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Carotenoid pathway engineering in carrot and functional characterization of cytochrome P450 carotenoid… Kim, Ji-Eun 2007

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Carotenoid Pathway Engineering in Carrot and Functional Characterization of Cytochrome P450 Carotenoid Hydroxylases By Ji-Eun Kim A THESIS SUBMITTED IN PARTIAL FULFILLMENT O F THE R E Q U I R E M E N T S FOR THE D E G R E E O F DOCTOR O F PHILOSOPHY in THE F A C U L T Y O F G R A D U A T E STUDIES (Animal Science) THE UNIVERSITY OF BRITISH COLUMBIA August 2007 ©Ji-Eun Kim, 2007 Abstract Carotenoid pigments are synthesized by bacteria, fungi and plants for photosynthesis and photoprotection. The ketocarotenoid astaxanthin is a strong anti-oxidant and has also been exploited as fish feed additive to provide the attractive red colour in salmon flesh. Astaxanthin cannot be produced efficiently from natural sources and has to be chemically synthesized. I explored the feasibility of synthesizing ketocarotenoids in transgenic carrots. Using light and transmission electron microscopy, I confirmed that the biosynthesis and accumulation of carotenoids mostly occurred in the chromoplasts in the phloem tissue of carrot roots, and also found that increased accumulation of /3-carotene in roots of the High Carotene Mass (HCM) carrot variety was achieved by increased ^caro tene content in the chromoplasts and not by increasing the number of chromoplasts. An Agrobacterium binary vector system with the C a M V 3 5 S promoter directing expression of a crtO ketolase gene (isolated from the algal Haematococcus pluvialis) was used to transform H C M carrots. P C R analysis of genomic DNA from regenerated plants confirmed the insertion of the crtO gene in the carrot genome. Further work by our research team has detected novel ketocarotenoids: Astaxanthin, adonixanthin, adonirubin, canthaxanthin, ^-cryptoxanthin, and echinenone in the transformed carrots. It is of interest to examine the working relationship between the ketolase and hydroxylases in carotenoid biosynthetic pathways. I started by generating a single gene crtO transgenic line in Arabidopsis, three single gene carotenoid hydroxylase overexpression lines (AtB1, C Y P 9 7 A 3 , and CYP97C1) , and a single gene candidate carotenoid hydroxylase overexpression line (CYP97B3). HPLC-aided profiling of the carotenoid products in the leaves of crtO ketolase transgenic plants indicated no ketocarotenoid production. The three hydroxylase overexpression lines confirmed and also clarified the roles of the three hydroxylases in the carotenoid synthesis pathways in Arabidopsis. While I found that C Y P 9 7 B 3 has /3-ring hydroxylase activity and allowed biosynthesis of zeaxanthin and fi-ll cryptoxanthin in E.coli, the C Y P 9 7 B 3 overexpression line provided evidence for the first time that C Y P 9 7 B 3 gene product has hydroxylase activity primarily in the ocarotene pathway. I also co-transformed the crtO gene with each of the four hydroxylase genes to generate four co-transformants. In all co-transformants, hydroxylase activities in. the ^-carotene pathway were enhanced at the cost of hydroxylation activities in the /2-carotene pathway. iii Table of Contents Abstract ii Table of Contents iv List of Tables viii List of Figures • ix List of Abbreviations • xii Acknowledgements • xiii Co-authorship statement •• • xv Dedication — xvi CHAPTER 1 INTRODUCTION AND LITERATURE REVIEW 1 1.1 CAROTENOIDS • • 1 1.1.1 Carotenoids and animal health 2 1.1.2 Carotenoids in salmon aquaculture 2 1.1.2.1 Source of astaxanthin for fish feed supplement • 3 1.1.3 Carotenoids in higher plants 4 1.2 BIOSYNTHESIS OF CAROTENOIDS IN HIGHER PLANTS 4 1.2.1 Precursors of carotenoids 4 1.2.2 Carotene Synthes is 6 1.2.3 Xanthophyll Synthesis 11 1.2.3.1 Hydroxy lases 11 1.2.3.1.1 Non-heme di-iron monooxygenases in plants 12 1.2.3.1.2 Cytochrome P450-type hydroxylase and plant carotenoid biosynthesis 13 1.2.3.2 Ketolase modification of carotenoids 16 1.2.3.3 Epoxidase, de-epoxidase and nepxanthin synthase 19 1.2.4 Carotenoid Biosynthesis in Plastids • 20 1.2.4.1 Function of plastids in higher plants 20 1.2.4.2 Regulation of carotenoid biosynthesis in chloroplasts and chromoplasts 21 1.3 METABOLIC ENGINEERING OF CAROTENOIDS IN PLANTS 24 1.3.1 Genetic engineering of carotenoid biosynthesis in crop plants 25 1.3.2 Genetic engineering of ketocarotenoids in higher plants 27 1.4 Objectives and Organization of the Thesis • 28 1.5 REFERENCE • • 30 CHAPTER 2 ULTRASTRUCURE AND CAROTENE ACCUMULATION IN CHROMOPLASTS FROM DIFFERENT CARROT VARIETIES 48 2.1. Introduction 48 2.2. Materials and Methods •. 50 2.2.1. Plant, materials • 50 2.2.2. Light Microscopy 50 2.2.3. Transmission Electron Microscopy • 50 2.2.4. Quantitative analysis 52 2.3. Resu l ts •••• • • 53 2.3.1. Light microscopy and carotene accumulation in three carrot varieties 53 2.3.2. Comparative ultrastructure and carotene accumulation 55 2.4. D iscuss ion 61 2.5. References • 65 CHAPTER 3 AGROBACTERIUM-MEDIATED TRANSFORMATION OF THE ALGAL CRTO KETOLASE GENE INTO CARROT {DAUCUS CAROTA L.) 69 3.1. Introduction • • • 69 3.2. Materials and Methods 72 3.2.1. Sequence analysis 72 3.2.2. Binary vector construction 73. 3.2.3. Plant materials 76 3.2.4. Preparation of M S media 76 3.2.5. Agrobacterium-mediated crtO gene transformation into H C M carrot v 77 3.2.6. Regeneration of transformed plants (single C a M V 35S) and P C R confirmation 77 3.3. Results • 78 3.3.1. Confirmation of crtO gene cDNA sequence 78 3.3.2. Confirmation of the bar gene as a suitable selection marker in H C M carrot 79 3.3.3. P C R analysis of regenerated plant (single C a M V 35S-crtO) 81 v 3.3.4. Phenotype of transgenic carrot callus derived from Agro£>acter/'ivm-mediated transformation • • 83 3.4. Discussion • 85 3.5. References • 88 CHAPTER 4 CO-EXPRESSION OF ARABIDOPSIS THALIANA CYTOCHROME P450 ENZYME AND NADPH-CYTOCHROME P450 REDUCTASE IN ESCHERICHIA COLI: Testing the function of candidate ^carotene hydroxylase 93 4.1. Introduction 93 4.2. Materials and Methods 94 4.2.1. Plasmid construction and bacterial growth conditions 94 4.2.2. Extraction of carotenoid and H P L C analysis 94 4.2.3. S D S - P A G E and Western blot • • 95 4.3. Results • •  95 4.4. Discussion • 97 4.5. References 99 CHAPTER 5 FUNCTIONAL CHRACTERIZATION OF CYP450 CAROTENOID HYDROXYLASES AND /7-CAROTENE KETOLASE IN ARABIDOPSIS THALIANA • 100 5.1. Introduction - 1 0 0 .5.2. Materials and Methods 103 5.2.1. Binary vector construction • • 103 5.2.2. Plant transformation and selection 106 5.2.3. Genomic DNA 107 5.2.4. RNA isolation - 108 5.2.5. cDNA synthesis and semi-quantitative R T - P C R analysis 108 5.2.6. Pigment extraction and H P L C analysis • 109 5.3. Resu l t s 112 5.3.1. Select ion of transformed plants • 112 5.3.2. Transcript level of single and co-transformed plants 113 5.3.3. Carotenoid profiling •• 116 5.4. Discussion 123 vi 5.5 References : 129 CHAPTER 6 GENERAL DISCUSSION AND CONCLUSION 133 6.1 References -138 APPENDICES , 141 Appendix 1 •• 141 A1. Methods for Carotenoid Extraction from Carrot Root (from Chapter 3) 141 A1.1 Preparation of carrot root samples 141 A1.2 Extraction with two different solvent systems 141 A1.3 Carotenoid analysis 142 A1.4 Authentic standards for H P L C analysis 142 A1.5 References 143 Appendix 2 • 145 A2. Heterologous expression of P450 hydroxylases in E.coli (from Chapter 4) 145 A2.1 Cloning strategy 145 A2.2 Transformation strategy •••••• • 146 v i i List of Tables Table 2.1 Comparison among the three varieties of carrots in chromoplast numbers and carotene conecentration • 57 Table 3.1 Primers for sequencing of the double 35S-crtO construct (pKB-1) 76 Table 5.1 Sources of genes used in this study • 104 Table 5.2 Primers for gene cloning for the fusion binary vector constructs 105 Table 5.3 Primers for P C R confirmation of transgene presence in transformed Arabidopsis lines • 105 Table 5.4 Primers for R T - P C R measurement of transgene expression 109 Table 5.5 Profiling of carotenoids derived from ^-carotene in Arabidopsis wild-type plants and over expression lines 117 Table 5.6 Profiling of carotenoids derived from /r-carotene in Arabidopsis wild-type plants and over expression lines '• 118 Table 5.7 Summary of changes in carotenoid product accumulation in single hydroxylase over expression lines and hydroxylase overexpression lines co-transformed with the crtO ketolase • 119 Table A2.1 The gene specific primers used to isolate the coding region of each gene from cDNA of Arabidopsis : • • 147 Table A2.2 Primers for pETDUET-ATR1, pETDUET-97A3, 97B3 and 97C1 vector construction 148 v i i i List of Figures Figure 1.1 Biochemical structure of C40 carotenoid derived from C 5 isoprene 1 Figure 1.2 Cytosolic MVA pathway and Plastid M E P isoprenoid biosynthentic pathway 5 Figure 1.3 Four consecutive desaturation reactions in carotenoid biosynthetic pathway (plants and bacteria) • .• 8 Figure 1.4 Xanthophyll biosynthetic pathways 10 Figure 1.5 Ketocarotenoid biosynthetic pathway in Haematococcus. pluvialis 18 Figure 2.1 Light microscopy comparison of carotene accumulation between orange carrot and H C M carrot 54 Figure 2.2 Light microscopy comparison of carotene accumulation between orange carrot and H C M carrot (phloem regions) • 55 Figure 2.3 Cellular ultrastructure of carrot varieties 58 Figure 2.4 Chromoplasts in the phloem of three carrot varieties and amyloplasts in white carrot • •• 60 Figure 3.1 Biochemical pathway of astaxanthin biosynthesis - —•• 71 Figure 3.2 Single C a M V 35S-crtO binary vector construct 73 Figure 3.3 Schematic diagram of double C a M V 35S-crtO gene construct 75 Figure 3.4 Amino acid sequences of the Keto2 clone 79 Figure 3.5 Preliminary test of bar selection with H C M carrot 80 Figure 3.6 Effect of phosphinothricin (BASTA) on [CaMV35S:crtO] transformed callus 80 ix Figure 3.7 Regeneration of transgenic lines with CaMV35S-cr tO ; 82 Figure 3.8 P C R screening of CaMV35S-cr fO transgenic lines 83 Figure 3.9 Compar isons of color between C a M V 35S-crtO transformed callus and non-transformed callus • 84 Figure 3.10 Comparisons of color between double C a M V 35S-crtO transformed callus and non-transformed callus 85 Figure 4.1 Western blot analysis of CYP97A3 , C Y P 9 7 B 3 , and C Y P C 1 accumulation in E.coli strain C43 • • •' 96 Figure 4.2 H P L C analysis of carotenoid extracts 97 Figure 5.1 Representation of the xanthophyll pathways showing hydroxylated products derived from a- and ^ -carotene • 101 Figure 5.2 Transgene expression levels in individual lines harbouring single hydroxylase or ketolase transgenes 114 Figure 5.3 Transgene expression levels in lines harbouring both hydroxylase and ketolase t ransgenes 115 Figure 5.4 The potential enzyme function of ketolase in o«-carotene pathway in Arabidopsis transformed with crtO 128 Figure A1.1 The H P L C results of steamed carotenoid extraction method from carrot root 143 Figure A2.1 Cloning strategy for expression of three P450 genes (97A3, 97B3 and 97C1) and NADPH-cytochrome p450 reductase (ATR1) 145 x Figure A2.2 Transformation strategy for co-expression of CYP: :ATR1 in E.coli strain C43 producing /^-carotene as a substrate in vivo : 146 xi List of Abbreviations y#-Ohase1 Arabidopsis /?-ring carotenoid hydroxylase AtB Arabidopsis /3-ring carotenoid hydroxylase ATR Arabidopsis cytochrome P450 reductase bp base pair CrtR-b beta carotene hydroxylase CrtO beta carotene ketolase CaMV35S cauliflower mosaic virus 35S promoter cDNA complimentary deoxyribonucleic acid P450 cytochrome P450 CYP cytochrome P450 CPR cytochrome P450 reductase 2,4D 2,4-dichlorophenoxyacetic acid DNA deoxyribonucleic acid DW dry weight e.coli Escherichia coli HPLC hi performance liquid chromatography HPF hi pressure freezing HCM high carotene mass hr hour IPTG Isopropyl )8-D-1-thiogalactopyranoside kb kilobase kD kilodalton LB Luria-Bertani min minute NADPH nicotinamide adenine dinucleotide phosphate OD optical density PPT phosphinothricin PAGE polyacrylamide gel electrophoresis PCR polymerase chain reaction PVDF polyvinylidene difluoride RNA ribonucleic acid rpm rotation per minute s second SDS sodium dodecyl sulphate SE standard error TEM transmission electronic microscopy UV ultraviolet w/v weight per volume w/w weight per weight Acknowledgements When I started to write the acknowledgements, my memory of the past 6 and half years clearly went through my mind, and I became emotional and short of words (English words in particular) to express my appreciation. I have two mentors. Drs Kim Cheng and Carl Douglas. To me they are much more than thesis supervisors. Without Kim and Carl 's supports and protective care, I will not be at this stage of my thesis to enable me to write the acknowledgements. I would like to express my true respect and appreciation to Kim, my thesis supervisor. During the first year of my PhD program, culture shock and language problems made my university life very difficult. I must have used up several boxes of Kim's Scotties to wipe out my tears. Kim has a big, warm heart and endless patience. He has spoiled me and allowed me to be childish for the past 6 years, but I want to "grow up" to be just like him. I often question how I can handle my life in the future without his support. A very sincere appreciation to Carl, my research supervisor. One of luckiest things in my life is to meet Carl and to be able to work in his lab. I have learnt from Carl logical thinking and open mindedness. I regret that I have been slow in getting his scientific point of view and critical comments, and tested his patience by repeatedly asking the same questions. From him I learnt multi-tasking and how to enjoy life and put it beside the scientific world. Thanks Carl, for your North American way of ethical justice and your faith in me to carry out my research projects successfully. A million thanks to my thesis committee members, Drs Bob Devlin and Dave Higgs. Bob had brought up the critical idea of the carotenoid study for my PhD thesis. Dr Higgs gave me positive way of thinking and nice encouragements. Without their expertises and support, I could not have carried out this project. I would also like to express my thanks to the members of Carl's lab. This group of friends was so diverse and dynamic and I consider myself lucky to be one of them. I often stayed late in the lab just because of them. Special thanks to Dr Bjoern Hamberger, who shared lots of motivating discussions with me during working on the Arabidopsis experiment; To my lab bench mates, Bahram, Dave, Clarice and Eryang, thanks for your nice warm hearts, honesty and friendship. I will never forget our hiking and cross-country ski trips. Thanks to the "old xiii boys", Lee, Michael, Juergen and Weiya, and the "new boys", Sungsoo, Apurv and Jun. Carry on with the lab spirit. Since my project was broad-based, I had the opportunity to work at different labs. Thus, I also want to express my thanks to people whom I worked with in Botany, Animal Science at U B C , Department of Fisheries and Oceans Canada, and Simon Fraser University. I would like to thank Dr Andrew Riseman and Peter Kalynyak (Botanical Garden, UBC) for allowing access to the tissue culture facilities and for their kind assistance. Many thanks to Dr Mark Smith (UBC) who provided advice for my E.coli expression work. For carrying out my research project, Dr P. Simon (ARC/USDA, Wisconsin State Univ.) provided white and H C M carrot seeds, Dr R. Peters (Iowa State Univ.) the Arabidopsis P450 reductase clone, Dr N. Misawa (Biotechnology Institute Co) the pACCR16Acr tX, Dr Raju Datla (National Research Council Canada) the double CaMV35S promoter, Dr S. Mansfield (UBC) the pSM-1 vector and Dr J . Jayaraj (SFU) the single CaMV35S-ketolase construct. My thesis research was partly funded by an N S E R C Strategic Grant to Drs. Zamir Punja (Simon Fraser Univ), Carl Douglas and Kim Cheng (UBC) and a U B C Graduate Fellowship. Above all, I would like to share my joyful moments with my sisters and brothers in Korea. Their priceless love and care have been guiding me here. Thanks to my sweetie Froaddy for his love and understanding. It would have been a long and lonesome journey in a foreign country for me if not for the nice friendship of my Korean friends at U B C , Sooyeun, Romee, Seara, Jiyoung and Daekyun. xiv Co-authorship statement The chapters in this thesis have been written up as manuscripts to be submitted for publication. The manuscripts will have multiple authors and their contributions are as follows: Dr Kim Rensing (Biolmaging Laboratory, UBC) adapted the High Pressure Freezing technique to carrot root and carried out the specimen preparation process for the Transmission Electronic Microscopy work. I generated, analyzed, and interpreted the T E M images. Dr Neal Craft (Craft Technologies, North Carolina) carried out the H P L C analyses of the Arabidopsis transgenic lines, and supplied the data to me. Dr Bjoern Hamberger (Michael Smith Laboratory, UBC) provided valuable suggestions for laboratory procedures and contributed to research ideas throughout my thesis research. Dr Zamir Punja (Simon Fraser University) contributed to project development at its early stages and provided funding for some of the studies. xv Devote to my parents and younger brother, Sang-woo CHAPTER 1 Introduction and Literature Review 1.1 CAROTENOIDS Carotenoids are among the most widely distributed natural pigments. These fat-soluble red, yellow and orange pigments are synthesized by algae, higher plants and photosynthetic bacteria, where they are involved in the photosynthetic process. They also occur in some non-photosynthetic bacteria, yeasts, and fungi, where they may have a protective function against damage by light and oxygen (Taylor and Ramsay, 2005). More than 600 carotenoids have been characterized and defined by their chemical structures. The majority of carotenoids are derived from a 40-carbon polyene chain, which can be considered the backbone of the molecule (Figure 1.1) (Britton, 1995; Cunningham and Gantt, 1998). C 5 ^ ^ - C H 2 O P P » -^J-^" CH 2OPP DMAPP , IPP C40 Figure 1.1 Biochemical structure of C40 carotenoid derived from C 5 isoprene. C10 and C20 is assembled from molecules of the C5 (DMAPP and IPP) and C40 is derived by a head-to-head condensation of two C20. DMAPP: dimethylallyl pyrophosphate, IPP: isopentenyl diphosphate, IPI: Isopentenyl diphosphate isomerase (Cunningham and Gantt, 1998). 1 1.1.1 Carotenoids and animal health Although animals appear to be incapable of synthesizing carotenoids, they incorporate carotenoids from dietary sources. Carotenoids provide bright coloration, serve as antioxidants, and can be a source of human nutrients (Ong and Tee 1992; Britton, 1995; Frank eta/ . , 1996; Deming and Erdman Jr. 1999; Garcia-Asua et al., 1998; Sandmann, 2001). As a vital part of human diets, /^-carotene plays an essential role as the main dietary source of vitamin A and retinols (Bartley and Scolnik, 1995). Deficiency in vitamin A leads to xerophthalmia, blindness and premature death (Mayne, 1996). More than 100 million children are vitamin A-deficient and over five hundred thousand become blind every year (www.who.int/nut/vad.htm). Other carotenoids alleviate age-related disease when taken in sufficient quantities in the diet because of their powerful lipophilic antioxidant functions. For example, zeaxanthin and lutein protect against age-related macular degeneration (Seddon et al., 1994; Thomson et al., 2002) and high intake of red tomatoes which contain lycopene reduces the chance of prostate cancer and DNA damage in white blood cells (Giovannucci, 1999; Chen et al., 2001). Therefore, dietary carotenoids fulfill essential requirements for human health. However, overdoses can cause pathogenic conditions (Bendich and Olson, 1989). 1.1.2 Carotenoids in salmon aquaculture Salmon is an economically important aquaculture species and a valuable natural resource. Consumers prefer salmon with deep reddish pink flesh colour and are willing to pay a much higher price for the colour. The reddish pigments in salmon are flesh ketocarotenoids, astaxanthin (3,3'-dihydroxy-4,4'-diketo-y£carotene) and canthaxanthin (4,4'-diketo-y#-carotene), which occur in the natural diet of many aquatic species, including salmon, trout, shrimp and crustaceans through the food chain. Astaxanthin is absorbed and deposited in the flesh of salmon more readily than canthaxanthin (Torrissen et al., 1989; Choubert et al., 1994). 2 Therefore, it is necessary to supply astaxanthin in the salmon diet. In addition to flesh color, astaxanthin in the diets of salmonid fish (40-80 mg/kg of feed) provides protection of eggs from damage by UV irradiation and higher survival and grown rate of juveniles (Torrissen, 1984). 1.1.2.1 Source of astaxanthin for fish feed supplement In farmed salmon, where their natural food containing astaxanthin is not available, the reddish pigment of its flesh is currently achieved by the inclusion of chemically-synthesized carotenoid into commercial salmon diets as feed supplements, a practice that adds approximately 20-25% to the cost of feed (Torrissen et al., 1995). The demand for astaxanthin is annually increasing in the aquaculture industry. Annual sales of synthetic astaxanthin are estimated at more than $185 million for salmon and trout production (www.aquasearch.com). There is therefore a need to produce astaxanthin in a cheaper way for fish farms. In nature, astaxanthin biosynthesis has been observed in a limited number of organisms, e.g. in some marine bacteria, in the yeast Xanthophyllomyces dendrorhous (formally Phaffia rhodozyma) and some green algae (Johnson and Schroeder, 1995). While astaxanthin can be extracted from krill oil and X. dendrorhous (0.03 mg-0.8 mg/g, DW), the unicellular green alga Heamatococcus pluvialis has the highest level of astaxanthin accumulation (10-30 mg/g, DW) and seems to be the most suitable natural source. However, H. pluvialis is very expensive to cultivate because of restrictive culture conditions and a slow growth rate (Bubrick, 1991; Lorenz and Cysewski , 2000). High production costs of natural astaxanthin preclude it as a commercial product. An alternative cheaper source of astaxanthin will need to be found to decrease the feed cost in the salmon industry. 3 1.1.3 Carotenoids in higher plants In higher plants, carotenoids are synthesized and accumulate in plastids, where they are essential for plant viability because they function in the photosynthetic pigment-protein complexes of the reaction centers. Carotenoids play additional roles as accessory pigments in thermal dissipation of excess light energy (Demmig-Adams and Adams, 1992 and 1996; Ronen et al., 1999). Carotenoids also serve functions in photoprotection by quenching triplet-state chlorophyll molecules, singlet oxygen and other reactive species (Siefermann-Harms, 1987). In addition, carotenoids bring color to flowers and fruits as attractants to pollinators (Bramely, 2002). 1.2 BIOSYNTHESIS OF CAROTENOIDS IN HIGHER PLANTS Genes encoding the carotenogenic enzymes for nearly every step for the carotenoid pathway in higher plants have now been identified, sequenced and characterized (Cunningham, 2002). 1.2.1 Precursors of carotenoids Carotenoids are isoprenoids and generally consist of eight isoprene units joined together. The isoprenoid biosynthetic pathway provides intermediates for the synthesis of a multitude of natural products which serve numerous biochemical functions in plants (Lange and Ghassemian, 2003). All isoprenoids are derived from the basic C5-isoprene units, Isopentenyl diphosphate (IPP) and its isomer dimethylallyl pyrophosphate (DMAPP). Isopentenyl diphosphate isomerase (IRI) can reversibly interconvert IPP and D M A P P (Lichtenthaler, 1999) (Figure 1.1). Both IPP and D M A P P are derived from the cytosolic mevalonic (MVA) acid pathway and the plastidic nethylerythritol phosphate pathway, respectively (Figure 1.2). 4 A. Cytosol: MVA pathway Acetyl-CoA 1 HMG-CoA i MVA • DMAPP « • IPP 1 GPP (C10) I FPP (C15) I Sterols Brassinosteroids dolichol polyterpenes sesquiterpenes cytokinin B. Plastid: MEP pathway GA3 + Pyruvate 1 DXP i MEP DMAPP « • IPP • GPP (C10) 1 GGPP (C20) Gibberellins Tocopherols plastoquinones phylloquinones Chlorophylls Carotenoids (C40) 1 abscisic acid Figure 1.2 Cytosolic MVA pathway and Plastid M E P isoprenoid biosynthentic pathway. A . MVA pathway in cytosol. H M G - C o A , 3-hydroxy-3-methylglutaryl-coenzyme A; HMGR, hydroxymethylglutaryl-CoA reductase; MVA, mevalonic acid. B. Pathway for the biosynthesis of MEP-derived isoprenoids in plastids. GA-3P, glyceraldehyde 3-phosphate; M E P , methylerythritol phosphate; D X S , DXP synthase; DXR, DXP reductoisomerase; D X P , 1-deoxy-D-xylulose 5-phosphate; IPP, isopentenyl diphosphate; D M A P P , dimethylallyl pyrophosphate; G P P , geranyl diphosphate; F P P , farnesyl pyrophosphate; G G P P , geranylgeranyl diphosphate Several studies have demonstrated that IPP and D M A P P are used as common precursors in both the MVA and M E P pathways (Lichtenthaler et al., 1997; Zeidler et al., 1998). In the cytosol, MVA derived IPP and D M A P P are incorporated into brassinosteroids, triterpenes, sesquiterpenes, polyterpenes, dolichol, and the isoprenyl groups used for protein prenylation and cytokinin biosynthesis. In plastids, the M E P derived IPP and D M A P P are incorporated into carotenoids and chlorophylls, tocopherols, plastoquinones, phylloquinones, gibberellins and abscisic acid (Sauret-Gueto et al., 2006) (Figure 1.2). Carotenoids are exclusively synthesized in the central isoprenoid pathways within plastids* (chloroplast and chromoplast), while carotenoid biosynthetic enzymes are nuclear encoded (Taylor and Ramsay, 2005). As the first step in the biosynthesis of carotenoids in plastids, generanylgeranyl pyrophosphate ( G G P P ; C 2 0 ) is formed from IPP and D M A P P by generanylgeranyl diphosphate synthase (GGPS) . G G P P is used as a common substrate for biosynthesis of C 4 0 isoprenoids (Burkhardt et al., 1997) (Figures 1.1 and 1.2). Carotenoids are mainly composed of carotenes and xanthophylls. Carotenes typically contain only carbon and hydrogen such as ^-carotene, /^-carotene and lycopene. Xanthophylls are often modified by various oxygen-containing functional groups to generate compunds such as lutein and zeaxanthin (Zaripheh and Erdman, 2002). 1 .2 .2 Carotene Synthes is The first committed step for the C 4 0 carotenoid biosynthesis is the head-to-head condensation of two molecules of G G P P (C 2 0 ) catalyzed by the enzyme phytoene synthase (PSY) to produce the first carotenoid, phytoene (Figure 1.3). This pool of G G P P represents the metabolic link between the other C 4 0 isoprenoids and biosynthesis of carotenoids (Sandmann et al., 2006). Flux through the isoprenoid branch pathways such as those leading to carotenoids and gibberellins compete for G G P P substrate pools, and the Psy-1 gene plays a 6 key role at this bottleneck (Fray etal., 1995; Cunningham, 2002) (Figure 1.2). Fray et al. (1995) showed that overexpression of the PSY gene in tomato plants enhances lycopene production. In contrast, it results in a 30-fold reduction of flux into the gibberellin pathway and causes severe dwarfism due to the depletion of the endogenous precursor pool of G G P P , leading to decreased gibberellin levels. This study is a good example of the competition that occurs between different branch pathways and how enzyme levels can be used to redirect metabolic flux (Cunningham, 2002). The synthesis of phytoene takes place in the stroma, whereas later steps, the downstream pathways of carotenoid biosynthesis take place in the thylakoid membrane (Cunningham and Gantt, 1998; Cunningham, 2002). Genes encoding phytoene synthase have been identified and isolated from many plants: pepper (Kuntz et al., 1992), melon (Karvouni et al., 1995), Arabidopsis (Lange and Ghassemian, 2003), daffodil (Schledz et al., 1996), maize (Buckner et al., 1996) and marigold (Moehs et al., 2001). Two phytoene synthase (Psy-1 and Psy-2) genes have been found in tobacco (Busch et al., 2002) and tomatoes (Bartley and Scolnik, 1993; Fray and Grierson, 1993). Phytoene, a colorless compound, is unable to absorb light at visible wavelengths because of its conjugated double bond system. The number of conjugated bonds in carotenoids affects their light absorption properties and color. Usually, carotenoids with 7 or more conjugated double bonds can absorb UV light and give pigment colors of yellow, orange or red (Cunningham and Gantt, 1998). In the carotenoid desaturation pathway, phytoene undergoes four sequential desaturation steps, to produce phytofluene, ^-carotene, neurosporene and lycopene (Figure 1.3). The formation of the first two products is catalyzed by phytoene desaturase (PDS) and the latter two are catalyzed by ^-carotene desaturase (ZDS) in photosynthetic organisms. These desaturase enzymes catalyze similar dehydrogenation reactions by introducing four double bonds and thereby transform the colorless phytoene into the red-colored lycopene (Nievelsten 7 etal., 1995; Cunningham and Gantt, 1998) in its c/'s-form (Bartley etal., 1999). Recently, genes encoding carotene isomerase (CRTISO) have been isolated from Arabidopsis (Park et al., 2002) and tomatoes (Isaacson etal., 2002). Phytoene (C40) GGPP (C20) + GGPP (C20) PSY Plant Bacteria Phytofluene ^-carotene Neurosporene PDS Lycopene ZDS CRTI Figure 1.3 Four consecutive desaturation reactions in carotenoid biosynthetic pathway (plants and bacteria). P S Y : phytoene synthase, P D S : phytoene desaturase, ZDS : zeta desaturase, CRTI: bacterial phytoene desaturase (Cunningham and Gantt, 1998) 8 CRTISO catalyzes the conversion of c/s-lycopene to the transform of lycopene in a further enzymatic step. However, in non-photosynthetic organisms, the desaturation of phytoene directly to frans-lycopene is catalyzed by a single enzyme, bacterial phytoene desaturase (Crtl) (Hirschbergef a/., 1997) (Figure 1.3). The subsequent cyclization of lycopene is an important branching point in the formation of ocarotene and /2-carotene (Figure 1.4), catalyzed by lycopene f-cyclase (LCY-e) and lycopene /f-cyclase (LCY-b), respectively. These enzymes introduce cyclic end groups, ft- and e-rings to lycopene. Two /2-rings are ubiquitous, one y3- and one f-ring are common and two e-rings are uncommon. LCY-b catalyzes the introduction of the first /2-ring to the ends of lycopene to produce ^carotene (0-,) followed by the sequential addition of the second /3-ring to produce /2-carotene (73-, ft-), which can be subsequently is further processed to xanthophylls: zeaxanthin, violaxanthin, neoxanthin. ft-nngs of /^-carotene and its derivatives are the most abundant in nature, ^-carotene can be cleaved to form vitamin A, which is essential in the human diet for eye health (Cuttriss and Pogson, 2004). As the alternative branch, both LCY-b and LCY-e are required to convert lycopene to a-carotene (ft-, £-). First, LCY-e catalyzes the introduction of the f-ring at one end of lycopene to form ^-carotene (e-,) before a ft ring is added to the other end by /^-cyclase to form /^carotene. The f, /2-cyclic branch typically terminates in the formation of the oxygenated a-carotene derivative, lutein. So far, Lcy-b genes have been identified from Arabidopsis (Cunningham and Gantt, 2001), daffodil, tobacco and pepper (Hugueney et al., 1995). In tomato, two genes encoding lycopene-/i-cyclase have been isolated: Lcy-b1 and Lcy-b2. The latter shows chromoplast-specific expression and encodes a protein with an amino acid sequence that is 53% identical to that of LCY-b1 (Ronen et al., 2000). Lcy-e genes have been isolated from Arabidopsis (Cunningham and Gantt, 2001) and tomatoes (Ronen^era/., 1999). 9 Lycopene j?-Cryptoxanthin Antheraxanthin ZEP | | VDE Violaxanthin Neoxanthin Figure 1.4 Xanthophyll biosynthesis pathways. L C Y - e : lycopene e-cyclase, L C Y - b : lycopene ^ c y c l a s e , Cr tR-b: fi-nng hydroxylase, Cr tR-e: e-ring hydroxylase, Z E P : zeaxanthin epoxidase, NSY: neoxanthin synthase, V D E : violaxanthin de-epoxidase (DellaPenna and Pogson, 2006; Kim and DellaPenna, 2006) 10 Both LCY-b and LCY-e predicted proteins show significant identity at the amino acid level, with 30% identity. However, the LCY-e in romaine lettuce has the unique ability to catalyze the formation of bicyclic e, e-carotene: lactucaxanthin (Phillip and Young, 1995; Cunningham and Gantt, 2001). These carotenoids with cyclic end groups are essential to all known oxygenic photosynthesis organisms, and plants can apportion substrate to either the /?,/?-carotenoids that are essential for photoprotection or the /jU-carotenoids that serve primarily to capture light energy for photosynthesis (Cunningham et al., 1996; Pogson et al., 1996). 1.2.3 Xanthophyll Synthesis In higher plants, a-carotene and /?-carotene are further modified to produce xanthophylls, which are oxygenated derivatives of carotenes. Xanthophylls include lutein, zeaxanthin, violaxanthin and astaxanthin containing hydroxyl, epoxy and keto groups (Tian and DellaPenna, 2001; Cunningham and Gantt, 2005) (Figure 1.4). Xanthophylls comprise most of the carotenoid pigments in the thylakoid membrane of plants (Cunningham and Gantt, 1998; Cunningham, 2002). 1.2.3.1 Hydroxylases Hydroxylation of the ft- and e-rings of a- and /^-carotene are carried out by two hydroxylase enzymes: /3-ring hydroxylase and ^-hydroxylase (Cunningham and Gantt, 1998; Tian and DellaPenna 2004; Quinlan et al., 2007). Lutein (3R,3'R-ft, £-carotene-3,3'-diol), the most abundant of the xanthophylls in all plant photosynthetic tissues, is the major product derived from ^-carotene (ft, a-carotene). Zeaxanthin {3R,3'R-ft, y3-carotene-3,3'-diol), a component of non-photochemical quenching, is formed by hydroxylation of two /2-rings from ft carotene (ft, /J-carotene) (DellaPenna, 2001) (Figure1.4). Two types of monooxygenases have 11 been shown to be involved in carotenoid hydroxylation: (i) non-heme di-iron monooxygenase: /?-ring hydroxylase 1 and /?-ring hydroxylase 2 and (ii) cytochrome P450-type: /?-ring hydroxylase and ^ -ring hydroxylases. 1.2.3.1.1 Non-heme di-iron monooxygenases in plants: Non-heme di-iron monooxygenase type /^-hydroxylases have been cloned from bacteria, algae and higher plants (Hundle et al., 1993; Misawa et al., 1994; Sun et al., 1996; Masamoto et al., 1998). These all contain conserved histidine regions originally identified in membrane-localized fatty acid desturases, suggesting a membrane integral location in vivo (Cunningham and Gantt, 1998). These regions are signature motifs for non-heme di-iron monooxygenases (Shanklin et al., 1994). In these enzymes, the histidine motifs play a mandatory role in fatty acid desaturation since they are likely involved in iron coordination (Arnold and Haymore, 1991). Bouvier et al. (1998) also demonstrated that ferredoxin-dependent enzymes belong to the di-iron protein family and that all ten of the conserved iron-coordinating histidines are required for activity in pepper /^-hydroxylase. Higher plant /^-hydroxylases have been isolated and functionally characterized using E.coli heterologous expression or overexpression, knockout and gene silencing in planta in the following plants: Arabidopsis (Sun et al., 1996; Tian et al., 2001; Davison et al., 2002; Tian et al., 2004), Adonis (Cunningham and Gantt, 2005; Yu etal., 2006), pepper (Bouvier etal., 1998), tomato (Hirschberg, 2001; Galpaz et al., 2006), citrus (Kim et al., 2001). However, e-hydroxylase was genetically identified in only Arabidopsis (Pogson et al., 1996; Tian et al., 2003; Tian etal., 2004). The two copies of genes encoding /^-hydroxylase in Arabidopsis, /^-hydroxylase 1 (B1) and /^-hydroxylase 2 (B2) are expressed in all t issues, although B1 has higher expression levels than B2 (Tian and DellaPenna, 2001). Overexpression of endogenous B1 enhances 12 stress tolerance in Arabidopsis (Davison et al., 2002). Expression levels of hydroxylase genes are modulated by different intensities of white light during tobacco de-etiolation (Woitsch and Romer, 2003) and are strongly induced by excess light in Arabidopsis leaves (Rossel et al., 2002) . 1.2.3.1.2 Cytochrome P450-type hydroxylases and plant carotenoid biosynthesis: Another monooxygenase class has been suspected to function in carotenoid hydroxylation in plants. Cytochrome P450 enzymes, which are one of the largest superfamilies found in all organisms (Chappie, 1998; Archakov etal., 2001), were first detected from animal liver extracts as membrane-bound and heme-containing enzymes that show a characteristic absorption maximum at 450nm after binding to carbon monoxide in the dithionite-reduced form (Omura and Sato, 1964). These P450 enzymes contain two highly conserved domains: a conserved oxygen binding signature, and a conserved heme-thiolate binding signature (FXXGXXXCXG) that binds a single heme group with one iron atom in the center (Poulos, 1995; Tian and DellaPenna, 2004; Quinlan et al., 2007). Typical eukaryotic P450 enzymes contain single N-terminal transmembrane anchoring sequences which are usually bound to the endoplasmic reticulum or inner mitochondrial membranes in eukaryotes. In contrast, P450s in prokaryotes are found as soluble enzymes (Chappie, 1998; Danielson, 2002; Werch-Reichhart and Feyereisen, 2000). Many animal P450s are inducible enzymes, involved in detoxification of xenobiotics and have relatively broad substrate specificities (Bolwell etal., 1994; Bundock etal., 2003) . The biochemical reactions catalyzed by P450s are based on activation of molecular oxygen with insertion of one of its atoms into the substrate and reduction of the other to form H 2 0 , and most P450s require NAD(P)H for 0 2-dependent hydroxylation reactions (Werch-Reichhart and Feyereisen, 2000). During the cytochrome P450 catalytic cycle, N A D P H -cytochrome P450 reductase (CPR) is required and essential for the transfer of electrons. There 13 is only one C P R found in animals but are at least two C P R forms in plants (Ortiz de Montellano, 1995). Genes encoding Arabidopsis cytochrome P450 N A D P H reductases (ATR; ATR1 and ATR2) have been cloned and ATR1 was identified using a functional approach by complementation of a yeast strain deficient in ATR with an Arabidopsis cDNA library (Urban et al., 1997). ATR proteins contain signature FMA-, FAD- and NADPH-binding domains (Urban et al., 1997; Hull and Celenza, 1999). Plant P450s are generally classified in two main clades: the A-type and the non-A type (Durst and Nelson, 1995; Paquette etal., 2000). The P450s in the A-type clade are involved in the biosynthesis of diverse secondary metabolites or natural products found in plants such as phenylpropanoids, alkaloids, terpenes, lipids, cyanogenic glycosides, glucosinolates, brassinosteroids, gibberellins (Schuler, 1996; Bak et al., 2001; Ro et al., 2002 and 2005). The A-type P450s originate from, a single common ancestral gene (Durst and Nelson, 1995). In contrast, the non-A type clade is a much more divergent group of sequences consisting of several individual clades that often show more similarity to non-plant P450s than to other plant P450s (Paquette et al., 2000). In sequenced plant genomes, 272 cytochrome P450-encoding genes (246 CYP450 genes and 26 pseudogenes) have been identified in Arabidopsis (Nelson et al., 2004; http:Zwww.biobase.dk/p450) and approximately 455 cytochrome P450-encoding genes (356 C Y P 4 5 0 genes and 99 pseudogenes) in rice (Oryza sativa). However, the functions of most of these cytochrome P450s are still unknown (Nelson et al., 2004). Schoefs et al. (2001) demonstrated that cytochrome P450-type hydroxylases could be involved in carotenoid synthesis in photosynthetic organisms and suggested that the biosynthesis of astaxanthin might involve a cytochrome P450 enzyme because astaxanthin biosynthesis in the alga Haematococcus pluvialis requires common substrates and co-factors for cytochrome P450 (molecular oxygen, N A D P H and Fe 2 + ) . P450s with carotenoid hydroxylase function have been isolated from various organisms, including the bacteria Aphanocapsa 14 (Sandmann and Bramley, 1985), Thermus thermophilus (Blasco et al., 2004) and Brevibacterium linens A T C C 9175 (Dufosse and de Echanove, 2005), the yeast Xanthophyllomyces dendrorhous (Alvarez et al., 2006), the red alga Cyanidioschyzon merolae (Cunningham et al., 2007), and the higher plants: Arabidopsis (Tian et al., 2004; Kim and DallaPenna, 2006) and rice (Quinlan et al., 2007). Lutein b iosynthes is : In Arabidopsis, genetic studies suggest that the C Y P 9 7 family gene CYP97C1 has an £-ring hydroxylase function (Tien et al., 2004) and the related gene CYP97A3 has /3-ring hydroxylase function (Kim and DellaPenna, 2006; Fiore et al., 2006). The functions of f-ring specific hydroxylase (CYP97C1) and /?-ring hydroxylase (CP97A3) were demonstrated by isolation of mutants at the LUT1 and LUT5 loci, respectively. Tien at al. (2004) observed that while the /3, /r-xanthophyll derivatives in knockout mutants of B1, B2 and LUT1 (CYP97C1) are reduced by 80% relative to wild type, the monohydroxy Orcarotene derivative zeinoxanthin still accumulates to high levels. They suggested that there might be an unknown fourth hydroxylase enzyme with activity toward the /brings of p, e-and ft, /J-carotenoids. This result suggested the hypothesis that C Y P 9 7 A 3 which shares 49% amino acid identity to LUT1 (CYP97C1) might be such a hydroxylase. Consistent with this hypothesis, Kim and DellaPenna (2006) found that /3-ring hydroxylase activity of C Y P 9 7 A 3 (LUT5) contributes major in vivo activity toward the / k i n g hydroxylation of o-carotene fj3,e- carotene) and minor activity toward the /3-ring of /3-carotene (/?,/3- carotene) as well as /?,/?-xanthophylls (primarily violaxanthin and neoxanthin), and these were reduced by 35% in the Iut5 mutant versus a 76% reduction in the b1b2 genotype. Based on this result, B1 and B2 appeared to mainly prefer /^-carotene as the substrate, are able to hydroxylate the fi-r\ng of ^carotene as in the Iut5/lut1 double mutant, in which zeinoxanthin is still present at normal levels. Fiore et al. (2006) provided support for the results of the previous genetic studies by generating triple knockout b1b2lut5 mutants, which completely lack /?-xanthophylls (zeaxanthin, antheraxanthin, violaxanthin and neoxanthin). 15 In contrast, double b1b2 mutants show an approximately 70% reduction of the same compounds in leaves. Recently, other C Y P 9 7 family genes (CYP97A4, CYP97B4 and CYP97C2) have been isolated from rice (Oryza sativa) and used in functional complementation experiments to test their enzyme activities in E.coli(Quinlan era/ . , 2007), in which the endogenous bacterial P450 N A D P H reductase was used instead of rice C P R . This study reported that rice C Y P 9 7 A 4 (same clan as Arabidopsis 97A3) shows /^-hydroxylase activity and rice C Y P 9 7 C 2 (same clan as Arabidopsis 97C1) shows ^-hydroxylase activity, respectively. This result is similar to that described in one of the previous Arabidopsis genetic mutant studies (Tian et al., 2004; Kim and DellaPenna, 2006). 1.2.3.2 Ketolase modification of carotenoids In addition to modification by hydroxylation, addition of a keto group is necessary to form ketocarotenoids, a family of xanthophyll derivates of /^-carotene. Ketolase genes encoding /2-carotene ketolase (^-C-4-oxygenase) have been cloned from green algae and bacteria (Britton, 1998). The algal Haematococcus pluvialis ketolase gene is designated crtO or bk1 whereas the bacterial ketolase genes are generally designated crtW (Lotan and Hirschberg, 1995; Kajiwara et al., 1997; Huang et al., 2005). H. pluvialis crtO exclusively participates in the secondary carotenoid pathway to produce astaxanthin in cytoplasmic lipid vesicles outside the plastid (Grunewald et al., 2001), while its precursor /^-carotene is synthesized in chloroplast (Grunewald et al., 2000). It has been proposed that phytoene desaturase might transport intermediates from the site of early biosynthetic steps in the chloroplast to the site of oxygenation and accumulation in cytoplasmic lipid vesicles (Grunewald et al., 2001). Three crtO genes have been identified from H. pluvialis (Huang et al., 2005). Most notably in higher plants, the flowers of Adonis spp. accumulate.ketocarotenods at levels of up to 1% of dry 16 weight (Renstrom et al., 1981). Recently, the Adonis ketolase (Adoketo) gene has been cloned from the flower of A. aestivalis and the enzyme function characterized in both E.coli and Arabidopsis (Cunningham and Gantt, 2005; Yu ef al., 2006). The Adonis ketolase gene which contains transit peptide sequences enables to ketocarotenoid synthesis in the plastid (Cunningham and Gantt, 2005). Like the Arabidopsis /3-ring hydroxylases and the Adonis ketolase, the H. pluvialis ketolases are members of a large class of membrane-intergral, di-iron oxygenase enzyme (Cunningham and Gantt, 1998 and 2005). The protein sequence of H. pulvialis CrtO displays similar histidine motifs and this homology suggests that ketolase is mechanistically related to carotenoid hydroxylase (Bouvier et al., 1998). The key ketocarotenoid end product, astaxanthin (3,3'-dihydroxy-4,4'-diketo-/3/3-carotene), which is used as an additive in feed for pigmentation of fish and crustaceans, is formed from /3-carotene via two alternative pathways, by the actions of two enzymes: ketolase and /3-ring hydroxylase as discussed above (Cunningham and Gantt, 1998) (Figure 1.5). In one alternative pathway, ketolase catalyzes the addition of a keto group at the number 4 position of one or both /brings of the yellow /3-carotene to produce the reddish-orange to red pigments, echinionone (4-keto-/3,/3-carotene) and canthaxanthin (4,4'-diketo-/3,/3-carotene). A /3-ring hydroxylase then catalyzes the further addition of two hydroxyl groups at the 3, 3' positions to produce astaxanthin. The other pathway requires that the /3-ring hydroxylase first catalyzes the addition of a hydroxyl group at the 3 and 3' positions of one or both /3-ring(s) of /3-carotene to produce /3-cryptoxanthin (3,3'-hydroxy-/3,/3-carotene) and zeaxanthin (3,3'-dihydroxy-#/3-carotene). In this case, ketolase functions only to catalyze the addition of a carbonyl at carbon 4 of both /3-rings of zeaxanthin to synthesize astaxanthin (Lotan and Hirschberg, 1995; Cunningham and Gantt, 1998; Mann et al., 2000). Several studies showed that ketolase enzymes are active against substrates consistent with its participation in the late stages of astaxanthin biosynthesis. 17 ^Caro tene /S-Cryptoxanthin CrtO CrtR-b o- Echinenone CrtR b 3-hy droxyechi neno ne 3' - Hy d r o xy echinenone Astaxanthin Figure 1.5 Ketocarotenoid biosynthetic pathway in Haematococcus pluvialis. CrtO: /2-carotene ketolase, CrtR-b: /3-ring hydroxylase. CrtO and CrtR-b have important functions to produce astaxanthin and other ketocarotenoids (Lotan and Hirschberg, 1995; Mann etal., 2000). 18 In the flowers of Adonis aestivalis, the appropriate substrate for the Adonis ketolase (Adoketo) produced in recombinant form in E.coli appears to be the dihydroxy carotenoid zeaxanthin rather than /3-carotene (Cunningham and Gantt, 2005). In contrast, H. pluvialis recombinant CrtO enzymes expressed in E.coli are unable to utilize the zeaxanthin as a substrate (Lotan and Hirschberg 1995; Breitenbach etal., 1996), suggesting that the alternative route to astaxanthin proceeds with /3-carotene and echineone as the preferential substrates. 1.2.3.3 Epoxidase, de-epoxidase and neoxanthin synthase In the xanthophyll cycle of higher plants, light plays a very important role in further carotenoid metabolism: epoxidation and de-epoxidation. This consists of interconversion of zeaxanthin, antheraxanthin (5,6,-epoxy-5,6-dihydro-/3,/3-carotene-3,3'-diol) and violaxanthin (5,6,5',6'-diepoxy-5,6,5',6'-tetrahydro-/3,/3-carotene-3,3'-diol) (Figure 1.4). Under normal light conditions, when light can be safely utilized for photosynthetic electron transport, zeaxanthin epoxidase (ZEP) converts zeaxanthin to violaxanthin via antheraxanthin by introducing 5,6-epoxy groups to the 3-hydroxy-/3-rings. The ZEP gene from pepper is a monooxygenase that requires 0 2 and is active at approximately pH 7.5. The recombinant pepper Z E P protein expressed in E.coli requires N A D P H and ferrodoxin. Z E P localizes to the stromal side of the thylakoid membrane and is constitutively active (Bouvier et al., 1996). Under excessive light, the reverse reaction, addition of an de-epoxy group at the same 5, 6'-position of violaxanthin is rapidly activated to form zeaxanthin by the violaxanthin de-epoxidase (VDE), which is localized in the lumen of thylakoids and becomes activated by acidification of the lumen (Pfundel and Bilger, 1994; Hager and Holocher, 1994). Zeaxanthin is effective in the thermal dissipation of excess excitation energy in the light-harvesting antennae and thus plays a key role in protecting the photosynthetic system from damage by strong light. The inter-conversion of zeaxanthin and violaxanthin is known as the 'xanthophyll cycle'. Lack of 1 9 the xanthophyll cycle in the Arabidopsis mutant npql (non-photochemical quenchingl), owing to a null mutation in VDE, increases the sensitivity of the plants to intense light (Niyogi et al., 1998). In the last step of xanthophyll synthesis from /3-carotene, the conversion of violaxanthin to neoxanthin is performed by neoxanthin synthase (NSY). Genes that encode enzymes with limited N S Y activity were originally identified in tomato and potato based on the similarity to the capsanthin-capsorubin synthase (CCS) , which catalyzes the conversion of antheraxanthin and violaxanthin, respectively (Al-Babili etal., 2000; Bouvier etal., 2000). Surprisingly, the predicted amino acid sequence of N S Y from tomato is 99% identical to that of lycopene /3-cyclase (LCY-b), which is a bi-functional enzyme capable of converting both lycopene to /3-carotene and violaxanthin to neoxanthin (Bouvier ef al., 2000). In ripening pepper fruit, antheraxanthin and violaxanthin are further converted into the red ketocarotenoids, capsanthin and capsorubin by capsanthin-capsorubin synthase (CCS) (Bouvier et al., 1994). As an apocarotenoid, the phytohormone abscisic acid (ABA) is synthesized by cleavage of carotenoids. A B A plays roles in mediating seed maturation and dormancy and stress responses in plants (Tan era/. , 1997). 1.2.4 Carotenoid Biosynthesis in Plastids 1.2.4.1 Function of plastids in higher plants As mentioned above, carotenoid biosynthesis occurs in plastids (Rodriquez-Concepcion and Boronat, 2002). Plastids are semiautonomous organelles whose diverse functions include photosynthesis and biogenesis of micro and macromolecules (Bouvier et al., 1998). As such, they are able to transcribe and translate their own genome but are strongly dependent on imported proteins that are encoded in the nuclear genome, translated in the cytoplasm and imported into plastids (Joyard etal., 1998). 20 Depending on their morphology and functions, structurally distinct plastid types can be defined. Colorless leucoplasts, or amyloplasts, contain starch grains as storage reservoirs in roots and tubers. Elaioplasts are lipid-storing plastids. Chloroplasts, specialized for photosynthesis in leaves and green stems, and chromopjasts in non-photosynthetic organs can synthesize and accumulate carotenoids (Cuttriss et al., 2006). The plastids are able to inter-convert and can differentiate and de-differentiate into other plastid types accompanied by dramatic morphogenic changes. For example, in ripening tomato fruit, chloroplasts differentiate into chromoplasts (Ronen et al., 2000). In contrast, chromoplasts can also de-differentiate and revert back to chloroplasts under the proper storage and lighting conditions such as in pumpkin fruit (Devide and Ljubesic, 1974; Bramley, 2002). In other instances, etioplasts that are found in plants grown in the dark develop into chloroplasts via light-dependent mechanisms (Welsch et al., 2000). Amyloplasts differentiate into chromoplasts in tobacco nectaries (Horner ef al., 2007). 1.2.4.2 Regulation of carotenoid biosynthesis in chloroplasts and chromoplasts Carotenoid biosynthetic enzymes are plastid-localized but nuclear-encoded proteins, and are post-translationally imported into plastids (Rodriquez-Concepcion and Boronat, 2002; Sandmann, 2002). In higher plants, carotenoids accumulate and are sequestered within chloroplasts and chromoplasts. In chloroplasts, essentially all carotenoids are associated with light-harvesting complexes in green leaves, acting as accessory pigments by transferring energy to the photosystem reaction centers and also acting as quenchers of triplet excited states in chlorophyll molecules that are generated during photosynthesis (Demmig-Adams et al., 1996; Taylor and Ramsay, 2005). Since carotenoids are an essential part of the pigment-protein complexes in thylakoid membranes (Thayer and Bjorkman, 1992), the regulation of carotenoid biosynthesis in green tissues is therefore likely related to and integrated with chloroplast 21 development, chlorophyll formation, the level of carotenoid binding proteins, and thylakoid membrane lipid composition. Light is known to be a key factor in the regulation of carotenoid biosynthesis in chloroplasts. For example, in Arabidopsis and tomato plants, transfer from low light to high light induces a shift in the ratio between lycopene p- and e- cyclase m R N A levels (Hirschberg, 2001). Carotenoid biosynthesis in chromoplasts has been more extensively studied than in chloroplasts (Bramley, 2002). Large quantities of carotenoids accumulate in chromoplasts, where they provide color to plant organs such as the red, orange, and yellow colors found in many flowers, fruits and roots (Goodwin and Britton, 1988). As mentioned above, chromoplasts are often derived from fully developed chloroplasts during fruit ripening and flower development. Chromoplast biogenesis involves chlorophyll degradation, carotenoid accumulation, and the appearance of chromoplast-associated proteins (Kreuz eta/ . , 1982; Marano era/ . , 1993; Libal-Weksler era/ . , 1997). Regulation of carotenoid biosynthetic gene expression has been studied during tomato and pepper fruit ripening, flower development in daffodil (Al-Babili et al., 1996; Schledz et ai, 1996), tomato and marigold (Moehs et al., 2001), as well as in citrus fruit development and ripening (Ikoma et al., 2001). These studies showed that transcription of carotenogehic genes is both up-and down-regulated in association with carotenoid biosynthetic activity. Tomato is an important model plant for the study of carotenoid biosynthesis in chromoplast-containing tissues. During dramatic color changes of the fruit development, the induction of lycopene accumulation coincides with up-regulation of upstream carotenogenic genes (Psy, Pds, Zds and CRTSO) (Pecker etal., 1992 and 1996; Giuliano etal., 1993; Ronen et al., 1999). The increase in carotenoid biosynthesis in tomato flowers is correlated with up-regulation of Psy1, Pds and Lcy-b genes. However, the Lcy-e gene is expressed at a very low level in petals and anthers, resulting in the accumulation of very small amount of lutein 22 (Giuliano era/., 1993; Corona etal., 1996; Ronen etal., 2000). Tomato contains two Psygenes, PsyA which is up-regulated in chromoplasts during flower and fruit ripening, and Psy-2 which is up-regulated in chloroplasts of green tissues (Fraser et al., 1999). Using mutants, Galpaz et al (2006) reported that two tomato /3-ring hydroxylase genes (CrtR-b1 and CrtR-b2) are differently regulated: CrtR-b1 is constitutively expressed in leaves, whereas CrtR-b2 is exclusively up-regulated during flower development. These findings demonstrate a role for differential expression of duplicated genes in the chromoplast-specific carotenoid biosynthetic pathway. Marigold (Tagetes erecta L.) flower petals synthesize and accumulate carotenoids at levels greater than 20 times that in leaves (Moehs et al., 2001). Similarly, carotenoid accumulation in pepper fruits is correlated with increased mRNA levels of several carotenoid biosynthetic genes (Hugueney et al., 1996). In addition to regulation according to developmental stage (Bugos et al., 1999) and plant species (Romer ef al., 1993), carotenoid gene expression during chromoplast differentiation is also affected by environmental factors such as oxidative stress, which has also been shown to up-regulate expression of carotenoid genes in pepper (Bouvier et al., 1998) and redox potential (Bouvier et al., 1996; Audran et al., 1998; Steinbrenner and Linden, 2001; Cuttriss and Pogson, 2004). In addition to control of gene expression at the transcriptional level, post-transcriptional regulation of carotenogenic enzymes also occurs. In daffodil flowers, both P S Y and P D S were detected in inactive forms in the soluble fraction, but became active when they were bound to the membrane (Al-Babili etal., 1996; Schledz et al., 1996). Carotenoid-associated and binding proteins play important roles in the sequestration of carotenoids and post-transcriptionally affect the accumulation of carotenoids within chromoplasts (Vishnevetsky era/., 1999). Chromoplasts have developed unique mechanisms to sequester carotenoids within specific lipoprotein structures (Bartely and Scolnik, 1995; Vishnevetsky et al., 1999). It was suggested that these lipoprotein complexes are formed to sequester the carotenoids away from other structures to 23 avoid possible harmful affects (Deruere ef al., 1994). For example, pepper chromoplasts sequester carotenoids associated with the fibrillin protein as fibrils (Deruere et al., 1994), and tomato chromoplasts appear to sequester lycopene in crystalline form (Camara et al., 1995). During marigold flower development, ultrastructural changes occur during plastid differentiation and carotenoids accumulate in lipidic vesicles (Del Vil lar :Martinez et al., 2005). Vasquez-Caicedo et al. (2006) demonstrated that carotenoids are deposited in plastoglobular substructures in mango chromoplasts due to overproduction of lipids. Furthermore, this study suggested that o-carotene and /^-carotene accumulate in chromoplasts of carrot root primarily as frans-isomers in crystalline form with minor amounts of c/'s-isomers in plastoglobuli. It is assumed that the crystalline structure is responsible for the stability of the trans-carotene in carrot root. As a non-photosynthetic storage organ with high carotene content, carrot roots are a good model for studying ultrastructure and carotenoid accumulation in chromoplasts. 1.3 METABOLIC ENGINEERING OF CAROTENOIDS IN PLANTS Efforts for metabolic engineering of plants to produce novel compounds or for the improved production of endogenous compounds have made significant progress since the mid-1980s (DellaPenna, 2001). Plant metabolic engineering has relied on efficient protocols for /Agrobacfer/'um-mediated transformation of plants. The introduction of the carotenogenic genes of the bacteria Erwinia uredovora and Erwinia herbicola (Misawa et al., 1990, 1991, 1993 and 1995) has contributed to the dissection of plant carotenoid pathways. These bacterial carotenogenic gene clusters have been successfully introduced into plants (Shewmaker et al., 1999; Romer et al., 2000; Ducreux et al., 2005) and into noncarotenogenic microorganisms such as E. coli (Misawa eta/ . , 1990 and 1991) and Saccharomyces cerevisiae (Yamano et al., 1994). In addition to higher plants, algae and cyanobacteria are also good eukaryotic carotenoid models (Harker and Hirschberg, 1997; Liang et al., 2006). Therefore, a broad and 24 diverse array of carotenogenic genes from prokaryotic and eukaryotic organisms is available to introduce novel new carotenoid branch pathways into plants and microorganisms. However, there still remain questions regarding the functional interactions between the proteins encoded by endogenous genes in higher plants and proteins derived form exogenous (prokaryotic or eukaryotic) carotenogenic genes, as discussed by Sandmann et al. (2006). 1.3.1 Genetic engineering of carotenoid biosynthesis in crop plants With the awareness of the important roles played by carotenoids in improvement of plant nutritional quality for human health and enhanced plant tolerance to abiotic stress, genetic engineering of carotenoid biosynthesis has been successfully applied to staple crop plants (Romer and Fraser, 2005). Metabolic engineering of plant carotenoids has been carried out through different approaches: 1) "Overexpression" in higher plants of a bacterial, algal and plant carotenogenic gene encoding a rate-limiting step in the pathway (Burkhardt etal., 1997; Shewmaker etal., 1999; Fraser etal., 2002; Ducreux etal., 2005); 2) Pathway manipulation by the silencing of a competing biosynthetic step using antisense or co-suppression of targeted genes (Giuliano et al., 2000; Busch et al., 2002; Romer et al., 2002). Recently, rice (Ye et al., 2000), tomatoes (Romer etal., 2000), canola (Shewmaker etal., 1999) and potato (Morris etal., 2006) have all been successfully engineered for the enhancement of carotenoids with improved nutritional value. As model plants, tobacco and Arabidopsis are used for better understanding the regulation and biosynthesis of carotenoids (Mann et al., 2000; Tian et al., 2004; Kim and Dellapenna, 2006). A remarkable example of metabolic engineering of carotenoids in crop plants is "golden rice", engineered as an improved source of vitamin A (Ye et al., 2000). This result represents a breakthrough not only in the biotechnology of carotenoids but also in the public recognition of the potential of improvement of food nutritional value. Generally, mature rice endosperm is 25 capable of synthesizing G G P P but completely lacks carotenoids. To achieve /3-carotene biosynthesis, the daffodil phytoene synthase and lycopene /3-cyclase cDNAs under the control of the glutelin promoter to achieve endosperm-specific expression, and the E. uredovora phytoene desaturase [crtl) under constitutive control (CaMV35S promoter) were introduced into transgenic rice. The level of carotenoids formed in the transgenic endosperm was estimated at 1.6 ug/g endosperm which would provide 10-20% of the recommended daily allowance of p-carotene in 300 g of rice. Tomato represents a principal dietary source of the carotenoid lycopene which accumulates in chromoplasts during fruit ripening (Bramley, 2002; Carrari and Fernie, 2006). Expression of the bacterial E. uredovora phyteone synthase (crtB) gene in a fruit specific manner-resulted in a 2-4 fold increase in tomato fruit carotenoids (Fraser et al., 2002). When the E. uredovora phytoene desaturase (crtl) gene was constitutively expressed in tomato, however, /3-carotene content increased about three fold but the total carotenoid level was reduced. (Romer et al., 2000). Expression of the Arabidopsis Lcy-b gene under the control of the tomato Pds promoter in tomato led to a 5-fold increase in /3-carotene and orange colored ripen fruit (Rosati et al., 2000). When the Lcy-b and CrtR-b genes under the control of the fruit-specific Pds promoter were overexpressed in tomato, fruits of the transformants showed significant increases in /3-carotene, /3-cryptoxanthin and zeaxanthin levels (Dharmapuri et al., 2002). In contrast, when the endogenous tomato Lcy-b was placed in the antisense orientation under the control of the Pds promoter control, tomato transformants showed up to a 50% down-regulation of Lcy-b expression in ripening fruits and lycopene content of the fruit increased slightly while leaf carotenoids were unaffected (Ronen etal., 2000). Potato is the fourth most important source of calories after wheat, rice and maize (www.cipotato.org). /3-carotene (pro-vitamin A) and xanthophylls such as zeaxanthin and lutein are important for human nutrition. However, potatoes contain very low levels of these 26 carotenoids. In order to provide a better supply of carotenoids in potato, several critical studies have been carried out to improve the amount of total carotenoids and modify /J-carotene and xanthophyll content (Romer etal., 2002; Morris etal., 2006; Ducreux etal., 2005; Diretto etal., 2006). In potato tubers, zeaxanthin concentration could be increased up to 130-fold and total tuber carotenoids increased up to 5.7-fold using antisense inhibition of zeaxanthin epoxidase (Romer et al., 2002). When Lcy-e was silenced in a tuber-specific manner by antisens, p-carotene content increased up to 14 fold and total carotenoids up to 2.5-foid, however lutein levels decreased (Diretto etal., 2006). Carotenoid content in canola was improved using bacterial carotenogenic genes. A single gene transformation with the E. uredovora phytoene synthase (crtB), under control of the seed-specific napin promoter from Brassica rapa, resulted in a 50-fold increase in carotenoids (Kridl et al., 1991; Shewmaker et al., 1999). Genetic manipulation of carotenoid biosynthesis in carrot has been very rarely reported. However, E. herbicola carotenogenic genes were introduced into carrot, resulting in a 2-5-fold increase in /^-carotene content in the root (Hauptmann et al., 1997). 1.3.2 Genetic engineering of ketocarotenoids in higher plants The carotenoid pathway in higher plants can be used as a source of precursors for the synthesis of high-value ketocarotenoids (e.g. canthaxanthin, 4-ketozeaxanthin and astaxanthin). Mann et al. (2000) successfully engineered the nectary of tobacco flowers to accumulate ketocarotenoids, the first study to accomplish this in a higher plant. This study showed that expression of the Haematococcus pluvialis crtO gene under the control of the tomato Pds promoter into tobacco via y4arabactenj/'m-mediated transformation resulted in the accumulation of ketocarotenoids including astaxanthin. This result showed that the ketocarotenoid biosynthetic pathway can be engineered into non-ketocarotenoid producing plants. 27 Stalberg et al. (2003) showed that an Arabidopsis line over-expressing the endogenous Psy gene crossed with Arabidopsis expressing crtO under the seed storage promoter napA produced seeds of the cross containing ketocarotenoids (mainly 4-keto-lutein, canthaxanthin and adonirubin but very low levels of astaxanthin) (Lindgren etal., 2003). The accumulation of astaxanthin in plant non-photosynthetic storage organs has been rarely achieved. Studies to engineer ketocarotenoids in potato tubers have been carried out. When H. pluvialis crtO and bacterial crtB were co-transformed into potato, ketocarotenoids (ketolutein and astaxnathin) were synthesized (Morris et al., 2006). In another approach to engineer ketocarotenoids in the potato tuber, Gerjets and Sandmann (2006) transformed the ketoalse (crtW) gene from the cyanobacterium Synechocystis under control of the C a M V 35S promoter into a transgenic potato line which accumulated zeaxanthin due to an inactivated zeaxanthin epoxidase gene (Romer et al., 2002), resulting in accumulation of ketocarotenoids in leaves and tubers. Ralley et al. (2004) demonstrated that two astaxanthin biosynthetic genes (/3-hydroxylase: crtZ and /3-ketolase: crtW from the marine bacterium Agrobacterium aurantiacum) synthesize ketocarotenoids in both tobacco and tomato. Ketocarotenoids of transgenic tobacco leaves accumulated up to levels of 800 ug/g (DW), whereas in tomato, 50-fold lower levels were found in transgenic leaves and the maximal level in tomato fruit was 10 ug/g (DW). The major ketocarotenoid in both plants was echinenone. 1.4 Object ives and Organizat ion of the Thes is My thesis stemmed out of a project aimed at finding an economic way to improve flesh pigmentation of salmon. It was hypothesized that by transferring the /3-carotene ketolase gene into carrot, which is a rich source of the /3-carotene substrate, ketocarotenoids such as astaxanthin and canthaxanthin could be biosynthesized economically. The specific objective of my thesis research was therefore to examine ways to introduce a ketocarotenoid biosynthetic 28 pathway in carrot, using with both exogenous algal /5-carotene ketolase and plant cytochrome P 4 5 0 /J-hydroxylases. In Chapter 2, I examined the ultrastructure and ^-carotene accumulation in chromoplasts of the H C M (High Carotene Mass) carrot using light microscopy and transmission electronic microscopy. The H C M carrot accumulates high concentration of /^-carotene in its root and is the variety that I used for my transgenic study. I confirmed that the H C M carrot accumulates /r-carotene the same way as does the wild type orange carrot, and I explored different fixation protocols to improve the quality of the specimen preparation. This study was carried out with Dr Kim Rensing of the Bioimaging Laboratory at the University of British Columbia. In Chapter 3, I performed a pilot study of >4grobacterium-mediated transformation of an algal /2-carotene ketolase gene into carrot. The H C M carrot variety was used in this study. With the goal of engineering the production of novel ketocarotenoids in higher plants, C a M V 3 5 S -ketolase and double CaMV35S-ketolase constructs were made and used to test transformation efficiency. Single CaMV35S-ketolase transgenic plants were regenerated. Results from these studies indicated that the transformation of the ketolase gene alone was capable of modifying the H C M carrot to produce astaxanthin. In Chapter 4, I carried out functional testing of candidate /2-carotene cytochrome P 4 5 0 hydroxylase genes (CYP97 genes: C Y P 9 7 A 3 , C Y P 9 7 B 3 and CYP97C1) by heterologous expression in the non-carotenogenic organism Escherichia coli. Each P450 hydroxylase and NADPH-cytochrome P450 reductase were co-expressed in an E.coli strain engineered to produce /3-carotene. The result of this study was presented at the 14 t h International Conference on Cytochrome P450 and extended abstract was published in the Proceedings of the conference (Kim etal., 2005). 29 In Chapter 5, I tested the hypothesis that the Arabidopsis P450 hydroxylases can efficiently hydroxylate /3-carotene using an in planta system. The objectives of the this part of my thesis.were (1) to characterize hydroxylase function through overexpression of single genes (2) to examine if the ketocarotenoid astaxanthin would be produced by co-transformation of the P450 hydroxylases with ketolase and (3) to determine if there were any crosstalk between carotenoid biosynthesis pathways. 1.5 REFERENCES Al-Babili S, Von Lintig J, Haubruck H, Beyer P. 1996. A novel, soluble form of phytoene . desaturase from Narcissus pseudonarcissus chromoplasts is Hsp70-complexed and competent for flavinylation, membrane association and enzymatic activation Plant Journal 9,601-612. 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Inhibition of the non-mevalonate 1-deoxy-D-xylulose-5-phosphate pathway of plant isoprenoid biosynthesis by fosmidomycin. Z Natudorsch 53, 980-986. 47 CHAPTER 2 Ultrastructure and carotene accumulation in chromoplasts from different carrot varieties1 2.1. Introduction Carotenoid biosynthesis is an area of study of great interest due to the roles that carotenoids play as pharmaceuticals with cancer-reducing potential and in the prevention of cardiovascular and eye diseases (Van den Berg ef al., 2000). Domestic carrots (Daucus carota L.) contribute high amounts of dietary carotenes and have a pleasant flavor and colour. Orange carrot roots contain predominantly /^-carotene (up to 80% of total carotenoids) and provide up to 30% of the provitamin A consumed in the United States (Simon, 2000). There is a great deal of phenotypic diversity in cultivated carrots, and there have been numerous studies related to the nutrients, carotenoid content, volatiles, and sugars in different varieties of carrot roots (Alasalvar et ai, 2001; Surles et al., 2004). A High Carotene Mass (HCM) carrot variety has been genetically selected for producing high levels of carotene (>500 ug/g: Simon et al., 1985 and 1989; Simon and Wolff, 1987). Studies using reversed-phase high performance liquid chromatography and thin layer chromatography revealed that H C M carrot roots have more than three times the ^caro tene content than orange carrots (Simon and Wolff, 1987). H C M and a "carotenoid-free" white carrot variant (D. carota var. carota) were also instrumental for QTL (quantitative trait loci) analysis of genes controlling carotenoid content and genetic mapping of genes involved in carotenoid biosynthesis (Santos and Simon, 2002; Just et al., 2007). 1 A version of this chapter will be submitted for publication. Kim JE, Rensing K, Douglas CJ, Cheng KM. 48 While it has been postulated that carotenoids are synthesized and accumulated in chromoplasts in flowers, fruits, and roots (Whatley and Whatley, 1987), these postulations were based on very limited studies that were published over two decades ago (Frey-Wyssling and Schwegler, 1965; Ben-Shaul et al., 1968; Grote and Fromme, 1978) when improved transmission electron microscopy (TEM) techniques were not available. The fibrous nature of the carrot root and the extraction of carotenoids by the fixation solvents have been providing challenges to sample preparation for T E M studies. Recent developments in approaches for T E M sample preparation, for example high pressure freezing (HPF) (Fuchigama era/ . , 1997; Van Buggenhout et al., 2005) may offer the opportunity to reexamine the subcelluar localization of carrot root carotenoids, and the potential ultrastructural differences between different carrot varieties. Vasquez-Caicedo et al. (2006) examined utrastructural changes of chromoplasts in ripening mango mesocarp using light microscopy and T E M , and only briefly compared these results with chromoplast structure in carrot roots. A s far as we are aware, there has been no study that examines the relationship between chromoplast number and carotene accumulation in carrot roots, or carries out quantitative analyses of chromoplasts in carrot varieties with different carotene contents. The objectives of my study were to reexamine the ultrastructure of carotene-accumulating carrot root cells using HPF , which improves preservation of subcellular structures, prior to T E M , and to establish correlations between carotenoid levels in three carrot varieties and differences in cellular and chromoplast ultrastructure. My results confirm that the major form of carotene deposition is in the crystalline form in chromoplasts of cells in the root phloem, and that the white carrot variety is greatly deficient in chromoplast differentiation. The major apparent difference between H C M and orange carrot varieties was in carotene content per chromoplast, rather than differences in chromoplast number. 49 2.2. Materials and Methods 2.2.1. Plant materials Orange carrot (Scarlet Nantes, Daucus carota var. sativus) seeds were obtained from Garden Corner Company (Etobicoke, Ontario, Canada). Seeds of the H C M and white carrot (D. carota var. carota) were acquired from USDA (ARS, University of Wisconsin, Madison, USA). Seeds of the three varieties were planted in potting soil (Sunshine Mix 5; SunGro, Seba Beach, Alberta, Canada) at the same time and grown on a window bench top under natural light. 2.2.2. Light Microscopy Cross sections 0.5 pm in thickness that included phloem, xylem and cortex were cut from fresh root samples of H C M and orange carrots using a vibratome (Leica VT1000S, Leica, Germany). The sections were fixed in fixation buffer (2.5% glutaraldehyde in 0.1 M cacodylate buffer) and jnounted on the glass slides, observed and photographed with a Zeiss Axioplan 2 fluorescent light microscope (Carl Zeiss Microimaging Inc. US). 2.2.3. Transmission Electron Microscopy Three-and-a-half month old roots from orange, H C M , and white carrot varieties were bisected radially. Due to the lipophillic nature of carotene, all previously published electron micrographs showed substantial extraction of carotene through the normal EM fixation process of glutaraldehyde and osmium treatment (UBC Biolmaging lab protocol www.emlab.ubc.ca/p_em.htm). None of these electron micrographs was able to demonstrate carotene in situ. In this study, different methods for fixation were attempted in order to retain the carotene in situ. Microwave treatment (Hopwood et al., 1984; Boon et al., 1986; Giberson and Demaree Jr., 1999) to assist fixation, dehydration, and infiltration to reduce the negative 50 effects of each of these processes was tried. However, the carrot root tissue did not dehydrate fully, hampering infiltration and subsequent polymerization of the root tissue. This resulted in poor thin sections, with chatter and tears similar to those observed in earlier electron micrographs published by Frey-Wyssling and Schwegler (1965). Further attempts to use LR white and LR gold for embedding (Bozzola and Russell, 1999) did not alleviate the extraction problem. Both room temperature and low temperature embedding of these resins was not sufficient to retain carotene. In the final attempt to address the problem of extraction of carotene, high pressure freezing (HPF) was used (Rensing et al., 2002). Approximately 0.5 mm-thick radial longitudinal slices containing portions of phloem were hand-cut from each half and immediately immersed in 0.2 M sucrose. The slices in sucrose were placed under vacuum (20 inches Hg) for 15 minutes then high-pressure-frozen using a Bal-Tec HPM010 (Bal-Tec, Liechtenstein). Frozen samples were freeze-substituted with 2% osmium tetroxide in dry acetone for 120 hours, using a dry-ice and acetone bath which equilibrated at -80°C. The tissues were then warmed to -20°C in a freezer for 4 hours and to 4°C in a refrigerator for 4 hours, after which they were brought to room temperature. The tissues were transferred to fresh, dry HPLC-grade acetone for 1 hour. The samples were transferred through 25%, 50%, and 75% Spurr's resin in acetone for 2 hours each, then 3 changes of 100% Spurr's resin over 24 hours. They were polymerized overnight at 60°C. The colour of the carrot root was retained. Blocks were sectioned at 70 nm thickness for T E M . Sections mounted on Formvar-coated grids were stained with aqueous 2% uranyl acetate for 20 minutes, followed by Reynold's lead citrate for 10 minutes, then observed and photographed on a Hitachi H7600 T E M (Hitachi High-Technologies Canada, Inc). While the ultrastructural morphology was improved and sectioning artifacts reduced compared to published carrot root electron micrographs, /3-carotene was no more evident than it was through conventional preparation techniques. 51 This work was carried out at the Biolmaging Laboratory (University of British Columbia) with the collaboration of Dr Kim Rensing, who adapted the High Pressure Freezing technique to carrot root for specimen preparation for the T E M work. 2.2.4. Quantitative analysis For each carrot variety, T E M images of good quality phloem areas were selected. To quantify chromoplast numbers, grids were randomly selected from these sections. Random starting points at the left edge of the grid were chosen and 10 sequential frames (at 15,000X magnification, the area of each frame was 10 microns x 10 microns) from left to right moving horizontally were photographed. A total of 15 sets of frames were photographed for each variety. From the electron micrographs, the number of chromoplasts and the number of amyloplasts in each frame, were recorded. Chromoplasts were identified by sub-organelle features such as crystal shaped spaces, lipid droplets and plastoglobuli, and with reference to previously published electron micrographs (Grote and Fromme, 1978; Frey-Wyssling and Schwegler, 1965; Vasquez-Caicedo et al., 2006). Amyloplasts (non-pigmented plastids for starch storage) on the other hand, were filled with starch grains, with very little other observable sub-organelle structure and totally void of crystal shaped spaces (Frey-Wyssling and Schwegler, 1965). To quantify carotenoid content in chromoplasts, the 5 sets showing a suitable number of chromoplasts from the total of 15 sets of each variety were selected. Three to five chromoplasts from each set were photographed (at 30,000 to 100,000X). From the electron micrographs, the area occupied by each chromoplast, and the area occupied by the extracted carotenoid crystals within each chromoplast were measured using the Image J software (version 1.37, National Institutes of Health, USA). Carotenoid content was normalized per chromoplast unit area in each chromoplast to obtain an estimate of carotenoid concentration in chromoplasts, as follows: 52 Concentration = Area of crystal spaces / area of chromoplast The differences among the three varieties in carotene content per chromoplast (concentration) and in chromoplast number per unit area were analyzed by Least Squares Analysis of Variance using the J M P software (version 5.1.2, S A S Institute, Cary, NC). Before analysis, chromoplast data were square-root transformed and the carotene concentration data were arc-sine transformed to maximize normality. 2.3. Results 2.3.1. Light microscopy and carotene accumulation in three carrot varieties Carotene concentration in the phloem of carrot roots is reported to be higher than that in the xylem of the root (Booth, 1951; Wu and Salunkhe, 1971; also see Figure 2.1). In order to obtain an overview of the distribution of carotenoids in carrot roots, I prepared thin cross-sections from fresh roots, and visualized carotenoid location by light microscopy, comparing the normal orange carrot variety (Scarlet Nantes) with a variety with much higher carotenoid content (HCM variety). Visual inspection of these images could not determine that carotene content, visualized by the orange color of carotene, was higher in the H C M carrot than the orange carrot (Figures 2.1 A and 2.1 B), but higher levels of carotene were evident in the phloem than in other parts (xylem and cortex) of the root in both orange (Figures 2.1C and 2.1 E), and H C M (Figures 2.1 D and 2.1 F) carrots. Higher magnification of the phloem regions (Figures 2.2) showed the colour and structure typical of carotene (Frey-Wyssling and Schwegler, 1965; Zhou etal., 1996; Vasquez-Caicedo et al., 2006) in both varieties. Thus, my light microscopy study confirmed that the H C M carrot variety has the same pattern of carotene distribution in the root tissue as the orange carrot variety. 53 Figure 2.1 Light microscopy comparison of carotene accumulation between orange carrot and H C M carrot. A, C, E, orange carrot; B, D, F, H C M carrot. A, B; cross section of root (phloem, xylem and cortex) (4x), C, D; phloem and cortex (10x); E, F; xylem (40x). Cr: cortex, Xy: Xylem, Ph: Phloem. Images are at 4X, 10X, 40X magnification. 54 < r B is 4* 4 i f , I m Yi1 Figure 2.2 Light microscopy comparison of carotene accumulation between orange carrot and H C M carrot (phloem regions) A. orange carrot and B. H C M carrot are shown. Images are at 40x magnification 2.3.2. Comparative ultrastructure and carotene accumulation Based on the light microscopy results, I focused my further attention on carotene accumulation in the phloem of all three carrot varieties (orange, H C M , and white), using transmission electron microscopy (TEM) to visualize the ultrastructure of carotene-containing cells in the phloem. Previous studies on the ultrastructure of such cells relied on chemical fixation of samples before visualization by T E M . The highly lipophylic carotene molecules are unavoidably extracted during fixation. Therefore, clearly defined non-staining spaces in T E M images left by such carotene removal have been accepted as presence of carotene (for example, Frey-Wyssling and Schwegler, 1965; Vasquez-Caicedo et al., 2006). We used an alternative method, high pressure freezing (HPF), during the fixation process but failed to retain carotene in the cells. Nevertheless, the use of H P F can minimize damage to sub-cellular structure, which may occur with chemical fixation (Fuchigami etal., 1997). 55 Images at low magnification (3,000x and 15,000x), provided an overview of the carrot root phloem tissue and cell morphologies, respectively. Cell morphology in the phloem from the orange (Figure 2.3A) and H C M carrots (Figure ,2.3C) was similar. Both the orange and H C M cells showed very well organized subcellular organelles including chromoplasts. The white carrot cells appeared to lack this organization but large starch grains were observed (Figure 2.3E). White carrot had very few chromoplasts and they were very small and electron dense compared to those of the H C M and orange carrots. In their place, there were numerous amyloplasts, which appeared to be filled with starch.(Figure 2.3F). The ultrastructure of cells in the phloem from the orange and H C M carrots was similar (Figures 2.3B and 2.3D). There was no evidence of crystal shaped spaces outside of the chromoplasts in all three varieties. To determine if the difference in carotene content between the orange and H C M carrot varieties is correlated with chromoplast number, I compared the relative chromoplast numbers in the three carrot varieties. As described in Materials and Methods, chromoplasts were counted in sets of captured frame images of equal area within randomly chosen grids, and results expressed as number of chromoplasts per frame (100 nm 2). While there was variation in chromoplast number between the three varieties, analysis of variance showed that there was no significant difference (P > 0.05) between H C M carrot (0.41 ± 0.05 chromoplasts per 100 pm 2 ; 0.28 ± 0.03 after data transformation) and orange carrot (0.23 ± 0.06 chromoplasts per 100 pm 2 ; 0.20 ± 0.04 after data transformation) in chromoplast numbers per unit area. However, white carrot (0.04 ± 0.06 chromoplasts per 100 pm 2 ; 0.04 ± 0.04 after data transformation) had significantly (P < 0.05) fewer chromoplasts than the other two varieties (Table 2.1). Amyloplasts filled with starch grains were observed in white carrot but not in H C M and orange carrots (see below). Chromoplasts in all three carrot varieties usually contained a single but sometimes a few starch grains. 56 Table 2.1 Comparison among the three varieties of carrots in chromoplast numbers and carotene concentration . Carrot Varieties White Orange H C M Mean number of Chromoplasts/frame (100 pm 2) 0.04 ± 0.06 b 0.23 ± 0.06 a 0.41 ± 0 . 0 5 a Mean number of Chromoplasts + Amyloplasts /frame (100 pm 2) 0.34 ± 0.08 a 0.23 ± 0.07 a 0.41 ± 0 . 0 7 a 1 Carotene concentration (Area of crystals/area of chromoplast) 0.01 ± 0.009 0 0.05 ± 0.007 b 0.10 ± 0.007 a Within each row, means followed by the same letter were not significantly different. Means followed by different letters (a,b,c) were significantly (P< 0.05) different by Least-squares Analysis of Variance. Since both chromoplasts and amyloplasts develop from proplastids (Horner et al., 2007), the number of chromoplasts and amyloplasts were pooled in the white carrot and entered again into the analyses. The number of chromoplasts plus amyloplasts per frame was not significantly different among the three carrot varieties. These data suggest that the primary location of carotene in cells of the phloem is in chromoplasts but the high amount of carotene in the H C M carrot did not result in higher number of chromoplasts in the cells. The data also suggest that high carotene content in the H C M carrot is more strongly associated with differences in chromoplast carotene content in the H C M carrot. To investigate this in more detail, I generated and analyzed higher magnification images of the chromoplasts of each variety. 57 Figure 2.3 Cellular ultrastructure of carrot varieties. Transmission electron micrographs of the phloem region of orange, H C M , and white carrot varieties are shown. A, C , E, magnification at 3,000x; B, D, F, magnification at 15,000x. A, B, orange, C, D, H C M , and E, F, white varieties. Sc : Single cell, C m : Chromoplast, Am: Amyloplast, S: starch (ACE, bar: 10 um, BDF, bar: 2pm) Representative high magnification images of chromoplasts from orange, H C M , and white carrot varieties are shown in Figure 2.4. In chromoplasts of the orange carrot, crystal-shaped spaces derived from the apparent removal of crystalline carotene were clearly observed (Figure 2.4A). In the chromoplasts of H C M carrots (Figure 2.4B), similar crystal-shaped areas were seen. In contrast, the few chromoplasts from the white carrot were small and lacked such crystal-shaped void areas, consistent with a lack of carotene in this variety (Figure 2.4C). In chromoplasts from both orange and H C M carrots, plastoglobuli were also observed, but they were relatively small in size and number. Plastoglobuli accumulate p-carotene c/'s-isomers and have been found, in abundance in mango fruit. In carrot root, however, plastoglobuli are very small in number and carry a minute amount of carotene cis-isomers (Vasquez-Caicedo et al., 2006). Chromoplasts of carrot root accumulate predominantly crystalline carotene, which are trans-isomers (Vasquez-Caicedo et al., 2006). As well, in well developed chromoplasts from H C M and orange varieties, a few starch granules were usually observed. This was in contrast to the abundant amyloplasts in the white variety, which were filled with abundant starch grains (Figure 2.4D). As a measure of carotene content, I calculated the area of crystal-shaped void areas, presumed to be the locations of crystalline carotene deposition (Vasquez-Caicedo et al, 2006), in the chromoplasts of all three varieties (see Materials and Methods). To obtain estimates of carotene concentration in chromoplasts, I normalized these data to chromoplast cross-sectional surface area and expressed content as a proportion of the chromoplast cross-sectional surface area (see Materials and Methods). Based on these measurements, Table 2.1 indicated that the H C M chromoplasts had significantly (P < 0.05) higher carotene concentration (0.10 ± 0.007 of the surface area occupied by the chromoplast) than chromoplasts from the orange carrot (0.05 ± 0.007 of the surface area). 59 Figure 2.4 Chromoplasts in the phloem of three carrot varieties and amyloplasts in white carrot. A. chromoplast of orange carrot, B. chromoplast of H C M carrot, C. chromoplast of white carrot, and D. amyloplast of white carrot (bar: 500nm) S: Starch, P: plastoglobuli, * : space left by apparent removal of crystalline carotene 60 Both varieties had significantly (P < 0.05) higher carotene concentration than chromoplasts from the white carrot (0.01 ± 0.009 of the surface area). Because in each section, the chromoplasts were sectioned at various angles, and the magnification also varied, I could not estimate the volume of the chromoplasts. 2.4. Discussion Carrots are a well-known natural source of /3-carotene, which serve as provitamin A (retinol) compounds in the human diet. Carotene has been postulated to accumulate in root chromoplast in crystalline form (Straus, 1950; Frey-Wyssling and Schwegler, 1965; Grote and Fromme, 1978). However, it has not been well established whether the root chromoplasts are the sole sites of synthesis and accumulation. of carotene. Furthermore, the relationship between amount of carotene and carotene crystalline accumulation in chromoplasts of carrot roots has been questionable. In more recent studies on /^-carotene accumulation in carrot roots, Baranski et al. (2005) and Baranska et al. (2006) used near-infrared (NIR) excited Fourier transform (FT) Raman spectroscopy to localize and differentiate different carotenoids at the tissue level but not at the sub-cellular level. Carrot root chromoplasts are classified as crystalline chromoplasts (Frey-Wyssling and Schwegler, 1965; Paolillo ef al., 2004; Vasquez-Caicedo et al., 2006) and accumulate carotene predominantly in its frans-isomer crystalline form based on observations made using polarized microscopy (Frey-Wyssling and Schwegler, 1965). My study confirmed that a variety of orange carrot (Scarlet Nantes), with normal levels of carotene, accumulates highest levels of carotene cells of the carrot root phloem, and shows that H C M carrots share this property. Within the phloem tissue, both orange and H C M varieties contain numerous chromoplasts, 61 distinguished by void crystal shaped areas that are the apparent sites of crystalline carotene accumulation. It is interesting to note that with the application of HPF , the orange and H C M root tissue remained yellowish after fixation, unlike those fixed with conventional chemical fixation, which were bleached white. Yet crystalline carotene was still not observable. Our data (Figure 2.4) show that carotene accumulates in chromoplasts of the H C M variety the same way that orange carrot does. Vasquez-Caicedo et al. (2006) suggested that in carrot root, low amounts of cis J3-carotene accumulate in the plastoglobuli. My study also showed that there were very few plastoglobuli in chromoplasts of H C M and orange carrots, and none was observed in white carrot chromoplasts. In contrast, mango chromoplasts showed numerous plastoglobuli varying in size and electron density. They were the main site of carotenoid accumulation, which contributed to the partial solubilization of the pigments in lipid droplets (Vasquez-Caicedo etal., 2006). It is assumed that the frans-isomer carotene is more stable (Marx etal., 2003) and both the nature of the structure in which carotenes accumulate and the crystalline state of the pigments are crucial for the stability of the trans configuration (Vasquez-Caicedo et al., 2006). Cells in the phloem of white carrots had a very small number of chromoplasts and these chromoplasts seem to be small and electron dense. Klein and Ben-Shaul (1967) postulated that the dense inclusions in the chromoplasts of white carrot.were residues of disintegrating membranes. Recently, Surles et al. (2004) detected very small amounts of (3-carotene inwhite carrot roots (0.06 pg/mg compared with 128 pg/mg in orange carrot). The /5-carotene content of white carrot roots was therefore about 0.05% of that of orange carrot root. My study supports the findings of Surles et al. (2004). I also observed crystal shaped void spaces in chromoplasts of white carrot and found that 1% of the cross-section area of white carrot chromoplast was taken up with such crystal-shaped spaces (compared with 5% in orange carrots). 62 When I took into consideration that white carrot phloem tissue has 0.04 chromoplasts per 100 pm 2 and orange carrot phloem tissue has 0.23 chromoplast in the same area, I estimated that carotene content of white carrot root was 3% of that of orange carrot. The correlation of the lack of crystal-shaped spaces with the very low carotene content of the white carrot roots supports the interpretation that these void crystal-shaped areas in the chromoplasts of orange and H C M roots represent the sites of crystalline carotene accumulation. I observed a large number of amyloplasts in the cells in the phloem of white carrot but not in H C M or orange carrots. Frey-Wyssling and Schwegler (1965) observed amyloplasts in very young orange carrot roots but then during the root development, the starch grains and the amyloplasts started to disappear when crystalline carotene started to accumulate. Grote and Fromme (1978) observed that starch grains and amyloplasts almost completely vanished when carrot roots were stored. In tobacco floral nectaries, amyloplasts were converted to chromoplasts when /3-carotene started to accumulate in them (Horner et al., 2007). Santos et al. (2005), using a statistical approach to analyse the correlation among various compounds in the carotene biosynthesis pathway in white carrot, suggested that the synthesis of phytoene, a precursor of /3-carotene, was blocked. It is therefore likely that in orange and H C M carrot roots, both chromoplasts and amyloplasts differentiate from proplastids and the amyloplasts are then converted to chromoplasts later during development when carotene starts to accumulate. On the other hand, white carrot root has very little carotene and most of their amyloplasts continue to develop for starch storage rather than developing into chromoplasts. These data suggest that a developmental signal is derived from carotene accumulation in plastids, and that this signal directs a developmental transition from amyloplasts into chromoplasts. 63 My quantitative analysis of chromoplast number and carotene concentration shows increased carotene accumulation in the chromoplasts of H C M carrot roots, but there were not more chromoplasts in the H C M variety compared with the orange variety. Thus, my data suggest that a key difference in the two varieties with respect to carotene content is the level of crystalline carotene per chromoplast. Like the data on chromoplasts of the white carrot variety, the comparative chromoplast ultrastructural data from orange vs. H C M carrot varieties support the interpretation that the void crystal-shaped areas in chromoplasts represent the sites of crystalline carotene accumulation, since there is a strong correlation between the areas of these structures and known ^-carotene content of the two varieties. Wu and Salunkhe (1970) predicted that any increase in carotene content in carrot roots would be due to increases in the number and size of carotene-containing crystals and plastoglobuli in chromoplasts. It is therefore unlikely that selective breeding for increase production of carotene (Stein and Nothnagel, 1995) would alter the number of chromoplasts after the cells are displaced from the meristem. On the other hand, while evolution favoured the clustering of related pathway loci in carotenoid biosynthesis (Santos and Simon, 2002), the expression of these genes may be more easily regulated (Wurbs et al., 2007). Thus, selective breeding may have up-regulated the expression of genes in the pathway to increase carotene biosynthesis, resulting in the increased crystalline carotene content of the H C M chromoplasts. My data on the localization of /^-carotene in carrot root cells may be significant relative to metabolic engineering of carotenoid biosynthesis. The localization of the site of /^-carotene accumulation in H C M carrot to chromoplasts suggests that novel proteins introduced into carrots for metabolic engineering of the /3-carotene biosynthetic pathway should be targeted to chromoplasts (Daniell et al., 2005; Galpaz et al., 2006; Ruhlman et al., 2006; Wurbs et al., 2007). 64 2.5. References Alasalvar C, Grigor JM, Zhang D, Quantick PC, Shahidi, F. 2001. 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Carotenoid biosynthesis structurural genes in carrot (Daucus carota): isolation, sequence-characterization, single nucleotide polymorphism (SNP) markers and genome mapping. Theoretical and Applied Genetics 114, 693-704. Klein S, Ben-Shaul Y. 1967. Development of carotene bodies in the carrot root. Project A10-CR-14 Final Report submitted to USDA. pp49. Marx M, Stuparic M, Schieber A, Carle R. 2003. Effects of thermal processing on trans-cis-isomerization of /3-carotene in carrot juice and carotene-containing preparations. Food Chemistry 83, 609-617. Paolillo DJ Jr, Garvin DF, Parthasarathy MV. 2004. The chromoplasts of Or mutants of cauliflower [Brassica oleracea L. var. botrytis). Protoplasma 224, 245-253. 66 Rensing KH, Samuels, AL, Savidge RA. 2002. Ultrastructure of vascular cambial cell cytokinesis in pine seedlings preserved by cryofixation and substitution. Protoplasma 220, 39-49. Ruhlman T, Lee SB, Jansen RK, Hostetler JB, Tallon LJ, Town CD, Daniell H. 2006. Complete plastid genome sequence of Daucus carota: implications for biotechnology and phytogeny of angiosperms. BMC GenomicsZA, 222.1-222.13. Santos CAF, Simon PW. 2002. QTL analyses reveal clustered loci for accumulation of major provitamin A carotenes and lycopene in carrot roots. Molecular Genetics and Genomics 268, 122-129. Santos CAF, Senalik D, Simon PW. 2005. Path analysis suggests phytoene accumulation is the key strop limiting the carotenoid pathway in white carrot. Genetics and Molecular Biology 28, 287-293. Simon PW, Wolff XY, Peterson CE. 1985. Selection for high carotene content in carrots. HortScience 20, 586. Simon PW, Wolff XY. 1987. Carotenes in typical and dark.orange carrots. Journal of Agricultural Food Chemistry 35, 1017-1022. Simon PW, Wolff XY, Peterson CE,. Kammerlohr DS, Rubatzky VW, Strandberg JO, Bassett MJ, White JM. 1989. High Carotene Mass carrot population. HodScience 24, 174. Simon PW. 2000. 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Journal of the American College of Nutrition 15, 84-91. 68 CHAPTER 3 Agrobacterium-med\ate6 transformation of the algal crtO ketolase gene into carrot (Daucus carota L.) 3.1. Introduction Among 600 naturally occurring carotenoids, astaxanthin (3,3'-dihydroxy-4,4'-diketo-/3-carotene) is one of the ketocarotenoids that is beneficial to both human health and salmon aquaculture. Astaxanthin is a strong anti-oxidant and has immune functions in the visual and cardiovascular systems (Van den Berg et al., 2000). The ability to biosynthesize astaxanthin is limited to the yeast Xanthophyllomyces dendrorhdus (Andrewes et al., 1976), marine bacterium Agrobacterium aurantiacum (Misawa et al., 1995) and some green algae, particularly Haematococcus pluvialis (Boussiba and Vonshak, 1991; Johnson and Schroeder, 1995). In higher plants, only Adonis aestivalis was found to produce astaxanthin in its flowers (Mawson, 1994, US Patent 5,453,565; Cunningham and Gantt, 2005). The green alga H. pluvialis has the highest astaxanthin accumulation (4-5% dry weight; Yuan and Chen, 2000) and seems to be the most suitable natural source (Lorenz and Cysewski, 2000). Astaxanthin bio-accumulates in the food chain and provides the red pigmentation for the flesh of salmonid fishes. Salmonids preferentially deposit astaxanthin, followed by the related ketocarotenoid compound canthaxanthin in their flesh (Bowen et al., 2002). Since salmonids fish grown in aquaculture cannot acquire astaxanthin naturally, it must be added to the formulated diet (FDA approval granted in U S A in 1995). There is no efficient way to cultivate green algae to harvest astaxanthin (Bubrick, 1991; Lorenz and Cysewski , 2000). The source of feed astaxanthin is primarily through chemical synthesis (by Hoffman-La Roche, as Carophyll Pink, or B A S F Ltd., as Lucanthin Pink, at a cost of US$ 2,000-2,500/kg) and as a result, it constitutes an expensive component of salmon farming, accounting for almost 20-25% of total feed costs (Torrissen et al., 1995). 69 The annual worldwide market for astaxanthin is around US$ 200 million (Lorenz and Cysewski , 2000). There is therefore an incentive to produce astaxanthin in a more economical way. Astaxanthin is formed from ^caro tene by the introduction of keto and hydroxyl moieties at the 4,4' and 3,3' positions of the /?-ionone rings through two alternative pathways (Figure 3.1). The two critical enzymes that involved in this pathway are /^-carotene ketolase and /2-carotene hydroxylase. Except for the flowers of Adonis, there is no evidence that higher plants have genes encoding /f-carotene ketolase (Cunningham and Gantt, 2005). Lotan and Hirschberg (1995) first isolated the algal ketolase (crtO) gene from H. pluvialis and its heterologous expression in Escherichia coli and cyanobacteria has been tested (Breitenbach etal., 1996; Harker and Hirschberg, 1997; Kajiwara et al., 1997). These studies contributed to the characterization of crtO and its function in the ketocarotenoid pathway. In addition to the alga crtO, a cyanobacterjm crtO gene has been cloned from Synechocystis sp. P C C 6803 (Fernandez-Gonsalez et al., 1997), and a /2-carotene ketolase (crtW) gene was isolated from marine bacterium, Agrobacterium aurantiacum (Misawa et al., 1995) and Brevundimonas sp. SD212 (Choi et al., 2007). Mann et al. (2000) successfully transformed the H. pluvialis crtO to tobacco {Nicotiana tabacum) and opened up the possibility of engineering astaxanthin < biosynthetic pathway in the nectary. This research has recently triggered work on the metabolic genetic engineering of ketocarotenoid biosynthesis in plants. Synthesis of ketocarotenoids in transgenic plants has been demonstrated in the leaves and flowers of tomatoes and tobacco (Ralley et al., 2004), the tubers of potatoes (Gerjets and Sandmann, 2006; Morris et al., 2006), the flowers of Lotus japonicus (Suzuki et al., 2007) and the seeds of Arabidopsis(Stalberg etal., 2003). ' 70 Figure 3.1 Biochemical pathway of astaxanthin biosynthesis. /3-carotene ketolase (CrtO) and /3-carotene hydroxylase (CrtR-b) are important enzymes alternatively to produce ketocarotenoids (Mann etal., 2000). 71 In carrot roots, /^-carotene accumulates at high levels and is considered to be a good substrate for testing the ability of the crtO gene product to generate high levels of ketocarotenoids in planta. The main purpose of this study was to test the efficiency of ^orobacferium-mediated transformation of the H. pluvialis crtO gene in carrot and to see whether the constitutive promoters, C a M V 3 5 S and double C a M V 3 5 S are suitable for expression of the crtO gene at high enough levels in this plant to provide a potential source of astaxanthin or other ketocarotenoids for use salmon aquaculture and other purposes. Previous research reported that the untranslated leader sequence from alfalfa mosaic virus RNA4 combined with a duplicated-enhancer C a M V 3 5 S promoter (designated as double C a M V 3 5 S or C a M V 2x35S) can drive high levels of the transgene expression in tobacco and tomatoes (Kay et al., 1987; Datla et al., 1993; Krasnyanski et al., 2001). In transgenic tobacco plants, the steady-state expression of GUS under the control of a double C a M V 3 5 S promoter was elevated about 8-fold relative to levels using a native single C a M V 3 5 S promoter (Datla et al., 1993). The study reported here was carried out concurrently and in co-ordination with that of Dr. J . Jayaraj (Simon Fraser Univ. Canada). 3.2. Materials and Methods 3.2.1. Sequence analysis Two potential crtO cDNA clones (Ketol and Keto2) were cloned by R A C E from H. pluvialis by J . Maltby, Drs. D. Higgs and R. Devlin (Department of Fisheries and Oceans, Canada) and the sequences were analyzed by automated Prism Cycle Sequencing (ABI, Sunnyvale, CA) at the University of British Columbia Nucleic Acid-Protein Service Unit. A transit peptide sequence was not found using chloroplast transit peptide prediction software, Ch loroPv1.1 . (Emanuelsson, 1999; http://www.cbs.dtu.dk/services/ChloroP). 72 Hence, there was need to synthesize a Rubisco small subunit transit peptide sequence during binary vector construction (see below). 3.2.2. Binary vector construction In this study, two vector systems with constitutive promoters, C a M V 3 5 S and double CaMV35S were used to control crtO gene expression. A bar gene (encoding phosphinothricin acetyltransferase) cassette was used as a selectable marker and linked with the crfO gene cassette in tandem. The two cassettes were joined using an EcoRI enzyme site (Figure 3.2). The first binary vector construct with the C a M V 35S promoter upstream of the crtO gene, [CaMV35S-TP::crtO::Nos]-[CaMV35S-bar::Nos] was obtained from Dr. J . Jayaraj (Simon Fraser Univ. Canada). Since the crtO gene does not contain the plastid target sequences, the Rubisco small subunit transit peptide sequence (TP) was synthesized and fused with crtO gene. This transit peptide is needed to target gene expression in the plastid. Hindlll BamHI SphI PstI EcoRI BamHI Bglll EcoRI 1 I I 1 i 1 1 1 • C a M V 35 S T P Ketolase N o s CaMV35S j bar Nos | Figure 3.2 Single C a M V 35S-crtO binary vector construct (Jayaraj et al., 2005). CaMV 35S: promoter, crtO: /J-carotene ketolase gene, bar. phosphinothricin acetyltransferase gene, TP: transit peptide, Nos: nos terminator 73 The second vector system with a double CaMV35S promoter, [CaMV 2x35S-TP::crfO::Nos]-[CaMV35S-bar::Nos] was developed from the previous CaMV35S-cr fO vector system by me. The pSM-3 binary vector (a gift from Dr. S. Mansfield, UBC) , which is modified from pCambia1390, was used for this double CaMV35S-cr fO vector construct. The fragments of two gene cassettes, TP::crfO::Nos and CaMV35S-£>ar::Nos were excised by restriction enzymes, EcoRI and SamHI, respectively (Figure 3.3A-1 and 2). Since there were two BamHI sites in the CaMV35S-oar: :Nos gene cassette, it was first excised by EcoRI digestion (Figure 3.3A-1) and then TP::crfO::Nos was excised by SamHI digestion (Figure 3.3A-2). The TP::crfO::Nos fragment was ligated between SamHI and EcoRI sites downstream of the double C a M V 3 5 S promoter in pSM-3 (Figure 3.3A-3). Sequentially, pSM-3 was digested with EcoRI (Figure 3.3A-4) and then the CaMV35S-£>ar::Nos cassette was inserted (Figure 3.3A-5). Finally, the [CaMV2x35S-TP::crfO.\Nos]-[CaMV35S-£>ar::Nos] cassettes were added (Figure 3.3B). Using double C a M V 35S and crtO gene specific primer sets, the insertion of genes was confirmed by P C R . The primers used were: forward, ( 5 ' - T G A C G C A C A A T C C C A C T A T C C T T C G C A A G A C C C T - 3 ' ) and reverse ( 5 ' - G T T G T G G T G C T C C C A A T G C T T G C G G - 3 ' ) . To confirm the insertion of CaMV35S and bar gene cassette by P C R , forward ( 5 ' - C A G G A C T A A C T G C A T C A A GAACAC-3 ' ) , and reverse ( 5 ' - G A C T T C A G C A G G T G G G T G T A G A - 3 ' ) primers were used. The P C R cycling profile was 94°C 2 min, 94°C 45 sec, 58°C 30 sec, 72°C 1 min (25 cycles), 72°C 7 min, 10°C 5 min. The completed double 35S-crfO vector construct (pKB-1) was confirmed by DNA sequencing analysis in N A P S Unit (UBC) and the sequencing primers are listed in Table 3.1. 74 A. HIII BamHI 1 I © EcoRI I EcoRI © 1 C a M V 35 S TP crtO N o s C a M V 3 5 S bar N o s EcoRI EcoRI C a M V 3 5 S bar N o s ligation © BamHI 1 © TP EcoRI I crtO N o s ligation 1 © BamHI EcoRI C a M V 2 x 3 5 S . © BamHI C a M V 2 x 3 5 S EcoRI Cut-with E c o R I B. Hindi II BamHI 1 • EcoRI EcoRI C a M V 2 x 3 5 S TP cHO Nos C a M V 3 5 S bar Nos Figure 3.3 Schematic diagram of double CaMV35S-cr fO gene construct. A. Single CaMV35S-cr fO containing bar gene cassette, bar cassette and crtO cassettes were digested with restriction enzymes (E.coRI and BamHI), respectively. Two cassettes were ligated to pSM-3 vector. B. Reconstruction of CaMV2x35S-cr fO. crtO. /3-carotene ketolase, bar: phosphinothricin acetyltransferase, TP: Transit peptide sequences, Nos: nos terminator 75 Table 3.1 Primers for sequencing of the double 35S-crtO construct (pKB-1) Name Orientation Primer sequences 2X35S 1 F 2 5 ' - T G A C G C A C A A T C C C A C T A T C C T T C G C A A G A C C C T - 3 ' Keto_SQR1 R 3 5 ' - C C C A G G A G C C G A T G A C A G C T A G C G - 3 ' Keto_SQF2 F 5 ' -ACATCG C C G T A G T C T T C I I I G T C C T G G - 3 ' Keto_SQF3 F 5 ' - G C C G C C G C C T G T C T G G C C G A G G T C - 3 ' Bar_SQF1 F 5 - C A G G A C T A A C T G C A T C A A G A A C A C - 3 ' Bar_SQF2 F 5 ' -TTCTG G C A G C T G G A C T T C A G C C T - 3 ' 1 double C a M V 35S promoter forward sequences 2 F: forward, 3 R: reverse 3.2.3. Plant materials > The seeds of High Carotene Mass (HCM) carrot (Simon et al., 1985) were obtained from Dr. P. Simon (USDA/ARS, Univ. of Wisconsin, Madison, USA). Seeds were dipped in 70% ethanol for 1 min and soaked in a 20% solution of bleach (Javax 6.25% sol.) for 15 min, and then washed over 3 times in sterile distilled water. Seeds were grown in germination medium for one month and the petioles (1cm long) were excised and used for Agrobacterium mediated transformation. 3.2.4. Preparation of MS media Murashige and Skoog Basal Medium mix (Sigma, Canada) and liquid vitamin mix were used for all M S media. For germination of the seeds and regeneration of plants, 0.25% sucrose and 0.01% myo-inositol were added to half-strength MS medium (1/2MS, pH 5.8). 76 For callus inducing MS media, 3% sucrose, 0.01% myo-inositol and 1mg/L dichlorophenoxyacetic acid (2,4-D) were supplemented (MS1D, pH 5.8). After autoclaving and cooling of the MS medium, 1 and 10mg/L of B A S T A (phosphinothricin; Sigma-Aldrich, Canada) and 300mg/L of Timentin (SmithKline Beecham Co, UK) were added. One-tenth strength MS liquid medium (1/1OMS, pH 5.2) was used for Agrobacterium mediated transformation, to which 0.2% sucrose and 0.1% glucose were added. 3.2.5. Agrobacterium-mediated crtO gene transformation into HCM carrot The binary vectors carrying the crtO gene were transformed to Agrobacterium tumefaciens strain LBA4404 by electroporation. Agrobacterium strains carrying binary vectors were grown in LB liquid media (2mL) with streptomycin (100mg/mL) and kanamycin (50mg/mL) at 28-30°C for 2 days at rotary shaker at 200rpm. The culture was centrifuged for 5 minutes at 7,000 x g and the pellet was resuspended in 1/10 M S liquid media to an O D 6 0 0 of 0.3, and acetosyringone (200uM) was added to the resuspended culture, which was co-cultivated with one month-old petioles of H C M carrot for 10 minutes. The explants were blotted on filter paper (Whatman 3MM) briefly and transferred to 1D MS medium (MS with 2,4-D, 1mg/L). After incubation at room temperature for 3 days in the dark, the explants were transferred to 1D MS media with 1mg/L BASTA plus 300 mg/L Timentin. After 2-3 weeks, they were transferred to 1D MS with 10mg/L BASTA. To facilitate normal growth, the small calli were transferred to fresh 1D M S media every two weeks. 3.2.6. Regeneration of transformed plants (single CaMV 35S) and PCR confirmation To regenerate single CaMV35S-cr tO transformed plants, surviving calli on the 1 DMS with BASTA (10mg/L) and Timentin (300mg/L) were transferred to 2,4-D hormone and BASTA free 1/2MS solid media. 77 Embryos generated from callus were individually transferred to 1/2MS solid media in a bigger container to become individual putative single CaMV35S-cr tO transformed plant. Once transformed plants were obtained and genomic DNA was extracted from leaf samples using the modified method of Sambrook et al. (1998), or a Nucleon PhytoPure kit (Amersham Biosciences, Quebec, Canada). The crtO gene specific primers used for P C R amplification of crtO sequences from genomic DNA templates were: Keto2F, G C A A T G G T G G A A G A G T A A A G T G C , Keto2R, G T T G T G G T G C T C C C A A T G C T T G C G G . P C R cycling parameters were: 94°C 2 min, followed by 94°C for 45 s, 58°C for 30 s, 72°C for 40 s (30 cycles), and a final extension at 72°C for 7 min. In a parallel study, regeneration of double CaMV35S-cr fO transformed plants for confirmation was carried out by Dr J . Jayaraj (Jayaraj era/. , unpublished). 3.3. Results 3.3.1. Confirmation of crtO gene cDNA sequence Before carrying out transformation experiments with ^caro tene ketolase clones, it was first necessary to verify their sequences. DNA sequences of two crtO cDNA clones (designated as Ketol and Keto2) from H. pluvialis, obtained from Dr. R. Devlin were analyzed. Ketol had two C nucleotides missing at 587bp and 773bp that caused a frame shift. Keto 2 had no missing nucleotides. The amino acid sequence of Keto2 indicated a putative ketolase protein of 328 amino acids (Figure 3.4). The predicted crtO polypeptide shared overall identity of 94% with the ketolase sequence published by Harker and Hirschberg (1997; GenBank [CAA60478]) and 78% with the sequence published by Kajiwara et al. (1995; GenBank [Q39982]). The sequence analysis confirmed that the Keto2 clone encodes the /2-carotene ketolase (CrtO), and therefore it was used for crtO transformation experiments described in this chapter and in Jayaraj et al. (2005). 7 8 Keto2 MQLAATVMLEQLTGSAEAFKEKENVAGSSDVLRTWATQYSLPSEESDAARPGLKNAYKPPPSDTKG Keto2 I T M A L A V I G S W A A V F V H A I F Q I K L P T S L D Q L H W L P V S D A T A Q L V G G S S S L M H I A W F F V L E F L Y T G Keto2 LFITTHDAMHGTIAMRNRQLNDFLGRVCISLYAWFDYNMLHRKHWEHHNHTGEVGKDPDFHRGNPG Keto2 IVPWFVSFMSSYMSWQFARLAWWTVVMQLLGAPlylANLLVF^4AAAPILSAFRLFYFGTYMPHKPEP Keto2 SATSGSSSVVMNWWKSRTSKASDLVSFLTCYHFDLHWEHHRWPFAPWWELPNCRRLSGRGLVPA Figure 3.4 Amino acid sequences of the Keto2 clone. Keto2 (crtO gene encoding ketolase) was isolated from Haematoccous pluvialis. This was used for /4a/robacter/'um-mediated transformation in carrot. 3.3.2. Confirmation of the bar gene as a suitable selection marker.in HCM carrot I tested the use of the phosphomannose isomerase (pmi) gene as a selection marker in H C M carrots but found that it was not suitable (data not shown). In the test to see if the bar gene is a suitable marker, the calli from non-transformed explants showed 20% survival at 1mg/L B A S T A but were completely inhibited at 5mg/L BASTA. For explants transformed with the bar gene (derived from co-cultivation with an Agrobacterium suspension harboring the pCambia-tlp vector, obtained from Dr. Z. Punja, Simon Fraser Univ.) placed on 1DMS with 10mg/L BASTA solid medium, the survival rate was 92.4% (Figure 3.5). This is consistent with previous results in normal orange carrot, in which the bar gene had been previously tested and shown to be a suitable selective marker (Chen and Punja, 2002). Based on this test, I established the following B A S T A selection protocol for crtO transformation of carrots: Transformed explants were placed on 1DMS with 1mg/L B A S T A as a preliminary selection. After 2-3 weeks, surviving healthy calli were transferred to 1DMS with 10mg/L B A S T A solid medium as a final selection. Figure 3.6 shows that transformed calli derived from co-cultivation with an Agrobacterium strain harboring the crtO gene in the T-DNA vector were able to grow on 1DMS solid medium containing BASTA (10mg/L), while none of the non-transformed calli (as a negative control) survived. 7 9 Figure 3.5 Preliminary test of bar selection transformed and transformed explants. A . transformants. Selection was carried out (10mg/mL). (bar: 170mm) with H C M carrot. Comparison between non-Explants of non-transformants B. Callus of on 1D media supplemented with B A S T A Figure 3.6 Effect of phosphinothricin (BASTA) on [CaMV35S:crtO] transformed callus. Petioles of wild-type (left) and plants transformed with [CaMV35S:crfO] are shown. Non-transformants completely died and transformed explants produced healthy callus on MS medium with BASTA (10mg/l_). (bar: 170mm) Both single and double CaMVSSS-cr fO transformed calli placed under selection on 1DMS with BASTA (10mg/L) showed similar survival patterns. For single CaMV35S-cr tO transformation, 201 calli survived from 255 explants (78.8%). For double CaMV35S-cr fO transformation, 185 calli survived from 230 explants (80%). 3.3.3. PCR analysis of regenerated plant (single CaMV 35S-crtO) BASTA resistant calli (Figure 3.7A) putatively transformed with the single C a M V 3 5 S -crtO construct were used to regenerate carrot plants (Figure 3.7B, C and D). Genomic DNA was extracted from leaves of the regenerated single 35S-crtO plants (Figure 3.7D) and P C R analysis confirmed the insertion of the transgene using crtO gene specific primers. Out of 138 regenerated putative single CaMV35S-cr fO plants, 21 plants showed crtO gene P C R products (Figure 3.8). Thus, while transformation could only be confirmed in 15% of B A S T A seedlings, this was sufficient to generate adequate numbers of transgenic lines. 81 Figure 3.7 Regeneration of transgenic lines with CaMV35S-cr fO. A. callus B. embryo from callus C. embryo (bar: 5mm) D. regenerated plant (bar: 130mm) L 1 2 3 4 5 6 7 8 9 10 11 12 Figure 3.8 P C R screening of CaMV35S-cr fO transgenic lines. L. reference ladder, Lane 1: positive control, Lane 12: negative control (non-transformed plant), Lanes 2, 6, 8, 10, 11: regenerated plants showing crtO, Lanes 3, 4, 5, 7, 9: regenerated plants without crtO. 3.3.4. Phenotype of transgenic carrot callus derived from 4grobacfer/um-mediated transformation The calli derived from non-transformed carrot explants, grown on 1DMS media without selection, had a yellowish colour (Figure 3.9 D). However, the putatively transformed carrot calli in both the single CaMV35S-cr fO (Figure 3.9) and the double CaMV35S-c r fO transgenes (Figure 3.10) produced distinctively reddish pink calli on 1DMS media with selection. For single CaMV35S-cr fO transformation, 40 calli from 180 calli showed reddish pink colour (22%). For double CaMV35S-cr tO transformation, 45 calli from 191 calli showed reddish pink colour (23%). Jayaraj et al. (2005) also found reddish coloured calli in lines of crtO transformed carrots using the double C a M V 3 5 S promotor, in work carried out in parallel. H P L C analyses showed the presence of novel ketocarotenoids in these calli. Since Jayaraj et al. (2005) have demonstrated that reddish colour was a dependable indicator of crtO gene expression in carrots, it was deemed unnecessary for us to duplicate the H P L C analysis in our study. 83 Figure 3.9 Comparisons of color between sinlge CaMV35S-crfO transformed callus and non-transformed callus. A-C, single CaMV35S-c/tO transformed callus: putative crtO red and pink callus, D: non-transformed callus normal yellow callus (bar: 5mm) Figure 3.10 Comparisons of color between double CaMV35S-cr rO transformed callus and non-transformed callus. A-C: putative crtO red and pink callus, D: non-transformed normal yellow callus (bar: 5mm) 3.4. Discussion The red coloured ketocarotenoid astaxanthin (3,3'-dihydroxy-4,4'-diketo-/?,/3-carotene) is used widely as an additive in feed for the pigmentation of fish and crustaceans and is also frequently included in human nutritional supplements. Because of the high cost of chemical synthesis, there is considerable interest in developing a plant-based biological production process for astaxanthin. 85 Although higher plants synthesize carotenoids, they do not possess the ability to form ketocarotenoids (with the exception of Adonis spp.; Cunningham and Gantt, 2005). In order to modify higher plants to make them capable of synthesizing astaxanthin, one strategy is to express heterologous genes responsible for ketocarotenoid biosynthesis, such-as the H. pluvialis crtO gene encoding carotene 4,4' oxygenase (used in this study) in target transgenic plants, to generate the biochemical pathway shown in Figure 3.1. According to Figure 3.1, in some cases, it may also be necessary to introduce a gene encoding the 3,3' hydroxylase as well. In our study, we selected the HCM carrot as our target plant because of its high fi-carotene content in the root, which could supply a higher concentration of substrate for p-carotene ketolase and hydroxylase enzymes, potentially resulting in higher ketocarotenoid yields than in other plants or organs with lower /3-carotene levels. As a source of ketolase activity, we selected the previously characterized H. pluvialis crtO gene which has recently been shown to encode a functional /3-carotene ketolase enzyme by heterologous expression in a /3-carotene-producing E.coli strain, where it was capable of generating /3-carotene ketocarotenoid derivatives astaxanthin, canthaxanthin, and echinenone (Jayaraj et al., 2005). To generate a vector construct for crtO gene expression in carrot, the protein coding region was fused to an N-terminal transit peptide specifying protein import into plastids. The targeting of the crtO expression to the plastids was necessary since the carotenoid biosynthetic pathway takes place within plastids (Del Villa-Martinez etal., 2005), the sites of /3-carotene accumulation (Chapter 2). Similar targeting strategies and engineering of ketocarotenoid metabolism have been demonstrated in tobacco (Mann etal., 2000; Rally etal., 2004), tomatoes (Rally et al., 2004; Wurbs et al., 2007), potatoes (Gerjets and Sandmann, 2006; Morris et al., 2006) and Arabidopsis (Stalberg et al., 2003). 86 The Agrobacterium-med\a\ed transformation protocol using resistance to B A S T A conferred by the bar gene was tested. This protocol proved to be efficient in generating transformation into H C M carrot lines harbouring either the C a M V 3 5 S or the double C a M V 3 5 S promotor driving crtO expression. Feeneya and Punja (2003) used the phosphomannose isomerase (pmi) gene as a non-antibiotic selection marker in hemp (Cannabis sativa L.) transformation studies and found it to be effective. However, I found that it was not effective in carrots because of high survival rate in the non-transformed plants (data not shown). Therefore, the pmi gene was not a suitable selection marker for carrot transformation. I was able to regenerate 138 carrot lines after transformation with a single CaMV35S-crtO construct and showed that the transgene was present in 21 lines with P C R analysis. Jayaraj et al. (2005) examined 98 regenerated carrot lines after transformation with a double CaMV35S-cr fO construct and found 45 lines with ketolase expression. Wally et al. (2005) compared the efficiency of several promotors fused to the Gus reporter gene for carrot transformation and found that the double C a M V 3 5 S promotor shows higher expression efficiency than the C a M V 3 5 S promotor. Therefore, while the single CaMV35S-cr tO transgenic calli I generated appeared to produce the desired ketocarotenoid product, based on their reddish pink color, it was decided that the double C a M V 3 5 S promotor would be used for further transformation studies by our collaborators (Jayaraj et al., in prep) and I did not pursue further work on regeneration of double CaMV35S-crfO lines or their analysis. The visual phenotypes of some calli transformed with both single CaMV35S-crfO and double CaMV35S-crfO constructs, in which red pigments consistent with ketocarotenoid accumulation were observed (Figures 3.9 and 3.10), suggest that the ketocarotenoid accumulation was achieved in these putatively transgenic cells. While l did not pursue further molecular and biochemical analyses of these calli or transgenic lines derived from them, similar phenotypes were observed in calli derived from transformation with the double 87 CaMV35S-cr tO construct (Jayaraj et al., 2005; J . Jayaraj and Z. Punja, personal communication). H P L C analysis of the roots and leaves of 5 transformed carrot lines derived from these calli (Jayaraj era/ . , in prep) demonstrated the presence of novel ketocarotenoids: Astaxanthin (the major ketocarotenoid), adonixanthin, adonirubin, canthaxanthin, and echinenone. Stalberg et al. (2003) expressed the crtO gene in seeds of Arabidopsis using a seed storage protein promotor napA. The major ketocarotenoids in their transgenic plants were: 4-keto-lutein, adonirubin and canthaxanthin. 4-Keto-lutein differs from adonixanthin only in the position of one double bond. In summary, I carried out the first important steps in genetically modifying H C M carrots to make them capable of synthesizing astaxanthin and canthaxanthin. This study laid down the ground work that enabled subsequent studies (Jayaraj et al., in prep) to demonstrate the feasibility of introducing a new ketocarotenoid pathway into transgenic carrots. 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Cold Spring Harbor, New York: Cold Spring Harbor Laboratory Press. Stalberg K, Lindgren O, Ek B, Hoglund A-S. 2003. Synthesis of ketocarotenoids in the seed of Arabidopsis thaliana. The Plant Journal 36, 771-779. Suzuki S, Nishihara M, Nakatsuka T, Misawa N, Ogiwara I, Yamamura S. 2007: Flower color alteration in Lotus japonicus by modification of the carotenoid biosynthetic pathway. Plant Cell Report 26, 951 -959. Torrissen OJ, Christiansen R, STRUKSN/ES G and Estermann R. 1995. Astaxanthin deposition in the flesh of Atlantic Salmon, Salmo salar L., in relation to dietary astaxanthin concentration and feeding period. Aquaculture Nutrition 1, 77-84. Yuan JP, Chen F. 2000. Purification of trans-astaxanthin from a high yielding astaxanthin ester-producing strain of the microalga Haematoccous pluviallis. Food Chemistry 68, 443-448. 91 Van den Berg H, Faulks R, Fernando Granado H, Hirschberg J, Olmedilla B, Sandmann G, Southon S, Stahl W. 2000. The potential for the improvement of carotenoid levels in foods and the likely systemic effects. Journal of the Science of Food and Agriculture 80, 880-912. Wally O, Jayaraj J, Punja ZK. 2005. Comparative analysis of tissue and wound induced transgene expression levels in carrot (Daucus carota L.) using 7 different promoter constructs. Presented at the American Society for Plant Biology meeting in Seattle, WA, July, 2005. Wurbs D, Stephanie R, Ralph, B. 2007. Contained metabolic engineering in tomatoes by expression of carotenoid biosynthesis genes from the plastid genome. The Plant Journal 49, 276-288. 92 CHAPTER 4 Co-expression of Arabidopsis thaliana cytochrome P450 enzymes and NADPH-cytochrome P450 reductase in Escherichia coli: Testing the functions of candidate /J-carotene hydroxylases2 4.1. Introduction In plants, cytochrome P450 (CYP) monooxygenases play important roles in the biosynthesis of natural products (e.g. fatty acids, phenylpropanoids, alkaloids, terpenoids, and carotenoids), but of the at least 272 P450 genes identified in Arabidopsis, most are of unknown biochemical function. In green plants, zeaxanthin (3R,3'R-/3,/3-carotene-3,3'-diol), is an important carotenoid synthesized transiently under conditions of excess light energy, where it participates in photoprotection. It may also function as a starter molecule for the biosynthesis of other carotenoid derivatives in plants. A crucial enzymatic step required for the.formation of zeaxanthin and its precursor/J-cryptoxanthin is /3-ring hydroxylation of the precursor, /3-carotene, carried out by the action of /3-carotene hydroxylase. An Arabidopsis /3-carotene hydroxylase gene was identified by Sun et al. (1996), but the C Y P 9 7 family in Arabidopsis is suspected of encoding additional enzymes with /3-carotene hydroxylase activity (Tian etal., 2004). However, not all of these genes have been experimentally demonstrated to have this role, and there is no data on the enzymatic activity of any of these P450s. In this study, we have investigated the roles of three Arabidopsis family C Y P 9 7 genes in P450 dependant hydroxylation of /3-carotene by expressing recombinant enzymes in a /3-carotene producing E.coli strain and measuring • recombinant protein accumulation, and product accumulation by H P L C . 2 A version of this chapter has been published. Kim JE, Punja ZK, Douglas CJ. 2005. In P450 Systems and Regulation, Proceedings of 14 th International Conference on Cytochromes P450, Dallas, TX, 2005: Medimond S.r.l, Italy. pp115-120. 93 4.2. Materials and Methods 4.2.1. Plasmid construction and bacterial growth conditions Full length cDNAs of three cytochrome p450 genes (97A3, 97B3 and 97C1) were isolated from Arabidopsis by polymerase chain reaction (PCR) using gene-specific P C R primers (Appendix 2) and cDNA clone of Arabidopsis NADPH-cytochrome P450 reductase (ATR1) was obtained from Dr. R. Peters (Ohio State Univ. USA). CYP: :ATR1 co-expression constructions were made in the bacterial expression vector pETDUET-1 (Novagen) (pETDUET-97A3, 97B3, 97C1). P C R was applied to introduce restriction enzyme sites flanking the ATR1 cDNA for fusion with N-terminal 6His-tag in pETDUET. Sequentially, three P450 genes were inserted to generate tandem C-terminal S-tag fusion proteins in the same vector. Constructs were transformed into /3-carotene producing E.coli C43 (DE3) (Miroux and Walker, 1996) which,harbors pACCARI6Acr tX (Misawa e ta / . , 1990). After induction of gene expression with 0.1 mM IPTG, cultures were grown at 30°C for 48 hours in LB media supplemented with chloramphenicol (30mg/L) and ampicillin (100mg/L). Cloning and transformation strategies were shown in Appendix 2. 4.2.2. Extraction of carotenoid and HPLC analysis For extraction of the carotenoid pigments from bacterial cultures* overnight cultures were harvested and the pellet fractions were resuspended with [chloroform:methanol:water (20:20:8, v/v/v)]. The yellowish lower phase fraction was dried out and dissolved in the mobile phase for H P L C analysis (53% methyl tertiary-butyl ether, 43% methanol, 4% water (v/v/v)). A C30 reversed-phase column (Waters, 4.6 mm x 250 mm) was used for H P L C analysis with a flow rate of 1.0 mL/min. Absorption at 450 nm was measured and retention times of peaks compared to those of authentic standards (CaroteNature). 94 4.2.3. SDS-PAGE and Western blot Crude E.coli extracts were separated on 10% (w/v) polyacrylamide gels and transferred to P V D F membrane (Amersham) for western blot analysis. Immunodetection of fusion proteins was carried out with the E C L detection system (Amersham). An anti-His monoclonal antibody and an anti-mouse peroxidase-conjugated IgG (Amersham) were used as primary and secondary antibodies for immunodetection of His tag::ATR1. For detection of C Y P : : S tag fusion proteins, an S-protein H R P conjugated antibody (Novagen) was employed. The P D V F membrane was developed using the E C L Plus buffer system according to the manufacturer's recommendations, and analyzed with a Storm Image 860 phosphorimaging system (Amersham). 4.3. Results I generated E.coli C43 strains, each co-expressing one of three candidate C Y P 9 7 family ^caro tene hydroxylase genes and the cytochrome p450 reductase A T R 1 . In order to facilitate detection of the expressed proteins in E.coli extracts, ATR1 was expressed as an N-terminal His-tag fusion, and C Y P 9 7 proteins as C-terminal S-tag fusions. After induction of strains with IPTG, protein extracts were prepared, fractionated by S D S - P A G E , and western blots developed with anti-His tag and anti S-tag antibodies. All strains showed accumulation of ATR1 , at roughly equal levels (Figure 4,1). Western blot analysis also confirmed accumulation of CYP97A3 , C Y P 9 7 B 3 and CYP97C1 proteins, with predicted sizes of just under 70kD (Figure 4.1). While C Y P 9 7 B 3 was well expressed, as measured by S tag-CYP97B3 accumulation, the expression levels of C Y P 9 7 A 3 and CYP97C1 were several-fold lower, for unknown reasons. i 95 C Y P 9 7 A 3 CYP97B3 C Y P 9 7 C 1 E A T R 1 Figure 4.1 Western blot analysis of CYP97A3 , C Y P 9 7 B 3 , and CYP97C1 accumulation in E.coli strain C43. Crude E.coli extracts were separated by S D S - P A G E , and an anti-S tag antibody was used to detect fusion proteins. ATR1 was expressed in C43 and detected by His-tag antibody. The migration of a 70kD protein standard is shown. E: empty vector (negative control) I next prepared carotenoid extracts from each of the strains, and fractionated extracts by H P L C . As a control, I generated identical extracts from a strain expressing the non-P450 p-carotene hydroxylase described by Sun et al. (1996) in E.coli strain C43 . A s expected from previous results (Sun et al., 1996), the latter strain was able to produce products that co-migrate with both zeaxanthin and its singly hydroxylated precursor ,3-cryptoxanthin, as judged by co-migration with authentic standards (Figure 4.2; standards not shown). In contrast, a C43 strain harbouring the empty vector produced /2-carotene only (Figure 4.2). The dual CYP97B3-ATR1 expressing C43 strain also produced products co-migrating with zeaxanthin and p-cryptoxanthin, although at levels lower than those reduced by the known hydroxylase gene. UV spectra for the peaks were obtained, and that they were consistent with the proposed chemical nature of the compounds (/f-cryptoxanthin and zeaxanthin). Neither of the strains co-expressing C Y P 9 7 A 3 and CYP97C1 with ATR1 showed any detectable accumulation of zeaxanthin or /J-cryptoxanthin (data not shown). 96 Empty vector AtB1 C Y P 9 7 B 3 Figure 4.2 H P L C analysis of carotenoid extracts. Extracts of strain C43 transformed with the constructs indicated below were subjected to H P L C analysis as described in Materials and Methods. Carotenoids were detected at 450nm. C Y P 9 7 B 3 was co-expressed with A T R 1 . AtB1: /3-ring hydroxylase (Sun et al., 1996). b-car: /3-carotene, zea: zeaxanthin, b-crypt: /3-cryptoxanthin 4.4 . Discussion This study demonstrated that the Arabidopsis C Y P 9 7 B 3 gene, of previously unknown function, encodes an enzyme with apparent /3-carotene hydroxylase function, making it a new /3-carotene hydroxylase gene. Several researchers have suspected that, in addition to the previously described non-heme di-iron monooxygenase /3-carotene hydroxylase enzyme (Sun et al., 1996), cytochrome P450 enzyme activity might be involved in hydroxylation of carotenoids (Cunningham and Gantt, 1998; Schoefs etal., 2001; Inoue, 2004). Furthermore, 97 Balsco et al. (2004) cloned CYPA75A1 from Thermus thermophilus HB27 and found that it has /3-carotene hydroxylase function. My study provides biochemical evidence that plant C Y P enzymes do indeed have the ability to hydroxylate /3-carotene, and may play important roles in carotenoid metabolism. Based on phylogenetic reconstructions (Tian etal., 2004), C Y P 9 7 B 3 is most closely related to C Y P 9 7 A 3 and C Y P 9 7 C 1 . These authors used genetic analyses to show that CYP97C1 has a function in ^-carotene hydroxylation (Tian etal., 2004). However, no functional test of GYP97C1 enzyme activity was reported. While my data strongly suggest that C Y P 9 7 B 3 , in a C-terminal S-tag version, has /3-carotene hydroxylase activity, it was surprising that neither C Y P 9 7 A 3 nor CYP97C1 converted /3-carotene to zeaxanthin via /3-cryptoxanthin in my E.coli system. However, due to relatively low expression of these proteins in my E.coli strains, the data on C Y P 9 7 A 3 and CYP97C1 remain inconclusive at this point. As shown in Figure 4.1, expression levels of theses two proteins were much lower in the C43 strain than C Y P 9 7 B 3 , making it possible that enzyme levels were too low to produce detectable hydroxylated products. I am currently optimizing expression of all three C Y P 9 7 gene family members in E.coli. Also surprising were the relatively low levels of /3-cryptoxanthin and zeaxanthin produced in the CYP97B3-expressing strain, in comparison to the known non-CYP /3-carotene hydroxylase expressed in the same C43 background. Although the presence of a C-terminal His-tag has been shown not to significantly affect the activity of the Populus C Y P 7 3 (cinnamate-4-hydroxylase) enzyme in yeast cells (Ro et al., 2001), the presence of C-terminal S-tags on C Y P 9 7 B 3 and the other C Y P 9 7 fusion proteins could interfere with their activity in the E.coli system. Thus, it may be desirable to express them as native proteins. Further work oh the expression of C Y P 9 7 family proteins in E.coli should clarify their roles in plant carotenoid metabolism. 98 4.5. References Blasco F, Kauffmann I, Schmid RD. 2004. CYP175A1 from Thermus thermophilus HB27, the first beta-carotene hydroxylase of the P450 superfamily. Applied Microbiology and Biotechnology 64, 671-674. Cunningham Jr. FX, Gantt E. 1998. Genes and enzymes of carotenoid biosynthesis in plants. Annual Review of Plant Physiology and Plant Molecular Biology 49, 557-583. Inoue K. 2004. Carotenoid hydroxylation--P450 finally! Trends in Plant Science 9, 515-517. Miroux B, Walker JE. 1996. Over-production of proteins in Escherichia coli: mutant hosts that allow synthesis of some membrane proteins and globular proteins at high levels. Journal of Molecular Biology 260, 289-298. Misawa N, Nakagawa M, Kobayashi K, Yamano S, Izawa Y, Nakamura K, Harashima K. 1990. Elucidation of the Erwinia uredovora carotenoid biosynthetic pathway by functional analysis of gene products expressed in Escherichia coli. Journal of Bacteriology 172, 6704-6712. Ro DK, Mah N, Ellis BE, Douglas CJ. 2001. Functional characterization and subcellular localization of poplar (Populus trichocarpa X Populus deltoides) cinnamate 4-Hydroxylase. Plant Physiology 126, 317-329. Schoefs B, Rmiki N-E, Rachadi J, Lemoine Y. 2001. Astaxanthin accumulation in Hematococcus requires a-cytochrome P450 hydroxylase and an active synthesis of fatty acids. FEBS Letters 500, 125-128. Sun ZR, Gantt E, Cunningham Jr FX. 1996. Cloning and functional analysis of the /3-carotene hydroxylase of Arabidopsis thaliana. Journal of Biological Chemistry 271, 24349-24352. Tian L, Musetti V, Kim J, Magallanes-Lundback M, DellaPenna D. 2004. The Arabidopsis LUT1 locus encodes a member of the cytochrome P450 family that is required for carotenoid f-ring hydroxylation activity, Proceedings of the National Academy of Sciences USA 101, 402-407. 99 CHAPTER 5 Functional Characterization of CYP450 Carotenoid Hydroxylases and /3-carotene Ketolase in Arabidopsis thaliana3 5.1. Introduction Xanthophylls are oxygenated derivatives of carotenes that perform critical roles in the photosynthetic apparatus of higher plants (Niyogi, 1999). Lutein (3R,3'R-/3,£-carotene-3,3'-diol) is the most abundant xanthophyll in all plant photosynthetic tissues, where it plays an important role in light harvesting complexes (Tian et al., 2004). Zeaxanthin (3R,3'R- /3,/3-carotene-'3,3'-diol) is a structural isomer of lutein and a component of the non-photochemical quenching mechanism (Cunningham and Gantt, 1998). Both lutein and zeaxanthin are dihydroxy xanthophylls. Lutein is derived by the addition of hydroxyl groups to the 3, 3' positions of the e- and /3-rings of «?-carotene (/?,£-ring carotene) with the help of e- and /3-ring hydroxylases. Zeaxanthin is generated by the addition of hydroxyl groups to the 3, 3' position of both /3-rings of /3-carotene (fi, /3-ring carotene) with the help of /3-hydroxylases. (DellaPenna, 2001) (Figure 5.1). In addition to Arabidopsis (Sun et al., 1996; Tian and DellaPenna, 2001), /3-ring hydroxylases from photosynthetic and nonphotosynthetic bacteria, green algae and other plants have been cloned and functionally characterized by heterologous expression (Misawa etal., 1990; Hundle etal., 1994; Misawa era/ . , 1995; Bouvier etal., 1998; Cunningham and Gantt, 1998; Masamoto etal., 1998; Linden, 1999; Galpaz etal., 2006). In plants, cytochrome P450 (CYP) monooxygenases play important roles in biosynthesis of secondary metabolites, but most are of unknown biochemical function. 3 A version of this chapter will be submitted for publication. Kim J E , Cheng KM, Craft N, Hamberger B, Douglas C J . 100 Two cytochrome P450 type monooxygenases with activity against carotenoids have been identified from the phenotypes of knockout mutants of Arabidopsis. CYP97C1 (LUT1 locus) has £-ring hydroxylase activity (Tian et al., 2004), and C Y P 9 7 A 3 (LUT5 locus) encodes a fi-ring hydroxylase with major activity towards the /3-ring of ocarotene and minor activity on the /brings of /?-carotene (Kim and DellaPenna, 2006). Lycopene / \ CYP97C1 / \ CYP97A3 AtB1 / \ (CYP97B3) * o-Carotene ^•Carotene o-Cryptoxanthin CYP97A3\ zeinoxanthin AtB1 /5-Cryptoxanthin HO Lutein I HO' I I Zeaxanthin Antheraxanthin, violaxanthin, Neoxanthin Figure 5.1 Representation of the xanthophyll pathways showing hydroxylated products derived from a- and /2-carotene. The proposed sites of action of /3-ring hydroxylase (AtB1) and P450 hydroxylases (CYP97A3, C Y P 9 7 B 3 , CYP97C1) in a-, fi-fmg modification, based on the results of this study, are shown (Kim and DellaPenna, 2006) 101 Recently, two cytochrome P450-type carotenoid hydroxylases (CYP97A4 and CYP97C2) have been isolated from Oryza sativa and characterized in a /3-carotene producing E.coli strain (Quinlan et al., 2007). C Y P 9 7 A 4 and C Y P 9 7 C 2 , which are grouped in the same clan as C Y P 9 7 A 3 and C Y P 9 7 C 1 , were shown to have e-ring hydroxylase and /3-ring hydroxylase activities, respectively. Furthermore, CYP175A1 , cloned from the thermostable bacterium Thermus thermophilus HB27, shows /3-ring hydroxylase activity in E.coli. C Y P 1 7 5 A 1 , hydroxylating both /3- rings of /3-carotene to form zeaxanthin (Balsco et al., 2004). Therefore, there is clear evidence that P450 type monooxygenases, in addition to non-heme hydroxylases such as AtB1, are involved in carotenoid hydroxylase pathways! As mentioned above, four carotenoid hydroxylases have been identified in Arabidopsis. However, previous genetic studies suggest that additional unknown carotenoid hydroxylase(s) could be involved in the biosynthesis of the /3-,/3-ring xanthophylls or fi-,e-r\r\g xanthophylls, since residual accumulation of hydroxylated carotenoids was observed in mutants lacking activity of these known hydroxylases (Tian et al., 2004; Fiore era/. , 2006; Kim and DellaPenna, 2006). Based on phylogenic relationship, C Y P 9 7 B 3 shares 42% amino acid identity with CYP97C1 (Tian et al., 2004), and so I hypothesized that C Y P 9 7 B 3 could have /3-carotene hydroxylase activity (Kim et al., 2005; see also Chapter 4). In that study, a role for C Y P 9 7 B 3 in hydroxylation of /3-carotene was demonstrated by expressing the recombinant enzyme in a /3-carotene producing E.coli strain. This work showed that C Y P 9 7 B 3 could generate /3-cryptoxanthin and zeaxanthin from /3-carotene. Interestingly, predicted subcelluar locations of C Y P 9 7 A 3 and CYP97C1 are in the chloroplasts and these are among the nine cytochrome P450s in Arabidopsis predicted to be chloroplast-targeted. C Y P 9 7 B 3 is predicted to be targeted to the mitochondria (Schuler and Werch-Reichhart, 2003; Tian et al., 2004). These locations have not been experimentally verified. In my current study, I have further investigated the hypothesis that C Y P 9 7 B 3 and other P450 type carotenoid hydroxylases play 102 different roles in the hydroxylation of the /3-ring of ocarotene and /3-carotene in plants by determining the metabolic consequences of overexpressing the corresponding genes in transgenic Arabidopsis. To test this hypothesis, I generated Arabidopsis overexpression lines transformed using a binary vector harboring AtB1, CYP97A3, CYP97B3, CYP97C1 transgenes under the control of a double C a M V 3 5 S promoter. With the additional goal of further exploring alternative ways of engineering the production of novel ketocarotenoids in higher plants (see Chapter 3), I also generated a double CaMV35S-ketolase construct. I used a co-transformation approach to generate transgenic Arabidopsis lines expressing ketolase alone, and double transgenic lines expressing ketolase and each of the candidate carotenoid hydroxylases. The objectives of the this study were to (1) obtain evidence about P450 hydroxylase activities by examining the metabolic consequences of their overexpression as single transgenes in plants, (2) examine co-transformed lines for production of ketocarotenoids, and (3) test overexpression lines for evidence of cross-talk between carotenoid metabolic pathways. 5.2. Materials and Methods 5.2.1. Binary vector construction Hydroxylases: The coding regions of /3-carotene hydroxylase (AtB1) and C Y P 9 7 family proteins (97A3, 97B3 and 97C1) were isolated from Arabidopsis cDNA using gene specific P C R primers (Table 5.1 and Appendix 2). The genes were placed under the control of the double C a M V 35S promoter in the pSM-3 vector containing a hygromycin resistance (hygR) gene as a plant selectable marker (See Chapter 3; a gift from Dr. S. Mansfield, U B C ) by P C R amplification of cDNAs using gene-specific primers containing restriction enzyme 103 recognition sites (Table 5.2). The P C R products were digested with each enzyme and ligated into the corresponding site of pSM-3. The orientation of genes was confirmed by P C R and DNA sequencing (Table 5.3). These fusion constructs were named as pAtB1 (double CaMV35S-AtB1) , p97A3 (double CaMV35S-97A3) , p97B3 (double CaMV35S-97B3) and p97C1 (double CaMV35S-97C1) . Table 5.1 Sources of genes used in this study Arabidopsis Gene Origin gene GenBank Reference designation accession no. AtB1 Arabidopsis thaliana At4g25700 U58919 Sun etal. (1996) CYP97A3 Arabidopsis thaliana At1g31800 . AY142017 Kim & Dellapenna (2006) CYP97B3 Arabidopsis thaliana At4g15110 BT002582 Kim et al (2005) CYP97C1 Arabidopsis thaliana At3g53130 AY091083 Tian et al (2004) crtO Heamatococcus X86782 Harker & Hirschberg (1997) pluvials - . This study 1 1 A gift from Dr. R. Devlin, Department of Fisheries and Ocean, Canada 104 Table 5.2 Primers for gene cloning for the fusion binary vector constructs Primer Construct Primer sequence AtB1 (Xbal) F 1 pAtB1 5 ' - C A G G C T T C T A G A A T C A T G G C G G C A G G A C T C T - 3 ' 3 AtB1 (Kpnl) R 2 pAtB1 5 ' - G A G C C A G G T A C C T C A A G A A C T C G A A C T C G A C C - 3 ' 97A3 (Xmal) F p97A3 5 ' - T C C G T C C T C C C G G G T A T G G C T A T G G C C I I ICCT-3 ' 97A3 (Xmal) R p97A3 5 ' - A C G T A G G G T C C C G G G T T A A G A A A G A G C A G ATG-3 ' 97B3 (BamHI) F p97B3 5 ' - T G C C C C A G G ATCCG C A A T G G T A G C A G C C ATG G -3' 97B3 (Kpnl) R p97B3 5'- G C C C G T C G G T A C C T C A C I I I G A T C T T C T C T T T A G T T - 3 ' < 97C1 (Xbal) F p97C1 5'- C A C G T C C G C T C T A G A A T G G A G T C T T C A C T C T T T - 3 ' 97C1 (BamHI) R p97C1 5 ' - G T G T A C C G G A T C C T T A C C I I I G G C T C A C C T T C - 3 ' 1 F: forward, 2 R: reverse 3 Enzyme recognition site underlined and in bold Table 5.3 Primers for PGR confirmation of transgene presence in transformed Arabidopsis lines Primer name Orientation Primer sequence 2X 35S 1 F 2 5 ' - T G A C G C A C A A T C C C A C T A T C C T T C G C A A G A C C C T - 3 ' AtB1_R2 R 3 5 ' - T C T C A C C T C C C T C C A I I I G C C A A - 3 ' 97A3_R2 R 5 ' - C A A T G C A G G A A C A A T G G C A C G C C - 3 ' 97B3_R2 R 5 ' - T C C C A T A A T C G G C T C T A A G A T C T C A - 3 ' 97C1_R2 R 5 ' - C A A T C A C A G A C A A A T A C C T C C T G - 3 ' 1 double C a M V 35S promoter 2 F: forward, 3 R: reverse 105 Ketolase: The double CaMV35S-ketolase construct (pKB-1) harboring the crtO ketolase and bar selectable marker genes (Chapter 3) was modified for this study. To facilitate selection of Arabidopsis co-transformants with each P450 hydroxylase transgene, the hygromycin resistance gene (hygR), in pKB-1 was eliminated by Xho\ digestion and re-ligation. P C R with gene specific primers (forward: 5 ' - A T G A A A A A G C C T G A A C T C A C C G C G - 3 ' and reverse: 5 ' - T C T A C A C A G C C A T C G G T C C A G A C - 3 ' ) confirmed excision of the gene. This double CaMV35S-ketolase construct without hyg R was called pKB-2. Agrobacterium tumefaciens strain GV3101 was transformed with each fusion constructs (pAtB1, p97A3, p97B3, p97C1 and pKB-2) by heat shock. Transformants were selected on Luria-Bertani media containing rifampicin (25mg/L), gentamycin (25mg/L) and kanamycin (50mg/L), and the presence of the fusion constructs were verified by P C R . 5.2.2. Plant transformation and selection Agrobacterium strain GV3101 carrying the fusion constructs was grown at 28-30°C and transformed into Arabidopsis thaliana (Columbia ecotype: WT), using the floral dip method (Clough and Bent, 1998) with 0.05% (v/v) Silwet L-77 (Lelhle Seeds, Round Rock, TX) and 5% sucrose. Agrobacterium strains with AtB1 and P450 hydroxylase constructs were named GV3101/pAtB1, GV3101/p97A3, GV3101/p97B3, and GV3101/p97C1, respectively and the Agrobacterium strain with the ketolase (crtO) construct was named GV3101/pKB-2. For co-transformation, GV3101/pKB-2 was mixed with equal volume of the appropriate GV3101/hydroxylase strain. Dipped plants were covered with black plastic wrap (for 24 h) to maintain high humidity and were kept in the potting room for drying after seed set. Seeds were harvested and sterilized with 70% and 95% ethanol. Single transformed plants were designated as AtB1, 97A3, 97B3, 97C1 and KB, respectively. The seedlings (T1) of primary hydroxylase 106 transformed plants (T0) were selected on Murashige-Skoog (MS) media containing hygromycin (50mg/L; Invitrogen, Ontario, Canada). KB seedlings (T1) were selected either on MS media with BASTA (glufosinate-ammonium, 50mg/L; Sigma-Aldrich, Canada) or by spraying with 0.1% (v/v) BASTA (glufosinate Final EV150, AgrEvo EH, Paris, France). For selection of co-transformed plants (designated as A tBKB, 97AKB, 97BKB and 97CKB), seedlings (T1) of primary co-transformed plants (T0) were first selected on M S media containing hygromycin (50mg/ml_), transferred to soil mix (Sunshine Mix 5; SunGro, Seba Beach, Alberta, Canada) and grown to maturity. Instead of using B A S T A selection, T1 cotransformed lines harboring both ketolase and hydroxylase T-DNA insertions were identified by P C R analysis of genomic DNA. The seeds of T1 plants were harvested and selected on M S media supplemented with hygromycin. The surviving seedlings (T2) were transferred to soil and T2 double transformant lines sprayed with BASTA every 7 days after potting. Plants were grown under a 12-h photoperiod (100-120 umol/m' 2 S" 1 ; 22°C) and 18°C at night. To identify homozygous lines, the segregation pattern of hygromycin and B A S T A resistances of T2 lines was determined by growing T3 seedlings on MS media (MS, MS with 50mg/L hygromycin, and/or MS with 50mg/L BASTA) . T3 homozygous plants were grown on MS media without selection and leaves were harvested for H P L C analysis. 5.2.3. Genomic DNA Plant genomic DNA was extracted from 4 week old leaves (0.1g) of wild type Arabidopsis Col-0 plants and transgenic T1 lines with a Nucleon PhytoPure kit (Amersham Biosciences). P C R conditions were 94°C for 2 min, 94°C for 45 sec, 57°C for 30 sec, 72°C 45 sec (30 cycles), 72°C 7 min, 10°C 5 min. Gene specific primers as shown in Table 5.3 were used. 107 5.2.4. RNA isolation Four to five week old leaves from each T2 line were used for transgene transcript analysis to select lines with high transcript levels. Total RNA was extracted from leaf samples (0.1g) using the RNeasy Plant Mini Kit (Qiagen, Canada) and was treated with RNase-free DNase kit (Qiagen) to ensure the complete removal of DNA. Total RNA was quantified in a UV-visible spectrophotometer (Biospec-1601, Shimadzu) and the quality evaluated on an agarose gel (1%). 5.2.5. cDNA synthesis and semi-quantitative RT-PCR analysis The first-strand cDNA synthesis was performed using 2 pg of total RNA/20 pi reaction with the Superscript II Reverse Transcriptase (Invitrogen). To normalize for variation in cDNA concentrations, actin transcripts were P C R amplified in parallel from each sample (actin primers are shown in Table 5.4). The P C R condition was: 94°C for 3 min, 94°C for 30 s, 63°C for 30 s, and 72°C for 1min, followed by a final extension at 72°C for 3 min (30 cycles). After normalizing the amount of cDNA of each gene with actin, the transcript level of each sample was compared. The gene specific P C R primers are shown in Table 5.4. The P C R conditions were:' 94°C for 2 min, 94°C for 45 s, 56-58°C for 30 s, and 72°C for 40s, followed by a final extension at 72°C for 3 min (30 cycles). P C R products were separated by electrophoresis on a 1% agarose gel and visualized under ultraviolet light after staining with ethidium bromide. The P C R band intensities were measured using Alpha Imager software. 108 Table 5.4 Primers for R T - P C R measurement of transgene expression Primer name Orientation Primer sequence AtB1_RT F 1 5 ' - G A G A A C G A T G A G A G A C C G G A - 3 ' AtB1_RT R 2 5 ' - T C G A A C T C G A C C C G G A G C C C G A -3' 97A3_RT F. 5 ' - C G T A C T C T C G G A G A T C G A A C - 3 ' 97A3_RT R 5 ' -TATC A A G T G T C A A A C G A G A G - 3' 97B3_RT F 5 ' - G T T G T C A T C T C A G A T C C C A T T A T - 3 ' 97B3_RT R 5 ' - G C C G G T C A T C A A T G T C A A C A C - 3 ' 97C1_RT F 5 ' - T G G A G T C T T C A C T C I I I I C T C C - 3 ' 97G1_RT R 5 ' -TACC I I I G G C T C A C C T T C A T A T A C - 3 ' Actin F 5 ' - G C G A C A A T G G A A C T G G A A T - 3 ' Actin R 5 ' - G G A T A G C A T G T G G A A G T G C A T A C C - 3 ' Keto F 5 - G C A A T G G T G G A A G A G T A A A G T G C - 3 ' Keto R 5 ' - G T T G T G G T G C T C C C A A T G C T T G C G G - 3 ' 1 F: forward, 2 R: reverse 5.2.6. Pigment extraction and HPLC analysis Two homozygous lines per each transformant or co-transformant with high transgene expression were chosen for biochemical analysis. Only one homozygous 97BKB line was available and it was supplemented with a heterozygous line. Two individual plants (T3) from each line were sampled for H P L C analysis. Four to five week old leaf samples were collected and quickly frozen in -80°C until analysis. 109 The following protocol, communicated to me and written by Dr. Craft, was carried out by Craft Technologies on leaf samples that I supplied. Approximately 50 to 150 mg of leaf tissue was weighed into 15 mL glass tubes with Teflon lined caps. The leaf material was extracted with 5 mL of 50% methanol/50% THF. The samples were then saponified by adding 400 L of 10% pyrogallol as an antioxidant and 1 mL of 40% methanolic K O H . Samples were topped with nitrogen gas and incubated at 60°C for 30 min. 3 mL of water was added to each tube and the samples were extracted twice with 4 mL of hexane. The extracts were brought to 10 mL with hexane. The hexane extracts were measured in a spectrophotometer to calculate the approximate total carotenoids. (1) Portions of the hexane extracts were dried under vacuum in a SpeedVac™. The extracts were redissolved in ethyl acetate and diluted in 90% acetonitrile/10% isopropanol. The H P L C system consisted of a computer data system, an autosampler maintaining samples at 20°C, a column heater at 31 °C, a programmable ultraviolet visible detector and a fluorescence detector (Thermo Separation Products, Fremont, CA). The separation was performed isocratically on a Spherisorb O D S 2 column (3 pm, 4.0 x 250 mm with titanium frits, E S Industries, West Berlin, NJ) protected by a Javelin™ guard column containing a similar stationary phase (Thermo Electron Corp, Bellefonte, PA). The mobile phase consisted of acetonitrile/dioxane/isopropanol /triethylamine (80/15/5/0.1) at a flow rate of 1.0 mL/min. The alcohol component contained 100 mM ammonium acetate. The diode array detector with light pipe flow cell was programmed to monitor carotenoids at 450 nm and scanned from 280 nm to 520 nm. (2) Another portion of the hexane extracts were dried under vacuum in a SpeedVac™. The extract was redissolved in hexane. An aliquot was injected onto a Chromegabond Diol normal-phase column (3 urn, 4.0 x 150 mm, E S Industries, West Berlin, NJ) protected by an 110 Amino SecurityGuard™ guard column (Phenomenex, Torrence, CA) and eluted isocratically with 96% hexane/4% isopropanol. The flow rate was 1.5 mL/min. The carotenoids were monitored at 450 nm using a programmable wavelength detector. A 15 |xL injection was used. Extracted samples that had not been saponified were also checked by normal-phase H P L C to check for the presence of astaxanthin. The analytes all possess absorbance which is proportional to their concentration in solution; therefore these properties are used for quantitative analysis. Chromatograms were recorded using a computer data system. Analytes are quantified by external standard quantification using neat standards to calculate response factors based on the peak area of the analyte. For confirmation of peak identities, samples representative of a particular treatment group were concentrated and separated by C30 gradient H P L C with diode array detection. This permitted the collection of visible spectra even of small peaks. Samples were refrigerated at 15° C and the column was maintained at 35° C . Carotenoids were monitored at 450 nm and scanned from 280 to 500 nm. The analytical column was a Y M C C30 (3[xm, 4.6x250mm, Waters, Milford, MA) with a C18 guard. Mobile phase A consisted of 80 THF/20 water with 20 mM ammonium acetate and 0.05% triethylamine. Mobile phase B was acetonitrile containing 0.05% triethylamine. Mobile phase C was ethyl acetate containing 0.05% triethylamine. The gradient held 15% of A throughout. The initial conditions of 75% B and 10% C were held for 2 min. A gradient to 50% B and 35 % C occurred over the next 23 min. Next was a linear gradient to 17% B and 70 % C over 7 min. Then a shallow linear gradient occurred to 10% B and 75% C over 18 min. The final conditions were held for 2 min then returned to initial conditions over 8 min. The column was equilibrated for 10 min before the next injection. The flow rate was 1.0 mL/min. Individual carotenoids were calibrated using peak areas by external standard calibration with multiple concentrations of neat standards. Calibration curves were generated by a computer data system using peak areas. I l l Carotenoids for which no neat calibration standard was available, were calculated using the closest eluting major carotenoid for which a standard was available. All methods were developed from Craft and Furr (2004). H P L C data were analyzed by Least-squares Analysis of Variance (JMP-IN version 5.1.2, S A S Institute, Cary, NC 2004) with the following statistical model: Y i j k = p + Ci + L(C)ij + E i J k Where Y is one of the carotenoid products detected, C denotes all the treatments to be compared (WT, AtB1, AtBKB, 97A3, 97AKB, 97B3, 97BKB, 97C1, 97CKB, and KB), and L(C) denotes the two lines nested in each treatment. Mean separation of significant factors was done by Student's t-tests. Except for 97CKB, the values given in the Tables are the means of 4 measurements: 2 lines, with 2 independent leaf samples per line (Each sample value is the mean of two injections). The mean of the 97CKB represents the mean of 2 measurements (1 line with 2 samples). 5.3. Results 5.3.1. Selection of transformed plants The morphological phenotypes of T1 plants harboring a single transgene were similar to that of wild type (data not shown). T1 plants co-transformed with hydroxylases or candidate hydroxylases and the ketolase (crtO) were also similar to wild type. Seedlings of co-transformed T1 lines (T2 generation) were selected sequentially for hygromycin, and B A S T A resistance to identify co-transformed progeny in this generation. Surprisingly, these plants when grown to maturity showed severe dwarfism (data not shown). 112 However, this phenoty'pe was not inherited in the T3 generation in which seedlings were not subjected to hygromycin and B A S T A selection. This indicates that dwarfism was likely caused by stress from the application of two selection agents and was not due to transgene expression. I obtained between 10 and 31 transgenic lines containing each of the following single double 35S driven transgenes with the given designation. 2x35S::AtB1 (AtB1), 2x35S: :CYP97A3 (97A3), 2x35S: :CYP97B3 (97B3), 2x35S: :CYP97C1 (97C1) and 2x35S::KB (KB). In addition, I obtained between 8 and 10 lines harboring each hydroxylase or putatative hydroxylase plus the ketolase (crtO), both under control of the double 35S promoter. These double transformation lines were designated AtBKB, 97AKB, 97BKB and 97CKB. The presence of the transgene(s) was verified by P C R analysis of genomic DNA isolated from T1 plants (data not shown), which were allowed to self pollinate to generate T2 lines. 5.3.2. Transcript level of single and co-transformed plants Semi-quantitative R T - P C R was performed using RNA isolated from hygromycin or hygromycin plus B A S T A resistant T2 plants to reveal the transcript levels of the transgene in each line and to the amount of hydroxylase and hydroxylase candidate gene relative to wild-type plants. This analysis was carried out for lines harboring all nine single and double transgenes. Figures 5.2 and 5.3 show examples of this analysis, showing examples of high and low transcript levels in representative single and co-transformed lines. I used actin to normalize for variations in cDNA amount of each gene. Relative expression levels were deduced from band intensities. While actin transcript levels were constant between cDNA samples, levels of transgene transcripts varied greatly, with both high and low expressing lines evident (Figures 5.2 and 5.3). As expected, no ketolase expression was detected in wild-type plants (Figure 5.2E), and it was possible to identify AtB1 and P450 candidate hydroxylase lines with greatly elevated expression relative to endogenous gene expression. 113 Single and double transformed lines with the highest transcript levels of one or both genes were chosen for carotenoid metabolite profiling by H P L C . Figure 5.2 Transgene expression levels in individual lines harboring single hydroxylase or ketolase transgenes. R T - P C R was carried out on RNA isolated from 12 lines, and from a wild-type control line. P C R cycle numbers were adjusted so that amplification actin transcripts were was not saturated. An actin control was included for each cDNA sample, to normalize for variations in cDNA amount. Expression in representative lines for each construct are shown. A. AtB1 single transgenic lines, B. CYP97A3 single transgene lines, C. CYP97B3 single transgene lines, D. CYP97C1 single transgenic lines, E. ketolase single transgenic lines. WT. wild-type control line Line 1-3, individual T2 lines for each transgene. High expression lines (bold fonts) were used to H P L C analysis. 114 Figure 5.2 shows single high expression transgene lines (AtB-1 and AtB-3, 97A3-1 and 97A3-2, 97B3-1 and 97B3-3, 97C1-2 and 97C1-3, KB2 and KB3) chosen for H P L C analysis. actin CYP97B3 ketolase WT AtBKBI A tBKB2 AtBKB3 WT 97BKB1 97BKB2 97BKB3 WT 97AKB1 97BAKB2 WT 97CKB1 97CKB2 Figure 5.3 Transgene expression levels in lines harboring both hydroxylase and ketolase trangenes. R T - P C R was carried out on RNA isolated from T2 lines, and from a wild-type control line. P C R cycle numbers were adjusted so that amplification actin transcripts were was not saturated. An actin control was included for each cDNA sample, to normalize for variations in cDNA amount. Expression in representative lines for each construct and double transformants are shown. A. AtBKB: AtB1 and ketolase double trangene lines, B. 9 7 A K B : C Y P 9 7 A 3 and ketolase double transgene lines, C. 9 7 B K B : C Y P 9 7 B 3 and ketolase transgene lines, D. 9 7 C K B : CYP97C1 and ketolase transgene lines. WT. wild-type control line Line 1-3, individual T2 lines for each double transgene. High expression lines (bold fonts) were used to H P L C analysis. 115 Figure 5.3 also shows double transformed lines (AtBKB2 and AtBKB3, 97AKB1 and 97AKB2, 97BKB1 and 97BKB2, and 97CKB1) with highest expression levels chosen for H P L C analysis. I was able to obtain only one 97CKB high expression line, 97CKB1. 5.3.3. Carotenoid profiling For most of the nine single or double transgene combinations, two homozygous lines with high transcript levels were chosen and leaves from two individual T3 plants (samples) from each line were harvested for H P L C analysis of carotenoid content. Only a single high expression 97CKB line was identified and homozygous plants of this genotype were analyzed. Each sample was run twice and the mean of the two runs was used for statistical analysis. Due to the inability to identify two homozygous 97BKB lines with high expression, one heterozygous 97BKB line for both transgenes (with high expression for both genes) was used. Carotenoid extraction and H P L C analysis was carried out by Craft Technolgies, Inc. (North Carolina, USA) , following carotenoid extraction. Concentrations of 21 carotenoids (including isomers) were determined; frans-lycopene, c/s-lycopene, phytoene, phytofluene, echinenone, canthaxanthin, frans-astaxanthin, c/'s-astaxanthin, ^-carotene, ocryptoxanthin, zeinoxanthin, frans-lutein, c/'s-lutein, frans-/?-carotene, c/'s-/?-carotene, /3-cryptoxanthin, trans-zeaxanthin, rrans-violaxanthin, c/s-violaxanthin, frans-neoxanthin, and c/s-neoxanthin. Of these, concentrations of echinenone, canthaxanthin, frans-astaxanthin, c/s-astaxanthin, frans-lycopene, c/s-lycopene, phytoene, and phytofluene were below detectable levels in all samples. Tables 5.5 and 5.6 show summaries of the carotenoid concentrations measured in the wild-type and the nine single or double transformation (the mean represents 2 lines with 2 samples per line.). The data for each transgene combination is presented below. 116 Table 5.5 Profiling of carotenoids derived from a-carotene in Arabidopsis wild-type plants and over expression lines. Means ± SE followed by the same letters are not significantly different (P > 0.05) by Least-squares Analys is of Var iance. In addition, values highlighted in yellow are significantly different between transgenic lines in wild type. Lines are abbreviated as follows: WT, wild-type; AtB1, 2x35S: :AtB1; AtBKB, 2x35S::AtB1 and 2x35S::cr tO; 97A3, 2x35S: :97A3; 97AKB, 2x35S::97A3 and 2X35S::cr fO; 97B3, 2x35S::97B3; 97BKB, 2x35S: :97B3 and 2x35S::crfO; 97C1, 2x35S: :97C1; 97CKB, 2x35S::97C1 and 2x35S::crfO; KB, 2x35S::crtO. Line Carotenoid amount (pg/g of leaf tissue) a-carotene a-cryptoxanthin zeinoxanthin frans-lutein c/s-lutein WT 1.27 ± 0.12 a 0.27 ± 0.04 a 0.24 ± 0.04 0 88.8 ± 7.28 a 6.33 ± 1.03 D c AtB1 1.06 ±0.08 a b 0.13 ± 0.03 ^ 0.29 ± 0.03 c d 69.0 ± 5.15 a 8.19 ± 0 .73 a b AtBKB 0.84 ± 0.08 b c 0.19 ± 0 .03 a b c 0.34 ± 0.03 a b c 75.4 ± 5.15 a 6.63 ± 0.73 b c 97A3 0.84 ± 0.08 0.23 ± 0.03 a b 0.30 ± 0.03 80.6 ± 5.15 a 8.04 ± 0.73 a b 97AKB 0.75 ± 0.08 c 0.15 ± 0.03 0.37 ± 0.03 a b c 75.2 ± 5.15 a 8.72 ± 0.73 a b 0.16 ±0.03 b c d 7.15 ±0.73 a b 97B3 0.79 ± 0.08 c 0.40 ± 0.03 a 73.5 ± 5.15 a 0.16 ±0.03 0.37 ± 0.03 a b 97BKB 0.80 ± 0.08 G 80.6 ± 5.15 a 9.26 ± 0.73 a 0.20 ± 0.03 a b c 0.34 ± 0.03 a b c 6.62 ± 0.73 b c 97C1 0.79 ± 0.08 c 91 .2±5 .15 a 8.60 ± 1.03 a b 97CKB 0.60 ± 0.12 c 0.08 ± 0.03 d 0.39 ± 0.04 a b 88.0 ± 7.28 a KB 0.72 ± 0.08 c 0.19 ±0.04 a b c 0.23 ± 0.03 d 75.2 ± 5.15 a 4.97 ± 0.73 0 Table 5.6 Profiling of carotenoids derived from p -carotene in Arabidopsis wild-type plants and over expression lines. Means ± SE followed by the same letters are not significantly different (P > 0.05) by Least-squares Analysis of Variance. In addition, values highlighted in yellow are significantly different between transgenic lines in wild type. Lines are abbreviated as follows: WT, wild-type; AtB1, 2x35S: :AtB1; AtBKB, 2x35S::AtB1 and 2x35S::crtO; 97A3, 2x35S: :97A3; 97AKB, 2x35S: :97A3 and 2X35S::cr fO; 97B3, 2x35S: :97B3; 97BKB, 2x35S: :97B3 and 2x35S::crtO; 97C1, 2x35S: :97C1; 97CKB, 2x35S::97C1 and 2x35S::crtO; KB, 2x35S::crfO. Line Carotenoid amount (ug/g of leaf tissue) /?-carotene /3-cryptoxanthin zeaxanthin frans-violaxanthin c/s-violaxanthin frans-neoxanthin c/s-neoxanthin WT 41.8 ± 2.70 a 0.39 ± 0.04 D 0.79 + 0.23 D c a 14.7*1 .17° 2.60 ± 0.23 M 14.3 ± 7 . 2 8 ° 12.0 ± 1.03 3 AtB1 38.2 ± 1.91 3 0.51 ± 0.02 3 1.58 ±0 .16 3 31.0 ± 0 . 8 3 3 6 . 4 5 ± 0 . 1 6 3 30.6 ± 5.15 a 6.6 ± 0 . 7 3 3 AtBKB 38.1 ± 1.91 3 0.32 ± 0 .02 b c d 0.54 ± 0.16 d 14.5 ± 0.83 b 2.76 ± 0.16 b c d 14.2 ± 5.15 b 8.9 ± 0 . 7 3 3 97A3 41.7 ± 1.91 a 0.36 ± 0.02 b c 1.62 ± 0 . 1 6 3 15.0 ± 0.83 b 3.00 ± 0.16 b c 14.6 ±5 .15 6 9.2 ± 0 . 7 3 3 97AKB 36.9 ± 1.91 a 0.30 ± 0.02 b c d 1.55 ±0 .16 3 13.2 ± 0.83 b 2.82 ± 0.16 b Q d 12.8 ± 5.15 b 6.7 ± 0 . 7 3 3 97B3 36.7 ± 1.91 a 0.22 ± 0.02 6 1.33 ± 0.16 3 b 14.6 ± 0.83 b 2.83 ± 0.16 b c d 14.0 ± 5.15 b 8.8 ± 0 . 7 3 3 97BKB 44.5 ± 1.91 a 0.29 ± 0.02 b c d e 0.66 ±0 .16 0 6 14.7 ± 0.83 b 3.21 ±0 .16 b 14.3 ± 5.15 b 7.5 ± 0 . 7 3 3 97C1 41.3 ± 1.91 a 0.28 ± 0 . 0 2 d e 1.47 ± 0.16 3 b c 14.1 ± 0 . 8 3 b 2.73 ± 0.16 b c d 13.6 ± 5.15 b 11.3 ± 0.73 a 97CKB 37.6 ± 2.70 a 0.28 ± 0.04 b o d e 1.19 ± 0 . 2 3 a b c 13.8 ± 1.17b 2.60 ± 0.23 c d 13.6 ± 7.28 b 7.6 ± 1.03 3 KB 39.8 i 1.91 a 0.34 ± 0.02 b c d 0.77 ±0 .16 "* 13.8 ± 0.83 b 2.46 ± 0.16 d 13.4 ± 5.15 b 10.1 ±0 .73 3 Table 5.7 Summary of changes in carotenoid product accumulation in single hydroxylase over expression lines and hydroxylase over expression lines co-transformed with the crtO ketolase. Increases or decreases in abundance (j. or t) relative to levels in wild-type plants are shown, using data from Tables 5.5 and 5.6. Hydroxylase Single hydroxylase transformants Double transformants (hydroxylase with ketolase) a-carotene pathway /3-carotene pathway a-carotene pathway /3-carotene pathway AtB1 i a-cryptoxanthin t /2-cryptoxanthin t zeaxanthin t trans-violaxanthin t cis-violaxanthin t trans-neoxanthin J. a-carotene t zeinoxanthin CYP97A3 I a-carotene f zeaxanthin i a-carotene t zeaxanthin i a-cryptoxanthin t zeinoxanthin CYP97B3 i a-carotene i a-cryptoxanthin t zeinoxanthin J, /3-cryptoxanthin I a-carotene fcis-violaxanthin i a-cryptoxanthin t zeinoxanthin f cis-lutein CYP97C1 I a-carotene t zeinoxanthin i y3-cryptoxanthin t zeaxanthin I a-carotene I a-cryptoxanthin t zeinoxanthin WT: H P L C analysis of WT plants showed that lutein and /^-carotene were the major products in WT leaves (Tables 5.5 and 5.6). As expected, no ketocarotenoids (astaxanthin, canthaxanthin, echienone) detected. AtB1: Compared with WT, overexpression of AtB1 resulted in significant increases in products derived from /5-carotene. AtB1 lines over expressing AtB1 alone had significantly (P< 0.05) higher concentrations of /3-cryptoxanthin (0.51 pg/g of leaf vs 0.39 pg/g for WT), trans-zeaxanthin (1.58 pg/g leaf vs. 0.79 pg/g for WT), frans-violaxanthin (31.02 pg/g vs. 14.7 pg/g for WT), c/s-violaxanthin (6.45 pg/g vs. 2.60 pg/g) and frans-neoxanthin (30.6 pg/g vs. 14.3 pg/g). AtB1 over expressing transgenic plants also had significantly lower concentration of ce-cryptoxanthin (0.13 pg/g vs. 0.27 pg/g for WT). There were no significant differences between these carotenoid concentrations in the two high expression AtB1 lines. In the AtB1 and crtO, co-transformed lines, no ketocarotenoid products were detected. Furthermore, the significant increases in all /3-carotene derivatives (,3-cryptoxanthin, trans-zeaxanthin, frans-violaxanthin, c/s-violaxanthin, and frans-neoxanthin) seen in the single gene (Table 5.7). AtB1 overexpressers were not longer detected; the concentrations of these products in the AtBKB plants were not significantly different from those in WT plants (Table 5.6). Instead, there were changes in accumulation of <£-carotene and products derived from o-carotene (Table 5.5). In particular, there was a significant decrease in or-carotene' concentration (0.84 pg/g vs. 1.27 pg/g for WT) and a significant increase in zeinoxanthin concentration (0.34 pg/g vs. 0.24 pg/g for WT) in the AtBKB lines. CYP97B3: Compared with the WT, overexpression of CYP97B3 resulted in significant shifts in the abundance of ocarotene and its derivatives, there were significant decreases in a-carotene (0.79 pg/g vs. 1.27 pg/g for WT) and ^cryptoxanthin (0.16 pg/g vs. 0.27 pg/g for WT) concentrations, while a significant increase in zeinoxanthin concentration (0.40 pg/g vs. 0.24 pg/g for WT) was observed (Table 5.5). 97BKB lines expressing the crtO ketolase gene and overexpresssing CYP97B3 showed similar changes the concentrations of 120 a-carotene and its derivatives relative to wild-type (Table 5.7), but differences between the 97B3 {CYP97B3 overexpression plants) and 97BKB plants were only slight. In addition to the decreased a-carotene and a-cryptoxanthin concentrations and increased zeinoxanthin concentrations similar to 97B3 plants, 97BKB plants also had a significantly higher c/s-lutein concentration (9.26 pg/g) compared to the W T (6.33 ug/g). Accumulation of /3-carotene derivatives in the CYP97B3 overexpressing lines (97B3 and 97BKB) was less affected: 97B3 plants had a significantly lower /3-cryptoxanthin concentration (0.22 pg/g) than W T plants (0.39 pg/g) but the two independent 97B3 overexpression lines behaved differently in that only one of the two was significantly different from WT (data not shown). Although zeaxanthin concentration seemed higher in 97B3 lines (1.33 pg/g vs. 0.79 pg in WT), this difference was not statistically significant, and there were no significant differences in the accumulation of other /3-carotene-derived products in either 97B3 and 97BKB lines except for c/'s-violaxanthin, which was significantly higher in 97BKB (3.21 pg/g) than WT (2.60 pg/g). CYP97C1: Overexpression of CYP97C1 resulted in changes in abundance of both the a- and /3-carotene pathway products. In the a-carotene pathway, CYP97C1 overexpression resulted in significantly decreased a-carotene concentration (0.79 pg/g vs 1.27 pg/g for WT). While there was no overall significant difference in a-cryptoxanthin levels (0.20 pg/g vs 0.27 pg/g for WT), one over expression line had a significantly lower a-cryptoxanthin concentration (0.09 pg/g vs 0.27 pg/g for WT) while the other did not. The lines had a significantly higher zeinoxanthin concentration (0.34 pg/g) than the W T (0.24 pg/g). In the /3-carotene pathway, 97C1 lines exhibited a significant decrease in /3-cryptoxanthin concentration (0.28 pg/g vs 0.39 pg/g for WT). While CYP97C1 overexpression resulted in significantly higher zeaxanthin concentration, there was a significant line difference in that one transgenic line had significantly higher zeaxanthin concentration (2.24 pg/g) than WT (0.79 pg/g) while the other had not (data not shown). 121 The same line with a zeaxanthin concentration similar to W T also had a significantly lower a-cryptoxanthin concentration than the WT, while the line that had a significantly higher zeaxanthin concentration exhibited no increase in a-cryptoxanthin (data not shown). Co-transformation of the CYP97C1 gene with crtO enhanced the effect, of CYP97C1 on the a-carotene pathway but eliminated the effect of CYP97C1 overexpression on the p-carotene pathway (see Table 5.7) CYP97A3: Overexpression of the CYP97A3 gene significantly decreased a-carotene (0.84 pg/g vs 1.27 pg/g for WT) levels but otherwise had no detectable effect on accumulation of a-carotene derivatives. 97A3 plants also had a significantly higher zeaxanthin concentration (1.62 pg/g) compared to W T plants (0.79 pg/g). Co-transformation of the CYP97A3 gene with crtO seemed to affect the a-carotene pathway more so than the /3-carotene pathway. Similar to 97A3 plants, 97AKB plants had a significantly lower a-carotene concentration (0.75 pg/g vs 1.27 pg/g for WT). In addition, 97AKB plants had a significantly lower a-cryptoxanthin concentration than wild-type (0.15 vs 0.27) but higher levels of zeinoxanthin (0.37 pg/g vs 0.24 pg/g in WT). Both 97A3 and 97AKB plants had similar concentrations of /3-carotene derivatives (both had significantly higher zeaxanthin concentrations than WT plants). crtO: Arabidopsis lines expressing the crtO ketolase gene alone (KB) did not yield any detectable ketocarotenoid products. The only difference between KB lines and WT was in o-carotene concentration, which was significantly lower (0.72 pg/g vs. 1.27 pg/g in WT). This indication that crtO activity affects the a-carotene pathway is supported by results from lines in which crtO and hydroxylase genes were co-expressed, as discussed above, where expression of the crtO gene together with overexpression of a hydroxylase generally had a greater effect on the accumulation of a-carotene or its hydroxylation products than overexpression of the hydroxylases alone (See Table 5.7). 122 5.4. Discussion Carotenoids are 40-carbon isoprenoids that are integral and essential components of the photosynthetic membranes in all plants. In Arabidopsis and other plants, carotenoid products are produced either from a- or /3-carotene, via hydroxylation of the e- and /3-rings of a-carotene, or hydroxylation of both /3-rings of /3-carotene (Cunningham and Gantt, 1998). Previous evidence suggested that non-heme di-iron hydroxylases AtB1 and AtB2 are mostly responsible for hydroxylation activities of the /3-rings in the /3-carotene pathway (Sun et al., 1996; Tian and DellaPenna, 2001; Davison et al., 2002) whereas the hydroxylation of the e-and /3-rings of a-carotene is carried out by two cytochrome P450 hydroxylases, C Y P 9 7 A 3 and CYP97C1 (Tien et al., 2004; Kim and Dellapenna, 2006). Tian et al. (2003 and 2004) used double knockout loss of function mutants of AtB1 and AtB2 to show that AtB1 and AtB2 have specific functions in /3-carotene hydroxylation. However, these mutants still contain /3-ring hydroxylated /3-carotene derivatives, suggesting that other hydroxylases activities can also act on /3-carotene. While CYP97C1 is mainly responsible for f-ring hydroxylation, there is evidence that /3-ring hydroxylation deficiency in AtB1 and AtB2 can be partially compensated by up-regulation of CYP97C1 (Tien et al., 2004). Furthermore, while CYP97A3 encodes a /3-ring hydroxylase with major activity towards the /3-ring of a-carotene, the same hydroxylase also has minor activity on the-/3-rings of /3-carotene (Kim and DellaPenna,> 2006). The roles of the P450 hydroxylases in the carotenoid pathways therefore appear not rigid but depend on the relative abundance of the different types of carotenoid hydroxylases and the availability of substrates. 123 In my overexpression study, I measured product accumulation to indirectly deduce enzyme functions and activities. When AtB1 was overexpressed in Arabidopsis, I found significant increases of /2-carotene pathway products (Table 5.6). These results support the hypothesis of Tien et al. (2003) that AtB1 and AtB2 play major roles in /?-carotene ring hydroxylation, and show that my approach of testing in vivo functions of Arabidopsis carotenoid hydroxylases by overexpression in transgenic plants is an effective one. In the AtB1 overexpression lines I also detected a significant decrease in ocryptoxanthin. This is an indication that AtB1, when overexpressed, can also hydroxylate the /J-ring of ocryptoxanthin, but the activity may be relatively minor and not. enough to cause a detectable increase in the large lutein pool. Overexpression of CYP97C1 led to significant decreases in c^carotene and a-cryptoxanthin (in one line) concentrations but an increase in the zeinoxanthin concentration. This is an indication that increased f-ring hydroxylation by CYP97C1 in these lines may have caused lutein production to shift more to the a^cryptoxanthin route and away from the normally major zeinoxanthin route. Alternatively, CYP97C1 may have /3-ring hydroxylase function on both a-carotene and /2-carotene. Our overexpression lines also had significantly lower /3-cryptoxanthin and higher zeaxanthin concentrations, supporting the finding of Tien et al. (2004) that up-regulation of CYP97C1 will cause the enzyme to hydroxylate the /3-rings of P-carotene as well. My two overexpression lines also behaved differently with respect to / k i n g hydroxylation, suggesting that such activity may vary in with due to uncontrolled difference in expression levels or other factors. When I overexpressed CYP97A3, I found a significant decrease in ocarotene concentration and a significant increase in zeaxanthin concentration, thus supporting the notion that C Y P 9 7 A 3 has activity towards the /J-ring of ocarotene as well as minor activity on the /3-rings of /5-carotene (Kim and DellaPenna, 2006). 124 Apparently, overexpession of CYP97A3 is able to direct metabolic flux from the small pool of a-carotene towards its hydroxylated derivatives, and into zeaxanthin in the /3-carotene pathway. My overexpression study complements the knockout studies by Tien et al. (2003 and 2004) and Kim and Dellapenna (2006). While double and triple mutation combinations involving the AtB1, AtB2, CYP97A3 and CYP97C1 were successful in completely blocking /3-carotene hydroxylation, /3-ring hydroxylation of a-carotene was still observed (Tian et al., 2004; Kim and DellaPenna, 2006; Fiore et al., 2006) and the existence of a fifth hydroxylase, acting on the /3-ring of a-carotene but not /3-carotene is suspected (Fiore et al., 2006). The Arabidopsis genome contains another C Y P 9 7 family member, C Y P 9 7 B 3 , which is 42% identical to CYP97C1 protein (Tian ef al., 2004). Overexpression of CYP97B3 led to significant decreases in a-carotene and o-cryptoxanthin concentrations and a significant increase in zeinoxanthin concentration. These results provide evidence that C Y P 9 7 B 3 is the suspected additional /3-ring hydroxylase in the o-carotene pathway, with activity in /3-ring hydroxylation of a-carotene, since overexpression seems to preferentially shift the pathway to the production of the /3-ring intermediate (zeinoxanthin) and away from the £-ring intermediate (a-cryptoxanthin). However, I also found that overexpression of CYP97B3 significantly decreased /3-cryptoxanthin concentration, suggesting possible activity in /3-ring hydroxylation. This is supported by the finding (Chapter 4; Kim et al., 2005) that expression of C Y P 9 7 B 3 in a /3-carotene producing E.coli strain could lead to generation of /3-cryptoxanthin and zeaxanthin from /3-carotene. This is in contradiction to the prediction of Fiore et al. (2006) that the additional fifth hydroxylase would not act on the /3-rings of /3-carotene. Furthermore, Schuler and Werck-Reichhart (2003) predicted that C Y P 9 7 B 3 is targeted to the mitochondria and not chloroplasts. Further studies are therefore needed to evaluate the role of C Y P 9 7 B 3 in the carotene pathways. 1 2 5 Ketolase activity catalyzes the addition of a keto group to the 4 position of one or both rings of the /2-carotene to produce ketocarotenoids like echinenone and canthaxanthin. Ketolase genes and activity in higher plants have, to date, only been found in Adonis paleastina. However, synthesis of ketocarotenoids in transgenic plants has been demonstrated in leaves and flowers of tomatoes and tobacco (Ralley et al., 2004), tubers of potatoes (Gerjets and Sandmann, 2006; Morris et al., 2006), flowers of Lotus japonicus (Suzuki et al., 2007), roots of carrot (See Chapter 3) and seeds of Arabidopsis (Stalberg et al., 2003) by expression of heterologous ketolase genes. I transformed the ketolase gene crtO from Haematococcus pluvialis (see Chapter 3) into Arabidopsis but found no detectable levels ketocarotenoids in leaves of 4 week old transgenic plants. Stalberg et al. (2003) found that the yield of ketocarotenoids in their transgenic Arabidopsis plants were low, even with the co-transformation of a phytoene synthase construct to boost the carotenoid content. They speculated that the efficient hydroxylation of both the a- and /3-rings made the hydroxylated carotenes less attractive for carbonylation. Substrate availability and differences in temporal expression of the introduced gene in relation to endogenous biosynthesis of carotenoids are all factors contributing to the low yield of ketocarotenoids in the transgenic plants, which may have been below the level of detection in my ketolase transgenics. The only difference in carotenoid content in the ketolase transgeneic was an unexpected decrease of ^-carotene content in our transgenic plants. Stalberg et al. (2003) and Morris et al. (2006) transformed the same algal ketolase that I used in my study and detected 4-keto-lutein (3,3'-dihydroxy-/?,£-carotene-4-one), one of o-carotene derivatives, in the seeds of Arabidopsis and potato tubers, respectively. Although 4-keto-lutein was not detected in my transgenic lines, other /3-ring derivatives for which we have no available standards could have been produced (Figure 5.4). In order to generate a metabolic change by transfer of a ketolase transgene, it is often necessary to co-transform it with a hydroxylase gene (Ralley et al., 2004). I co-transformed the cdO gene with each of the 4 carotenoid hydroxylase genes, but no ketocarotenoids were 126 detected in any of four co-transformed lines, A tBKB, 97AKB, 97BKB, and 97CKB. However, in all co-transformants, the hydroxylation activity in the a-carotene pathway appeared to be altered, based on the accumulation of hydroxylated carotenoid products in single versus double transformants. This effect is most obvious in the A tBKB lines, where all of the increases in /3-carotene products resulting from overexpression of AtB1 are abolished by co-expression of the ketolase. At the same time, a-carotene product accumulation increased. Comparisons of the similarities between AtB1 and crtO ketolase showed that both enzymes belong to membrane-integral, di-iron oxygenase enzymes which share common amino acid residues such as histidine motifs and transmembrane helix (Cunningham and Gantt, 1998). CrtO ketolase has been suggested to be mechanistically related to AtB1 since both enzymes require the presence of conserved histidine residues for activity in pepper fruits and Haematococcus pluvialis (Bouvier et al., 1998; Ye et al., 2006). Another example of similarity between ketolase and hydroxylase proteins is evident in the Adonis ketolase polypeptide, which has more than 6 0 % similarity to Arabidopsis /3-carotene hydroxylases. Sun et al. (1996) speculated that the AtB1 normally associates with a second /3-hydroxylase (or with e-hydroxylase) to form a dimer and that a portion of the N terminus mediates these subunit interactions. One possible explanation for the effect of the ketolase expression on hydroxylasese activity in my transgenics is that when AtB1 and ketolase are integral to the plastid membrane, the ketolase might form a herterodimer with AtB1, altering the substrate preference of the hydroxylase. Therefore, the interaction between these two enzymes might have shifted enzyme activities from the /3-carotene to the a-carotene pathway. Further investigation along this line would be worthwhile. My interpretations of enzyme functions based on product accumulation in overexpressing lines assume that interconversion between a-carotene and /3-carotene products do not take place in vivo. As far as I am aware, there is no evidence that such interconversions can take place, suggesting that this is a valid assumption. 127 In summary, this overexpression study has confirmed the roles that the three carotenoid hydroxylases, AtB1, CYP97A3 , and C Y P 9 7 C 1 , play in the carotenoid pathways. I have also for the first time demonstrated the hydroxlase function of C Y P 9 7 B 3 in the carotenoid pathways, and also opened up questions regarding (1) cross-over activities of hydroxylases in the a- and /3-carotene pathways, (2) the interactions between the crtO transgene and the hydroxylases in carotenoid biosynthesis. CrtO? or-Carotene CYP97C1 a-Cryptoxanthin / \ CYP97A3, CYP97B3 Zeinoxanthin CYP97A3 \ , J CYP97C1 Lutein AtB1 CrtO n ^ /5-Carotene H(r*^ ° F rh inonr tna ACryptoxanthin AtBI Echinenone CrtO Zeaxanthin CrtO Canthaxanthin AtB1 I-Adonixanthin Adonirubin CrtO AtB1 Astaxanthin Figure 5.4 The potential enzyme function of ketolase in a-carotene pathway in Arabidopsis transformed with CrtO. Ketolase is functional to produce ketocarotenoids with /3-ring hydroxylase (AtB1). C Y P 9 7 A 3 and C Y P 9 7 B 3 showed the major /3-ring hydroxylase function toward a-carotene. CYP97C1 has £-ring hydroxylase funtion. AtB1 showed /3-ring activity toward /3-carotene. Co-transformation with CrtO boosted the activity of a-carotene pathway 128 5.5. References Alvarez V, Rodriguez-Saiz M, de la Fuente JL, Gudina EJ, Godio RP, Martin, JF, Barredo JE. 2006. The crtS gene of Xanthophyllomyces dendrorhous encodes a novel cytochrome-P450 hydroxylase involved in the conversion of /3-carotene into astaxanthin and other xanthophylls. Fungal Genetics and Biology 43, 261-272. Blasco F, Kauffmann I, Schmid RD. 2004. CYP175A1 from Thermus thermophilus HB27, the first beta-carotene hydroxylase of the P450 superfamily. Applied Microbiology and Biotechnology 64, 671-674. Bouvier F, Backhaus RA, Camara B. 1998. Induction and control of chromoplast-specific carotenoid genes by oxidative stress. Journal of Biological Chemistry 273, 30651-30659. Clough SJ, Bent AF. 1998. Floral dip: a simplified method for Agrobacterium-med\a\ed transformation of Arabidopsis thaliana. The Plant J16, 735-743. Craft NE, Furr HC. 2004. Improved H P L C analysis of retinol and retinyl esters, tocopherols, and carotenoids in human serum samples for the N H A N E S . FASEB Journal 18, A534. Cunningham Jr FX, Gantt, E. 1998. Genes and enzymes of carotenoid biosynthesis in plants. Annual Review of Plant Physiology and Plant Molecular Biology 49, 557-583. Cunningham Jr FX, Lee H, Gantt E. 2007. Cartenoid biosynthesis in the primitive red alga Cyanidioschyzon merolae. Eukaryotic Cell 6, 533-545. Davison PA, Hunter CN, Horton P. 2002. Overexpression of beta-carotene hydroxylase enhances stress tolerance in Arabidopsis. Nature 418, 203-206. DellaPenna, D. 2001. Carotenoid synthesis and function in plants: Insights from mutant studies in Arabidopsis thaliana. In. The Photochemistry of Carotenoid. ed. pp21-37 Kluwer Academic, Dordrecht. Dufosse L, de Echanove MC. 2005. The last step in the biosynthesis of aryl carotenoids in 129 the cheese ripening bacteria Brevibacterium linens A T C C 9175 (Brevibacterium aurantiacum sp. nov.) involves a cytochrome P450-dependent monooxygenase. Food Research International 38, 967-973. Fiore A, Dall'Osto L, Fraser FD, Bassi R, Giuliano G. 2006. Elucidation of the ,3-carotene hydroxylation pathway in Arabidopsis thaliana. FEBS Letters 580, 4718-4722. Galpaz N, Ronen G, Khalfa Z, Zamir D, Hirschberg J. 2006. A chromoplast-specific carotenoid biosynthesis pathway is revealed by cloning of the tomato white-flower locus. Plant Cell 18, 1947-60. Gerjets T, Sandmann G. 2006. Ketocarotenoid formation in transgenic potato. Journal of Experimental Botany 57, 3639-3645. Hundle BS, Alberti M, Nievelstein V, Beyer P, Kleinig H, Armstrong GA, Burke GA, Kleing H, Hearst JE. 1994. Functional assignment of Erwinia herbicola Eho10 carotenoid genes expressed in Escherichia coli. Molecular Genetics and Genetics. 245, 406-416. Inoue K. 2004.Carotenoid hydroxylation~P450 finally! Trends in Plant Science. 9, 515-517. Niyogi KK. 1999. Photoprotection revisited: genetic and molecular approaches. Annual Review of Plant Physiology and Plant Molecular Biology 50, 333-359. Kim JE, Punja, ZK, Douglas CJ. 2005. Co-expression of Arabidopsis thaliana cytochrome P450 enzymes and NADPH-cytochrome P450 reductase in Escherichia coli. Testing the function of candidate /3-carotene hydroxylases.: In P450 Systems and Regulation, Proceedings of 14 t h International Conference on Cytochromes P450, Dallas, TX, 2005: Medimond S.r.l, Italy. pp115-120. Kim J, DellaPenna D. 2006. Defining the primary route for lutein synthesis in plants: the role of Arabidopsis carotenoid / k i n g hydroxylase C Y P 9 7 A 3 . Proceedings of the National Academy of Sciences USA 103, 3474-3479. Kay R, Chan A, Daly M, McPherson J. 1987. Duplication of C a M V 35S promoter sequences 130 creates a strong enhancer for plant genes. Science 236, 1299-1300. Linden H. 1999. Carotenoid hydroxylase from Haematococcus pluvialis: cDNA sequence, regulation and functional complementation. Biochimica et Biophysica Acta. 1446, 203-212. Masamoto K, Misawa N, Kaneko T, Kikuno R, Toh H. 1998. /J-Carotene hydroxylase gene from the Cyanobacterium Synechocystis sp. PCC6803 . Plant and Cell Physiology. 39, 560-564. Misawa N, Nakagawa M, Kobayashi K, Yamano S, Izawa Y, Nakamura K, Harashima K. 1990. Elucidation of the Erwinia uredovora carotenoid biosynthetic pathway by functional analysis of gene products expressed in Escherichia coli. Journal of Bacteriology 172, 6704-6712. Misawa N, Satomi Y, Kondo K, Yokoyama A, Kajiwara S, Saito T, Ohtani T, Miki, W. 1995. Structure and functional analysis of a marine bacterial carotenoid biosynthesis gene cluster and astaxanthin biosynthetic pathway proposed at the gene level. Journal of Bacteriology 177, 6575-6584. Morris WL, Ducreux LJM, Fraser PD, Millam S, Taylor MA. 2006. Engineering ketocarotenoid biosynthesis in potato tubers. Metabolic engineering 8, 253-263. Ralley L, Enfissi EMA, Misawa N, Schuch W, Bramley PM, Fraser PD. 2004. Metabolic engineering of ketocarotenoid formation in higher plants. The Plant Journal 39, 477-486. Quinlan RF, Jaradat TT, Wurtzel ET. 2007. Escherichia coli as a platform for functional expression of plant P450 carotene hydroxylases. Archives of Biochemistry and Biophysics 458, 146-157. Schoefs B, Rmiki N-E, Rachadi J, Lemoine Y. 2001. Astaxanthin accumulation in Hematococcus requires a cytochrome P450 hydroxylase and an active synthesis of fatty acids. FEBS Letters 500,125-128. Schuler MA, Werck-Reichhart D. 2003. Functional genomics of P450s. Annual Review of 131 Plant Biology 54, 629-667. Stalberg K, Lindgren O, Ek B, Hoglund A-S. 2003. Synthesis of ketocarotenoids in the seed of Arabidopsis thaliana. The Plant Journal 36, 771-779. Sun Z, Gantt E, Cunningham Jr. FX. 1996. Cloning and functional analysis of the beta-carotene hydroxylase of Arabidopsis thaliana. Journal of Biological Chemistry 271, 24349-24352. Suzuki S, Nishihara M, Nakatsuka T, Misawa N, Ogiwara I, Yamamura S. 2007. Flower color alteration in Lotus japonicus by modification of the carotenoid biosynthetic pathway. Plant Cell Report 26, 951-959. Tian L, DellaPenna D. 2001. Characterization of a second carotenoid /^-hydroxylase gene from Arabidopsis and its relationship to the LUT1 locus. Plant Molecular Biology 47, 379-388. Tian L, Magallanes-Lundback M, Musetti V, DellaPenna D. 2003. Functional analysis of /J-and e- ring carotenoid hydroxylase in Arabidopsis. The Plant Ce//15,1320-1332. Tian L, Musetti V, Kim J, Magallanes-Lundback M, DellaPenna D. 2004. The Arabidopsis LUT1 locus encodes a member of the cytochrome P450 family that is required for carotenoid £-ring hydroxylation activity. Proceedings of the National Academy of Sciences USA 101, 402-407. Ye RW, Stead KJ, Yao H, Heo H. 2006. Mutational and Functional Analysis of the /3-Carotene Ketolase Involved in the Production of Canthaxanthin and Astaxanthin. Applied and Environmental Microbiology 72, 5829-5837. 132 CHAPTER 6 GENERAL DISCUSSION AND CONCOUSION Carotenoid pigments are integral and essential components of the photosynthetic membranes in all plants, algae and cyanobacteria. Carotenoids serve a wide variety of important functions in plants (Cunningham and Gantt, 1998). In carrot roots, ,3-carotene accumulates at high levels and it is considered to be a good model for examining carotenoid biosynthesis. While it has been postulated that carotenoids are synthesized and accumulate in chromoplasts in flowers, fruits, and roots (Whatley and Whatley, 1987), studies on carrot root cell chromoplast ultrastructure and the sites of /3-carotene accumulation in carrot roots have been limited and were published over two decades ago (Frey-Wyssling and Schwegler, 1965; Ben-Shaul et al., 1968; Grote and Fromme, 1978). Problems were encountered mainly because of the fibrous nature of the carrot root and the extraction of carotenoids by the fixation solvents and these provide challenges to sample preparation for transmission electron microscopy (TEM) studies. I attempted several fixation techniques (Chapter 2) including microwave treatment (Giberson and Demaree, 1999) of the root tissue during the fixation process, the use of LR white and LR gold for embedding (Bozzola and Russel l , 1999), and High Pressure Freezing (Walther and Muller, 1999) to alleviate the carotene extraction problem but to no avail. I also explored the use of confocal and polarized light microscopy because tissue preparation for light microscopy would not extract carotene. Between the two, polarized light microscopy showed some promise because polarized light seems to show the carotene crystals in cells and may offer the ability to create three-dimensional stacks for volumetric analysis. Further exploration along this line seems warranted. While the use of High Pressure Freezing did not stop the extraction of carotene, it did result in high definition electron micrographs that clearly depict crystalline shaped spaces, assumed to be left by extracted carotene crystals, in the chromoplasts. By comparing the amount of these spaces in three varieties of carrots, H C M (high carotene), orange (regular carotene level), and white (very low carotene), I provided evidence to confirm that these spaces were indeed occupied by 133 the carotene crystals before they were extracted by the fixation process. The confirmation that carotenoid biosynthesis is carried out in the chromoplasts was foundational work for the transformation study that is described in Chapter 3. Carotenoids in algae and higher plants not only are involved in harvesting light for photosynthesis, but also in photoprotection by quenching the reactive oxygen species and other oxidizing molecules generated during the process of photosynthesis (Niyogi, 1999). As such, some carotenoid products are strong anti-oxidants. Carotenoids also function as coloring agents in flowers and fruits to attract pollinators and agents of seed dispersal, respectively (Cunningham and Gantt, 1998). There has been considerable interest in the manipulation of carotenoid content and composition in plants to improve their nutritional value for human and animal consumption. Astaxanthin (3,3'-dihydroxy-4,4'-diketo-/3-carotene) is a strong anti-oxidant that has immune function in the visual and cardiovascular system (Van den Berg et al., 2000). It has also been exploited as fish feed additive to provide the attractive red colour in salmon flesh. Astaxanthin is formed from /^-carotene by the introduction of keto and hydroxyl moieties at the 4,4' and 3,3' positions, respectively, of the /3-ionone rings. The two critical enzymes involved are /3-carotene ketolase and /3-carotene hydroxylase. While /3-carotene hydroxylase is common in higher plants, /3-carotene ketolase is not. Except for Adonis spp, ketocarotenoids have not been described in green plants. I transferred a /3-carotene ketolase gene isolated from algae to carrots (Chapter 3) to examine the possibility of increasing astaxanthin production in carrots, which could be mass-produced to provide a cheap source of astaxanthin. Synthesis of ketocarotenoids in transgenic plants has been demonstrated in leaves and flowers of tomato and tobacco (Ralley et al., 2004), tubers of potato (Gerjets and Sandmann, 2006; Morris et al., 2006), flowers of Lotus japonicus (Suzuki et al., 2007) and seeds of Arabidopsis (Stalberg et al., 2003). Similar to these previous attempts, our research team was also successful in finding astaxanthin and canthaxanthin in transgenic carrots. 134 It remains to be determined whether such transgenic carrots will grow normally, and whether the ketocarotenoids can be extracted from the carrot root efficiently in high enough quantities for economical biological production of astaxanthin. Xanthophylls are oxygenated derivatives of carotenes that perform critical roles in the photosynthetic apparatus of higher plants (Niyogi, 1999). Lutein (3fl,3'/ :?- /3,£--carotene-3,3'-diol) is the most abundant xanthophyll in all plant photosynthetic tissues (Tian ef al., 2004) and zeaxanthin (3fl,3'fl-/3,/3-carotene-3,3'-diol) is a structural isomer of lutein that is involved in thermal dissipation and photoprotection (Niyogi, 1999). Both lutein and zeaxanthin are dihydroxy xanthophylls. Lutein is derived by the addition of hydroxyl group to the 3, 3' position of the e- and /3-rings of ^-carotene with the help of e- and /3-ring hydroxylases. So far, four carotenoid hydroxylases have been identified in Arabidopsis: non-heme di-iron monoxygenases, AtB1 (Sun ef al., 1996) and AtB2 (Tian and Del laPenna, 2001) and cytochrome P450 type monooxygenases, CYP97C1 (Tian ef al., 2004) and C Y P 9 7 A 3 (Kim and Del laPenna, 2006). AtB1 and AtB2 are thought to specialize on hydroxylation of the /3-rings of /3-carotene (Tian etal., 2004) whereas the C Y P hydroxylases, 97A3 and 97C1, are primarily active in the ^-carotene pathway, but have minor activity in the /3-carotene pathway as well (Kim and Del laPenna, 2006; Fiore ef al., 2006). To further study the roles that these hydroxylases play in the carotene pathways, I isolated the AtB1, CYP97A3, and CYP97C1 hydroxylase genes from Arabidopsis and performed single gene transformations of these genes back into Arabidopsis to create overexpression lines for each of these three genes (Chapter 5). I found that, while AtB1 hydroxylase was mostly active in the /3-carotene pathway, it had minor cross-over activity in the ^-carotene pathway when overexpressed. Because of its importance for maintaining photosynthesis, and ultimately for the survival of photosynthetic organisms in a variety of environments, a functional photoprotection mechanism is essential. Carotenoids involved in photoprotection, such as lutein and zeaxanthin, are derived from both the o> and ,3-carotene pathways. 135 It therefore makes sense that evolution would allow cross-over activities of these carotenoid hydroxylases so that they can compensate for each other under extraordinary circumstances. Results from gene knockout studies predict the existence of a fifth yet unknown /3-ring hydroxylase with primary (and maybe exclusive) activity in the a-carotene pathway. The Arabidopsis genome also contains another C Y P 9 7 family member, C Y P 9 7 B 3 , which is 42% identical to the C Y P 9 7 C 1 protein (Tien ef al., 2004). I conducted a heterologous expression study of this gene using an E.coli strain that was previously engineered for endogenous B-carotene production (Chapter 4) and found that C Y P 9 7 B 3 has hydroxylase activity and allowed biosynthesis of zeaxanthin and /3-cryptoxanthin in E.coli, although this activity appeared weaker than that of AtB1. I also created an overexpression line in Arabidopsis (Chapter 5) and demonstrated that C Y P 9 7 B 3 is a new /3-ring hydroxylase in Arabidopsis. Further studies are needed to define the role of this hydroxylase in the carotene pathways. Contrary to ketolase transformation in tobacco and tomato, the co-transformation of a hydroxylase gene into the carrot was not necessary for ketcarotenoid production. Substrate availability and differences in temporal expression of the introduced gene in relation to endogenous biosynthesis of carotenoids may be all factors contributing to the variation in yield of ketocarotenoids in the transgenic plants of different species. It is of interest to examine the working relationship between the ketolase and the hydroxylases in the carotenoid synthesis pathways. I used Arabidopsis thaliana as a model because the crtO ketolase and AtB1 hydroxylase (endogenous in Arabidopsis) are members of the same class of membrane-integral, di-iron oxygenase enzymes and share many common amino acid residues (Bouvier et al., 1998). I transformed the algal crtO gene to Arabidopsis and also co-transformed the same crtO gene with each of the four carotenoid hydroxylase genes (AtB1, CYP97A3, CYP97B3, and CYP97C1) to create 4 co-transformants (Chapter 5) and examined their carotenoid product profiles. No ketocarotenoids were detected in the single gene transformants or in the 4 co-transformants. However, the production of other ketolase derivatives in the a-carotene and 136 /J-carotene pathways could not be ruled out. The visible spectra of our H P L C analyses showed several minor unidentified peaks. The logical next step would be to isolate enough of each to perform L C / M S to obtain molecular weight as Stalberg et al. (2003) have done. In all the co-transformed transgenic plants, hydroxlase activities in the a-carotene pathway were enhanced at the cost of hydroxylation activities in the /3-carotene pathway. Bouvier et al. (1998) also predicted that /3-carotene ketolases and carotenoid hydroxylases are mechanistically related because of the homology in their histidine motifs. Another example of similarity between ketolase and hydroxylase is that the Adonis ketolase polypeptide has more than 60% similarity with Arabidopsis /3-carotene hydroxylases. In case of our co-transformants, the ketolase may interact with the hydroxylase and alter the substrate preference of the hydroxylase. I believe these co-transformants will provide valuable models for studying the working relationship between /3-carotene ketolases and carotenoid hydroxylases. Future research directions could be along the following lines: (1) Functional analyses using C Y P 9 7 B 3 single knockout mutant, double and triple knockout mutants with C Y P 9 7 A 3 and CYP97C1 (Tien et al., 2004), (2) heterologous expression of the C Y P hydroxylases in an a-carotene producing E.coli strain (Farmer and Liao, 2000), and (3) using hydroxylase-GFP fusions (Ro et al., 2002) or antibody detection to define the subcellular localization of the C Y P proteins studied. In summary, my thesis research made contributions in new basic information about the carotenoid hydroxylases and a /3-carotene ketolase, as well as facilitating the application of some of the information to the production of astaxanthin in carrot roots. 137 6.1. References Ben-Shaul Y, Treffry T, Klein S. 1968. Fine structure studies of carotene body development. Journal of Microscopie 7, 265-274. Bourvier F, Keller Y, D'Harlingue A, Camara B. 1998. Xanthophyll biosynthesis: molecular and functional characterization of carotenoid hydroxylases from pepper fruits (Capsicum annuum L ) . Biochimica et Biophysica Acta 1391, 320-328. Bozzola JJ, Russell LD. 1999. Electron Microscopy: Principles and Techniques for Biologists. 2nd edn. Boston, Jones & Bartlett Publishers. Cunningham Jr. FX, Gantt E. 1998. Genes and enzymes of carotenoid biosynthesis in plants. Annual Review of Plant Physiology and Plant Molecular Biology 49, 557-583. Farmer WR, Liao JC. 2000. Improving lycopene production in Escherichia coli by engineering metabolic control. Nature Biotechnology IB, 533-537. Fiore A, Dall'Osto L, Fraser FD, Bassi R, Giuliano G. 2006. Elucidation of the /3-carotene hydroxylation pathway in Arabidopsis thaliana. FEBS Letters 580, 4718-4722. Frey-Wyssling A, Schwegler F. 1965. Ultrastructure of the chromoplasts in the carrot root. Journal of Ultrastructure Research 13, 543-559. Gerjets T, Sandmann G. 2006. Ketocarotenoid formation in transgenic potato. Journal of Experimental Botany 57, 3639-3645. Grote M, Fromme HG. 1978. Electron Microscopic studies in cultivated plants. II. Fresh and Stored Roots of Daucus Carota L. Z. Lebensm.Unters.-Forsch 166, 74-79. Giberson RT, Demaree RS Jr. 1999. Microwave processing techniques for electron microscopy: A four-hour protocol. In: Electron Microscopy Methods and Protocols. N. Hajibagheri, ed. Humana Press, Inc., Totowa, NJ . Kim, J, DellaPenna D. 2006. Defining the primary route for lutein synthesis in plants: the role 138 of Arabidopsis carotenoid beta-ring hydroxylase C Y P 9 7 A 3 . Proceedings of the National Academy of Sciences 103, 3474-3479. Morris WL, Ducreux LJ, Fraser PD, Millam S, Taylor MA. 2006. Engineering ketocarotenoid biosynthesis in potato tubers. Metabolic Engineering 8, 253-263. Niyogi KK. 1999. Photoprotection revisited: genetic and molecular approaches. Annual Review of Plant Physiology and Plant Molecular Biology 50, 333-359. Ralley L, Enfissi EMA, Misawa N, Schuch W, Bramley PM, Fraser PD. 2004. Metabolic engineering of ketocarotenoid formation in higher plants. The Plant Journal 39, 477-486. Ro DK, Ehlting J, Douglas CJ. 2002. Cloning, functional expression, and subcellular localization of multiple NADPH-cytochrome P450 reductases from hybrid poplar. Plant Physiology 130, 1837-1851. Stalberg K, Lindgren O, Ek B, Hoglund A-S. 2003. Synthesis of ketocarotenoids in the seed of Arabidopsis thaliana. The Plant Journal 36, 771-779. Sun ZR, Gantt E, Cunningham Jr FX. 1996. Cloning and functional analysis of the /3-carotene hydroxylase of Arabidopsis thaliana. Journal of Biological Chemistry 271, 24349-24352. Suzuki S, Nishihara M, Nakatsuka T, Misawa N, Ogiwara I, Yamamura S. 2007. Flower color alteration in Lotus japonicus by modification of the carotenoid biosynthetic pathway. Plant Cell Report 26, 951-959. Tian L, DellaPenna D. 2001. Characterization of a second carotenoid beta-hydroxylase gene from Arabidopsis and its relationship to the LUT1 locus. Plant Molecular Biology 47, 379-388. Tian L, Musetti V, Kim J, Magallanes-Lundback M, DellaPenna D. 2004. The Arabidopsis LUT1 locus encodes a member of the cytochrome P450 family that is required for carotenoid e-r\r\g hydroxylation activity, Proceedings of the National Academy of Sciences USA 101, 402-407. 139 Van den Berg H, Faulks R, Fernando Granado H, Hirschberg J, Olmedilla B, Sandmann G, Southon S, Stahl W. 2000. The potential for the improvement of carotenoid levels in foods and the likely systemic effects. Journal of the Science of Food and Agriculture 80, 880-912. Walther P, Miiller, M. 1999. Biological ultrastructure as revealed by high resulution cryo S E M of block faces after cryo-sectioning. Journal of Microscopy 279-287. Whatley JM, Whatley FR. 1987. When is a chromoplast? New Phytologist 106, 667-678. 140 A P P E N D I C E S Appendix 1 A1. Methods for Carotenoid Extraction from Carrot Root (from Chapter 3) For carotenoid extraction from carrot roots, raw, steamed and freeze-drying methods were tested with two different solvent systems. The optimal extraction method was used for analysis of the ketolase transgenic carrot lines (Chapter 3). A1.1 Preparation of carrot root samples Carrot roots were treated 3 ways before extraction: 1) Fresh carrot root was cut by hand (1 cm 2) and homogenized, 2) Carrots were put in a steamer and steamed for 25 min. Around 5 g of the steamed carrot were homogenized, and 3) Carrot root was in -20°C freeze-dryer for 2 days before homogenizing. Samples were weighed before and after freezing to measure moisture loss. A1.2 Extraction with two different solvent systems Samples were extracted with: 1) [chloroform: methanol: water (20:20:8, v/v/v)]. Samples were homogenized with [chloroform and methanol] for 30 s. Water was added and homogenized for additional 20 s. After the two phases were well separated, the chloroform phase containing the extracted pigment was collected using a separatory funnel. This process was repeated two to three 141 times. The chloroform phase was evaporated to complete dryness. 2) [acetone: petroleum: water (50:50:10, v/v/v)] by a homogenizer. After homogenizing the samples with acetone, they were transferred to the vacuum filter. Acetone was added to samples on the filter paper until the orange color disappeared. The pigment in acetone was collected and transferred to the funnel. Petroleum and water were added to the acetone and allowed to settle into two separate layers. Since the petroleum containing pigment was upper layer of the two phases, it was not easy to collect pigments in upper layer. The upper layer was evaporated completely. It is dissolved in [acetonitrile: ethyl acetate (70:30, v/v)] (Mann et al., 2000). An extract is filtered by 0.22 pm filter membrane and store at -40°C before H P L C analysis. A1.3 Carotenoid analysis Carotenoid analyses were conducted using reverse-phased H P L C and a Spherisorb O D S 2 column (4.6 mm x 250 mm) (Waters Co. , USA). Solvent gradient of (A) Acetonitril (70%) (B) Ethyl acetate (30%) were used. The flow rate is set at 1.0ml min-1. Compunds detected at 450 nm wavelength were identified and quantitified using authentic standards. (Sigma, Co. , Canada and Prodemex Co. , Mexico). H P L C analysis showed clear peak separation of the major carotenoids in carrots (Figure 1.1) using the procedure developed. The steam method and chloroform extraction showed the best result. A1.4 Authentic standards for HPLC analysis Commercial authentic standards, a-carotene and /3-carotene (Sigma-Aldrich Canada Ltd., Oakville, Ontario), canthaxanthin (Hoffmann LaRoche, Basle, Switzerland) and lutein and zeaxanthin (Prodemex Co. , Los Mochis, Mexico), were used. For preparation of standards, a-, /3-carotene, canthaxanthin, astaxanthin, 0.03 g of standard is weighed. Butylated hydroxytoluene (BHT), as an antioxidant and toluene are added. It is massed up to volume 1 4 2 100ml of [hexane: acetone (80:20, v/v)] and further diluted for sample analysis. For lutein and zeaxanthin, 0.03 g of standard is added to ethanol and acetone and made up to a volume of 100ml of Hexene. For working solution, it is diluted with hexane (Weber, 1995). Aliquots of these standard solutions were evaporated to dryness and then a mixture of 70:30 acetonitrile/ethyl acetate was added to each of the dry standards so that they were suitable for use with a reverse phase column. Results for standards are shown in Figure 1. Samples from carrots were run on a Waters LC1 equipped with a UV detector. Standard 1 2 Carotenoids i u n s A u x i> ^ 400 Mill tiles 1100 1400 Carrot Carotenoids J 3 00 10.00 Minutes 1200 1400 1800 Figure A1.1 The H P L C results of steamed carotenoid extraction method from carrot root. Major carotenoid compounds in carrot root. 1. cathaxanthin, 2. zeaxanthin, 3. lutein, 4. a-carotene, 5. /2-carotene A1.5 References Mann V, Harker M, Pecker I, Hirschberg J. 2000. Metabolic engineering of astaxanthin production in tobacco flowers. Nature Biotechnology\8, 888-892. Kiessling A, Dosanjh B, Higgs D, Deacon G, Rowshandeli N. 1995. Dorsal aorta cannulation: a method to monitor changes in blood levels of astaxanthin in voluntarily feeding Atlantic salmon, Salmo salarL. Aquaculture nutrition 1, 43-50. 143 Weber S. 1995. Determination of Xanthophylls, lutein and zeaxanthin in complete feeds and xanthophylls blends with H P L C . Hoffman-La, Roche, Ltd. pp 83 144 Appendix 2 A2 . Heterologous express ion of P450 hydroxy lases in E.coli (from Chapter 4) A2.1. Cloning strategy N'- terminus His-tag ColEl Sail Natl ATR1 pETDUET-1 (5,42 Obp) C-terminus Amp R 1 97A3 Stag V 97B3 Stag •HI Sg 97C1 Stag Figure A2.1 Cloning strategy for expression of three P450 genes (97A3, 97B3 and 97C1) and NADPH-cytochrome p450 reductase (ATR1). Each CYP97: :S- tag gene cassette was cloned at C'-terminal part of His-tag:ATR1, respectively. 145 A2.2. Transformation strategy ji -carotene Figure A2.2 Transformation strategy for co-expression of CYP: :ATR1 in E.coli strain C43 producing /3-carotene as a substrate in vivo. A m p R : Ampicillin resistance gene, C a p R : Chloramphenicol; Co lE, p15A: replicon 146 Table A2.1 The gene specific primers used to isolate the coding region of each gene from cDNA of Arabidopsis Hydroxylase Genes or Candidate 3 Arabidopsis gene designation Origin Primer sequence AtB1 At4g25700. F 1 5 ' - C A C C A T G G C G G C A G G A C T C T C A A C - 3 ' R 2 5 ' - T C A A G A A C T C G A A C T C G A C C C - 3 ' CYP97A3 At1g31800 F 5 ' - C A C C C A T G G C T A T G G C C I I ICCTCTT-3 ' R 5 ' - T T A A G A A A G A G C A G A T G A A A C - 3 ' CYP97B3 At4g15110 F 5 ' - C A C C A T G G T A G C A G C C A T G G C I I I -3 ' R 5 ' -TCAC I I I G A I C I I C I C I I IAG-3 ' CYP97C1 At3g53130 F 5 ' - C A C C A T G G A G T C T T C A C T C I I I ICT-3 ' R 5 ' -TTACC I I IG G C T C A C C T T C A T - 3 ' 1 F: forward, 2 R: reverse 3 Gene source: Arabidopsis thaliana 147 Table A2.2 Primers for pETDUET-ATR1 , pETDUET-97A3, 97B3 and 97C1 vector construction Construct Primer sequences pETDUET- ATR1 (Sail) F 1 5 ' - C T A C G G G T C G A C A T G A C T T C T G C I I I G T A T G C T - 3' 3 pETDUET-ATR (Notl) R 2 5 ' - C G A A G C A C T T G C G G C C G C T C A C C A G A C A T C T C T - 3 ' p E T D U E T - C Y P 9 7 A 3 (Bgl l l ) F 5 ' - A T C C T G G C T A G A T C T C A T G G C T A T G G C C T T T C C T C T - 3 ' p E T D U E T - C Y P 9 7 A 3 (Muni) R 5 ' - A C G T A C C T C A A T T G A G A G A A A G A G C A G A T G A A A C T T C - 3 ' p E T D U E T - C Y P 9 7 B 3 (Muni) F 5 ' - T C G T C G G A G G T C A A T T G T A T G G T A G C A G C C A T G G C T T - 3 ' p E T D U E T - C Y P 9 7 B 3 (Fsel) R 5 ' - G C A G T C A C G G C C G G C C C A C T T T G A T C T T C T C T T T A G - 3 ' pETDUET-CYP97C1 (Bglll) F 5 ' - G T C G C C T C A G A T C T A A T G G A G T C T T C A C T C I I I I CT-3 ' pETDUET-CYP97C1 (Sgfl) R 5 ' - A C T G A T G C G C G A T C G C C C T T T G G C T C A C C T T C A T A - 3 ' 1 F: forward, 2 R: reverse 3 Enzyme restricted site underlined and in bold 148 

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