usage in YFP and CFP compared to the cassava codon usage table ...... 310

4.7 YFP and CFP codon optimized sequence ...... 311

4.8 Gateway recombination vectors ...... 313

4.9 Restriction digestion analysis of Arabidopsis promoters Gateway vectors ...... 315

4.10 PCR analysis of putative transgenic A14:CFP, E40:CFP, E49:CFP and S8:CFP plants ...... 319 4.11 Analysis of YFP and CFP fluorescence in A14:CFP, E40:CFP, E49:CFP and S8:CFP fibrous roots ...... 321

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4.12 Comparison of insertion number vs fluorescence in A14:CFP, E40:CFP, E49:CFP and S8:CFP lines ...... 323

4.13 Semiquantitative RT-PCR analysis of A14:CFP, E40:CFP, E49:CFP and S8:CFP lines ...... 325

4.14 Confocal microscopy analysis of A14:CFP, E40:CFP, E49:CFP and S8:CFP plants ...... 327 4.15 Z-stack scan of A14-60 fibrous root ...... 329

4.16 Semiquantitative RT-PCR analysis of greenhouse grown A14:CFP, E40:CFP, E49:CFP and S8:CFP lines ...... 332

4.17 Real-time RT-PCR analysis of CFP expression driven by the E49 promoter .....334

4.18 Real-time RT-PCR analysis of CFP expression driven by the S8 promoter ...... 335

4.19 Real-time RT-PCR analysis of CFP expression driven by the A14 promoter .....338

4.20 Real-time RT-PCR analysis of CFP expression driven by the E40 promoter .....339

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LIST OF TABLES 2.1 Primers designed for the synthesis of mutated codon optimized NIA2...... 49 2.2 Fusion PCR primer pairs ...... 50

2.3 Linamarin standard curve ...... 74

3.1 T-DNA insertions of VL and SVL lines ...... 194

3.2 Storage root individual free amino acids in wild type, VL and SVL lines ...... 233

3.3 Leaf individual free amino acids in wild type, VL and SVL lines ...... 235

3.4 Total nitrogen in the storage roots of WT, VL and SVL lines ...... 254

3.5 Summary of results from CNIA, VL and SVL lines ...... 267

4.1 Expression tissues of A14, E40, E49 and S8 promoters ...... 270

4.2 Total promoter lines regenerated ...... 317

4.3 Representative cis-motifs of the A14 and E40 promoters ...... 342 4.4 Representative cis-motifis of the E49 and S8 promoters ...... 343

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CHAPTER 1 INTRODUCTION

1.1 BIOLOGY OF CASSAVA

Cassava (Manihot esculenta Crantz) is a shrubby woody perennial belonging to the

Euphorbiaceae family (Fig. 1.1). It was originally domesticated in South America

(Allem, 2002) although is now cultivated around the world. It is known by several common names such as cassava, yucca, mandioca and manioc. Cassava cultivation takes place between 30˚ North latitude and 30˚ South latitude. It grows in a broad range of temperatures from 18 to 35˚C but the ideal temperature is 25 to 28˚C. The annual rainfall requirement averages 1000-1500 mm, however, cassava is drought tolerant and will produce a crop in regions with precipitations as low as 600 mm/year such as the environments of Northeast Brazil. It will also grow in wet tropical areas with precipitation as high as 4000 mm/year, e.g., in Indonesia (Embrapa, 2005).

Figure 1.1 Images of Manihot esculenta Crantz (A) Line art drawing of cassava

(Reproduced with permission from Wikimedia Commons). (B) Cassava in a field in

Tanzania (Reproduced with permission from Kevin H. photography)

Both the leaves and storage roots are consumed but there is marked preference for consumption of the root in most societies. The cassava storage root system starts out similar to that of other dicots, a fibrous tap root system. As it matures it converts into a storage root associated with the swelling of fibrous roots (DeSouza, 2001) (Fig. 1.2A).

The formation of storage roots involves both radial and elongated growth (De Souza et al., 2004). Cassava storage roots can be divided into three tissue systems (De Souza,

2001) (Fig. 1.2B). The Tissue System I (TSI) is comprised of the phellogen and phelloderm, this system is also known as the peel of the storage root. The Tissue System

II (TSII) is composed of secondary phloem and vascular cambium, and Tissue System III

(TSIII) is comprised of secondary xylem and a large volume of starchy parenchyma cells.

The TSII and TSIII are collectively known as the pulp of the root.

The shoot of the cassava plant can grow up to five meters in height. The size of the leaves and petioles vary depending on the cultivar, age and environmental factors but commonly leaves will have five to seven lobes and follow a 2:5 phyllotaxy ratio (Embrapa 2005).

Figure 1.2 Cassava root images (A) Whole cassava storage roots (reproduced with permission of Captain of Green and White photography) and (B) diagram of a cassava storage root cross section. TSI= Tissue system I also known as the peel, TSII=Tissue

System II and TSIII= Tissue System III, TSII and TSIII are collectively known as the pulp.

Cassava is most frequently propagated clonally using stem cuttings (Fig. 1.3). An advantage of its clonal propagation is higher yield. Cassava plants grown from stem cuttings have the formation of one or more storage roots at the meristems of each of the nodes buried in the soil, while plants grown from true seeds commonly have one storage root only. (McKey 2010, review). As a result, farmers have selected for thick stemed varieties having enlarged meristems which are conducive to clonal propagation (McKey,

2010 review).

Figure 1.3 Images of Cassava stems 1 Cassava stem cuttings are used for propagation

(Reproduced with permission of International Institute of Tropical Agriculture).

1.2 CASSAVA IN AFRICA

The domestication of cassava took place in South America (Allem, 2002) and cassava continues to be grown in South and Central America as well as the Caribbean. During the colonial times the Portuguese introduced cassava into the African continent (Rogers and

Appan, 1973) and during colonial times. In some countries such as Zambia, cassava cultivation was required by the government as a famine reserve or food security crop in case maize or other crops failed due to drought (Jones, 1959). However, throughout the years, cassava has gained importance in Sub-Saharan Africa for many reasons. Cassava is more drought tolerant, has higher starch content and herbivore resistance (due to the presence of cyanogenic glucosides) than other major staple crops. In addition cassava roots may remain underground for several years providing food security in case of drought or times of conflict (Nweke et al., 2002; Haggblade and Zulu, 2003). More recently the increase of HIV/AIDS throughout the African continent has reduced the adult labor force resulting in increased cultivation of cassava which has reduced labor inputs relative to other crops (Haggblade and Zulu, 2003).

The African continent accounts for the largest percentage (47.5%) of global cassava production. Asia is second at 30.99%, and the Americas produce 13.1% of the total global yield (FAOSTAT, 2009) (Fig. 1.4, blue bar graph). Cassava yields per hectare in

Africa, however are, the lowest in the world at 10.1 metric tons/hectare while in Asia yields average 20.2 metric tons/ hectare (FAOSTAT, 2009) (Fig. 1.4, green bar graph).

Figure 1.4 Worldwide production of cassava. Graph depicting the cassava production on each continent as a percentage of the total (blue bar graph and left y-axis). (green bar graph and right y-axis).

Cassava production in Sub-Saharan Africa has steadily grown, over the last 50 years. The total area under cultivation for cassava in Sub-Saharan Africa grew by 221% (FAOSTAT

2009) (Fig. 1.5A) while the population of this region has increased by 250%. The Sub-

Saharan country of Nigeria is the world’s largest producer of cassava. Over the last 50 years cassava cultivation increased by 484% in Nigeria (Fig. 1.5B). The per capita consumption of cassava in Nigeria is one of the highest in the world at 114 kg/year

(2007) providing 252 kcal/day/person (FAOSTAT, 2007). The Democratic Republic of

Congo is second largest total producer of cassava in Sub-Saharan Africa, but the highest per capita consumer at 287 kg/year providing 861 kcal/day/person (FAOSTAT, 2007).

Interestingly in Malawi the area under cassava cultivation has increased by 871%, associated in part with the food security it provides (FAOSTAT, 2007).

Figure 1.5 Sub-Saharan Africa cultivation of cassava. (A) Hectares cultivated in Sub-

Saharan Africa over the last 50 years. (B) Hectares cultivated in Nigeria (black) and

Democratic Republic of Congo (red) over the last 50 years. Data obtained from

FAOSTAT, www.faostat.fao.org.

1.3 CYANOGENESIS OF CASSAVA

Nearly 2500 plant species produce cyanogenic glucosides, but cassava is the most important cyanogenic staple crop among these (Jones 1998). Cyanogenic glycosides are glucosides of ɑ-hydroxinitriles derived from amino acids (Vetter 2000). All tissues of cassava, with the exception of the seeds, are cyanogenic (McMahon et al. 1995, review)

Cassava contains two cyanogenic glucosides linamarin and lotoaustralin, derived from valine and isoleucine, respectively: linamarin accounts for 95% of the total cyanogenic glucoside content (Cock 1985, Balogopalan et al. 1988). Cassava varieties can be classified according to the cyanogenic potential of their storage roots. A fresh cassava storage root is considered as low cyanogenic or “sweet” if it contains less than 100 mg cyanogens /kg fresh weight and as high cyanogenic or “bitter” if they contain 100 – 500 mg cyanogens/kg fresh weight (Wheatley et al. 1993). Values over 500 mg cyanogens/kg fresh weight are considered “very bitter” (Montagnac et al. 2009). The leaves contain 5 to

20 times higher cyanogen levels than storage roots and can reach up to up to 5 g cyanogens/kg fresh weight (Montagnac et al. 2009).

The biosynthesis pathway of linamarin and lotoaustralin proceeds in a manner similar to that of other cyanogenic glucosides such as dhurrin, the cyanogenic glucoside present in sorghum. In sorghum two membrane bound cytochromes P450, named CYP79A1 and

CYP71E1, and a soluble UDPG-glycosyltransferase, are involved in the synthesis of dhurrin from tyrosine (McFarlene et al. 1975, Conn 1980, Jones et al 1999). In cassava two similar CYP79 enzymes named CYP79D1 and CYP79D2 were identified by

Andersen et al. (2000) as the enzymes responsible for synthesizing the oxime from the amino acid (Fig. 1.6). Recently, Jorgensen et al. (2010) identified CYP71E7 which codes

for the enzyme responsible for converting the oximes into their corresponding cyanohydrins.

Figure 1.6 Synthesis of linamarin in cassava. The cyanogenic glucoside linamarin is derived from the amino acid L-valine. CYP79D1/D2 are responsible for catalyzing the conversion of L-valine to its corresponding oxime, 2-methyl propanol oxime. The

CYP71E7 cytochrome P450 is involved in the synthesis of acetone cyanohydrin from the oxime, finally a UDPG-glucosyltransferase condenses acetone cyanohydrin with glucose to obtain linamarin.

The release of cyanide from linamarin is a two-step process (Fig. 7). The first step involves the hydrolysis of linamarin by the β-glucosidase, linamarase. Linamarase is a 65 kDa enzyme (Mkpong et al. 1990) with three isoforms (Eksittikul and Chulavatnatal,

1988) that belongs to the family of ɑ-1 glycoside hydrolases (Henrissat and Bairoch,

1996). Linamarase catalyzes the hydrolysis of β-glucosidic linkages of single glucosides such as linamarin. The hydrolysis of linamarin yields acetone cyanohydrin and glucose.

The second step of cyanogenesis involves the hydrolysis of acetone cyanohydrin to cyanide and acetone (Fig. 1.7). This reaction occurs spontaneously at pH > 5 or high temperatures >35˚C or by catalysis by hydroxynitrile lyase (HNL). HNL enzyme is a

28.5 kDa protein expressed predominantly only in leaves (White et al. 1998). In contrast, linamarase is expressed at high levels in all tissues.

In cassava, as in other cyanogenic crops, there is compartmentalization between the cyanogenic glucosides and their corresponding β-glucosidase and hydroxynitrile lyase

(Grunhert et al., 1994; White et al,. 1994). The physical separation between the cyanogenic glucoside and the enzymes involved in their degradation limits the occurrence of cyanogenesis (Robinson, 1930; Conn, 1981). In cassava the linamarin is stored in the vacuole (White et al. 1994); as for the enzymes, both linamarase and HNL have been localized to the apoplast and laticifers (Mkpong et al., 1990; Pancoro et al.,

1992; White et al., 1998).

Figure 1.7 Cyanogenesis of linamarin. The release of cyanide from linamarin occurs by the hydrolysis of linamarin by the enzyme linamarase, followed by either the spontaneous degradation of acetone cyanohydrin (high pH>5, high T> 35˚C) or by action of the enzyme hydroxynitrile lyase (HNL).

1.4 LINAMARIN TRANSPORT

The synthesis of linamarin was proposed to occur primarily in the leaves, where studies have shown that the incorporation of 14C-valine to 14C-linamarin occurs more efficiently in leaves rather than roots (Bedakio et al. 1981). Active linamarin synthesis is possible in roots but the magnitude of its contribution to root steady-state linamarin levels was still in question (McMahon et al. 1994, Du et al. 1995). If linamarin is synthesized in the leaves of cassava this would require the transport of linamarin to the roots. Evidence of the transport of linamarin was indicated by the detection of linamarin in phloem exudates, as well as evidence from stem girdling experiments in which linamarin accumulated at the site above the girdle (De Bruijin, 1975; Nambisan et al., 1996). In addition Makame et al.

(1987) observed that in reciprocal stem grafting experiments of cassava varieties of high and low cyanide the cyanogenic glucosides accumulated in the root were due at least in part to transport from the leaves. The latest evidence of linamarin transport was provided by Siritunga et al. (2003). By blocking the synthesis of linamarin in the leaves by RNAi technology, the linamarin content in the root tissues dropped to 1% of that of wild-type.

A possible explanation is that the reduction of linamarin synthesis in the leaves results in reduced transport of linamarin to the roots, and therefore lowers linamarin accumulation in the storage roots.

The presence of linamarase and HNL in the apoplast of leaf tissues restrictions possible transport routes for linamarin. One possible transport pathway is symplastic transport of linamarin (Nambisan et al., 1996); the other is via the linustatin pathway (Selmar et al.,

1988). The transport of linamarin via the linustatin pathway has been shown to occur in

Hevea brasilensis (Lieberie et al., 1985; Selmar 1988). Linustatin is the glucoside of

linamarin, and its proposed pathway involves the conversion of linamarin to linustatin in the leaves. This diglucoside cannot be hydrolyzed by cassava linamarase (Yeoh and Woo,

1992), and so linustatin could be transported from the leaves to the roots apoplastically without being degraded by linamarase. Once linustatin arrives at the root it could be sequentially or simultaneously hydrolyzed by a diglucosidase (Selmar, 2010) (Fig. 1.8).

The cleavage of one glucose from linustatin generates linamarin, while simultaneous cleavage of linamarin produces gentobiose and acetone cyanohydrin. The later product can be degraded spontaneously at high pH or high temperatures, which would result in the release of cyanide. The released cyanide could potentially enter cyanide assimilation or detoxification pathways, resulting in the synthesis of aspartate and ammonia, or thiocyanate, respectively. (See Section 1.5 Cyanide Assimilation). There is some evidence for the transport of linamarin in cassava through the linustatin pathway (Selmar,

1994), however, to date, however, no linustatin was detected in cassava in other studies

(Siritunga 2002).

Figure 1.8 The linustatin degradation pathways. Simultaneous degradation releases acetone cyanohydrin that may be further degraded to cyanide and acetone. Sequential degradation releases linamarin and glucose.

1.5 CYANIDE ASSIMILATION AND DETOXIFICATION PATHWAYS

Cyanide can potentially inhibit a variety of important enzymes including, cytochrome C oxidase, Cu/Zn superoxide dismutase, catalase, nitrate/nitrite reductase, nitrogenase, and peroxidase (Grossmann 2003). Importantly, in mitochondria cyanide binds to the heme iron of cytochrome-c oxidase blocking reduction of oxygen to water (Jones et al., 1984;

Donato et al., 2007). In cassava cyanogenic glucosides are not the only source of cyanide since in all plants the synthesis of ethylene results in the production of cyanide (Peiser et al., 1984). To protect against cyanide toxicity, plants resort to two pathways, the cyanide assimilation pathway, or the cyanide detoxification pathway.

The first-dedicated step in the cyanide assimilation pathway is catalyzed by β- cyanoalanine synthase (CAS) (Blumenthal et al., 1968) (Fig. 1.9) which condenses cyanide and cysteine to produce β-cyanoalanine. This is followed by hydration of β- cyanoalanine to asparagine by β-cyanoalanine hydrase (Castric et al. 1972), and finally the deammination of asparagine by asparaginase results in the production of aspartate and ammonia (Michalska et al. 2006). The CAS enzyme is located in the mitochondria

(Wurtele et al. 1985, Hartzfeld et al. 2000), a site of high sensitivity to cyanide given the presence of cyanide-sensitive enzymes such as cytochrome-c oxidase. The activity of

CAS has been studied in cyanogenic and non-cyanogenic plants, and has been shown to be present in all plant tissues analyzed to date from cyanogenic plants (Miller and Conn,

1980; Elias et al., 1997). Miller and Conn (1980) correlated the cyanide potential of a plant to the activity of CAS.

Figure 1.9 The cyanide assimilation pathway or β-cyanoalanine synthase pathway. The first step of this reaction involves the condensation of cyanide and cysteine by β- cyanoalanine synthase to β-cyanoalanine; this is followed by hydrolysis of the β- cyanoalanine by β-cyanoalanine hydrase to produce asparagine. The de-ammination of asparagine generates aspartate and ammonia.

Cyanide assimilatory enzymatic activity including CAS, β-cyanoalanine hydrase and asparaginase, has been evaluated in cassava leaves roots and stems (Nambisan and

Sundaresan, 1994; Elias et al., 1997; Zidenga, 2011). CAS activity was detected in all tissues analyzed (Elias et al. 1997). Roots, had a CAS activity of 13.7 µmol H2S/ mg protein/ min, while in leaves it was about three-fold lower (5.1 µmol H2S/ mg protein/ min) (Zidenga, 2011). β-cyanoalanine hydrase activity in roots (25 µg cyanide liberated/min/100 mg protein) was also three times higher than in leaves, (8.8 µg cyanide liberated/min/100 mg protein) (Elias et al., 1997). Asparaginase activity was also greater in leaves compared to the roots. This correlates to prior observations, in which 49% of the carbon assimilated as radiolabeled 14CN in cassava roots, was found as asparagine while only 6% was present in aspartate (Nartey, 1969). These results suggest that in cassava storage roots asparagine is not rapidly metabolized into aspartate and ammonia.

Cyanide can also be detoxifided by rhodanese (Chew , 1973) (Fig. 1.10). The presence of the rhodanese is not widespread in plant species (Miller and Conn, 1980). However, in cassava, rhodanese activity was detected in the leaves with an activity of 4.19 µmol thiocyanate/ mg protein/ min (Zidenga, 2011). This value was comparable to the CAS activity in the leaves. Significantly, rhodanese activity was not detected in roots, indicating that CN is most likely assimilated by CAS in roots (Nambisan and Sundaresan,

1994; Zidenga, 2011).

Figure 1.10 The cyanide detoxification or rhodanese pathway. A single reaction is present in the rhodanese pathway: the production of thiocyanate and sulfite from cyanide and thiosulfate.

1.6 FUNCTION OF CYANOGENIC GLUCOSIDES

There are several proposed functions for cyanogenic glucosides. The most recognized function of cyanogens is as herbivore deterrents (Vetter, 2000). It has been shown that the cassava burrowing bug (C.bergi) prefers feeding on low-cyanogenic varieties of cassava versus high-cyanogenic varieties (Belloti and Arias, 1993). However, not all herbivores are susceptible to cyanogenic glucosides. In some cases cyanide acts as a phagostimulant

(Gleadow and Woodrow, 2002).

Cyanogenic glucosides may also function as a transportable form of reduced nitrogen

(Lieberei et al., 1985; Selmar et al., 1993; Selmar, 1994; Siritunga, 2002; Selmar, 2010).

In Hevea brasilensis upon germination, linamarin is converted into linustatin and transported to the seedling (Selmar, 1988). The linustatin is simultaneously hydrolyzed in to gentobiose and acetone cyanohydrin, the later can subsequently decompose to generate cyanide, which can be assimilated by CAS to produce aspartate or asparagine (Selmar,

1988; Selmar, 1993). The cyanogenic glucoside in the seed provides reduced nitrogen for amino acid synthesis in the seedling. Recently, it has been proposed that cyanogenic glucosides from Hevea brasilensis provide not only nitrogen for amino acid synthesis but carbon for latex production (Kongsawadworakul et al., 2009).

In cassava there are several reports indicating that linamarin is a transportable form of reduced nitrogen. Calatayud et al. (1996) observed that in cassava phloem, the linamarin nitrogen contribution was 2.4 fold higher than that present from other nitrogen-containing compounds. Non-protein nitrogen in cassava accounts for 30-50% of the total nitrogen

(Yeoh and Truon, 1996). And finally it was observed by Siritunga (2002) that when

linamarin synthesis is blocked exclusively in the leaves, not only is the linamarin content reduced in the roots, but roots require a supply of reduced nitrogen to develop.

1.7 DIETARY CYANIDE CONSUMPTION

Cassava foods may contain cyanogens, including linamarin and acetone cyanohydrin

(Bradbury and Holloway, 1988). The quantity of cyanogens will depend on several factors including the cassava variety and how the root is processed to remove cyanogens

(Montagnac et al., 2009). The lethal dose range of cyanide for humans is 0.05 – 3.5 mg/per kg body weight, which for a 60 kg adult amounts to 30-120 mg HCN

(Solomonson, 1981). In a study conducted by Onabolu et al. (2000) the total cyanogens

(mg HCN equivalents/kg dry weight) in cassava processed foods ranged from 0 to 62 mg

HCN equivalents/kg dry weight, with 19% of the analyzed processed foods containing more than the recommended intake by the FAO of 10 mg HCN equivalents/kg of dry weight. After consumption, the cyanide liberated can be detoxified by rhodanese to produce thiocyanate (Westley, 1981). There are limits to the amounts of CN that can be detoxified, however, and the dietary protein (cysteine) status can impact the detoxification pathway (Tylleskar et al.; 1992). Several diseases have been related so far to consumption of cassava dependent on the amount of cyanogen exposure; they are listed below:

1.7.1 Konzo

Konzo was first described by Trolli (1938) as a distinct form of tropical myelopathy characterized by abrupt onset of spastic paraparesis. It was observed that the diet of patients who succumb to Konzo in all countries where it has been described, is uniformly similar consisting of poorly processed cassava (Tylleskar et al. 1991). Konzo is correlated with high blood cyanide levels associated with consumption of improperly processed cassava and is acerbated by diets deficient in sulfur-containing amino acids or low protein consumption (Tylleskar et al. 1992). More recently other etiologies for Konzo have been described. Banea-Mayumba et al. (2000) proposes a specific neurotoxic effect of linamarin rather than damage due directly to cyanide exposure, and Adamolekum et al.

(2010) has postulated that Konzo is brought on by a severe thiamine deficiency that occurs on individuals with marginal thiamine deficiency plus inadequate sulfur amino acid levels brought upon by the need to detoxify the blood of cyanide.

1.7.2 Tropical Ataxic Neuropathy

The symptoms for Tropical Ataxic Neuropathy (TAN) comprise bilateral optic atrophy, bilateral perceptive deafness, myelopathy with or without polyneuropathy, and high frequency of sensory ataxia that results in an uncoordinated gait (Osuntokun, 1981).

Clark (1936) was the first to associate the cyanogens found in cassava as the cause of

TAN. Although TAN and konzo are both classified as myeloneuropathies derived from consumption of dietary cyanide their clinical features and history make them distinguishable (Oluwole et al., 2000). In addition, certain cases of TAN may be improved by reducing the cassava component of the diet (Moore, 1930), whereas Konzo

results from acute cyanogen exposure and is often immediate in its manifestation after eating a highly toxic cassava meal. The frequency of TAN in Southwest Nigeria was said to have decreased to 2.4% in 1969, however, recently, Oluwole et al. described a prevalence of up to 6%, presenting evidence for the endemic nature of this disease in the region.

1.7.3 Goiter

Iodine is absorbed from food and taken up by the thyroid gland, a deficiency of iodine in the diet causes the development of goiter which is enlargement of the thyroid gland

(Bradbury and Holloway, 1988). The accumulation of iodine in the thyroid can be further complicated by the presence of anionic inhibitors; one of these is thiocyanate (Goodman and Gilman, 1985), which is a product of cyanide detoxification. Excessive thiocyanate levels in the blood stream, as a result of consumption of insufficiently processed cassava and possibly reduced protein intake, lead to reduced iodine uptake and the development of goiter (Delange et al., 1983).

1.8 CASSAVA FOOD PROCESSING

Many different processing techniques are used to eliminate the cyanogenic glucosides from cassava before consumption. The most common methods are peeling, boiling, sun- drying, fermentation and grating/crushing.

1.8.1 Peeling

The peel from the storage root contains between 7 to 17 times more cyanogenic glucosides than the root pulp (Santana et al. 2002). Therefore removing the peel removes a large amount of the cyanogenic glucosides reducing the cyanogenic potential of the root

(Bokanga, 2003) (Fig. 1.11A)

1.8.2 Boiling

Cooke and Maduagwu (1978) reported the reduction of cyanogens by 50% after 25 min of boiling, however, if the ratio of root weight to water volume is increased from 1:1 to

1:10 of the 78% cyanogens are removed by boiling (Montagnac et al. 2009, review)

1.8.3 Sun-drying

Natural sun-drying is another common processing technique (Fig. 1.11B). Factors which affect its efficiency for cyanogen removal are the thickness of the cassava chips (pieces) and length of drying time. The greatest amounts of cyanogen removal required 17 days of drying (Mlingi and Bainbridge, 1994). Addition of other techniques such as crushing the roots prior to sun-drying improves the detoxification, reducing the cyanogens by 98% presumably associated with better mixing of linamarin with linamarase (Montagnac et al.,

2009).

1.8.4 Fermentation

Fermentation of cassava by lactic bacteria is a common processing method in Africa. The fermentation of previously grated or soaked cassava can result in the removal of 60-94% of the cyanogens (Westby and Choo, 1994) (Fig. 1.11C).

1.8.5 Grating and Soaking

The grating and soaking of the cassava roots prior to the use of other detoxification methods improves the cyanogen removal efficiency. If cassava roots are soaked before sun-drying up to 98.7% of the cyanogens are removed (Oke, 1994).

Figure 1.11 Different cassava processing techniques. (A) A woman peels cassava (Photo reproduced with permission from the International Institute for Tropical Agriculture) (B)

Women set cassava chips on a platform to sun-dry (Photo reproduced with permission from Ken Wiegand photography) (C) Peeled cassava roots during fermentation (Photo reproduced with permission from the International Institute for Tropical Agriculture).

1.9 PROTEIN CONTENT IN CASSAVA

The protein content per unit dry weight for cassava storage roots is among the lowest of the world’s major crops. The protein levels in cassava storage roots vary according to the variety as well as time of harvest, but generally fresh roots have a protein content of 0.7-

3% (Ceballos et al., 2006; USDA, 2010). The processing method will further impact the protein levels. For example gari, which is a commonly eaten processed cassava food in

Nigeria, contains an average of 0.2% (w/dry weight) protein, while flour contains 0.16 to

0.22% protein (Favier, 1977; Montagnac et al., 2009b).

In contrast to the storage roots, the leaves are high in protein and reach concentrations five to ten times higher than those of the root (Montagnac et al., 2009b). A prepared cassava leaf meal has an average protein content of 28.1% (Wobeto et al., 2006).

Unfortunately, the cassava leaves are not as widely consumed as the storage root; therefore the populations that consume cassava as a staple food may be more prone to protein deficiency (Stephenson et al. 2010). One of the drawbacks of diets depending on a single food source, for example cassava storage roots, is that deficiencies found in the food source will be exacerbated and consequently malnutrition may result (Sun 2008).

Protein-energy malnutrition (PEM) arises when insufficient protein is acquired in the diet over a prolonged period of time, followed or in conjunction with deficiencies in carbohydrates and fat. Two varieties of PEM are distinguished: a) marasmus, which is defined as severe wasting (wasting = severely reduced weight and height); and b) kwashiorkor, which is defined as malnutrition with edema (edema = swelling caused by fluid accumulation in the body) (Muller and Krawinken, 2005). A combined form of

these two diseases known as marasmic kwashiorkor can also manifest itself particularly in small children (Brabin and Coulter, 2003).

In the Sub-Saharan African countries where cassava is consumed, severe deficiencies in both protein and caloric consumption may exist (Fig. 1.12). If the top five producers of cassava in Sub-Saharan Africa and Malawi (included due to recent high increase in cassava production) are compared for their average per capita consumption of protein and calories, only Nigeria and Ghana have a per capita consumption of calories above the daily requirement of 2500 kcal. However, none of these countries has sufficient protein consumption/capita to meet the recommended daily minimum. Additionally, Brazil, a top producer of cassava, and the USA, were included in the analysis. Among the countries included in this study, Brazil is the only cassava producing and consuming country with adequate input of protein and calories; this is possibly due to greater variety in the diet including animal protein.

The availability of cassava with increased protein may help reduce protein malnutrition in

Sub-Saharan Africa countries that depend on cassava as a staple crop.

Figure 1.12 Protein and energy consumed per capita. The protein consumption per capita per day in relation to the quantity of energy consumed is depicted in the graph. The top six cassava producing countries in Sub-Saharan Africa along with Malawi a country with a recent high increase in cassava production were included, in addition Brazil, one of the top producers of cassava in the world as well as the USA were included for reference. All data was obtained at FAOSTAT and collected in 2008. MZ= Mozambique, DRC=

Democratic Republic of Congo.

Several strategies have been proposed to target protein deficiency in cassava as well as in other plants. One approach is to overexpress a native storage protein that contains adequate essential amino acids. This strategy may prove difficult to apply in cassava as there are currently no identified endogenous root storage proteins (Shewry et al., 2003;

Sheffield et al., 2006; Li et al., 2010). The second strategy is expression of a heterologous or artificial protein with a high content of the desired essential amino acids. The first attempt to express an artificial protein was reported by Zhang et al. (2003), and the artificial protein 1 or ASP1 was constitutively expressed in cassava using the 35S gene promoter. The total protein content of the storage roots was unchanged while the protein content in the leaves was increased by 30% (Zhang et al., 2004). It was speculated that the lack of certain amino acids may have limited root protein accumulation (Stupak et al.,

2006). A third strategy is manipulation of the free amino acid pool, this strategy has not been tested previously in cassava. The regulation of the amino acid biosynthetic pathways is complex and therefore it may only be possible to target select amino acids and not a broad amino acid increase. An increase in the total free amino acid pool may be achieved through increased nitrate assimilation, however. Previously Lea et al. (2006) observed that the expression of a de-regulated nitrate reductase with increased activity in

N. plumbaginifolia resulted in the doubling of the free amino acid pool.

1.10 GENETIC TRANSFORMATION OF CASSAVA

Transgenic cassava has been produced for the last 15 years with the first reports of successful transformation occurring in the mid 1990’s (Konan et al., 1994; Sarria et al.,

1995; Schopke et al., 1996; Raemakers et al., 1996; Li et al., 1996). The development of transgenic cassava relies on the development of tissue culture systems that generate embryogenic tissues and the regeneration ability of those tissues to mature and germinate

(Taylor et al,. 2004). Four different transformation systems have been described by

Taylor et al. (2004) for the generation of transgenic cassava (Fig. 1.13).

Figure 1.13 Transformation systems for cassava. The four different transformation systems along with the two different gene transfer methods (Agrobacterium mediated or particle bombardment) have been used for the production of transgenic cassava. There is currently a marked preference for the use of Agrobacterium mediated transformation in comparison to particle bombardment so as to reduce gene copy number integration events. Image reproduced with the permission of the journal of Plant Molecular Biology from Taylor et al. (2004).

Three of the transformation systems depend on the generation of embryogenic structures through somatic embryogenesis. Somatic embryogenesis is the process of developing embryos from tissues that would normally not produce embryos (von Arnold, 2008).

Somatic embryos resemble zygotic embryos structurally (von Arnold, 2008) and have the ability to mature, germinate and develop into plants (Szabados et al., 1987). Somatic embryos have been obtained from a variety of cassava tissues, such as meristem explants, zygotic embryos or floral tissue (Raemakers et al., 1997). Somatic embryogenesis and regeneration of plantlets has also been achieved using immature leaves and leaf lobes

(Szabados et al., 1987; Raemakers et al., 1993; Mathew et al., 1993). Only one transformation system utilizes somatic embryos directly for transformation (Siritunga and

Sayre, 2003), the other two somatic embryo transformation systems depend on the generation of additional tissues for transformation. One approach is to generate friable callus, which is formed by many embryogenic units, also known as friable embryogenic callus or FEC (Taylor et al. 1996). Alternatively, somatic embryos are used to develop cotyledons that are then used for transformation. Immature leaf explants may also be directly transformed (Arias-Garzon and Sayre, 2000).

Two approaches have been used to deliver transgenes to cassava. Particle bombardment involves the introduction of the DNA of interest in to the plant cell by propelling a microparticle coated with the DNA into the tissue (Hansen and Wright 1999, review).

Cassava transformation by particle bombardment was first reported by Schopke et al.

(1996) and Raemakers et al. (1996), and cassava was transformed with the reporter genes

GUS and luciferase, respectively.

A second method for cassava transformation is Agrobacterium-mediated T-DNA transfer.

In this method the ability of Agrobacterium to transfer a portion of its Ti-plasmid, known as the T-DNA, to the host plant’s genome is exploited (Winans et al., 1992; Hansen and

Wright, 1999). The T-DNA segment may be engineered to include selectable markers and the gene of interest. Transformation of cassava via Agrobacterium has been applied to selection and reporter genes such as GUS (Li et al., 1996; Puonti-Kaerlas et al., 1997;

Schreuder et al., 2001), hph and nptII (Schreuder et al., 2001). The final step in any transformation system is the regeneration of plants which may occur via somatic embryogenesis involving maturation and germination of the embryos (Mathews et al.,

1993; Taylor et al., 1996; Ma et al., 1998; Raemakers et al., 2000; Siritunga and Sayre,

2003; Taylor et al., 2004) or via organogenesis (system #2) for shoot regeneration (Li et al., 1996, 1998; Mussio et al., 1998).

There is much debate over the success of the different transformation systems; in particular over the material used for transformation or the methods of regeneration

(Taylor et al. 2004). However, there seems to be a consensus over the use of

Agrobacterium-mediated transformation versus biolistiscs. Agrobacterium-mediated transformation has produced more transgenic lines with valuable traits such as herbicide resistance (Sarria et al., 2000), reduced linamarin synthesis (Siritunga and Sayre, 2003), virus resistance (Chellappan et al., 2004), increased cyanide detoxification during processing by over expression of hydroxynitrile lyase (Siritunga et al,. 2004) and increased starch production (Ihemere et al,. 2006). The focus of cassava transformation has more recently focused on engineering farmer preferred cultivars from Africa and

Brazil (Atehnkeng et al., 2006; Hankoua et al., 2006; Ibrahim et al., 2008).

The production of transgenic cassava may be hindered by the limited number of molecular tools available (Delmer, 2005). There are only a few gene promoters with confirmed expression in cassava storage roots: the 35S promoter ( Li et al., 1996; Puonti-

Kaerlas et al., 1997; Schreuder et al., 2001; Zhang et al., 2003b; Siritunga et al., 2004), the patatin class I promoter from potato (Kim et al., 1994; Siritunga and Sayre, 2003), the p15 and p54 two cassava promoters that were described with activity in the storage roots as well as vascular tissues (Zhang et al., 2003a) and the Pt2L4 promoter from cassava with activity in cassava storage roots and stem vascular tissues (Beltran et al. 2010).

1.11 OBJECTIVES

Cassava is a major source of calories for millions of people many of whom live in Sub-

Saharan Africa. Cassava cultivation has grown extensively in the last 50 years, particularly in Africa. It has increased popularity due to its drought tolerance, ability to grow on poor soils and requirement for low human labor input. However, cassava storage roots do not contain adequate levels of protein. In addition, all cassava tissues with the exception of the seed, store cyanogenic glucosides. The consumption of improperly processed cassava with high quantities of cyanogens has also been linked to diseases such as konzo and tropical ataxic neuropathy. In recent years there have been increasing efforts to improve the nutritional quality of cassava through genetic transformation. Our broad objective was to use plant transformation technologies as a means of improving the protein content of cassava storage root. To achieve this broad objective required the development of new molecular tools to control transgene expression in cassava roots.

Our specific objectives were:

1) Increase the root free amino acid pool and protein content by increasing the

nitrate assimilation through the expression of a more stable nitrate reductase;

2) Reduce linamarin content and increase of the free amino acid pool in roots for

increased protein synthesis by utilizing the nitrile group of linamarin for amino

acid synthesis.

3) Identify new root-specific gene promoters for cassava storage roots with the aim

of increasing the available molecular tools for cassava transformation.

CHAPTER 2

EXPRESSION OF A MUTATED NITRATE REDUCTASE TO

INCREASE THE FREE AMINO ACID AND PROTEIN CONTENT IN

CASSAVA STORAGE ROOTS

2.1 INTRODUCTION

Cassava is a staple crop for over 600 million people in the world (International Fund for

Agricultural Development, 2008). In the Republic of Congo and Nigeria, the two highest producers and consumers of cassava in Sub-Saharan Africa, consumption of cassava amounts to 287 and 114 kg/person/year, respectively (FAOSTAT, 2007). However, the low protein value of cassava storage roots potentially can lead to protein deficiencies in the diet if not supplemented with other protein-rich foods (Stephenson et. al 2010). The protein composition of cassava storage roots varies with age, geographical location and variety (Ceballos et al. 2006). On average cassava storage roots contain 0.7-3% protein, substantially less than corn (9.42%), soybean (12.95%) and potato (3.02%) (USDA

2010).

Potential strategies to increase cassava root protein levels include the expression of non- native storage proteins and engineering the amino acid metabolism to increase the supply of available amino acids for protein synthesis (Stupak et al. 2006). Constitutive expression of the engineered storage protein ASP1 (Kim et al. 1992), whose expression was driven by the CaMV 35S promoter in cassava resulted in no increase in storage root protein but a 30% increase in leaf protein levels (Zhang et al. 2004). Additionally, the storage roots of ASP1 plants had reduced free amino acid levels particularly of the essential amino acids asparagine, alanine and methionine (Stupak et al. 2006). Over- expression of native or non-native storage proteins in plants has been suggested to be dependent on in situ availability of an adequate pool of free amino acids and on properties of the expressed protein such as its folding and accumulation (Beauregard and

Hefford, 2006, Sands et al. 2009). It is possible that low levels of certain amino acids hinder the accumulation of protein in the root of ASP1 expressing cassava, in addition the decrease in essential amino acids observed may indicate that expression of ASP1 has an indirect effect on amino acid metabolism because of its high essential amino acid content

(Zhang et al. 2003, Stupak et al. 2006). It may be possible to bypass these constraints by increasing the amounts of reduced nitrogen for amino acid synthesis. In the current literature the majority of reports of increases in free amino acid pool sizes relate to essential amino acids only. Changes in steady state amino acid levels associated with targeted increases in specific amino acid biosynthesis have only been observed for one or two amino acids at a time (Ufaz and Galili 2008). A generalized increase in the free amino acid pool could be considered a better strategy to increase protein synthesis but the biosynthesis pathway for each amino acid are highly regulated (Joshi and Jander 2010). It

has been observed that uptake and metabolism of nitrate is linked to total free amino acid pool size (Matt et al. 2002, Lea et al. 2006) and protein content (Maldonado et al. 1996,

Martre et al. 2003). The rate limiting step in nitrogen metabolism is the reduction of nitrate (Beevers and Hageman 1969, Campbell 2001).

Nitrate reductase is tightly regulated at both the transcriptional and post-translational levels. Previous attempts to enhance nitrate reduction by over-expression of nitrate reductase, however, did not result in increased total free amino acid pools (Foyer et al.

1994, Quillere et al. 1994).

The post-translation regulation of nitrate reductase is complex. Deletion of a 56 amino acid N-terminal domain and phosphorylation of a key regulatory serine followed by binding of a 14-3-3 protein have been shown to be involved in post-translation regulation of NR levels. Nitrate reductase without the 56 amino acid N-terminal domain (ΔN-NR) was initially linked to light regulation. Removal of the N-terminal 56 amino acids was believed to eliminate a regulatory phosphorylation/14-3-3 binding (Nussaume et al.

1995). However, further research revealed that the ΔN-NR form of the enzyme could be phosphorylated and bind to a 14-3-3 protein (Lillo et al. 1997). The ΔN-NR was shown to be less stable than full-length nitrate reductase and it is suggested the N-terminal region functions as a stability domain (Provan et al. 2000). ΔN-NR has been expressed constitutively in potato (Djenanne et al. 2002, and 2004) and N. plumbaginifolia (Lea et al. 2006), however, there was no increase in the free amino acid levels.

The regulation of nitrate reductase by phosphorylation/14-3-3 binding is a two step process that begins with the phosphorylation of a key serine residue (serine 521 in tobacco, serine 534 in Arabidopsis) located in the Hinge 1 region of the protein (Su et al.,

1996). Phosphorylation of this serine is followed by the binding of a 14-3-3 protein which causes the inactivation of the nitrate reductase by blocking the transfer of electrons from heme to nitrate (Kanamuru et al., 1999; Lambeck et al., 2010). Other aspects of nitrate reductase post-translational regulation involve the phosphatase and kinase enzymes responsible for its phosphorylation and dephosphorylation in response to calcium and photosynthesis metabolites (Harmon et al., 2005; Ali et al., 2007; Lillo,

2008, review). From all the different post-translational regulatory factors controlling nitrate reductase, it has been suggested that phosphorylation of NR and associated 14-3-3 binding has the greatest impact on NR activity (Lillo et al., 2003).

In 2006 Lea et al. expressed a modified nitrate reductase in Nicotiana plumbaginifolia with a mutation in the phosphorylation site involved in 14-3-3 binding. The regulatory serine 521 (S521), was replaced with an aspartate which cannot be phosphorylated. This mutated S521D nitrate reductase proved to have a higher activity than wild-type and ΔN-

NR forms of the enzyme; where the activity state is defined as nitrate reductase activity in the presence of Mg+2 and 14-3-3 proteins as percentage of nitrate reductase activity in the presence of EDTA (total activity (Lillo et al. 2003). Expression of the mutated nitrate reductase in N. plumbaginifolia resulted in the accumulation of 2 fold more free amino acids and a decrease in the total nitrate of the leaves (Lea et al. 2006).

We have expressed in cassava storage roots an Arabidopsis nitrate reductase gene (NIA) with a mutation in the regulatory serine (S534D) site that prevents its phosphorylation and subsequent 14-3-3 protein binding. The results in this chapter demonstrate the ability to modify nitrate assimilation in storage roots to increase the free amino acid content.

2.2 MATERIALS AND METHODS

2.2.1 E.coli and Agrobacterium strains

The Escherichia coli strain DH5ɑ (Invitrogen, Carlsbad, CA, USA) was used for all gene cloning. Agrobacterium tumefaciens strain LBA4404 (Invitrogen) was used for cassava transformation. E.coli cells were grown in Luria-Bertani (LB) (Sezonov et al., 2007) media at 37 ˚C and Agrobacterium cells were grown in YM media (Msikita et al., 2006) at 28 ˚C. Agrobacterium cells growing for transformation of friable embryogenic callus were grown on YM media at 28 ˚C to an OD600 of 0.7.

2.2.2 Chemical reagents and supplies

The chemical reagents used in all experimental procedures were purchased from Fisher

Scientific, Hampton, NH unless otherwise noted. All plastic products such as microfuge tubes and PCR tubes were purchased from USA Scientific, Ocala, Florida.

2.2.3 Cassava cultivar, in vitro and greenhouse growth

The cassava cultivar TMS60444 was used for genetic transformation by the method of friable embryogenic callus (Taylor et al. 1996)(See section Generation of Transgenic

Callus).

Cassava plantlets were grown in vitro on Murashige and Skoog media (1962) supplemented with 20g/L sucrose, MS Vitamins (Sigma Aldrich, USA) and 2.4 g/L of the gelling agent phytagel. This media will be subsequently known as MS2. All media used for in vitro propagation of cassava was sterilized through autoclaving. The growth

chamber conditions were set at a temperature of 28 ˚C, a light intensity of 75 to 100

µmol photons m-2 s-2 and a 16 hr day/8 night cycle.

Six week old in vitro grown cassava plantlets were then transferred to the greenhouse.

They were potted in 4” by 4” in pots with Fafard 51 soil mix (Fafard, Agawam, MS) and supplemented with 15:16:17 (N:P:K) fertilizer or Jack’s 10:30:20 (JR Peters, Allentown,

PA) twice a week. At 4 weeks after initial transfer to soil the plantlets were transferred to

6” by 6” pots.

2.2.4 Nitrate reductase gene

The Arabidopsis nitrate reductase gene (NIA2) was obtained from the Arabidopsis

Biological Resource Center in Columbus, OH (www.abrc.osu.edu). The NIA2 gene was received as an insert in a pUni51 based vector named U09319.

2.2.5 General molecular techniques

All the PCR reactions were run in a BioRad Mycycler thermal cycler (BioRad, Hercules,

CA). All the cloning was done using E.coli DH5ɑ sub-cloning efficiency cells according to the protocol given by the supplier Invitrogen (Catalog # 18265017). Plasmids were isolated using the Qiagen Qiaquick Mini Prep kit following the instructions provided by the manufacturers. The PCR products and recovered plasmids were confirmed in 1%

(w/v) agarose gel in TAE buffer (40mM Tris-acetate, 1mM EDTA) and purified using

Qiaquick Gel Extraction kit according to the instructions set by the manufacturer

(Qiagen, Valencia, CA). The concentration of the final purified product was determined using a Nanodrop 2000 Spectrophotometer (Fisher Scientific) for nucleic acids, with one

unit of absorbance at 260 nm equivalent to 50 ng/µL for double stranded DNA and 40 ng/µL for single stranded RNA. The quality of the product was also determined by 260 nm/230 nm and 260 nm/280 nm ratios between 1.8 and 2.3. All digestions were carried out using 500 ng of purified template with 20 units of specific restriction enzyme in its preferable buffer. All ligations were carried out using a 1:3 ratio, being 50 ng of vector to

150ng of insert, using 400 units of T4 DNA ligase (NEB) in 1X T4 DNA ligase buffer.

2.2.6 Codon optimization

The Arabidopsis nitrate reductase gene was analyzed for codon usage bias using the

Graphical Codon Usage Analyzer (GCUA) in its relative adaptiveness modality

(www.gcua.schoedl.de). The GCUA program was able to identify problematic codons based on the cassava codon usage table available from Kasuza

(http://www.kazusa.or.jp/codon/).

The codon optimization of the gene sequence was done by fusion PCR (Fig. 2.1). This was accomplished by designing primers for the regions of the NIA2 gene that contained problematic codons, the primer sequences used for each fusion PCR reaction include any necessary codon changes or in the case of serine 534 the mutation required to change it to aspartate.

Figure 2.1 Outline of a fusion PCR reaction used for codon optimization. The primers are designed for regions of the gene that require codon optimization; the three products are amplified and then fused together. The fusion occurs due to the overlapping sequence of the primers. The obtained fusion product contains the sequence changes found in the primers.

The sequence of all primers used in the fusion PCR reactions can be found in Table 2.1.

Further detail of primer pairs used in each round of fusion PCR can be found on Table

2.2.

Table 2.1 Primers designed for the synthesis of mutated codon optimized NR. Name and sequence of primers used in the fusion PCR

reactions

N1 ATG GCT GCC TCT GTA GAT AAT AGA CAA TAC G N5 CCT GTC CCA TTG CAC TAC GTT N5R AAC GTA GTG CAA TGG GAC AGG N6 GC GAA TGG ACT GTC GAG GTG N6R CAC CTC GAC AGT CCA TTC GC N7 GA TTC GTC AAA AGA CCC ATG AAA TTC ACC N7R GGT GAA TTT CAT GGG TCT TTT GAC GAA TC N8 GCC GCT ACG CTA GTC TGC N8R GCA GAC TAG CGT AGC GGC N9 GC GCT GGG AAC AGA CGT AAG N9 R CT TAC GTC TGT TCC CAG CGC N10 GGG TCT GAG GAT CTT CCA GGC N10R GCC TGG AAG ATC CTC AGA CCC N11 CAC GGT TTT CCA GTT AGA ATC ATC ATC CC N11R GG GAT GAT GAT TCT AAC TGG AAA ACC GTG N12 GT GGC AGA ATG GTT AAA TGG TTG AAA AGA ATC N12R GAT TCT TTT CAA CCA TTT AAC CAT TCT GCC AC N13 CTA AAC ATA AAC TCC GTG ATT ACT ACT CCA TGT C N13R GAC ATG GAG TAG TAA TCA CGG AGT TTA TGT TTA G N14 GGA TGG ATG GCT AAG GAA CGT CAC N14R GTG ACG TTC CTT AGC CAT CCA TCC N15 GT CAC CTC GAA AAA TCT GCT GAC N15R GTC AGC ATA TTT TTC GAG GTG AC N16 GCT GAC GCT CCT CCT AGT CTA AAG N16R CTT TAG ACT AGG AGG AGC GTC AGC

49 N17 G AAG TCT GTC GAT ACT CCA TTT ATG AAC ACA AC N17R GT TGT GTT CAT AAA TGG AGT ATC GAC AGA CTT C

N18 CA ACT GCT AAG ATG TAC TCT ATG TCC GAG N18R CTC GGA CAT AGA GTA CAT CTT AGC AGT TG N19 GA CAT ATC TAT GAT TGT ACA AGA TTC CTT ATG GAT CAC N19R GTG ATC CAT AAG GAA TCT TGT ACA ATC ATA GAT ATG TC N20 G GAT CAC CCA GGT GGT TCT GAT TC N20R GA ATC AGA ACC ACC TGG GTG ATC C N21 GCT GGT ACT GAT GTG ACT GAG GAG TTT G N21R C AAA CTC CTC AGT ACA ATC AGT ACC AGC N23 G TTC TCT CTG TTG GCT CCA ATT GG N23R CC AAT TGG AGC CAA CAG AGA GAA C N24 GA GAG GCT ACT CCA GTT AGG AAC C N24R G GTT CCT AAC TGG AGT AGC CTC TC N25 G GTT AAT CCA AGA GCT AAA GTC CCA GTT C N25R G AAC TGG GAC TTT AGC TCT TGG ATT AAC C N26 CGT AAA TTC AGA TTT GCT TTA CCA GTT GAG G N26R C CTC AAC TGG TAA AGC AAA TCT GAA TTT ACG N27 GAG GAT ATG GTT CTT GGC TTA CCA GTT G N27R C AAC TGG TAA GCC AAG AAC CAT ATC CTC N28 GGT AGT TTC ACT GTT CAC GGT AAA CC N28R GG TTT ACC GTG AAC AGT GAA ACT ACC N29 GT GGA ACC GGA ATT ACT CCA GTT TAC C N29R G GTA AAC TGG AGT AAT TCC GGT TCC AC N30 C GTC ATT TAT GCT AAC AGA ACC GAG GAA G N30R C TTC CTC GGT TCT GTT AGC ATA AAT GAC G N31 GAG CAA TAC CCA GAC AGA TTA AAG GTT TG N31R CA AAC CTT TAA TCT GTC TGG GTA TTG CTC N35 G ATG CAA TAT AAC ATC AAG GAG GAT TTC TTG ATA TTC TAG N35R CTA GAA TAT CAA GAA ATC CTC CTT GAT GTT ATA TTG CAT C N36 CCA AGA GCT AAA GTC CCA GTT CAA CTC N36R GAG TTG AAC TGG GAC TTT AGC TCT TGG N37 GTT CGT AAA TTC AGA TTT GCT TTA CCA GTT GAG N37R CTC AAC TGG TAA AGC AAA TCT GAA TTT AGC AAC N38 GGT GAA CTT ATC ACC ACT GGT TAT TCC TCT GAC TCT TC N38R GAA GAG TCA GAG GAA TAA CCA GTG GTC ATA AGT TCA CC N39 GTT GAT GTG GTT GGC TAC TTC GAA CTT GTG G N39R CCA CAA GTT CGA AGT AGC CAA CCA CAT CAA C N1EL CAG CGC GAA TTC ATG GCT GCC TCT GTA GAT AAT AG N35RHL GCA GCA AAG CTT GAA TAT CAA GAA ATC CTC CTT GAT GTT ATA TTG

Table 2.2 Nitrate Reductase Fusion PCR primer pairs reactions. In each fusion PCR reaction three to five different PCR products were fused together. Presented in the table above are the primer pairs for each individual PCR reaction that was ran to obtain products for each fusion PCR round (roman numeral).

Fusion PCR reaction Primer pairs number I N1/N11R, N11/N21R and N21/N35R II N1/N9R, N9/N13R, N13/N19R, N19/N28R and N28/N35R III N1/N10R, N10/N14R, N14/N27R and N27/N35R IV N1/N8R, N8/N15R, N15/N25R, N25/N29R and N29/N35R V N1/N12R, N12/N16R, N16/N24R and N24/N35R VI N1/N7R, N7/N17R, N17/N23R, N23/N30R and N30/N34R VII N1/N6R, N6/N18R, N18/N26R and N26/N35R VIII N1/N5R, N5/N20R, N20/N31R and N31/N35R IX N1/N13R, N13/N37R and N37/N35R X N1/N13R, N13/N36R and N36/N35R XI N1/N38R, N38/N39R and N39/N35R

All the fusion PCR reactions were run using a similar protocol. Described below is a detailed description of the first fusion PCR reaction. In the first fusion PCR reaction three products were fused, these products were amplified using vector U09319 as a template and the Platinum Taq High Fidelity DNA Polymerase (Invitrogen) enzyme. The primers used were N1 (located at the 5’ end of the gene starting at base 1) paired with N11R, N11 paired with N21R, and N21 paired with N35R (3’end primer from end of gene) (see

Table 2.1 for primer sequences). The three PCR reactions to amplify the N1/N11R,

N11/N21R and N21/N35R products were using 500 ng of vector U09319, one unit of the enzyme Platinum Taq High Fidelity DNA Polymerase (Invitrogen, Carlsbad, CA), 10 µL of the supplied 10X High Fidelity PCR Buffer, 2 µL of 50 mM MgSO4, 2µL of 10 mM dNTP mixture and 2.5 µL of 10µM solution of each primer. The reaction conditions were the following: one cycle of 5 min at 94˚C, followed by 35 cycles of amplification (30s at

94˚C, 30s at 55 ˚C, and 60s at 72˚C), and a final cycle of 5 min at 72˚C. The three PCR products N1/N11R, N11/N21R and N21/N35R were gel purified, and 500 ng of each purified product was used for the fusion PCR reaction along with one unit of the enzyme

Platinum Taq High Fidelity DNA Polymerase (Invitrogen), 10 µL of the supplied 10X

High Fidelity PCR Buffer, 2 µL of 50 mM MgSO4 and 2 µL of 10 mM dNTP mixture, no primers were used in the fusion PCR reaction. The fusion PCR followed the conditions: one cycle of 5 min at 94˚C followed by 36 cycles of amplification (30 s at 94˚C, 45 s at

50˚C, and 60 s at 72˚C), and a final cycle of 5 min at 72˚C. The fusion PCR product was used directly as a template for a PCR reaction using the N1EL and N35RHL primers

(Table 2.1); these primers are from the 5’ and 3’ end of the gene and include restriction enzyme sites EcoRI (NEB, Ipswich, MA) and HindIII (NEB) respectively. The

amplification was carried out in a BioRad Mycycler Thermal cycler using 100 ng of fusion PCR product, 1 unit of Platinum Taq High Fidelity enzyme (Invitrogen), 10 µL of the 10X High Fidelity PCR Buffer (supplied with the enzyme), 2 µL of 50 mM MgSO4 and 2 µL of 10 mM dNTP mixture and the following cycling conditions: one cycle of 5 min at 94˚C followed by 32 cycles of amplification (30 s at 94˚C, 30 s at 61˚C, and 3 min at 72˚C), and a final cycle of 5 min at 72˚C. The product of this reaction was gel purified

(section 2.2.5). The purified fusion PCR product was then digested with EcoRI (NEB) and HindIII (NEB) simultaneously with NEB Buffer #2 at 37 ˚C for 1 hour, after which time the digestion product was purified (section 2.2.5), and ligated in to a similarly digested pUC19 vector (Invitrogen). The ligation was carried out at room temperature for one and a half hours, and the ligation product of pUC19 and the fusion PCR product were used directly to transform E.coli DH5ɑ sub-cloning efficiency cells (section 2.2.5). After transformation the cells were plated in LB media containing ampicillin (100µg/mL) and incubated for 12 hours at 37˚C. The resulting putative transformed colonies were grown in liquid LB media containing ampicillin (100 µg/mL) for 12 hours at 37˚C. The plasmid

DNA was isolated (section 2.2.5) was used as a template for PCR amplification with the

N1EL and N35RHL primers to confirm the cloning insertion. The plasmid was then sequenced at The Ohio State University Plant-Microbe Genomics facility (Columbus,

OH). The vector containing the confirmed first fusion PCR product was renamed pUC-I-

NIA and used in a subsequent round of fusion PCR as a template. This process was repeated 11 times (Table 2.2 for rounds and primers used), the vector containing the final codon optimized and mutated NIA2 gene was named pUC-XI-NIA and the codon optimized NIA2 was renamed XI-NIA.

2.2.7 Cloning of XI-NIA in to the pCAMBIA2300 binary vector

The pUC-XI-NIA vector was used as a template to amplify XI-NIA using the NIEZH and

N35REZH primers, these primers include restriction enzyme sites for SmaI and SstI restriction enzymes, respectively.

N1EZH: TCG GCT CCC GGG ATG GCT GCC TCT GTA GAT AAT AG

N35REZH: TGG GCT GAG CTC CTA GAA TAT CAA GAA ATC CTC CTT GAT

GTT ATA TTG CAT CTT CTC C

The reaction was set up with 50 ng of pUC-XI-NIA vector, one unit of the enzyme

Platinum Taq High Fidelity DNA Polymerase (Invitrogen, Carlsbad, CA), 10 µL of the supplied 10X High Fidelity PCR Buffer, 2 µL of 50 mM MgSO4, 2 µL of 10 mM dNTP mixture, 2.5 µL of 10 µM of each primer and autoclaved double distilled water up to a volume of 50 µL. The PCR amplification reaction was ran with the following program: one cycle of 5 min at 94˚C followed by 32 cycles of amplification (30 s at 94˚C, 30 s at

61˚C and 3 min at 72˚C for extension) and a final cycle of 5 min at 72˚C. The 3DVL vector is a modified pBI121 (Invitrogen) vector with a patatin promoter: vacuolar linamarase: nos terminator cassette (See Chapter 3 for 3DVL vector map) and the gel purified XI-NIA PCR product were digested sequentially with SmaI (Invitrogen) and SstI

(Invitrogen) (section 2.2.5), first in React 4 ® buffer (Invitrogen) at 30 ˚C for one hour, then the partially digested XI-NIA product was purified and digested with SstI in React 2

® buffer (Invitrogen), at 37 ˚C for one hour.

The purified digested 3DVL vector was ligated with the digested XI-NIA gene at room temperature for one and a half hours and then at 14˚C over night. The ligation product was used directly to transform E.coli DH5ɑ sub-cloning efficiency cells (Invitrogen).

After transformation the cells were plated in LB media containing kanamycin (50 µg/mL) and incubated for 12 hours at 37˚C. The resulting putative transformed colonies were grown in liquid LB media containing kanamycin (50 µg/mL) for 12 hours at 37˚C. The plasmid DNA was isolated and correct integration of the XI-NIA gene was confirmed by

DNA sequencing using the NIEL and N35RHL primers and digestion with SmaI and SstI

(section 2.5.5). The vector was renamed 3D-XINIA.

The patatin promoter: XI-NIA: nos terminator cassette from 3D-XINIA was amplified in a PCR reaction using the patNNN and nosNNN primers.

patNNN: GAGACTGCAGTTGTAGTTAATGCGTATTAGTTTTAGC

nosNNN: TCTCGGTACCGATCTAGTAACATAGATGACACCGCG

The reaction was set up with 500 ng of template vector, one unit of the enzyme Platinum

Taq High Fidelity DNA Polymerase (Invitrogen, Carlsbad, CA), 1X High Fidelity PCR

Buffer, 2 µL of 50 mM MgSO4, 2 µL of 10 mM dNTP mixture and 2.5 µL of 10 µM solution of each primer. The PCR amplification consisted of: one cycle of 5 min at 94˚C followed by 32 cycles of amplification (30 s at 94˚C, 30 s at 58˚C and 4 min at 72˚C for extension) and a final cycle of 5 min at 72˚C. The pCAMBIA2300 binary vector and the purified patatatin promoter: XI-NIA: nos terminator amplified product were digested with

PstI (NEB) and KpnI (NEB) restriction enzymes simultaneously using NEB Buffer #2at

37 ˚C for 1 hour after which time the digestion products were purified and ligated at room temperature for one and a half hours. The product from the ligation was used to transform

E.coli DH5ɑ sub-cloning efficiency cells (Section 2.2.5). After transformation the cells were plated in LB media containing kanamycin (50µg/mL) and incubated for 12 hours at

37˚C. The resulting putative transformed colonies were grown in liquid LB media containing kanamycin (50 µg/mL) for 12 hours at 37˚C. The plasmid DNA was isolated and correct integration of the patatin promoter: XI-NIA gene: nos terminator cassette was confirmed by DNA sequencing of the plasmid DNA using the NIEL, N35RHL, patNNN and nosNNN primers. The vector was renamed pCNIA.

2.2.8 Agrobacterium transformation

Transformation of Electromax LB4404 Agrobacterium cells was done through electroporation. A total of 20 ng of pCNIA vector was added to a 1.5 mL microcentrifuge tube. A volume of 20 µL of thawed cells was added to the microcentrifuge tube and tapped gently to mix. The cell/DNA mix was pipetted in to a 0.1 cm electroporation cuvette (Fisher Scientific, Hampton, NH) and electroporated under the following conditions: 2.0 kV, 200 Ω, 25 μF. Immediately after electroporation a volume of 500 µL of pre-warmed (at 28 ˚C) YM media was added and the cells were pipetted gently up and down. The total volume was removed and added to 500 µL of pre-warmed YM media.

The cell solution was incubated at 28 ˚C for 3 hours and then plated in YM media containing Kanamycin (50µg/mL) and Streptomycin (100µg/mL). The plates were incubated at 28 ˚C for 3 days after which time the formed colonies were picked and grown in liquid YM media containing Kanamycin (50µg/mL) and Streptomycin (100

µg/mL) for 3 days. The insert was confirmed by PCR using the plasmid DNA isolated above as a template and the patNNN and nosNNN primers under the following conditions in a BioRad Mycycler (BioRad): one cycle of 5 min at 94˚C followed by 32 cycles of amplification (30 s at 94˚C, 30 s at 58˚C and 4 min at 72˚C for extension) and a final cycle of 5 min at 72˚C. The reaction was set up with 500 ng plasmid, one unit of the enzyme Platinum Taq High Fidelity DNA Polymerase (Invitrogen, Carlsbad, CA), 10 µL of the supplied 10X High Fidelity PCR Buffer, 2µL of 50 mM MgSO4, 2 µL of 10 mM dNTP mixture and 2.5 µL of 10µM solution of each primer. The product was confirmed by sequencing at the Ohio State Plant Microbe Genomics Facility using the primers patNNN, nosNNN, N1EL and N35RHL.

2.2.9 Generation of Friable Embryogenic Callus

The production of FEC material was carried out based on the procedures previously outlined by Taylor et al. (1996, and personal communication 2008). All the media used for in vitro cassava tissue culture was sterilized through autoclaving. In vitro cassava plantlets were subcultured and grown in sterilized Murashige and Skoog media (1962) supplemented with 20 g/L sucrose, MS Vitamins (Sigma Aldrich, USA) and 2.4 g/L of the gelling agent phytagel, this media will be subsequently known as MS2. After 6 weeks apical leaves from the plantlets were carefully removed and placed on Driver and

Kuniyuki media (1984) supplemented with 20 g/L sucrose, 50 µM picloram and 2.4 g/L of phytagel, this media was named DKW2 (Fig. 2.2A). The apical leaves remained in

DKW2 media for 4 weeks at the following environmental conditions: temperature of 28

˚C, a reduced light intensity of 20 to 35 µmol photons m-2 s-2 and a 16 hr day/8 night

cycle. After 4 weeks in DKW2 media the visible organized embryogenic structures

(OES) (Fig. 2.2B) were removed and placed in fresh DKW2 media. The OES remained in the second DKW2 media for 3 weeks at the same environmental conditions

(temperature of 28 ˚C, a reduced light intensity of 20 to 35 µmol photons m-2 s-2 and a 16 hr day / 8 night cycle) After three weeks the OES were placed in Gresshoff and Doy media (1974) supplemented with 20 g/L sucrose, 50 µM picloram and 2.4 g/L phytagel, this media was named GD2P50. After 3 weeks in GD2P50 the OES may form friable embryogenic callus (FEC) structures, these were removed and placed in fresh GD2P50 for 3 more weeks (Fig. 2.2C). Finally the growing FEC tissues were removed from the second GD2P50 and proliferated for one more round in GD2P50 for three weeks. All three growth cycles in GD2P50 media were done at the following environmental conditions: temperature of 28 ˚C, a reduced light intensity of 20 to 35 µmol photons m-2 s-2 and a 16 hr day/8 hr night cycle. After the third round of proliferation in GD2P50 media the FEC were used in transformation.

Figure 2.2 Cassava transformation tissue culture images. (A) Apical leaves from 6 week old plants were placed in DKW2 media for the development of organized embryogenic structures (B) After four weeks organized embryogenic structures were visible (C)

Friable embryogenic callus developed after the organized embryogenic structures were placed on GD2P50 media. (D) Some of the apical leaves placed in DKW2 media never developed organized embryogenic structures.

2.2.10 Transformation of cassava

Cassava transformation was carried out using friable embryogenic callus (FEC). The subsequent transformation steps were carried out as outlined by Taylor et al. (1996 and personal communication 2008). Briefly, the generated FEC material was co-cultivated with Agrobacterium transformed with pCNIA. After inoculation the FECs were placed in

GD media (Gresshoff and Doy, 1974) containing 50 µM picloram, 500 mg/L carbenicillin and 27.5 µM paramomycin for 28 days. After this period of time the FEC were transferred to Stage 1 regeneration media which consisted of MS2 media supplemented with 5 µM NAA, 250 mg/L carbenicillin and 45 µM paramomycin. After

21 days growing tissues were transferred to Stage 2 regeneration media (MS2 medium plus 0.5 µM NAA and 45 µM paramomycin). The preferred tissues transferred from

Stage 1 to Stage 2 were any FEC clusters showing early torpedo stage or beginning cotyledon stage embryos. At 21 days of growth in Stage 2 the embryos that had developed in to a mature cotyledon stage were placed in the germination stage media

(MS2 medium supplemented with 2µM BAP and 45 µM paramomycin). Embryos that germinated were micropropagated into MS2 media. Each germinated embryo is considered an individual putative transgenic event. The transgenic lines were grown and maintained in MS2 media.

2.2.11 Cassava DNA extraction and transformation validation with PCR

Genomic DNA was extracted from 3-4 leaves of 6 week old in vitro grown cassava plantlets using the DNeasy Plant Mini Kit (Qiagen). Total DNA concentration was quantified using a NanoDrop 2000 Spectrophotometer. The genomic DNA was used as a

template for amplification of a 700 bp region of the XI-NIA gene, primers used where

N37 and N34R (Table 1). The following conditions were used in the PCR: one cycle of 5 min at 94ºC followed by 36 cycles of amplification (30 s at 94 ºC, 60 s at 54 ˚C and 45 s at 72 ºC) and a final cycle of 5 min at 72 ºC. The ChoiceBlue Taq DNA Polymerase

(Denville Scientific, Metuchen, NJ) was used in all reactions, each reaction contained:

400 -1000 ng of genomic DNA, 1 unit of ChoiceBlue Taq DNA Polymerase, 2.5 µL of

10X Choice PCR Buffer, 2.5 µL of each 10 µM primer solution and autoclaved double distilled water up to a volume of 25 µL.

2.2.12 Dot blot analysis of putative transgenic plants

The samples for the dot blots were done in triplicates with 100 ng aliquots of genomic

DNA (20ng/µL) from each putative transgenic line and from four control lines. The control lines have a known copy number of the 35S promoter:nptII:nost terminator cassette as determined by Southern blot, the control lines used in this experiment had

0,1,2 and 3 insertions. All samples of 100 ng of genomic DNA were denatured by adding

150 µl of 0.4 M NaOH followed by boiling for 5 min. A Nylon Hybond N+ membrane was prepared by wetting in 2X SSC buffer (0.3 M NaCl, 0.03 M sodium citrate pH = 7), then placed on top of a moist sheet of Whatman 3MM filter paper, and these two were then placed in between the two layers of the dot blot apparatus. To each sample after denaturation a volume of 150 µL of 2XSSC buffer was added, the total volume of each sample was applied to the wells of the dot-blot apparatus and a gentle vacuum was used to draw the samples in to the membrane. The DNA was immobilized to the membrane using a Stratalinker UV-Crosslinker (Stratagene) set at an energy setting of 120,000

microjoules/cm2 for 15 seconds; the membrane was then turned 90 degrees and crosslinked again at the same conditions and time. A digoxenin (DIG)-labeled probe targeted to the 35S promoter was synthesized using the Roche DIG Probe Synthesis Kit

(Roche, Mannheim, Germany). The primers used were,

35SF: CAC ATC AAT CCA CTT GCT TTG AAG

35SR: CAT GGT GGA GCA CGA CACT

The pCambia2300 vector was used as a template and the reaction was set up using reagents provided in the kit: 5 µL of 10X PCR Buffer with MgCl2, 5 µL of PCR DIG

Probe Synthesis mix, 5 µL each of 10 µM forward and reverse primers, 60 pg of template, 0.75 µL of enzyme mix and autoclaved double distilled water up to 50 µL. The

PCR reaction parameters were as follows: one cycle at 95 ˚C for 5 min, 40 cycles of amplification (95 ˚C for 30 sec, 59 ˚C for 30 sec and 72 ˚C for 15 sec) and a final cycle of

72 ˚C for 5 min. After synthesis and before first use the probe was denatured by boiling for 5 min and then leaving on ice for 5 min. The probe solution was then prepared by using 20 mL of DIG Easy Hyb solution (Roche) and adding 25ng of probe per mL of solution.

The dot blot membrane was pre-hybridized in 8mL of Roche DIG Easy Hyb solution

(Roche) for 1 hour, the solution was discarded and replaced with the 35S probe solution and left over night in the rotary hybridization oven at 40 ˚C. After hybridization the 35S probe solution was removed and stored at -20 ˚C for later use and the membrane was washed twice for 5 min in 2X SSC / 0.1% (w/v) SDS buffer (0.3M NaCl, 0.03M sodium citrate, 0.1% w/v SDS, pH = 7). The membrane was then washed twice in 0.1X SSC /

0.1% SDS (0.015 M NaCl, 0.0015M sodium citrate, 0.1% SDS, pH = 7) for 30 min at 65

˚C in the rotary hybridization oven.

To detect the probe the DIG High Prime DNA Labeling and Detection Starter Kit II was used. The following buffers and solutions were prepared: 1) Maleic acid buffer consisting of 0.01M Maleic acid, 0.15M NaCl, pH=7.5, 2) Detection buffer consisting of 0.1M Tris-

HCl, 0.1M NaCl, pH=9.5, 3) Blocking solution consisting of 10% w/v 10X Blocking solution (provided in kit) in maleic acid buffer (prepared in no.1 above), 4) Antibody solution containing Anti-Digoxigenin-Alkaline Phosphatase antibody diluted 1:10,000 in blocking solution (prepared above in no. 3) for 20mL of total solution 2 µL of antibody was added, 5) Washing buffer consisting of 0.1 M Maleic acid, 0.15 NaCl, 0.3% v/v

Tween-20, pH = 7.5.

The detection procedure was as follows: the dot blot membrane was incubated for 30 min in 100 mL of Blocking solution at room temperature, after this time the membrane was incubated for 30 min in 20 mL of Antibody solution at room temperature in a rotary oven, this was followed by two washes with 75 mL of washing buffer for 15 min each.

The membrane was then incubated for 5 min in Detection buffer, removed and placed face up on a sheet of Saran wrap. A volume of 1 mL of CSPD-ready-to-use chemiluminescent reagent was applied and incubated for 5 min at room temperature. The excess liquid was removed and the membrane was placed inside a plastic development envelope. The membrane was exposed to X-ray film for 5 to 25 min, the X-ray film was then developed in a Kodak 2000A X-ray film developer (Kodak, Rochester, NY)

The developed films were scanned using an Epson Scanner and the Adobe Photoshop program. The dot blot images were analyzed using ImageJ (www.nih.gov) to quantify the

integrated density for each dot-blot spot, the integrated density is the product of the area

(in square millimeters) and the mean grey value (sum of all the gray values of all the pixels in the spot divided by the number of pixels). An average integrated density value from the three replicates was obtained for each line and the controls. Using the integrated density for the controls a standard curve equation was obtained of copy number vs. spot intensity. The equation was used to calculate the copy number in the transgenic lines.

2.2.13 Cassava root RNA extraction

The extraction of RNA from storage root pulp was done using a LiCl extraction method based on the methodology developed by Manickavelu et al. (2007). A ground sample of

50-100 mg of storage root pulp dry tissue was ground inside a microfuge tube using a mini plastic pestle (Fisher Scientific), this was followed by extraction of the tissue with

900 µL of LiCl buffer (100 mM LiCl, 1% SDS (w/v), 100 mM Tris-HCl, 100 mM EDTA and 1% β-mercaptoethanol) and 900 µL of 125 phenol: 24 chloroform:1 isoamyl alcohol for 60 min at room temperature. The samples were centrifuged at 12,400 RCF for 20 min in a microfuge, the aqueous upper phase was transferred to a fresh tube and a volume of

300 µL of 8 M LiCl was added to precipitate the RNA from the tissue samples. The root tissue RNA samples were precipitated overnight at 4 ˚C; the next day the precipitated nucleic acid was pelleted by centrifuging at 16,800 RCF for 10 min at a temperature of 4

˚C. The RNA pellet was washed twice by adding 100 µL of ice cold 80% ethanol and centrifuging at 16,800 RCF at a temperature of 4 ˚C, the ethanol solution was then removed and the RNA pellet was air dried for 10 min at room temperature. The pellets of nucleic acid were resuspended in 40 µL of RNase-free water (Promega, Madison, WI)

and treated with RQ1 RNase-free DNase (Promega) as follows: 40 µL of RNA solution,

4 µL of RQ1 10X Reaction Buffer and 2 µL of RQ1 DNase; the reaction was incubated at 37 ˚C for 30 min, to stop the reaction 4 µL of RQ1 Stop Solution were added to the sample and then incubated at 65 ˚C for 10 min. The RNA was then re-precipitated from the solution with 100 µL of chilled ethanol and 25 µL of sodium acetate. The RNA solution was then incubated at -20 ˚C for 3 hours and later on pelleted by centrifugation at 16800 RCF, the ethanol/sodium acetate solution was removed by pipetting. The remaining total RNA was washed once with 100 µL of 80% ethanol and then centrifuged again at 16800 RCF, the ethanol was removed and the RNA pellet was resuspended in 50

µL of RNase-free water. The concentration and quality of the RNA was obtained using

NanoDrop (section 2.2.5).

2.2.14 Semiquantitative RT-PCR of the XI-NIA gene

The cDNA for wild-type and transgenic storage roots was synthesized using 1µg of extracted RNA (2.2.13), 4µL of Quantas qScript cDNA SuperMix (Quanta Biosciences,

Gaithersburg, MD) and RNase-free water (Promega) up to a volume of 20 µL. The reaction was run using the following program: 5 min at 25 ˚C, 30 min at 42 ˚C, 5 min at

85˚C and hold at 4 C˚. The cDNA was stored at -20 ˚C.

The RT-PCR reaction was carried out using the first-strand cDNA for wild-type and transgenic storage roots as a template. Gene specific primers for XI-NIA (N37 and N34R,

Table 2.1) were used along with primers for the cassava tubulin gene (5’TUB and

3’TUB), which served as control for the cDNA reaction and the overall quality of the

RNA extraction.

5’TUB: GAT CCT ACT GGG AAG TAC ATT GG

3’TUB: CTG CAT TCT CCA CCA ACT GA

The following conditions were used in the RT-PCR reaction: one cycle of 5 min at 94º C followed by 32 cycles of amplification (30 s at 94º C, 30 s at annealing temperature and

45 s at 68ºC) and a final cycle of 5 min at 68º C. The annealing temperature for the XI-

NIA primers was 54º C and for the tubulin primer pair it was 55º C. The Platinum Pfx

DNA Polymerase enzyme (Invitrogen) was used in all reactions as follows: 100 ng of cDNA, 1 unit of Platinum Pfx DNA Polymerase, 5 µL of 10X Pfx Amplification Buffer,

0.5 µL of 50 mM MgSO4, 1.5 µL of 10 mM dNTP solution, 1.5 µL of each 10 µM primer solution and autoclaved double distilled water up to 25 µL. The amplification products were separated in 1% (w/v) agarose gels.

2.2.15 Real-time RT-PCR analysis of XI-NIA expression

The real-time RT-PCR reaction was carried out using an Applied Biosystems Step One

Plus Real Time PCR system (AB, Carlsbad, CA). The first-strand cDNA’s where used as a template, the cDNA reaction product was diluted to a concentration of 1 ng/µL and in four replicates 5 ng were loaded of each sample in a 96-well plate (Axygen PCR

Microplate, Axygen, Union City, CA). In addition the following was added to each reaction well: 10 µL of SYBR Green ROX (Quanta BioSciences, Gaithersburg, MD),

1µL each of the promoter forward and reverse primer for XI-NIA (RTNIA5 and

RTNIA3) or the housekeeping control tubulin (RTTUB5 and RTTUB3) and 3 µL of

0.1% (v/v) dyethylpyrocarbonate (DEPC) treated water. The forward and reverse primers used where,

RTNIA5: CTG CGG GAT CTT TAG CCG AAA AGG

RTNIA3: CTA ACT GGA AAA CCG TGG TCT GGT G

RTTUB5: GTG GAG GAA CTG GTT CTG GA

RTTUB3: TGC ACT CAT CTG CAT TCT CC

The real-time RT-PCR reaction was carried out using an Applied Biosystems Step One

Plus Real Time PCR system (AB, Carlsbad, CA) programmed to calculate the threshold cycle (Ct) values for each target gene, the Ct value is defined as the number of cycles required for the fluorescent signal to cross the background fluorescent signal. Analysis of gene expression was done using the Comparative Ct Method or 2^-ΔΔCt method (Bulletin

#2 Applied Biosystems Sequence Detection System). Using the comparative Ct method the data can be presented as the fold difference in gene expression normalized to an endogenous reference gene and relative to a reference sample.

The calculations were as follows: the average Ct value of XI-NIA for each CNIA sample was normalized to the average Ct value of the endogenous tubulin gene in the same

CNIA sample to obtain the ΔCt values,

The ΔCt values of the different samples are then compared to a reference sample, in many situations the reference sample is an untreated control or the native gene, however in our case the wild-type samples do not express XI-NIA. Therefore our reference sample

was selected as the lowest expressor according to ΔCt values, in this case it was CNIA-

12. To calculate the ΔΔCt the following formula was used,

Finally the relative expression was calculated, where the relative expression is the amount of target normalized to the endogenous housekeeping gene and relative to a reference sample, the relative expression of CNIA-12 using the formula presented below is 1, the remaining CNIA samples are expressed as fold higher than CNIA-12.

The standard errors for the relative expression were calculated according to the SDS RQ

Manager 1.1 Software Manual (Applied Biosystems).

2.2.16 Linamarin determination in storage root and leaf

Linamarin levels of leaves and storage root pulp were determined by GC-MS using the internal standard method in a similar manner as previously described by Siritunga et al.

(2003). Analysis was done using an Agilent 5975C inert XL MSD with Triple Axis detector (Agilent Technologies, Santa Clara, CA). The column used was an Agilent DB-5 of 30m length, internal diameter 0.32 mm and film thickness of 0.25 µm (Agilent

Technologies). The GC-MS was operated at a pressure control mode at a flow of 1 mL/min. The GC-MS oven program was as follows, 50˚C for 1 min, ramp at 30˚C/min to

185˚C, ramp at 6˚/min to 230˚C, ramp at 12˚C/min to 280˚C and 3 min at 290˚C for column cleaning.

2.2.16.1 Standard curve of linamarin and internal standard

A standard curve was prepared using variable amounts of linamarin and a set quantity of

5 µg of the internal standard phenyl-β-glucopyranoside (PGP). The stocks of linamarin and PGP were prepared in acetonitrile at a concentration of 2µg/µL. The six samples prepared for the standard curve all contained the 5µg of PGP and either 1µg, 2µg, 3µg,

4µg or 6µg of linamarin. All standard samples had a final volume of 50 µL in acetonitrile. In addition two samples were prepared containing only 5 µg linamarin or 5

µg PGP these were used to determine the mass ion to be monitored and the retention time of each of these two compounds.

2.2.16.2 Tissue sample extraction for linamarin quantification

A total of 5 mg and 25 mg of dry weight were taken from leaf and storage root pulp respectively of 5 month old greenhouse grown cassava. A volume of 600 µL of acetonitrile and 2.5µL of the internal standard phenyl-β-glucopyranoside (PGP) stock solution (2µg/µL) was added to each sample and this was followed by a 40 min extraction in a vortex set at 1,200 rpm. The samples were then centrifuged at 16,800 RCF for 10 min. The supernatant was removed and stored in a new 2 mL microcentrifuge tube.

The remaining pellet was re-extracted with 600 µL of acetonitrile for 40 min in a vortex set at 1,200 rpm. Samples were centrifuged once more at 16,800 RCF and the supernatant was combined with the previous one. The total extract obtained from root pulp samples was completely dried in a Centrivap speedvac. An aliquot of 100 µL was taken from the total extracts for leaf and dried completely in a Centrivap speedvac. After drying the pellets were suspended in 150 µL of water, followed by addition of 150 µL of chloroform

to remove lipids and other contaminants. The samples were vortexed at high speed for 10 seconds and centrifuged at 16,800 RCF for 10 min. The aqueous phase was transferred to a new 2 mL microcentrifuge tube and dried completely in a Centrivap speedvac, after drying the remaining pellet was resuspended in 50 µL of acetonitrile.

2.2.16.3 Derivatization of linamarin and internal standard

A volume of 50 µL of N-Methyl-N-(trimethylsilyl) trifluoroacetamide with 1% trimethylchlorosilane (MSTFA + 1% TMCS) (Sigma-Aldrich, St. Louis, MO) and 10 µL of extra dry pyridine (Sigma-Aldrich) were added to all samples (from tissue or standards). They were incubated at 65˚C for 30 min on a dry heating block, and were run on the Agilent 5975C inert XL MSD with Triple Axis detector GC-MS (Sigma-Aldrich,

St. Louis, MO) immediately after derivatization.

2.2.16.4 Selection of ion for single ion monitoring of linamarin content

To determine the retention time and mass ion to be monitored for linamarin and phenyl-

β-glucopyranoside (PGP), these two compounds were ran individually in the GC-MS.

The retention times were determined to be 12.7 min for linamarin (Fig. 2.3A) and

15.15min for PGP (Fig. 2.3B). Using the analytical software of the Agilent 5975C, we obtained the mass spectrometer scan for the linamarin peak (Fig. 2.4A). The most abundant ion had a mass-to-charge ratio (m/z) of 204.1 followed by 73.1 m/z. For PGP the most abundant ion was found at 73.1 m/z (Fig. 2.4B), while the second most abundant was located at 361.2 m/z. The 73.1 m/z peak is common to compounds containing a

glucose moiety (www.webbook.nist.gov) and due to its commonality in both linamarin and PGP, the 361.2 m/z ion was selected for monitoring in PGP.

Figure 2.3 GC-MS analysis of Linamarin (A) and phenyl-β-glucopyranoside (B).The retention time of linamarin was 12.702min (A)

and that of phenyl-β-glucopyranoside was 15.15 min (B).

Figure 2.4 Mass spectrometer scan of Linamarin (A) and phenyl-β-glucopyranoside (B). The most abundant ion for linamarin

was at 204.1 m/z , which was selected for monitoring (A). However the most abundant 73.1 m/z ion derived from the glucose

moiety of phenyl-β-glucopyranoside it is also present in linamarin, therefore the second most abundant ion at 361.2 m/z was

selected for monitoring phenyl-β-glucopyranoside (B).

2.2.16.5 Linamarin Standard curve

A set of five standards containing 1 to 6 µg of linamarin and a constant 5µg of PGP were run in the GC-MS using the aforementioned program and column. The areas of the linamarin and PGP peaks were determined using the Agilent Analytical Software (Table

2.3).

Table 2.3 Linamarin standard curve. Five standards containing 1 to 6 µg of linamarin and

5 µg of phenyl-β-glucopyranoside (PGP) as our internal standard were analyzed through

GC-MS. The areas of the peaks of each compound were determined based on the ion

204.1 m/z for linamarin and 361.1 m/z for PGP.

Linamarin PGP Linamarin PGP

Standard µg µg Area Area Wlin/WPGP Alin/APGP 1 1 5 169328 2377377 0.2 0.071225 2 2 5 1051527 2128878 0.4 0.493935 3 3 5 2527934 2245280 0.6 1.125888 4 4 5 4102372 2172206 0.8 1.888574 5 6 5 8184673 2419392 1.2 3.382946

A standard curve was plotted with the x-axis values corresponding to the ratio of linamarin/PGP weight in µg and the y-axis corresponding to the ratio of their peak areas.

Analysis of the data gave a linear trend line (Fig. 2.5) with a 0.9906 correlation coefficient using the following equation:

Were WLin = Linamarin weight in µg

WPGP = PGP weight in µg

ALin = Linamarin peak area

APGP = PGP peak area

The equation can be used to calculate the total µg of linamarin (WLin) in each of our tissue samples, when a known quantity of PGP (as internal standard) is added to the sample. The equation used in our further calculations was,

Figure 2.5 Standard Curve of the weight and area ratios of linamarin and phenyl-β- glucopyranoside (PGP). Linear correlation analysis of the weight and area ratios obtained for linamarin and our internal standard PGP gave a correlation coefficient of 0.9906, indicating a positive correlation between the weight ratio of linamarin and PGP and the area ratio of linamarin and PGP.

2.2.17 Free amino acid extraction and analysis from storage root and leaf

Fresh tissue samples were freeze dried in a FTS FlexiDry Lyophilizer (SP Scientific,

Warminster, PA). A total of 20 mg of root pulp and 15 mg of leaf dry weight for each line were extracted twice with 600 µL of Amino Acid Extraction Solution (3 water:5 chloroform:12 methanol(v/v). The two extraction volumes were combined and 400 µL of water and 350 µL of chloroform were added to the extract. The samples were vortexed at high speed and centrifuged at 16,800 RCF for 5min, the upper phase was removed and dried completely in a Centrivap speedvac. The pellets were sent for free amino acid analysis at the Proteomics and Mass Spectrometry Facility of the Donald Danforth Plant

Science Center.

2.2.18 Total protein analysis of storage root pulp and leaf

Fresh tissue samples were freeze dried in a FTS FlexiDry Lyophilizer (SP Scientific).

Analysis of total protein was done on a dry weight basis and all steps of the extraction were performed at 4˚C. Samples of 15 mg dry weight from leaf and storage root pulp were placed in a microcentrifuge tube and ground to a powder using a mini pestle (Fisher

Scientific). The powder was then placed in a 1.5 mL microcentrifuge tube with 500 µL of cold protein extraction buffer (200mM NaCl, 1mM EDTA, 0.2% Triton X-100, 100mM

Tris-Cl pH 7.8, 4% 2-mercaptoethanol and Roche cOmplete Protease inhibitor cocktail).

The sample was vortexed at 1200 rpm for 10min and then centrifuged at 8,600 RCF for 5 min, the supernatant was saved in a new microcentrifuge and the pellet was re-extracted with 500 µL of protein extraction buffer and processed in a similar manner. The two supernatants were combined and kept at 4˚C.

The protein concentration was determined using CB-X kit (G Biosciences, Maryland

Heights, MO) according to manufacturer instructions. The protocol provided by the manufacturer was followed briefly, a volume of 10-100 µL of crude protein extract was transferred to a 1.5 mL centrifuge tube, one mL of chilled CB-X ™ (at -20 ˚C) was added and the sample was vortexed for 30 seconds and then centrifuged at 16,800 RCF for 10 min at 4 ˚C. The supernatant was discarded and the pellet was resuspended in 50 µL of

Solubilization Buffer I and 50 µL of solubilization buffer II (provided in the kit), this was followed by addition of 1mL of CB-X Assay Dye, vortexing for 5 sec and incubating at room temperature for 5 min. The absorbance of the samples was then measured at 595 nm. The concentration of total protein was calculated according to the absorbance/protein concentration standard table provided by G-Biosciences in each CB-X kit, the standard table contains a BSA based protein concentration vs. absorbance curve.

2.2.19 Western blot of nitrate reductase in CNIA and wild-type plants

Protein was extracted from 2-3 g fresh weight of storage root pulp by grinding the tissue in liquid nitrogen in a mortar and pestle. A volume of 5 mL of MOPS buffer (100mM

MOPS, 0.08% (w/v) cysteine, 1% (w/v) polyvinylpyrrolidone and 1 tablet of Roche cOmplete protease inhibitor cocktail for every 50mL of extraction buffer) was added to the ground tissues and mixed with the mortar and pestle, the tissue and buffer were then placed in a 15 mL falcon tube and vortexed at 1000 RCF for 15min. The samples were then centrifuged at 3400 RCF and the supernatant was collected in a fresh 15mL falcon tube. The samples were kept at 4˚C during all stages of the protein extraction. The proteins were precipitated by the addition of acetone using a similar protocol to the one

detailed in the PIERCE Technical Resource: Acetone precipitation of proteins. The procedure was done as follows: A volume of 7-10 mL of acetone (Fisher Scientific,

Pittsburgh, PA) was added to the protein extract and it was left overnight at -20˚C to precipitate the extracted proteins. The next day the samples were centrifuged for 10 min at 3800 RCF. The supernatant was discarded and the pellet was resuspended in 200 µL of

100 mM sodium phosphate buffer pH 7.5 with mini cOmplete protease inhibitor cocktail

(Roche, 1 tablet for every 10mL of buffer). Protein concentration was determined using the CB-X kit following all guidelines provided by the supplier.

An amount of 20 to 50 µg of protein was denatured by adding 15 – 25 µL of loading dye

(0.06M Tris-HCl, pH 6.8, 10% (v/v) glycerol, 2% (w/v) SDS, 5% (v/v) 2- mercaptoethanol, 0.0025% (w/v) bromophenol blue) and incubating at 95˚C for 10min and then placing on ice for 2min. After denaturing the samples were ready to load on an

SDS protein gel. A Bio-Rad pre-cast ready gel of 7.5% polyacrylamide ( Bio-Rad

Catalog # 116-1154) was used to run the samples in an Mini Protean electrophoresis apparatus (Bio-Rad) using 1X running buffer (30mM Tris base, 200mM glycine and

0.1% w/v of SDS per liter). The gel was run at 25 mAmps until completion. Three

Whatman filter paper pieces previously wetted with transfer buffer (20 mM Tris-Cl, 192 mM glycine, 20% v/v methanol, 0.01% SDS) were placed on a dry blot transfer apparatus, followed by the protein gel, Immobilon nylon membrane and three more

Whatman filter paper layers, all previously wetted with transfer buffer. The dry blot transfer apparatus was run at a constant current of 1.9-2.5mA/cm2 of gel area for 4 hours.

After the transfer the nylon membrane was placed in blocking buffer (20 mM Tris-HCl pH 7.5, 500 mM NaCl, 1% (w/v) bovine serum albumin) for 1 hour. Then the membrane

was incubated overnight at 4˚C with Arabidopsis nitrate reductase rabbit antibody (10

µg/µL) (Agrisera, Vannas, Sweden) at a 1:10,000 dilution in Antibody Buffer (20mM

Tris-Cl pH 7.5, 500 mM NaCl, 0.05% Tween-20, 1% (w/v) bovine serum albumin). The following day the blot was washed three times for 15 min each in TTBS buffer (20mM

Tris-Cl pH 7.5, 500 mM NaCl, 0.05% v/v Tween-20) and then incubated in with the secondary antibody (10 µg/µL of anti-rabbit horseradish peroxidase) at 1:10,000 in

Antibody Buffer for 2 hours at room temperature. The blot was then washed three times with TTBS buffer for 15 min and one time with TBS buffer (20mM Tris-HCl, 500mM

NaCl, pH = 7.5) for 5 min. The western blot was then developed using chemiluminescence with luminol in a protocol similar to that developed by Yakunin and

Hallenbeck (1998). The detection solution was prepared in the dark with the following reagents: 7 mL of Solution A (100mM glycine-NaOH buffer pH=9.6), 7 mL of Solution

B (30% v/v H2O2 in water), 1 mL of luminol reagent (45mM luminol in dimethyl sulfoxide) and 1 mL of 4-iodophenol reagent (750 mM 4-iodophenol in dimethyl sulfoxide). The western blot membrane was placed in the detection solution for 1 min then washed in TBS buffer (20 mM Tris-HCl, 500 mM NaCl, pH = 7.5) for 5 seconds and placed in a plastic developing envelope. The membrane was exposed to X-ray films for 30 seconds to 5 min and developed in a Kodak 2000A X-Ray developer (Kodak).

+ 2.2.20 Growth experiment on free NH4 media

Two different types of growth media in addition to the control MS2 media were prepared for in vitro culture of cassava. The MS2 media had a concentration of total N of 60 mM

- + with 39.4mM in the form of NO3 and 20.61 mM in the form of NH4 . The media named

NH4 free #1 contained no ammonia ions and had a total N content set at 39.4 mM, all in the form of nitrate. A second media named NH4 free #2, similarly had no ammonia content and had a total concentration of nitrogen in 18.8mM of nitrate. All three media contained all other nutrients present in MS2 media (Murashige and Skoog media (1962) supplemented with 20 g/L sucrose, MS Vitamins (Sigma Aldrich, USA) and 2.4 g/L of the gelling agent phytagel).

Three plates of each media with three plants each of wild type and transgenic lines

CNIA-11, CNIA-12 and CNIA-15, were grown for 6 weeks under in vitro conditions at a temperature of 28 ˚C, a light intensity of 75 to 100 µmol photons m-2 s-2 and a 16hr day/8 hr night cycle. After this time the plantlets were removed intact from the media and dried in a FTS Flexi Dry Lyophylizer (SP Scientific) for 2 days. The plantlets were weighed individually and the average dry weight per line per media was obtained.

2.2.21 Analysis of calorie content in cassava storage roots

Samples from 5 month old greenhouse grown cassava storage roots were dried in an FTS

Flexi Dry Lyophilizer (SP Scientific). Analysis of the calorie content was done for samples of 0.25 g dry weight using a Parr 6300 Automatic Isoperibol Calorimeter with an oxygen flow at a pressure of 450 psi.

2.2.22 Statistical analysis

All analyses were done in triplicate with the exception of the growth experiment in which nine samples of each line in each media were measured. The samples for all 5 month old greenhouse grown plants came from three different plants of the same line grown under the same conditions but placed randomly in the greenhouse space by staff. To determine statistical significance a two-tailed t-test analysis was done to obtain a p-value. If the p- value obtained was less than 0.05 the values compared were considered statistically significantly different.

2.3 RESULTS

2.3.1 Codon optimization and mutation of Arabidopsis NIA2

The Arabidopsis NIA2 gene was codon optimized and serine 534 was mutated to an aspartate by fusion PCR. Eleven fusion PCR reactions were needed to optimize a total of

50 codons and modify two SstI enzyme sites (Fig. 2.6). Successful modification of the sequence was confirmed by DNA sequencing and sequence alignment between the

Arabidopsis NIA2 gene and the cassava optimized and mutated XI-NIA gene (Fig. 2.7) along with the translated protein sequences (Fig. 2.8).

I-NIA

II-NIA

V-NIA

Figure 2.6 Fusion PCR reaction. (A) First Fusion PCR reaction. Three products were amplified using the plasmid U093 as template and using three different primer pairs: N1/N11R, N11/N21R and N21/N35R. The three PCR products were fused together and the product obtained was amplified (far right shown) and renamed I-NIA. (B) Second Fusion PCR reaction. The product obtained after the first fusion PCR, also known as I- NIA was used as a template to amplify five different regions of the gene using the following primer pairs: N1/N9R, N19,13R, N13,N19R, N19/N28R and N28/N35R. The five PCR products were fused together and the product obtained was amplified (far right) and renamed II-NIA. (C) Fifth fusion PCR reaction. The IV-NIA (product of the fourth fusion PCR) was used as a template, the four PCR products were amplified using the following primer pairs: N1/N12R, N12/N16R, N16/N24R and N24/N35R. The resulting amplified products were fused together and the resulting gene was renamed V-NIA. (far right).

* 20 * 40 * 60 * 80 * 100 * 120 * 140 * XI-NIA : ATGGCTGCCTCTGTAGATAATAGACAATACGCTCGTCTCGAGCCAGGTTTGAACGGCGTGGTTCGTTCTTACAAACCTCCCGTTCCAGGCCGGTCCGATTCCCCTAAGGCGCACCAGAACCAAACCACCAACCAAACCGTGTTCTTGAAA : 150 NIA2 : ATGGCGGCCTCTGTAGATAATCGCCAATACGCTCGTCTCGAGCCAGGTTTGAACGGCGTGGTTCGTTCTTACAAACCTCCCGTTCCAGGCCGGTCCGATTCCCCTAAGGCGCACCAGAACCAAACCACCAACCAAACCGTGTTCTTGAAA : 150

160 * 180 * 200 * 220 * 240 * 260 * 280 * 300 XI-NIA : CCAGCCAAGGTTCATGACGATGACGAAGACGTGTCGAGCGAAGACGAGAACGAGACACACAACAGCAACGCCGTGTACTACAAGGAGATGATAAGAAAATCCAACGCCGAGCTTGAACCGTCTGTTTTGGACCCGAGGGACGAATACACG : 300 NIA2 : CCAGCCAAGGTTCATGACGATGACGAAGACGTGTCGAGCGAAGACGAGAACGAGACACACAACAGCAACGCCGTGTACTACAAGGAGATGATAAGAAAATCCAACGCCGAGCTTGAACCGTCCGTTTTGGACCCGAGGGACGAATACACG : 300

* 320 * 340 * 360 * 380 * 400 * 420 * 440 * XI-NIA : GCTGATAGCTGGATCGAGCGTAACCCTTCCATGGTACGTCTCACAGGGAAACATCCCTTCAACTCCGAGGCGCCTCTTAACCGTTTAATGCACCACGGGTTTATCACCCCTGTCCCATTGCACTACGTTCGTAACCACGGCCACGTCCCT : 450 NIA2 : GCTGATAGCTGGATCGAGCGTAACCCTTCCATGGTACGTCTCACAGGGAAACATCCCTTCAACTCCGAGGCGCCTCTTAACCGTTTAATGCACCACGGGTTTATCACCCCTGTCCCGTTGCACTACGTTCGTAACCACGGCCACGTCCCT : 450

460 * 480 * 500 * 520 * 540 * 560 * 580 * 600 XI-NIA : AAAGCCCAATGGGCCGAATGGACTGTCGAGGTGACCGGATTCGTCAAAAGACCCATGAAATTCACCATGGACCAGCTCGTCTCCGAGTTTGCTTACCGCGAGTTCGCTGCTACCCTAGTCTGCGCTGGGAACAGACGTAAGGAACAGAAC : 600 NIA2 : AAAGCCCAATGGGCCGAATGGACGGTCGAGGTGACCGGATTCGTCAAACGGCCCATGAAATTCACCATGGACCAGCTCGTCTCCGAGTTTGCTTACCGCGAGTTCGCCGCGACGCTAGTCTGCGCGGGGAACCGCCGTAAGGAACAGAAC : 600

* 620 * 640 * 660 * 680 * 700 * 720 * 740 * XI-NIA : ATGGTGAAGAAGTCAAAGGGATTCAACTGGGGATCCGCCGGAGTTTCCACCTCCGTGTGGCGTGGTGTCCCTCTTTGCGACGTACTGCGTCGCTGCGGGATCTTTAGCCGAAAAGGCGGCGCTCTCAACGTCTGCTTCGAAGGGTCTGAG : 750 NIA2 : ATGGTGAAGAAGTCAAAGGGATTCAACTGGGGATCCGCCGGAGTTTCCACCTCCGTGTGGCGTGGTGTCCCTCTCTGCGACGTACTGCGTCGCTGCGGGATCTTTAGCCGAAAAGGCGGCGCTCTCAACGTCTGCTTCGAAGGGTCGGAG : 750

760 * 780 * 800 * 820 * 840 * 860 * 880 * 900 XI-NIA : GATCTTCCGGGTGGTGCCGGAACTGCTGGTTCCAAATACGGAACGAGCATCAAGAAGGAATATGCCATGGATCCATCAAGAGACATCATTTTGGCTTATATGCAAAACGGAGAGTATCTAACACCAGACCACGGTTTTCCAGTTAGAATC : 900

85 NIA2 : GATCTTCCGGGCGGTGCCGGAACTGCTGGTTCCAAATACGGAACGAGCATCAAGAAGGAATATGCCATGGATCCATCAAGAGACATCATTTTGGCTTATATGCAAAACGGAGAGTATCTAACACCAGACCACGGTTTTCCGGTTCGGATC : 900

* 920 * 940 * 960 * 980 * 1000 * 1020 * 1040 * XI-NIA : ATCATCCCCGGTTTCATTGGTGGCAGAATGGTTAAATGGTTGAAAAGAATCATTGTCACAACTAAAGAATCCGACAATTTCTACCATTTCAAGGACAACAGAGTTTTACCTTCTTTGGTAGACGCCGAACTCGCCGACGAAGAAGGTTGG : 1050 NIA2 : ATCATCCCCGGTTTCATTGGTGGCCGGATGGTTAAATGGTTGAAACGAATCATTGTCACAACTAAAGAATCCGACAATTTCTACCATTTCAAGGACAACAGAGTTTTACCTTCTTTGGTAGACGCCGAACTCGCCGACGAAGAAGGTTGG : 1050

1060 * 1080 * 1100 * 1120 * 1140 * 1160 * 1180 * 1200 XI-NIA : TGGTATAAGCCAGAGTACATAATCAACGAGCTAAACATAAACTCCGTGATTACTACTCCATGTCACGAGGAGATTCTTCCCATCAACGCTTTCACAACCCAAAGACCTTATACTTTAAAGGGTTACGCATATTCCGGAGGTGGAAAAAAA : 1200 NIA2 : TGGTATAAGCCAGAGTACATAATCAACGAGCTAAACATAAACTCCGTGATTACGACGCCATGTCACGAGGAGATTCTTCCCATCAACGCTTTCACAACCCAAAGACCTTATACTTTAAAGGGTTACGCATATTCCGGAGGTGGAAAAAAA : 1200

* 1220 * 1240 * 1260 * 1280 * 1300 * 1320 * 1340 * XI-NIA : GTGACCCGTGTGGAGGTCACGGTAGATGGTGGAGAGACATGGAACGTATGTGCACTTGACCATCAAGAGAAGCCAAACAAGTATGGGAAGTTCTGGTGTTGGTGTTTTTGGTCACTTGAGGTTGAGGTTTTGGACTTGCTTAGTGCCAAA : 1350 NIA2 : GTGACCCGTGTGGAGGTCACGGTAGATGGTGGAGAGACATGGAACGTATGTGCACTTGACCATCAAGAGAAGCCAAACAAGTATGGGAAGTTCTGGTGTTGGTGTTTTTGGTCACTTGAGGTTGAGGTTTTGGACTTGCTTAGTGCCAAA : 1350

1360 * 1380 * 1400 * 1420 * 1440 * 1460 * 1480 * 1500 XI-NIA : GAGATTGCTGTTCGTGCATGGGACGAGACTCTCAACACGCAGCCCGAGAAAATGATATGGAATCTCATGGGGATGATGAATAACTGCTGGTTTAGAGTGAAGACTAACGTGTGCAAGCCACACAAGGGAGAGATTGGGATTGTGTTCGAG : 1500 NIA2 : GAGATTGCTGTTCGTGCATGGGACGAGACTCTCAACACGCAGCCCGAGAAAATGATATGGAATCTCATGGGGATGATGAATAACTGCTGGTTTAGAGTGAAGACTAACGTGTGCAAGCCACACAAGGGAGAGATTGGGATTGTGTTCGAG : 1500

* 1520 * 1540 * 1560 * 1580 * 1600 * 1620 * 1640 * XI-NIA : CATCCGACGCTTCCTGGTAATGAATCTGGTGGATGGATGGCTAAGGAACGTCACCTCGAAAAATCGGCTGACGCTCCTCCTAGTCTAAAGAAGTCTGTCGATACTCCATTTATGAACACAACTGCTAAGATGTACTCTATGTCCGAGGTC : 1650 NIA2 : CATCCAACGCTTCCTGGTAATGAATCTGGTGGATGGATGGCGAAGGAACGTCACCTCGAAAAATCGGCTGACGCGCCTCCTAGTCTAAAGAAGTCTGTCTCGACGCCGTTTATGAACACAACTGCGAAGATGTACTCGATGTCCGAGGTC : 1650

Figure 2.7 Gene sequence alignment of Arabidopsis Nitrate Reductase (NIA2) and cassava codon optimized Nitrate Reductase (XI- NIA). The nucleotide pairs highlighted in gray belong to the modified codons due to codon optimization. The Serine 534 mutation to aspartate is highlighted in yellow. continued next page

Figure 2.7

1660 * 1680 * 1700 * 1720 * 1740 * 1760 * 1780 * 1800 XI-NIA : AAGAAGCATAATTCGGCTGACTCTTGCTGGATCATTGTCCATGGACATATCTATGATTGTACAAGATTCCTTATGGATCACCCAGGTGGTTCGGATTCAATCTTGATCAATGCTGGTACTGATTGTACTGAGGAGTTTGAAGCCATTCAC : 1800 NIA2 : AAGAAGCATAATTCGGCTGACTCTTGCTGGATCATTGTCCATGGACATATCTATGATTGTACACGATTCCTTATGGATCACCCGGGTGGTTCGGATTCAATCTTGATCAATGCTGGTACGGATTGTACGGAGGAGTTTGAAGCCATTCAC : 1800

* 1820 * 1840 * 1860 * 1880 * 1900 * 1920 * 1940 * XI-NIA : TCGGATAAAGCCAAGAAGATGCTTGAGGATTACCGTATCGGTGAACTTATCACCACTGGTTATTCCTCTGACTCTTCTTCGCCTAACAACTCGGTTCACGGTTCATCCGCCGTGTTCTCTCTGTTGGCTCCAATTGGAGAGGCTACTCCA : 1950 NIA2 : TCGGATAAAGCCAAGAAGATGCTTGAGGATTACCGTATCGGTGAGCTCATCACCACTGGTTATTCCTCTGACTCTTCCTCGCCTAACAACTCGGTTCACGGTTCATCCGCCGTGTTCTCGCTGTTGGCTCCCATTGGAGAGGCGACTCCG : 1950

1960 * 1980 * 2000 * 2020 * 2040 * 2060 * 2080 * 2100 XI-NIA : GTTAGGAACCTCGCTTTGGTTAATCCAAGAGCTAAAGTCCCAGTTCAACTCGTCGAAAAGACTTCCATTTCTCATGATGTTCGTAAATTCAGATTTGCTTTACCAGTTGAGGATATGGTTCTTGGCTTACCGGTTGGTAAGCACATTTTC : 2100 NIA2 : GTTAGGAACCTCGCTTTGGTTAATCCCCGGGCTAAAGTCCCGGTTCAACTCGTCGAAAAGACTTCCATTTCTCATGATGTTCGTAAATTCCGGTTTGCTTTACCGGTTGAGGATATGGTTCTAGGCTTACCGGTTGGTAAGCACATTTTC : 2100

* 2120 * 2140 * 2160 * 2180 * 2200 * 2220 * 2240 * XI-NIA : CTTTGCGCCACCATCAATGACAAGCTCTGCCTCAGAGCTTACACACCAAGCAGCACCGTTGATGTGGTTGGCTACTTCGAACTTGTGGTCAAGATTTACTTTGGCGGTGTCCACCCAAGATTCCCTAACGGCGGGCTCATGTCTCAGTAC : 2250 NIA2 : CTTTGCGCCACCATCAATGACAAGCTCTGCCTCAGAGCTTACACACCAAGCAGCACCGTTGATGTGGTTGGCTACTTCGAGCTCGTGGTCAAGATTTACTTTGGCGGTGTCCACCCAAGATTCCCTAACGGCGGGCTCATGTCTCAGTAC : 2250

2260 * 2280 * 2300 * 2320 * 2340 * 2360 * 2380 * 2400 XI-NIA : CTAGACTCTTTGCCTATAGGGTCAACTTTGGAGATTAAAGGACCATTGGGTCACGTTGAGTATCTCGGCAAGGGTAGTTTCACTGTTCACGGTAAACCAAAGTTTGCTGATAAATTGGCAATGTTGGCAGGTGGAACCGGAATTACTCCA : 2400 NIA2 : CTAGACTCTTTGCCTATAGGGTCAACTTTGGAGATTAAAGGACCATTGGGTCACGTTGAGTATCTCGGCAAGGGTAGTTTCACGGTTCACGGTAAACCAAAGTTTGCTGATAAATTGGCAATGTTGGCAGGTGGAACCGGAATAACTCCG : 2400

* 2420 * 2440 * 2460 * 2480 * 2500 * 2520 * 2540 * XI-NIA : GTTTACCAAATTATCCAAGCCATTCTCAAGGATCCAGAGGATGAGACTGAAATGTACGTCATTTATGCTAACAGAACCGAGGAAGATATTCTCCTAAGGGAGGAACTGGATGGTTGGGCAGAGCAATACCCAGACAGATTAAAGGTTTGG : 2550 NIA2 : GTTTACCAAATTATCCAAGCCATTCTCAAGGATCCAGAGGATGAGACTGAAATGTACGTCATTTATGCTAACCGGACCGAGGAAGATATTCTCCTAAGGGAGGAACTGGATGGTTGGGCAGAGCAATACCCGGACCGGTTAAAGGTTTGG : 2550

* 20 * 40 * 60 * 80 * 100 * 120 * 140 * XI-NIA : MAASVDNRQYARLEPGLNGVVRSYKPPVPGRSDSPKAHQNQTTNQTVFLKPAKVHDDDEDVSSEDENETHNSNAVYYKEMIRKSNAELEPSVLDPRDEYTADSWIERNPSMVRLTGKHPFNSEAPLNRLMHHGFITPVPLHYVRNHGHVP : 150 NIA2 : MAASVDNRQYARLEPGLNGVVRSYKPPVPGRSDSPKAHQNQTTNQTVFLKPAKVHDDDEDVSSEDENETHNSNAVYYKEMIRKSNAELEPSVLDPRDEYTADSWIERNPSMVRLTGKHPFNSEAPLNRLMHHGFITPVPLHYVRNHGHVP : 150

160 * 180 * 200 * 220 * 240 * 260 * 280 * 300 XI-NIA : KAQWAEWTVEVTGFVKRPMKFTMDQLVSEFAYREFAATLVCAGNRRKEQNMVKKSKGFNWGSAGVSTSVWRGVPLCDVLRRCGIFSRKGGALNVCFEGSEDLPGGAGTAGSKYGTSIKKEYAMDPSRDIILAYMQNGEYLTPDHGFPVRI : 300 NIA2 : KAQWAEWTVEVTGFVKRPMKFTMDQLVSEFAYREFAATLVCAGNRRKEQNMVKKSKGFNWGSAGVSTSVWRGVPLCDVLRRCGIFSRKGGALNVCFEGSEDLPGGAGTAGSKYGTSIKKEYAMDPSRDIILAYMQNGEYLTPDHGFPVRI : 300

* 320 * 340 * 360 * 380 * 400 * 420 * 440 * XI-NIA : IIPGFIGGRMVKWLKRIIVTTKESDNFYHFKDNRVLPSLVDAELADEEGWWYKPEYIINELNINSVITTPCHEEILPINAFTTQRPYTLKGYAYSGGGKKVTRVEVTVDGGETWNVCALDHQEKPNKYGKFWCWCFWSLEVEVLDLLSAK : 450 NIA2 : IIPGFIGGRMVKWLKRIIVTTKESDNFYHFKDNRVLPSLVDAELADEEGWWYKPEYIINELNINSVITTPCHEEILPINAFTTQRPYTLKGYAYSGGGKKVTRVEVTVDGGETWNVCALDHQEKPNKYGKFWCWCFWSLEVEVLDLLSAK : 450

460 * 480 * 500 * 520 * 540 * 560 * 580 * 600 XI-NIA : EIAVRAWDETLNTQPEKMIWNLMGMMNNCWFRVKTNVCKPHKGEIGIVFEHPTLPGNESGGWMAKERHLEKSADAPPSLKKSVDTPFMNTTAKMYSMSEVKKHNSADSCWIIVHGHIYDCTRFLMDHPGGSDSILINAGTDCTEEFEAIH : 600 NIA2 : EIAVRAWDETLNTQPEKMIWNLMGMMNNCWFRVKTNVCKPHKGEIGIVFEHPTLPGNESGGWMAKERHLEKSADAPPSLKKSVSTPFMNTTAKMYSMSEVKKHNSADSCWIIVHGHIYDCTRFLMDHPGGSDSILINAGTDCTEEFEAIH : 600

* 620 * 640 * 660 * 680 * 700 * 720 * 740 * XI-NIA : SDKAKKMLEDYRIGELITTGYSSDSSSPNNSVHGSSAVFSLLAPIGEATPVRNLALVNPRAKVPVQLVEKTSISHDVRKFRFALPVEDMVLGLPVGKHIFLCATINDKLCLRAYTPSSTVDVVGYFELVVKIYFGGVHPRFPNGGLMSQY : 750 NIA2 : SDKAKKMLEDYRIGELITTGYSSDSSSPNNSVHGSSAVFSLLAPIGEATPVRNLALVNPRAKVPVQLVEKTSISHDVRKFRFALPVEDMVLGLPVGKHIFLCATINDKLCLRAYTPSSTVDVVGYFELVVKIYFGGVHPRFPNGGLMSQY : 750

87 760 * 780 * 800 * 820 * 840 * 860 * 880 * 900 XI-NIA : LDSLPIGSTLEIKGPLGHVEYLGKGSFTVHGKPKFADKLAMLAGGTGITPVYQIIQAILKDPEDETEMYVIYANRTEEDILLREELDGWAEQYPDRLKVWYVVESAKEGWAYSTGFISEAIMREHIPDGLDGSALAMACGPPPMIQFAVQ : 900 NIA2 : LDSLPIGSTLEIKGPLGHVEYLGKGSFTVHGKPKFADKLAMLAGGTGITPVYQIIQAILKDPEDETEMYVIYANRTEEDILLREELDGWAEQYPDRLKVWYVVESAKEGWAYSTGFISEAIMREHIPDGLDGSALAMACGPPPMIQFAVQ : 900

* 920 XI-NIA : PNLEKMQYNIKEDFLIF*-- : 917 NIA2 : PNLEKMQYNIKEDFLIF*-- : 917

Figure 2.8 Protein sequence alignment of Arabidopsis Nitrate Reductase (NIA2) and cassava codon optimized Nitrate Reductase (XI-

NIA). The program GeneDoc was used for the sequence alignment of the translated protein sequences of NIA2 and XI-NIA. The

yellow star marks the residue belonging to Serine 534 in NIA2 and mutated to aspartate in XI-NIA. No other differences were

observed in the remaining residues.

2.3.2 Construction of the pCNIA vector

Construction of the final transformation vector was done in two steps. An initial cloning of XI-NIA in to the 3DVL vector, a pBI121 derived vector containing the patatin promoter and nos terminator. The patatin promoter: XI-NIA: nos terminator cassette was then cloned in to the pCAMBIA2300 binary vector giving the pCNIA vector (Fig. 2.9)

Figure 2.9 T-DNA region of pCNIA vector. This vector was constructed using pCAMBIA2300 and the patatin:CNIA:nos terminator cassette.

2.3.3 Production of transgenic cassava

To generate plants expressing the mutated nitrate reductase, Agrobacterium harboring the pCNIA vector was used to transform cassava friable embryogenic callus (FEC). In addition to the patatin promoter:XI-NIA:nos terminator cassette, the T-DNA included a constitutively expressed nptII gene for antibiotic selection of transformants. The FEC clusters incubated with Agrobacterium were matured and regenerated into plantlets using paramomycin selection to select for transformants (Fig. 2.10). A total of 399 friable embryogenic structures were initially transformed but only 68 matured and produced embryogenic structures under paramomycin selection. Out of those 68 paramomycin resistant embryos only 46 germinated and regenerated fully into plantlets. Each plant originated from an independent transformation event, all the transgenic lines were named

CNIA along with a number from 1 to 46.

Figure 2.10 Schematic of cassava plantlet regeneration from transformed FEC. The transformed FEC were grown in Stage 1 and Stage 2 embryo maturation media under the selection of 45 µM paramomycin, after Stage 2 the embryos that developed cotyledons were placed in the Embryo germination media under the selection of 45 µM paramomycin. After germination the plantlets were subcultured in MS2 media, no selection was used.

2.3.4 PCR and dot-blot analysis

The 46 CNIA lines were screened by PCR amplification for the XI-NIA gene. The presence of the XI-NIA gene was analyzed using the N37/N34R primer pair (Table 2.1) to amplify a 750 bp band (Fig. 2.11, select lines are shown). The product was confirmed to be XI-NIA by DNA sequence analysis. A total of 38 lines were confirmed positive by

PCR analysis for a transformation efficiency of 9.5%.

Dot blot analysis was used to determine the number of T-DNA integrations and to further confirm the results of our PCR screening. Dot blot analysis revealed a total of 15 lines with a single insertion, 10 lines with two insertions and 13 lines with three or more insertion. A total of 15 lines with varying number of insertions were chosen for further analysis and were grown in the greenhouse.

Figure 2.11 PCR Screening of in vitro transgenic plantlets. The results of PCR amplification of a 750 bp band from the XI-NIA gene using the primers N37 and N34R

(Table 2.1) are shown for a select number of the CNIA lines.

2.3.5 Analysis of cassava storage roots by semiquantitative RT-PCR and real-time RT-

PCR

The abundance of the XI-NIA gene transcript was analyzed in the storage root pulp of 15

CNIA lines and wild-type plants using 5 month old greenhouse grown plants. A semiquantitative RT-PCR analysis using the N37/N34R primer pair amplified the XI-

NIA gene transcript in all 15 CNIA line storage roots but not in wild type (Fig. 2.12).

Amplification of tubulin was used as a control. Through semiquantitative RT-PCR the

CNIA-11 line appeared to be the highest expressor followed by the CNIA-06 lines. The lowest apparent expressor was CNIA-14.

Figure 2.12 RT-PCR analysis of XI-NIA expression in storage roots of transgenic CNIA and wild-type plants. Amplification of a 750 bp band confirms expression of the XI-NIA gene in the storage roots of the plants. The positive control used was the pCNIA plasmid and the negative control was a reaction containing deionized water as a template. The housekeeping gene tubulin was used a control for our RNA extraction and cDNA reaction.

To determine the NIA expression levels quantitatively, real-time RT-PCR was used (Fig.

2.13). Included in our experiment were wild-type samples to be used as a control for null expression, wild-type was not used to calculate the relative expression of XI-NIA in the

CNIA lines instead the lowest expressor based on ΔCt (Ct tubulin – Ct XI-NIA) was selected. The CNIA-12 line was the lowest expressor and assigned a relative expression value of 1, in comparison to CNIA-12 the line CNIA-11 had 52 times the expression of

CNIA-12 and was the highest expressor obtained. These two lines along with three intermediate expression lines, CNIA-06, -14 and -15, were selected for further analysis.

d c b

Figure 2.13 Real-time RT-PCR analysis of the 5 selected CNIA lines. CNIA-12 had the lowest expression of the XI-NIA gene relative to tubulin and it was given a relative expression of 1. The expression of all other lines is relative to CNIA-12. CNIA-11 had the highest relative expression with a value of 52. (n=6, the error bars represent the standard error, a letter was used to denote statistically significant differences among the different lines, a similar letter indicates a p-value of > 0.05, while a different letter indicates a p-value of < 0.05)

2.3.6 Harvest weight

The primary source of nitrogen for cassava growing in the field is typically nitrate

(Moraes et al. 1981, Molina and El-Sharkawy 1995, Cruz et al. 2004). Prior to our work, there was no report on the effect of improved nitrate assimilation on storage root yield in cassava. To determine if expression of the NIA gene had an effect on yield it was important to analyze the yield of both storage root and shoots of wild-type and transgenic

CNIA lines.

Cassava wild-type and CNIA-06, -11, -12, -14 and -15 lines were grown for 5 months in the greenhouse. At harvest time the fresh weight of both the shoot and storage root was measured to determine the yield of the storage root and any effect on the shoot weight.

The storage roots of three CNIA lines (11, 12 and 14) had a 2 fold increase in fresh weight relative to wild-type, in addition a fourth line CNIA-06 had a 1.7 fold increase compared to wild-type roots (Fig. 2.14). Only CNIA-15 had no significant increase (Fig.

2.14). Line CNIA-11 which had the highest level of NIA expression based on RT-PCR also had the highest increase in root fresh weight accumulation.

* * * *

Figure 2.14 Storage root harvest weight of 5 month old greenhouse grown wild-type and transgenic CNIA lines. The storage root harvest yield was increased in several of the transgenic CNIA lines. (n=3 different plants of the same line, error bars indicate the standard error, an * indicates statistically different with a p-value < 0.05)

All CNIA lines had a reduction in their shoot fresh weights, however, four of those lines

(CNIA-06, 11, 12 and 14) did not have statistically significant decreases and only CNIA-

15 with a 50% reduction relative to wild-type had a significant reduction in shoot fresh weight (Fig. 2.15).

The total fresh weight (shoot + root fresh weight) of CNIA plants was increased from 3 to

44% with the exception of CNIA-15 which was reduced by 40% in overall fresh weight.

Only one line, CNIA-14, had a significant increase in total fresh weight. The ratios of shoot to root weight were reduced in CNIA lines relative to wild type. Wild-type lines have a shoot to root weight ratio of 4.6 while CNIA lines are close to 2. These results indicate the possibility of a shift in biomass accumulation occurring in CNIA plants. The increased yield of storage roots however may be an important agronomical quality for the later stages of use of these lines.

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Figure 2.15 Shoot harvest weight of 5 month old greenhouse grown CNIA and wild-type lines. No significant difference was observed between the majority of the CNIA lines and wild type shoot fresh weight (n=3 different plants of the same line, error bars indicate the standard error, an * indicates statistically different with a p-value < 0.05).

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Figure 2.16 Photos of cassava storage roots from wild type and transgenic CNIA lines. Representative pictures of the storage roots of

the five CNIA lines selected and wild-type at harvest time after 5 months of greenhouse growth.

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2.3.11 Western Blot

Expression of the mutated nitrate reductase protein in the cassava storage roots was confirmed by western blot using a specific antibody for the Arabidopsis nitrate reductase protein. The highest expressor according to the real-time RT-PCR was CNIA-11 and similarly this line appears to have the highest protein accumulation while the remaining lines had lower protein accumulation (Fig. 2.17).

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110 kDa

Figure 2.17 Western blot analysis of 20µg of crude protein from storage roots of CNIA lines and wild-type. The positive control used was crude protein extract from Arabidopsis leaves. A distinct band can be observed for the nitrate reductase protein in CNIA-11 with faint bands in CNIA-14 and 15.

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2.3.7 Storage Root Free Amino Acid Pool analysis

The expression of a mutated nitrate reductase lead to an increase in the free amino acid

(FAA) pool of N. plumbaginifolia (Lea et al. 2006). The total free amino acid levels of storage root pulp of 5 month old greenhouse grown cassava plants were analyzed and compared to wild-type plants to determine if there was a similar change in FAA (Lea et al., 2006). In relation to wild-type, all five transgenic lines had significant increases in the total FAA levels. Two of the lines, CNIA-06 and CNIA-14 had twice the FAA levels of wild type, these two lines interestingly were not the highest expressors according to the real-time RT-PCR experiment, they were considered as middle expressors. The highest expressor according to the real-time RT-PCR experiment was CNIA-11 and it had only a

1.5 fold increase in root FAA, the two lowest expressors CNIA-15 and CNIA-12 had the lowest increase in FAA among the CNIA lines, however, they were still significantly higher than wild type (Fig. 2.18).

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** **

Figure 2.18 Total FAA content was measured in the storage root pulp of the wild-type and the five CNIA lines. All five CNIA lines presented significantly increased total free amino acid content. (n=3 from 3 different plants, error bars indicate standard error, an * indicates statistically different as determined by a p-value of <0.05, an ** indicates statistically different as determined by a p-value of < 0.01)

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Lea et al. (2006) reported the analysis of the accumulation of individual essential amino acids in N. plumbaginifolia when the mutated nitrate reductase was constitutively expressed. The essential amino acids are defined as those amino acids that have to be provided from external sources in to the diet of humans. An increase was observed in all essential amino acids quantified by Lea et al. (2006) (only Leu and Phe were decreased).

Free glutamine, asparagine and proline had the highest accumulations. In the cassava storage roots of CNIA lines and wild type the individual free amino acid accumulations were quantified to determine if expression of a de-regulated nitrate reductase would result in an overall increase in all amino acids or if in cassava there would be any trends towards synthesis of a specific amino acid. All amino acids including essential and non essential were quantified in cassava storage roots.

For discussion, the amino acids were divided into 6 groups in relation to their biosynthetic pathways: glutamate family, aspartate family, alanine/valine/leucine group, serine/glycine/cysteine group, aromatic group and histidine.

The glutamate family of amino acids is derived from the common precursor ɑ- ketoglutarate and in addition to glutamate this group encompasses arginine, glutamine and proline. The free amino acid levels in transgenic plants were compared to wild type to determine the fold increase or decrease (Fig. 2.19).

Arginine was the most abundant free amino acid in the storage root FAA pool for wild- type and CNIA transgenic roots. The arginine levels in wild type averaged 43.8 nmol/mg

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dry weight while the CNIA lines reached levels as high as 80.8 nmol/g dry weight for

CNIA-06 or as low as 33.89 nmol/mg dry weight for CNIA-12 (Fig. 2.19). Four lines

(CNIA-06,11,14 and 15), had significantly higher arginine levels relative to wild-type.

The lowest NIA expressing transgenic line, CNIA-12, had a small non-significant decrease in arginine concentration. In wild type, arginine represented 57% of the total free amino acid pool. The high accumulation of arginine and its high contribution to the total free amino acid pool is not uncommon. Free arginine is a form of storable nitrogen in many plants (Guak et al., 2007; Llacer et al., 2008; Gao et al., 2009), in conifers arginine accounts for 46% of the total free amino acids (Canton et al., 2005) and in chicory tubers it is 70% of the total free amino acids (Ameziane et al., 1997). In CNIA lines arginine represented 36 to 55% of the total free amino acid pool.

Of special interest was the increase in glutamate and glutamine due to their participation in primary nitrogen assimilation. When nitrate is acquired from the soil it is reduced to nitrite which is further reduced to ammonium, the ammonium can then be assimilated by action of the glutamine synthase (GS)/glutamine:2-oxyoglutarate amidotransferase

(GOGAT) cycle. In the GS/GOGAT cycle ammonium and glutamine are condensed into glutamine. The amino group can then be transferred to oxoglutarate producing an additional molecule of glutamate. It was possible that the expression of XI-NIA would cause a marked increase in these two amino acids as a result of increased reduced nitrogen metabolism. All five CNIA lines had significant increases in the level of glutamine ranging from 2.2 to 4.5-fold increase compared to wild type, however, only a few lines (CNIA-06 and CNIA-14) had increased glutamate content.

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Proline was strikingly higher in the N. plumbaginifolia lines described by Lea et al.

(2006) in a similar manner proline was significantly increased in all 5 CNIA lines by up to 18.2 times that of wild type. Proline is one of the lower abundance amino acids in wild-type cassava and represents only 0.13% of the total amino acids, however, out of the total amino acids analyzed proline had one of the most significant increases.

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Figure 2.19 Analysis of the free amino acids from the glutamate family in storage root pulp. The pool sizes for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted on top of each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates p-value < 0.01)

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Figure 2.19

111 ** ** ***

** **

111

The second amino acid group evaluated was the aspartate family which includes aspartate, asparagine, methionine, threonine, isoleucine and lysine. The common precursor for all amino acids in this group is oxaloacetate, however, aspartate is the carbon skeleton donor for the synthesis of all other amino acids (Joshi and Jander 2010) with the exception of asparagine. Asparagine is also produced via the β-cyanoalanine synthase cyanide assimilation pathway (see Chapter 1). Importantly the asparate derived amino acids methionine, threonine, isoleucine and lysine are essential amino acids. Even more extensive research has been done to obtain higher methionine and lysine content in other crops where they are considered limiting such as cereals (lysine deficient) and legumes (methionine deficient) (Ufaz and Galili, 2008).

The quantification of free aspartate revealed a small but significant increase in four CNIA lines (CNIA-06, 11,12 and 15) (Fig. 2.20) Similarly, asparagine levels increased in four

CNIA lines (CNIA-06, 11,12 and 15) (Fig. 2.20). Overall, the levels of all four aspartate derived amino acids: methionine, threonine, isoleucine and lysine were increased significantly in all five CNIA lines.

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Figure 2.20 Analysis of the free amino acids from the aspartate family in storage root pulp. The level of each amino acid is presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value

< 0.05 and an ** determines p-value of < 0.01)

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Figure 2.20

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The third group analyzed was the alanine/valine/leucine group these three amino acids share the common precursor, pyruvate. All three amino acids were found in low abundance in cassava wild-type storage roots. Free alanine, valine and leucine represent only 0.8%, 0.3% and 0.08% of the total free amino acid pool. Alanine was significantly elevated in four CNIA lines (CNIA-06, 11, 14 and 15) in particular CNIA-14 had a 10.7 fold increase relative to wild type (Fig. 2.21). Importantly, valine and leucine are essential amino acids that are present in low abundance in wild-type cassava but were found to be significantly increased in all five CNIA lines by up to 17 fold for valine and almost 40 fold for leucine (Fig. 2.21).

The amino acids serine, glycine and cysteine share the precursor 3-phosphoglycerate.

Among these three amino acids, only serine was detected in our samples. It is possible that the abundance of cysteine and glycine is so low that it was not detected by the derivatization analysis used and therefore these amino acids are missing from our results.

In relation to serine accumulation all five CNIA lines were significantly increased (Fig.

2.21) by up to a 3.5 fold increase in relation to wild type.

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Figure 2.21 Analysis of the free amino acids from the alanine/valine/leucine group and serine in the storage root pulp. The levels for each amino acid are presented as a fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and ** indicates p-value < 0.01)

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Figure 2.21

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The aromatic amino acid group consists of tyrosine, phenylalanine and tryptophan (Fig.

2.21). Tyrosine was the least abundant detectable amino acid in our analysis of wild-type storage roots, representing only 0.05% of the total pool. Tyrosine levels were substantially increased in all five CNIA lines. Tyrosine levels increased 52 fold relative to wild type in line CNIA14. Phenylalanine and tryptophan levels were also elevated in multiple transgenic lines (Fig. 2.22).

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Figure 2.22 Analysis of the free amino acids from the aromatic group and histidine in cassava storage root pulp. The levels for each amino acid are presented as a fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates p-value < 0.01)

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Figure 2.22

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In general there was an overall increase in all detectable free amino acids in the storage roots of CNIA lines, although the magnitude of the increase varied between different transgenic lines possibly reflective of the non-uniform means by which the transgenes are integrated into the genome and differential patterns of expression. Additionally leucine and tyrosine the two amino acids with the lowest detectable abundance in wild-type roots had the greatest changes in abundance in transgenic plants relative to wild-type plants.

The two transgenic lines (CNIA-06 and CNIA-14) that had the highest total FAA levels also accumulated the highest levels for each of the 20 amino acids with the exception of

CNIA-14 which had a very marked increase in isoleucine, methionine, alanine, valine, leucine and tyrosine relative to CNIA-06. Furthermore, line CNIA-14 did not have a substantive increase in arginine while CNIA-06 did. The reason for the differential amino acid accumulation in these transgenic lines is not clear.

2.3.8 Leaf Total Free Amino Acid Pool

Amino acids are a common nitrogen currency in plants and throughout the different growth stages of plants amino acid transport occurs (William and Lee, 2001). In CNIA lines we observed an increase in the total storage root FAA pool by up to a 2 fold relative to wild type. The leaf FAA pool was measured to determine, the effect of the increase in

FAA in the storage roots on the total FAA of leaves.

The total free amino acid content was measured in the leaves of 5 month old wild-type and CNIA lines. The total FAA concentration in wild-type leaves was 56.3 nmol/mg fresh weight (Fig. 2.23). In lines 06, 14 and 15 the total FAA content was 40-60% greater than wild type (Fig. 2.23). The remaining two CNIA lines had no significant difference to

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wild-type in their FAA pool concentrations (Fig. 2.23). It is possible that the observed

FAA increases in the leaves are due to increased amino acid transport from the storage roots.

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Figure 2.23 Leaf total free amino acid content in cassava leaves of 5 month old greenhouse grown plants. (n=3 from three different plants of the same line, error bars indicate the standard error, an * indicates statistically different as determined by a p-value

< 0.05)

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2.3.9 Protein analysis

As part of the hypothesis it was proposed that an increase of the free amino acid pool in the storage root may lead to an increase in the protein content of this organ. The protein content in wild-type and CNIA lines storage roots was measured to determine if the increases observed in free amino acids translated in to higher protein. Since the FAA content was increased in the leaves, possibly due to transport from the storage roots, the protein content was measured in the leaf too.

The protein content of leaves and storage root pulp were analyzed in 5 month old greenhouse grown cassava. The protein levels vary significantly from one organ to the other, in wild-type leaves the average total protein was 2.3mg protein/mg dry weight, almost 10 times the value of total protein in the wild-type storage root pulp which was

24.8 µg protein/mg dry weight. This significant difference between leaf and storage root protein was expected and has been observed in cassava consistently where the leaves are a good source of protein while the storage roots are very limited (Diasolua et al., 2003;

Alves et al., 2005; Montagnac et al.,2009)

In our analysis of the CNIA lines, we did not observe a significant difference between the protein content of the storage root pulp of the transgenic lines and wild type (Fig. 2.24).

We did, however, observe significant differences in the protein levels in the leaves of three of the CNIA lines, (CNIA-11, 12 and 14), ranging from 20-40% greater than wild type (Fig. 2.25). The protein and amino acid increase observed in the leaves of CNIA lines are indicative of amino acid transport, it is likely that the amino acid transport is occurring based on sink-source relationships of amino acid and protein (Morandi 2005).

In sink-source relationships the source is the tissue of assimilation or synthesis while the

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sink is the point of utilization (Sonnewald and Willmitzer, 1992). The leaves are a natural major nitrogen sink (Hirel et al. 2005; Yasumura et al., 2007) therefore it would be natural to observe the transport of the free amino acids from the storage roots and subsequent integration in to protein as was observed in CNIA lines. The lack of increased protein synthesis in CNIA roots may be due to the lack of presence of an adequate nitrogen sink in this organ such as vegetative storage proteins (Muntz, 1998) which up to date have not been described in cassava storage roots (Shewry 2003).

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Figure 2.24 Analysis of the total soluble protein content of cassava storage root pulp in 5 month old greenhouse grown plants. The total soluble protein content from wild-type storage roots and transgenic CNIA lines was not significantly different. (n=3 from three different plants of the same line, an * indicates statistically different as determined by a p-value < 0.05)

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Figure 2.25 Analysis of the total soluble protein content of cassava leaves at 5 months of greenhouse growth. The crude protein content of leaves from transgenic CNIA lines was compared to that of wild-type, three lines CNIA-11, 12 and 14 had significantly greater

(20 to 40%) protein than wild type. (n=3 from three different plants of the same line, an * indicates statistically different as determined by a p-value < 0.05)

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2.3.10 Linamarin analysis

Linamarin has been found to be a transportable source of nitrogen in Hevea brasilensis

(Lieberei et al. 1985, Selmar et al. 1988) and it is presumed that in cassava linamarin may play a role as well in nitrogen metabolism (Siritunga 2002). It is hypothesized that linamarin acts as a source of reduced nitrogen transported from the leaves to the roots

(Siritunga 2002). If linamarin is involved in the overall nitrogen metabolism of cassava it is possible that the increase in reduced nitrogen in the form mainly of FAA may affect linamarin transport and accumulation the roots. The linamarin content was quantified in the CNIA lines to determine if expression of the XI-NIA gene in storage roots altered linamarin pool sizes in leaves or storage root pulp of 5 month old greenhouse grown wild-type and CNIA lines.

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In the storage root pulp the linamarin content of the wild type plants averaged 0.94 µmol linamarin/ mg dry weight, this value is in accordance to those reported in the literature for low cyanogenic or “sweet” cassava varieties (Mkpong et al., 1990; Santana et al., 2002).

The CNIA lines storage root pulp linamarin had significant decrease in cyanogenic glycoside content in the range of 34-44% compared to wild-type (Fig. 2.26). The reduction of linamarin content in the storage roots may indicate a lower rate of transport from the leaves to the storage roots perhaps in response to an increased availability of reduced nitrogen in the form of free amino acids.

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Figure 2.26 Linamarin content of storage root pulp from 5 month old greenhouse grown transgenic and wild-type cassava. (n=3 from three different plants of the same line, error bars indicate standard error, an * indicates statistically different as determined by a p- value < 0.05)

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The leaf linamarin content in wild-type averaged 31 µmol linamarin/mg dry weight. The linamarin content of leaves of the CNIA lines was significantly reduced (12.5 – 17.4

µmol linamarin/mg dry weight) representing a reduction of up to 60% (Fig. 2.27).

Linamarin synthesis takes place primarily in the leaves and then it is transported to the roots (Bediako et al., 1981; Andersen et al., 2000; Siritunga and Sayre, 2003). The decrease of linamarin observed in the leaves can therefore be related to a possible increase in linamarin transport to the roots or a decrease in linamarin synthesis. The decrease of linamarin content in the storage roots likely indicates that increased transport of linamarin from the leaves to the roots is not present but rather that the decrease observed in the leaf linamarin content is related to a decrease in linamarin synthesis. If linamarin is part of the nitrogen metabolism and acts as a reduced source of nitrogen for the storage roots it is expected that linamarin synthesis and transport be regulated along with the total nitrogen status of the plant.

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Figure 2.27 Linamarin content of leaves from 5 month old greenhouse grown transgenic and wild-type cassava. (n=3 from three different plants of the same line, error bars indicate standard error, an * indicates statistically different as determined by a p-value <

0.05)

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2.3.12 Growth experiment in ammonia free media

In past reports a marked decrease in linamarin synthesis in the leaves resulted in a high decrease in linamarin levels in the roots (<1% of wild type) (Siritunga and Sayre, 2003) It was discovered that transgenic plants having reduced capacity to produce linamarin in their leaves required a reduced source of nitrogen (such as ammonia) in their growth media in order to survive (Siritunga 2002). It was proposed that linamarin made in the leaf served as a source of reduced nitrogen for roots. To further test this hypothesis, we determined whether the CNIA lines, having presumably enhanced ability to produce reduced nitrogen in roots, would have altered patterns of growth relative to wild type in media without a reduced nitrogen source. Cassava plantlets were grown in MS2 and two

- ammonia free media, one media contained 39.4 mM NO3 and the other contained 18.8

- mM NO3 (Fig. 2.28).

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Figure 2.28 In vitro growth experiment of wild-type, CNIA-11, CNIA-12 and CNIA-15 in MS2 media and two different ammonium

- - free media. NH4 free media #1 contains 39.4 mM NO3 and NH4 free media #2 contains 18.8 mM NO3 .

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The three CNIA lines studied in this experiment had fresh or dry weights comparable to wild type when grown in MS2 media (Fig. 2.28). When CNIA and wild-type plantlets

+ were grown in NH4 free #1 media the average total dry weight of the plantlets dropped by 39% to 71% in comparison to the same line growing on MS2 media (Fig. 2.29), therefore there is a generalized effect on wild-type and CNIA lines regarding growth on media without reduced ammonia. However, comparison of only wild-type and CNIA

+ plantlets growing in NH4 free #1 media reveals that three of the CNIA lines had on average 1.4 to 2.2 times higher dry weight than wild-type when grown on media without reduced nitrogen, indicating that CNIA lines were able not only to grow adequately in media without ammonia but in fact better than wild type when supplied only with nitrate.

+ + The second media used in this experiment was NH4 free #2, this media similarly to NH4 free #1 does not contain a major source of reduced nitrogen but in difference to the first media it contains half the quantity of nitrate and therefore half of the total nitrogen source. The CNIA lines growing in this lower nitrate content media had greater dry

+ weights than those grown in NH4 free #1 media, comparing CNIA lines in media #2 with those grown in regular MS2 media there is no significant difference in growth, indicating once more the ability of the CNIA lines to grow without a reduced source of nitrogen. (Fig. 2.29).

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* a c* * b a

Figure 2.29 Average dry weight of each plantlet grown in the three different media for wild-type and three CNIA transgenic lines. (n=3 from three different plants of the same line, error bars indicate standard error, a letter indicates statistical significance among the plants of the same line grown in different media, an * indicates statistically different values comparing the dry weight of CNIA lines in media #1 to wild type grown in the same media, statistical significance was considered as determined by a p-value < 0.05)

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2.3.13 Analysis of calorie content in cassava storage roots

Cassava is an important source of calories for the millions of people that consume it regularly and it has one of the highest energy yields (kJ/ha) among the important crops of the world (El-Sharkawy 2004). The nutritional improvement of cassava should not be at the recourse of diminishing the caloric content of the crop considering the importance of cassava as an energy source.

The calorie content of cassava storage root pulp was determined for wild-type and CNIA lines, the obtained values ranged from 3896 cal/g dry weight to 4069 cal/g dry weight.

There was no significant difference between wild-type and the five transgenic lines in energy content (calories/g dry weight) (Fig. 2.30).

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Figure 2.30 Calorie content of cassava storage roots. Total calorie amounts from wild- type and CNIA lines storage root pulp were measured and no significant difference was observed between them. (n=3 from three different plants of the same line, error bars indicate standard error, an * indicates statistically different as determined by a p-value <

0.05)

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2.3.14 Summary of CNIA lines

In transgenic cassava CNIA lines a mutated nitrate reductase (S534D) was expressed using the patatin promoter, a promoter previously used for expression in cassava storage roots. In Figure 2.31 a summary of the overall impact of the expression of this mutated nitrate reductase is presented. In all lines a significant increase in total free amino acid content in the storage roots was observed. There was up to a 2 fold increase relative to wild-type in some lines. The increase in free amino acid content may be related to an increase in the reduction of nitrate leading to increased ammonia accumulation which can be incorporated into amino acid biosynthetic pathways. In CNIA lines no protein increase was observed in the storage roots. In the leaves of CNIA lines an increase in free amino acid and protein content was observed. This was likely due to amino acid transport.

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Figure 2.31 Summary of the effect on protein, free amino acid and linamarin content in

CNIA lines. The free amino acid content was increased in the storage roots of CNIA lines but no protein increase wasobserved. An increase in free amino acid and protein content was observed in the leaves of CNIA lines. All bold arrows with orange outline indicate increase (pointing up) or decrease (pointing down). (FAA= free amino acid, N = nitrogen)

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2.4 DISCUSSION

Cassava storage roots are widely consumed in Sub-Saharan Africa due to the food security it provides. However, the protein content of cassava storage roots is among the lowest of the major staple crops. Protein content in cassava varies but it can be as low as

0.7% (Ceballos et al. 2006).

- + In plants the assimilated nitrogen (as NO3 or NH4 ) influences the biosynthesis and accumulation of amino acids and protein (Maldonado et al. 1996, Matt et al. 2002, Martre et al. 2003, Lea et al. 2006). We proposed to increase the free amino acid and protein content in cassava storage roots by expressing a mutated Arabidopsis nitrate reductase enzyme (NIA) to increase the assimilation of nitrate (Lea et al. 2006). In the mutated

NIA2 the Serine-534 residue is mutated to aspartate. The Ser-534 site is a key post- translational regulatory site that when phosphorylated mediates 14-3-3 protein binding to

NIA2 (Kanamuru et al. 1999, Lambeck et al. 2010). The binding of the 14-3-3 protein inactivates NIA2 (Lillo et al. 1997, Kanamaru et al. 1999) eliminating the reduction of nitrate to nitrite. Past reports have shown that exchange of the key regulatory serine for an aspartate residue causes the activity state of the nitrate reductase to increase and remain constant (Lillo et al. 2003, Lea et al. 2006). The S534D mutated Arabidopsis nitrate reductase was codon optimized for expression in cassava and named XI-NIA; the transgenic cassava lines carrying this gene are identified as CNIA lines.

In the transgenic cassava CNIA lines the root-specific expression of the mutated nitrate reductase resulted in as much as a 2-fold increase in the free amino acid pool of the storage roots. Lea et al. (2006) observed a similar result, a 2 fold increase in the leaf free

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amino acid pool by constitutively expressing using the 35S promoter the N. plumbaginifolia nitrate reductase bearing a mutation in its regulatory serine (Serine-521).

Lea et al. (2006) observed a 30% decrease in nitrate content and an increase in ammonia by 500% indicative of increased nitrate reduction. Interestingly, among the different

CNIA transgenic cassava lines, the CNIA-11 line, the highest transgene expressor as determined by real-time RT-PCR (52 fold higher than lowest expressor) and western blot analysis did not have the highest free amino acid levels. The lines CNIA-06 and CNIA-

14 had the greatest increase in the free amino acid pool (2 fold). How overall nitrogen status in cassava is controlled is not understood. The regulation of nitrate uptake by nitrate transporters (Tsay et al. 2007), amino acid transport and recycling between the xylem and phloem (Forde et al. 2002) as well as the feedback inhibition loops found in amino acid biosynthetic pathways (Ufaz and Galili, 2008; Jander and Joshi, 2010) are all possible key regulatory points involved in reduced nitrogen homeostasis in plants.

Therefore the high expression present in the CNIA-11 line may not necessarily elicit the highest accumulation in free amino acids.

It was hypothesized that an increase in the storage root FAA pool would result in an increase in root protein levels, however, no significant difference in root protein abundance was observed between wild-type and the CNIA transgenic lines. Upon further analysis of the CNIA lines it was observed that several lines had increased FAA and/ or protein in the leaves. Amino acids are a preferred transportable form of organic nitrogen

(Wipf et al. 2002) and transport of amino acids occurs at all growth stages (William and

Lee 2001). The transport of amino acids presumably is regulated by the source-sink relationships that apply to many metabolites. A source tissue is considered as the site of

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assimilation and synthesis of a metabolite. Sink tissues can be categorized as: 1) utilization sinks that are highly metabolically active in growing tissues, and 2) storage sinks such as seeds and tubers (Sonnewald and Willmitzer 1992). It is proposed throughout the literature that green photosynthetically active and developing leaves, meristems and reproductive organs are all major nitrogen utilization sinks (Ortiz-Lopez,

2000, Hirel et. al. 2005, Yasumura et al., 2007). In cassava it is therefore likely that the leaf tissues, in particular photosynthetically active and developing leaves, are a major nitrogen sink. The observed increase in free amino acid content in the leaves may be due to mobilization of storage root amino acids to satisfy the needs of the leaf nitrogen sink

(Ortiz-Lopez, 2000).

The increase of FAA and protein in CNIA leaves along with no increase in storage root protein may be due to the absence of a major nitrogen sink. To date there are no known storage proteins in cassava storage roots (Shewry, 2003; Sheffield et al., 2006; Li et al.,

2010). Storage proteins represent an important amino acid sink (Muntz et al. 1998) and are present in other starchy storage roots such as sporamin in sweet potato (Shewry,

2003).

Interestingly the accumulation of the cyanogenic glucoside linamarin was reduced in both leaves and storage roots of transgenic plants. A role for linamarin in the nitrogen metabolism of cassava has been suggested (Siritunga, 2002; Siritunga and Sayre, 2003).

Linamarin as a transportable source of reduced nitrogen was first observed in Hevea brasilensis seedlings (Lieberei et al., 1985; Selmar et al., 1988). In cassava, linamarin is synthesized in the leaves and then transported to the root system (Bediako et al., 1981;

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Andersen et al., 2000; Siritunga and Sayre, 2003). In cassava plants with reduced leaf linamarin synthesis the root linamarin was reduced to < 1% of wild type indicating that the root linamarin content depended on the transport of this cyanogenic glucoside from the leaves (Siritunga and Sayre, 2003). Importantly, these plants required an external source of reduced nitrogen to survive (Siritunga 2002). It was hypothesized that in cassava linamarin functions as both a transportable and a storable source of reduced nitrogen (Siritunga, 2002). If linamarin is integrated in to the overall nitrogen metabolism of cassava its synthesis and transport would be subjected to control by the overall N- status of the plant. The sensor(s) that regulate N-status have yet to be identified (Miller et al. 2007). It is in fact possible that N-status signaling is species specific (Miller et al.

2007). For example, in Arabidopsis and barley nitrogen uptake and assimilation seem to be correlated to amino acid levels (Fan et al. 2006, Miller et al. 2007, review). However, in Brassica napus amino acid pool sizes do not seem to be correlated with nitrate uptake

(Laine et al. 1995). The N-status sensors are not known in cassava however it is possible that the increase in reduced nitrogen as free amino acids in the storage roots was responsible for the decrease in linamarin observed in leaves and storage roots.

The past report in N. plumbaginifolia of expression of a similarly mutated nitrate reductase (Lea et al. 2006) identified an increase in the total FAA pool. Furthermore the individual free essential amino acid quantities were evaluated and an increase in the majority of the essential amino acids in the pool was observed (Lea et al., 2006).

Similarly, in CNIA plants the majority of the free amino acids were increased.

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Of particular interest were lysine and methionine. These two amino acids have been identified as the most limiting essential amino acids in major sources of human food and animal feed (Ufaz and Galili, 2008). Lysine is limited in cereals while methionine is in legumes (Ufaz and Galili, 2008). Past attempts at improving the accumulation of these two amino acids have focused on their biosynthetic pathways (Karchi et al., 1994; Zhu and Galili, 2004; Avraham et al. 2005; Frizzi et al., 2008; Lee et al., 2008; Ufaz and

Galili, 2008). Interestingly we observed in CNIA lines through the general increase of

FAA amino acid that a comparable accumulation of free lysine and free methionine can be obtained as when specific biosynthetic pathways are modified. Karchi et al. (1994) increased the lysine content in tobacco seeds up to 1.5 µmol lysine/ g dry weight through the expression of a feedback insensitive bacterial dihydrodipicolinate synthase and aspartate kinase. Interestingly the transgenic cassava lines (CNIA-06 and 14) that accumulated the highest FAA, had free lysine levels comparable to those obtained by

Karchi et al. (1994), the CNIA lines had free lysine levels in the storage roots of 1.5 µmol lysine/ g dry weight and 1.8 µmol lysine/g dry weight, respectively. Lee et al. (2008) reduced the activity of homocysteine methyltransferase in Arabidopsis and reported levels of methionine of 1800 nmol methionine/ mg dry weight. In one of the high CNIA accumulating lines, CNIA-14, the storage root had nearly double that reported by Lee et al. (2008) reaching 3573 nmol methionine/mg dry weight. These results suggest that enhancing overall nitrate reduction rather than engineering specific amino acid biosynthetic pathways may be an alternative approach for increasing overall free amino acid accumulation.

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Among the individual free amino acids analyzed free arginine was identified as the most abundant free amino acid in storage roots of wild-type and transgenic lines. In wild type plants it accounted for 57% of the total free amino acids accumulated. Free arginine is a form of storable nitrogen in plants (Guak et al., 2007; Llacer et al. 2008; Gao et al.,

2009). Arginine has been observed to be abundant in cassava storage roots as part of protein bound amino acids (14.5% of total) (Crawford et al., 1970; Nassar and De Soussa,

2007; Stupak 2008). A previous report mentions the abundance of arginine in the cassava storage root free amino acid pool, however, the amount of increase was not provided

(Morante et al., 2004). This is the first report of quantification of free arginine in cassava storage roots and its analysis of abundance in relation to the total free amino acid pool. It is likely given the evidence that free arginine in cassava storage roots serves as a form of nitrogen reserve. Importantly free arginine was increased in the storage roots of several

CNIA lines, however, the contribution of free arginine to the overall free amino acid pool of the storage roots in CNIA lines was only 36 to 55%. This may indicate that the increase in the total free amino acid pool in CNIA lines is not through increased storage of reduced nitrogen in the form of free arginine but rather is due to an overall increase in the synthesis of all amino acids. This is an important consideration for de novo protein synthesis as it has been linked to the availability of amino acids in situ (Sands et al.

2009).

Other amino acids found in abundance and of particular interest were glutamate and

+ glutamine, these two amino acids are involved in the primary assimilation of NH4 in to amino acids by the glutamate synthase (GS) / glutamine 2-oxoglutarate amido transferase

(GOGAT) cycle. It has been observed that when the nitrogen metabolism of a plant is

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affected the free glutamine levels but not those of free glutamate become a good marker for increased nitrate reductase activity and possible increased nitrate assimilation (Foyer et al. 2003). In N. plumbaginifolia plants with altered NR activity, the free glutamine content fluctuated in response to the NR activity but glutamate levels remained constant

(Foyer et al. 1994, Gojon et al. 1998). In a similar manner over expression of a deregulated nitrate reductase bearing a mutation in its regulatory serine (S521D) in N. plumbaginifolia leading to increased nitrate reductase activity was associated with increased free glutamine (9 fold higher than wild type) but not glutamate (1.5 fold higher than wild type) (Lea et al. 2006). All five transgenic cassava CNIA lines had significantly increased (2.2 to 4.5 fold) free glutamine levels relative to wild type while the glutamate levels were not significantly increased in the majority of the lines (CNIA-11, 12 and 15).

The increased glutamine levels in CNIA plants are consistent with past reports of higher glutamine levels caused by modification in the nitrate reductase expression and may be an indicator of greater nitrate assimilation (Foyer et al., 1994; Gojon et al., 1998; Foyer et al., 2003; Lea et al., 2006)

In addition to the aforementioned strategies to increase free lysine and methionine, other attempts at engineering amino acid levels have produced increased concentrations of select free amino acids such as histidine by over-expression of the ATP- phosphoribosyltransferase in Arabidopsis (Rees et al. 2009), asparagine by over- expression of asparagine synthase in tobacco (Brears et al. 1993), tryptophan by expression of a feedback insensitive ɑ subunit of anthranilate synthase in rice, potato and soybean (Yamada et al. 2004, Ishimoto et al. 2010). Synthesis of branched amino acids by impairment of isovaleryl-CoA-dehydrogenase in Arabidopsis (Gu et al. 2010) and of

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cysteine and methionine jointly by over expression of cystathionine γ-synthase in

Arabidopsis (Avraham et al. 2005). The CNIA lines differ from these past reports in that it is the total free amino acid content in cassava storage roots that were increased and not a single or few amino acids, Rademacher et al. (2002) also observed a general increase in

FAA by constitutively expressing a modified phosphoenol pyruvate carboxylase (PEPC) that was less sensitive to inhibition by malate in potato. This enzyme is involved in carbon metabolism and its deregulation may have an impact on amino acid biosynthesis

(Yanagisawa et al. 2004). However, the transgenic potato plants expressing the modified

PEPC had retarded growth, and a decrease in tuber yield and starch content. The CNIA lines on the contrary, had a high storage root yield not associated with an increase in energy density in the biomass.

In summary it was possible to express a de-regulated Arabidopsis nitrate reductase to increase the free amino acid pool in cassava storage roots. However, this increase in FAA did not result in an increase in root protein levels. Increased protein and free amino acid content in the leaves of the transgenic plants expressing NR in roots suggests that additional reduced nitrogen was transported to the leaves which are a strong amino acid sink. It is possible that by combining the increased synthesis of amino acids in cassava storage roots with the expression of a storage protein the protein level may increase in the storage root. Such strategy is known as a binary or a sink-source strategy (Beauregard and Hefford 2006), this has been proposed for methionine (Beach and Tarckzynski 2000,

Kirihara et al. 2001), lysine (Falco et al. 1998, Guenoune et al. 2002) and tryptophan

(Kita et al. 2010) in storage protein. Co-expression of a storage protein and the mutated

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nitrate reductase may result in the retention of free amino acids as protein in cassava storage roots. However, results presented in this thesis in Chapter 3 may indicate that the sole expression of a storage protein does not provide a strong storage nitrogen sink in comparison to the leaf. Further discussion is presented in Chapter 3.

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CHAPTER 3

EXPRESSION OF A VACUOLAR TARGETED LINAMARASE AND

THE STORAGE PROTEIN SPORAZEIN TO INCREASE THE FREE

AMINO ACID POOL AND PROTEIN CONTENT IN CASSAVA

STORAGE ROOTS

3.1 INTRODUCTION

Cassava is an important staple crop in many regions of the world especially in Sub-

Saharan Africa. Cassava has many valuable agronomical traits particularly for subsistence farmers including; high productivity and starch content, tolerance to drought, ability to grow in poor soils, and herbivore resistance (McMahon et al., 1995 review).

However, cassava is a poor source of protein, it has one of the lowest protein to energy ratios among the world’s major staple crops . Cassava roots contain only 0.7-3% (g/g) protein (Ceballos et al.,1996; USDA, 2010) while other staples such as maize and soybean contain 9.4% and 12.4% protein, respectively (USDA, 2010).

Another constraint for cassava foods is the high concentration of cyanogenic glucosides present in all tissues except the seeds (McMahon et al., 1995 review). Fresh cassava storage roots can be classified as low cyanogenic or “sweet” if they contain less than 100 mg cyanogens/kg fresh weight and as high cyanogenic or “bitter” if they contain 100 –

500 mg cyanogens/kg fresh weight (Wheatley et al. 1993), where cyanogens are defined

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as any compound capable of releasing cyanide. In cassava leaves generally contain 5 to

10 times the cyanogenic glucoside levels of storage roots. Leaves can have up to 5 g cyanogens/kg fresh weight. (Wheatley et al., 1993; Montagnac et al., 2009). Linamarin represents 95% of the cyanogenic glucosides in cassava (Balogopalan et al., 1988).

Linamarin synthesis takes place primarily in the leaves and then it is transported to the roots. Siritunga et al. (2003) reported that in plants where linamarin synthesis had been inhibited exclusively in the leaves had reduced root linamarin contents that were < 1% that of wild type. In addition, analysis of phloem exudates and stem girdling experiments have shown the presence of linamarin in the phloem and the accumulation of linamarin at the site above the girdle (DeBrujin, 1971; Nambisan, 1996). These results point towards the transport of linamarin from the site of synthesis in the leaf to the root.

Cyanogenesis, or the generation of cyanide from linamarin, takes place by two sequential reactions; the first is deglycosylation of linamarin by the enzyme linamarase to produce acetone cyanohydrin and glucose. The second is the cleavage of acetone cyanohydrin to release cyanide and acetone either spontaneously at high pH (pH>5), high temperature

(>35 ˚C) or by the enzyme hydroxynitrile lyase (HNL) (White et al., 1998). In cassava it has been shown that there is compartmentalization of the cyanogenic glucosides and the enzymes responsible for its degradation and release of cyanide. Linamarin is stored in the vacuoles (White et al., 1994) and both linamarase and HNL are found in the apoplast and laticifers of leaves (Mkpong et al., 1990; Santana et al., 1992; White et al., 1994).

Interestingly, HNL is not expressed in cassava roots (White et al., 1998). A rapid release of cyanide would generally only occur after a tissue disruption that results in the interaction of the β-glucosidase with linamarin (Nahrstedt, 1985).

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Linamarin is not the only possible source of cyanide in cassava. All plants produce cyanide as a by-product of ethylene biosynthesis (Peiser et al., 1984) and have the ability to fix this toxic compound via two pathways. One pathway is mediated by β- cyanoalanine synthase (CAS) (Blumenthal et al., 1968) and the other is initiated by rhodanese (Chew , 1973) (Fig. 1). To date, all plant tissues have detectable CAS activity while rhodanese activity is far less common in other plant tissues (Miller and Conn,

1980; Elias et al., 1997). In cassava, the storage roots do not have any detectable rhodanese activity while CAS activity in the roots is 3 times higher than in leaves.

(Zidenga, 2011). It is possible that in cassava storage roots only the CAS pathway is present. In the CAS pathway cyanide is initially condensed with cysteine via β- cyanoalanine synthase (CAS) to synthesize β-cyanoalanine. Subsequently, β- cyanoalanine hydrase synthesizes asparagine from β-cyanoalanine and water. Finally, in the last step of the CAS pathway asparagine is deaminated by asparaginase to release asparate and ammonia. The CAS pathway is considered the cyanide assimilation pathway because the carbon and nitrogen atoms of the cyanide molecule are incorporated in to the overall metabolism of the plant through the synthesis of amino acids. On the other hand the rhodanese pathway detoxifies cyanide.

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Figure 3.1 Cyanide assimilation and detoxification pathways in plants. The β- cyanoalanine synthase (CAS) pathway involves two more enzymes a β-cyanoalanine synthase and asparaginase and the end products are aspartate and ammonia. The rhodanese pathway involves only the rhodanese enzyme producing as end products thiocyanate and sulfite which is presumably re-assimilated.

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Several functions of cyanogenic glucosides have been proposed for plants. It is generally accepted that cyanogens provide defense against herbivore predation (Vetter, 1999;

Gleadow and Woodrow, 2002). In some plants, it has also been demonstrated that cyanogens are a transportable form of reduced nitrogen. In Hevea brasilensis linamarin is accumulated in the seed endosperm (Lieberei et al. 1985). During germination the seed cyanogenic content is greatly reduced while at the same time no release of cyanide is observed (Lieberei et al. 1985). It was shown that linamarin through the formation of an intermediary diglucoside known as linustatin is transported throughout the seedling

(Selmar et al., 1988). However, the cyanogens present in the seedling do not account for the total cyanogens lost in the seed. In addition the enzymes capable of the catabolism of linamarin such as those from the CAS pathway were highly active in the cotyledons of

Hevea brasilensis (Lieberie et al. 1985). It was therefore suggested that the cyanogenic glucosides were incorporated into the primary metabolism in the developing Hevea brasilensis plant as a nitrogen resource compound (Selmar et al., 1988; Selmar 2010).

More recently, cyanogenic glucosides have been proposed to serve as a renewable source of nitrogen during latex regeneration (Kongsawdworakui et al., 2009). In cassava it was observed by Siritunga (2002) that transgenic lines with reduced linamarin synthesis in the leaves required reduced nitrogen in their growth media or the plantlets would not form roots and eventually perish. These results strongly suggested that linamarin plays a role as a transportable form of reduced nitrogen in growing and mature cassava. In addition, it is proposed that in cassava storage roots the CAS pathway functions as a cyanide assimilation pathway potentially for amino acid synthesis.

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Our objective was to enhance the ability to utilize the nitrile group (-CN) of linamarin as a source of reduced nitrogen in cassava storage roots with the aim to increase the free amino acid pool and protein content in this organ. Our strategy was to increase the deglycosylation of linamarin in vivo resulting in the production of acetone cyanohydrin and consequentially increasing the release and re-assimilation of cyanide into aspartate and ammonia via the CAS pathway. Aspartate is the precursor of a family of amino acids, including the essential amino acids threonine and methionine (Jander and Joshi, 2010 review). In addition, ammonia can be incorporated in to primary nitrogen metabolism for the synthesis of amino acids (Potel et al., 2009). The increased production of aspartate and ammonia would potentially lead to an increase in the free amino acid pool available for protein synthesis. To increase the hydrolysis of linamarin in vivo it was proposed to target the linamarase to the cytoplasm by removing the signal peptide that targets this enzyme to the cell wall. During linamarin transport to its storage organelle the vacuole the cyanogenic glucoside could potentially be hydrolyzed by cytoplasmic linamarase.

Another proposed strategy was to target linamarase directly to the vacuole by addition of a vacuolar targeting domain from barley lectin shown to be sufficient for protein targeting to the vacuole (Bednarek et al. 1991). Linamarin is stored in the vacuole which also has a lower pH (4.5) than the cytoplasm (7.5) and so the spontaneous rate of conversion of acetone cyanohydrin to cyanide in the two targeting strategies for linamarase would presumably differ significantly (White et al., 1994).

Amino acids and protein interact in a source –sink relationship. A source tissue is defined as the site of synthesis or assimilation of a metabolite (Sonnewald and Willmitzer, 1992) and a sink is where the metabolite is utilized, for example specialized storage proteins

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such as sporamin of sweet potato are a sink for amino acids (Muntz, 1996). Up to this moment there have been no storage proteins identified in cassava storage roots (Shewry,

2003; Sheffield et al., 2006; Li et al., 2010). Therefore, it was a concern as to whether increasing root free amino acid levels by the expression of a cytoplasmic or vacuolar targeted linamarase would elevate root protein levels or rather if without a strong amino acid sink in the roots the amino acids or reduced nitrogen would be transported to nitrogen sink tissues such as young developing leaves (Sonnewald and Willmitzer, 1992).

Our second objective was to determine if the root-specific expression of a storage protein in conjunction with the vacuolar linamarase would increase the retention of free amino acids in the storage root and increase the protein content. The novel storage protein sporazein (Abhary et al., 2011 in press) was selected. It is derived from sporamin a sweet potato storage protein (Shewry, 2003) and β-zein a maize protein (Bagga et al., 1997).

This novel protein has been shown to form stable protein bodies in tobacco (Abhary et al., 2011) and in cassava (Abhary, personal communication).

The results in this chapter demonstrate the feasibility of augmenting the free amino acid pool through an increase on in vivo linamarin deglycosylation. The increase in free amino acids in plants expressing a vacuolar linamarase with or without co-expression of a storage protein resulted in a decrease in linamarin in both storage roots and leaves. In addition, it points to the presence of a link between the cyanogenic glucosides and nitrogen metabolism in cassava, similar to what has been proposed in Hevea brasilensis

(Kongsawadworakul et al., 2009; Selmar, 2010). The protein content of cassava storage roots was only increased in plants co-expressing sporazein indicating perhaps the need for adequate sinks in cassava for protein increase and that perhaps the ability of de novo

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protein synthesis in the cassava storage roots limits overall root protein nutritional improvement of this crop.

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3.2 MATERIALS AND METHODS

3.2.1 E.coli and Agrobacterium strains

3.2.2 Chemical Reagents and supplies

The chemical reagents used in all experimental procedures were purchased from Fisher

Scientific, Hampton, NH unless otherwise noted. All plastic products such as microfuge tubes and PCR tubes were purchased from USA Scientific, Ocala, Florida.

3.2.3 Cassava cultivar and in vitro and greenhouse growth

The cassava cultivar TMS60444 was used for genetic transformation by the method of friable embryogenic callus (Taylor et al. 1996)(See section Generation of Transgenic

Callus).

Cassava plantlets were grown in vitro on Murashige and Skoog media (1962) supplemented with 20 g/L sucrose, MS Vitamins (Sigma Aldrich, USA) and 2.4g/L of the gelling agent, phytagel. This media will be subsequently known as MS2 . All media used for in vitro propagation of cassava was sterilized through autoclaving. The growth

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chamber conditions were set at a temperature of 28 ˚C, a light intensity of 75 to 100 µmol photons m-2 s-2 and a 16 hr day/8 hr night cycle.

Six week old in vitro grown cassava plantlets were then transferred to the greenhouse.

They were potted in 4” by 4” in pots with Fafard 51 soil mix (Fafard, Agawam, MS) and supplemented with 15:16:17 (N:P:K) fertilizer or Jack’s 10:30:20 (JR Peters, Allentown,

PA) twice a week. At 4 weeks after initial transfer to soil the plantlets were transferred to

6” by 6” pots.

3.2.4 General molecular techniques

All the PCR reactions were run in a BioRad Mycycler thermal cycler (BioRad, Hercules,

(w/v) agarose gel in TAE buffer (40 mM Tris-acetate, 1 mM EDTA, pH=8) and purified using Qiaquick Gel Extraction kit according to the instructions set by the manufacturer

(Qiagen, Valencia, CA). The concentration of the final purified product was determined using a Nanodrop 2000 Spectrophotometer (Fisher Scientific) for nucleic acids, with one unit of absorbance at 260 nm equivalent to 50 ng/µL for double stranded DNA and 40 ng/µL for single stranded RNA. The quality of the product was also determined by 260 nm/230 nm and 260 nm/280 nm ratios between 1.8 and 2.3. All digestions were carried out using 500 ng of purified template with 20 units of specific restriction enzyme in its

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preferable buffer. All ligations were carried out using a 1:3 ratio, being 50 ng of vector to

150 ng of insert, using 400 units of T4 DNA ligase (NEB) in 1X T4 DNA ligase buffer.

3.2.5 Cloning of the linamarase gene

Leaves from in vitro grown cassava plantlets were flash frozen in liquid nitrogen and then ground to a powder. The leaf tissue was then used for RNA extraction using the Qiagen

Plant RNeasy kit and following all instructions provided by the manufacturer (Qiagen,

Valencia, CA). The final elution volume was 40 µL, the concentration, 260/230 nm ratio and 260/280 nm ratio were determined using a Nanodrop 2000 Spectrophotometer

(Invitrogen).

The purified RNA was treated with DNase I Amplification grade (Invitrogen, Carlsbad,

CA) to eliminate any DNA present in solution. Briefly, to a total of 4-6 µg of RNA the following was added: 5 µL of 10X DNaseI Reaction Buffer (supplied by the manufacturer Invitrogen with the DNase I enzyme), 2.5 µL of DNaseI and 2.5 µL of

0.1% (v/v) dyethylpyrocarbonate (DEPC) treated water. The sample was then incubated for 15 min at room temperature and the DNaseI enzyme was inactivated by addition of

2.5 µL of EDTA and heating for 10 min at 65 ˚C. After DNaseI treatment the RNA concentration, 260/280 nm ratio and 260/230 nm ratio were determined using a

Nanodrop 2000 Spectrophotometer (Fisher Scientific).

A total of 1µg of RNA was used to synthesize cDNA using the Super Script III First-

Strand Synthesis System (Invitrogen); all instructions provided by the manufacturer were followed. Briefly in a 1.5 mL microcentrifuge tube the following was added: 1 µg of

RNA, 1µL of 50 µM oligo(dT)20, 1 µL of 10 mM dNTP mix and up to a volume of 10 µL

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with RNase free water (Invitrogen), the RNA and reagents mix was incubated at 65 ˚C for 10 min and then on ice for at least 1 min. After incubation on ice to each tube were added 10 µL of cDNA synthesis mix (2µL of 10X RT buffer (provided with the kit), 4µL of 25 mM MgCl2, 2µL of 0.1 M dithiothreitol, 1µL of RNase OUT and 1µL of

Superscript Script III enzyme), followed by incubation at 50 ˚C for 50 min The reaction was terminated by incubation at 85 ˚C for 5 min. The leaf cDNA obtained was used as a template to amplify the linamarase gene using the primers that encompass the full length linamarase gene and the 5’UTR and 3’UTR regions, the primer 5fls begins at the linamarase gene 5’UTR and contains a SmaI restriction enzyme site and the 3fls primer sequence is located in the 3’UTR and contains an SstI restriction enzyme site. The sequence of the 5fls and 3fls primers were the following,

5fls: CAG CCC GGG ACA ACT TTC TTC AGC TAT CAG

3fls: GAG CTC ACA TAT GCT AGA TCA TTG G

The amplification reaction contained the following: 100 ng of cDNA, 1 unit of Platinum

Pfx DNA Polymerase, 5µL of 10X Pfx Amplification Buffer, 0.5 µL of 50 mM MgSO4,

1.5 µL of 10 mM dNTP solution, 1.5 µL of each 10 µM primer solution (5fls and 3fls) and autoclaved double distilled water up to 25 µL. The following cycle program was used: 94 ˚C for 5 min, 32 cycles of amplification (94 ˚C for 30 sec, 59 ˚C for 45 sec and

68 ˚C for 1:30 min) and a final cycle of 68 ˚C for 5 min. The linamarase gene PCR product was gel purified. The linamarase gene and the 3D vector were digested sequentially with SmaI (Invitrogen) and SstI restriction (Invitrogen) digestion enzymes.

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The 3D vector has the pBI121 vector backbone and contains a patatin promoter: GlgC gene : nos terminator cassette. The GlgC gene was removed from the 3D vector by digesting with SmaI and SstI . The SmaI digestion of the linamarase PCR product and the

3D vector was done in 1X React 4 ® buffer (Invitrogen) at 30 ˚C for one hour (See

Section XX). After digestion the two products were purified using the Qiaquick PCR cleanup kit I (Qiagen) using the protocol provided by the manufacturer. The SstI digestion reaction was done in 1X React 2 ® buffer (Invitrogen) at 37 ˚C for one hour.

The two digested products were gel purified; from the 3D vector SmaI/SstI digestion. The band corresponding to the vector backbone was purified. A second band corresponding to the GlgC gene was visible but not purified. The linamarase gene was ligated in to the 3D vector using the T4 DNA ligase enzyme (NEB). The ligation was carried out at room temperature for one and a half hours and then at 14˚C over night.

The ligation product was used to transform subcloning efficiency DH5ɑ cells by heat shock as described by the supplier (Invitrogen). The plasmid DNA was isolated and sent for sequencing at the Ohio State Plant-Microbe Genomics Facility

(http://www.biosci.ohio-state.edu/~pmgf/) The confirmed plasmid was renamed 3D-fls

(Fig. 3.2)

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Figure 3.2 T-DNA region of the 3Dfls plasmid. The linamarase sequence including the

5’UTR and 3’UTR region were cloned in to the 3D vector. nos= nopaline synthase terminator, 35S = CaMV35S promoter and nptII= neomycin phosphotransferase.

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3.2.6 Determination of the signal peptide

The SignalP 3.0 Server (http://www.cbs.dtu.dk/services/SignalP/) was used to predict the signal peptide cleavage site of linamarase.

3.2.7 Cloning of an ΔN-ter linamarase

The following primers 5LINA2 and 3LINA2 were designed so that the signal peptide region of the linamarase gene would not be part of our amplified sequence (ΔN-ter linamarase), additionally a new start codon (ATG) was integrated in to the sequence of the 5LINA2 primer. The 5LINA2 and 3LINA2 primer sequences were the following:

5LINA2: AAT CCC GGG ATG ACT GAT GAT GAT GAT GAT ATT CCT GAC G

3LINA2: CGG GAG CTC CTA CAT CAC ATA GAA TTT GCC AAC CTT

The 3D-fls plasmid was used as a template and the amplification reaction contained the following: 50 ng of 3D-fls plasmid, 1 unit of Platinum Pfx DNA Polymerase, 5 µL of

10X Pfx Amplification Buffer, 0.5 µL of 50 mM MgSO4, 1.5 µL of 10 mM dNTP solution, 1.5 µL of each 10µM primer solution (5LINA2 and 3LINA2) and autoclaved double distilled water up to 25 µL. The amplification program was as follows: 94˚C for 5 min, 32 cycles of amplification (94 ˚C for 30 sec, 57 ˚C for 45 sec and 68 ˚C for 1:30 min) and a final cycle of 68 ˚C for 5 min. The ΔN-ter linamarase gene and the 3D vector were digested sequentially with SmaI (Invitrogen) and SstI restriction (Invitrogen) digestion enzymes. The SmaI digestion of the ΔN-ter linamarase PCR product and the 3D vector was done in 1X React 4 ® buffer (Invitrogen) at 30 ˚C for one hour. After

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digestion the two products were purified using the Qiagen PCR Cleanup kit (Qiagen) following the instructions according to the manufacturer. The SstI digestion reaction was done in 1X React 2 ® buffer (Invitrogen) at 37 ˚C for one hour. The ΔN-ter linamarase gene was ligated into the 3D vector. The ligation reaction was carried out at room temperature for one and a half hours and then at 14 ˚C over night. The ligation product was used to transform subcloning efficiency DH5ɑ cells by heat shock as described by the supplier (Invitrogen). The plasmid DNA was isolated and confirmation of correct integration and sequence of the ΔN-ter linamarase gene was carried out by sequencing at the Ohio State Plant Microbe Genomics Facility. The plasmid containing the ΔN-ter linamarase gene was renamed 3Dls (Fig. 3.3).

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Figure 3.3 T-DNA region of the 3Dls plasmid. The signal sequence targeting linamarase to the cell wall was removed. This sequence was named ΔN- ter linamarase. The ΔN-ter linamarase was cloned in to the 3D plasmid. nos= nopaline synthase terminator, 35S=

CaMV 35S promoter and nptII= neomycin phosphotransferase.

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3.2.8 Cloning of a vacuolar targeted linamarase

The vacuolar targeting domain from the barley lectin (Bednarek et al. 1991) was fused to the linamarase gene by PCR. The protein sequence of the vacuolar targeting domain is

VFAEAIAANSTLVAE after processing within the cell the vacuolar targeting domain is cleaved. The nucleotide sequence of the vacuolar targeting domain was codon optimized based on the Codon Usage Table (http://www.kazusa.or.jp/ codon/) of cassava. The DNA sequence of the vacuolar targeting domain optimized for cassava is GTT TTT GCT

GAA GCT ATT GCT GCT AAT TCA ACT CTT GTT GCT GAA. The vacuolar targeting domain was added to a 3’ end primer (3LIVAC) which contained in addition part of the 3’ end sequence of linamarase without the stop codon, a stop codon was added after the vacuolar targeting domain. The sequence of the primers used to amplify linamarase and fuse a vacuolar targeting domain were named 5LIVAC2 and 3LIVAC2, their sequences are included below.

5LIVAC2: CGG CCC GGG ATG ACT GAT GAT GAT GAT GAT AAT ATT CCT

GAC GAT TTT AGC CGT AAA

3LIVAC: CAG AGC TCC TAT TCA GCA ACA AGA TGG GAA TTA GCA GCA

ATA GCT TCA GCA AAA ACC ATC ACA T

The 3Dls plasmid was used as a template and the 5LIVAC2 and 3LIVAC primers were used to amplify the linamarase gene and fuse the vacuolar targeting domain at the 3’ end.

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The amplification reaction contained the following: 50 ng of 3Dls plasmid, 1 unit of

Platinum Pfx DNA Polymerase, 5 µL of 10X Pfx Amplification Buffer, 0.5 µL of 50 mM

MgSO4, 1.5 µL of 10 mM dNTP solution, 1.5 µL of each 10µM primer solution

(5LIVAC2 and 3LIVAC) and autoclaved double distilled water up to 25 µL. The amplification program had the following cycles: 94 ˚C for 5 min, 32 cycles of amplification (94 ˚C for 30 sec, 62 ˚C for 45 sec and 68 ˚C for 1:30 min) and a final cycle of 68 ˚C for 5min. The vacuolar linamarase PCR product was gel purified. The vacuolar linamarase PCR product and the 3D vector were digested sequentially with SstI

(Invitrogen) and SmaI (Invitrogen). The SmaI digestion of the vacuolar linamarase PCR product and the 3D vector was done in 1X React 4 ® buffer (Invitrogen) at 30 ˚C for one hour. After digestion the vacuolar linamarase and 3D vector were purified using the

Qiaquick PCR Cleanup kit according the manufacturer’s instructions (Qiagen). The SstI digestion reaction was done in 1X React 2 ® buffer (Invitrogen) at 37 ˚C for one hour.

The linamarase gene was ligated into the 3D vector using the T4 DNA ligase enzyme

(NEB). The ligation was carried out at room temperature for one and a half hours and then at 14˚C over night. The ligation product was used to transform subcloning efficiency

DH5ɑ cells by heatshock as described by the supplier (Invitrogen). The plasmid DNA was isolated and confirmation of correct integration and sequence of the ΔN-ter linamarase gene was carried out by sequencing at the Ohio State Plant Microbe

Genomics Facility. The plasmid containing the vacuolar targeted linamarase gene was renamed 3DVL (Fig. 3.4).

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Figure 3.4 T-DNA region of the 3DVL plasmid. The vacuolar targeting domain (VTD) was added through PCR to the linamarase gene. nos= nopaline synthase terminator, 35S=

CaMV 35S promoter and nptII= neomycin phosphotransferase.

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3.2.9 Cloning of the pCambia2300-VL and pCambia2300-SVL vectors

The cloning of the pCambia2300-VL and pCambia2300-SVL was done by our collaborators at the ILTAB lab from the Donald Danforth Plant Science Center.

Briefly, the patatin promoter was cloned from the 3DVL vector in to the PstI/SacI site of the pMON999 shuttle vector. Next the vacuolar linamarase gene was cloned into the same vector at the SacI/KpnI sites, the new vector was renamed pMON-VacLam. The patatin promoter:vacuolar linamarase:nos terminator cassette was digested out of pMON-

VacLam with NotI and SmaI. The NotI end was blunt ended with Mung Bean

Exonuclease (New England Biolabs, Ipswich, MA) and ligated into a pCambia2300 vector digested with SmaI, this vector was renamed pCambia2300-VL. The patatin promoter:sporazein:nos terminator cassette was digested out of pMON-Sporazein

(property of ILTAB) with NotI and SmaI, the NotI end was made blunt by action of the

Mung Bean Exonuclease (NEB) and cloned in to the SmaI site of the pCambia2300-VL vector; the double construct vector was renamed pCambia2300-SVL.

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3.2.10 Generation of Friable Embryogenic Callus

Kuniyuki media (1984) supplemented with 20 g/L sucrose, 50 µM picloram and 2.4 g/L of phytagel, this media was named DKW2. The apical leaves remained in DKW2 media for 4 weeks at the following environmental conditions: temperature of 28 ˚C, a reduced light intensity of 20 to 35 µmol photons m-2 s-2 and a 16 hr day /8 hr night cycle. After 4 weeks in DKW2 media the visible organized embryogenic structures (OES) were removed and placed in fresh DKW2 media. The OES remained in the second DKW2 media for 3 weeks at the same environmental conditions (temperature of 28 ˚C, a reduced light intensity of 20 to 35 µmol photons m-2 s-2 and a 16 hr day/8 hr night cycle ) After three weeks the OES were placed in Gresshoff and Doy media (1974) supplemented with

20 g/L sucrose, 50 µM picloram and 2.4 g/L phytagel, this media was named GD2P50.

After 3 weeks in GD2P50 the OES may form friable embryogenic callus (FEC) structures, these were removed and placed in fresh GD2P50 for 3 more weeks. Finally the growing FEC tissues were removed from the second GD2P50 and proliferated for one more round of three weeks in GD2P50. All three growth cycles in GD2P50 media were done at the following environmental conditions: temperature of 28 ˚C, a reduced light

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intensity of 20 to 35 µmol photons m-2 s-2 and a 16 hr day/8 hr night cycle. After the third round of proliferation in GD2P50 media the FEC were used in transformation.

3.2.11 Transformation of cassava

After inoculation the FECs were placed in GD media (Gresshoff and Doy, 1974) containing 50 µM picloram, 500 mg/L carbenicillin to eliminate any remaining

Agrobacterium and 27.5 µM paramomycin for antibiotic selection during 28 days. After this period of time the FEC were transferred to Stage 1 regeneration media which consisted of MS2 media supplemented with 5 µM NAA, 250 mg/L carbenicillin and 45

µM paramomycin. After 21 days growing tissues were transferred to Stage 2 regeneration media (MS2 medium plus 0.5 µM NAA and 45 µM paramomycin). The preferred tissues transferred from Stage 1 to Stage 2 media were FEC clusters showing early torpedo stage or beginning cotyledon stage embryos. At 21 days of growth in Stage 2 media the embryos that had developed in to a mature cotyledon stage were placed in the germination stage media (MS2 medium supplemented with 2µM BAP and 45 µM paramomycin). Embryos that germinated were micropropagated into MS2 media, each germinated embryo is considered a potential individual transgenic event. The transgenic lines were grown and maintained in MS2 media. The lines transformed with 3Dls were

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named ΔN-linamarase lines, those transformed with pCambia2300-VL were named VL lines, and those transformed with pCambia2300-SVL were named SVL lines.

3.2.12 DNA extraction and PCR of putative transgenic ΔN-linamarase cassava lines

The genomic DNA from putative ΔN-linamarase lines was used as a template for amplification of a 1700 bp region including the linamarase gene and flanking regions of the patatin promoter and nos terminator. The designed primers annealed at the patatin promoter (PATCHECK) and nos terminator (NOSCHECK)

PATCHECK: TTT CTC AAC TTG TTT ACG TGC C

NOSTCHECK: CCG GCA ACA GGA TTC AAT CTT AAG

The following conditions were used in the PCR: one cycle of 5 min at 94ºC followed by

40 cycles of amplification (30 s at 94 ºC, 60 s at 52 ˚C and 60 s at 72 ºC) and a final cycle of 5 min at 72 ºC. The ChoiceBlue Taq DNA Polymerase (Denville Scientific, Metuchen,

NJ) was used in all reactions, each reaction contained: 400 -1000 ng of genomic DNA as template, 1 unit of ChoiceBlue Taq DNA Polymerase, 2.5 µL of 10X Choice PCR

Buffer, 2.5 µL of each 10 µM primer solution and autoclaved double distilled water up to a volume of 25 µL. The products of each PCR reaction were gel purified and confirmed by sequencing.

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3.2.13 DNA extraction and PCR of putative transgenic VL and SVL cassava lines

The genomic DNA from putative VL and SVL lines was used as a template for amplification of a 900 bp region of the vacuolar targeted linamarase gene, primers used where MID5L and VACCHECK. The MID5L primer sequence is located in the middle of the linamarase gene and the VACCHECK primer is located at the end of the vacuolar targeting domain.

MID5L: CTC CGA CAG TAA AGT TGA TGT CGA ACG AG

VACCHECK: GCA GCA ATA GCT TCA GCA AAA ACC ATC ACA

The following conditions were used in the PCR: one cycle of 5 min at 94ºC followed by

40 cycles of amplification (30 s at 94ºC, 60 s at 57 ˚C and 60 s at 72 ºC) and a final cycle of 5 min at 72 ºC. The ChoiceBlue Taq DNA Polymerase (Denville Scientific, Metuchen,

NJ) was used in all reactions, each reaction contained: 400 -1000 ng of genomic DNA as template, 1 unit of ChoiceBlue Taq DNA Polymerase, 2.5 µL of 10X Choice PCR

Buffer, 2.5 µL of each 10 µM primer solution and autoclaved double distilled water up to a volume of 25 µL. The products of each PCR reaction were gel purified and confirmed by sequencing.

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3.2.14 Dot blot analysis of putative VL and SVL transgenic cassava plants

The genomic DNA solutions from VL and SVL(obtained in Section X DNA extraction and PCR) were diluted to a concentration of 20 ng/µL. The samples for the dot blots consisted of three 100 ng aliquots of genomic DNA from each putative transgenic line and from four control lines. The control lines have a known copy number of the

CaMV35S promoter from the 35S promoter: nptII: nos terminator cassette present in all

T-DNA as the selection marker. The copy number of the control lines was previously determined by Southern blot. The lines had between 3 to 0 copy numbers.

The dot blot protocol proceeded as presented in Section 2.2.12 of Chapter 2.

3.2.15 Harvest weight

Cassava plants were harvested after 4 months of greenhouse growth and their fresh weight was recorded. The shoot and the storage root tissues were weighed separately.

3.2.16 RNA extraction and cDNA synthesis from cassava storage roots

The extraction of RNA from storage root pulp was done using a LiCl extraction method based on the methodology developed by Manickavelu et al. (2007). A sample of 50-100 mg of storage root pulp dry tissue was ground inside a microfuge tube using a mini plastic pestle (Fisher Scientific). The RNA extraction proceeded following the protocol outlined in Section 2.2.13 RNA extraction on Chapter 2)

For all RNA samples the concentration, 260/280 nm ratio and 260/230 nm ratio of each sample were determined using a NanoDrop 2000 Spectrophotometer (Fisher Scientific).

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The cDNA for all samples was synthesized using 1µg of RNA, 4 µL of Quantas qScript cDNA SuperMix (Quanta Biosciences, Gaithersburg, MD) and RNase-free water

(Promega) up to a volume of 20 µL. The reaction was run using the following program: 5 min at 25 ˚C, 30 min at 42 ˚C, 5 min at 85˚C and hold at 4 C˚. The cDNA obtained was stored at -20 ˚C.

3.2.17 Semiquantitative RT-PCR analysis of vacuolar linamarase and sporazein expression

The RT-PCR reaction was carried out using the first-strand cDNAs as template. Gene specific primers for vacuolar linamarase (MID5L and VACCHECK, sequences listed in

Section 3.2.13 of Chapter 3) and Sporazein (SpoF/SpoR, see below for sequences) were used. As a control for the cDNA reaction and the overall quality of the RNA extraction a reaction for the cassava tubulin gene (5’TUB and 3’TUB, see below for sequences) was prepared and ran at the same time.

SpoF: TAACTTAATATCTAGAATGAAAGCCTTCACACTCGCTCTCTTC

SpoR: AATATTCAATTGGTACCACACATCGGTAGGTTTGATGACAAAAACG

5’TUB: GAT CCT ACT GGG AAG TAC ATT GG

3’TUB: CTG CAT TCT CCA CCA ACT GA

The following conditions were used in the RT-PCR reaction: one cycle of 5 min at 94º C followed by 32 cycles of amplification (30 s at 94º C, 30 s at annealing temperature and

45 s at 68ºC) and a final cycle of 5 min at 68º C. The annealing temperature for the

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vacuolar linamarase primers was 57º C, for the Sporazein primer pair it was 58º C and for the tubulin primer pair it was 55º C. The Platinum Pfx DNA Polymerase enzyme

(Invitrogen) was used in all reactions as follows: 100 ng of cDNA, 1 unit of Platinum Pfx

DNA Polymerase, 5 µL of 10X Pfx Amplification Buffer, 0.5 µL of 50 mM MgSO4, 1.5

µL of 10 mM dNTP solution, 1.5 µL of each 10µM primer solution and autoclaved double distilled water up to 25 µL. The amplification products were gel purified and confirmed by DNA sequencing at the Ohio State Plant Microbe and Genomics facility

(http://www.biosci.ohio-state.edu/~pmgf/).

3.2.18 Linamarin determination

Linamarin levels of leaves and storage root pulp were determined by GC-MS using the internal standard method in a similar manner as previously described by Siritunga et al.

Technologies). The GC-MS was operated at a pressure control mode at a flow of 1 mL/min. The GC-MS oven program was as follows, 50 ˚C for 1 min, ramp at 30 ˚C/min to 185˚C, ramp at 6 ˚C/min to 230 ˚C, ramp at 12 ˚C/min to 280 ˚C and 3 min at 290 ˚C for column cleaning.

3.2.18.1 Standard curve

The standard curve obtained in Chapter 2 was used for the analysis of the linamarin data from all transgenic lines.

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3.2.18.2 Tissue sample extraction

A total of 5 mg and 25 mg of dry weight were taken from leaf and storage root pulp respectively of 4 month old greenhouse grown wild-type and transgenic cassava . A volume of 600 µL of acetonitrile and 2.5 µL of the internal standard phenyl-β- glucopyranoside (PGP) stock solution (2µg/µL) was added to each sample and this was followed by a 40 min extraction in a vortex set at 1,240 RCF. The samples were then centrifuged at 16,800 RCF for 10 min. The supernatant was removed and stored in a new

2 mL microcentrifuge tube. The remaining pellet was re-extracted with 600 µL of acetonitrile for 40 min in a vortex set at 1,240 rpm. Samples were centrifuged once more at 16,800 RCF and the supernatant was combined with the previous one. The total extract obtained from root pulp samples was completely dried in a Centrivap speedvac. An aliquot of 100 µL was taken from the total extracts for leaf and dried completely in a

Centrivap speedvac. After drying the pellet was resuspended in 150 µL of water, this was followed by addition of a volume of 150 µL of chloroform to remove lipids and other contaminants, the samples were vortexed at high speed for 10 seconds and centrifuged at

16,800 RCF for 10 min. The aqueous phase was transferred to a new 2 mL microcentrifuge tube and dried completely in a Centrivap speedvac, after drying the remaining pellet was resuspended in 50 µL of acetonitrile.

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3.2.18.3 Derivatization of linamarin and internal standard

A volume of 50 µL of N-Methyl-N-(trimethylsilyl) trifluoroacetamide with 1% trimethylchlorosilane (MSTFA + 1% TMCS) and 10 µL of extra dry pyridine were added to all tissue samples and they were incubated at 65 ˚C for 30 min on a dry heating block.

Samples were run immediately after derivatization.

3.2.19 Free amino acid extraction and analysis

The protocol followed in Section 2.2.17 of Chapter 2 was used for free amino acid extraction from leaf and storage root pulp of wild-type and transgenic VL and SVL cassava.

3.2.20 Total protein analysis

Analysis of total protein was done on a dry weight basis. The protocol followed in

Section 2.2.18 of Chapter 2 was used for crude protein quantification from leaf and storage root pulp of wild-type and transgenic VL and SVL cassava.

3.2.21 Western blot

Protein was extracted from 2-3 g fresh weight of storage root pulp by grinding the tissue in liquid nitrogen in a mortar and pestle. A volume of 5 mL of MOPS buffer (100 mM

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then centrifuged at 3400 RCF and the supernatant was collected in a fresh 15 mL falcon tube. The samples were kept at 4 ˚C during all stages of the protein extraction. The proteins contained in the final extract were precipitated by the addition of acetone using a similar protocol to the one detailed in the PIERCE Technical Resource: Acetone precipitation of proteins. The procedure was done as follows: A volume of 7-10 mL of acetone (Fisher Scientific, Pittsburgh, PA) was added to the protein extract and it was left overnight at -20˚C to precipitate the extracted proteins. The next day the samples were centrifuged for 10 min at 3800 RCF. The supernatant was discarded and the pellet was resuspended in 200 µL of 100 mM sodium phosphate buffer pH 7.5 with mini cOmplete protease inhibitor cocktail (Roche, 1 tablet for every 10mL of buffer). Protein concentration was determined using the CB-X kit following all guidelines provided by the supplier.

An amount of 20 to 50 µg of protein was denatured by adding 15 – 25 µL of loading dye

(0.06 M Tris-HCl, pH 6.8, 10% (v/v) glycerol, 2% (w/v) SDS, 5% (v/v) 2- mercaptoethanol, 0.0025 % (w/v) bromophenol blue) and incubating at 95 ˚C for 10 min and then placing on ice for 2min. After denaturing the samples were ready to run. A Bio-

Rad pre-cast ready gel of 7.5% polyacrylamide ( Bio-Rad Catalog # 116-1154) was used to run the samples in an Mini Protean electrophoresis apparatus (Bio-Rad) using 1X running buffer (30 mM Tris base, 200 mM glycine and 0.1% (w/v) of SDS per liter, pH=8.3). The gel was run at 25 mAmps until completion. Three Whatman filter paper pieces previously wetted with transfer buffer (20 mM Tris-Cl, 192 mM glycine, 20%

(v/v) methanol, 0.01% (w/v) SDS) were placed one on top of the other inside a dry blot transfer apparatus, a top the last Whatman filter the protein blot was carefully placed

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followed by the Immobilon nylon membrane and three more Whatman filter paper layers, all previously wetted with transfer buffer. The dry blot transfer apparatus was run at a constant current of 1.9-2.5 mA/cm2 of gel area for 4 hours.

After the transfer the nylon membrane was placed in blocking buffer (20 mM Tris-HCl pH 7.5, 500 mM NaCl, 1% (w/v) bovine serum albumin) for 1 hour. Then the membrane was incubated overnight at 4˚C with either linamarase rabbit antibody (10 µg/µL )

(Agrisera, Vannas, Sweden) or β-zein rabbit antibody (10 µg/µL) (Genscript, Piscataway,

NJ) at a 1:10,000 dilution in Antibody Buffer (20mM Tris-Cl pH 7.5, 500 mM NaCl,

0.05% Tween-20, 1% (w/v) bovine serum albumin). The following day the blot was washed three times for 15 min each in TTBS buffer (20 mM Tris-Cl pH 7.5, 500 mM

NaCl, 0.05 % (v/v) Tween-20) and then incubated with the secondary antibody (10

µg/µL of anti-rabbit horseradish peroxidase) at 1:10,000 in Antibody Buffer. The western blot was developed using chemiluminescence using luminol in a protocol similar to that developed by Yakunin and Hallenbeck (1998). The detection solution was prepared in the dark with the following reagents: 7 mL of Solution A (100 mM glycine-NaOH buffer pH=9.6), 7 mL of Solution B (30% v/v H2O2 in water), 1 mL of luminol reagent (45 mM luminol in dimethyl sulfoxide) and 1 mL of 4-iodophenol reagent (750 mM 4-iodophenol in dimethyl sulfoxide). The western blot membrane was placed in the detection solution for 1 min then washed in TBS buffer (20 mM Tris-HCl, 500 mM NaCl, pH = 7.5) for 5 seconds and placed in a plastic developing envelope. The membrane was exposed to X- ray films for 30 seconds to 5 min and developed in a Kodak 2000A X-Ray developer

(Kodak).

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3.2.22 Total nitrogen content

The total moles of nitrogen contributed by protein, free amino acids and linamarin were calculated. The total moles of nitrogen from linamarin were equal to the number of moles of linamarin present as only one atom of nitrogen is present in linamarin.

For the protein initially the total grams of nitrogen were calculated. It is generally accepted that 16.5% of the weight of a protein is nitrogen, however, there is a small variation for cassava storage root proteins which contain 18.5% nitrogen (based on data from Yeoh and Truong, 1996). The leaf nitrogen derived from protein was considered as

16.5% of the protein weight while that for the roots was considered to be 18.5%. The grams of nitrogen were then converted to moles.

For the total nitrogen contributed by amino acids the individual moles of each amino acid and the quantity of nitrogen moles per mole of amino acid present were taken in to account to calculate the total nitrogen contribution from all amino acids.

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3.3 RESULTS

3.3.1 Cloning of a ΔN-ter linamarase and a vacuolar linamarase

The complete linamarase cDNA, including its 5’ and 3’UTR, was cloned into the 3D vector (Fig. XA), this vector was renamed 3Dfls. The linamarase enzyme is targeted normally to the cell wall. To identify the signal peptide responsible for the periplasmic targeting the linamarase protein sequence was analyzed using the SignalP 3.0 Server in its Eukaryote modality (http://www.cbs.dtu.dk/services/SignalP/) (Fig. X). Three scores were obtained, the first is the S-score; this predicts the signal peptide and the values obtained for each amino acid will be high if the amino acid is part of the signal peptide.

The S-score was high for amino acids 1 through 17 in the linamarase protein. (Fig. X, green line). The second score given is the C-score or cleavage site score. This will give the position where the signal peptidase cleaves. In the case of linamarase, it is between amino acids 17 and 18, with 17 being part of the signal peptide and 18 being the first amino acid of the mature protein (Fig. X, red line). In the case of some protein sequences multiple cleaving sites may be predicted and the Y-score may then be used to determine the most likely cleaving site. The Y-score is high where the slope of the S-score is steep and the C-score is high. In the case of linamarase only one cleavage site was predicted and all three scores converged on amino acid 17 as the end of the signal peptide. An ΔN- terminal linamarase without the 17 amino acid signal peptide was cloned in to the 3D vector which was renamed 3Dls.

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The C-terminal vacuolar targeting domain of a barley lectin is composed of fifteen amino acids (VFAEAIAANSTLVAE) and it has been shown to be sufficient to target a protein to the vacuole (Bednarek et al. 1991). The vacuolar targeting domain was fused in frame to the C-terminal region of the linamarase gene by using a 3’end primer containing the full length vacuolar targeting domain. The gene was named vacuolar linamarase and it was cloned in to the 3D vector which was renamed 3DVL (Fig. XC).

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Figure 3.5 Cloning of a ΔN-ter linamarase and a vacuolar targeted linamarase. (A) 3Dfls vector. This construct contained the full cDNA sequence of linamarase. (B) 3Dls vector.

The N-terminal signal peptide responsible for the targeting of linamarase to the cell wall was removed from the nucleotide sequence. This construct is designated ΔN-linamarase.

(C) 3DVL. A vacuolar targeting domain was added to the C-terminal of the linamarase gene. VTD = vacuolar targeting domain, nptII = neomycin phosphotransferase, 35S =

CaMV 35S promoter, nos = nopaline synthase terminator

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Figure 3.6 Prediction of the signal peptide of linamarase. The protein sequence of linamarase was analyzed using the SignalP 3.0 prediction software

(http://www.cbs.dtu.dk/services/SignalP/) to determine the location of the signal peptide that targets linamarase to the cell wall.

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3.3.2 Cloning of the pCambia2300-VL and pCambia2300-SVL vectors

The patatin promoter: vacuolar linamarase gene: nos terminator cassette was cloned into the pCambia2300 vector to originate the pCambia2300-VL (Fig. 3.7). The patatin promoter: sporazein: nos terminator cassette was cloned in to pCambia2300-VL to originate the double construct pCambia2300-SVL (Fig. 3.8)

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Figure 3.7 T-DNA region of the pCambia2300-VL vector. This vector was constructed by cloning the patatin promoter: vacuolar

linamarase:nos terminator cassette in to pCambia2300. VTD= vacuolar targeting domain from barley lectin, NPTII=

neomicynphosphotransferase, nos= nopaline synthase terminator, 35S= CaMV 35S Promoter, VTD= vacuolar targeting domain.

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Figure 3.8 T-DNA region of the pCambia2300-SVL vector. This vector was constructed by cloning the patatin promoter: vacuolar

linamarase:nos terminator cassette in to pCambia2300. VTD= vacuolar targeting domain from barley lectin, NPTII=

neomicynphosphotransferase, nos= nopaline synthase terminator, 35S= CaMV 35S Promoter, VTD= vacuolar targeting domain.

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3.3.4 Generation of transgenic cassava expressing ΔN-ter linamarase

It was hypothesized that the expression of a ΔN-ter linamarase in cassava storage root cell cytoplasm would putatively increase the hydrolysis of linamarin in vivo and through the assimilation of the released cyanide increase the free amino acid and protein content in the storage roots. Transgenic cassava was generated through Agrobacterium mediated transformation of FECs using the 3Dls vector. A total of 15 independent lines were regenerated after transformation. Only six of the lines were proved to be transformants by

PCR verification of the presence of the ΔN-linamarase gene (Fig. 3.9). However, the transcript of ΔN-ter linamarase was not detected in any of the six lines; no lines confirmed by PCR and RT-PCR were recovered from transformation of cassava with the

3Dls vector. These results suggested that cytoplasmic targeting of linamarase was toxic.

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Figure 3.9 PCR analysis of putative transgenic cassava lines transformed with the 3Dls vector. Select PCR positive lines are shown (1) Line 3Dls8a (2) Line 3Dls29c (3) Line

3Dls32 (+) amplification using the 3Dls plasmid as a template.

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3.3.5 Generation of transgenic cassava VL and SVL lines

It was hypothesized that by increasing hydrolysis of linamarin in vivo and its subsequent catabolism reactions through the CAS assimilation pathway the free amino acid and protein content would be increased. A vacuolar targeted linamarase was expressed in the linamarin storage organelle, the vacuole, to increase linamarin hydrolysis in vivo. The generated putative transgenic lines were named VL. In addition, the novel storage protein sporazein was co-expressed with the vacuolar linamarase with the objective of increasing the protein sink in the cassava storage roots. The generated putative transgenic lines with the double construct of vacuolar linamarase and sporazein were named SVL.

Transgenic cassava was generated by Agrobacterium mediated transformation of FEC tissues using either the pCambia2300-VL or the pCambia2300-SVL vectors. The T-DNA transfer cassette includes a constitutively expressed nptII gene for antibiotic selection.

The FEC clusters were selectively grown under paramomycin selection and allowed to mature and regenerate into plantlets.

After transformation with the pCambia2300-VL construct a total of 38 VL putative transgenic plants were regenerated. After transformation with the pCambia2300-SVL double construct a total of 49 putative transgenic lines were regenerated. All the putative

VL and SVL transgenic lines were analyzed by dot-blot to determine the number of T-

DNA insertions in the genomic DNA (Fig. 3.10) and by PCR to confirm they were transgenic.

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Figure 3.10 Dot blot analysis of SVL and VL lines. A standard curve of 0 to 3 T-DNA insertions was required to analyze the number of insertions in each putative transgenic line. (a) VL null line (b) SVL-05 with 5 insertions (c) SVL-06 with 1 insertion (d) VL-05 with 1 insertion.

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From the 49 regenerated SVL antibiotic resistant lines, 12% were null, and 41% had 3 T-

DNA insertions or more. From the 38 regenerated VL putative transgenics 5.5% were null lines and 28% had 3 or more T-DNA insertions in the genome. A total of 15 lines of

VL and SVL with varying number of insertions were selected for further analysis and grown in the greenhouse. The copy number distribution of the total number of plants and those selected for greenhouse growth can be found on Table 3.1.

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Table 3.1 T-DNA insertions of VL and SVL lines. The total number of regenerated VL and SVL lines is presented distributed by the number of T-DNA insertions. The number of plants for the number of T-DNA insertions that were grown in the greenhouse is indicated

VL SVL Number of Total Greenhouse Total Greenhouse T-DNA insertions 0 2 -- 6 -- 1 17 10 14 7 2 11 3 11 5 3 2 2 3 2 4 8 -- 11 -- 5 -- -- 6 1 Total 38 15 49 15

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3.3.6 Greenhouse growth of wild-type, VL and SVL lines

The future acceptance of any developed transgenic cassava lines will inevitably depend on the improvement of a given trait while inherent traits such as yield and starch content should remain unchanged (Taylor et al., 2004). It was therefore important to evaluate the growth of VL and SVL transgenic lines in relation to wild type. A total of five plants from each of 15 VL and 15 SVL lines along with five wild-type plants were grown for two months in the greenhouse in 4 inch pots. At two months the majority of the transgenic plants had not developed storage roots (personal evaluation). The plants were maintained in the small volume pots (4 in pots) to promote storage root formation. At 4 months the majority of the VL and SVL transgenic lines had very poor tuber development (Fig. 3.11) in some cases no plants out of the five planted for a line had produced tubers.

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Figure 3.11 Roots from three different transgenic lines and wild-type at 4 months of age. The two VL and the SVL line presented here

had very poor storage root development. The majority of the VL and SVL lines had poor storage root development in comparison to

wild type at 4 months of greenhouse growth under the same conditions.

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At 4 months of greenhouse growth only three lines of VL and three lines of SVL produced storage roots (Fig. 3.12). The three VL lines were VL-04, VL-05 and VL-15; the three SVL lines were SVL-06, SVL-09 and SVL-11. These six transgenic lines were chosen for further analysis. Out of these six transgenic lines VL-04, VL-05, SVL-06,

SVL-09 and SVL-11 are single copy lines and VL-15 is a double copy line.

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Figure 3.12 Storage root images from wild-type, VL-04, 05 and 15 and SVL-06, 09 and 11. At four months of greenhouse growth the

VL lines 04, 05 and 15 and the SVL lines 06, 09 and 11 were the only six transgenic lines that produced storage roots in at least three

independent plants.

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The storage root weight of the VL lines was reduced by up to 64% compared to wild type while the SVL lines were reduced by up to 75%. (Fig. 3.13). The shoot fresh weight of the VL transgenic lines was significantly reduced by up to 28% in VL-04 and VL-05 in comparison to wild type. In SVL lines the shoot fresh weight was reduced significantly in

SVL-06 and SVL-09 by up to 20% compared to wild type. (Fig. 3.14). No significant differences were observed in storage root or shoot fresh weight between the single copy lines VL-04 and VL-05 and the double copy line VL-15.

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Figure 3.13 Storage root fresh weight. The fresh weight of the storage roots from wild- type and VL and SVL transgenic lines was measured at harvest time after 4 months of greenhouse growth. All three VL and SVL lines were significantly reduced in comparison to wild type in their storage root fresh weight. (n=3, error bars represent the standard error, asterisk represents statistical significance difference based on a p-value <

0.05 and an ** indicates p-value < 0.01)

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Figure 3.14 Shoot fresh weight. The fresh weight of the shoot from the different VL and

SVL lines as well as wild type was measured at harvest time after 4 months of growth in the greenhouse. Two VL lines (VL-04 and VL-05) as well as two SVL lines (SVL-06 and

SVL-09) were significantly reduced in comparison to wild type in their shoot fresh weight.(n=3, error bars represent the standard error, asterisk represents statistical significance difference based on a p-value < 0.05)

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3.3.7 Analysis by semiquantitative RT-PCR of vacuolar linamarase and sporazein expression.

The abundance of the vacuolar linamarase and sporazein gene transcript was analyzed in the storage root pulp of the three VL and three SVL lines and wild type at 4 months of greenhouse growth. A semiquantitative RT-PCR analysis amplified the vacuolar linamarase gene in all VL but not wild type (Fig. 3.15A), and amplified sporazein transcript as well as vacuolar linamarase transcript in SVL lines (Fig. 3.15B).

Amplification of tubulin was used as a control. Interestingly, through semiquantitative

PCR the double copy VL-15 had lower apparent expression levels than the single copy

VL-04 and VL-05 lines. In fact the VL-05 lines had higher apparent expression than VL-

15.

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Figure 3.15 Semiquantitative RT-PCR analysis of VL and SVL lines. (A)

Semiquantitative RT-PCR analysis of VL lines for amplification of the vacuolar linamarase. (B) Semiquantitative RT-PCR analysis of SVL lines for amplification of the vacuolar linamarase and sporazein. The positive control in (A) was the pCambia2300 VL plasmid and in (B) it was the pCambia2300 SVL

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3.3.8 Western blot

Expression of the vacuolar linamarase and the sporazein storage protein was confirmed in

4 month old storage roots by western blot analysis. As expected, the sporazein band was only present in the SVL lines (Fig. 3.16), however, the band corresponding to linamarase was present in both VL and SVL samples (Fig. 3.16). It is worth mentioning that the linamarase antibody used was not specific for vacuolar linamarase and so it was expected to detect a band on the wild-type lane as well, however, the band was not detected in wild-type; in fact the modified linamarase protein was detected in the VL and SVL lines only after loading 50 µg of protein. This could be due to various factors such as low content of linamarase in wild type, another is the presence of linamarase in the apoplast; the crude protein preparation protocol may have not been conducive to adequate linamarase extraction.

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Figure 3.16 Western blot analysis of VL and SVL lines. (A) 60444, (B) VL-04, (C) VL-

05 (D) VL-15 (E) SVL-06 (F) SVL-09 and (G) SVL-11. 50µg of crude protein from storage roots was loaded in each lane.

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3.3.9 Linamarin content analysis of storage root and leaf

The root-specific expression of a vacuolar targeted linamarase in VL and SVL lines would potentially increase the hydrolysis of linamarin in vivo and the assimilation of cyanide via the CAS pathway. Putatively this would not only result in an increase in the free amino acid content of the storage root but also on a decrease of the linamarin content in this organ. To determine the effect of the expression of a vacuolar linamarase on the overall linamarin accumulation of the storage roots the linamarin content was evaluated by Gas-Chromatography Mass-Spectroscopy. In addition because the linamarin present in roots is transported from the leaves (Bediako et al., 1981; Andersen et al., 2000, Siritunga and Sayre, 2003) the linamarin content was measured in leaves too (discussed below).

Linamarin content was quantified through GC-MS analysis using the internal standard method with phenyl β-glucopyranoside (PGP). The GC-MS analysis was done using single ion monitoring. The select ion for linamarin was 204.1 while for the internal standard it was 361.1. An example of a single ion chromatogram for a wild-type storage root sample is presented for both linamarin (Fig. 3.17) and the internal standard PGP

(Fig. 3.18). Similarly an example of a single ion chromatogram for a VL-04 storage root sample for both linamarin (Fig. 3.19) and PGP (Fig. 3.20).

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Figure 3.17 Single ion chromatogram for the linamarin monitored ion 204.1 from a sample from wild-type storage root; linamarin

was previously identified as the peak eluting at 12.68 min. The area of the peak as calculated by the Agilent software is shown by the

peak. There were additional peaks present in the chromatogram, with two with more abundance at 15.15 min and 18.04 min; these

additional peaks did not interfere with the measurement of the linamarin peak.

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Figure 3.18 Single ion chromatogram for the β-phenyl glucopyranoside monitored ion 361.1 from a sample from wild-type storage

root; β-phenyl glucopyranoside was previously identified as the peak eluting at 15.14 min. The area of the peak as calculated by the

Agilent software is shown by the peak.

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Figure 3.19 Single ion chromatogram for the linamarin monitored ion 204.1 from a sample of VL-04 storage root; linamarin was

previously identified as the peak eluting at 12.69 min. The area of the peak as calculated by the Agilent software is shown by the peak.

There were additional peaks present in the chromatogram, with two with more abundance at 15.14 min and 18.04 min; these

additional peaks did not interfere with the measurement of the linamarin peak.

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Figure 3.20 Single ion chromatogram for the β-phenyl glucopyranoside monitored ion 361.1 from a sample from VL-04 storage root;

β-phenyl glucopyranoside was previously identified as the peak eluting at 15.14 min. The area of the peak as calculated by the Agilent

software is shown by the peak.

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The wild-type storage root pulp had an average of 1.03 µmol linamarin/g dry weight, previous reports have seen linamarin accumulations between 0.27 to 5.2 µmol linamarin / g dry weight in cassava storage root pulp (Mkpong et al., 1990; Santana et al., 2002;

Siritunga and Sayre, 2003; Montagnac et al., 2009) with the lower cyanogenic or “sweet” varieties ranging from 0.27 to 2.96 µmol linamarin /g dry weight (Mkpong et al., 1990;

Santana et al., 2002). The linamarin concentrations in storage root pulp of the wild-type

(60444) variety used in these experiments are within the range for lower cyanide or

“sweet” cassava varieties. The storage root pulp linamarin content was significantly reduced in all three VL and three SVL lines (Fig. 3.21). The VL lines had 32 to 44 % lower linamarin concentrations than wild type while the SVL lines had a 22 to 45% reduction. The reduction of linamarin in the storage root of VL and SVL lines is likely linked to the increased catabolism of linamarin through the expression of a vacuolar linamarase. The linamarin reduction in roots observed in VL and SVL lines, however, were less than those achieved by inhibiting linamarin synthesis in leaves (Siritunga et al,

2003).

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Figure 3.21 Storage root pulp Linamarin from wild-type and the VL and SVL transgenic lines. The storage root pulp linamarin content was significantly reduced in all VL and

SVL transgenic lines. (n=3, error bars represent the standard error, asterisk represents stastical significant difference based on a p-value < 0.05 and an ** indicates p-value <

0.01)

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The leaf linamarin content of wild-type was 23.6 µmol linamarin/g dry weight, this is well within the observed values for linamarin content in the literature. Leaf linamarin has been reported in the ranges of 15 to 80 µmol linamarin/g dry weight (Mkpong et al.,

1990; Santana et al., 2002; Siritunga and Sayre, 2003; Montagnac et al., 2009). The VL lines had 36-45% lower linamarin content relative to wild-type leaves and the SVL-09 and SVL-11 lines linamarin levels were both reduced by 25% relative to wild type (Fig.

3.22). The linamarin reductions observed in VL and SVL lines is less than that reported by Siritunga and Sayre (2003), where the linamarin content in leaves was reduced 60 to

94% relative to wild type when linamarin synthesis was reduced in a leaf specific manner.

In both VL and SVL lines there was a generalized decrease of linamarin content in leaves and storage root. No significant difference in the reduction of linamarin was observed between the VL and the SVL lines.

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Figure 3.22 Leaf linamarin of wild-type and VL and SVL transgenic lines. There was a significant decrease in all three VL lines and in SVL-09 and SVL-11. (n=3, error bars represent the standard error, asterisk represents statistical significant difference based on a p-value < 0.05 and an ** indicates a p-value < 0.01)

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3.3.10 Total free amino acid content in leaf and storage roots

The total free amino acid levels of the cassava storage root pulp was measured to determine the effect of expression of a vacuolar linamarase and sporazein on the total accumulation of free amino acids. It was hypothesized that through the CAS pathway the cyanide produced from linamarin hydrolysis and acetone cyanohydrin decomposition would be assimilated to produce aspartate and ammonia. If that is the case the total free amino acid pool would be expected to increase. In addition, the expression of sporazein as a protein sink could potentially have an effect on the accumulation of free amino acids.

The total free amino acid levels were measured in the pulp of the storage root. All three

VL and all three SVL lines had significant increases in their free amino acid pool in comparison to wild-type. The VL lines had a 1.65 to 2.25 fold increase in total free amino acids in relation to wild-type and the SVL lines had a 1.91 to 2.4 fold increase in relation to wild-type (Fig 3.23). There was no significant difference between the VL and SVL lines in their total storage root free amino acid pool increase. This was unexpected as in accordance to many sink-source relationships (Morandini, 2009) the availability of a sink such as sporazein would expected to impose a demand on the free amino acid pool.

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Figure 3.23 Total free amino acid content of the storage root pulp. The total free amino acid pool of four month old greenhouse grown plants was nearly doubled in the majority of the VL lines, similarly all three SVL lines were significantly increased in their total free amino acid content in the storage root pulp. (n=3, error bars represent the standard error, asterisk represents statistical significance difference based on a p-value < 0.05 and an ** indicates a p-value < 0.01)

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The free amino acid levels of the leaves of the same plants were measured as well, free amino acids can be transportable forms of nitrogen (Wipf et al. 2002) therefore the effect on the free amino acid pool in the leaves from the increase in free amino acids in the storage root was evaluated.

The free amino acid content increased in all three VL lines by 1.4 to 1.5 fold relative to wild type. In the transgenic lines, however, only VL-15 was statistically significantly different than wild type (Fig. 3.24). Similarly all three SVL lines had increased leaf total free amino acid levels but only SVL-06 and a SVL-11 had statistically significant differences by up to 2.3 fold relative to wild type (Fig. 3.24). The increase of the free amino acid pool in the leaves of VL and SVL lines may be an indication of amino acid transport from the storage roots to the leaves.

217

Figure 3.24 Total free amino acid content in the leaf. The leaves from four month old greenhouse grown wild-type and VL and SVL transgenic lines were analyzed to quantify the total free amino acid content. Only one VL line, VL-15 was significantly increased by

1.5 fold relative to wild-type. Two SVL lines, SVL-06 and SVL-11, were significantly increased by up to 2.3 fold. No significant difference was observed between the increase observed in the VL line and that of the SVL lines. (n=3, error bars represent the standard error, asterisk represents statistical significance difference based on a p-value < .0.05)

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3.3.11 Individual free amino acid analysis in the storage root pulp

The total free amino acid pool was increased in the storage roots of VL and SVL transgenics. It is possible this was due to increased assimilation of cyanide through the

CAS pathway. The products of the CAS pathway are aspartate and ammonia, both of which may be utilized for further amino acid biosynthesis. The individual free amino acid quantities were evaluated in storage wild-type, VL and SVL storage root. Of particular interest in the individual free amino acid analysis was the group of amino acids commonly known as the aspartate family, this group of amino acids is derived from aspartate (Joshi and Jander, 2010).

The amino acids were divided according to their biosynthesis precursors for discussion purposes. The six amino acid groups include: glutamate family (precursor ɑ- ketoglutarate), aspartate family (precursor oxoloacetate), the alanine/valine/leucine group

(precursor pyruvate), the serine/cysteine/glycine group (precursor 3-phosphoglycerate), the aromatic group (precursors phosphoenolpyruvate and erythrose 4-phosphate), and histidine (precursor ribose 5-phosphate).

The amino acids arginine, glutamate, glutamine and proline are members of the glutamate family (Fig. 3.21). The amino acid arginine has been observed to be an abundant free amino acid in cassava storage roots (Stupak 2008, Chapter 2 of this thesis). In our analyses free arginine levels (14.5 nmol of arginine/ mg dry weight) corresponded to 37% of the total free amino acid content. In the VL lines the free arginine levels were significantly increased by 1.8 to 4.2 fold relative to wild type (Fig. 3.25). Similarly all three SVL lines had significant increases in free arginine levels by 2.1 to 3.6 fold relative to wild type (Fig. 3.25). The free arginine accounted for 43 to 66% of the total free amino

219

acids in the VL transgenic storage roots and for 33 to 67% in the SVL lines. The increase in free arginine observed in all VL and SVL lines contributed significantly to the general increase in the free amino acid pool of these transgenic lines in comparison to wild type.

It has been proposed that variations in the free glutamine content correlate with increased nitrogen assimilation possibly through the glutamine synthase (GS)/ glutamine:2- oxyoglutarate amidotransferase (GOGAT) cycle (Foyer et al., 1994; Gojon et al., 1998;

Foyer et al., 2003; Lea et al., 2006). In the GS/GOGAT cycle ammonium and glutamate are condensed into glutamine. The amino group from glutamine is transferred to oxoglutarate, synthesizing an additional molecule of glutamate. The released ammonium from the CAS pathway could potentially be assimilated through the GS/GOGAT cycle for incorporation in to other amino acid biosynthetic pathways. If there is increased ammonium assimilation through the GS/GOGAT cycle in the VL and SVL lines it is likely that the free glutamine levels will be significantly increased (Foyer et al., 1994;

Gojon et al., 1998; Foyer et al., 2003). All three VL lines had significant increases in their free glutamine content ranging from 1.7 to 6.5 fold greater than wild type. The majority of the SVL lines (SVL-06 and SVL-09) had significant increases in their free glutamine content while SVL-11 was reduced by 20% (Fig. 3.25). These results suggest that in the VL and SVL transgenics there is increased ammonium assimilation through the GS/GOGAT cycle.

220

Figure 3.25 Analysis of the free amino acids from the glutamate family in the VL and

SVL lines storage root. The obtained accumulations for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates a p-value <

0.01)

221

Figure 3.25

222

The individual amino acids of the aspartate family pathway were of particular interest to our analysis (Fig. 3.26). In the CAS pathway the condensation of cyanoalanine and water produces asparagine which in turn can be de-amminated by asparaginase to produce aspartate and ammonia. Both asparagine and aspartate are members of the asparate family; additionally aspartate is the precursor for the synthesis of the amino acids lysine, methionine, threonine and isoleucine. In the VL lines, VL-05 and VL-15 had significant increases in free aspartate by 1.6 to 2.4 fold greater than that of wild type. Two SVL lines, SVL-06 and SVL-09, also had significant increases, 2.1 and 4.0 fold, respectively relative to wild type (Fig. 3.26). High flux of aspartate towards the synthesis of other amino acids is a possible reason for some lines not exhibiting an increase in aspartate accumulation.

Very interestingly the levels of asparagine and methionine were reduced in all three VL lines and majority of SVL lines. Asparagine as part of the CAS pathway is the precursor of aspartate through catalysis by asparaginase. In past studies (Elias et al. 1997) it was determined that the asparaginase activity of cassava roots was lower than that of leaves by 40% and the affinity for the substrate of cassava root asparaginase was 16 times less than that of leaf asparaginase. In 1969, Nartey demonstrated that 49% of the radiolabeled carbon in 14CN applied to roots was incorporated into asparagine and 6% into aspartate.

These results suggested that low asparaginase activity impeded the deammination of asparagine to asparate and ammonium. In contrast, we observed a drop in asparagine in

VL (up to 80%) and SVL lines (up to 70%) along with the increase in aspartate (Fig.

3.26). These results indicate that there may be a strong pull for amino acid metabolism through the CAS pathway.

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As mentioned above methionine levels were significantly reduced in all VL lines (30-

40%) and in SVL-06 (40%) and SVL-11 (50%) (Fig. 3.26). The reduced methionine pools may be linked to reduced availability of the substrates required for its synthesis.

The enzyme cystathione gama-synthase (CGS) is the first-dedicated enzyme in methionine synthesis and its substrates are cysteine and O-phosphohomoserine (Kim et al., 2006). Cysteine is required for the proposed cyanide assimilation pathway therefore, its availability for methionine synthesis may be limiting in VL and SVL lines.

Free methionine pools are linked to the regulation of threonine through a product of methionine catabolism, S-adenosylmethionine (SAM). The threonine synthase (first enzyme dedicated for the synthesis of threonine) is allosterically regulated by SAM

(Hesse et al., 2004). Generally the synthesis of threonine is activated when the levels of methionine and SAM are high (Hesse and Hoefgen, 2003). Threonine synthesis is reduced but proceeds in the absence of SAM to regulate threonine synthase (Lee et al.,

2005). While methionine levels were low in all VL and select SVL lines threonine levels increased in both VL and SVL transgenic lines. All VL lines and all SVL lines had significant increases in their threonine levels from 2.1 to 6.6 fold for VL lines and 3.6 to

3.7 fold for SVL lines, compared to wild type (Fig. 3.26). These results suggest that threonine synthesis proceeded in cassava without high free methionine levels in the FAA pool.

224

Figure 3.26 Analysis of the free amino acids from the aspartate family in the VL and

0.01)

225

Figure 3.26

226

In the Alanine/Valine/Leucine group the free alanine content increased in two VL lines

(VL-05 and VL-15) by up to 2.5 times that of wild type (Fig. 3.27). There were no important changes or trends observed in valine or leucine content in the VL and SVL lines relative to each other and wild type.

In the Serine/Cysteine/Glycine group the amino acids serine and cysteine were detected, however, glycine was not. This was possibly due to low levels of this free amino acid in cassava storage roots. Cysteine takes part in the initial cyanide assimilation reaction catalyzed by CAS, therefore the levels of cysteine in the free amino acid pool of VL and

SVL lines were of particular interest. The levels of cysteine and serine were significantly reduced in VL lines. In difference SVL lines had reduced serine levels but cysteine levels with two SVL lines (SVL-06 and SVL-11) were similar to wild type (Fig. 3.27). The amino acid serine is the carbon skeleton donor for the biosynthesis of cysteine. In past reports where a marked reduction of serine was observed it was related to increased cysteine catabolism (Avraham et al. 2005). In addition methionine pools were reduced in

VL lines (40% reduction) and SVL lines (50% reduction) indicating that if cysteine catabolism is taking place it is primarily not for the synthesis of methionine but rather for cyanide assimilation.

227

Figure 3.27 Analysis of the free amino acids from the alanine/valine/leucine group and serine/cysteine/glycine group in the VL and SVL storage root pulp. Glycine was not detected in the four month old storage roots from greenhouse grown cassava. The obtained accumulations for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05)

228

Figure 3.27

229

The final amino acid group analyzed was the aromatic amino acid group and histidine.

Interestingly the accumulation of tryptophan was significantly increased in all SVL lines by 1.7 to 4.8 times that of wild type, while no such increase was observed in VL lines

(Fig. 3.28). On the other hand histidine was significantly reduced in all VL lines (up to

70%) but not in SVL lines (Fig. 3.28). It is not clear what the significance of these results is.

The values of the amino acid concentrations in pmoles / mg dry weight in the free amino acid pool from storage roots of wild type and VL and SVL transgenic lines are presented in Table 3.2. Past evaluations of the amino acid content in cassava storage roots have only focused on the protein bound amino acids (Nassar and Sousa, 2007; Montagnac et al., 2009) however similarities between the free amino acid pool profile and that of protein bound amino acids can be observed particularly in the arginine content. Arginine was found to be the most abundant free amino acid in wild-type storage roots and it is the second most abundant protein bound amino acid constituting 11.0% of all protein bound amino acids. (Montagnac et al., 2009). The two amino acids that were found in low abundance in the free amino acid pool of wild-type storage roots were tyrosine (90.2 ±

26.9 pmoles / mg dry weight, Table 3.2) and tryptophan (112.7 ± 11.7 pmoles/ mg dry weight, Table 3.2), these two amino acids are similarly found in low abundance as protein bound amino acids representing 0.4% (tyrosine) and 0.5% (tryptophan) of the total protein bound amino acids (Montagnac et al., 2009).

230

Figure 3.28 Analysis of the free amino acids from the aromatic group and histidine in the

VL and SVL storage root pulp. The obtained accumulations for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates a p-value <

0.01)

231

Figure 3.28

232

WT VL-04 VL-05 VL-15 SVL-06 SVL-09 SVL-11 Arg 18781.2 ± 4217.1 56343.6 ± 4981.3 33806.2 ± 3913.8 78881.0 ± 8357.2 39440.5 ± 4682.1 52587.4 ± 7923.5 67612.3 ± 6812.5 Glu 4201.7 ± 659.5 3361.3 ± 369.2 6722.7 ± 826.4 5462.2 ± 324.8 14705.8 ± 2305.1 7142.8 ± 981.3 5042.0 ± 824.6 Gln 3650.4 ± 606.6 22997.5 ± 982.4 9491.0 ± 873.2 6205.7 ± 328.5 5475.6 ± 735.1 12411.4 ± 1356.8 2920.3 ± 237.4 Pro 212.2 ± 47.2 191.0 ± 26.9 679.0 ± 165.4 127.3 ± 43.2 275.9 ± 92.4 191.0 ± 24.6 63.7 ± 14.5 Asp 5797.9 ± 821.6 5821.5 ± 734.1 13914.9 ± 988.4 9276.6 ± 300.4 23191.5 ± 1530.1 12175.5 ± 603.1 5923.6 ± 665.1 Asn 262.1 ± 83.2 104.9 ± 35.2 52.4 ± 13.8 124.8 ± 42.7 550.5 ± 87.3 183.5 ± 10.4 78.6 ± 15.7 Thr 516.5 ± 128.8 3409.1 ± 301.4 1136.4 ± 141.0 1084.7 ± 73.9 1911.2 ± 176.4 1792.7 ± 409.6 1859.5 ± 200.1 lle 148.7 ± 5.3 401.6 ± 40.3 327.2 ± 39.7 208.2 ± 12.6 580.1 ± 76.4 297.5 ± 43.8 178.5 ± 39.4 Met 109.9 ± 28.2 77.0 ± 3.5 76.5 ± 5.2 66.0 ± 2.8 66.5 ± 5.2 98.9 ± 11.2 55.0 ± 3.2

233 Lys 308.5 ± 72.5 246.8 ± 30.2 277.6 ± 60.3 250.4 ± 45.3 771.2 ± 232.5 462.7 ± 120.5 339.3 ± 76.4

Ala 543.9 ± 167.0 435.1 ± 67.5 1359.7 ± 122.1 924.6 ± 65.3 1957.9 ± 876.9 979.0 ± 144.1 652.6 ± 67.8 Val 302.2 ± 129.0 90.7 ± 11.1 181.3 ± 67.3 180.3 ± 47.3 544.0 ± 178.4 272.0 ± 45.2 151.1 ± 31.4 Leu 177.3 ± 44.3 212.8 ± 34.5 141.9 ± 32.1 106.4 ± 36.1 248.3 ± 45.3 230.5 ± 56.1 53.2 ± 4.2 Ser 1988.5 ± 573.8 994.2 ± 124.6 795.4 ± 113.2 1012.4 ± 125.3 795.4 ± 120.3 1391.9 ± 356.7 804.2 ± 78.3 Cys 253.3 ± 56.4 177.3 ± 34.6 169.2 ± 13.5 173.4 ± 18.3 202.7 ± 33.1 177.3 ± 31.6 304.0 ± 67.1 Phe 169.9 ± 25.5 85.0 ± 16.7 186.9 ± 32.1 172.3 ± 43.1 458.8 ± 156.8 203.9 ± 45.3 254.9 ± 68.9 Trp 112.7 ± 11.7 101.5 ± 12.4 115.4 ± 20.4 157.8 ± 43.2 349.5 ± 67.4 191.7 ± 23.2 541.2 ± 54.7 Tyr 90.2 ± 26.9 54.1 ± 7.1 89.2 ± 15.4 27.1 ± 3.2 72.2 ± 25.6 36.1 ± 4.5 27.1 ± 2.5 His 640.3 ± 69.7 256.1 ± 31.1 192.1 ± 24.5 320.1 ± 32.1 1024.4 ± 187.5 1856.8 ± 177.2 704.3 ± 64.3

Table 3.2 Storage root individual free amino acids. Free amino acid concentration in pmoles/mg dry weight of storage roots from

wild-type, VL and SVL transgenic lines.

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3.3.12 Individual free amino acid analysis in leaves

The total free amino acid pool in the leaves from VL and SVL transgenics was increased in all lines and significantly in VL-15 (1.5 fold that of wild type) , SVL-06 and SVL-11

(up to 2.3 fold relative to wild type). It is likely that the total increase in leaf free amino acid pools is due to amino acid transport (Wipf et al., 2002) from the roots. The individual amino acids were analyzed to determine if a select group of amino acids had been increased or if there were any relationships between the observed increases or decreases in the free amino acid pool of storage roots to that of the leaf. The concentration (pmoles / mg dry weight) of each amino acid detected in the free amino acid pool of leaves from wild type, VL and SVL lines is presented in Table 3.3. In past reports the protein bound amino acid content has been measured in cassava leaves

(Nassar and Sousa, 2007; Montagnac et al., 2009) but not the concentrations of free amino acids. In the data presented here in the free amino acid pool of cassava leaves the amino acids glutamate (Table 3.3) and aspartate (Table 3.3) were the most abundant. In the previously reported values of protein bound amino acids in cassava leaves the glutamate ( 7.1% of total protein bound amino acids) and aspartate (8.7 % of total protein bound amino acids) are the second and third most abundant protein bound amino acids

(Montagnac et al., 2009), however leucine which is the most abundant protein bound amino acid (9.7%) (Montagnac et al., 2009) was only the 8th most abundant free amino acid (Table 3.3). Therefore there seems to be a relation between abundance in protein bound content and that in the free amino acid pool only for some amino acids.

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WT VL-04 VL-05 VL-15 SVL-06 SVL-09 SVL-11 His 534.9 ± 11.3 600.8 ± 90.9 541.6 ± 32.0 692.6 ± 58.0 692.6 ± 58.0 460.9 ± 52.7 436.6± 42.3 Asn 799.0 ± 117.2 2368.0 ± 310.5 3883.4 ± 202.7 2844.6 ± 110.7 2844.6 ± 110.7 1262.6 ± 203.4 1933.0± 109.5 Ser 4141.9 ± 254.6 5485.9 ± 841.0 5189.6 ± 182.2 5498.3 ± 475.4 5498.3 ± 475.4 4885.7 ± 538.0 6706.9± 1089.6 Gln 2109.2 ± 288.8 9561.1 ± 873.3 8520.6 ± 487.1 5461.0 ± 264.8 5461.0 ± 264.8 6392.1 ± 635.5 4792.9 ± 843.8 Arg 1293.4 ± 65.5 3022.0 ± 299.3 770.0 ± 83.4 902.1 ± 18.1 902.1 ± 18.1 1157.8 ± 15.5 646.3 ± 33.6 Gly 633.1 ± 56.7 720.7 ± 93.4 1903.9 ± 450.9 2525.3 ± 163.1 2525.3 ± 163.1 548.3 ± 105.1 805.6 ± 18.2 Asp 8496.2 ± 771.6 8606.7 ± 1323.6 8546.3 ± 445.8 9423.6 ± 451.2 9423.6 ± 451.2 12898.2 ± 777.3 16296.3 ± 2274.1 Glu 10417.1 ± 1805.5 15204.8 ± 1966.3 15099.7 ± 1204.8 15843.1 ± 1759.3 15843.1 ± 1759.3 19490.2 ± 1896.8 16092.6 ± 688.8 Thr 1526.1 ± 180.4 1270.2± 157.0 1730.2 ± 406.7 2350.1 ± 315.1 2350.1 ± 315.1 1289.8 ± 212.0 2473.6 ± 139.1 235 Ala 4848.4 ± 529.7 6752.0 ± 237.1 5060.0± 175.4 5473.4 ± 321.3 5473.4 ± 321.3 6324.1 ± 285.9 8956.4± 1119.5

Pro 704.8 ± 67.6 1003.4 ± 61.4 1253.7 ± 337.9 1965.8 ± 92.7 1965.8 ± 92.7 896.7 ± 145.6 1683.1 ± 21.1 Cys 453.7 ± 76.7 224.8 ± 3.2 472.9 ± 19.4 603.9 ± 68.1 603.9 ± 68.1 412.3 ± 173.2 714.0 ± 34.8 Tyr 381.7 ± 64.4 403.4 ± 246.6 610.1 ± 152.5 408. ± 21.5 408.2 ± 21.5 401.9 ± 190.6 465.3 ± 19.9 Met 298.1 ± 30.4 171.6 ± 4.5 176.9 ± 9.2 159.2 ± 14.8 159.2 ± 14.8 311.1 ± 22.7 227.1 ± 7.9 Val 829.3 ± 110.1 943.9 ± 79.3 1766.2 ± 290.2 2470.6 ± 147.1 2470.6 ± 147.1 1051.7 ± 139.5 1427.7 ± 74.0 lle 521.2 ± 81.6 242.7 ± 22.5 701.6 ± 105.4 960.7 ± 65.8 960.7 ± 65.8 456.7 ± 155.2 769.9 ± 11.5 Leu 1164.4 ± 207.7 291.9 ± 11.7 976.9 ± 89.9 1502.7 ± 174.9 1502.7 ± 174.9 826.3 ± 429.8 1755.4 ± 58.2 Phe 670.4 ± 95.2 454.8 ± 82.2 741.8 ± 480.4 1886.4 ± 101.3 1886.4 ± 101.3 652.2 ± 201.1 912.8 ± 16.3 Trp 355.5 ± 44.4 377.0 ± 72.2 356.4 ± 14.9 1047.1± 435.4 1047.1 ± 435.4 200.9 ± 25.1 287.8 ± 66.1

Table 3.3 Leaf individual free amino acids. Concentrations of each individual amino acid in pmoles / mg dry weight in the leaf free

amino acid pool from wild type and VL and SVL transgenic lines after 4 months of greenhouse growth. The variation indicates the

standard error.

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The values obtained for wild type were compared to those for each of the transgenic VL and SVL lines. The amino acids quantified were divided according to their biosynthesis precursors. The six amino acid groups were: glutamate family (precursor ɑ-ketoglutarate)

(Fig. 3.29), aspartate family (precursor oxoloacetate) (Fig. 3.30), the alanine/valine/leucine group (precursor pyruvate) (Fig. 3.31), the serine/cysteine/glycine group (precursor 3-phosphoglycerate) (Fig. 3.32), the aromatic group (precursors phosphoenolpyruvate and erythrose 4-phosphate) (Fig. 3.33) and histidine (precursor ribose 5-phosphate) (Fig. 3.34). Select amino acids are discussed below.

Interestingly arginine was not the most abundant amino acid in the leaves; free glutamate was the most abundant free amino acid. In wild-type it accumulated to a concentration of

10.4 nmol free glutamate/ mg dry weight, representing 26% of the total free amino acid pool. Only in SVL-09 and SVL-11 was the accumulation of free glutamate significantly increased. In comparison to wild-type these transgenic lines had 90 and 50 % more glutamate than wild type, respectively.

Similar to roots the free glutamine content of leaves was elevated in the transgenic lines.

In fact glutamine was the only FAA whose levels increased significantly in the leaves of all three VL and three SVL lines. Glutamine increased up to 4.5 fold in VL lines and up to 3.4 fold in SVL lines in relation to wild type.

In contrast to roots, free asparagine was significantly increased in all three VL lines.

Asparagine also increased in all SVL lines, however, only SVL-06 and SVL-11 had significantly different increases. Asparagine is a widely used form of transportable

236

nitrogen in plants (Lea et al., 2007; Gaufichon et al., 2010) and therefore an increase in asparagine in the leaves may only indicate a general increase in amino acid transport.

In the storage roots methionine levels were reduced in all VL and a majority of SVL lines

Similarly, in the leaves of all three VL lines there were reductions in methionine levels while for the SVL lines methionine levels were similar to wild type. The decrease of methionine has implications not only related to the metabolism of cysteine and serine but also from a nutritional aspect. Methionine is one of the essential amino acids and it is an amino acid that is found in limiting quantities in select crops such as legumes (Ufaz and

Galili, 2008). A significant decrease in the methionine free amino acid pool in VL lines in storage roots and leaves may limit the quantity of methionine available for protein synthesis and ultimately affect the nutritional quality of the crop.

Cysteine and serine levels were unchanged in both VL and SVL plants possibly indicating a greater demand for cysteine as part of the CAS assimilation pathway in roots unlike leaves.

237

Figure 3.29 Analysis of the free amino acids from the glutamate family in the VL and

SVL leaf. The levels for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05)

238

Figure 3.29

239

Figure 3.30 Analysis of the free amino acids from the aspartate family in the leaf of VL and SVL lines. The obtained accumulations for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates p-value < 0.01)

240

Figure 3.30

241

Figure 3.31 Analysis of the free amino acids from the alanine/valine/leucine group and serine/cysteine/glycine group in the VL and SVL leaf. The obtained accumulations for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates a p-value < 0.01)

242

Figure 3.31

243

Figure 3.32 Analysis of the free amino acids from the aromatic group in the VL and SVL leaf. The obtained accumulations for each amino acid are presented as fold increase or decrease relative to the values obtained for wild type. The fold increase for each amino acid is depicted a top each bar of the graph. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates p-value < 0.01)

244

Figure 3.32

245

3.3.13 Protein content

The expression of a vacuolar linamarase and the resulting increase of linamarin catabolism is likely the cause for the observed increases in the total free amino acid pool of the storage roots and leaves of VL and SVL lines. It was hypothesized originally that an increase in the free amino acid pool would result in an increase in protein. This was not observed in the VL lines. The protein content in the storage root pulp of the three VL lines was not significantly different than wild type (Fig. 3.33). In addition, we co- expressed a potentially strong protein sink in roots in the SVL lines. In contrast the SVL lines expressing the double construct with sporazein, had substantial increases in their root storage protein levels by 1.6 to 1.9 times that of wild type (Fig. 3.29). It appears as if the expression of an amino acid sink in the form of the novel protein sporazein has a positive impact on protein accumulation, and that increase of the free amino acid pool in cassava storage roots such as was observed in VL lines does not result in protein increase.

That being said, the free amino acid pool sizes in SVL lines were still 1.9 to 2.4 fold greater than those in wild type suggesting that sporazein expression was insufficient to take full advantage of the additional free amino acids. It would have been informative to express sporazein by itself and determine the free amino acid pool size to see if enhanced sink strength alone had an impact on FAA.

246

Figure 3.33 Storage root pulp protein analysis. The root pulp crude protein content was measured in the wild-type and transgenic lines at 4 months old. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates p-value <

0.01)

247

The leaf protein content was also analyzed in the 4 month old greenhouse grown wild- type, VL and SVL plants. There was a significant protein increase observed in VL-05 and

VL-15 by up to 3.2 fold relative to wild type (Fig. 3.34). Similarly the leaf protein was increased in all SVL lines in the range of 1.5 to 3.2 times that of wild type (Fig. 3.34).

The protein and free amino acid increase in the leaves of VL and SVL lines are possible indicators that the leaves act as a strong sink tissue for amino acids for protein synthesis.

The observed increases in the SVL lines additionally present a problem as the expression of sporazein in the storage root is possibly not a strong sink in comparison to younger developing leaves which are considered strong sink tissues (Sonnewald and Willmitzer,

1992) for many metabolites.

248

Figure 3.34 Leaf protein analysis. The leaf crude protein content was measured in the wild-type and transgenic lines at 4 months old. (n=3 from three different plants of the same line, the error bars indicate the standard error, an * indicates statistically different as determined by a p-value < 0.05 and an ** indicates p-value < 0.01)

249

3.3.14 Total Nitrogen analysis of leaves and storage root

To determine if the transgenic lines had a net increase in nitrogen concentration, we summed the total N per unit dry weight for roots and leaves. The total µmoles of nitrogen per mg dry weight was calculated for wild type, VL and SVL lines in storage roots and leaves.

The reduced nitrogen concentration in the storage roots was increased significantly in all three VL lines up to 5.5 times that of wild type while SVL lines were increased by up to

7.1 fold relative to wild type (Fig. 3.35). In the VL lines the marked increase in reduced nitrogen content was due to an increase in the free amino acid pool, in particular arginine.

For each mole of arginine there are four moles of nitrogen. In all VL and SVL lines there was a pronounced increase in the free arginine content which contributed substantially to the increase in total in total reduced nitrogen in the roots of VL and SVL lines. In the leaves of VL-05, VL-15 and all SVL lines the increase in total nitrogen was due to increased protein (Fig. 3.35).The greatest net gains in N were observed in the leaves of the transgenic lines, however.

250

Figure 3.35 Total moles of reduced nitrogen in wild-type and VL and SVL transgenic lines. Stacked graph representing the total nitrogen from protein, free amino acids and linamarin of leaf and storage root from wild-type, VL and SVL cassava plants. Protein contribution to the total nitrogen is in dark grey, free amino acid pool contribution to the total Nitrogen is in light grey and linamarin contribution to the total nitrogen is in white.

In general the nitrogen content of the storage roots of all VL and SVL lines was increased relative to wild-type. With the exception of the VL-04 line the remaining VL lines and all

SVL lines had an increase in their total nitrogen content in the leaves relative to wild- type.

251

Figure 3.35

252

To determine if the yield of total reduced N was increased in transgenic lines, we multiplied the nitrogen concentration times the total biomass for storage roots (Table

3.4). The total reduced nitrogen in VL storage roots was increased by up to 3.7 fold relative to wild type while that of the SVL storage roots was increased by up to 4 fold.

It is clear from the total nitrogen data that in the storage root of the SVL lines the free amino acid pool contributed the majority of the reduced nitrogen and not the increased protein content observed in the roots of these lines (Fig. 3.35). It is interesting in SVL plants that the expression of a storage root protein did not affect the overall reduced nitrogen distribution or the yield of total reduced nitrogen. In SVL plants the “source” or reduced nitrogen in the form of amino acids for the “sink” or sporazein are present however it appears as if other mechanisms are in place that control the synthesis of sporazein and hinder a higher accumulation of this storage protein.

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Table 3.4 Total nitrogen in the storage roots of wild type, VL and SVL lines.

Total µmol N in storage Line roots Relative to WT WT 1620.7 1.0 VL-04 5955.6 3.7 VL-05 4338.3 2.7 VL-15 4020.7 2.5 SVL-06 6474.3 4.0 SVL-09 3720.1 2.3 SVL-11 3895.0 2.4

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3.3.15 Summary of VL and SVL lines

In the VL transgenic cassava lines a vacuolar linamarase was expressed driven by the root-specific patatin promoter while in the SVL lines in addition to the vacuolar linamarase a novel storage protein known as sporazein was expressed. In both VL and

SVL lines there was an observed decrease in linamarin content in the storage roots (Fig.

3.36 section 1) possibly due to the increased hydrolysis of linamarin by the vacuolar linamarase. Linamarin content was similarly reduced in the leaves. Linamarin synthesis takes place in the leaves and from there it is transported to the roots. It is possible that the linamarin decrease observed in VL and SVL lines is due to an increased flux of linamarin from the leaves to the roots. The total free amino acid content of VL and SVL lines was doubled (Fig. 3.36 section 2) with a remarkable increase in total free arginine content of up to 4.2 fold in VL lines and 3.6 fold in SVL lines. In the case of VL and SVL lines an increase in free amino acid content is likely related to an increase in assimilated cyanide via the cyanide assimilation pathway to produce aspartate and ammonia which may be used for the biosynthesis of other amino acids. An increase in total free amino acid content in the storage roots of VL lines did not result in an increase in protein in the storage roots (Fig. 3.36 section 3) while in the SVL lines a protein increase of nearly double was observed (Fig. 3.36 section 4). The increase in SVL lines was likely due to the expression of the storage protein sporazein. Finally an increase in both total free amino acid and protein content was observed in select VL and SVL lines (Fig. 3.36 section 5).

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Figure 3.36 Summary of the overall effect of expression of a vacuolar linamarase with

(VL lines) or without the storage protein sporazein (SVL lines) in cassava transgenic lines. Section 1 depicts the decrease of storage root linamarin. Section 2 depicts the increase in total free amino acids. Section 3 represents only VL lines which did not present an increase in storage root protein content. Section 4 represents only SVL lines which presented an increase in storage root protein content. Section 5 depicts the increase in free amino acid and protein content in the leaves of some VL and SVL lines.

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Figure 3.36

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3.4 DISCUSSION

Cassava in Sub-Saharan Africa has evolved from a farmer emergency crop (Jones, 1959) to a staple food and an important calorie source (Montagnac et al., 2009a). However, one of the drawbacks of cassava is the low protein content of the storage roots. With less than

3% (w/v) protein (Ceballos et al., 2006) the cassava storage roots are significantly lower in protein than other staple crops such as maize (9.4%) (USDA, 2010).

Protein-energy malnutrition is widespread in many regions were cassava is consumed

(FAO, 2010), for example in the Dem. Rep. of Congo the daily protein intake is only 25 g of protein / per person (FAOSTAT, 2009) a third of the daily recommended intake by

FAO of 75 g of protein/ per person a day (FAO, 2010). In addition, it has been suggested that high consumption of cassava places those populations at even greater risk of protein malnutrition. Stephenson et al. (2010) observed that when cassava is consumed as a staple food and the diet is not supplemented with additional protein sources there is an inverse correlation between cassava intake and protein intake in the diet of the children evaluated. The improvement of the protein content of cassava storage roots would potentially impact the diet of millions of people by providing a more nutritional food source.

There are several proposed strategies in the literature for protein and/or free amino acid increase in plants. It has been proposed to overexpress native proteins or to increase the available free amino acids through the modification of amino acid biosynthetic pathways

(Stupak et al., 2006; Ufaz and Galili, 2008). However in this chapter, we propose to utilize the nitrile group (-CN) from the cyanogenic glucoside linamarin as a reduced

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nitrogen source that could potentially lead to an increase in free amino acids and protein.

It has been hypothesized in past reports that linamarin is involved in the nitrogen metabolism of cassava, as it has been shown to be involved in Hevea brasilensis

(Lieberei et al., 1985; Selmar et al., 1988). Evidence for the involvement of linamarin as a nitrogen resource in cassava was provided by Siritunga (2002) when it was observed that cassava transgenic plants with reduced linamarin synthesis in the leaves resulted in decreased linamarin accumulation in the root and an inability of those transgenic lines to grow without a reduced source of nitrogen in the media. It was hypothesized then that linamarin transported from the leaves to the roots provided reduced nitrogen source for the roots. The incorporation of nitrogen from linamarin requires the hydrolysis of linamarin to release acetone cyanohydrin which in turn degrades (at pH>5 or temperature

> 35 ˚C) to produce cyanide. This is followed by the assimilation of cyanide through the

CAS pathway (Fig. 3.1) leading to the synthesis of aspartate and ammonia. Furthermore, aspartate and ammonia can be incorporated in to the biosynthesis of other amino acids

(Jander and Joshi, 2010). Nevertheless in cassava the hydrolysis of linamarin is limited due to the spatial separation between the cyanogenic glucoside and its β-glucosidase linamarase. Furthermore, linamarin in cassava is stored in the vacuole (White et al.,

1998) while linamarase is found in the apoplast (Mkpong et al., 1990). It was proposed that the targeting of linamarase to the cytoplasm or to the vacuole could potentially eliminate the spatial limitations for linamarin hydrolysis. Expression of linamarase in the cytoplasm would increase linamarin hydrolysis during the transport of linamarin to its storage organelle, the vacuole, while expression of linamarase in the vacuole would increase hydrolysis of linamarin at its storage location. Additionally, we proposed to

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express a storage protein with the vacuolar linamarase, to act as a protein sink for the increased free amino acid pool.

Interestingly, no positive transformants were recovered when the linamarase was targeted to the cytoplasm, a few lines were positive for the T-DNA insertion containing the ΔN-ter linamarase gene but no lines were recovered where the transcription of this cytoplasmic linamarase was active. A possible explanation is related to the conditions required for acetone cyanohydrin degradation. Acetone cyanohydrin is released after linamarin hydrolysis and as mentioned before can be spontaneously degraded at high pH >5 (White et al. 1998). The cytoplasmic pH is slightly alkaline pH 7.4-7.5 (Roberts et al., 1980;

Gout et al., 1992) at this pH any acetone cyanohydrin present in the cytoplasm would rapidly degrade releasing cyanide. The enzymes responsible for the assimilation of cyanide have been shown to be present in storage root tissues of cassava (Elias et al.,

1997), however, the efficiency of cyanide assimilation under in vivo conditions has not been reported for cassava. It is possible that an accelerated rate of cyanide production may be toxic to the cell. Cassava somatic embryos are non-cyanogenic (Joseph et al.,

1999), however, developing cotyledons and germinating embryos are cyanogenic (Joseph et al., 1999). Any transgenic lines where ΔN-ter linamarase was translated possibly died during the regeneration stages after transformation due to the high release of cyanide. In contrast, the expression of linamarase in the vacuole may not result in as rapid a release of cyanide. The pH in the vacuole fluctuates in the range of pH 3 to 6 (Lodish et al.,

2004), the fluctuation of pH within the vacuole would cause similar fluctuations in the degradation of acetone cyanohydrin and consequentially on the release of cyanide. This

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may have prevented sudden high increases in cellular cyanide and thus cassava plants expressing a vacuolar linamarase were successfully regenerated.

Transgenic cassava were regenerated expressing a vacuolar targeted linamarase, these lines were named VL. Additionally transgenic lines were recovered expressing a vacuolar targeted linamarase as well as the storage protein sporazein, these lines were named SVL.

The expression of a vacuolar targeted linamarase in cassava storage roots substantially affected total linamarin accumulation of VL and SVL lines. Root linamarin levels were reduced significantly in the storage root of VL lines (by up to 44%) and of SVL (by up to

45%). In addition linamarin levels were significantly reduced in the leaves of VL lines

(by up to 45%) and in SVL lines (by 25%). Linamarin synthesis takes place primarily in the leaves and then it is transported to the roots (Bediako et al., 1981; Andersen et al.,

2000; Siritunga and Sayre 2003). The reduction of linamarin in the leaves may indicate either a reduction of linamarin synthesis or an increase in the linamarin flux to the roots.

The reduction of linamarin synthesis in the leaves as previously reported by Siritunga and

Sayre (2003) reduced the linamarin content in the storage roots to less than 1% that of wild type. In the case of VL and SVL lines the storage root linamarin was not reduced as markedly as that observed by Siritunga and Sayre (2003). Reduction of linamarin synthesis in the leaves of VL and SVL lines along with the proposed increased hydrolysis of linamarin in the storage root through expression of vacuolar linamarase would be expected to render a larger reduction of storage root linamarin than what was observed.

Therefore, it is likely that the reduced linamarin content in the leaves is due to an increase in linamarin flux to the storage roots and as discussed below assimilation into free amino acids.

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As was predicted, the free amino acid pool in the storage roots of VL and SVL lines was substantially increased by 2.2 fold in VL lines and up to 2.4 fold in SVL lines. In SVL storage roots there was also a 2 fold increase in protein whereas VL storage roots exhibited no substantial change in protein content relative to wild type. The protein increases observed in SVL lines are possibly due to the expression of the novel storage protein sporazein. With this in mind it was expected that the increase of protein in the storage root of SVL lines would have an effect on the free amino acid pool of this organ and that SVL lines in comparison to VL lines would have a lower free amino acid pool than the VL lines. Interestingly no difference was observed between the free amino acid pool of VL lines and those of SVL lines. Analysis of the total reduced nitrogen content

(including only linamarin, free amino acids and protein) in the storage roots of VL and

SVL lines revealed a marked increase of 5.5 fold in VL lines and 7.1 fold for SVL lines in comparison to wild type. However, once again no significant differences were observed between VL and SVL lines. It is of particular interest to note that the main contribution of reduced nitrogen in the VL and SVL storage roots is in the form of free amino acids. In past reports it has been thought that available free amino acids can readily be incorporated in to storage proteins (Ufaz and Galili, 2008). If such is the case the expected increase in protein in the storage root of SVL lines in relation to the available free amino acids would have been larger than the obtained near 2 fold increase. A possible explanation is given through the analysis of the individual free amino acid contributions to the free amino acid pool of storage roots. Upon analysis it is observed that a large quantity of the free amino acid pool of VL lines (66%) and SVL lines (67%) is found in the form of free arginine. It is possible that the high increase observed in free

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arginine but not of other free amino acids limited the synthesis of sporazein as in situ limitations of free amino acids are known to hinder de novo protein synthesis (Sands et al., 2009). Alternatively, the large increases in arginine may be involved in reduced nitrogen transport to leaves which exhibited the greatest net changes in protein levels.

This hypothesis is consistent with leaves being the major N sink in cassava. Previous investigations have concluded that green photosynthetically active and growing leaves are a major N sink (Hirel et al., 2005; Yasumura et al., 2007) and amino acids are an important form of transportable nitrogen from source to sink organs (William and Lee,

2001). The leaf free amino acid pool in the VL lines increased up to 1.5 fold and in the

SVL lines up to 2.3 fold. The protein levels of VL and SVL lines in the leaves increased by up to 3.2 fold. Since there was no increase in root protein levels in the VL lines it is suggested that the additional reduced nitrogen in amino acids was transported to and used by the leaf for protein synthesis. This process apparently also occurs in SVL lines expressing sporazein indicating that leaves are the major nitrogen sink of cassava and in turn influence protein accumulation in roots.

The analysis of the individual free amino acid pool in the storage roots revealed decreased accumulation of cysteine, methionine and serine in VL and SVL lines.

Cysteine plays an important role in the CAS pathway as it is the substrate along with cyanide for β-cyanoalanine synthase (Miller and Conn, 1980). If the CAS cyanide assimilation pathway is increased in VL and SVL lines the demand for cysteine would similarly be increased. This correlates with the decrease in cysteine accumulation observed. Furthermore, the first dedicated enzyme for methionine synthesis is cystathione

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gama-synthase (CGS) and its substrates are cysteine and O-phospho-L-homoserine

(OPH) (Kim et al., 2006). Without adequate free cysteine pools the synthesis of methionine would be limited as well. Finally, serine is linked to these two amino acids as the carbon skeleton donor for cysteine biosynthesis (Bogdanova et al., 1995, 1997). A marked reduction in serine has been observed before when cysteine catabolism was increased by over expression of CGS (Avraham et al., 2005), similarly in VL and SVL lines cysteine catabolism is increased but not towards the synthesis of methionine but rather towards the assimilation of cyanide. The reductions in methionine, serine and cysteine are in accordance with an increase utilization of cysteine for cyanide detoxification.

In summary, in this chapter we have provided evidence for the integration of the nitrile group of linamarin in to the nitrogen metabolism as evidenced by the increased free amino acid pool in VL and SVL lines. In addition, it was observed that an increase in the free amino acid pool in cassava storage roots does not result in an increase in protein content. Furthermore, while it was observed that the expression of a storage protein improves the overall protein content of cassava storage roots; it was observed that limitations in the free amino acid pool (high arginine concentrations) and the presence of the leaves as a major N sink likely limit root protein increases. Transgenic plants expressing a novel root storage protein did not provide a strong enough nitrogen sink.

Interestingly, other starch containing tissues such as the potato and yam tubers as well as the sweet potato storage root are known to accumulate storage proteins therefore acting as a stronger nitrogen sink (Maeshima et al., 1985; Sonnewald et al., 1989; Conlan et al.,

1998; Shewry et al, 2003). However, the potato and yam tubers as well as the sweet

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potato storage root are able to act as propagules (Lebot, 2009), thus requiring storage proteins to be broken down during sprouting as a source of nitrogen (Shewry, 2003). In comparison the cassava storage root is not able to act as a propagule (Alves, 2005).

Another characteristic of the storage proteins of potato, sweet potato and yam is their localization to vegetative protein storage vacuoles (Shewry, 2003), it is not known if cassava storage roots possess vegetative protein storage vacuoles. These factors may be taken in to consideration to determine specific factors that may be modified to produce a strong nitrogen sink in cassava storage roots.

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3.5 SUMMARY OF STRATEGIES FOR THE INCREASE OF FREE AMINO

ACID AND PROTEIN CONTENT IN CASSAVA STORAGE ROOTS

In CNIA lines the strategy was to increase the reduction of nitrate in cassava roots through the expression of a mutated nitrate reductase. It was proposed that through the increased reduction of nitrate an increase would be obtained for ammonium and consequentially free amino acids. Potentially an increase in free amino acids would lead to increased protein content. In the VL and SVL lines the strategy was to increase the cyanide assimilation into amino acids through the β-cyanoalanine pathway through increased hydrolysis of linamarin by the expression of a vacuolar targeted linamarase.

The increase in free amino acids in VL and SVL lines was similarly proposed to lead to an increase in protein. In difference to CNIA and VL lines, the SVL lines expressed sporazein (novel storage protein) in the storage roots.

In CNIA, VL and SVL lines an increase of up to 2 fold in the total free amino acid pool was obtained in cassava storage roots. This increase in free amino acids in the storage roots lead only to a significant protein increase in the storage roots of SVL linesbut not in

CNIA and VL lines. These results and those of leaf and storage root linamarin as well as leaf and protein free amino acid content are summarized in Table 3.3.

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Table 3.5 Overall summary of the results obtained for free amino acid (FAA), protein and linamarin content in transgenic cassava

CNIA, VL and SVL lines. Highlighted in yellow are the increases observed in the storage roots of CNIA, VL or SVL lines; red

highlight indicates a decrease observed in the storage roots of CNIA, VL or SVL lines.

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CHAPTER 4

ALTERNATIVE ROOT GENE PROMOTERS FOR EXPRESSION

OF TRANSGENES IN CASSAVA STORAGE ROOTS

4.1 INTRODUCTION

In recent years efforts have increasingly focused on improving the nutrient content of cassava storage roots through genetic modification (Taylor et al. 2004, Thomson 2008).

Unfortunately, unlike many other staple crops there is a limited set of molecular tools available for use with cassava (Delmer 2005). An example is the low number of gene promoters with confirmed expression in cassava storage roots. Recently, Beltran et. al

(2010) reported on the activity of the promoter from the glutamic-acid rich protein gene

Pt2L4. When this promoter was fused to the GUSPlus gene a strong expression in the cassava storage roots and vascular stem tissues was observed. In other previous reports

Siritunga et al. (2003) showed that the patatin class I promoter from potato (Kim et. al

1994) had root-specific activity in cassava and Zhang et al. (2003) described two cassava promoters p15 and p54 with activity in storage roots and vascular tissues. However, to date there are no reports of promoters with tissue-type specific expression in cassava roots. The availability of such promoters in addition to the set of currently known promoters could potentially impact the nutrition engineering and industrial applications of cassava by opening new opportunities for expression of transgenes. While at the moment much of the focus is on individual genetic traits introduced into cassava it is clear that the need for gene stacking will eventually be reached. However ,if identical or similar promoters are used there is a risk of Transcriptional Gene Silencing (TGS) by

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promoter homology (Que et. al 1998, Mette et. al 2000, Halpin 2005). The TGS effect is an epigenetic inactivation arising when multiple copies of a sequence are present in the genome and involves increased promoter methylation that may activate chromatin components responsible for transcriptional silencing (Kooter et al. 1999). With the addition of novel promoters a variety of traits for nutrition enhancement, virus resistance among others may be stacked in cassava with reduced risk of TGS.

In 2006, Lee et al. reported on the activity of 61 different promoters of transcription factors expressed in an enriched manner in Arabidopsis roots. We sought to examine the strength and expression pattern of four of these promoters: A14, E40, E49 and S8. These promoters were selected based on the expression patterns in different root tissues (Table

4.1 and Figure 4.1).

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Gene name Gene TAIR # Promoter Size Expression

A14 AT5G43040 2.8 kb Epidermis, enriched in

atrichoblast

E40 AT5G24380 4.0 kb Endodermis and stele

from late maturation

zone

E49 AT3G05150 2.4 kb Cortex in elongation

zone and above

S8 AT5G60200 3.1 kb Pericycle in phloem from

elongation zone and up

Table 4.1 The four selected promoters have different tissues of expression in Arabidopsis roots.

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Figure 4.1 Expression maps of A14, E40, E49 and S8 obtained from the Arabidopsis

Gene Expression Database (AREX, www.arexdb.org) (Cartwright et al. 2009)

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The A14 promoter (Fig. 4.2) regulates expression of a putative DC-1 domain containing protein gene found in the epidermis of the root. The E40 promoter (Fig. 4.3) regulates a

Yellow-Stripe Like 2 gene (YSL). It has been shown that the YSL2 protein acts as a metal transporter for Fe and Cu chelated with nicotianamide acid (DiDonato et. al 2004), this promoter effects expression in the endodermis and stele.

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Figure 4.2 A14 promoter sequence. The promoter sequence starts at the 17265621 bp of the 5th chromosome of Arabidopsis and ends at 17268403 bp. The Transcription Start Site

(TSS) is double underlined and highlighted in yellow, the start codon (ATG) is highlighted in yellow and predicted TATA boxes are underlined. The TSS and TATA boxes were predicted using the SoftBerry TSSP/ Prediction of PLANT Promoters tool

(http://linux1.softberry.com/berry.phtml?topic=tssp&group=programs&subgroup=promoter) The promoter sequence is numbered according to the start codon position and the start codon position and sequence beyond is from the Arabidopsis AT5G4340 gene.

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Figure 4.2

-2783 CCTGAAAACAATATCCAACAAACCAAGAAGTTTATTTCGTGCAACAATTTCTACTCTCTTTTCATTGCCATTTA -2709 TAATAAACCATATTCATCATTGGTTGCTGCAAAAAACTGTAACTCAAATCTTACACTACTTAAGAAAAAATTAA -2635 TAATGCAACGCAATTATATTAAACTCCAATGGAGATATAAATTGGCTTCGAAATTGAGGTAATTGCGATCAACT -2561 CTTGAAATTAGGGTTACATTTAGGAAATAAAGGTGAAGTATCTATACTATGAAAATCAATACAGCCATGGGAAA -2487 AGGAAATACCAACACGAGGGAGATCGAAGAAGATTGCTTGGAACGAATTAATGGGAGCCAACAAGGGATTGAAG -2413 CCGTTTTTCAAAGTCTGGATTTTTGTTTAACAAAGGGAAGAAGATGAGAAAAATCGAAGAGGAAGCCACTGAAT -2339 TGGGCCTTCAACTGGGTCGAATAGTTTAGATTAGTTAGTTAATTACTATCCAAATTTTCACTATTTCAGAGAAG -2265 ATTCGAATTTTGGATCAGAAATATTGGATTATTTTGGGAAATTTTAAGTATTATATTTGTTGTGTTGTATATTG -2191 TTGGGCCAATGGGCCTACTTTAGATATCAACGATTCAAAAGTTAAATACACGAAGATCTAACACTTTCATTCAA -2117 TTGGAGGTTTCTACACTTCCCGTAAAAAACATAAAGATATTTTTCCATATTTTTCAGTTAAGCTGTGGTCCGGA -2043 TTTTTTTCTTTTTTTGAGTTTAATGCTAATATCCGATAATTCTAAATCTGGAAAAACCCACAAAAACTCGCCAT -1969 TAACTTACGTTTTGATGAACCGGTTAAACTGGATGGATGTGGATTTTAAAAAAGATATTTGATTTTAGAATAAA -1895 ATATTGTATTTTAAACTTTATATAGTTATTTTTACAAATTAATTTACTTTTTAAACTCATTTGATGTTTTAGAT -1821 TTTGGTAAAAACTTGACAATGTCGTCTTAAAAGAATGAATGTAATGGTGAGAGAGACAACCGATATTAGTTGTT -1747 CAAATTCGCAATGTTTTTTTAGTTTATTTTAGACATTGGTGATTTTCTACTATATTTGTTACGGTTTTAAACTT -1673 TAAAGTTGGATTTAAAAATTTTAGTCACTTTTTTTTATTGTTTAATTTCATATTTTGTTAAACATGATATGATT -1599 GTTTTTGTTTTTTTTTTATATATTTATAAAACCTATATCATCTCTATTAATATTAAATCTATTACATAAATTTA -1525 AACACCCAAAAATCATCTTGTTTTTTGTTTTCCATTTCTAAACTCTTTCTTTTATTCCAACAAGCATAAAAACC -1451 TAAAAATATAAAAAAAATCACCAAAAATCATCTTGTTTTTACTTAAAAGGTTCAAATTTCGGTATTAAAAAAAT -1377 TAATAAACTGATGAAAATATAAACTCTATATGTTAAAATACTTGCATTGAACTTATTGAAGAAAAATGTTTCAA -1303 CTGTTTAATTAACCCAATTAGGATGTTGAAAACTAAACAAAAAAAATAACAAAGATTTTATGGTGTTGAATCTA -1229 TTTAACATACTATTTTTTGTTTTTTGGTTCGCACATGTCATGTGTCACCATTTTATTCTGTTTGTTTTTTTTTG -1155 GTCAAAAACAATAATTATTGAAAAATTGATACTTCTCAAGTCTTCAAATACTTGTTTTCCATCTTTCTTTCTTT -1081 TTAATATCATTTTCGTCATTTTAGTGCCAATAACCTGAATATAATTCTTCAGAGTAGAAATACTTATAAACACT -1007 CGAGACCATATAAAATGTTTTCCAAAACCATATAGAAATCATAGTGAAAACAAATAAAAATAAAATGTTAAATA -933 AAATCAAAAGATAAAATTATTTATTAATCATCTCCGATTTGGCAACTGCAAGAACCACACAAACCTCTTGAAAC -859 CATGAAACTTTTCAAATTGGAATATCATTTCAAATTTTGTTTGTACAATTACTAAACCACGCATGCAAGTTTAT -785 CATACATTCATACTAGTATTTGAAGGTCATATTTTCTGCAGCAAAATTTAAGTTGAATGAATTAGTATCCAAAC -711 ATACAATTCTGTGAATCTGAGTTTTAAATCAATTTTTAATACATAGGAAAGTAATAAATCAAATAATTGTTTTA -637 GTAATATGCTAAATAACTGTACTTTCATTATGTGAAATACATCAACTCTCACCAAAAAAAAAAGGAATTACATC -563 GATTATTATACTATTTTCATTTTTATAATAGAATAAAATTTAATATTTAATGACCGATTGTAAAATTATTCTAC -489 GCTTCGAGACATGTGGTTACAGCTATCACCCATTAATATACAATATCAACTTGTTTAATCTCGGTTGATTATAA -415 TTAATATACTCTTAAAATTTCGTTGACATTAGGGGACAGCACGAGAGCATAGTTGTTATCAGATCCAATAAAAC -341 TTAACATACAGCTAGAAATACTGAGCTAGAATTAAATATGATGACCAACTGTAAAATATATTCTTCAATAGATA -267 CTTTTAAAATTATACTAGTGTTTAGGGACATGGTTACAGCTTTCACCCATTTATATACTTGTTCAAAATTAATA -193 GACTCTTAAATTTTCGTTGACTTAAATTTTCGTTGGCCTAAATTTTCGTTGGCCTAAATTTTAAAAGCTGAAGG -119 AAATTGAGTATTAATGAATCACCACATGACTTTGCCTTTATCTTTATATATATAAGCACAATGGTTGTGGCAAA -45 TCAATACATATCCTCACCCCCAAAGTTTTTTGATATCTACCATTAATGGAAGAGATAGGTGAAACACATGAGAG +30 TGCTGATGGTGGAGTTCTGTCTCCGTTTCAC

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Figure 4.3 E40 promoter sequence. The promoter sequence starts at the 8324097 bp of the 5th chromosome of Arabidopsis and ends at 8320146 bp. The Transcription Start Site

(http://linux1.softberry.com/berry. phtml?topic= tssp&group=programs&subgroup=promoter)

The promoter sequence is numbered according to the start codon position and the start codon position and sequence beyond is from the Arabidopsis AT5G24380 gene.

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Figure 4.3

-3972 ATGGCTCACTATATGAATATCCCGAAACCCGAAATTTTATCCAAAAATCCGGATATTTACCCAAAAAATCTGGA -3898 TATTTACCCAAAAATTCAGATATCGGGCCTGGAAATCTATATCCGAACCGATCCGAACCGAAACCAAACCGATA -3824 TTTTAGATTTACCCTATTGGGTCCTAAACTTCTCAATCTGAAAGACCCGTACCCGATTGGGTTTTACCCGAACC -3750 CGACCCGCTTACCCAAATTCCCACCCCTATTGAAAATCATACATATATTTATGGTTTTTTCTAACTAACTTACT -3676 ATTTTTTTTTTTGATGTTGTTTATCTAGCAAACCAGCATAACCATTTAAAAATAATGTTTTAAAGCTATATAAT -3602 GATTACAAATGAAAAATAATGTTTTATCTAACAAATGAAAATAATGATTACTCCACATAACATTTTTCTTAAAC -3528 TAATAAAAGTTTTAAGCATATATTTGTAGTTAATACTTAAAGTTTGTCATATGTTGTCACTCTTGTTTTTTAGG -3454 GTTCTCTATAAGATCAATGCAGGAAAAAATTGGATGTTTCATTTAACAAAATGGCCGTTTAGTTATAAAATTTG -3380 CTCCTTTAGTTTCCTTTAAGCTAAGATACTATAGTTATTGATTTCGATTTAGCTTACAAACTTTGCTCCTTTTA -3306 TTTCCTTGAAGGTATATATACATAGTTATTGATTTCGATTTAGCTTACAAAATTTGCTCCTTTTATTTCCTTTA -3232 AGATACATATACATAATTAATGGTTTCGATTTGGTATAGTAATCTGAGTATAAATTTATATGCTTTTGTACGTC -3158 TTCGTAAAGCATTTACATCACACACCGAAATGGCTCACTATATGAATATGAATCTGATCATCACTAGTCTTGTG -3084 ATGATCTTGATTTCACATTTGCCTTTTTCCTCAACTGTGAATCCAGACAAACCAACAAAGTCTATAATCGACAG -3010 AATCTGCAACCAAACAGTGAACTTCAATGAGTGTGAATTGATTGTCACGTCACAGCTACACTCGCCGCACGCAA -2936 ATATAGCGACCATAACGGAAGTGATGACAAAAAGAGCTCTGACGTTTGCTACGGAGACGGTTTCTCAAATCCAA -2862 GACTATCTTCTCCCAAACGCTACAGATACTCAGGACAAAGCCGTTTTCTCCGCTTGCGAGATCGCGTACAAAGC -2788 CGTGGTTTCTCGTCTTCAGAGTGCTTATACGTCGATGAAGAACAGTGATTACGTGGCGATGAAGGCACAGCAGA -2714 ATCAAGCACTTGGGTATATTGATGTGTGTGTTGAGAGGACTAACTTCTTTCGCCGGACTCCGATGGTAGCAGCC -2640 AACTATTATGTGAGGTTGACGGCGAAGATTGCTTCCATTGCAGGCCAGATCCTTGCTCCTAGTACGACCACGTC -2566 CCTGTCCCCTTGAACAAAATATTATTTTGTATTTTTGTTAGAGAAAAACTCGAGCTTTGATTCATCAATAATTA -2492 TTTATTTATTTATTAATTTTCGTTGTCTTTTCTCATAGAAACAAGACAAACCACAAAATTTAGCATTTCAGAAA -2418 TTTGTAGAATTCAGTGAATTTTGGAAATTTTCTCACAAGAGATTCGTAGAATTCAGTCCACTAAGTTTTCTATG -2344 AATGATGTGGTTATTAAGATAAGATTTGTTGTGTGCAAGAAAGTGAGTTACCGAACAAACAGCCGTCACTTACA -2270 CCAAAAAATTCAATATATATCATATATGGTGTAAGAAATTACACACTCCCAAAGTCCAGCTTGTGACGAGAGGT -2196 GCCTTTGAACTTATAAACCATCTGTTATTCTTTTTTCTTTCCAATGGTCTTAGGCCCGGTCAAATAAACGAACC -2122 AAATGTACCGAATCAAAACCAAAACAAAAATAATCGAATCGAATCAAATCGAAAATTGAAAAATAACTGAATGG -2048 TTACTAAATTTTATAACCAAAAGAACCGAAACCGAATGATTATGTATTGAAATATTTCAGGTAACCAAAAATAT -1974 CTCAAATATAAATATATTTTCAAAAATATTAGTTATTTCTAGACTAAATAATTAAAAATGGTTGAAAAAAATAT -1900 ATATTATAGTAAAAATATTCAAAAATAACCAAAAATATCCAAAATATCTGAAAATATTCAAAACAAAATAACCG -1826 AATAGATACTAAATTTTAAAATCGAAAGAACTGGAACCGAACCGAATCGAACCGAACTTTCAAAATAACCGAAT -1752 GGATACTAAAGTACCGAAAAAACCGATAACCGAATAAATACAACCGAAACCAAATGGATAACCGAACGTACAAG -1678 AATATGTGATCTTCTTTTAACATTTACACCCAAAAAAAATGTGTGATCTTATTTCCTCCAAATGAAATATTATT -1604 ATTTAGGCCCTTATGAATACCCGCCATTTTGAGCTCGGACGTCCACGTCGGTCTAATCCAATTGCCTACGTTAC -1530 GCGGTTCTTTCTGCGTGAATTATGTCACTCTTTCTCTTTTTTGGGGTGTTCATTAGAGCAGCACCAATGGTGGT -1456 CTCTCTTATGAGTCCTTTATCATAAAAATAAAGAAAAATAATTAAAAAGAGAGGGACAGAAGAGAGAGGAGCTC -1382 CAAAAACCCTTATTTAAGGGACAGTCTCAACCCCAAATATACACGTGTAGTTCTTTTAGTAGTCAAACTCTATT -1308 AAAAAACTAAAATATAATAACAACATTAAAATATTATTATTATTTTGGTTTTTAGGGACTCATAAGAGAGACCA -1234 CCATTGCTGCTCTTAGTACACTAGACAAACAAATATACTCTATTTCAAAGTAATACTTATCTAGAACTATGTAT -1160 ACAATTATTACATATTGCAATACGATAAAAAAGAGTATCATATCTAGAAAAGTGGAATGGTCTATACGTTTGTT

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Figure 4.3 continued.

-1086 GATGCTTAAAGTAAAATTTTAATTTATGACATAATTTGGTAGAATTCTCATCTGAATATATAATAGTAGTTGTT -1012 AGAAATTAATTAACTTGAAGGTAAAGTGTGCAAGTCATAATTTTATGTTACCTCCTAACTTACACTTCCCAAAA -938 TAAATTACATTCGTAAAGTTTCGTATAATACGTAATCCATACGAATCAGACCAAAAAAAAAACCAAAACTCAAC -864 TGTGAAAAAAATATTTTTGATAATAAATATTGTTGTTGTGTAACAGTGTTTTTTTATATATATATAACAAATGG -790 ATGGATTATGGAATAATTTTAACAGAATATATGGGTCAAACTTTTAATATTATGGGAATAGAAGCTGACAACAC -716 ACACATCAAAAAGCTGGGACACATCAGGTAGTAACTGAAGTCAAACATAGACACCAAACTCAACTAAACCCACT -642 CTTTTTCAAAAGTCAGTGCTCTTTCTCTTTGTGGGGTTTGTTGTTTACTATCGACACTTTCAAGGGACTCCATT -568 TTTTTCCTATTTTTATAATTATTACCAACACCAACTTCACAATATTTGTTCTGTTTTTGTTAATATACAACTTT -494 ACCGGATTTGTACTACCATCTTCTTATTATTCTTTTTTTCGGATCATATATTAAAATATAAAAAATTGCACGGT -420 TAGTGAGTTTCAGACCAACAACCATATATCAACCCGCTCTGTCTTTTATCTGAAAAACGTGTGTTTTTTCCACA -346 TTTGGTTGAATCTGAATCACATAATATATGTACTCTCAATTACAATGCAAACCCTAATTTGTACTTCTGTGCAG -272 TACACTTTCTCTCTCTCTTCATTCATTTCTCTTCTTCTATAAAAAAAAATTCTTCAATTCAATATCACTAATAT -198 GCGATTCTCTCTTCTTCTTCTTCTTCTTCCATCTTACGCCAAACCTCGAGTCTGCGAATCATTCGATTCTCAGA -124 TTCGCATATTTTCTTCGAATTCGCTAGGCTCAAAACGTTGTTTCCGTGTACAAGAAGCAACCTAACGATTTGGG -50 GCCAGGTGAAGGAAGAGCTGATTTAGGTTATTGAGATTCGTTTTCTTCAAATGGAAAACGAAAGGGTTGAGAGA +25 GAACAGAGCCAATTTCAGGAAGACGAGTTTATCGAT

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The E49 promoter (Fig. 4.4) and S8 promoter (Fig. 4.5) regulate the expression of a putative sugar transporter protein (Marshall et al. 2003 and Price et al. 2004) and a Dof1 type zinc finger domain containing protein (Yanagisawa et al. 2002), respectively. The expression pattern conferred by E49 is in the cortex region of the Arabidopsis root and for S8 it is in the pericycle in the phloem.

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Figure 4.4 E49 promoter sequence. The promoter sequence starts at the 1440215 bp of the 3rd chromosome of Arabidopsis and ends at 1437802 bp. The Transcription Start Site

(http://linux1.softberry.com/berry.phtml?topic=tssp&group=programs&subgroup=promoter).

The promoter sequence is numbered according to the start codon position and the start codon position and sequence beyond is from the Arabidopsis AT3G05150 gene.

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Figure 4.4

-2414 AGAAACGCGTTGGTTCTTTTATGTTTTGACGTATGATTTATGTCGTTTGGATCTTCTTCAGTGACATAACTTGA -2340 ATATTTAGAGTTGTACCTGCCTCGGGAGATTCAAAAGAAAAGAACAGACTAAACATGTCGTTCTTATGTAAATT -2266 TTGCGGGAAATTAAGAGAAAATCGGTGAAAATTCAGGGAAAGTTCGTTAATTCTGCTTAAACGTTCGTTAAGTA -2192 TCAATGAAAATCATAACAGAACAGAAGATAAACTTCTAGACATTAATATCATGTCTACCAAAAAAAAACTCAAA -2118 TCAAATATAAGAACAAACAGAGAAAAAAGTGATTTGATTGAATGATTACATTGGAAGAGCACAACACACGGGAG -2044 AAGCTTCTTTGCTTTGGTCTACCGCCATGAAAGAGTGTTGGTAGGACACTTTTAAGATTTTCTCAGAAACCTGA -1970 AAACCTAAAAATAGAGAAGAGGATCATAGAAAGGAAATAGAAAGAAGTTAACATTTAAACCAAAAATAAATAAA -1896 AATAGAAAGAAGTTTTGAAGTTGGTGTTCGTCTCGAGACACCCGAAAGAAAAGGTTTTGTCTGTGTGTTTAAAA -1822 TTTGATTTACTAATTTAATTGTTTTCCGTTGACCAATGGTATATCTACTAGTAGTTTTTTTGTTAAGCTAATAG -1748 TTAGACTCTAAAATGACTAAATCCTATATTTGGCAATATGCTTTTGATTTTCTCTTTTTGGATTGAACTCAGAG -1674 GCATTCTCTTAGCTTCATTCATTTATGGCAGAAAGGGTTAAGGGTCCTTAAAAATCGCTTCATCTGTCGGTCTC -1600 GTTATCCAAACTCATATTCCTAATTTAGGTCTTTTAAGGTTTTAAAAAACTACTTTCTTTATCTTGTTGATAAG -1526 GAATTTATAAAAGATCAAACTTGAGTAATCTAAGTAGTTTTAGTTTTACTAATTTCATTTACTGTTACTAATCA -1452 ATTGTAGATATTGACATTCTTGTATATATTGCGACCTGATAGTAATTATCTTTAATTTTCTATATGTTTTTATT -1378 CAAGTAGTACTCTGTTTTGTATTTTCTTTCTTGTGAATAGAATATTTTCTGTGTGTATATGATGCAGAAAGCCT -1304 GCATTAAAACCTGCATTGGCTTTATTGCTTTGCACCGTTTATTTATTTTTATGATTCTTATTTTTGTGATTAAC -1230 AAATATTATTTCAAGTGACTCTTATATAAGTTTTGGATAGTTAAACCACGGACCACAATCATGTTGGAAGTGAT -1156 TGTTCAATACTTCATTGTTGTATAAGAGAGGGGACATCGATCATTTATGTAAATCTTGTTAGGTACTGTTCTGT -1082 TCTATCAATCTATCAATAGTCCCTACGCTAACTTTCTGTACCCTCGATTAAATAGGTTTTTGCTCAATGACGCT -1008 GGGTCCAAATTGTAATGACTCAGTTCAACTTCCGCGACAAAATCATTTTCATAGATTTTGATGATTTCTATAGT -934 GGGCGGTTTTTTCTATACATCTTACATTCGCAAGAATTTGAATGATATTAGCCCATATGACGAGAAGAAGAATA -860 ATTCATTACTTATGTGGTTAATGCGGAGTGGAAACGAATATCTCATGGCCAACTACAAATCCAAATACTGTCTT -786 TTGAATCCCTGTAGTTTTATACTATATGATCTAGAGTTTTTTTTGGATTTTGTCTCCCAAATTAGAAATTATTA -712 TAGTATCAACTATCCATCACATTTATAGAAAAAAAAAAAAAAAAAAACTATCCATCACAGCTTGGATCTTGCGG -638 TGGGCCAGGAAAAAACACATGAGCATCGGATCTGTGGAAAACAACTGTCTTATCGGGTCAAACCTTTGAGTCAA -564 TTACACGTTACTGTGCATGTCATAAACCATCACCAAAAAGGAAAACAAAATAACAAACCATGCATCTTACTTAC -490 CCAGTAACACAATGCATCCATAATGTACATAGTATTTGTGAATTTTTTGAGGAAAGTATTAGTGAATATTATTA -416 CTATGAAATACCATGTAATTACTTCTGAATTATCTTACTTAAGAAAATTCAAGTTTAGTCAATCAACATTAAAA -342 CTATAGAGAACCCCCAACGTGTAATAATGCATTGGGAGTTGGGACTTTCATAAAATCTCGAATAAATTAATTCA -268 AATTGTCTAATTCGTCCAAAATTATCCTGCATGGGTTTTAGCTAAGCCAGCCTAGCTACATGTACTGATTTATA -194 CAGAGATAAATATAGACATGATATATATCAATAAATTAGGAACAAATCAGAGTTTTTAGATCTAAAAACACGGT -120 GCCTTCTCATTTCCTCCGGTGAGTTATCCAAATCTTGTGTTTCTTATCATATATCCTATATTTTATCTACATAA -46 AAATTGATAAGTATTCGTTGAATTCAAAGGGCATTTTACGAAGAAAATGGAGACAAGAAAAGACGACATGGAGA +29 AAAGGAACGACAAATCCGAGCCGTTGCTGCTG

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Figure 4.5 S8 promoter sequence. The promoter sequence starts at the 24241077 bp of the 5th chromosome of Arabidopsis and ends at 24237996 bp. the Transcription Start Site

(http://linux1.softberry.com/berry.phtml?topic=tssp&group=programs&subgroup=promoter).

The promoter sequence is numbered according to the start codon position and the start codon position and sequence beyond is from the Arabidopsis AT5G60200 gene.

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Figure 4.5

-3082 CCGAATTAGCATTAGGTCGAATAGGAATTTTCTAAAAGTGTTATATTGGTGTAAACAATATAACACTTACAGTA -3008 ACGATCAACAAAAATTACAGTAACGATCACTCTATAAACTATAATAAAAATGACTAGTACCATCATGTCGAGAT -2934 AGATTGCAATATGATTCATATATATGATGACATGAGAGAGAGCTCTCTTATCTATACGATCTGCTTTAACTTCT -2860 TTTTATTGCCCATGAAATTCTAGTTTTGTCACTTGTTGTCTTAAGACACATTATAAAGCAACTGAAAATATTAT -2786 AACTTTGACTGGTCAACCCAAAACTCAATGTCTTTTTCTTCCATTCCTATTTTCCAGTGGTCCCACCACTTCAG -2712 TTATTATAATGTTCGATTCCTGAATTTATGATTTTGAATGGAAGATTCGTGTTATGGATCACACTCTCATTCCT -2638 GTTTCTTCATCTCCACTTCTTTCGAAAATTGATCTATGTTAAGATTTATAAATTAGTTTGGTATTAACCTGTAT -2564 ACATTTGTCTTTTAAACCTGCCAATATAATTCTTTTGAATTACGTTCTTGCCTGTTTAAAAGTTTGGTGGTCAT -2490 CAAATTTTTGGCTTAATATCGTTTCGATTTGGTATGAATTGAATAGAATAAACCAAGTCATTGGATGATGTACA -2416 ATTAGGTTTCTCCATTTCGCTATTAGTAAACTAAATGATTAAAAGGTGTCTGAACTGTTAGTGCTAAGAAAAAG -2342 ACTTAGTGCTATAATATTTTTAATAAACACAAATCATTTACATAAATTGAACTTAAAATTTAAAGATGGCCATC -2268 ATTATTGGGATAAAATTTGGGAGCCACAAGTAGGTAGGGTCCAAAGGGAAAATATCCATATAATGATCATCATC -2194 AGTTATATTTGGAGACCAATGATTGTTTGTGAATGGATCACTAAATTGTTGTAAAAAAGAGAAAGAACGAATGA -2120 ATCACTATTTTCCTGGCGGGAATTGGTAATAACAAAATGATTTTTTTTTTTTTTCCATCCTATCAAACATGAAA -2046 CGAAAGAAAGAGTATAGTAAGTGGGATTTATTGAATGATTGATCCGTCTCTAGCTACTTCCATTGTCACATTAA -1972 CGTTTTCGTTTTCGTAGTAAACAAAGAAACATGTTTTCAACTAAGTAGTAAGACAGAAGAAAAAAATTCTAGAG -1898 GAACAAGTAACGATCTTATTATGATATATACTAACATGGGTTACTCATTTTTTTTCTCCACGACACAAGATTTA -1824 ACATAAATCGTATACTATATAAAGATTACGTAAACAATAATATAAAATATACTGTATAATGTTTTGTGTGGATA -1750 TGGTCCTTAGACCAAATCTCGGTCTTTCATAGAAGTCTCTTTCCCGAAAACCAATGTGGTAGTGTGATTTGTGT -1676 CTCTATGAACTTTAGATCCCCATCAGATAATGCGAGCACATCATGAGCTACACATTGTCCGGCATATTCTTTGC -1602 TTTTACTTCATCATGTGTTCTATTTACATATATACTTATGATTATTTATAAAACAATGTAAGCGTATATTATAT -1528 CCACAATTTTTTAATTTAGATTTCAGAGTAGAATTGTAGTTTTATTCAACAAATTATATAAAGTATAATAGAGT -1454 ATAAATTTAGAATGAACAAAAAATTGTAGCCCAATATGTCTTGACAAAAATAGAAACATTTGTGGTCCTATCAA -1380 ATAAAGAGTGTTGGTCCAAGATGATTTTTTTTTCTTTTTGACAGCAATGACATTATATTTGCTTGATTTTGAAA -1306 TATATGTGTTTAAAATTAAGGCAATGGGAATATGGAGGACAAATTTAAAATCTCTTTGGTAAAATTTTCGATAA -1232 AAGAAACAAGACATTATCGAATATTTTTTATGCGCTTTACAATTCACATAATCCTATCAAATCACATAAAGAAA -1158 CCAACTCGACAAATACAAATTTTAGTATTAAAACACTTGATTTTGCTTTAAACATCGAATTAGAATGTACTCAT -1084 ATATGACTGATCGAAATAGAAAATCACATATATTTGTTCAATCCTCTAGAGTACGAAGGAACGGTACGTTAGGG -1010 TCCACAAGGAAGAACAAGACATTGCGATGAGTATGGGACCCTGGTGGTGACAACTCAACGGACCATTGAGACAG -936 CAACACAGAAATGACAAAGAAGTTGAGTGCGTGGGAATCCGACGTCGAACATTATCTCTCATCCATCATCGTAC -862 GGTTACTATTCATCTTAATCAAATAGATCCGATTTCACATTATTGATAACGATTGCAAGTTTGCAACTCAACAT -788 TTTTTTATTTTTTTTTATCGCTTGAAGAAATTGACCGATACATTATTGTGGAATGTGGAGTGACGAAGCAAGCT -714 TAGTACATATTATATCAGACTATAGTTATAGATTTTAAGCTACAAAGGTTTTATATAGATCTCAAGTTTTTGAT -640 GTGGTTATAGGTTTATAGCTTTACTCATAACACTACAATTGAGTAGTAAGTTTAGTTGTTTTTATGAGATTATA -566 TAGTAAAAAGATTTGGTAAATGTTATATAAAAAAAATTATGAGAATTAATTTACACTAGAAAATATATTTTAAA -492 ACTTACTTCATTTTCAAATTTTAGATATCAAATCATACAAAATGTTGTTTTATGAAACTAATTAATATATAACC -418 TTGTTTGAAGTAGCAATTTTTAACAAGTCTTCAAACATTAATATATGAAGAGATAATAACATTATTAGGTTATA -344 CTAATTTCCACAGGCTATCAAATGTAGATTTTTTGTTCGCATTATAAATTTTTTAATAAACGGCGGCTTTATGT -270 CTGTTCCAAGTCACGTTCCGAATTGGTTCACATTATTCTTCTTAGCATTGTTTGAAATATTAAACTAATAATAT -196 GAGATCGAGGCCACAACAACAATTGTGTGTCTCTCTCCTCCCACATTCCCATCCCATGTATATATACAGATGCT -122 TGGCTTTTCCTTTTGCTTCACTTTTAATTTCTACTCATATCCATCATTCTCTCTGATCTCAAGAAATAATCTCA -48 AATCTCTTACACTCATAGTCTTCAAGCAACTTCTCAGATTCAGCTCTTATGGATCATTTGTTACAACACCAGGT +27 AAACTACAATTCAAGATCCACAGTATCTAACATC

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Our goal was to determine the functionality and pattern of expression of these heterologous promoters in cassava. While cassava leaves and fibrous roots have typical tissue distribution the cassava storage root can be divided into the peel and the pulp. The pulp consists of two main tissue systems, TSII (phloem and vascular cambium) and TSIII

(starchy parenchyma cells and xylem) (de Souza et al. 2001). It is the pulp that is consumed by humans and processed for industrial uses such as additives and biofuels.

The peel is formed by only one main tissue system, TSI (epidermis and cortical parenchyma cells) and in difference to the pulp it contains no starchy parenchyma. The peel, however, does contain higher levels of cyanogenic glucosides in relation to the pulp.

Previous analyses have revealed that linamarin concentrations in the peel range from 1.7 fold that of the pulp (Dufour, 1988) to 17 fold (Santana et al. 2002) depending on the variety and age of the root. Given the differences in composition and uses of the root pulp and peel it is of particular interest to obtain tissue specific promoters to better utilize their properties and so these two tissues from the storage roots where evaluated separately.

To evaluate efficiently all promoters a system compatible with the Gateway Cloning

System (Invitrogen) was developed. In the designed double expression vector the 35S promoter was fused to the Yellow Fluorescent Protein (YFP) and any one of the four promoters controlled the expression of the Cyan Fluorescent Protein (CFP). The

Arabidopsis promoters were introduced in to each of their vectors by Gateway recombination, in this manner this system may be used in the future to evaluate other promoters of interest and to compare their expression levels to that of the 35S promoter.

The YFP and CFP reporter genes were chosen because in addition to evaluating their

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transcript accumulation levels their protein accumulation levels may be evaluated due to the fluorescent properties of their proteins (Shaner et al., 2007). Therefore, providing the opportunity for different analysis techniques for promoter expression. In addition, the two reporter genes were codon optimized for cassava usage. The importance of codon optimization of reporter genes has been reported previously by Rouwendal et al. (1997) with the green fluorescent protein (GFP) gene optimized for expression in tobacco and more recently by Sattarzadeh et al. (2010) who reported the optimization of blue fluorescent protein, GFP and YFP for expression in maize, and Sastalla et al. (2009) who reported optimization of CFP and YFP for expression in gram-positive bacteria.

In this chapter, we demonstrate the functionality of two new promoters for cassava storage root expression and the effectiveness of using a Gateway Cloning System based strategy to evaluate multiple promoters in cassava. The A14 promoter had activity in cassava storage roots more specifically has increased expression in the storage root peel.

In addition, the E40 promoter shows strong expression in the pulp of the storage roots.

Both the A14 promoter and the E40 promoter have limited activity in the leaves which is ideal for cassava roots biofortification. The two remaining promoters, E49 and S8, did not show significant expression in leaves or storage roots of greenhouse grown plants, however, in young plantlets the E49 promoter had fibrous root specific expression and the S8 promoter had leaf-specific activity. To our knowledge this is the first report where a variety of promoters were analyzed for cassava storage root expression with direct comparison to a constitutive promoter and the first to present tissue specific promoters.

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4.2 MATERIALS AND METHODS

4.2.1 E.coli and Agrobacterium strain

Two Escherichia coli strains were used for gene cloning, the DH5ɑ (Invitrogen,

Carlsbad, CA,USA) was used for all Gateway Cloning reactions and the OneShot TOP10 strain (Invitrogen) was used with the pCR8/GW/TOPO® TA Cloning Kit.

Agrobacterium tumefaciens strain LBA4404 (Invitrogen) was used for cassava transformation. E.coli cells were grown in Luria-Bertani (LB) media and Agrobacterium cells were grown in YM media at 28 ˚C. Agrobacterium cells growing for transformation of friable embryogenic callus were grown on YM media at 28 ˚C to an OD600 of 0.7.

4.2.2 Chemical Reagents

The chemical reagents used in all experimental procedures were purchased from Fisher

Scientific, Hampton, NH unless otherwise noted. All plastic products such as microfuge tubes and PCR tubes were purchased from USA Scientific, Ocala, Florida.

4.2.3 Cassava cultivar and in vitro and greenhouse growth

The cassava cultivar TMS60444 was used for genetic transformation. Cassava plantlets were grown in vitro on Murashige and Skoog media (1962) supplemented with 20 g/L sucrose, MS Vitamins (Sigma Aldrich, USA) and 2.4 g/L of the gelling agent phytagel, this media will be subsequently known as MS2. The growth chamber conditions were set at a temperature of 28 ˚C, a light intensity of 75 to 100 µmol photons m-2 s-2 and a 16hr day/8 hr night cycle.

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Six week old in vitro grown cassava plantlets were then transferred to the greenhouse.

They were potted in 4” by 4” in pots with Fafard 51 soil mix (Fafard, Agawam, MS) and supplemented with 15:16:17 (N:P:K) fertilizer or Jack’s 10:30:20 (JR Peters, Allentown,

PA) twice a week. At 4 weeks after initial transfer to soil the plantlets were transferred to

6” by 6” pots.

4.2.4 Bioinformatic analysis of the Arabidopsis promoters

The sequences of the four Arabidopsis promoters were obtained courtesy of Dr. Philip

Benfey (Duke University, North Carolina, USA). The Transcription Start Site (TSS) and

TATA boxes of each promoter were predicted using the SoftBerry TSSP/ Prediction of

PLANT Promoters tool (http://linux1.softberry.com/berry.phtml? topic=tssp&group= programs &subgroup=promoter) (Solovyev and Shahmuradov 2003). The sequence of each of the promoters was analyzed using the database of Plant Cis-acting Regulatory DNA

Elements (PLACE) (www.dna.affrc.go.jp/PLACE) (Higo et al. 1999).

4.2.5 Arabidopsis promoters and entry vector

The four Arabidopsis root-expressed promoters were obtained courtesy of Dr. Philip

Benfey (Duke University, North Carolina, USA), each promoter came in a modified pGreen II 0229 vector (Lee et al. 2006). Each promoter was amplified using the individual pGreen plasmids as templates and the Platinum© Taq DNA Polymerase High

Fidelity enzyme (Invitrogen), primers used to amplify each promoter are found below,

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A14 F: CCTGAAAACAATATCCAACAAACCAAG

A14 R: AAGCCTGCTTTTTTGTACAAACTTGTAATG

E40 F: ATGGCTCACTATATGAATATCCCGAAAC

E40 R: GCTTTTTTGTACAAACTTGTTGAAGAAAAC

E49 F: AGAAACGCGTTGGTTCTTTTATGTTTTG

E49 R: TTTCTTCGTAAAATGCCCTTTGAATTC

S8 F: CCGAATTAGCATTAGGTCGAATAGG

S8 R: AAGAGCTGAATCTGAGAAGTTGCTTG

The amplification reaction for the individual promoters was set up with 15ng of plasmid

DNA (corresponding pGreen vector for each promoter), one unit of the enzyme Platinum

Taq High Fidelity DNA Polymerase (Invitrogen, Carlsbad, CA), 10 µL of the supplied

10X High Fidelity PCR Buffer, 2 µL of 50 mM MgSO4, 2 µL of 10 mM dNTP mixture,

2.5 µL of 10µM solution of each primer and autoclaved double distilled water to a volume of 50 µL. The PCR reactions were performed in a BioRad Mycycler (Bio-Rad,

Hercules, CA) under the following conditions: one cycle of 5 min at 94˚C followed by 32 cycles of amplification (30s at 94˚C, 30s at 53˚C and 3min or 4min at 72˚C for extension) and a final cycle of 5min at 72˚C. The extension time used for the PCR reactions of the

A14, E49 and S8 promoter was 3 min and for E40 it was 4 min. The resulting reaction

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products were separated on 1% (w/w) agarose gel in TAE buffer (40mM Tris-acetate,

1mM EDTA). The correctly sized amplified bands were gel purified from the agarose gel using the QiaQuick Gel Extraction kit (Qiagen, Valencia, CA, USA). From the purified fresh PCR product 3 µL were taken for TOPO® Cloning. The TOPO® Cloning reaction was set up according to the manufacturers recommendations found in the pCR8/GW/TOPO® TA Cloning kit using 3 µL fresh PCR product plus 1 µL of Salt

Solution (200mM NaCl and 10mM MgCl2), 1 µL of autoclaved deionized H2O and 1 µL of linear pCR8/GW/TOPO®. The mix was incubated at room temperature for 5 min after which it was added to a thawed vial of OneShot TOP10 (Invitrogen) cells and mixed gently. The cells were incubated on ice for 30 min and immediately submitted to a heat shock treatment at 42˚C for 30 s. Next, a volume of 250 µL of SOC media (2% (w/v) tryptone, 0.5% (w/v) yeast extract, 10 mM NaCl and 2.5 mM KCl) was added and the cells where further incubated for 1 more hour at 37 ˚C. The cells were spread on pre- warmed LB plates containing 100 µg/mL of spectinomycin and incubated overnight. The following day 5 colonies were picked from each plate and grown overnight in 5 mL of

LB media at 37˚C. Using the Qiaprep Spin Miniprep Kit (Qiagen) the plasmid product was purified and used as a template for PCR amplification. The previously noted primers were used to amplify their respective promoters, the reactions were set up as follows:

15ng of plasmid DNA (corresponding pGreen vector for each promoter), one unit of the enzyme Platinum Taq High Fidelity DNA Polymerase (Invitrogen, Carlsbad, CA), 10 µL of the supplied 10X High Fidelity PCR Buffer, 2 µL of 50 mM MgSO4, 2 µL of 10 mM dNTP mixture, 2.5µL of 10µM solution of each primer and autoclaved double distilled water to a volume of 50 µL. The PCR reactions were performed in a BioRad Mycycler

288

(Bio-Rad, Hercules, CA) under the following conditions: one cycle of 5 min at 94˚C followed by 32 cycles of amplification (30s at 94˚C, 30s at 53˚C and 3 min or 4 min at

72˚C for extension) and a final cycle of 5 min at 72˚C. The extension time used for the

PCR reactions of the A14, E49 and S8 promoter was 3min and for E40 it was 4 min. The products were fractioned on 1% (w/w) agarose gel in TAE buffer and gel purified using the QiaQuick Gel Extraction Kit (Qiagen). The purified promoter PCR products were sequenced in both directions at the Ohio State University Plant-Microbe Genomics facility. The primers A14F, A14R, E40F, E40R, E49F, E49R, S8F and S8R along with two more internal primers for each promoter were used in the sequencing reactions. The internal primers used for sequencing were,

A14 Internal Forward Primer: TTTCTACACTTCCCGTAAAA

A14 Internal Reverse Primer: GGTTGTCTCTCTCACCATTACATTCATTC

E40 Internal Forward Primer: CTCCTTTTATTTCCTTGAAG

E40 Internal Reverse Primer: AATTCACACTCATTGAAGTTCACTGTTTGG

E49 Internal Forward Primer: GACTAAATCCTATATTTGGC

E49 Internal Reverse Primer: CTATCAGGTCGCAATATATACAAGAATGTC

S8 Internal Forward Primer: CCATTTCGCTATTAGTAAAC

S8 Internal Reverse Primer: CATTTTGTTATTACCAATTCCCGCC

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The positively confirmed plasmids were named pCR8-A14, pCR8- E40, pCR8- E49 and pCR8-S8.

4.2.6 Codon optimization and gene synthesis

The cassava codon usage table was obtained through Kasuza (www.kasuza.or.jp/codon/)

(Nakamura et al. 2000). The table was used in the Graphical Codon Usage Analyzer

(www.gcua.schoedl.de) (Fhurmann et al. 2004) to evaluate the codon usage in the YFP and CFP sequences and determine any problematic codons for expression in cassava. The codons with low usage in cassava found in the YFP and CFP sequences were exchanged to higher usage codons as determined by the information of the Kasuza codon usage table. The codon optimized YFP and CFP were synthetically made by Epoch Biolabs

(Sugarland, TX, USA). The codon optimized CFP gene was delivered in the pUC19 vector and this vector was renamed pUC-CFP. The codon optimized YFP gene was delivered in the pBluescript II KS (+) and this vector was renamed pBS-YFP.

4.2.7 Destination vector

The codon optimized CFP gene was digested out of the pUC-CFP vector sequentially with KpnI (Invitrogen) and XhoI (Invitrogen) enzymes. The KpnI digestion reaction was done under the following conditions: 100ng of the pUC-CFP vector, 1µL of KpnI enzyme, 3 µL of 10X React 4 ® buffer (Invitrogen) and autoclaved double distilled water up to a volume of 30 µL, the digestion was carried out at 37 ˚C for one hour. After digestion the partially digested pUC-CFP product was purified using the Qiaquick PCR

Cleanup kit according the manufacturer’s instructions (Qiagen), the final elution volume

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was 30µL. The XhoI digestion reaction was done under the following conditions: 30µL of purified partially digested pUC-CFP product, 1µL of XhoI enzyme and 3.5 µL of 10X

React 2 ® buffer (Invitrogen), the digestion was carried out at 37 ˚C for one hour. The product of digestion was run in a 1% (w/v) agarose gel and the band corresponding to

CFP was gel purified using a Qiagen Qiaquick Gel Extraction kit (Qiagen). The pBluescript II KS(+) vector was digested sequentially with KpnI and XhoI following the same procedures as stated above. The digested CFP was ligated with the digested pBluescript II KS(+) vector to make the pBS-CFP vector. The ligation reaction conditions were as follows: 1 µL of T4 DNA ligase (NEB), 2 µL of 10X T4 DNA ligase buffer, 50 ng of digested vector, 150 ng of digested CFP product and autoclaved double distilled water up to a volume of 20 µL. The ligation product of CFP and the pBluescript

II KS(+) product was used directly to transform E.coli DH5ɑ sub-cloning efficiency cells according to the protocol given by the supplier Invitrogen (Catalog # 18265017). After transformation the cells were plated in LB media containing ampicillin (100 µg/mL) and incubated for 12 hours at 37˚C. The resulting putative transformed colonies were grown in liquid LB media containing ampicillin (100 µg/mL) for 12 hours at 37˚C. The plasmid

DNA was isolated using the Qiagen Qiaquick Mini Prep kit following the instructions provided by the manufacturers and sent for sequencing to the Ohio State Plant Microbe

Genomics Facility (Columbus, OH)

The NOS terminator was amplified from the pBI121 vector with XhoI and ClaI sites at the 5’end and 3’end using primers 5’NOSXhoI and 3’NOSClaISal,

5’NOSXhoI: CCGGCTCGAGCGAATTTCCCCGATCGTTCAAAC

3’NOSClaISal: CCGGATCGATGTCGACCCCGATCTAGTAACATAGATGACACCG

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The amplification reaction was set up with 15 ng of pBI121 plasmid DNA, one unit of the enzyme Platinum Taq High Fidelity DNA Polymerase (Invitrogen, Carlsbad, CA),

10µL of the supplied 10X High Fidelity PCR Buffer, 2 µL of 50 mM MgSO4, 2 µL of 10 mM dNTP mixture, 2.5 µL of 10 µM solution of each primer and autoclaved double distilled water to a volume of 50 µL. The PCR reactions were performed in a BioRad

Mycycler (Bio-Rad, Hercules, CA) under the following conditions: one cycle of 5 min at

94˚C followed by 32 cycles of amplification (30s at 94˚C, 30s at 54˚C and 45s at 72˚C) and a final cycle of 5min at 72˚C. The product was fractioned on 1% agarose gel in TAE buffer and gel purified using the QiaQuick Gel Extraction Kit (Qiagen).

The pBS-CFP-NOS vector was constructed by inserting the PCR amplified NOS terminator in the XhoI/ClaI sites of the pBS-CFP vector, briefly 500ng of PCR amplified

NOS terminator and 300 ng of pBS-CFP vector were digested individually in a reaction containing 1 µL of ClaI enzyme, 1 µL of XhoI enzyme, 3µL of 10X React 4 ® buffer

(Invitrogen) and autoclaved double distilled water up to a volume of 30µL, the digestion was carried out at 37 ˚C for one hour. After digestion both products were run in a 1%

(w/w) agarose gel in TAE buffer and gel purified using the Qiaquick Gel Extraction kit according the manufacturer’s instructions (Qiagen), the final elution volume was 30µL.

The ligation of the NOS terminator and pBS-CFP were as follows: 1 µL of T4 DNA ligase (NEB), 2 µL of 10X T4 DNA ligase buffer, 50 ng of digested pBS-CFP, 150 ng of digested NOS product and autoclaved double distilled water up to a volume of 20 µL.

The ligation product of NOS and the pBS-CFP vector was used directly to transform

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E.coli DH5ɑ sub-cloning efficiency cells according to the protocol given by the supplier

Invitrogen (Catalog # 18265017). After transformation the cells were plated in LB media containing ampicillin (100 µg/mL) and incubated for 12 hours at 37˚C. The resulting putative transformed colonies were grown in liquid LB media containing ampicillin (100

µg/mL) for 12 hours at 37˚C. The plasmid DNA was isolated using the Qiagen Qiaquick

Mini Prep kit following the instructions provided by the manufacturers and sent for sequencing to the Ohio State Plant Microbe Genomics Facility (Columbus, OH), the confirmed product was named pBS-CFP-NOS.

The 35S-GUS-NOS cassette was digested out from pBI121 by simultaneously with

HindIII and EcoRI, the digestion reaction was as follows, 500 ng of pBI121 vector, 1 µL of HindIII enzyme (NEB), 1 µL of EcoRI enzyme (NEB), 3µL of 10X NEB Buffer 2 and autoclaved double distilled water up to a volume of 30µL, the digestion was carried out at

37 ˚C for one hour. After digestion the reaction products were ran in a 1% (w/w) agarose gel in TAE buffer and the band corresponding to the 35S: GUS: NOS cassette was gel purified using the Qiaquick Gel Extraction kit according the manufacturer’s instructions

(Qiagen), the final elution volume was 30µL. The HindIII/EcoRI digested 35S: GUS:

NOS cassette was ligated in to a similarly digested pUC19 vector. The ligation of these two products was as follows: 1 µL of T4 DNA ligase (NEB), 2 µL of 10X T4 DNA ligase buffer, 50 ng of digested pUC19, 150 ng of digested 35S: GUS: NOS cassette and autoclaved double distilled water up to a volume of 20 µL. The ligation product of was used directly to transform E.coli DH5ɑ sub-cloning efficiency cells according to the protocol given by the supplier Invitrogen (Catalog # 18265017). After transformation the

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cells were plated in LB media containing ampicillin (100 µg/mL) and incubated for 12 hours at 37˚C. The resulting putative transformed colonies were grown in liquid LB media containing ampicillin (100 µg/mL) for 12 hours at 37˚C. The plasmid DNA was isolated using the Qiagen Qiaquick Mini Prep kit following the instructions provided by the manufacturers and sent for sequencing to the Ohio State Plant Microbe Genomics

Facility (Columbus, OH), the confirmed product was named pSP102.

The GUS coding sequence was removed from pSP102 by digestion with XbaI (NEB) and SacI (NEB), the digestion reaction was as follows: 500 ng of pSP102 vector, 1 µL of

XbaI enzyme (NEB), 1 µL of SacI enzyme (NEB), 3µL of 10X NEB Buffer 4 and autoclaved double distilled water up to a volume of 30µL, the digestion was carried out at

37 ˚C for one hour. After digestion the reaction products were ran in a 1% (w/w) agarose gel in TAE buffer and the band corresponding to the remaining digested pSP02 vector was gel purified using the Qiaquick Gel Extraction kit according the manufacturer’s instructions (Qiagen), the final elution volume was 30µL.

The YFP gene was codon optimized and delivered in pBluescript II KS (+); this vector was named pBS-YFP. The YFP gene was digested out from pBS-YFP with XbaI and

SacI, the digestion reaction was set up with the digestion reaction was as follows: 500 ng of pBS-YFP vector, 1 µL of XbaI enzyme (NEB), 1 µL of SacI enzyme (NEB), 3 µL of

10X NEB Buffer 4 and autoclaved double distilled water up to a volume of 30 µL, the digestion was carried out at 37 ˚C for one hour. After digestion the reaction products were ran in a 1% (w/w) agarose gel in TAE buffer and the band corresponding to the digested YFP gene was gel purified using the Qiaquick Gel Extraction kit according the manufacturer’s instructions (Qiagen), the final elution volume was 30 µL. The digested

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YFP gene was ligated in to similarly digested pSP102. The ligation of these two products was as follows: 1 µL of T4 DNA ligase (NEB), 2 µL of 10X T4 DNA ligase buffer, 50 ng of digested pSP102, 150 ng of digested YFP gene and autoclaved double distilled water up to a volume of 20 µL. The ligation product of was used directly to transform

E.coli DH5ɑ sub-cloning efficiency cells according to the protocol given by the supplier

µg/mL) for 12 hours at 37˚C. The plasmid DNA was isolated using the Qiagen Qiaquick

The 35S-YFP-NOS cassette was amplified from the pSP-YFP vector using primers that included a PstI enzyme site at the 5’ end and a HindIII site at the 3’end (35S5’PstI and

NOS3’HindIII primers).

35S5’PstI: ATGCCTGCAGCCCACAGATGGTTAGAGAGGC

NOS3’HindIII: CCGGAAGCTTCCCGATCTAGTAACATAGATGACACCG

The PCR amplified 35S-YFP-NOS was gel purified and the final product was digested with PstI (NEB) and HindIII (NEB), the digestion reaction was set as follows: 500 ng of

35S-YFP-NOS PCR product, 1 µL of PstI enzyme (NEB), 1 µL of HindIII(NEB) enzyme

(NEB), 3 µL of 10X NEB Buffer 4 and autoclaved double distilled water up to a volume

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of 30 µL, the digestion was carried out at 37 ˚C for two hours. After digestion the reaction product was ran in a 1% (w/w) agarose gel in TAE buffer and gel purified using the Qiaquick Gel Extraction kit according the manufacturer’s instructions (Qiagen), the final elution volume was 30 µL. The digested 35S:YFP:NOS was ligated in to similarly digested pCambia2300 vector. The ligation of these two products was as follows: 1 µL of T4 DNA ligase (NEB), 2 µL of 10X T4 DNA ligase buffer, 50 ng of digested pCambia2300, 150 ng of digested 35:YFP:NOS and autoclaved double distilled water up to a volume of 20 µL. The ligation product of was used directly to transform E.coli

DH5ɑ sub-cloning efficiency cells according to the protocol given by the supplier

Invitrogen (Catalog # 18265017). After transformation the cells were plated in LB media containing kanamycin (50 µg/mL) and incubated for 12 hours at 37˚C. The resulting putative transformed colonies were grown in liquid LB media containing kanamycin (50

µg/mL) for 12 hours at 37˚C. The plasmid DNA was isolated using the Qiagen Qiaquick

The CFP-NOS cassette was removed from pBS-CFP-NOS by sequential digestion with

KpnI (NEB) and SalI (NEB). The KpnI digestion reaction was done under the following conditions: 500ng of the pBS-CFP-NOS vector, 1µL of KpnI enzyme, 3 µL of 10X NEB

Buffer 1 and autoclaved double distilled water up to a volume of 30 µL, the digestion was carried out at 37 ˚C for one hour. After digestion the partially digested pBS-CFP-NOS product was purified using the Qiaquick PCR Cleanup kit according the manufacturer’s instructions (Qiagen), the final elution volume was 30 µL. The SalI digestion reaction

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was done under the following conditions: 30 µL of purified partially digested pUC-CFP product, 1 µL of SalI enzyme and 3.5 µL of 10X NEB Buffer 3, the digestion was carried out at 37 ˚C for one hour. The product of digestion was ran in a 1% (w/v) agarose gel and the band corresponding to the CFP:NOS casette was gel purified using a Qiagen

Qiaquick Gel Extraction kit (Qiagen). The pCAM-YFP vector was digested sequentially with KpnI and SalI following the same procedures as stated above. The digested

CFP:NOS cassette was ligated in to the pCAM-YFP vector to make the pBS-CFP vector.

The ligation reaction conditions were as follows: 1 µL of T4 DNA ligase (NEB), 2 µL of

10X T4 DNA ligase buffer, 50 ng of digested pCAM-YFP, 150 ng of digested CFP:NOS product and autoclaved double distilled water up to a volume of 20 µL. The ligation product was used directly to transform E.coli DH5ɑ sub-cloning efficiency cells according to the protocol given by the supplier Invitrogen (Catalog # 18265017). After transformation the cells were plated in LB media containing kanamycin (50 µg/mL) and incubated for 12 hours at 37˚C. The resulting putative transformed colonies were grown in liquid LB media containing kanamycin (50 µg/mL) for 12 hours at 37˚C. The plasmid

DNA was isolated using the Qiagen Qiaquick Mini Prep kit following the instructions provided by the manufacturers and sent for sequencing to the Ohio State Plant Microbe

Genomics Facility (Columbus, OH), the confirmed product was renamed pCAM-CFP-

YFP.

The pCAM-CFP-YFP vector was digested with EcoRI (NEB) under the following conditions: 500ng of the pCAM-CFP-YFP vector, 1µL of EcoRI enzyme, 3µL of 10X

NEB Buffer 1 and autoclaved double distilled water up to a volume of 30µL, the digestion was carried out at 37 ˚C for one hour. The product was purified using Qiagen

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PCR Cleanup Kit according to the instructions of the manufacturer (Qiagen) and treated with T4 DNA Polymerase (Invitrogen) to make the ends blunt, after alkaline phosphatase treatment the vector was ligated with the Gateway Reading Frame Cassette A (GrfA,

Invitrogen). The position of this last element is exactly before the CFP coding sequence.

The resulting plasmid was named pCAM-YCFP-DES and it was our destination vector.

4.2.8 Expression vector

The expression vectors were generated by LR recombination reactions (Gateway LR

Clonase Enzyme, Invitrogen) of pCAM-YCFP-DES (destination vector) and each of the pCR8 promoter vectors (entry vector). The LR recombination was set up as follows for each promoter: 300 ng of pCAM-YCFP-DES, 300 ng of pCR8 (entry vector), 4 µL of 5X

LR Clonase Reaction Buffer and water to a volume of 16 µL. From the product of the recombination reaction 1 µL was taken and used to transform DH5α cells (Invitrogen).

The obtained colonies were grown overnight at 37˚C under kanamycin selection in 5 mL of liquid LB media. Plasmid DNA was isolated from the bacterial cultures using a Qiagen

DNA miniprep kit and following instructions provided by manufacturer. The correct integration of the promoter by recombination was confirmed by PCR and digestion reactions. The restriction digestion reactions where XhoI and SacI for the A14 containing vector. They were KpnI/HindIII for E40 containing vector, and for the E49 and S8 vectors, HindIII and SacI. The final four expression vectors have an Arabidopsis promoter-CFP-NOS-35S-YFP-NOS cassette and they were named A14:CFP, E40:CFP,

E49:CFP and S8:CFP.

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4.2.9 Generation of Friable Embryogenic Callus

The production of FEC material was carried out based on the procedures previously outlined by Taylor et al. (1996 and personal communication 2008). In vitro cassava plantlets were subcultured and grown in Murashige and Skoog media (1962) supplemented with 20g/L sucrose, MS Vitamins (Sigma Aldrich, USA) and 2.4 g/L of the gelling agent Phytagel, this media will be subsequently known as MS2. After 6 weeks, apical leaves from the plantlets were carefully removed and placed on Driver and

Kuniyuki media (1984) supplemented with 20 g/L sucrose, 50 µM picloram and 2.4 g/L of Phytagel, this media was named DKW2. After 4 weeks in DKW2 media the visible organized embryogenic structures (OES) were removed and placed in fresh DKW2 plates. The OES remained in the second DKW2 plate for 3 weeks after which time they were placed in Gresshoff and Doy media (1974) supplemented with 20 g/L sucrose, 50

µM picloram and 2.4 g/L Phytagel, this media was named GD2P50. After 3 weeks in

GD2P50 the OES may form friable embryogenic callus (FEC) structures, these were removed and placed in fresh GD2P50 for 3 more weeks. Finally the growing FEC tissues are removed from the second GD2P50 and proliferated for one more round in GD2P50 for three weeks. After this the FECs were collected and used for transformation.

4.2.10 Transformation of cassava

Cassava transformation was carried out using friable embryogenic callus (FEC). The subsequent transformation steps were carried out as outlined by Taylor et al. (1996 and personal communication 2008). Briefly, the generated FEC material was co-cultivated with Agrobacterium containing one of the following vectors: A14:CFP, E40:CFP,

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E49:CFP and S8:CFP. After inoculation the FECs were placed in GD media (Gresshoff and Doy, 1974) containing 50 µM picloram, 500 mg/L carbenicillin and 27.5 µM paramomycin for 28 days. After this period of time the FECs were transferred to Stage 1 regeneration media which consisted of MS2 media supplemented with 5µM NAA, 250 mg/L carbenicillin and 45 µM paramomycin. After 21 days growing tissues were transferred to Stage 2 regeneration media (MS2 medium plus 0.5 µM NAA and 45 µM paramomycin). The preferred tissues transferred from Stage 1 to Stage 2 were any FEC clusters showing early torpedo stage or beginning cotyledon stage embryos. At 21 days of growth in Stage 2 the embryos that had developed into a mature cotyledon stage were placed in the germination stage media (MS2 medium supplemented with 2µM BAP).

Embryos that germinated were micropropagated in to MS2 media, each germinated embryo is considered an individual transgenic event. The transgenic lines were grown and maintained in MS2 media.

4.2.11 DNA extraction and PCR

YFP gene. The following conditions were used in the PCR: one cycle of 5min at 94ºC followed by 36 cycles of amplification (30 s at 94ºC, 60 s at annealing temperature and

45 s at 72º C) and a final cycle of 5 min at 72º C. The annealing temperature for the

NPTII primer set (NPT5 and NPT3) was 53˚ C and for the YFP primer set (M5YFP and

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3YFP) it was 58˚ C. The ChoiceBlue Taq DNA Polymerase (Denville Scientific,

Metuchen, NJ) was used in all reactions.

NPT5: GGA TTG CAC GCA GGT TCT CCG

NPT3: GCC ACA GTC GAT GAA TCC AGA AAA GC

M5YFP: CTT CGG CTA CGG CCT TCA GTG C

3YFP: CTT TTC ACT TAT CGT CAT CAT CCT TAT AAT CCT TGT ACA GCT

CGT C

4.2.12 Dot blot

Genomic DNA solutions were diluted to a concentration of 20 ng/µL. The samples for the dot blots consisted of three 100 ng aliquots of genomic DNA from each putative transgenic line and from four control lines. The control lines have a known copy number of the 35S promoter determined by Southern blot, the lines have 3, 2, 1 and 0 copy numbers. All samples were denatured by adding 0.4 M NaOH followed by boiling for 5 min. A Nylon Hybond N+ membrane was prepared by wetting in 2X SSC buffer (0.3 M

NaCl, 0.03 M Sodium citrate, pH = 7), it was then placed on top of a moist sheet of

Whatman 3MM filter paper, these two were then placed in between the two layers of the dot blot apparatus. To each sample a volume of 150 µL of 2X SSC buffer was added post-denaturation, the total volume of each sample was applied to the wells of the dot- blot apparatus and a gentle vacuum was used to draw the samples in to the membrane. To immobilize the DNA it was cross-linked to the membrane using a Stratalinker UV-

Crosslinker (Stratagene) set at an energy setting of 120,000 microjoules/cm2. A digoxenin (DIG)-labeled probe targeted to the 35S promoter was synthesized using the

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Roche DIG Probe Synthesis Kit. The dot blot membranes were hybridized and processed for detection of the probe according to the protocol of the DIG High Prime DNA

Labeling and Detection Starter Kit II. The final detection was done by chemiluminescence, each dot blot was incubated with the CDP-Star Reagent followed by exposure to X-Ray films. The films were scanned using an Epson Scanner and the Adobe

Photoshop program. The dot blot images were analyzed using ImageJ (www.nih.gov) to quantify the intensity of each spot. An average intensity value from the three replicates was obtained for each line and the controls. Using the intensity value for the controls a standard curve equation was obtained of copy number vs. spot intensity. The equation was used to calculate the copy number in the transgenic lines.

4.2.13 Microscopy

The excitation wavelength for YFP is 514 nm while the emission wavelength is 527 nm.

The excitation wavelength for CFP is 435 nm while the emission wavelength is 495 nm.

Initial screening of the transgenic lines was done using a Nikon Eclipse 800 Widefield

Epifluorescence microscope with mercury arc lamp as excitation source. The CFP filter settings used were 426-446 nm for excitation and 460-500nm for emission. The YFP filter settings used were 490-510 nm for excitation and 520-550 for emission.

Further imaging of in vitro fibrous roots and leaves was done using a Zeiss LSM 510 inverted confocal microscope equipped with a 488 nm Argon laser and a Ti-Sapphire multiphoton laser. Fluorescent images of YFP were obtained using the 488 nm laser for excitation, a main dichroic beam splitter of 700/488 and a band pass of 500-550nm.

Images of CFP fluorescence were obtained using the Ti-Sapphire laser set at 790 nm, a

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main dichroic of 650 nm and a band pass of 480-520 nm. All samples used were hand- sliced fresh cassava fibrous roots or leaves. Cassava leaves and roots display autofluorescence at the CFP excitation and emission wavelengths, to eliminate this background the laser intensity was lowered.

4.2.14 RNA extraction

RNA was extracted from the leaves of 1.5 and 4 month old cassava plants, from the fibrous roots of 1.5 month old plants and from the storage root cortex and storage root peels of 4 month old cassava. All tissues from 4 month old cassava were freeze dried prior to extraction using a Flexi-Dry MP freeze dryer (FTS Systems, Stone Ridge, NY,

USA). Extraction of leaf total RNA was done using a CTAB based extraction buffer and following procedures similar to those explained by Cota-Sanchez et al. (2006). Briefly, the CTAB buffer (2% CTAB, 100mM Tris-HCl, 20mM EDTA, 1.4M NaCl, 5% β- mercaptoethanol) was warmed to 65ºC and a 150-200 mg of ground leaf tissue was added to 700 µL of buffer. The samples were incubated at 65º C for 30 min then extracted twice with 700 µL of chloroform:isoamyl alcohol (24:1). The nucleic acids were precipitated by adding 350 µL of chilled isopropanol, followed by centrifugation at 13,000 rpm for 10 min. The isopropanol was decanted and the samples were washed twice with 500 µL of

70% ethanol. The samples were air-dried for 20 min, resuspended in 50 µL of RNase-free water and finally treated with RQ1 RNase-free DNase (Promega, Madison, WI, USA) as follows: 40 µL of RNA solution, 4 µL of RQ1 10X Reaction Buffer and 2 µL of RQ1

DNase; the reaction was incubated at 37 ˚C for 30 min, to stop the reaction 4 µL of RQ1

Stop Solution were added to the sample and then incubated at 65 ˚C for 10 min.

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The extraction of RNA from storage root pulp and peel was done using a LiCl extraction method based on the methodology developed by Manickavelu et al. (2007). A ground sample of 50-100 mg of dry storage root tissue was extracted with 900 µL of LiCl buffer

(100 mM LiCl, 1% SDS (w/v), 100 mM Tris-HCl, 100mM EDTA and 1% β- mercaptoethanol) and 900 µL of 125 phenol: 24 chloroform: 1 isoamyl alcohol for 60 min at room temperature. The samples were centrifuged at 12,400 RCF rpm for 20 min, the aqueous upper phase was transferred to a fresh tube and a volume of 300 µL of 8 M

LiCl was added to precipitate the RNA from the tissue samples. The root tissue RNA samples were precipitated overnight at 4 ˚C; the next day the precipitated nucleic acid was pelleted by centrifuging at 16,800 RCF for 10 min at a temperature of 4 ˚C. The

RNA pellet was washed twice by adding 100 µL of ice cold 80% ethanol and centrifuging at 16,800 RCF at a temperature of 4 ˚C, the ethanol solution was then removed and the RNA pellet was air dried for 10 min at room temperature. The pellets of nucleic acid were resuspended in 40 µL of RNase-free water (Promega, Madison, WI) and treated with RQ1 RNase-free DNase (Promega) as follows: 40 µL of RNA solution,

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again at 16800 RCF, the ethanol was removed and the RNA pellet was resuspended in 50

µL of RNase-free water.

The extraction of RNA from the leaves and fibrous roots of young cassava plants was done using the Qiagen RNeasy Plant Mini Kit (Qiagen, Valencia, CA,USA), all tissues were frozen in liquid nitrogen prior to extraction.

For all RNA from leaf, storage root and fibrous roots we determined the concentration,

260/280 nm ratio and 260/230 nm ratio of each sample using a NanoDrop 2000

Spectrophotometer. The value of the 260/280 nm ratio and the 260/230 nm ratio should be 2.0 or close to 2.0 (1.8-2.3), if the values were not in this range the RNA extraction was repeated.

The cDNA for all samples was synthesized using 1 µg of RNA, 4µL of Quantas qScript cDNA SuperMix (Quanta Biosciences, Gaithersburg, MD) and RNase-free water

(Promega) up to a volume of 20 µL. The reaction was run in BioRad Mycycler using the following program: 5 min at 25 ˚C, 30 min at 42 ˚C, 5 min at 85˚C and hold at 4 C˚. The cDNA obtained was stored at -20 ˚C.

4.2.15 Semiquantitative RT-PCR

The RT-PCR reaction was carried out using the first-strand cDNAs as a templates. Gene specific primers for YFP (5’MYFP and 3’YFP) and CFP (5’MCFP and 3’CFP) were used. As a control for the cDNA reaction and the overall quality of the RNA extraction a reaction for the cassava tubulin gene (5’TUB and 3’TUB) was prepared and ran at the same time.

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5’MYFP: CTTCGGCTACGGCCTTCAGTGC

3’YFP: CTTTTCACTTATCGTCATCATCCTTATAATCCTTGTACAGCTCGTC

5’MCFP: CTGACCTGGGGCGTGCAG

3’ CFP: GCTGTCCCCTCAAGCATAATCTGGAACATC

5’TUB: GAT CCT ACT GGG AAG TAC ATT GG

3’TUB: CTG CAT TCT CCA CCA ACT GA

The following conditions were used in the PCR: one cycle of 5 min at 94º C followed by

32 cycles of amplification (30 s at 94º C, 30 s at annealing temperature and 45 s at 68ºC) and a final cycle of 5 min at 68º C. The Platinum Pfx DNA Polymerase enzyme

(Invitrogen) was used in all reactions as follows: 1000 ng of cDNA, 1 unit of Platinum

Pfx DNA Polymerase, 5µL of 10X Pfx Amplification Buffer, 0.5 µL of 50 mM MgSO4,

1.5 µL of 10 mM dNTP solution, 1.5 µL of each 10µM primer solution and autoclaved double distilled water up to 25 µL. The annealing temperature for the YFP primers was

58º C, for the CFP primer pair it was 57º C and for the tubulin primer pair it was 55º C.

The amplification products were separated in 1% (w/v) agarose gels and stained with ethidium bromide. The amplification products from the CFP and YFP reactions were gel purified and sequenced to confirm amplification of the correct transcript.

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4.2.16 Real-time RT-PCR

The first-strand cDNA’s where used as a template, the cDNA reaction product was diluted to a concentration of 1 ng/µL and in four replicates 5 ng were loaded of each sample in a 96-well plate (Axygen PCR Microplate, Axygen, Union City, CA). In addition each well reaction was added 15 µL of RT-PCR reaction mix corresponding to

10 µL of SYBR Green ROX (Quanta BioSciences, Gaithersburg, MD), 1 µL each of the promoter forward and reverse primer for YFP (RTYFP5 and RTYFP3), CFP (RTCFP5 and RTCFP3) or the housekeeping control tubulin (RTTUB5 and RTTUB3) and 3 µL of

0.1% (v/v) diethylpyrocarbonate (DEPC) treated water. The forward and reverse primers used were,

RTYFP5: CAA CTC TCA CAA CGT CTA TAT CAT GG

RTYFP3: GCT CTT TTC ACT TAT CGT CAT CAT CC

RTCFP5: CTG ACC TGG GGC GTG CA

RTCFP3: CGG TGA TAT AGA CGT TGT GGC TGA TGT AG

RTTUB5: GTG GAG GAA CTG GTT CTG GA

RTTUB3: TGC ACT CAT CTG CAT TCT CC

The real-time RT-PCR reaction was carried out using an Applied Biosystems Step One

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The calculations were as follows: the average Ct value of YFP and CFP for each promoter tissue sample was normalized to the average Ct value of the endogenous tubulin gene in the same promoter tissue sample to obtain the ΔCt values for example,

The ΔCt values of the different samples are then compared to a reference sample, in many situations the reference sample is an untreated control or the native gene, however in our case the reference sample was YFP expression in each tissue.

To calculate the ΔΔCt the following formula was used,

The ΔΔCt is calculated for both CFP and YFP, the YFP value will be 0. Finally the relative expression was calculated, where the relative expression is the amount of target normalized to the endogenous housekeeping gene and relative to a reference sample, the relative expression of YFP using the formula presented below is 1, the CFP samples are expressed as fold higher or lower than YFP in their respective tissues of expression.

The relative expression formula is,

The standard errors for the relative expression were calculated according to the SDS RQ

Manager 1.1 Software Manual (Applied Biosystems).

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4.3 RESULTS

4.3.1 Arabidopsis promoters

The A14, E40, E49 and S8 promoters were chosen due to their different tissues of expression as it was of interest to obtain promoters for expression in different cassava root tissues. In Arabidopsis roots A14 is an epidermal promoter, E40 effects expression in the endodermis and stele, E49 is a root cortex promoter and S8 is a pericycle promoter.

4.3.2 Codon optimization

The gene sequences of YFP and CFP were submitted to the Graphical Codon Usage

Analyzer to evaluate any problematic codon usage bias of these genes in cassava. The codon usage for three amino acids: proline, serine and arginine was biased against optimal expression in cassava (Fig. 4.6) For proline in both YFP and CFP the only codon used was “CCC”, however, in cassava the proline codon is primarily “CCA” followed by

“CCT”. In the case of serine the abundantly used codons in cassava are “TCA” and

“TCT” while the sequences have a split usage between “AGC” and “TCC”. And finally for arginine the only codon present for this amino acid in YFP and CFP was “CGC”; while in cassava the preference is for “AGA”. Based on these results the sequences of

YFP and CFP were codon optimized, the differences between the non-optimized and codon-optimized sequences are presented in Fig.7.

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Figure 4.6 Comparison of the codon usage of (A) YFP and (B) CFP to the codon usage in cassava was done using the Graphical Codon Usage Analyzer. Three amino acids stood out with significant bias, proline, serine and arginine. In black bars are the % usage for each codon in cassava and in red bars are represented the % usage of each codon used in the individual sequences.

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YFP non : ATGGTGAGCAAGGGCGAGGAGCTGTTCACCGGGGTGGTGCCCATCCTGGTCGAGCTGGACGGCGACGTAAACGGCCACAAGTTCAGCGTGTCCGGCGAGGGCGAGGGCGATGCCACCTACGGCAAGCTGACCCTGAAGTTCATCTGCACC : 150 YFP codon : ATGGTGTCTAAGGGCGAGGAGCTGTTCACCGGGGTGGTGCCAATCCTGGTCGAGCTGGACGGCGACGTAAACGGCCACAAGTTCTCTGTGTCCGGCGAGGGCGAGGGCGATGCCACCTACGGCAAGCTGACCCTGAAGTTCATCTGCACC : 150

YFP non : ACCGGCAAGCTGCCCGTGCCCTGGCCCACCCTCGTGACCACCTTCGGCTACGGCCTGCAGTGCTTCGCCCGCTACCCCGACCACATGAAGCAGCACGACTTCTTCAAGTCCGCCATGCCCGAAGGCTACGTCCAGGAGCGCACCATCTTC : 300 YFP codon : ACCGGCAAGCTGCCAGTGCCATGGCCAACCCTCGTGACCACCTTCGGCTACGGCCTTCAGTGCTTCGCCAGATACCCAGACCACATGAAGCAGCACGACTTCTTCAAGTCCGCCATGCCAGAAGGCTACGTCCAGGAGAGAACCATCTTC : 300

YFP non : TTCAAGGACGACGGCAACTACAAGACCCGCGCCGAGGTGAAGTTCGAGGGCGACACCCTGGTGAACCGCATCGAGCTGAAGGGCATCGACTTCAAGGAGGACGGCAACATCCTGGGGCACAAGCTGGAGTACAACTACAACAGCCACAAC : 450 YFP codon : TTCAAGGACGACGGCAACTACAAGACCAGAGCCGAGGTGAAGTTCGAGGGCGACACCCTGGTGAACAGAATCGAGCTGAAGGGCATCGACTTCAAGGAGGACGGCAACATCCTGGGGCACAAGCTGGAGTACAACTACAACTCTCACAAC : 450

YFP non : GTCTATATCATGGCCGACAAGCAGAAGAACGGCATCAAGGTGAACTTCAAGATCCGCCACAACATCGAGGACGGCAGCGTGCAGCTCGCCGACCACTACCAGCAGAACACCCCCATCGGCGACGGCCCCGTGCTGCTGCCCGACAACCAC : 600 YFP codon : GTCTATATCATGGCCGACAAGCAGAAGAACGGCATCAAGGTGAACTTCAAGATCAGACACAACATCGAGGACGGCTCTGTGCAGCTCGCCGACCACTACCAGCAGAACACCCCAATCGGCGACGGCCCAGTGCTGCTGCCAGACAACCAC : 600

YFP non : TACCTGAGCTACCAGTCCGCCCTGAGCAAAGACCCCAACGAGAAGCGCGATCACATGGTCCTGCTGGAGTTCGTGACCGCCGCCGGGATCACTCTCGGCATGGACGAGCTGTACAAGGATTATAAGGATGATGACGATAAGTGA : 744 YFP codon : TACCTGTCTTACCAGTCCGCCCTGTCTAAAGACCCAAACGAGAAGAGAGATCACATGGTCCTGCTGGAGTTCGTGACCGCCGCCGGGATCACTCTCGGCATGGACGAGCTGTACAAGGATTATAAGGATGATGACGATAAGTGA : 744

CFP non : ATGGTGAGCAAGGGCGAGGAGCTGTTCACCGGGGTGGTGCCCATCCTGGTCGAGCTGGACGGCGACGTAAACGGCCACAAGTTCAGCGTGTCCGGCGAGGGCGAGGGCGATGCCACCTACGGCAAGCTGACCCTGAAGTTCATCTGCACC : 150 CFP codon : ATGGTGTCTAAGGGCGAGGAGCTGTTCACCGGGGTGGTGCCAATCCTGGTCGAGCTGGACGGCGACGTAAACGGCCACAAGTTCTCTGTGTCCGGCGAGGGCGAGGGCGATGCCACCTACGGCAAGCTGACCCTGAAGTTCATCTGCACC : 150

311 CFP non : ACCGGCAAGCTGCCCGTGCCCTGGCCCACCCTCGTGACCACCCTGACCTGGGGCGTGCAGTGCTTCAGCCGCTACCCCGACCACATGAAGCAGCACGACTTCTTCAAGTCCGCCATGCCCGAAGGCTACGTCCAGGAGCGCACCATCTTC : 300 CFP codon : ACCGGCAAGCTGCCAGTGCCATGGCCAACCCTCGTGACCACCCTGACCTGGGGCGTGCAGTGCTTCTCAAGATACCCAGACCACATGAAGCAGCACGACTTCTTCAAGTCCGCCATGCCAGAAGGCTACGTCCAGGAGAGAACCATCTTC : 300

CFP non : TTCAAGGACGACGGCAACTACAAGACCCGCGCCGAGGTGAAGTTCGAGGGCGACACCCTGGTGAACCGCATCGAGCTGAAGGGCATCGACTTCAAGGAGGACGGCAACATCCTGGGGCACAAGCTGGAGTACAACTACATCAGCCACAAC : 450 CFP codon : TTCAAGGACGACGGCAACTACAAGACCAGAGCCGAGGTGAAGTTCGAGGGCGACACCCTGGTGAACAGAATCGAGCTGAAGGGCATCGACTTCAAGGAGGACGGCAACATCCTGGGGCACAAGCTGGAGTACAACTACATCAGCCACAAC : 450

CFP non : GTCTATATCACCGCCGACAAGCAGAAGAACGGCATCAAGGCCAACTTCAAGATCCGCCACAACATCGAGGACGGCAGCGTGCAGCTCGCCGACCACTACCAGCAGAACACCCCCATCGGCGACGGCCCCGTGCTGCTGCCCGACAACCAC : 600 CFP codon : GTCTATATCACCGCCGACAAGCAGAAGAACGGCATCAAGGCCAACTTCAAGATCAGACACAACATCGAGGACGGCTCTGTGCAGCTCGCCGACCACTACCAGCAGAACACCCCAATCGGCGACGGCCCAGTGCTGCTGCCAGACAACCAC : 600

CFP non : TACCTGAGCACCCAGTCCGCCCTGAGCAAAGACCCCAACGAGAAGCGCGATCACATGGTCCTGCTGGAGTTCGTGACCGCCGCCGGGATCACTCTCGGCATGGACGAGCTGTACAAGTATCCATATGATGTTCCAGATTATGCTTGA : 747 CFP codon : TACCTGTCTACCCAGTCCGCCCTGTCTAAAGACCCAAACGAGAAGAGAGATCACATGGTCCTGCTGGAGTTCGTGACCGCCGCCGGGATCACTCTCGGCATGGACGAGCTGTACAAGTATCCATATGATGTTCCAGATTATGCTTGA : 747

Figure 4.7 (A) YFP and (B) CFP non-optimized DNA sequence (identified as “non”) alignment with the codon-optimized

(identified as “codon”) sequence.

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4.3.3 Expression system

The four expression vectors containing A14, E40, E49 and S8 were constructed using

Gateway recombination cloning technology (Fig. 4.8) All the vectors contain a

35S:YFP:nos cassette and an Arabidopsis promoter:CFP:nos cassette. This double construct would allow direct comparison between the Arabidopsis promoters and the 35S promoter.

The A14, E40, E49 and S8 promoters were cloned in to pCR8/GW/TOPO® vector using

TOPO® Cloning (Invitrogen), they were flanked by attL1 and attL2 recombination sites

(Fig. 4.8A). The destination vector contained a Gateway Reading Frame Cassette A with the recombination sites attR1 and attR2 (Fig. 4.8B) The final expression vectors were generated by LR recombination reactions using Gateway Vector Conversion System

(Invitrogen)(Fig. 4.8C).

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313

Figure 8. The final expression vector (c) was obtained thru Gateway recombination of the destination vector (a) and each promoter in

a TOPO® entry vector (b).

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The resulting vectors were checked by restriction digestion analysis (Fig. 9), PCR and sequencing. These final expression vectors contain the Arabidopsis root-specific promoter-CFP-NOS::CaMV35S-YFP-NOS. The 35S:YFP was used as a control to compare expression levels of the Arabidopsis promoters. The confirmed promoter constructs were named A14:CFP, E40:CFP, E49:CFP and S8:CFP.

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Figure 4.9 Restriction digestion analysis of A14:CFP vector with (1) XhoI and (2) SacI.

E40:CFP with (3) KpnI/HindIII. E49:CFP with (4) HindIII and (5) SacI. And S8:CFP with (6) HindIII and (7) SacI.

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4.3.4 Generation of transgenic cassava

Cassava friable embryogenic callus were transformed by incubation with Agrobacterium tumefaciens containing one of the following promoter constructs: A14:CFP, E40:CFP,

E49:CFP and S8:CFP. The friable embryogenic tissues were selected and regenerated in to plantlets under paramomycin selection. A total of 33, 28, 29 and 29 plants were regenerated for A14, E40, E49 and S8 promoter constructs respectively (Table 4.2).

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Promoter Total number of lines No. of lines expressing YFP

A14 38 6

E40 28 8

E49 29 8

S8 29 4

Table 4.2 Total number of lines per promoter that were PCR positive for NPTII and YFP gene and total number of lines exhibiting YFP fluorescence.

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4.3.5 Characterization of transgenics by PCR, dot-blot and widefield fluorescence

All regenerated plantlets were initially characterized for presence of the transgene by

PCR amplification of the NPTII and YFP genes using genomic DNA (Figure 4.10, select lines are shown). The copy number of all PCR positive lines was determined by dot-blot analysis.

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Figure 4.10 (A) PCR amplification of the NPTII gene from genomic DNA. 1: E40-57, 2:

A14-149, 3: E49-64, 4: E40-82, 5: E49-81, 6: A14-60, 7: E40-92, 8:A14-41, 9: S8-32,

10: A14-99, 11: A14-89, 12: A14-101, 13: A14-145, 14: E49-10, 15: S8-34, 16: E49-10,

17: E40-64, 18: S8-103, 19: E49-81, 20: E49-66, 21: E49-40, 22: A14-145, 23: E40-102,

24: E40-58, 25: E49-52, 26: A14-17, 27: E49-8, 28: S8-55. (B) PCR amplification of the

YFP gene from genomic DNA from 8 selected transgenic lines, wild-type cassava and as positive control the pBS-YFP plasmid. For the negative control reaction water instead of

DNA template was used.

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In addition, given the fluorescent properties of the YFP and CFP proteins, the initial characterization of cassava plantlets included evaluation of the YFP and CFP fluorescence in fibrous roots using a Nikon Eclipse 800 Widefield Epifluorescence

Microscope. However, background CFP autofluorescence was observed in wild-type fibrous roots (Fig. 4.11) impairing the ability to visualize CFP expression in the transgenic lines. Unexpectedly, a high percentage of the transgenic lines, ranging from

72%-86%, did not display YFP fluorescence, one such example is E40-57 (Fig. 4.11)

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Figure 4.11 Analysis of YFP and CFP fluorescence of PCR and dot-blot positive transformants using a Nikon Eclipse 800 Widefield Epifluorescence microscope with

CFP excitation filter of 426-446 nm and emission filter of 460-500nm and YFP excitation filter of 490-510 nm and emission filter of 520-550 nm. The wild-type fibrous roots exhibited CFP autofluorescence. Some lines such as E40-57 did not display YFP fluorescence.

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In Fig. 4.12 are shown the results of the YFP fluorescence analysis in relation to the line copy numbers. The results show that plants with higher transgene copy numbers as 5 and

6 had no YFP fluorescence. Out of the total number of plants presenting YFP fluorescence the majority were those of 1 and 2 gene copies. The CaMV 35S constitutive promoter controls the expression of YFP, therefore all lines were expected to have YFP fluorescence emission. Those lines without YFP fluorescence were discarded. After initial characterization 4 to 8 lines remained for each promoter type (Table 2). Two representative lines for each promoter, A14-17, A14-60, E40-92, E40-102, E49-8, E49-

66, S8-34 and S8-55 were selected for further analysis.

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Figure 4.12 Comparison of dot-blot positive transgenic plants displaying YFP (green) fluorescence and those without YFP (red) fluorescence. Lines with 5 and 6 copy numbers had no positive YFP fluorescence lines.

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4.3.6 Analysis of young cassava plants by semi-quantitative RT-PCR

The expression of YFP and CFP in the leaves and fibrous roots of 1.5 month old cassava plantlets was analyzed through semi-quantitative RT-PCR (Figure 4.13, only one line is featured for each promoter). YFP expression was used for normalization of expression and our results show YFP expression in all tissues with the exception of the wild-type leaves and fibrous roots. In the A14:CFP and E40:CFP lines, we observed CFP expression in both leaves and fibrous roots, however, in A14:CFP lines CFP expression was lower in the leaves while in E40:CFP transgenics CFP nearly equivalent expression was observed in leaves and fibrous roots. It appears that in young cassava plants the A14 and E40 promoters are not root-specific. In the E49:CFP lines CFP expression was observed in the fibrous roots but not in leaves while in S8:CFP transgenics, we observed the opposite with CFP expression present in leaves but not in fibrous roots. The S8 promoter was the only promoter that did not drive CFP expression in fibrous roots while the E49 promoter was the only promoter specifically expressed in cassava fibrous roots.

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Figure 4.13 Semiquantitative RT-PCR results from leaf (L) and fibrous root (FR) samples of 1.5 month old cassava.

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4.3.7 Analysis of young cassava plants by confocal microscopy

Tissue samples used for the confocal microscopy analysis came from one and a half month old cassava plantlets. For all lines, we analyzed, fully expanded leaves and roots.

All root sections were from the region above the root tip, and only the epidermal layer, and not the cross section, was imaged. The limitation in imaging root cross sections was caused by the thickness of the roots (1-2mm) and the type of tissue used. These factors prevented the reproduction of adequate images from the cortex and vascular tissues.

Another important aspect in our imaging evaluation was the presence of autofluorescence in both the leaves and fibrous roots of cassava at the CFP excitation wavelength. To avoid the background fluorescence the settings in the confocal microscope were reduced, however, this resulted in the need to decrease the laser intensity by 40%.

For each promoter line the YFP and CFP fluorescence was analyzed (Fig. 4.14). In the case of E40:CFP, E49:CFP and S8:CFP lines the fluorescence microscopy results were very similar. In all three promoter lines the 35S driven YFP fluorescence was observed in both leaf and root samples, the expression of the YFP protein was localized throughout the cytosol and in some cases the nucleus. However, when the same samples were submitted to CFP excitation no fluorescence was detected in any of them. The inability to detect CFP fluorescence in these samples implies that CFP did not accumulate in these transgenics. For the A14 promoter YFP expression was observed in both leaves and roots. However, CFP fluorescence was detected in the root but not in the leaf.

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Figure 4.14 Samples from cassava fibrous roots visualized in a LSM 510 Zeiss confocal microscope. YFP fluorescence observed by excitation with a 488 nm Argon laser. CFP fluorescence observed by excitation with a Ti-sapphire multiphoton laser set at 790 nm.

The thickness of the cassava root does not allow an adequate differential interference contrast (DIC) image. The CFP fluorescence observed in the A14 fibrous roots registered an intensity value (integrated density measured in Image J) that was 9.8 times that of the background fluorescence observed in wild-type fibrous roots. All fibrous roots were from

1.5 month old plants.

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A Z-stack scan was done using a root sample from the A14-60 line (Fig. 4.15). CFP fluorescence was present in the sample up to a depth of 13 µm. In comparison YFP fluorescence was detected until 34 µm. These results show that the A14 promoter drove

CFP expression in the epidermal layer of cassava fibrous roots. The A14 promoter is an epidermal promoter in Arabidopsis and a similar expression pattern is observed in these young cassava roots.

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Figure 4.15 A Z-stack scan from a 1.5 month old fibrous root of A14-60 was obtained using an LSM 510 Zeiss confocal microscope. An initial top section image taken at a depth of 9 µm clearly shows YFP and CFP fluorescence. The next imaging presented (B) was taken at 34 µm. At this point the CFP fluorescence is substantially reduced except at the margin of curvature while YFP fluorescence is present in all tissues.

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4.3.8 Analysis of cassava leaves and storage roots by semi-quantitative RT-PCR

Young cassava plants such as the ones that were analyzed at 1.5 months of age only have a fibrous root system, however, at later stages of growth the formation of storage roots occurs. The cassava storage roots have three main tissues (DeSouza et al 2001), tissue system I (TSI) consists of the epidermis and a layer of parenchyma cells, it is also known as the peel. TSII consists of the phloem and vascular cambium, and TSIII consists of parenchyma cells highly specialized for starch storage and secondary xylem. The TSII and TSIII tissues together are known as the pulp of the root. The storage roots of cassava are the agricultural and commercially important organ of the plant and so our analysis of the four Arabidopsis promoters encompassed evaluation of their different expression patterns in cassava storage root pulp and peel from 4 month old cassava and in leaves from the same plants.

We carried out semi-quantitative RT-PCR analysis of YFP and CFP in the leaves and storage root peel and pulp of 4 month old cassava, in each case the housekeeping gene tubulin served as a control and CFP expression was compared to YFP expression.

Significantly, we detected YFP transcript in all tissues from all 8 promoter lines. This was not the case for CFP as significant variations in levels and tissues of expression were observed between the A14, E40, E49 and S8 lines (Fig. 4.16). In both A14-17 and 60 there was low CFP expression in the root pulp and higher CFP expression the peel. In the leaves of these two lines there was low amplification of CFP. In the case of the two E40 lines the CFP transcript was present with the strongest expression in the storage root pulp

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and in the leaves, and with a lower intensity in the root peel. In the E49 promoter lines there is no CFP transcript detected in any tissue. A very similar result was obtained for

S8 lines 34 and 55. The CFP was amplified only in the leaves of S8-34. These results show that the A14 and E40 promoters have promising activity in cassava storage roots, while the two remaining promoters, S8 and E49 seem to not have any effect on the CFP gene and possibly no promoter activity in cassava storage roots.

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Figure 4.16 Semiquantitative RT-PCR analysis of 4 month old cassava plants. Three different tissues where analyzed (L) Leaf, (PP) storage root pulp and (PL) storage root peel.

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4.3.9 Real-time RT-PCR analysis

Real-time PCR analysis of each promoter line was done for storage root tissues and leaves. It was important to obtain ratios of YFP vs. CFP to determine the strength of the

Arabidopsis promoter versus the constitutive 35S promoter, also very important was the comparison done between CFP relative expression in the pulp (CFP-Pulp) and CFP relative expression in the peel (CFP-Peel), as it would distinguish any tissue specificity of each promoter.

In the semi-quantitative RT-PCR analysis the E49:CFP and S8:CFP lines had low to absent CFP expression in root tissues and in leaves. In the real-time RT-PCR experiment the CFP transcript was detected but in the majority of the tissues from E49 (Fig. 4.17) and S8 transgenic lines (Fig. 4.18) the CFP transcript abundance was low. Our results show that these two promoters have no significant activity in cassava storage roots in comparison to the 35S promoter. The relative expression levels of YFP range from 8 to

250 fold higher than the CFP levels in the pulp and peel of E49 and S8 lines. As previously observed in the semi-quantitative RT-PCR the S8-34 leaf had CFP expression.

We observed a relative expression level of 0.232 for CFP/YFP in our real-time RT-PCR experiment. No conclusion can be drawn from these results as the remaining S8 line did not have similar results.

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Figure 4.17 Real-time RT-PCR analysis of CFP expression driven by the E49 promoter relative to YFP expression driven by the 35S promoter in four month old cassava leaves and storage tubers. Relative CFP (blue) transcript abundance is normalized in relation to the YFP (yellow) transcript in each tissue, prior data analysis included normalization of both YFP and CFP to the endogenous housekeeping gene tubulin. Data shown represents mean values obtained from independent reactions (n=6) and the error bars indicate the upper and lower limit errors as determined by the formulas on the RQ Manager 1.1

Software Manual (Applied Biosystems)

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Figure 4.18 Real-time RT-PCR analysis of CFP expression driven by the S8 promoter relative to YFP expression driven by the 35S promoter in four month old cassava leaves and storage tubers. Relative CFP (blue) transcript abundance is normalized in relation to the YFP (yellow) transcript in each tissue, prior data analysis included normalization of both YFP and CFP to the endogenous housekeeping gene tubulin. Data shown represents mean values obtained from independent reactions (n=6) and the error bars indicate the upper and lower limit errors as determined by the formulas on the RQ Manager 1.1

Software Manual (Applied Biosystems)

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In the case of the A14:CFP lines (Fig. 4.19) our results initially indicated that there was detectable CFP expression. In our analysis for the A14-17 line we observed that YFP relative expression that is 10 fold higher than that of CFP in root pulp. In the peel of this same line there was no significant difference between the levels of expression of the two reporter genes. For the A14-60 line in the pulp the CFP expression was 1.3 fold that of

YFP ; in the peel CFP relative expression is 2.9 fold higher than that of YFP. To determine the tissue specificity of A14 driven gene expression in cassava storage roots, we calculated the CFP-Pulp: CFP-Peel ratio, for A14-17 and A14-60 which were 0.1 and

0.47 respectively. These lower ratios indicate that there is higher CFP expression in the peel of the storage root compared to the pulp. It is clear from these results and the previous experiment that there is CFP expression in the storage root with higher specificity to the peel, as determined by the A14 promoter.

In the E40:CFP lines (Fig. 4.20) we detected CFP transcripts by real-time PCR analysis.

For the E40-92 line the detected levels of YFP relative expression in the pulp are 1.2 fold that of CFP, thus both promoters drive similar expression levels since the transcripts are virtually identical. In the peel of E40-92 the level of YFP is 8.6 fold higher than the value obtained for CFP. In the case of the E40-102 line, in the pulp the CFP relative expression was 2.5 times that of YFP relative expression, while in the peel it is the YFP relative expression that is higher compared to CFP relative expression by a 3 fold increase. This is similar to the results obtained from the previous E40:92 line. In the E40:CFP lines we observed a result contrasting to that of A14:CFP lines when we evaluated the CFP-

Pulp:CFP-Peel ratio. The ratio was 7.3 for E40-92 and 7.4 for E40-102. In the plants

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expressing E40:CFP we observe a higher expression of CFP in the pulp in relation to the peel and in the pulp the E40 promoter strength is similar to the 35S promoter.

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Figure 4.19 Real-time RT-PCR analysis of CFP expression driven by the A14 promoter relative to YFP expression driven by the 35S promoter in four month old cassava leaves and storage tubers. Relative CFP (blue) transcript abundance is normalized in relation to the YFP (yellow) transcript in each tissue. Prior data analysis included normalization of both YFP and CFP to the endogenous housekeeping gene tubulin. Data shown represents mean values obtained from independent reactions (n=6) and the error bars indicate the upper and lower limit errors as determined by the formulas on the RQ Manager 1.1

Software Manual (Applied Biosystems)

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Figure 4.20 Real-time RT-PCR analysis of CFP expression driven by the E40 promoter relative to YFP expression driven by the 35S promoter in four month old cassava leaves and storage tubers. Relative CFP (blue) transcript abundance is normalized in relation to the YFP (yellow) transcript in each tissue, prior data analysis included normalization of both YFP and CFP to the endogenous housekeeping gene tubulin. Data shown represents mean values obtained from independent reactions (n=6) and the error bars indicate the upper and lower limit errors as determined by the formulas on the RQ Manager 1.1

Software Manual (Applied Biosystems)

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4.3.10 Identification of cis-acting elements and transcription start site

The Transcription Start Site (TSS) and the TATA boxes were determined with the

SoftBerry TSSP/Prediction of PLANT Promoters software for the A14 (Fig. 4.2), E40

(Fig. 4.3), E49 (Fig. 4.4) and S8 (Fig. 4.5).

The cis-acting elements were identified for each promoter using the PLACE database. A select number of them are presented in Table 4.3 for promoters A14 and E40 and Table

4.4 for promoters E49 and S8. For the purpose of this chapter the focus of the cis-element analysis was on determining the presence of root expression related motifs and due to the starch content in cassava roots on identifying the presence of sugar related motifs. The cis-element analysis focused as well on identifying elements found exclusively in each promoter.

All four promoters were found to contain the TAPOX1 motif present in multiple sites.

This motif is associated with root expression and is found in the root-specific rolD promoter from Agrobacterium rhizogenes (Elmayan et al. 1995). Another root motif present in all four promoters was the sulfur related element (SURE) present in root- expressed genes that respond to sulfur limitations in the plant (Maruyama et al. 2005). In the A14, E40 and S8 promoters the root hair expression related motif RHERPATEXPA7

(Kim et al. 2006) was found at least once. The A14, E40 and S8 promoters have the

SP8BF cis-element present in their sequences, this element is related to sucrose induced expression in sweet potato storage roots (Ishiguro et al. 1992) and is found in three storage root specific genes in sweet potato among them the sporamin storage protein.

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In difference to the other promoters the A14 promoter contained an enhancer element known as Q-element which is found to be responsible for increasing expression but not tissue specificity of pollen proteins in maize (Hamilton et al. 1998).

The E40 promoter contained a group of abscisic acid (ABA) related cis-elements named

ABRE (Choi et al. 2000), ABRE-like (Hattori et al. 2002) and ABRE-related (Kaplan et al. 2006) these were found in a total of 9 sites, the remaining promoters did not contain

ABA related elements in their sequences.

Three different motifs related to repression of expression by sugar were found in the S8 promoter, these motifs were the A-box (Toyofuku et al. 1998), CGACG motif (Hwang et al. 1998) and SRE (Lu et al. 1998). In addition, the S8 promoter was the only promoter to contain the SURE-1 motif, this motif is found in the potato tuber-specific patatin promoter (Grierson et al., 1994).

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Promoter Cis-element Sequence Position Reference A14 Q-Element AGGTCA -761(-) Hamilton et al. 1998 A14 RHERPATEXPA7 KCACGW -376(+),-2746(-) Kim et al. 2006 A14 TAPOX1 ATATT -283,-520,-756,-1549,- 1579,-1623,-1695,-1758, -1894,-1914,-2070,-2080, Elmayan et al. -2196,-2212,-2244,-2619, 1995 -2699 (+), -286,-306,-412, -447,-454,-521,-634,-838, -1043,-1078,-1361,-1446, -1550,-1895,-2015,-2245, -2773(-) A14 SP8BF TACTATT -1221,-554(+) Ishiguro et al. 1992 A14 SURE GAGAC -1768,-1005,-483 (+) Maruyama et al. 2005 E40 ABRE YACGTGGC -2738 (+) Choi et al. 2000 E40 ABRE-like ACGTG -363,-1339,-2734 (+) , -340, -1560,-2571,-2964 Hattori et al. 2002 (-) E40 ABRE-related MACGYGB -1340 (+) Kaplan et al. 2006 E40 RHERPATEXPA7 KCACGW -2965 (+) Kim et al. 2006 E40 TAPOX1 ATATT -118,-446,-526,-743,-837, -853,-1148,-1277,-1611, -1846,-1885,-1899,-1948, -1960,-1996,-2547,-2698, -3503,-3705,-3826,-3898, -3919 (+) , -202,-211, Elmayan et al. -323,-439,-506,-527,-744, 1995 -764,-838,-854,-1031, -1202,-1278,-1297,-1346, -1612,-1677,-1847,-1857, -1866,-1886,-1904,-1949, -1963,-1969,-1978,-1997, -2258,-2548,-2936,-3113, -3956 (-) E40 SP8BF TACTATT -3679 (+), -1024 (-) Ishiguro et al. 1992 E40 SURE GAGAC -1240,-2882 (+) , -1358,- Maruyama et al. 1457 (-) 2005

Table 4.3 Representative motifs found in the sequence of the A14 and E40 promoters as determined by the PLACE database. Root and sugar related motifs as well as motifs found only in a single promoter are included.

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Promoter Cis-element Sequence Position Reference E49 ABRE-like ACGTG -325 (+) , -560 (-) Hattori et al. 2002 E49 TAPOX1 ATATT -62,-424,-889,-1227, -1336,-1426,-1444,-1586, Elmayan et al. -1722,-2339 (+) , 1995 -185,-425,-823,-1228, -1337,-1713,-2114,-2147, -2340 E49 SRE TTATCC -96,-246,-1598 (+) Lu et al. 1998 E49 SURE GAGAC -1861 (+), -734,-1604, Maruyama et al. -1866 (-) 2005 E49 TATCCAY motif TATCCAY -663,-701 (+) Toyofuku et al. 1998 S8 A-BOX TACGTA -2517 (+)(-) Toyofuku et al. 1998 S8 CGACG motif CGACG -1616 (+), 1613 (-) Hwang et al. 1998 S8 RHERPATEXPA7 KCACGW -979 (-) Kim et al. 2006 S8 TAPOX1 ATATT -933,-1427,-1774,-1931, -2045,-2256,-2332,-2909, -3048,-3512,-3759 (+) , Elmayan et al. -934,-1098,-1148,-1224, 1995 -1932,-1997,-2027,-2141, -2498,-2505,-2937,-3049, -3195,-3261,-3516,-3646, -3745 (-) S8 SP8BF TACTATT -1578 (-) Ishiguro et al. 1992 S8 SRE TTATCC -2920 (-) Lu et al. 1998 S8 SURE-1 AATAGAAAA -1789 (+) Grierson et al. 1994 S8 SURE GAGAC -1662,-2902 (+) , -887,- Maruyama et al. 2397,-2435,-2720 (-) 2005 S8 TATCCAY motif TATCCAY -804,-2250,-2935 (+) Toyofuku et al. 1998

Table 4.4 Representative motifs found in the sequence of the E49 and S8 promoters as determined by the PLACE database. Root and sugar related motifs as well as motifs found only in a single promoter are included.

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4.4 DISCUSSION

Cassava storage roots are an important agricultural and industrial resource around the world and it is of particular interest to improve the qualities of this crop, the inclusion of a variety of promoters for transgene expression in the roots would provide new opportunities in cassava improvement. In this chapter the activity and tissue specificity of four Arabidopsis root promoters was evaluated in transgenic cassava. The four promoters: A14, E40, E49 and S8 are all expressed in different tissues of the Arabidopsis root (Table 1).

A Gateway Recombination Technology system was developed to easily evaluate the different Arabidopsis promoters and compare them to the Cauliflower Mosaic Virus 35S

(35S) promoter. The 35S promoter is a commonly used constitutive promoter in plants and it has been successfully used for strong constitutive expression in cassava of the uidA gene (Schopke et al. 1996, Gonzalez et al. 1998), the neomycin phosphotransferase gene

(Siritunga et al. 2003, Ihemere et al. 2006) and the hydroxynitrile lyase gene (Siritunga et al. 2004).

Two reporter genes were used to evaluate the relative expression of both the constitutive

35S promoter and the Arabidopsis promoters. The two variants of the Green Fluorescent

Protein known as Yellow Fluorescent Protein (Miyawaki et al., 1999) and Cyan

Fluorescent Protein (Cubitt et al., 1995) were selected to act as reporter genes. These two genes have been used extensively as reporters by taking advantage of the fluorescent properties of their proteins (Shaner et al. 2007, review) and analysis of the transcript levels of the genes (Smith et al. 2003 and Takenaka et al. 2007). The YFP and CFP

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reporter genes have been used in the past in plants in a variety of applications such as protein localization, promoter studies, fluorescent resonance energy transfer and more

(DeBlasio et al., 2010, review) and they differ sufficiently in their properties such as

DNA sequence (29 base pair differences in 780bp sequence) and wavelengths of fluorescent emission and excitation (YFP λexcitation= 514 nm and λemission= 527 nm; and

CFP λexcitation= 435 nm and λemission= 495 nm) that they may be expressed simultaneously and be distinguished from one another (Bauer et al., 2004; Kodama et al.,. 2009, examples). By introducing simultaneously the 35S promoter driving the expression of the

YFP gene and one of the Arabidopsis promoters driving the expression of the CFP gene, the expression evaluation system contains an internal standard in the form of 35S:YFP that allows a direct comparison of each Arabidopsis promoter to a strong constitutive promoter. In addition, it introduces a control that is subject to the same positional and copy number effects as the Arabidopsis promoter:CFP. It has been observed that copy number and positional effects dependent on the location of the integration of the T-DNA in the genome affect transgene expression (Halpin 2005).

Cassava plants possess two types of root systems, the fibrous root system is present from the early stages of growth and on and the storage root system is present in mature plants

(DeSouza et al. 2001). Evaluation of the four promoters was done on both fibrous and storage root tissues to determine the tissue specific expression. As our objective was to identify strong root promoters the leaves of the same plants were analyzed to determine organ specific expression.

The A14 promoter driven CFP expression was detected by RT-PCR in both fibrous root and leaves of 1.5 month old young cassava plants. The leaf expression appeared to be

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lower than that of the roots but it is possible that the A14 promoter is not root specific during the early stages of cassava growth. Importantly, the CFP fluorescence in the 1.5 month old A14:CFP plants was detected and localized through confocal microscopy to the epidermal layer of cassava fibrous roots. In the same plants there was no detectable

CFP fluorescence in the leaf epidermal tissue.

In cassava 4 month old storage roots of A14:CFP lines the analysis by real-time RT-PCR demonstrated CFP expression in both storage root peel and pulp, however, the CFP relative expression in the peel was 2 to 10 times that of the CFP relative expression in the pulp. In addition, real-time RT-PCR analysis of cassava storage root peel indicated that the A14 promoter has a strength similar to the constitutive 35S promoter. The presence of low levels of CFP transcript in the leaves of the 4 month old A14:CFP lines possibly indicates that this promoter is not entirely root specific.

While the storage root remains the most important organ of cassava it is also important to consider gene promoters for the fibrous root development phase as well, in particular for micronutrient (iron and zinc) biofortification as it has been observed that cassava fibrous roots have 10-100 times higher iron content than mature storage roots (Ihemere, personal communication). It is also known that the primary site of iron uptake in roots is the epidermal root hairs (Gilroy and Jones, 2000). However, root hairs are absent from storage roots and metal uptake is presumably not taking place in storage roots. A root epidermal promoter such as A14 may be capable of driving expression of a metal transporter during the fibrous root stage that would presumably increase metal uptake during the stage were accumulation occurs.

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The E40 promoter in Arabidopsis drives the expression of the Yellow-Stripe Like 2 gene

(YSL2), the YSL2 protein is a metal transporter responsible for lateral movement of metals in the vasculature (DiDonato et al. 2004). In 1.5 month old transgenic cassava of expressing the E40:CFP construct the CFP transcript was accumulated in both the fibrous roots and the leaves. Unlike A14:CFP plants the fluorescent CFP protein was not detected in the epidermal tissues of E40:CFP transgenics which correlates with the expected tissue-specific expression of this promoter in Arabidopsis (endodermis and stele). The tissue specificity of the E40 promoter in the cassava fibrous roots requires additional analysis.

In storage roots of 4 month old E40 transgenic plants the CFP transcript was expressed on average 7.35 times higher in the root pulp than in the peel. This may indicate a higher tissue specific expression of the E40 promoter in the storage root pulp. In addition, the

E40 promoter strength in the root pulp was comparable or higher to that of the constitutive 35S promoter.

Both the E40 and A14 promoters appear to drive transgene expression in cassava storage roots. In addition, these two promoters have different tissue specificity . The A14 promoter is localized to the peel and the E40 promoter is localized to the pulp. In either case there is a low expression in the leaves. However, with the exception of the A14-17 line the relative expression of these promoters is substantially less that 35S promoter driven transgene expression in the leaf.

The remaining two promoters E49 and S8 had no significant activity in cassava storage roots or leaves of four month old plants relative to the 35S promoter. Interestingly, in young 1.5 month old plants the E49 promoter was the single promoter with root specific

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expression. However in E49:CFP lines CFP fluorescence was not detected. These results suggest that there may also be temporal differences in gene expression that may not be easily detected at the protein level.

The bioinformatic analyses using the PLACE database (Higo et al. 1999) of the four promoters indicated that the A14, E40, E49 and S8 promoters had common as well as unique trans-acting factor binding domains, many of which are considered root-specific.

Interestingly the A14 promoter contained a unique Q-element (Hamilton et al. 1998) that has been shown to act as a promoter enhancer. Both the A14 and E40 promoters contain the RHERPATEXPA7 motif which is common to promoters of root hair expressed genes

(Kim et al. 2006) and the SP8BF motif which is found in sweet potato storage root expressed genes. These motifs may play a role in the root expression of these two promoters. The E40 promoter contained a total of 9 ABA related motifs. ABA is involved in a multitude of physiological processes including stress response (Giraudat et al., 1994) and it has been shown that ABA related cis-elements are sufficient to confer ABA- responsiveness in heterologous promoters (Skriver et al., 1991). In cassava ABA has been linked to the water deficit response (Alves et al. 2002 and 2004) however, no reports are available of cassava storage roots response to ABA. The ABA induced expression of the E40 promoter in cassava will require further characterization. Finally, the bioinformatic analysis determined that the S8 promoter contained a single copy of the

SURE-1 motif (Grierson et al. 1994) this motif is found twice in the patatin promoter, this promoter was mentioned before as one that is functional and specific in cassava storage roots (Siritunga et al. 2003, Ihemere et al. 2006), however, the patatin promoter not only contains two copies of the SURE-1 motif it has two copies of the SURE-2 motif,

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so it is possible that the presence of just one copy of a single motif is not sufficient for storage root expression in cassava. In addition, a set of three different sugar repressive elements were found in the S8 promoter sequence including the A-box, CGACG motif

(Toyofuku et al. 1998) and the SRE motif (Lu et al. 1998). The cassava storage roots have high starch content and it may be possible that the presence of these sugar repressive elements affected the ability of the S8 promoter to drive the expression of the

CFP reporter gene in cassava storage roots.

In summary, two new promoters for cassava storage root expression have been characterized, the A14 and E40 promoters from the Arabidopsis genes AT5G43040 and

AT5G24380, respectively. These two promoters are unique in their tissue-specific expression in cassava storage roots with the possibility of A14 being used for peel enriched expression and E40 for pulp enriched expression. In addition, the current available promoters for cassava root expression are native or are from tuber producing crops. In this study, we have confirmed the ability of heterologous root promoters from non-tuber crops to express in cassava. Such findings increase the number of promoter candidates that may be tested in the future for cassava root expression. And finally a

Gateway Technology based expression system was developed that may be used in the future to easily test additional promoters and compare them to the strong 35S constitutive promoter.

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