Biochemical and molecular investigations of benzoic acid metabolism in calycinum cell cultures

Von der Fakultät für Lebenswissenschaften

der Technischen Universität Carolo-Wilhelmina zu Braunschweig

zur Erlangung des Grades

einer Doktorin der Naturwissenschaften

(Dr. rer. nat.)

genehmigte

D i s s e r t a t i o n

von Poonam Singh aus Bijpur/Uttar Pradesh / Indien

1. Referent: Professor Dr. Ludger Beerhues 2. Referent: apl. Professor Dr. Robert Hӓnsch 3. Referent: Privatdozent Dr. Thomas Vogt eingereicht am: 22.07.2019 mündliche Prüfung (Disputation) am: 25.11.2019

Druckjahr 2020

„Gedruckt mit Unterstützung des Deutschen Akademischen Austauschdienstes“

Vorveröffentlichungen der Dissertation

Teilergebnisse aus dieser Arbeit wurden mit Genehmigung der Fakultät für Lebenswissenschaften, vertreten durch den Mentor der Arbeit, in folgenden Beiträgen vorab veröffentlicht:

Publikationen Gaid, M., Singh, P., El-Awaad, I., Nagia, M. & Beerhues, L. Biotechnological production of prenylated xanthones for pharmaceutical use. Pharmaceutical Biocatalysis: Chemoenzymatic Synthesis of Active Pharmaceutical Ingredients (2019; ISBN 9789814800808). Singh, P., Gaid, M., Preu, L., Beurle, T., Hӓnsch, R. & Beerhues, L. Benzoate-CoA : A missing link towards xanthone biosynthesis in cell cultures (in preparation). Singh, P., Kaufholdt, D., Hänsch, R., Beerhues, L. & Gaid, M. Molecular cloning and characterization of the first cytosolic benzaldehyde dehydrogenase involved in the biosynthesis of xanthones in Hypericum (in preparation).

Tagungsbeiträge A. Vorträge Singh, P., Gaid, M., Preu, L., Beuerle, T. & Beerhues, L.: Benzoate-CoA ligase: From Gene Discovery to Homology modeling. (Oral Presentation). I Euroindoamerican Natural Products Meeting (I EIAMNP), Madrid, Spain (29.05.2018 - 01.06.2018). Singh, P., Gaid, M., Preu, L., Hӓnsch, R. & Beerhues, L.: Benzoate-CoA ligase: a missing link toward xanthone biogenesis. (Oral presentation). Section Natural of DBG, Section Meeting, Warberg, Germany (01.10.2018 - 03.10.2018). B. Posterbeiträge Singh, P., Gaid, M., Beuerle, T. & Beerhues, L.: Activation of benzoic acid: A key step of xanthone biosynthesis in Hypericum root cultures. (Flash poster oral presentation). Green for Good IV- Biotechnology of Products, Olomouc, Czech Republic (19.06.2017 - 22.06.2017). Singh, P., Gaid, M., Beuerle, T., Hӓnch, R. & Beerhues, L.: Activation of benzoic acid: A key step of xanthone biosynthesis in Hypericum root cultures. (Poster). Botanikertagung, Kiel, Germany (17.09.2017 - 21.09.2017). Singh, P., Gaid, M., Preu, L., Beuerle, T. & Beerhues, L.: Benzoate-activating CoA ligase from Hypericum calycinum cell cultures: From Gene discovery to Homology modeling. (Poster). Plant Biology Europe, Copenhagen, Denmark (18.06.2018 - 21.06.2018).

Acknowledgement

Praise to the almighty for showering me with endless blessings.

Firstly, I would like to express my immense gratitude towards my supervisor Professor Dr. Ludger Beerhues for welcoming me in his research group and providing me with all the required resources and an interesting research topic. I thank him for his never-ending supervision, discussions, encouragement and support during the entire PhD. I learned a lot working in the research group. Also, I appreciate him for letting me travel to and Sweden whenever needed which helped me in ensuring a healthy work-life balance. Next, I would like to thank Professor Dr. Robert Hӓnsch for agreeing to be my second supervisor and for his guidance during Laser Scanning Microscopy sessions. The useful discussion with him regarding subcellular localization has increased my knowledge and interest in this field. I also appreciate Professor Dr. Ute Wittstock for agreeing to be the chairperson of my thesis committee.

Next, I would take the opportunity to appreciate the endless support that Dr. Mariam Gaid has offered me during the entire PhD. The useful and prompt discussions that I had with her have helped me grow as a researcher. I appreciate her sharing ideas with me to make my work be a good one. I would also like to thank Dr. Lutz Preu for his support in the field of Computational Biology. His prompt replies and discussions have helped in the timely completion of this project. I am also thankful to Dr. Till Beuerle for the LC-MS and GC-MS analyses he carried out and for the beneficial discussions. Lastly, I would like to thank Dr. Eline Biederman for helping in RNA extraction and Dr. David Kaufholdt for providing Nicotiana when urgently needed.

I thank Dr. Rainer Lindigkeit for the technical support during the PhD period. I appreciate the efforts of Dr. Benye Liu, Ines Rahaus, Doris Glindemann and Bettina Böttner. They have promptly helped in managing laboratory and administrative work. I would like to acknowledge all my current and former colleagues in the Institute of Pharmaceutical Biology for providing me with a wonderful working atmosphere and being so friendly, kind and helpful. Special thanks to Anna, Eline, Elena, Nicola, Svenja, Christian and Philip for being my German language translators for the four years. You guys helped me a lot. I enjoyed all the time I spent with you during game nights, dinners and Profturntables. I will cherish all the memories we created together.

I am also grateful to the German Academic Exchange Service (DAAD) Germany and India for granting me a long-term scholarship to support my PhD work and my stay in Germany.

All that I have achieved till now in life and will ever achieve is dedicated to my family. I cannot thank them enough for the continuous support, love and the unwavering faith they have put in me.

Last, but not the least I would like to sincerely thank my husband, Vikrant, who always stood by my side through thick and thin and pushed me forward to achieve what I deserve. He is my pillar of strength.

Contents

Contents List of Figures ...... vi List of Tables ...... ix List of Abbreviations ...... xi 1 Introduction ...... 1 1.1 Biosynthesis of BAs through the shikimate/chorismate pathway ...... 3 1.1.1 3-Dehydroshikimate as a precursor ...... 3 1.1.2 Chorismate/Isochorismate as a precursor...... 3 1.2 BA biosynthesis from L-phenylalanine via trans-cinnamic acid ...... 4 1.3 Genus Hypericum ...... 6 1.4 Xanthones ...... 9 1.5 The aim of the work ...... 14 2 Materials ...... 15 2.1 Biological Materials...... 15 2.1.1 Plant material ...... 15 2.1.2 Bacterial strains for cloning and expression ...... 15 2.2 Vectors ...... 15 2.3 Primers ...... 16 2.4 ...... 18 2.4.1 Enzymes used for reverse transcription reaction ...... 18 2.4.2 Enzymes used for Polymerase Chain Reaction ...... 18 2.4.3 Enzymes used for Gateway cloning ...... 18 2.4.4 Restriction endonucleases ...... 18 2.4.5 Miscellaneous enzymes ...... 18 2.5 Kits ...... 19 2.6 Ladder ...... 19 2.7 Chemicals ...... 19 2.8 Culture media ...... 22 2.8.1 Plant tissue culture medium and required phytohormones ...... 22 2.8.2 Bacterial culture media and required supplements ...... 23 2.9 Buffers and solutions ...... 24 2.9.1 Buffers for plasmid isolation from E. coli ...... 24 2.9.2 Agrobacterium DNA extraction buffer ...... 24 2.9.3 Agrobacterium activation medium ...... 24

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Contents

2.9.4 Agarose gel electrophoresis ...... 24 2.9.5 SDS PAGE ...... 25

2.9.6 His6-tagged protein affinity purification ...... 26 2.9.7 Bradford solution for determination of protein concentration ...... 26 2.10 Online tools and databases ...... 26 2.11 Softwares ...... 27 2.12 Equipment ...... 28 2.12.1 General equipment ...... 28 2.12.2 Equipment for HPLC and MS analyses ...... 29 3 Methods ...... 31 3.1 Cultivation of in vitro cultures ...... 31 3.1.1 Propagation and elicitation of H. calycinum cell suspension cultures ...... 31 3.1.2 Cultivation of N. benthamiana potted plants ...... 31 3.2 Extraction of xanthones from H. calycinum cell suspension cultures ...... 31 3.3 Molecular biology methodologies ...... 32 3.3.1 Isolation of nucleic acids ...... 32 3.3.1.1 Isolation of total RNA from H. calycinum cells ...... 32 3.3.1.2 Isolation of plasmid DNA from E. coli cells ...... 32 3.3.1.3 Isolation of total DNA from A. tumefaciens ...... 33 3.3.1.4 Isolation of DNA from agarose gel and purification of PCR and restriction digestion products 33 3.3.2 Quantification of nucleic acids ...... 33 3.3.3 Reverse transcription for first-strand cDNA synthesis ...... 34 3.3.4 Primer design ...... 36 3.3.5 Polymerase chain reaction (PCR) ...... 37 3.3.5.1 Standard PCR ...... 38 3.3.5.2 Touchdown PCR ...... 38 3.3.5.3 High-fidelity PCR ...... 39 3.3.5.4 PCR for analyzing gene expression ...... 40 3.3.5.4.1 Semi-quantitative PCR ...... 40 3.3.5.4.2 Quantitative-PCR ...... 41 3.3.6 Agarose gel electrophoresis ...... 42 3.3.7 Procedures for DNA modification ...... 43 3.3.7.1 Restriction digestion ...... 43 3.3.7.2 Dephosphorylation of the digested vectors ...... 43 3.3.7.3 Ligation ...... 44

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Contents

3.3.8 Gateway cloning technology for cloning of localization constructs ...... 44 3.3.9 Sequencing ...... 47 3.4 Microbiological methodologies ...... 47 3.4.1 Determination of the microbial growth ...... 47 3.4.2 Preparation of competent cells ...... 47 3.4.2.1 Preparation of E. coli competent cells by calcium chloride method ...... 47 3.4.2.2 Preparation of A. tumefaciens electrocompetent cells ...... 47 3.4.3 Transformation of competent cells ...... 48 3.4.3.1 Transformation of E. coli competent cells by heat-shock ...... 48 3.4.3.2 Transformation of A. tumefaciens by electroporation ...... 48 3.4.4 Preparation of stock cultures ...... 49 3.5 Biochemical methodologies...... 49 3.5.1 Heterologous expression in E. coli ...... 49

3.5.2 Extraction and purification of the hexahistidine (His6)-tagged recombinant protein ... 49 3.5.3 Gel filtration chromatography ...... 50 3.5.4 Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) ...... 50 3.5.5 Determination of protein concentration using the method of Bradford (1976) ...... 51 3.5.6 ALDH assay ...... 52 3.5.7 Analysis of ALDH assay incubation ...... 52 3.5.8 CoA ligase assay ...... 53 3.5.8.1 Incubations to be analyzed by HPLC ...... 53 3.5.8.2 Incubations to be analyzed by spectrophotometer ...... 53 3.5.9 Biochemical characterization of HcBD and HcBZL ...... 53 3.5.10 Assay for optimization of luciferase protein amount ...... 55 3.5.11 Luciferase-based specificity assay for CoA ...... 55 3.5.12 Agroinfiltration and transient expression in N. benthamiana ...... 56 3.6 Analytical methods ...... 56 3.6.1 High-performance liquid chromatography ...... 56 3.6.1.1 Method used for the analysis of xanthone-rich extracts ...... 56 3.6.1.2 Method used for the analysis of product formation catalyzed by HcBD ...... 57 3.6.1.3 Method used for the analysis of product formation catalyzed by HcBZL ...... 57 3.6.1.4 Calibration curves ...... 57 3.6.2 Liquid chromatography-Mass spectrometry (LC-MS) ...... 58 3.6.2.1 Sample preparation ...... 58 3.6.2.2 Sample analysis ...... 58

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Contents

3.6.3 Gas chromatography-Mass spectrometry (GC-MS) ...... 59 3.6.4 Phylogenetic analyses ...... 59 3.6.5 Laser scanning microscopy ...... 60 4 Results ...... 61 4.1 Changes in xanthone content of H. calycinum cell cultures upon elicitor treatment ...... 61 4.2 H. calycinum benzaldehyde dehydrogenase (HcBD) ...... 63 4.2.1 Cloning a full-length HcBD cDNA ...... 63 4.2.2 Biochemical characterization of HcBD ...... 64 4.2.3 Phylogenetic analysis of HcBD ...... 73 4.3 H. calycinum benzoate-CoA ligase (HcBZL) ...... 74 4.3.1 Bioinformatic analysis of H. perforatum transcriptomes to garner putative BZL sequences ...... 74 4.3.2 Cloning of HcBZL ...... 76 4.3.3 Preliminary screening for the determination of the substrate specificities of HcBZL and HcAAE1 ...... 81 4.3.4 Luciferase-based determination of the substrate-specificity of HcBZL ...... 83

4.3.4.1 Cloning, heterologous expression and His6-tagged luciferase purification ...... 83 4.3.4.2 Optimization of the luciferase amount for the luciferase-based substrate-specificity assay 85 4.3.4.3 Substrate-specificity determination of HcBZL ...... 85 4.3.5 Biochemical characterization of HcBZL ...... 87 4.3.6 Phylogenetic reconstitution of HcBZL and HcAAE1 ...... 92 4.4 Elicitor-induced accumulation of HcBD and HcBZL transcripts as well as xanthones in H. calycinum cell cultures ...... 93 4.4.1 RT-PCR analysis of HcBD expression ...... 94 4.4.2 RT-qPCR analysis of HcBZL expression ...... 94 4.5 Subcellular localization of HcBD and HcBZL ...... 96 4.5.1 Preparation of HcBD and HcBZL YFP fusion constructs for subcellular localization 96 4.5.2 Subcellular localization of HcBD ...... 102 4.5.3 Subcellular localization of HcBZL ...... 103 5 Discussion ...... 108 5.1 Xanthones in Hypericum cell suspension cultures ...... 108 5.2 ALDH [aldehyde: NAD(P)+ ]/ HcBD ...... 109 5.3 Adenylate-forming enzymes ...... 117 5.3.1 Acyl/Aryl activating enzymes from H. calycinum cell cultures ...... 118

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Contents

5.3.2 Validation of HcBZL substrate-specificity through homology modelling and docking of substrates and intermediates...... 122 5.3.3 Dual localization of HcBZL ...... 126 5.4 In vivo participation of HcBD and HcBZL in the CoA-dependent non-β-oxidative route to synthesize xanthones in Hypericum ...... 128 5.5 Proposed subcelullar trafficking of benzoic acid from the CoA-dependent non-β-oxidative route to xanthone biosynthesis ...... 128 5.6 Perspectives ...... 130 6 Summary ...... 131 7 References ...... 133 8 Appendix ...... 156 A Figures ...... 156 B Tables ...... 166 C Alignment ...... 168

v

List of Figuresontents

List of Figures Fig. 1.1: Representatives of plant benzoic acids...... 1 Fig. 1.2: Overview of benzoic acid biosynthesis in plants...... 2 Fig. 1.3: Benzoic acid biosynthesis from trans-cinnamic acid in planta based on Abd El-Mawla and Beerhues (2002)...... 5 Fig. 1.4: Flowers of H. perforatum (a) and H. calycinum (b)...... 9 Fig. 1.5: Structure of the xanthone nucleus (a) and its various partially saturated di-, tetra- and hexahydroxanthone derivatives (b)...... 10 Fig. 1.6: Biosynthesis of xanthones in plants...... 12 Fig. 1.7: Modification of the primary 1,3,7-trihydroxyxanthone in Hypericum species...... 13 Fig. 3.1: Schematic diagram representing the Gateway Technology...... 46 Fig. 4.1: HPLC analysis of methanolic extracts from elicitor-treated H. calycinum cell cultures 24 h (a, red) and 0 h (a, blue) post-treatment...... 62 Fig. 4.2: Elicitor-induced changes in the total xanthone content of H. calycinum cells...... 62 Fig. 4.3: Schematic representation for ligating the NheI/KpnI digested HcBD ORF with the NheI/KpnI linearized pRSETB vector...... 64 Fig. 4.4: SDS-PAGE of affinity-purified recombinant HcBD protein...... 64 Fig. 4.5: Stacked HPLC-DAD chromatograms showing HcBD activities. R, reference compound; S, substrate (benzaldehyde or trans-cinnamaldehyde); P, enzymatically formed product...... 65 Fig. 4.6: UV absorption spectra of the enzymatically formed products which were identical to those of authentic acid references...... 66 Fig. 4.7: Substrate specificity of the recombinant HcBD...... 67 Fig. 4.8: GC-MS chromatograms for HcBD incubations with the substrate benzaldehyde...... 68 Fig. 4.9: GC-MS analysis of HcBD incubations with trans-cinnamaldehyde...... 69 Fig. 4.10: Effect of variation in incubation parameters on the activity of HcBD...... 70 Fig. 4.11: Effect of addition of supplements on HcBD activity in the standard assay...... 71 Fig. 4.12: Effect of storage conditions on the activity of HcBD...... 72

Fig. 4.13: Determination of Michaelis-Menten constants (Km) and maximum velocities (Vmax) of HcBD with various substrates...... 73 Fig. 4.14: Neighbor-joining tree presenting the phylogenetic relationships between HcBD and other members of the ALDH2 family...... 74 Fig. 4.15: Tissue-specific expression of various putative BZL transcripts ...... 76 Fig. 4.16: Agarose gel electrophoresis of a RNA sample extracted 8 h after yeast extract-treatment of H. calycinum cultured cells...... 77 Fig. 4.17: Agarose gel electrophoresis of HcBZL (a) and HcAAE1 (b) as proofread PCR products. 78

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List of Figuresontents

Fig. 4.18: Schematic representation for constructing the recombinant HcBZL expression plasmid. .. 79 Fig. 4.19: Schematic representation for constructing the recombinant HcAAE1 expression plasmid.79 Fig. 4.20: SDS-PAGE of the affinity-purified recombinant HcBZL protein...... 80 Fig. 4.21: SDS-PAGE of the affinity-purified recombinant HcAAE1 protein...... 80 Fig. 4.22: Substrate-utilization profiles of HcBZL and HcAAE1 as determined by HPLC-DAD analysis...... 81 Fig. 4.23: UV absorption spectra of HcBZL catalyzed thioester products. Benzoyl-CoA (a); aliphatic- CoA thioesters (b)...... 82 Fig. 4.24: ESI-MS/MS analysis of HcBZL catalyzed benzoyl-CoA formation, represented by the molecular ion peak [M-H]– at m/z 870.4...... 82 Fig. 4.25: HPLC analysis of HcBZL activity with various substrates...... 83 Fig. 4.26: Schematic representation of the subcloning of the luciferase ORF from pRS413-GAL- luc*(-SKL) into pRSETB expression vector...... 84 Fig. 4.27: SDS-PAGE of affinity-purified recombinant P. pyralis luciferase...... 84 Fig. 4.28: Optimization of purified recombinant luciferase for its application in the luciferase-based substrate-specificity assay...... 85 Fig. 4.29: Principle underlying the luciferase-based substrate specificity assay (Schneider et al., 2005)...... 85 Fig. 4.30: Luciferase-based substrate utilization profile of HcBZL...... 86 Fig. 4.31: Effect of the variation in various incubation parameters on the activity of HcBZL...... 87 Fig. 4.32: Effect of addition of supplements to the incubation of HcBZL...... 88 Fig. 4.33: Effect of storage conditions on the activity of HcBZL...... 89

Fig. 4.34: Determination of the Michaelis-Menten constants (Km) and maximum velocities (Vmax) of HcBZL with various substrates and co-substrates...... 91

Fig. 4.35: Determination of the Michaelis-Menten constants (Km) and maximum velocities (Vmax) of HcBZL with various substrates...... 92 Fig. 4.36: Neighbor-joining tree representing the phylogenetic relationships between HcBZL, HcAAE1 and other plant AAEs...... 93 Fig. 4.37: Elicitor-induced changes in HcBD expression. HcBD transcript levels were normalized with respect to H2A (•••) and 18s rRNA (---) transcripts...... 94 Fig. 4.38: Changes in the normalized relative expression of HcBZL in H. calycinum cell cultures upon treatment with yeast extract...... 96 Fig. 4.39: Agarose gel images indicating bands corresponding to attB PCR products of HcBD (a) and HcBZL (b)...... 97 Fig. 4.40: Schematic representation of construction of HcBD entry clones...... 98 Fig. 4.41: Schematic presentation of the construction of HcBZL entry clones...... 99 Fig. 4.42: Schematic representation of construction of HcBD expression clones...... 100

vii

List of Figuresontents

Fig. 4.43: Schematic representation of construction of HcBZL expression clones...... 101 Fig. 4.44: Subcellular localization of HcBD fused to YFP in N. benthamiana leaf epidermis cells..102 Fig. 4.45: Cytoplasmic localization of YFP-HcBZL upon transient expression of the construct in N. benthamiana leaf epidermis cell...... 103 Fig. 4.46: Co-transformation of N. benthamiana leaf epidermis cells with HcBZL-YFP and either the peroxisomal marker CFP-PTS1 (a-d) or the dually localized peroxisomal and cytoplasmic marker eqFP611 (e-h)...... 104 Fig. 4.47: Expression clones generated using three truncated HcBZL attB PCR products (b,c,d) as presented in Fig. 4.48a...... 105 Fig. 4.48: Subcellular localization of truncated HcBZL proteins fused to YFP...... 106 Fig. 4.49: Co-transformation of N. benthamina leaf epidermis cells with truncated HcBZL proteins fused to YFP and the CFP-PTS1 peroxisomal marker...... 107 Fig. 5.1: Multiple sequence alignment of selected ALDH2 family members including HcBD...... 111 Fig. 5.2: The mechanism of ALDH catalysis...... 113 Fig. 5.3: Two-step reaction mechanism of adenylate-forming enzymes...... 118 Fig. 5.4: Multiple sequence alignment depicting the conserved motifs BOX I and BOX II of the adenylate-forming family (yellow) and the 12 amino acid residues proposed to determine the substrate specificity of A. thaliana 4CL isoform 2 (At4CL2; annotated by asterisks; Schneider et al., 2003)...... 119 Fig. 5.5: Two-step reaction catalyzed by benzoate-CoA ligase (BZL)...... 121 Fig. 5.6: 3D model of HcBZL (grey) generated by using RpBZL (PDB ID: 4EAT_A, blue) as a template...... 123 Fig. 5.7: Docking models of the HcBZL SBP with benzoic acid (pink, a) and hexanoic acid (yellow, a) as well as the intermediates benzoyl-AMP (grey, b) and hexanoyl-AMP (green, c)...... 124 Fig. 5.8: Docking models of the HcBZL SBP docked with 3-hydroxybenzoyl-AMP (pink, a); 3,5- dihydroxybenzoyl-AMP (grey, a); 4-hydroxybenzoyl-AMP (grey, b); 3,4,5-trihydroxybenzoyl- AMP (grey, c); cinnamoyl-AMP (pink, d) and 4-coumaroyl-AMP (green, d) intermediates. ... 126 Fig. 5.9: Proposed biosynthetic pathway depicting the contributions of HcBD and HcBZL to xanthone formation proceeding via the CoA-dependent non-β-oxidative pathway route of benzoic acid biosynthesis...... 129

viii

List of Tablesontents

List of Tables Table 3.1: Standard components and reaction conditions of a reverse transcription reaction ...... 35 Table 3.2: Components of a standard PCR ...... 38 Table 3.3: Thermocycling parameters associated with a standard PCR ...... 38 Table 3.4: Thermocycling parameters of a Touchdown PCR ...... 39 Table 3.5: Standard components of Phusion Hot Start II and Q5 Hot Start high-fidelity PCR ...... 39 Table 3.6: Thermocycling parameters associated with a high-fidelity PCR ...... 40 Table 3.7: Thermocycling parameters used for semi-quantitative PCR ...... 41 Table 3.8: Components for RT-qPCR setup ...... 41 Table 3.9: Thermal cycling parameters of the RT-qPCR analysis ...... 42 Table 3.10: Components of a single and a double restriction digestion reaction ...... 43 Table 3.11: Components of a dephosphorylation reaction ...... 44 Table 3.12: Components of a standard ligation reaction ...... 44 Table 3.13: Components of a BP reaction ...... 45 Table 3.14: Components of a LR reaction ...... 46 Table 3.15: Set up for a Bradford assay ...... 51 Table 3.16: Components of an ALDH assay for HPLC and spectrophotometric analyses ...... 52 Table 3.17: Components of a CoA ligase assay for HPLC- and spectrophotometer-based analyses .. 53 Table 3.18: Various parameters tested for optimization of enzyme activity ...... 54 Table 3.19: Components of the luciferase-based substrate specificity assay...... 55 Table 3.20: HPLC-DAD methods and detection wavelengths used for the quantification of various enzymatically formed acids ...... 57 Table 4.1: Optimum conditions selected for kinetic characterization of HcBD ...... 72 Table 4.2: Steady-state kinetic parameters of HcBD...... 73 Table 4.3: Closest NCBI and MPGR hits of selected CoA ligases ...... 75 Table 4.4: Sequences selected from the Onekp database for amplification of putative HcBZL CDS 77 Table 4.5: Optimum incubation conditions for HcBZL activity with benzoic acid as substrate, used to determine Michaelis-Menten kinetics...... 89 Table 4.6: Steady-state kinetic parameters of HcBZL...... 90 Table 4.7: Slope of the standard curves, correlation coefficient (r2) and amplification efficiency percentage generated by the software used for each primer pair and probe set ...... 95 Table 4.8: Predicted localization of HcBD and HcBZL ...... 96

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List of Tablesontents

Table B 1: Accession numbers and names of functional ALDH2 family members used for the phylogenetic reconstruction of HcBD...... 166 Table B 2: Functional AAEs used for phylogenetic reconstruction of HcBZL and HcAAE1...... 167

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List of Abbreviationsontents

List of Abbreviations µg: Microgram µm: Micrometer μl: Microliter µM: Micromolar 2,4-D: 2,4-Dichlorophenoxy acetic acid 35S: Promoter from Cauliflower mosaic virus 3BZL: 3-Hydroxybenzoate-CoA ligase 4CL: 4-Coumarate-CoA ligase 4HBD: 4-Hydroxybenzaldehyde dehydrogenase 4HBS: 4-Hydroxybenzaldehyde synthase AAE: Acyl-activating enzymes AAO4: Aldehyde oxidase 4 ADCL: Aminodeoxychorismate ADCS: Aminodeoxychorismate synthase ADT: Arogenate dehydratase AIM1: Abnormal inflorescence meristem ALDH: Aldehyde dehydrogenases AMADH: Aminoaldehyde dehydrogenases AMP: Adenosine-5'-monophosphate APS: Ammonium persulphate ATP: Adenosine-5'-triphosphate att: Attachment site BA: Benzoic acid BA2H: Benzoate 2-hydroxylase BD: Benzaldehyde dehydrogenase BLAST: Basic Local Alignment Search Tool bp: Base pair BPS: Benzophenone synthase BSA: Bovine serum albumin BZL: Benzoate-CoA ligase C4H: Cinnamate 4-hydroxylase CCL: Carboxy CoA ligase CDS: Coding sequence cDNA: Complementary deoxyribonucleic acid CFP: Cyan fluorescent protein CHD: Cinnamoyl-CoA hydratase-dehydrogenase CHL: Cinnamoyl-CoA hydratase-lyase cLSM: Confocal laser scanning microscopy CM: Chorismate mutase CNL: Cinnamate-CoA ligase

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List of Abbreviationsontents

CoASH/CoA: cps: Counts per second CTAB: Cetyl trimethyl ammonium bromide cTP: Chloroplastic transit peptide CYP: Cytochrome P450 enzyme DAD: Diode array detector DEL: Deleted stop codon dH2O: Deionized water DHD-SDH: 3-Dehydroquinate dehydratase-shikimate dehydrogenase DIPS: Dearomatized isoprenylated phloroglucinols DMSO: Dimethy sulfoxide dNTP: Deoxyribonucleotide triphosphate DTT: 1,4-Dithiothreitol

EDTA.2H2O: Ethylenediamine tetraacetic acid disodium salt dihydrate Eco: Escherichia coli FAD: Flavin adenine dinucleotide FPKM: Fragments per kilobase of transcript per million reads mapped g: Gram GC-MS: Gas chromatography-mass spectrometry GSP: Gene-specific primer h: Hour Hc: Hypericum calycinum HCHL: 4-Hydroxycinnamoyl-CoA hydratase-lyase HPLC: High performance liquid chromatography Hs: ICS: Isochorismate synthase IPTG: Isopropyl β-D-1-thiogalactopyranoside IUPAC: International union of pure and applied chemistry KAT: 3-Ketoacyl-CoA kDa: Kilodalton

Km: Michaelis-Menten constant Kpn: Klebsiella pneumoniae l: Liter LB: Luria Bertani LS: Linsmaier and Skoog MES: 2-(N-Morpholino)ethanesulfonic acid mg: Milligram min: Minute ml: Milliliter mM: Millimolar M: Molar MCS: Multiple cloning site

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List of Abbreviationsontents

MPGR: Medicinal plant genome resource MSTFA: N-methyl-N-(trimethylsilyl)-trifluoroacetamide NAA: 1-Naphthaleneacetic acid NAD: β- adenine dinucleotide NADH: β-Nicotinamide Adenine dinucleotide reduced NADP: β-Nicotinamide Adenine dinucleotide phosphate NCBI: National Center for Biotechnology Information Nco: Nocardia corallina ng: Nanogram Nhe: Neisseria mucosa heidelbergensis nm: Nanometer NRPS: Nonribosomal peptide synthetases OD: Optical density ORF: Open reading frame PAL: Phenylalanine ammonia lyase PAGE: Polyacrylamide gel electrophoresis PCR: Polymerase chain reaction PDT: Prephenate dehydratase pI: Isoelectric point PPA-AT: Prephenate aminotransferase PPi: Inorganic pyrophosphate PPY-AT: Phenylpyruvate aminotransferase PT: Prenyltransferase PTS1: Type 1 Peroxisomal targeting signal PTS2: Type 2 Peroxisomal targeting signal RACE: Rapid amplification of cDNA ends REF: Reduced epidermal fluorescence RF: Restorer of fertility RT: Reverse transcription RNA: Ribonucleic acid rpm: Revolutions per minute s: Second S3H: Salicylate 3-hydroxylase SAS: Salicylaldehyde synthase SBP: Substrate binding pocket SD: Standard deviation SDH: Shikimate dehydrogenase SDS: Sodium dodecyl sulfate SOC: Super optimal broth with catabolite repression

Ta: Annealing temperature

Tm: Melting temperature TE: Thioesterase

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List of Abbreviationsontents

TEMED: N,N,N',N'-Tetramethyl ethylenediamine Tris: Tris(hydroxymethyl)-aminomethane TXS: Trihydroxyxanthone UTR: Untranslated region

Vmax: Maximum velocity of an enzyme-catalyzed reaction X6H: Xanthone 6-hydroxylase YEB: Yeast extract broth YFP: Yellow fluorescent protein Zeo: Zeocin β-ME: 2-Mercaptoethanol ˚C: Degree celcius

Amino acids Name 3 letter code 1 letter code Name 3 letter code 1 letter code Alanine Ala A Arginine Arg R Asparagine Asn N Aspartic acid Asp D Cysteine Cys C Glutamine Gln Q Glutamic acid Glu E Glycine Gly G Histidine His H Isoleucine Ile I Leucine Leu L Lysine Lys K Methionine Met M Phenylalanine Phe F Proline Pro P Serine Ser S threonine Thr T tryptophan Trp W tyrosine Tyr Y Valine Val V

Nucleotides Purines Pyrimidines Name Code Name Code Adenine A Thymine T Guanine G Cytosine C

xiv

Introduction

1 Introduction Benzoic acid (BA) is a small yet fascinating aromatic carboxylic acid (Fig. 1.1a) whose biosynthesis in plants has perplexed researchers for more than two decades. Benzoic acid and its C6-C1 substituted forms are the building blocks of various primary and secondary metabolites which are indispensable for survival and fitness of plants (Widhalm and Dudareva, 2015). Salicylic acid (2-hydroxybenzoic acid; Fig. 1.1b) acts as an endogenous signal for activation of defense responses in plants upon pathogen attack (Vlot et al., 2009). It is also known to regulate flowering and thermogenesis in plants (Cleland et al., 1974; Rhoads and McIntosh, 1992; Martínez et al., 2004). Methyl salicylate (Fig. 1.1c) acts as an airborne defense signal that activates defense-related genes in neighboring plants and healthy tissues of the infected plant (Shulaev et al., 1997). 4-Hydroxybenzoic acid (Fig. 1.1d) contributes to the biosynthesis of ubiquinone, a respiratory (Block et al., 2014). It is also known to be a precursor as well as a stimulator for the production of the antimicrobial compound shikonin (Inouye et al., 1979; Yazaki et al., 1997; Brigham et al., 1999). 4-Aminobenzoic acid (Fig. 1.1e) is a precursor for the formation of folic acid which acts as a cofactor in one-carbon transfer reactions (Basset et al., 2004a; 2004b). Gallic acid (Fig. 1.1f) is the precursor of gallotannins and ellagitannins (Dewick, 1997). The coenzyme A (CoASH/CoA) thioester of benzoic acid participates in the biosynthesis of defense compounds like xanthones and biphenyls in plants (Schmidt and Beerhues, 1997; Franklin et al., 2009; Chizzali et al., 2013).

Fig. 1.1: Representatives of plant benzoic acids. Benzoic acid (a); salicylic acid (b); methyl salicylate (c); 4-hydroxybenzoic acid (d); 4-aminobenzoic acid (e); gallic acid (f); anthranilic acid (g); 2,3-dihydroxybenzoic acid (h); 3-hydroxybenzoic acid (i). Based upon the current state of knowledge, plant BAs are formed either through intermediates of the shikimate/chorismate pathway or from L-phenylalanine via C2 side chain shortening of trans- cinnamic acid (Fig. 1.2). The latter pathway consists of three routes namely, CoA-dependent β- oxidative; CoA-dependent non-β-oxidative and CoA-independent non-β-oxidative (Fig. 1.3; Mustafa and Verpoorte, 2005; Wildermuth, 2006; Widhalm and Dudareva, 2015). Single or multiple routes can co-exist in the same species (Ishikura et al., 1984; Boatright et al., 2004; Shine et al., 2016).

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Fig. 1.2: Overview of benzoic acid biosynthesis in plants. ADCS, aminodeoxychorismate synthase; ADCL, aminodeoxychorismate lyase; ADT, arogenate dehydratase; AS, anthranilate synthase; BA2H, benzoate 2-hydroxylase; BD, benzaldehyde dehydrogenase; BS, benzaldehyde synthase; CHD, cinnamoyl-CoA hydratase-dehydrogenase; CHL, cinnamoyl-CoA hydratase- lyase; CM, chorismate mutase; CNL, cinnamate-CoA ligase; C4H, cinnamate 4-hydroxylase; DHD-SDH, 3-dehydroquinate dehydratase-shikimate dehydrogenase; 4HBS, 4- hydroxybenzaldehyde synthase; 4HBD, 4-hydroxybenzaldehyde dehydrogenase; ICS, isochorismate synthase; KAT, 3-ketoacyl-CoA thiolase; PAL, phenylalanine ammonia lyase; PDT, prephenate dehydratase; PPA-AT, prephenate aminotransferase; PPY-AT, phenylpyruvate aminotransferase; SAS, salicylaldehyde synthase; SDH, shikimate dehydrogenase; S3H, salicylate 3-hydroxylase; TE, thioesterase. Dashed arrows indicate steps studied at the biochemical level only (Mustafa and Veerapoorte, 2005; Wildermuth, 2006; Widhalm and Dudareva, 2015).

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Introduction

1.1 Biosynthesis of BAs through the shikimate/chorismate pathway In this kind of biosynthesis, the carboxyl group of shikimic/chorismic acid is retained in the product. Direct origin of BAs from shikimate/chorismate is common in bacteria (Serino et al., 1995; Gehring et al., 1997; Fox et al., 2008; Pfleger et al., 2008). However, there are limited reports for plants.

1.1.1 3-Dehydroshikimate as a precursor Tracer experiments were used to suggest that 3-dehydroshikimic acid is a precursor for gallic acid (3,4,5-trihydroxybenzoic acid) production in young leaves of Acer and Rhus species (Dewick and Haslam, 1969; Ishikura et al., 1984). Later, feeding of 13C-labeled glucose to young leaves of Rhus typhina and 13C nuclear magnetic resonance spectroscopy followed by comparison of isotopomer patterns in gallic acid and tyrosine revealed that an early intermediate of the shikimate pathway, likely 3-dehydroshikimic acid, is the precursor of gallic acid (Werner et al., 1997). Another independent experiment involving measurement of oxygen isotope abundance supported the aforementioned idea (Werner et al., 2004). The first report on the enzyme came when dehydroshikimate dehydrogenase catalyzing the conversion of 3- dehydroshikimate to gallic acid was partially purified from Betula pubescens leaves (Ossipov et al., 2003). Recently, cDNAs encoding shikimate dehydrogenases have been cloned from Juglans regia and Vitis vinifera. Apart from performing the classical activity, they were also capable of converting 3-dehydroshikimic acid to gallic acid, hence contributing to gallic acid metabolism (Muir et al., 2011; Bontpart et al., 2016).

1.1.2 Chorismate/Isochorismate as a precursor In plants, at least four different enzymes that utilize chorismate and subsequently lead to production of BAs are known. Biosynthesis of anthranilic acid (2-aminobenzoic acid; Fig 1g) from chorismate is catalyzed by anthranilate synthase (AS; Niyogi and Fink, 1992). Anthranilate is a precursor for tryptophan biosynthesis and it also serves as a precursor for avenanthramide phytoalexin biosynthesis as determined by feeding of radiolabelled substrates in oat leaves (Ishihara et al., 1999). Aminodeoxychorismate synthase (ADCS) is another enzyme catalyzing the conversion of chorismate to aminodeoxychorismate which then undergoes pyruvyl side chain removal by aminodeoxychorismate lyase (ADCL) leading to the formation of 4-aminobenzoic acid which serves as a precursor for biosynthesis of the benzenoid ring of folic acid (Basset et al., 2004a; Basset et al., 2004b). The next enzyme using chorismate is isochorismate synthase (ICS) which catalyzes the formation of the isomeric compound, isochorismate. The first plant ICS cDNA was cloned from elicitor-treated cell cultures of Catharanthus roseus (van Tegelen et al., 1999). Isochorismate acts as a precursor for the biosynthesis of salicylic acid (Wildermuth et al., 2001), 2,3-dihydroxybenzoic acid (Fig. 1.1h; Moreno et al., 1994; Muljono et al., 2002) and ortho-succinylbenzoic acid which gets incorporated in phylloquinone ( K1; Gross et al., 2006). The fourth chorismate-accepting enzyme is chorismate mutase, which catalyzes prephenate formation and marks the first committed step in phenylalanine and tyrosine biosynthesis (Eberhard et al., 1993).

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Some other reports have also mentioned the formation of BAs from the shikimate/chorismate pathway. Based upon feeding of radiolabelled substrates, the analysis of radiolabelled xanthones and the basal level of phenylalanine ammonia lyase (PAL) activity in elicited-treated cell cultures of Centaurium erythraea it was suggested that 3-hydroxybenzoic acid (Fig. 1.1i) is formed directly from the shikimate pathway rather than from trans-cinnamic acid (Abd El- Mawla et al., 2001). During the same period Swertia chirata roots were also subjected to stable isotope feeding experiments and labeling patterns of amarogentin and 1,3,5,8- tetrahydroxyxanthone indicated that 3-hydroxybenzoic acid, the precursor of the aforementioned metabolites, originated from an intermediate of the shikimate pathway rather than from L- phenylalanine via trans-cinnamic acid or benzoic acid (Wang et al., 2001; 2003). Interestingly, the two systems were derived from the members of the Gentianaceae family.

1.2 BA biosynthesis from L-phenylalanine via trans-cinnamic acid L-Phenylalanine biosynthesis in plants occurs through the plastidial arogenate pathway (Graindorge et al., 2010; Maeda et al., 2010; Maeda et al., 2011) and the recently discovered microbial-like cytoplasmic phenylpyruvate pathway (Yoo et al., 2013; Qian et al., 2019). L- phenylalanine formed is converted to trans-cinnamic acid through a non-oxidative deamination reaction catalyzed by phenylalanine ammonia lyase (PAL; EC 4.3.1.5). This reaction marks the entry step that directs the carbon flux from the primary metabolism into the phenylpropanoid secondary metabolism (Vogt, 2010; Fraser and Chapple, 2011). trans-Cinnamic acid is converted to benzoic acid by various routes via C2 side chain shortening. Contrary to the preservation of the precursor carboxyl group in benzoic acid originating from an intermediate of the shikimate/chorismate pathway, the precursor carboxyl group in this case is not preserved in the benzoic acid product and originates from the β-carbon of the trans-cinnamic acid side chain. Multiple pathways have been suggested for the shortening of the propyl side chain of trans- cinnamic acid and its substituted forms.

Early evidence of the CoA-dependent β-oxidative biosynthetic route (Fig. 1.3, top) of plant benzoic and salicylic acids came from a stable isotope-labeled 3-hydroxy-3-phenylpropanoic acid feeding experiment by Jarvis et al. (2000) in Cucumis sativa and Nicotiana attenuata. The experiments indicated the incorporation of the labeled carbon into benzoic and salicylic acids but not in benzaldehyde. The CoA-dependent β-oxidative route was also reported to be functional in Lithospermum erythrorhizon cell cultures where 4-hydroxybenzoyl-CoA was formed from 4- coumaroyl-CoA followed by its conversion to 4-hydroxybenzoic acid by the activity of thioesterases (Loscher and Heide, 1994). Since then all the molecular players for this route have been identified and functionally characterized in Petunia hybrida. The pathway begins with a peroxisomal cinnamate-CoA ligase (CNL) that activates trans-cinnamic acid to cinnamoyl-CoA (Colquhoun et al., 2012; Klempien et al., 2012; Lee et al., 2012). A similar reaction is catalyzed by Arabidopsis thaliana peroxisomal CoA ligase At4g19010 for initiating the β-oxidative side chain shortening of 4-coumaric acid to 4-hydroxybenzoic acid (Block et al., 2014). In P. hybrida, cinnamoyl-CoA is then accepted by the bi-functional peroxisomal enzyme cinnamoyl-CoA

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Introduction hydratase-dehydrogenase (CHD; Qualley et al., 2012) to form 3-oxo-3-phenylpropanoyl-CoA that is converted to benzoyl-CoA by the peroxisomal 3-ketoacyl-CoA thiolase 1 (KAT1; Van Moerkercke et al., 2009). Recently, the presence of peroxisomal thioesterase (TE) which hydrolyzes benzoyl-CoA to benzoic acid has been reported for P. hybrida (Qualley et al., 2012; Adebesin et al., 2018). Through the analysis of β-oxidation mutants of A. thaliana with no or less accumulation of benzoylated glucosinolates, Abnormal Inflorescence Meristem (AIM1), a P. hybrida CHD homolog, and KAT2, a P. hybrida KAT1 homolog, were identified as potential players that are involved in the production of benzoic acid (Bussell et al., 2014).

Fig. 1.3: Benzoic acid biosynthesis from trans-cinnamic acid in planta based on Abd El-Mawla and Beerhues (2002). BD, benzaldehyde dehydrogenase; BS, benzaldehyde synthase; BZL, benzoate-CoA ligase; CHD, cinnamoyl-CoA hydratase-dehydrogenase; CHL, cinnamoyl-CoA hydratase-lyase; CNL, cinnamate-CoA ligase; KAT, 3-ketoacyl-CoA thiolase. Dashed arrows indicate steps studied at the biochemical level. In the CoA-dependent non-β-oxidative route (Fig. 1.3, intersecting pathway), cinnamoyl-CoA production by CNL is followed by its hydration and C2 side chain cleavage to generate benzaldehyde. Based on the detected activity in Hypericum cell cultures, the reaction was proposed to be catalyzed by a bi-functional cinnamoyl-CoA hydratase-lyase (CHL; Abd El-

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Mawla and Beerhues, 2002). The similar reaction mechanism was first observed in case of bacterial 4-hydroxycinnamoyl-CoA hydratase-lyase (HCHL) which accepted various hydroxylated cinnamoyl-CoA derivatives to produce their corresponding aldehydes but not cinnamoyl-CoA (Mitra et al., 1999). Chy1, a HCHL homolog from A. thaliana encoded a protein with in vitro activity (Ibdah and Pichersky, 2009). The next step involves the oxidation of benzaldehyde to benzoic acid by the enzyme benzaldehyde dehydrogenase (BD). Crude preparations of H. androsaemum, Sorbus aucuparia and Pyrus pyrifolia cells exhibited BD activity (Abd El-Mawla and Beerhues, 2002; Gaid et al., 2009; Saini et al., 2017). 4- Hydroxybenzaldehyde dehydrogenase (4HBD) activity was also detected in crude preparations of Daucus carota and L. erythrorhizon in vitro cultures (Yazaki et al., 1991; Schnitzler et al., 1992). A cDNA encoding a mitochondrial BD was cloned from Antirrhinum majus flowers which contribute to methyl benzoate production (Long et al., 2009). Multiple cytoplasmic and mitochondrial aldehyde dehydrogenases (ALDHs) from Zea mays are capable of oxidizing benzaldehyde to benzoic acid (Liu and Schnable, 2002; Končitíková et al., 2015). Benzaldehyde to benzoic acid conversion is also catalyzed by A. thaliana aldehyde oxidase 4 (AAO4) which contributes to the pool of benzoylated glucosinolates as determined by studying the aao4 null mutants (Ibdah et al., 2009). The last step of the route is the activation of benzoic acid to benzoyl-CoA which is catalyzed by benzoate-CoA ligase (BZL). BZL activity was previously reported in crude preparations of H. androsaemum cell cultures (Abd El-Mawla and Beerhues, 2002), Clarkia breweri flowers (Beuerle and Pichersky, 2002b) and P. pyrifolia yeast extract- treated cell cultures (Saini et al., 2019). A 3-hydroxybenzoate-CoA ligase (3BZL) activity was also reported in the crude extracts of C. erythraea (Barillas and Beerhues, 1997; 2000).

The third route which is the CoA-independent non-β-oxidative route (Fig. 1.3, bottom) begins with the conversion of trans-cinnamic acid to benzaldehyde through C2 side chain cleavage by benzaldehyde synthase (BS). BS activity has recently been reported in crude preparations of elicitor-treated P. pyrifolia cells in which it contributes to biphenyl production (Saini et al., 2019). Previously, 4-hydroxybenzaldehyde synthase (4HBS) activity was detected in methyl- jasmonate-treated hairy roots of D. carota and in Vanilla planifolia (Podstolski et al., 2002, Sircar and Mitra, 2008). Also, salicylaldehyde synthase (SAS) activity was detected in crude preparations of N. tabacum and Malus domestica converting 2-hydroxycinnamic acid into salicylaldehyde (Malinowski et al., 2007; Sarkate et al., 2018). A similar mechanism was reported for V. planifolia where vanillin synthase converted ferulic acid into vanillin (Gallage et al., 2014). Once benzaldehyde is formed an enzymatic cascade of BD and BZL leads to benzoyl- CoA production. The CoA-independent non-β-oxidative route was previously also detected in Solanum tuberosum (French et al., 1976), L. erythrorhizon (Yazaki et al., 1991), D. carota (Schnitzler et al., 1992; Sircar and Mitra, 2008) and V. planifolia (Podstolski et al., 2002).

1.3 Genus Hypericum Since the current work focused on the biosynthetic pathway present in Hypericum species this plant genus is introduced in more detail. The flowering plants within the family

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Introduction consist of nine genera, namely Cratoxylum, Eliea, Harungana, Hypericum, Lianthus, Santomasia, Thornea, Triadenum, and Vismia. The genus Hypericum contributes to approximately 80% of the diversity of the family and is represented by herbs, and seldom trees. The genus has been classified into 36 taxonomic sections. Hypericum species inhabit temperate regions and higher altitudes of the tropics. Eurasia and Andean South America are the major centers of species richness along with minor centers of diversity being , Southeast Asia and Africa (Crockett and Robson, 2011; Nürk and Crockett, 2011). Approximately 590 Hypericum species have been reported until 2016, which constitute ~0.2% of the total angiosperm species recorded (Christenhusz and Byng, 2016).

The term Hypericum is a derivative of the Greek words hyper and eikon which translates to ‘above’ and ‘icon/image’, respectively. In ancient period the blossoms of the plant were placed above holy figures in households to repel ‘evil spirits’ which in the current period is synonymous to Hypericum extracts being used as a potent remedy for the treatment of depressive ailments (Miller, 1998). The use of Hypericum species in traditional medicine is based upon the presence of bioactive specialized/secondary metabolites that belong to various chemically distinct classes. The major classes of compounds found include hypeforins, hypericins, flavonoids and xanthones (Beerhues, 2011). Additionally, benzophenone derivatives (Baggett et al., 2005), biphenyl derivatives (Bréard et al., 2018), essential oils/volatile compounds (Bertoli et al., 2011) and phenylpropanes (Kwiecień et al., 2015) are also present in Hypericum extract.

Hypericum species contain three different kinds of secretory structures, namely dark glands or black nodules, translucent glands or pale glands and secretory canals (Ciccarelli et al., 2001a, 2001b). Not all of these structures are found in all the species of the genus (Crockett and Robson, 2011). Secretory structures are the site of synthesis and/or accumulation of different secondary metabolites. The black nodules are the sites for accumulation of naphthodianthrones (Zobayed et al., 2006; Kucharíková et al., 2016). The amount of naphthodianthrones correlates positively with the size and the number of dark glands (Zobayed et al., 2006). Acylphloroglucinols like hyperforin and adhyperforin accumulate in the translucent glands (Soelberg et al., 2007; Kucharíková et al., 2016). Essential oils are present in translucent glands and secretory canals (Ciccarelli et al., 2001b).

A representative member of the genus Hypericum is H. perforatum also known as St. John’s wort. The common name of the plant is derived from the fact that it flowers around the St. John’s day which is the 24th of June. It is a perennial flowering herb native to Europe but has spread to temperate locations in north-western Africa, Asia, , and North and South America. The height of the herb typically ranges from 40 to 80 cm. The stems and branches are covered with oblong, smooth-margined leaves of a length ranging from 1 to 3 cm (Klemow et al., 2011). The spheroidal translucent glands in the leaves give it the perforated look (Ciccarelli et al., 2001b). The yellow flowers (1-3 cm in diameter; Fig. 1.4a) are borne in small clusters on the tip of the branches and consist of five petals with black dots on the edges. The flower contains numerous prominent (Klemow et al., 2011; https://keyserver.lucidcentral.org/weeds/

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Introduction data/media/Html/hypericum_perforatum.htm). It is a thoroughly investigated species whose extracts are used for the treatment of mild to moderate depression (Apaydin et al., 2016), and for wound healing (Süntar et al., 2011). Pharmacological studies also revealed the potential use of the extracts as antioxidant (Zou et al., 2004), antiviral (Axarlis et al., 1998) and antibacterial agents (Süntar et al., 2016). Another species whose antidepressant effect was suggested to be as potent as that of H. perforatum and can be possibly used for the treatment of depression is H. calycinum (Oztürk et al., 1996). H. calycinum (also known as Aaron's beard or creeping St. John's wort) is a deciduous , native to southeastern Europe, distributed naturally in the forested region of eastern Turkey and southeastern Bulgaria. This species is also cultivated and is naturalized in many other countries around the world (Nürk and Crockett, 2011). It is usually planted as ground cover, however, is considered as an environmental weed in Australia as it spreads extensively via creeping stems and competes with the native ground flora for nutrients and space. The rose-like ornamental yellow flowers (5-7 cm in diameter) of the plant contain five petals and bushy with reddish anthers (Fig. 1.4b). They appear singly or in groups of 2-3. The leaves are oval to oblong and can reach up to a size of 10 cm (https://keyserver. lucidcentral.org/weeds/data/media/Html/hypericum_calycinum.htm; http://www. missouribotanicalgarden.org/PlantFinder/PlantFinderDetails.aspx?kempercode=a520). A few studies have been conducted inspecting the secondary metabolites produced by H. calycinum and a detailed report was presented by Cirak et al. (2016). The plant contains traces of hyperforin and adhyperfoin (0.01 mg/g dry weight) in the flowers. Chlorogenic, neochlorogenic, 2,4- dihydroxybenzoic acids, hyperoside, isoquercitrin, quercitrin, quercetin, avicularin and rutin are present in flowers, leaves and stem with the highest amount being found in leaves. Similarly, I3- II8-biapigenin and (+)-catechin were also present in all three tissues with their amounts being highest in the flowers. (-)-Epicatechin was present equally in flowers and leaves. Hypericin, pseudohypericin and caffeic acid were not detected in any of the tested tissues. Previously, trace amounts of hyperforin (~0.03 mg/g dry weight) and adhyperforin (~0.3 mg/g dry weight) were also detected in cultured cells of H. calycinum (Klingauf et al., 2005). Along with the flavonoids, the yellow flowers of H. calycinum also contain dearomatized isoprenylated phloroglucinols (DIPS) which attract pollinators. DIPS (e.g. hypercalin A) are present in high concentrations in the anthers and the ovarian wall of the flowers and are defense compounds that discourage herbivory (Gronquist et al., 2001). The major essential oils found in the aerial part of the plant varied depending upon the location from which the plant material was collected. While α- terpineol and β-pinene were the major components in Asian H. calycinum, α-humulene and germacrene D were the major components of Italian H. calycinum (Demirci et al., 2005; Maggi et al., 2010). Recently, prenylated xanthones have been identified in elicitor-treated cultured cells of H. calycinum (Gaid et al., 2012; Nagia et al., 2019). Previously, prenylated flavanones were extracted from the stem of the plant (Win et al., 2012). Extracts of the plants show cytotoxic (Decosterd et al., 1989; Win et al., 2012), antioxidant (Kirmizibekmez et al., 2009; Win et al., 2012), antimicrobial (Maggi et al., 2010; Nogueira et al., 2013), fungicidal and antimalarial activities (Decosterd et al., 1991).

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(a) (b)

Fig. 1.4: Flowers of H. perforatum (a) and H. calycinum (b). In the current work, H. calycinum cell suspension cultures were used for studying the biosynthesis of benzoic acid which enters the metabolism of xanthones. For this reason, this class of constituents is presented in more detail.

1.4 Xanthones Xanthones (IUPAC name 9H-xanthen-9-one) are polyphenolic secondary metabolites found in fungi, lichens and a few higher plant families. The name “xanthone” has been derived from the Greek word “xanthos” which stands for “yellow” (Masters and Brӓse, 2012). A xanthone consists of two benzene rings connected by a carbonyl group and an ether bridge giving rise to a planar dibenzo-γ-pyron skeleton (Fig. 1.5a). The symmetrical nature of the xanthone scaffold combined with the mixed biogenetic origin of its carbons in higher plants made it necessary to number the carbons according to a biosynthetic convention. Carbons 1-4 and 5-8 were allocated to the acetate-derived ring A and the shikimate-derived ring B, respectively (Bennett and Lee, 1989). When only ring B is oxygenated the lowest numbers are assigned to its carbons according to IUPAC nomenclature rules. However, when discussing the biosynthetic origin of xanthones, the above-mentioned numbering and ring designations should be used (Bennett and Lee, 1989). In the fully aromatized xanthone nucleus, one of the aromatic rings can be partially or completely saturated giving rise to di-, tetra- or hexahydroxanthone derivatives (Fig. 1.5b; Maters and Brӓse, 2012).

Xanthones are “privileged structures” as they are capable of interacting with various target biomolecules, thereby producing various desirable biological effects (Masters and Brӓse, 2012). This class of compounds exhibits a broad spectrum of biological activities (El-Seedi et al., 2010). In general, xanthones have been classified into six major groups, namely simple oxygenated xanthones, xanthone glycosides, prenylated and related xanthones, xanthonolignoids, bis- xanthones and miscellaneous xanthones. Depending upon the degree of oxygenation, simple oxygenated xanthones are further subdivided into non-, mono-, di-, tri-, tetra-, penta-, and hexaoxygenated xanthones (Peres and Nagem, 1997; Vieira and Kijjoa, 2005; Negi et al., 2013).

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Fig. 1.5: Structure of the xanthone nucleus (a) and its various partially saturated di-, tetra- and hexahydroxanthone derivatives (b).

As per a recent report, 168 species of plants belonging to 58 genera of 24 families have been reported to contain xanthones. The families Clusiaceae, Gentianaceae, and Hypericaceae are the major representatives of xanthone producers among plants (Ruan et al., 2017). As of January 2016, the Dictionary of Natural Products indicated the existence of 1940 natural xanthones including the reduced derivatives. The source of ~80% of the xanthones is plants. The remaining 20% is constituted majorly by fungi (~15%) and lichens (~5%; Le Pogam and Boustie, 2016).

Different biosynthetic pathways exist for the formation of xanthones in fungi, lichens and plants. In the former two taxa, the xanthone scaffold is derived solely from acetate while in plants xanthones originate from the acetate-shikimate pathway. Thus, both the shikimate and the malonate routes contribute to the biosynthesis of the plant xanthone scaffold. Formation of the xanthone core requires starter and extender molecules. In some plant families the preferred starter molecule is benzoyl-CoA, which is generated from L-phenylalanine via trans-cinnamic acid, while in other families 3-hydroxybenzoyl-CoA is the preferred starter substrate which is derived from an intermediate of the shikimate pathway (Abd El-Mawla et al., 2001). In Hypericum, trans-cinnamic acid gets converted to benzoyl-CoA through the CoA-dependent non-β-oxidative route mentioned earlier (Abd El-Mawla and Beerhues, 2002; Gaid et al., 2012). Benzoyl-CoA is the preferred starter substrate in Hypericum species and Garcinia mangostana while 3-hydroxybenzoyl-CoA is the preferred starter substrate in C. erythraea. Subsequently, benzophenone synthase, a type III polyketide synthase, catalyzes the iterative decarboxylative condensation of benzoyl- and 3-hydroxybenzoyl-CoA with three molecules of malonyl-CoA to generate a linear tetraketide which undergoes cyclization to produce 2,4,6- trihydroxybenzophenone and 2,3',4,6-tetrahydroxybenzophenone, respectively (Beerhues, 1996; Schmidt and Beerhues, 1997; Liu et al., 2003; Huang et al., 2012; Nualkaew et al., 2012; Tocci

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Introduction et al., 2018). In Hypericum species the recently cloned CYP81AA1 and CYP81AA2 are bifunctional enzymes that catalyze (i) the hydroxylation of 2,4,6-trihydroxybenzophenone to 2,3',4,6-tetrahydroxybenzophenone and (ii) the subsequent regioselective C-O phenol coupling reactions para and ortho to the 3'-hydroxy group, giving rise to 1,3,7-trihydroxyxanthone and 1,3,5-trihydroxyxanthone, respectively (Fig. 1.6; Peters et al., 1998; Schmidt and Beerhues, 1997; El-Awaad et al., 2016).

The isomeric 1,3,7- and 1,3,5-trihydroxyxanthones are considered to be the precursors of all plant xanthones (Bennett and Lee, 1989; Peters et al., 1998). They can be further hydroxylated, glycosylated or prenylated to form an array of xanthones. Simple, oxygenated xanthones are found in both Hypericaceae and Gentianaceae families and are comparatively more oxygenated in the Gentianaceae. Also, O-glycosylxanthones are common in the Gentianaceae whereas Hypericaceae are rich source of prenylated xanthones which are not reported in the Gentianaceae (Bennett and Lee, 1989; Jensen and Schripsema, 2002). 1,3,7- and 1,3,5-Trihydroxyxanthones are hydroxylated at C-6 by a P450 monooxygenase, xanthone 6-hydroxylase (X6H) to yield 1,3,6,7- and 1,3,5,6-tetrahydroxyxanthones, respectively (El-Awaad et al., unpublished). In C. erythraea (Gentianaceae), X6H exclusively hydroxylated 1,3,5-trihydroxyxanthone while in H. androsaemum 1,3,7-trihydroxyxanthone was preferentially hydroxylated. Hydroxylation at C-6 is the first xanthone-modifying step in both cell cultures (Schmidt et al., 2000b). Further stepwise hydroxylations in Centaurium species give rise to tetra-, penta- and hexaoxygenated xanthones which later get methoxylated and O-glycosylated (Beerhues and Berger, 1995)

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Fig. 1.6: Biosynthesis of xanthones in plants. BPS, benzophenone synthase; BZL, benzoate-CoA ligase; 3BZL, 3-hydroxybenzoate-CoA ligase; CoASH/CoA, coenzyme A; CYP81AA1 and CYP81AA2, trihydroxyxanthone synthases. Dashed arrows (black) indicate reactions that have been reported at the biochemical level only (Adapted from Gaid et al., 2019). Prenylation of xanthones has been elucidated in Hypericum species. The first plant xanthone prenyltransferase referred to as HcPT was cloned from cell suspension cultures of H. calycinum and it catalyzed the regiospecific C-8 prenylation of 1,3,6,7-tetrahydroxyxanthone to produce 1,3,6,7-tetrahydroxy-8-prenylxanthone. In hyperxanthone E formation, 8-prenylation is followed by intramolecular cyclization leading to the formation of the pyran ring (Fig. 1.7; Fiesel et al., 2015). Recently, new C-8 prenyltransferases have been cloned from H. calycinum and H. sampsonii referred to as Hc/HsPT8px and Hc/HsPTpat which sequentially add prenyl groups at C-8 of 1,3,6,7-tetrahydroxyxanthone and 1,3,6,7-tetrahydroxy-8-prenylxanthone, respectively, and thus participate in the biosynthetic pathway of the gem-diprenylated xanthone, patulone (Fig. 1.7; Nagia et al., 2019).

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Fig. 1.7: Modification of the primary 1,3,7-trihydroxyxanthone in Hypericum species. DMAPP, dimethylallyl pyrophosphate, Hc, H. calycinum; Hs, H. sampsonii; PT, prenyltrasferase; X6H, xanthone 6-hydroxylase (El-Awaad et al., unpublished). The dashed arrow indicates a step proposed by Nagia et al. (2019).

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Introduction

1.5 The aim of the work Multiple genes contributing to upstream and downstream steps of the xanthone biosynthetic pathway have been cloned from H. calycinum cell suspension cultures. With the xanthone biosynthetic route being completely elucidated at the molecular level it became important to understand how the biosynthetic precursor, i.e. benzoyl-CoA, is formed. Biosynthesis of benzoyl-CoA has been completely established at the biochemical level in H. androsaemum cell suspension cultures. Of the four enzymes involved only CNL has so far been studied at the gene level.

The objectives of the current work were to carry out cDNA cloning, functional characterization and subcellular localization of benzaldehyde dehydrogenase (BD) and benzoate-CoA ligase (BZL) from elicitor-treated H. calycinum cell suspension cultures. The work involved the following steps:

• Investigation of the xanthone profile of H. calycinum cell suspension cultures before and after yeast extract treatment • Making use of the sequence information about functionally characterized CoA ligases to carry out a blast and to fetch putative sequences from publicly available H. perforatum transcriptomes • Cloning of putative BZL ORFs from yeast extract-treated H. calycinum cell cultures and obtaining their 5'- and 3'-UTRs by using a combination of gene-specific and UTR- specific primers and rapid amplification of cDNA ends (RACE), respectively, with a previously cloned putative BD being included for subsequent analysis • Expression analysis of the retrieved genes using quantitative and semi-quantitative PCR techniques • Heterologous expression of HcBD and HcBZL coding sequences in E. coli BL21, followed by crude protein extraction and His6-tagged protein purification • Testing the activity of the purified recombinant proteins with various substrates • Characterization of the active recombinant proteins with respect to kinetic properties • Construction of various full-length and truncated expression clones of HcBD and HcBZL using the Gateway cloning technology • Determination of the subcellular localization of HcBD and HcBZL by transient expression of their YFP fusions in leaves of N. benthamiana.

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2 Materials

2.1 Biological Materials

2.1.1 Plant material The plant material used in the current study included H. calycinum cell suspension cultures and N. benthamiana potted plants propagated at the Institute of Pharmaceutical Biology (IPB), Technische Universität Braunschweig (TU-BS, Germany). H. calycinum cell suspension cultures were used for metabolite profiling and cloning and expression analysis of HcBD and HcBZL. N. benthamiana potted plants were used for studying the subcellular localization of HcBD and HcBZL in the plant cell.

2.1.2 Bacterial strains for cloning and expression Escherichia coli Strain name Genotype - - + DH5α F Φ80lacZΔM15 Δ(lacZYA-argF) U169 recA1 endA1 hsdR17 (rk , mk ) phoA supE44 thi-1 gyrA96 relA1 λ- - - - DB3.1 F gyrA462 endA1 ∆(sr1-recA) mcrB mrr hsdS20(rB , mB ) supE44 ara14 galK2 lacY1 proA2 rpsL20(Smr) xyl5 ∆leu mtl1 - - - r BL21(DE3)pLysS F ompT hsdSB (rB , mB ) gal dcm λ(DE3) pLysS (Cam ) Agrobacterium tumefaciens Strain name Genotype C58C1 C58C1(Rifr), pMP90 (Gentr)

2.2 Vectors Name Purpose Source pJET1.2 Positive selection cloning vector conferring Thermo Scientific ampicillin resistance. The vector contains a lethal gene which is disrupted by the ligation of the gene of interest into the cloning site. Thus, only cells containing the recombinant plasmid are able to propagate. pRSET B Expression vector with N-terminal polyhistidine Invitrogen (6xHis) tag pRS413•GAL1•luc Source of Photinus pyralis luciferase ORF Dr. Michael Benton *(•SKL) Addgene pDONR/Zeo Gateway donor vector Invitrogen pEarley-104 Gateway destination vector, N-terminal YFP construct production pEarley-101 Gateway destination vector, C-terminal YFP construct production EqFP611 Cytoplasmic and peroxisomal marker construct Prof. Robert (Forner and Binder, 2007) Hӓnsch’s Group,

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Helper plasmid Reduces the effect of post-transcriptional gene- Institute of Plant p19 silencing (Voinnet et al., 2003) Biology, TU CFP-SNL Peroxisomal marker construct (Nowak et al., Braunschweig 2004)

2.3 Primers All the primers were synthesized in HPSF (High Purity Salt-Free) quality at Eurofins Genomics (Ebersberg, Germany).

Primer name Sequence (5'→3') Sequencing primers pJET 1.2 Forward CGACTCACTATAGGGAGAGCGGC pJET 1.2 Reverse AAGAACATCGATTTTCCATGGCAG T7 TAATACGACTCACTATAGGG T7 term CTAGTTATTGCTCAGCGGT M13 Forward GTAAAACGACGGCCAG M13 Reverse CAGGAAACAGCTATGAC First-strand cDNA synthesis primers 5'-RACE CDS (T)25VN N = A, C, G, or T Clontech (5'-CDS) V = A, G, or C Laboratories 3'-RACE CDS AAGCAGTGGTATCAACGCA (3'-CDS) GAGTAC(T)30VN Random Hexamer Thermo Scientific 5'- and 3'-RACE PCR primers RACE Long CTAATACGACTCACTATAGGGCAAGCAGTGGT Clontech ATCAACGCAGAGT Laboratories RACE Short CTAATACGACTCACTATAGGGC Primers designed using transcriptome 2026015FNheI1 ATTGCTAGCATGGAAGGAGTTGTCAAGTGC 2026015RKpnI1 ATTGGTACCTTACAGTTTGCTGGCACTTGG 2009069FNheI1 ATTGCTAGCATGGAAGACCTCAAGCCCC 2009069REcoRI1 ATTGAATTCTCAGAGTCTACTAACTTTAGAAG 2019579FNheI1 ATTGCTAGCATGGGAGACATAGACGATCTC 2019579RKpnI1 ATTGGTACCTTATAACTTGCTCTTTGGGATTG HcBZLNheIF ATTGCTAGCATGGAGGGGCTGATGAGG HcBZLNcoIR ATTCCATGGTCAAGAAAGGCTGCCCATAG HcendF2 GGACATTATAATCTCTGGGGG RT-HpR3 ACTAGCTGCAGATGAATCATC UTRF3 GAGGCAAGGGATCATCATCA GSPR13 ACGTGGCCTGGGGAAATG Primers for cloning of luciferase ORF Luc_FNheI ATTGCTAGCATGGAAGACGCCAAAAAC Luc_RKpnI ATTGGTACCTTAAAGCTTCTTTCCGCC Primers for full-length HcBZL ORF attB product generation*

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Materials

HcLOCF GGGGACAAGTTTGTACAAAAAAGCAGGCTTAACCATGGAG GGGCTGATGAGG HcLOCrter GGGGACCACTTTGTACAAGAAAGCTGGGTCTCAAGAAAGG CTGCCCATAG HcLOCr GGGGACCACTTTGTACAAGAAAGCTGGGTCAGAAAGGCTG CCCATAGCC Primers for truncated HcBZL sequence attB product generation* HcLocSmall_R GGGGACCACTTTGTACAAGAAAGCTGGGTCGAAGTGCAGC TCGTACATC HcLocLongFragF GGGGACAAGTTTGTACAAAAAAGCAGGCTTAACCATGTAC GAGCTGCACTTCGC HcLocEndFragF GGGGACAAGTTTGTACAAAAAAGCAGGCTTAACCATGGCT CCGCGAAGTGTC Primers for full-length HcBD attB product generation* LocBDF GGGGACAAGTTTGTACAAAAAAGCAGGCTTAACCATGGCT GACCACACTAATG LocBDR GGGGACCACTTTGTACAAGAAAGCTGGGTCGAGCCAAGGA GAATTGTAAATG LocBDterR GGGGACCACTTTGTACAAGAAAGCTGGGTCTCAGAGCCAA GGAGAATTG Primers for screening positive Gateway expression clones eYFP F TGAACTTCAAGATCCGCCAC eYFP R AAGGTGGTCACGAGGGTG Primers for Real-time quantitative PCR (RT-qPCR) analysis of HcBZL qRTHcBZL_F2 ATGGTTCAGGAGCGGTGATC qRTHcBZL_R2 AACCGCTGCCTCGAGTATTG Reference gene primers for RT-qPCR analysis Histone-H2A-QF AACATCTACTCTTTGGACGACTTG El-Awaad et al., 2016 Histone-H2A-QR AATTGCTGGAGGTGGAGTTATTC Actin-QF CGGCAGTGGTTGTGAACAT Actin-QR TCTCGCTGGTCGTGATCTG Primers for semi-quantitative PCR of HcBD HcBD_RT_for ATGGCTGACCACACTAATGG HcBD_RT_rev AACCTTCACGAAGAACATCATGG Reference gene primers for semi-quantitative PCR Hp_H2A RTPCRF TCAGAACAGCTCCATCAAACC Hp_H2A RTPCRR GGAAGTCGACCAAGGACAAG Hp_18S_F4 TGATGGTATCTACTACTCGG Gaid et al., 2012 Hp_18S_R4 AATATACGCTATTGGAGCTGG 1Numbers indicate the Onekp sequence identifier assigned to the contig used for designing the primer. 2Used in combination with RACE Long and RACE Short primers to obtain the 3'-UTR of HcBZL 3Designed for cloning 5'-UTR of HcBZL 4The primer pair was used additionally to confirm the success of reverse transcription reaction

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Restriction sites are shown in bold *attB1 and attB2 sites are annotated in orange and purple colour, respectively

2.4 Enzymes

2.4.1 Enzymes used for reverse transcription reaction Enzyme name Manufacturer RevertAid H Minus Reverse Transcriptase Thermo Scientific RiboLock RNase Inhibitor Thermo Scientific

2.4.2 Enzymes used for Polymerase Chain Reaction Enzyme name Manufacturer peqGOLD Taq DNA polymerase Peqlab Q5 Hot Start High-Fidelity DNA Polymerase New England Biolabs Phusion Hot Start II High-Fidelity DNA Thermo Scientific Polymerase

2.4.3 Enzymes used for Gateway cloning Enzyme name Manufacturer BP ClonaseTM enzyme mix Invitrogen LR ClonaseTM enzyme mix Invitrogen Proteinase K Invitrogen

2.4.4 Restriction endonucleases Enzyme name Restriction Recognition Manufacturer site (5'→3') NheI G^CTAGC Thermo Scientific KpnI GGTAC^C Thermo Scientific EcoRI G^AATTC Thermo Scientific NcoI C^CATGG Thermo Scientific EcoRV(Eco32I) GAT^ATC Thermo Scientific

2.4.5 Miscellaneous enzymes Name Manufacturer FastAP Thermosensitive Alkaline Phosphatase Thermo Scientific T4 DNA ligase Thermo Scientific RNase A Thermo Scientific RNase-Free DNase I Qiagen Proteinase K Thermo Scientific

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2.5 Kits Name Purpose Manufacturer InnuPREP DOUBLEpure DNA extraction from agarose gel Analytik Jena slices, purification and concentration of PCR products InviTrap Spin Plant RNA Total RNA isolation Stratec Molecular Mini Kit qPCRBIO SyGreen Real-time RT-qPCR analysis Nippon Genetics Mix Lo-ROX

2.6 Ladder Name Manufacturer GeneRuler 1 kb DNA ladder Thermo Scientific GeneRuler DNA Ladder Mix Thermo Scientific Unstained Protein MW Marker Thermo Scientific PageRuler Unstained Protein Ladder Thermo Scientific

2.7 Chemicals Name Manufacturer Acetaldehyde (≥ 99.5%) Roth Acetic acid (99.5-100%) J.T.Baker Acetonitrile (HPLC) Fisher Chemicals Acetosyringone (3,5-dimethoxy-4-hydroxy-acetophenone) Sigma Aldrich Acrylamide/Bisacrylamide (30%) Bio-Rad Agar Kobe AppliChem Ammonium acetate J.T. Baker APS Roth Ampicillin sodium salt Roth 2-Aminobenzoic acid (Anthranilic acid, 98%) Aldrich ATP disodium salt (≥ 98%) Roth Beef extract Roth Benzaldehyde (≥ 80%) Roth Benzoic acid (≥ 99.5%) Roth Benzoyl-CoA lithium salt (≥ 90%) Sigma Aldrich Bovine serum albumin (BSA) Sigma Aldrich Bromophenol blue Sigma Aldrich Butyric acid (≥ 99%) Aldrich CaCl2.2H2O Merck Chloramphenicol AppliChem Chloroform Fisher Chemical trans-Cinnamaldehyde (99%) Aldrich trans-Cinnamic acid (> 99%) Merck CoCl2.6H2O Riedel-de Haën

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Materials

CoA trilithium salt (≥ 94%) Calbiochem Coomassie blue (G-250 and R-250) Sigma 4-Coumaric acid (≥ 98%) Sigma CTAB Roth CuCl2 Fluka CuSO4.5H2O Riedel-de Haën Dichloromethane (> 99.9%, GC) Sigma Aldrich 2,4-D Fluka 3,4-Dihydroxybenzaldehyde (Protocatechualdehyde, 97%) Acros Organic 2,3-Dihydroxybenzoic acid (99%) Aldrich 2,5-Dihydroxybenzoic acid (Gentisic acid, 99%) Acros Organic 3,5-Dihydroxybenzoic acid (97%) Acros Organic 3,4-Dihydroxycinnamic acid (Caffeic acid, ≥ 98%) ROTICHROM 3,5-Dimethoxy-4-hydroxycinnamic acid (Sinapic acid, ≥ Roth 98%) DTT Roth EDTA.2H2O Roth DMSO Roth Ethanol VWR Chemicals Ethyl acetate Merck Sigma Aldrich FeCl2 Roth FeSO4.7H2O Riedel-de Haën FAD sodium salt (≥ 95%) Sigma Formic acid Roth Gentamycin sulfate Roth Glucose Roth Glycerol (86%) Roth Glycine Roth H3BO3 Merck HCl (37%) VWR Chemicals Hexanoic acid (≥ 98%) Roth Heptanoic acid (96%) Sigma-Aldrich 2-Hydroxybenzaldehyde (Salicylaldehyde, ≥ 98%) Sigma-Aldrich 3-Hydroxybenzaldehyde (≥ 99%) Aldrich 4-Hydroxybenzaldehyde (98%) Aldrich 4-Hydroxy-3-methoxybenzaldehyde (Vanillin, > 99% ) Merck 2-Hydroxybenzoic acid (Salicylic acid, ≥ 99%) Sigma 3-Hydroxybenzoic acid (≥ 98%) Fluka 4-Hydroxybenzoic acid (99%) Fluka 3-Hydroxybenzoyl-CoA Laboratory collection 4-Hydroxy-3-methoxybenzoic acid (Vanillic acid, 97%) Aldrich Chemical 4-Hydroxy-3-methoxycinnamaldehyde (Coniferyl aldehyde, Aldrich 98%) 4-Hydroxy-3-methoxycinnamic acid (Ferulic acid, 99%) Sigma Isobutyric acid (99%) Sigma

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Materials

Isobutyryl-CoA lithium salt (≥ 85%) Sigma Aldrich Isopropanol VWR Chemicals IPTG Roth Kanamycin sulfate Roth KCl Roth KH2PO4 Roth KI Riedel-de Haën KNO3 Roth KOH Roth LiCl Sigma D-Luciferin (> 99%) Roth Malic acid Roth D (-)-Mandelic acid (≥ 99%) Fluka β-ME Roth Methanol (LC-MS) Roth Methanol (HPLC) Fisher Chemicals 2-Methoxybenzaldehyde (o-Anisaldehyde, 98%) Sigma 2-Methoxybenzoic acid (o-Anisic acid, 99%) Aldrich MES potassium salt Sigma MgCl2.6H2O Calbiochem MgSO4.7H2O Roth Midori Green Nippon Genetics MnCl2.4H2O Roth MnSO4.H2O Merck Myo-inositol Roth NaCl Roth Na2MoO4.2H2O Merck NaOH Roth NAA Fluka NH4Cl Riedel-de Haën NH4NO3 Roth NiCl2 Roth NAD GERBU NADH disodium salt reduced GERBU NADP disodium salt Roth MSTFA Alfa Aesar Octanoic acid (≥ 99.5%) Roth Peptone from casein Roth peqGold Universal Agarose Peqlab Phenol Roth L-Phenylalanine (> 99%) Merck ortho-Phosphoric acid Roth Potassium acetate Roth Propanoic acid (≥ 99.5%) Sigma-Aldrich Rifampicin Roth

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Materials

Shikimic acid (99%) Sigma SDS Roth Streptomycin sulphate Roth Sucrose Diadem TEMED Bio-Rad HCl Sigma-Aldrich 3,4,5-Trihydroxybenzoic acid (Gallic acid, > 98%) Fluka Tris Roth Xanthone references (Hyperxanthone E and Patulone) Purified from cell cultures by Dr. Mariam Gaid Xylene Cyanol FF Sigma Yeast extract Roth ZnCl2 Fluka ZnSO4.7H2O Riedel-de Haën

2.8 Culture media

2.8.1 Plant tissue culture medium and required phytohormones LS (Linsmaier and Skoog; Duchefa Biochemie) Macro Elements mg/l mM CaCl2 332.02 2.99 MgSO4 180.54 1.50 KH2PO4 170.00 1.25 KNO3 1900.00 18.79 NH4NO3 1650.00 20.61 Micro Elements mg/l µM CoCl2.6H2O 0.025 0.11

H3BO3 6.20 100.27 CuSO4.5H2O 0.025 0.10 KI 0.83 5.00 FeNaEDTA 36.70 100.00 Na2MoO4.2H2O 0.25 1.03 ZnSO4.7H2O 8.60 29.91 MnSO4.H2O 16.90 100.00 mg/l µM Myo-Inositol 100.00 554.94 Thiamine HCl 0.40 1.19 Sucrose 30 g/l Phytohormone 2,4 D (1 mg/ml) 220 µl NAA (1 mg/ml) 186 µl 2,4 D and NAA powder were dissolved in ethanol LS powder (4.4 g) and sugar were dissolved in water, followed by the addition of phytohormones. The pH was adjusted to 6.0 and the volume was made up to 1 liter with

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Materials

millipore water. The medium was then autoclaved.

2.8.2 Bacterial culture media and required supplements LB Final concentration Yeast extract 0.5% Peptone from casein 1.0% NaCl 1.0% Agar (for solid medium, added before 1.5% autoclaving) For ampicillin-based antibiotic selection, sterile-filtered ampicillin was added to the autoclaved medium at a final concentration of 100 mg/l when the temperature of the autoclaved medium was ~55˚C. For ZeocinTM-containing medium, NaCl was reduced to 0.5%. YEB Final concentration Beef extract 0.5% Yeast extract 0.1% Peptone from casein 0.5% Sucrose 0.5% MgSO4.7H2O 0.049% Depending upon the recombinant A. tumefaciens to be cultivated, the desired antibiotics were added to the autoclaved medium when its temperature was ~55˚C. SOC Final concentration Peptone from casein 2.0% Yeast extract 0.5% KCl 2.5 mM NaCl 10 mM The pH was adjusted to 7.5 with NaOH. The medium was autoclaved. Once the medium cooled to ~55˚C the remaining components (mentioned below) were added as sterile-filtered solutions. MgSO4.7H2O 5.0 mM MgCl2 5.0 mM Glucose 20.0 mM Aliquots of the medium were prepared and stored at -20˚C Antibiotics Stock preparation Final concentration (µg/ml) Ampicillin* 100 mg powder was dissolved in 1 ml 100 double distilled water Chloramphenicol 30 mg powder was dissolved in 1 ml 30 pure ethanol Kanamycin* 50 mg powder was dissolved in 1 ml 50 double distilled water Rifampicin 35 mg powder was dissolved in 100 methanol Gentamycin* 50 mg powder was dissolved in 1 ml 100

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double distilled water Streptomycin* 50 mg powder was dissolved in 1 ml 100 double distilled water ZeocinTM Supplied by Invitrogen-100 mg/ml 50 *Sterile filtered using 0.22 µm filter (Roth) before use.

2.9 Buffers and solutions

2.9.1 Buffers for plasmid isolation from E.coli Buffer I Final concentration Tris HCl 50 mM Na2EDTA.2H2O 10 mM RNase A 100 µg/ml The pH was adjusted to 8 with HCl. RNase A was added just before use. Buffer II Final concentration NaOH 0.2 M SDS 1% (w/v) Buffer III Final concentration Potassium acetate 3 M The pH was adjusted to 5.5 with glacial acetic acid.

2.9.2 Agrobacterium DNA extraction buffer Component Final concentration Tris HCl (pH 8) 110 mM EDTA 55 mM NaCl 1.54 M CTAB 1.1% (w/v)

2.9.3 Agrobacterium activation medium Component Final concentration MES-buffer (pH 5.6, adjusted with KOH) 10 mM MgCl2 10 mM Acetosyringone (dissolved in DMSO) 150 μM

2.9.4 Agarose gel electrophoresis 50x TAE buffer Final concentration Tris base 2 M Na2EDTA.2H2O 50 mM The pH was adjusted to 8.0 using glacial acetic acid. For the preparation of agarose gel and running buffer, the 50x stock was diluted to 1x.

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1-2% agarose gels were prepared in 1X TAE buffer for a sample run. 6x Loading Dye Final concentration Tris HCl 10 mM Na2EDTA.2H2O 60 mM Xylene cyanol FF 0.03% (w/v) Bromophenol blue 0.03% (w/v) Glycerol 60% (v/v) The pH of the solution was adjusted to 7.6 with HCl and the solution was stored at -20˚C.

2.9.5 SDS PAGE Stacking Gel (5 %)* Volume required Water 2.72 ml Acrylamide/Bisacrylamide 30% (w/v) 664 µl Tris HCl 1M pH 6.8 504 µl SDS 10% (w/v) 40 µl APS 10% (w/v) 40 µl TEMED 4 µl Separating gel (12 %)* Volume required Water 2.30 ml Acrylamide/Bisacrylamide 30% (w/v) 2.8 ml Tris HCl 1.5 M pH 8.8 1.75 ml SDS 10% (w/v) 70 µl APS 10% (w/v) 70 µl TEMED 2.8 µl * The volumes are for the preparation of 2 gels. 2x Protein loading buffer Volume required Tris HCl 0.5 M pH 6.8 1.0 ml SDS 10% (w/v) 3.3 ml Bromophenol blue 0.5% (w/v) 0.5 ml Glycerol 2.0 ml β-Mercaptoethanol (added freshly) 0.5 ml Water 2.7 ml 10x Electrophoresis buffer (pH 8.3) Final concentration Tris base 80.6 mM Glycine 1.9 M SDS 1% (w/v) For the preparation of running buffer, the 10x stock was diluted to 1x Coomassie blue R250 stock solution Final concentration Coomassie blue R250 in methanol 1% (w/v) The solution is filtered prior to use. Staining solution Final concentration Coomassie blue (stock) 10% (v/v) Acetic acid 10% (v/v) Methanol 50% (v/v)

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Destaining solution Final concentration Acetic acid 10% (v/v) Methanol 15% (v/v)

2.9.6 His6-tagged protein affinity purification Lysis Buffer (pH 8) Final concentration (mM) NaH2PO4 50 NaCl 30 Imidazole 20 Wash buffer (pH 8) Final concentration (mM) NaH2PO4 50 NaCl 1.5 Imidazole 50 Elution buffer (pH 8) Final concentration (mM) NaH2PO4 50 NaCl 300 Imidazole 250

2.9.7 Bradford solution for determination of protein concentration Bradford dye solution Final concentration Coomassie Brilliant Blue G-250 0.01% (w/v) Ethanol absolute 5% (v/v) ortho-Phosphoric acid (85%) 10% (v/v) Coomassie Brilliant Blue G-250 was first dissolved in ethanol followed by the addition of ortho-phosphoric acid. The solution was then made to 1 l using double distilled water. The solution was then filtered prior to use and stored at 4˚C in an amber bottle.

2.10 Online tools and databases Databases, web address Description NCBI database Source of nucleotide and amino acid sequences https://www.ncbi.nlm.nih.gov/nuccore associated with CoA ligases and aldehyde https://www.ncbi.nlm.nih.gov/protein dehydrogenases MPGR database Transcriptomic and metabolomic data for 14 http://medicinalplantgenomics.msu.edu/ medicinal plant species including H. perforatum OneKP database Transcriptomic data for over 1000 plant species http://www.onekp.com/ including H. perforatum All the databases possess a BLAST tool for blasting a query sequence against a particular plant transcriptome or against all the plant transcriptomes. RT-qPCR primer designing and amplicon analysis tools Tool name Link Primer 3.0 http://primer3.ut.ee/ uMeltSM v2.0.2 https://dna.utah.edu/umelt/um.php

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Miscellaneous tools ExPASy translate tool http://web.expasy.org/translate/ DoubleDigest Calculator https://www.thermofisher.com/de/de/home/brand s/thermo-scientific/molecular-biology/thermo- scientific-restriction-modifying- enzymes/restriction-enzymes-thermo- scientific/double-digest-calculator-thermo- scientific.html Oligo Analysis Tool, Eurofin Genomics https://www.eurofinsgenomics.eu/en/dna-rna- oligonucleotides/oligo-tools/ Oligo Analyzer Tool, Integrated DNA https://eu.idtdna.com technologies Subcellular localization prediction tools# DeepLoc-1.01 http://www.cbs.dtu.dk/services/DeepLoc/ LocTree32 https://rostlab.org/services/loctree3/ Plant-mPLoc3 http://www.csbio.sjtu.edu.cn/bioinf/plant-multi/ PPero4 https://biocomputer.bio.cuhk.edu.hk/PP/ PSORT II5* https://psort.hgc.jp/form2.html Target Signal Predictor/ PTS1 http://216.92.14.62/Target_signal.php and PTS2 binding site4 #These tools can differentiate between various localizations of protein: 1Cell membrane, chloroplast, cytoplasm, endoplasmic reticulum, extracellular space, Golgi apparatus, mitochondria, nucleus, peroxisomes and vacuole 2,5Cytoplasm, cytoskeleton, endoplasmic reticulum (ER), extracellular space including cell wall, Golgi, mitochondria, nucleus, peroxisomes, plasma membrane, vacuole and vesicles of secretory system 3Cell membrane, cell wall, chloroplast, cytoplasm, endoplasmic reticulum, extracellular space, Golgi apparatus, mitochondria, nucleus, peroxisomes and vacuole 4Peroxisome *This prediction tool does not support input sequences from plants All listed websites have been successfully accessed on June 11, 2019

2.11 Softwares Name, Web address Use Lasergene-DNAStar 7.0 It is a package of multiple https://www.dnastar.com/cb/f-reg-submit.aspx programs used for nucleotide and protein sequence analysis Bio-Rad CFX Manager 3.1 Used for designing qPCR http://www.bio-rad.com/de-de/product/previous-qpcr- plates, calculation of the software-releases?ID=OO2BB34VY amplification efficiencies and data analysis MEGA 7.0 Used for building http://www.megasoftware.net/ phylogenetic trees (Kumar et al., 2016)

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ZEN 2.3 (Carl Zeiss) Used for visualization and https://www.zeiss.com/microscopy/int/products/microscope- analysis of images produced software/zen-lite.html by LASER scanning microscopy. Hyper32 data analysis https://hyper32.software.informer.com/download/ Image J 1.52a Image processing program for https://imagej.nih.gov/ij measuring the pixel intensity of the PCR bands SnapGene For simulating restriction and https://www.snapgene.com/try-snapgene/ Gateway cloning and visualizing DNA sequences and their features All listed websites have been successfully accessed on June 11, 2019

2.12 Equipment

2.12.1 General equipment Equiment Model Manufacturer Autoclave Systec VX-120 Systec Balance ABJ-NM/ABS Kern Kern 572 Kern Centrifuge Sigma 1-15 K Sigma Universal 32 R Hettich Centrifuge 5810 R Eppendorf Dry block heater Dri-Block DB-3D Techne Electrophoresis chamber Sub-Cell GT Agarose Gel System Bio-Rad (agarose gels) Electrophoresis chamber Mini-PROTEAN Tetra System Bio-Rad (PAGE) Gel documentation Infinity-3000 Vilber Lourmat Heating circulators- Water MW-4 Julabo Bath Incubator shaker Multitron Infors HT KF4 Laminar Air flow LaminAir HLB 2472 Heraeus Luminometer Victor2 multilabel counter; Perkin Elmer Wallac 1420 Lyophilizer Gamma -20 Christ Lyophilizing apparatuses GmbH Magnetic stirrer IKA-Combimag RET Janke & Kunkel

Microscope (confocal laser cLSM-510META connected Carl Zeiss

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scanning) to Axiovert 200M MicroPulser Bio-Rad pH meter pH 325 WTW (wissenschaftlich- technische-werkstätten) Power Pack Standard Power Pack P25 Biometra Power Pac 300 Bio-Rad Real-Time PCR Detection CFX Connect Bio-Rad System Thermocycler Tpersonal Biometra T-Professional basic gradient Biometra Ultra-Low Temperature MDF-U53V Sanyo Freezer (-80 °C) Ultrasonic Cell Disruptor Sonifier 250 Branson (G. Heinemann) UV-VIS Spectrophotometer UVmini-1240 Shimadzu

Ultrospec 1000 Pharmacia Biotech

SimpliNano GE Healthcare Life Sciences Vortex Vortex-Genie 2 Scientific Industries Vortex shaker Vortex genie 2 Digital Scientific industries Water purification system Arium 611 Sartorius

2.12.2 Equipment for HPLC and MS analyses Equipment Model Manufacturer HPLC Agilent 1260 Infinity System Agilent Pump G1311C Quaternary Pump VL Agilent Autosampler G1329B ALS Agilent TM Column C18 column (3.5 µm, 4.6 x 100 XBridge mm) for xanthone quantification and CoA ligase assay Extend-C18, 3.5 μm, 4.6 X 150 Agilent mm for BD assay Diode array detector G1315D DAD VL Agilent Software Agilent ChemStation for LC Agilent 3D System Rev. B.04.03(16) Mass spectrometer 3200 QTRAP LC/MS/MS System AB Sciex

Software Analyst vers. 1.4.2 AB Sciex GC-MS Agilent 6890 gas chromatograph Agilent Column ZB5-MS column (30 m, Phenomenex 0.25 mm i.d., 0.25 mm ft) Oven GC 5890 A Hewlett-Packard

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Company Mass spectrometer Agilent 5975B MSD Agilent Software Chemstation Agilent

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3 Methods

3.1 Cultivation of in vitro cultures

3.1.1 Propagation and elicitation of H. calycinum cell suspension cultures Cell cultures of H. calycinum were propagated in liquid LS medium (Linsmaier and Skoog, 1965; 2.8.1) as mentioned previously (Fiesel et al., 2015). Suction-dried H. calycinum cells (3g) were transferred every two weeks into 50 ml LS medium. The cultures were shaken in Erlenmeyer flasks (300 ml) at 120 rpm and 25 ± 1˚C in the dark. H. calycinum cells were collected every second day starting from the day of subculture for sixteen days and were lyophilized. The dry weight of the cells was plotted as a function of time to determine the growth curve of H. calycinum cells.

For elicitation of H. calycinum cell suspension cultures, yeast extract solution was prepared. Yeast extract (150 mg) was dissolved in 1 ml double-distilled water and autoclaved (120˚C, 20 min). The sterile stock solution was ready to use and could also be stored at 4˚C. Yeast extract solution (2 ml) was added to 100 ml homogenous H. calycinum cultures on the fourth day after subculture to give a final concentration of 3 g/l (Gaid et al., 2012). Post-elicitation, cell suspension cultures (2 ml) were collected at various intervals (0, 4, 8, 12, 16, 24, 28, 32, 36, 48, 72, 96, 120, 144 h). The collected samples were centrifuged at 5000 rpm for 5 min. The supernatant, which is the LS medium (1 ml), was collected in Eppendorf tubes and stored at - 20˚C until extraction (section 3.2). The sediment, which were the cells, was lyophilized for 2 days in the lyophilizer to get rid of the moisture content and was then stored at -20˚C until extraction (section 3.2). Both the cells and the medium were analyzed for the total xanthone content.

3.1.2 Cultivation of N. benthamiana potted plants Seeds of N. benthamiana were sprinkled on autoclaved and cooled flowering soil in a pot and cultivated in an acclimatization room at 24˚C with 65% humidity and a light period of 12 h (30 μmol/m2s). After 2-3 weeks, the seedlings were transferred to separate pots and grown under identical conditions for another 2-3 weeks.

3.2 Extraction of xanthones from H. calycinum cell suspension cultures Lyophilized cells (20 mg) mixed with 1 ml methanol (HPLC grade) and small amount of sea sand were vortexed for 15 min using a vortex shaker. The homogenate was centrifuged at 14000 rpm for 10 min. The supernatant was collected. The extract was filtered through a sterile filter (0.20 µm; Chromafil O-20/15 MS, Macherey-Nagel) to remove insoluble particles which could interfere with the chromatographic separation of the compounds. The culture medium was extracted with ethyl acetate. Culture medium (1 ml) was acidified with formic acid (1 μl) and mixed with 500 µl ethyl acetate. The mixture was intensively vortexed for 15 min. The organic phase was collected following centrifugation at 14000 rpm for 10 min. The process was repeated

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Methods and the total organic phase collected was pooled and evaporated to dryness. The residue was dissolved in 100 µl methanol (HPLC grade). The possible xanthone content for both cells and medium was analyzed via HPLC using aliquots of the methanolic extracts (50 µl) as discussed later under section 3.6.1.1.

3.3 Molecular biology methodologies

3.3.1 Isolation of nucleic acids

3.3.1.1 Isolation of total RNA from H. calycinum cells H. calycinum cultures elicited with yeast extract (3 g/l) were collected 8 h post-elicitation and total RNA was extracted from the cells using InviTrap Spin Plant RNA Mini Kit (Stratec Molecular; 2.5) following the manufacturer’s protocol. This procedure relies heavily on the presence of mineral carrier particles in the lysis solution, which not only helps in shearing of the plant material but also allows selective binding of genomic DNA (Stratec Biomolecular). Under the strong denaturing lysis conditions, the cells get quickly disintegrated, the RNases are simultaneously deactivated and the RNA is secured. The genomic DNA bound mineral carrier particles are then removed by centrifugation.

3.3.1.2 Isolation of plasmid DNA from E. coli cells The alkaline extraction procedure by Birnboim and Doly (1979) was employed for the isolation of plasmids from bacterial cells. It is based upon the selective denaturation of high molecular weight chromosomal DNA under alkaline conditions while the small circular plasmid DNA retains its native configuration. Upon neutralization of the conditions, large molecular weight chromosomal DNA and proteins form a precipitate while circular plasmid maintains its soluble state.

For plasmid isolation, a single E. coli colony was inoculated in a 5 ml LB tube containing the required antibiotic (ampicillin or ZeocinTM). The salt concentration of the LB medium was half of the standard amount when ZeocinTM was used for selection. The cultures were grown overnight at 37˚C with continuous shaking at 200 rpm. For the subsequent steps, 2 ml of the culture was transferred into a 2 ml Eppendorf tube and was pelleted by centrifuging at 5000 rpm for 5 min. The supernatant was discarded and the pellets were resuspended in 300 μl ice-cold buffer I (2.9.1) which was freshly supplemented with RNase A (2.4.5). For alkaline extraction, 300 μl buffer II (2.9.1) was added, followed by gentle mixing of the contents by flipping the Eppendorf tubes five times and incubating at room temperature for 5 min. Neutralization of the contents was done by the addition of 300 μl buffer III (2.9.1). Subsequently, the Eppendorf tubes were gently mixed by flipping five times and were stored in ice for 20 min. It resulted in precipitation of chromosomal DNA and proteins. The resulting mix was centrifuged at 14000 rpm for 10 min. 700 μl clear supernatant was acquired and was added to an equal volume of chloroform in a new Eppendorf tube. The immiscible mix was vortexed well and centrifuged at 14000 rpm for 10 min. 420 μl of the aqueous phase was transferred to a 1.5 ml Eppendorf tube

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Methods and mixed with an equal volume of isopropanol. The mixture was vortexed and centrifuged at 14000 rpm for 20 min. The supernatant was discarded and the pellet was washed with 500 μl of 70% ethanol. Following another round of centrifugation at 14000 rpm for 20 min, the supernatant was discarded and the pelleted plasmid was dried at 37˚C in an oven for 30 min. In the end, 30 μl of deionized water was added to the dried plasmid. Depending upon the downstream procedure the plasmid served as a template for restriction digestion (3.3.7.1), a template for amplification of the desired insert (3.3.5.1) or it was simply sent for sequencing (3.3.9).

3.3.1.3 Isolation of total DNA from A. tumefaciens A single A. tumefaciens colony was inoculated in 10 ml YEB medium supplemented with suitable antibiotics and was grown overnight at 28˚C with continuous shaking at 220 rpm. Culture medium (2 ml) was centrifuged at 6500 rpm for 1 min. The supernatant was discarded and the step was repeated. The pellets were resuspended in 500 μl of 100 mM Tris HCl (pH 8) and centrifuged at 6500 rpm for 1 min. The supernatant was discarded. These steps were repeated once more. The pellets were resuspended in 600 μl of Agrobacterium DNA extraction buffer (2.9.2) which was freshly supplemented with 50 μl of proteinase K (5 mg/ml; 2.4.5) and shortly vortexed. SDS (10%, 160 μl) was added to the suspension and mixed gently by flipping the tubes five times. The resulting suspension was incubated at 65˚C for one hour. Following incubation, the suspension was cooled to room temperature and phenol:chloroform (1:1, 500 µl) was added. The resulting mixture was vortexed intensively. It was centrifuged at 14000 rpm for 20 min at room temperature. The supernatant (650 µl) was collected and intensively mixed with 390 μl isopropanol. The mixture was centrifuged at 4˚C, 14000 rpm for 20 min. The supernatant was then decanted, while the pellets were washed with 500 μl of 70% ethanol and centrifuged at 4˚C, 14000 rpm for 10 min. The supernatant was again decanted and the pellets were dried at 37˚C in an oven for 30 min. The dried pellets were dissolved in 20 μl Tris HCl (10 mM, pH 8).

Total DNA extracted served as a template for PCR amplification (3.3.5.1) to detect the integration of the gene of interest in the Agrobacterium genome.

3.3.1.4 Isolation of DNA from agarose gel and purification of PCR and restriction digestion products Isolation of DNA from agarose gel, purification of PCR and restriction digestion products was done using the innuPREP DOUBLEpure Kit (Analytik Jena; 2.5) as per manufacturer’s protocol. In this technique, DNA was firstly bound selectively to the silica membrane of the spin column, washed with ethanol containing buffer and then eluted with an aqueous buffer.

3.3.2 Quantification of nucleic acids The concentration of nucleic acids was determined spectrophotometrically by recording the UV absorbance of a diluted sample (1:200) at 260 nm. The concentration of the sample can be calculated by the Beer-Lambert equation which is as follows:

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where C is the concentration of nucleic acid (ng/μl)

OD260 is the absorbance of the nucleic acid sample at 260 nm after normalization with respect to a blank l is the path length (1 cm)

ε is the standard coefficient for the nucleic acid being measured (50 µg/ml for double-stranded DNA, 33 µg/ml for single-stranded DNA and 40 µg/ml for RNA)

D is the dilution factor

Traditionally OD260/OD280 is used to assess the purity of DNA and RNA samples. In principle, the ratio is ~1.8 for pure DNA and ~2.0 for pure RNA. OD260/OD280 values lesser than the aforesaid values indicate the presence of protein, phenol or other contaminants in the sample which absorb at or near 280 nm. OD260/OD230 is another accessory value that can be used to support the purity of DNA and RNA samples. For pure samples, the value ranges between 2.0- 2.2. Lower values indicate the presence of contaminants which absorb at or near 230 nm like EDTA or guanidine HCl.

3.3.3 Reverse transcription for first-strand cDNA synthesis Reverse transcriptase is an RNA-dependent DNA polymerase catalyzing first-strand cDNA synthesis from total RNA. The primers for first-strand cDNA synthesis are of three types, namely oligo(dT) primer which binds to the polyadenylated tail of mRNA, random hexamers which improve cDNA yield and lastly gene-specific primers which offer specific priming and enrichment of the gene of interest. For best results, a combination of oligo(dT) and random hexamer primers should be used in reverse transcription. The synthesized cDNA can then be used for amplification of the desired genes. The standard components and reaction conditions of a reverse transcription reaction are listed in Table 3.1.

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Table 3.1: Standard components and reaction conditions of a reverse transcription reaction

Step Component Volume (µl) Template RNA X (1 μg) Oligo(dT) primer (10 μM) 1 dH2O Up to 12.5 Denaturation 65˚C for 5 min Primer annealing Store reaction mix on ice for 5 min The volume is made up to 20 μl by the addition of the components mentioned below: 5X reaction buffer 4 RiboLock™ RNase Inhibitor 0.5 (40 U/µl; 2.4.1) dNTP-Mix (each 10 mM) 2 RevertAid H Minus Reverse 1 Transcriptase (200 U/µl; 2.4.1) Reverse transcription 25˚C for 5 min* 42˚C for 60 min Enzyme deactivation 70˚C for 10 min *This step is added when a mixture of random hexamer primers is either used alone or in combination with oligo(dT) primer for reverse transcription.

Synthesis of the 3'-RACE-ready cDNA was carried out using the reverse transcription procedure with 3'-CDS primer, which is an oligo(dT) primer with an anchor sequence at its 3' end. After reverse transcription, the newly synthesized first-strand cDNA with the anchor sequence at its 5' end can serve as template for second strand synthesis using a gene-specific forward primer and an anchor-specific reverse primer (RACE Long). The specificity of the amplified product can be increased by another round of PCR using gene-specific forward primer and RACE Short as reverse primer. This amplification will provide the sequence information regarding the 3'-UTR region of the gene.

5'-RACE-ready cDNA can be produced by the combined action of SMARTScribeTM reverse transcriptase (Clontech) and SMART IITM A oligonucleotide (Clontech). SMARTScribeTM reverse transcriptase displays a terminal activity and adds 3-5 cytosine residues at the 3' end of the synthesized first-strand cDNA. The SMARTTM II A Oligo contains terminal G residues that pair with the cytosine-rich cDNA tail and serve as an extended template for the reverse transcriptase. SMARTScribeTM reverse transcriptase switches the template from the RNA to SMARTTM II A oligo generating the complete cDNA copy of the RNA with the added SMART sequence or the anchor at the 3' end. 5'-RACE-ready cDNA can serve as template for second strand synthesis using RACE Long as the forward primer and a gene-specific reverse primer. RACE Long is specific to the anchor sequence of the first-strand cDNA. Similar to 3' RACE PCR, the specificity of the amplified product can be increased by another round of PCR with RACE Short forward primer and gene-specific reverse primer. This amplification would

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Methods provide the sequence information regarding the 5' UTR region of the gene. In the current work, information on the 5' UTR region of the genes was obtained by setting up a PCR with 5'-CDS primed first-strand cDNA template and 5' UTR-specific forward and gene-specific reverse primers designed based on sequence information from the databases.

3.3.4 Primer design General points to be considered while designing primers are as follows:

• To ensure specific binding, ideally, the primer length should be 18-24 bases. • The GC content of the primer should range between 40-60 % with the 3’ end terminating with G or C to enhance binding. Primer binding to the template is further enhanced when the last 5 bases at the 3' end have at least 2 G or C bases. • Runs of 4 or more of a particular base and dinucleotide repeats should be avoided as its presence can lead to mispriming.

• Primers with melting temperature (Tm) in the range between 55-65˚C are suitable for most of the templates. The Tm of a primer pair should be close, with a maximum difference of 5˚C. • Intra-primer complementarity and inter-primer complementarity must be avoided to prevent self-dimers/hairpins and primer-dimer formation, respectively. IDT Oligo Analyzer and http://biotools.nubic.northwestern.edu/OligoCalc.html can be used for detection of secondary structure. Primer homology to other regions of the template should be avoided to ensure amplification of correct PCR product. • When designing primers with restriction sites at the 5' ends, 3 additional nucleotides (anchor) are added before the restriction digestion site to ensure efficient cutting by the enzyme. When calculating the Tm of these primers, the anchor and the restriction site sequences should not be taken into consideration.

Special attention should be given when designing primers for amplifying attB products employed in Gateway Cloning Technology (Invitrogen).

Forward primer: The 5' end of the forward primer for attB product amplification must contain four guanine residues followed by 25 bp attB1 sequence and at least 18-25 bp gene-specific sequence. If the gene of interest is needed to be fused with an N-terminal tag, two additional nucleotides must be added to the primer after the attB1 sequence to maintain the correct reading frame as attB1 ends with thymidine. attB1 Forward primer

5'-GGGG-ACA AGT TTG TAC AAA AAA GCA GGC TNN-(Gene-specific sequence)-3'

Orange = attb1 sequence; NN cannot be AA, AG, GA

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Reverse primer: The 5' end of a reverse primer must contain four guanine residues followed by the 25 bp attB2 sequence and at least 18-25 bp gene-specific sequence. For the fusion of the gene of interest to a C-terminal tag, one additional nucleotide must be added after the attB2 sequence to maintain correct reading frame and the in-frame stop codon must be removed. attB2 Reverse primer

5'-GGGG-AC CAC TTT GTA CAA GAA AGC TGG GTN-(Gene-specific sequence)-3'

Purple = attB2 sequence; N can be any nucleotide.

Amplification using the attB primers was done using a standard PCR (3.3.5.1). The PCR products were run on an agarose gel (3.3.6), excised and subsequently purified (3.3.1.4).

3.3.5 Polymerase chain reaction (PCR) PCR is a technique established by Kary Mullis, for which he was awarded the Nobel Prize in Chemistry in 1993. It relies on the principle of complementary nucleic acid hybridization and nucleic acid replication which are carried out a multiple number of times leading to the exponential increase of the target region of the DNA. The reaction consists of various components. It requires a DNA template which can be genomic DNA, cDNA or plasmid DNA, primers that bind to peripheries of the target region, deoxyribonucleotides (dNTPs), a thermostable DNA polymerase and reaction buffer.

PCR is a three-step process. The first step is denaturation at 95-98˚C for 30 s which leads to the melting of DNA i.e. disruption of hydrogen bonds between complementary bases and the separation of the two strands of a double-stranded DNA molecule. Each strand serves as a template for the next step, which is annealing where primers bind to their complementary sites on the single-stranded DNA at annealing temperature (Ta = Tm-5˚C or Ta = Tm+3˚C). The last step is extension which is carried out at 72˚C at which most DNA polymerases exhibit optimum activity. In this step, the DNA polymerase adds dNTP complementary to the base on the template strand at the 3' end of the primer. The duration of this step depends on the length of the target to be amplified and the speed at which the DNA polymerase adds dNTPs to the daughter strand being synthesized. The steps are repeated 20-30 times in an automated thermocycler leading to exponential accumulation of the desired DNA fragment. Also, a standard PCR consists of three phases. First is the exponential phase where with each cycle the amount of DNA doubles (assuming 100% reaction efficiency). All the reagents are in surplus amount. Next is the linear phase wherein because of the consumption of reagents the reaction slows down and the doubling of DNA is not observed. The last phase is the plateau phase or end point where product formation does not occur.

The amplified PCR products were then analyzed by agarose gel electrophoresis (3.3.6).

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3.3.5.1 Standard PCR In general, this method was used for screening plasmids for the presence of the target insert after ligation (3.3.7.3) or plasmid isolation (3.3.1.2). Thermus aquaticus (Taq) DNA polymerase was the DNA polymerase of choice for the screening experiments. The components of the standard reaction and thermocycling conditions are mentioned in Tables 3.2 and 3.3, respectively.

Table 3.2: Components of a standard PCR

Component Volume (25 μl) Water Up to 25 10X reaction buffer 2.5 dNTPs (10 mM each) 1 Forward primer (10 μM) 1 Reverse primer (10 μM) 1 Template DNA X (100-150 ng) Taq DNA polymerase (5U/μl; 0.25 2.4.2)

Table 3.3: Thermocycling parameters associated with a standard PCR

Step Temperature Time Cycles Initial Denaturation 95˚C 3 min - Denaturation 95˚C 30 s 30 Annealing Tm-5˚C 45 s Extension 72˚C 2 min 30 s Final Extension 72˚C 10 min - Storage 12˚C Pause

3.3.5.2 Touchdown PCR It is a variant of standard PCR employed for templates that were challenging for target amplification. This type of PCR ensures high product specificity and yield during amplification. The reaction components of this PCR are the same as standard PCR, however, the PCR cycling parameters are variable. The cycling program has two phases (Table 3.4). The first phase is the touchdown phase which begins with a Ta equal to the lower Tm value of the primer pair being used and transitions to lower Ta over successive cycles. With each cycle, a decrease in Ta occurred by -0.5˚C/cycle for 10 cycles till Ta = Tm-5˚C of the primers was reached. The second phase involved the exponential amplification of the enriched target at the final Ta reached in phase I. In this way, at high temperature in phase I primer-dimer formation and non-specific primer binding are prevented and only specific amplification can occur, however, the yield is compromised. As the temperature falls with each cycle the desired PCR products are selectively enriched and in phase II the yield of the desired enriched sequence is increased exponentially surpassing that of any non-specific sequence amplified during initial enrichment.

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Table 3.4: Thermocycling parameters of a Touchdown PCR

Phase I Step Temperature Time Cycle ΔT Initial Denaturation 95˚C 3 min - Denaturation 95˚C 30 s 10 -0.5˚C/ cycle Annealing Tm 30 s Extension 72˚C 2 min 30 s Phase II Step Temperature Time Cycle Denaturation 95˚C 30 s 30 Annealing Tm-5˚C 30 s Extension 72˚C 2 min 30 s Final extension 72˚C 10 min - Storage 12˚C Pause

3.3.5.3 High-fidelity PCR High-fidelity PCR relies on DNA polymerases which combines low dNTP misincorporation rate with proofreading activity (3'→5' exonuclease activity) to provide accurate replication of the target gene. High-fidelity DNA polymerases are employed in PCR where the correct DNA sequence is essential for the downstream experiment like cloning/subcloning of the target gene in an expression vector. Unlike Taq DNA polymerase products which possess 3' A overhangs due to the enzyme’s terminal transferase activity, proofreading polymerase products are blunt ended because of the 3'→5' exonuclease activity. The standard components and thermocycling conditons of a high-fidelity PCR are indicated in Tables 3.5 and 3.6, respectively.

Table 3.5: Standard components of Phusion Hot Start II and Q5 Hot Start high-fidelity PCR Table 3.5: Component Volume (μl) Component Volume (μl) dH2O Up to 20 5X Q5 Reaction buffer 5 5X Phusion HF Buffer 4 dNTPs (10 mM each) 0.5 dNTPs (10 mM each) 0.4 Forward Primer (10 μM 1.25 Forward Primer (10 μM) 1 Reverse Primer (10 μM) 1.25 Reverse Primer (10 μM) 1 Template DNA (10-50 ng) 1 Template DNA (10-50 1 Q5 Hot Start High-Fidelity 0.25 ng) DNA Polymerase; 2.4.2) Phusion Hot Start II 0.2 dH2O Up to 25 DNA polymerase (2U/μl; 2.4.2)

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Table 3.6: Thermocycling parameters associated with a high-fidelity PCR

Step Temperature Time Cycle Initial Denaturation 98 ˚C 30 s 1 Denaturation 98 ˚C 10 s 30 Annealing Tm+3˚C 30 s Extension 72˚C 1 min 30 s Final extension 72˚ C 10 min 1 Storage 12˚C Pause

3.3.5.4 PCR for analyzing gene expression In the current study the total RNA was extracted from untreated (0 h; control) and yeast extract- treated H. calycinum cells 1, 2, 4, 8, 12, 16, 20, 24, 36, 48 h post-elicitation using InviTrap Spin Plant RNA Mini Kit (Stratec Molecular; 2.5) following the manufacturer’s protocol. Genomic DNA contamination was removed from all samples through on-column digestion using RNase- Free DNase Set (Qiagen, 2.4.5). RNA concentration, 260/280 and 260/230 ratios were determined using the SimpliNano Spectrophotometer (GE Lifesciences). The cDNA corresponding to each time-point was synthesized using 1 µg RNA, 5 µM 5' CDS, 5 µM random hexamer primers and RevertAid H Minus Reverse Transcriptase (2.4.1). The cDNA produced was subjected to 1:50 dilution resulting in 1 ng/μl cDNA. For gene expression analysis, no reverse-transcriptase (NRT) control and no template control (NTC) were also prepared to check for genomic DNA contamination of cDNA and primer-dimer formation, respectively. In NTC reaction setup cDNA was replaced by an equal volume of water.

3.3.5.4.1 Semi-quantitative PCR Semi-quantitative PCR can be used for preliminary analysis of the expression pattern of a gene relative to a reference gene. Amplification cycle is a crucial parameter in this analysis. Optimization of the amplification cycle is vital and a cycle number should be selected such that the product is visible on an agarose gel and can be quantified by image processing tool. The optimal number of cycles should be in the same range for the target gene and the internal control. The components of a semi-quantitative PCR were the same as the standard PCR. The thermocycling parameters utilized for semi-quantitative PCR are mentioned in Table 3.7.

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Table 3.7: Thermocycling parameters used for semi-quantitative PCR

Step Temperature Time Cycle Initial denaturation 94˚C 3 min - Denaturation 94˚C 45 s 30 Annealing Tm-5˚C 45 s Extension 72˚C 1 min 30 s Final extension 72˚C 10 min - Storage 12˚C Pause

HcBD expression analysis was done by semi-quantitative PCR. The reference genes 18S rDNA and Histone 2A (H2A) were chosen as internal standards for normalizing the expression. Reaction components were the same as that for a standard PCR with the template amount being 4 ng in a 25 μl PCR for amplification of HcBD and H2A and 2 ng in a 25 μl PCR for amplification of 18S rDNA. After PCR, the products were run on a 1% (w/v) agarose gel containing Midori Green staining dye and were photographed by the gel-imaging system (Infinity-3000; Vilber Lourmat) with the exposure time being 1 s. Image J 1.52a software (2.11) was used for measuring the pixel intensity of the PCR bands corresponding to HcBD, HcH2A and Hc18S rDNA PCR products. HcBD expression level was calculated as relative integrated intensity normalized with respect to H2A and 18S rDNA.

3.3.5.4.2 Quantitative-PCR Contrary to semi-quantitative PCR, quantitative PCR (qPCR) is a highly sensitive technique that allows the amplification as well as the quantification of the nucleic acid target in real time. PCR products are detected by using fluorescent dyes that intercalate with the double-stranded DNA or by using fluorescently labeled sequence-specific probes. During each cycle of amplification, the amount of fluorescence is measured which is directly proportional to the amount of amplified product.

The RT-qPCR analysis was carried out for the determination of HcBZL expression. Actin and H2A served as the reference genes during the analysis (El-Awaad et al., 2015; Tocci et al., 2018, Nagia et al., 2019). The experiment was carried out with three biological repeats. For each biological repeat, 3 technical repeats were pipetted. The reaction components of a qPCR are mentioned in Table 3.8.

Table 3.8: Components for RT-qPCR setup

Component Volume (10 μl) cDNA 4 (1 ng) Forward primer (10 μM) 1 Reverse primer (10 μM) 1 2X qPCRBIO SyGreen Mix Lo-ROX buffer 4 (2.5)

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The reaction was pipetted into a 96 well plate (Bio-Rad) and sealed with a microseal ‘B’ adhesive sealing film (Bio-Rad). The plate was given a short spin to bring the contents at the bottom of the wells and was then placed in the thermocycler (Bio-Rad CFX Connect). The thermal cycling program mentioned in Table 3.9 was selected.

Table 3.9: Thermal cycling parameters of the RT-qPCR analysis

Step Temperature Time Cycle Initial denaturation 95˚C 2 min - Denaturation 95˚C 5 s 40 Annealing/ Extension 60˚C 30 s 95˚C 30 s - Melt curve 65˚C→95˚C; 0.05 s 0.5˚C increment

The device detected the fluorescence of the SYBR Green associated with the amplified DNA after each cycle. At the end of the amplification, melt curve analysis was performed in order to determine the specificity of the product. The amplified product was also run on a 2% (w/v) agarose gel. The PCR product was extracted from the agarose gel (3.3.1.4) and was sent for sequencing (3.3.9).

3.3.6 Agarose gel electrophoresis Electrophoresis is the migration of charged molecules under the influence of an electric field. Agarose gel electrophoresis is a traditional method used for the size-based separation of DNA fragments in a cross-linked agarose matrix. DNA is negatively charged and migrates towards the anode upon application of voltage. The main factors affecting the migration rate of DNA are the size of the DNA, agarose concentration and the conformation of DNA.

For the preparation of a 1% agarose gel which can be used for running 15–20 samples, 0.6 g of agarose was dissolved in 60 ml of 1X TAE buffer by heating the mix until a clear solution was obtained. Once the solution cooled to ~60˚C, 1.5 μl Midori Green was added. The solution was mixed well and applied on a suitable gel casting tray containing the desired comb. After the solidification of the gel, the comb was removed and the gel tray was transferred to the horizontal electrophoresis chamber. Samples were mixed with loading dye and were loaded in the wells of the gel. DNA ladder (5 µl, 2.6) was added in one of the wells for determination of the size of the PCR products. The gel was run in 1X TAE buffer for 30 min at 120 V and 400 mA. On completion of the run, the gel was visualized with the gel-imaging system (Infinity-3000; Vilber Lourmat). The desired DNA sample entrapped in the gel can be cut and extracted as mentioned earlier (3.3.1.4).

The above-mentioned steps are also applicable for running RNA for determining its purity and integrity. RNA (1μg) was mixed with RNase inhibitor (1 μl; 2.4.1) along with 6x loading dye (1 μl; 2.9.4) before loading in the wells to prevent its degradation.

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3.3.7 Procedures for DNA modification

3.3.7.1 Restriction digestion Type II restriction endonucleases (2.4.4) are DNA-modifying enzymes that recognize specific palindromic sequences of lengths 4 to 8 bp. They cut the double-stranded DNA within or close to the recognition sequence, thereby generating products with either blunt or sticky ends. Restriction endonucleases are used to modify vector and insert ends such that they become compatible for interaction in a ligation reaction (3.3.7.3). The restriction sites introduced in the primers during the primer designing were chosen such that they are present in the multiple cloning site (MCS) of the vector but are absent in the insert DNA. The components of the single and double restriction digestion reactions are mentioned in Table 3.10.

Table 3.10: Components of a single and a double restriction digestion reaction

Single digestion reaction Double digestion reaction Component Volume (µl) Component Volume (µl) Insert or Plasmid X (200-500 ng) Insert or Plasmid X (200-500 ng) 10X Reaction buffer 1 10X Reaction buffer 1 Restriction enzyme 0.5-1 Restriction enzyme 1 0.5-1 (10U/μl) (10 U/μl) dH2O Up to 10 Restriction enzyme 2 0.5-1 (10 U/μl) dH2O Up to 10

The plasmid pJET2.1 (2.2) served as the cloning vector for the work and pRSETB (2.2) was chosen for expression study. For the construction of the pJET2.1 vector, the plasmid was digested with EcoRV leading to the production of blunt ends. The linearized plasmid was dephosphorylated to prevent recircularization in the ligation reaction. It then served as a vector for ligating blunt-ended PCR products. The pRSETB plasmid was linearized by restriction enzymes whose restriction digestion sites were present in the MCS but were absent within the insert. It was also subjected to dephosphorylation reaction to prevent recircularization of the linearized plasmid during ligation reaction. The digestion reactions for the screening of plasmids for the presence of the desired insert were incubated for 1 h at 37˚C, while the vector and insert digestion reactions were incubated for 3 h at 37˚C.

Double digestion of DNA can be done in either one step or in two steps depending upon the compatibility of the enzymes and the reaction buffers used. This can be checked using the DoubleDigest Calculator (Thermo Scientific).

3.3.7.2 Dephosphorylation of the digested vectors It is a common step in cloning procedure to ensure that an empty linearized vector does not recircularize during ligation reaction. Removal of the 5' phosphate group using the FastAP Thermosensitive alkaline phosphatase reduces the chance of vector re-closure by intramolecular

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Methods ligation, hence leading to a reduction in background colonies after transformation. Reaction components of a dephosphorylation reaction are shown in Table 3.11. The contents of the reaction were mixed well, briefly centrifuged and incubated at 37˚C for 10 min. The reaction was stopped by the deactivation of the enzyme at 75˚C for 5 min. For the ligation of an insert with a dephosphorylated linearized vector, the insert should possess a 5' phosphate group.

Table 3.11: Components of a dephosphorylation reaction

Component Volume (μl) Linear DNA (Vector) X (1 μg) 10X reaction buffer for Alkaline Phosphatase 2 FastAP Thermosensitive Alkaline Phosphatase 1 (2.4.5) dH2O Up to 20

3.3.7.3 Ligation Ligation reaction is catalyzed by T4 DNA ligase which leads to a phosphodiester bond formation between the 5' phosphate group of one DNA fragment and the 3' hydroxyl group of the other. The required cofactors for the reaction are ATP and Mg2+. T4 DNA ligase carries out sticky and blunt end ligations and is capable of filling nicks in double-stranded nucleic acids. The components of a standard ligation reaction are shown in Table 3.12.

Table 3.12: Components of a standard ligation reaction

Component Volume (μl) Insert 6 Linearized vector 2 10X ligation buffer 1 T4 DNA ligase (2.4.5) 0.5 dH2O Up to 10

Ideally, the insert to vector molar ratio should be 3:1, however in the course of the study it ranged between 2:1 and 5:1. The ligation reaction was incubated at 4˚C overnight and 5 µl was either used immediately for transformation or was stored at -20˚C.

3.3.8 Gateway cloning technology for cloning of localization constructs It is a cloning strategy based upon the bacteriophage lambda site-specific recombination which enables the movements of the gene of interest flanked with attachment (att) sites in multiple vectors without disturbing the orientation and the reading frame of the sequence, thus allowing their use in protein expression or functional analysis. The reaction components include the DNA recombination sites (att sites) and the enzymes that facilitate the recombination reaction (ClonaseTM enzyme mix). Most of the Gateway vectors contain a unique cassette flanked by att

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Methods sites that leads to an efficient selection of recombinant clones. The cassette contains the ccdB gene for negative selection and chloramphenicol resistance gene (CmR) for counter selection. Recombination reactions that form the basis of the technology are BP and LR reactions (Fig. 3.1). Following BP and LR recombination reactions, the cassette containing ccdB and CmR genes are swapped by the gene of interest, producing entry clones and expression clones, respectively.

BP reaction: In this reaction, recombination occurs between an attB flanked substrate (attB flanked PCR product) and an attP flanked substrate (donor vector) to generate an attL flanked product (entry clone) in presence of the enzyme BP ClonaseTM (2.4.3). The components of a BP reaction are mentioned in Table 3.13. Once the components were mixed, 2 μl BP ClonaseTM enzyme mix (2.4.3) was added. The resulting reaction mix was vortexed twice (2 s each) and subsequently incubated at 25˚C for 2 h. Later, 1 μl proteinase K (2 μg/μl; 2.4.3) was added and the reaction mix was incubated at 37˚C for 10 min. The BP reaction product can be immediately used for transformation of competent E.coli DH5α cells or be stored at -20˚C up to 1 week. The transformation was done as mentioned in section 3.4.3.1. The transformed cells were plated and selected on low salt LB agar plates containing 50 μg/ml ZeocinTM. A low salt concentration LB media is required for ZeocinTM to be active. After the selection of positive transformants, plasmid isolation was done as mentioned in section 3.3.1.2. Positive entry clones can be sequenced using M13 sequencing primers.

Table 3.13: Components of a BP reaction

Component Volume (μl) attB PCR product X (150 ng) pDONRTM/Zeo (2.2) Y (150 ng) TE buffer (pH 8)* Up to 8 *10 mM Tris-HCl, pH 8.0, 1 mM EDTA

LR reaction: In this reaction, recombination occurs between an attL flanked substrate (entry clone) and an attR flanked substrate (destination vector) to create an attB product (expression clone) in presence of the enzyme LR ClonaseTM (2.4.3). Components of the LR reaction are mentioned in Table 3.14. LR ClonaseTM (2 µl; 2.4.3) enzyme mix was added to the reaction and shortly vortexed twice (2 s each). The resulting reaction mix was incubated at 25˚C for 3 hours. Post-incubation 1 μl proteinase K (2 μg/μl; 2.4.3) was added to the reaction mix and the same was incubated at 37˚C for 10 min. The LR reaction generated could be used for transformation (3.4.3.1) of competent E.coli cells (DH5α) or be stored at -20˚C up to a week. The transformants were plated and selected on LB agar plate containing 50 μg/ml kanamycin. Positive transformants selection and growth was followed by expression clone isolation as mentioned in section 3.3.1.2. The presence of the gene of interest in expression clone was verified by setting up a standard PCR (3.3.5.1) containing expression clone as the template and gene-specific forward and reverse primers.

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Table 3.14: Components of a LR reaction

Component Volume (μl) Entry clone X (150 ng) Destination vector (pEarleyGate 104/ pEarleyGate 101; Y (150 ng) 2.2) TE buffer (pH 8)* Up to 8 *10 mM Tris-HCl, pH 8.0, 1 mM EDTA

Since the expression clones were to be used in subcellular localization experiments to visualize the fluorescently labeled protein, they were also sequenced using YFP-specific primers designed close to the junction of YFP sequence and gene of interest. Both donor vector and destination vector were propagated and maintained in DB3.1TM E. coli competent cells (2.1.2) that are resistant to the lethal effect of CcdB protein encoded by the ccdB gene.

GOI GOI BP clonase attBPCR product Entry clone + +

ccdB-CmR ccdB-CmR

Donor vector By-product (a)

GOI GOI

Entry clone LR clonase Expression clone + +

ccdB-CmR ccdB-CmR

Destinationvector Donor vector (b)

attB1 attB2 attP1 attP2 attL1 attL2 attR1 attR1 Vector backbone

Fig. 3.1: Schematic diagram representing the Gateway Technology. BP reaction (a) LR reaction (b). GOI, Gene of interest; ccdB gene, control of cell death B gene; CmR, Chloramphenicol resistance gene. The ccdB gene is located on the F sex factor plasmid of E. coli and is part of a

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Methods toxin-antitoxin system encoded by the ccd operon, which is responsible for maintenance of plasmid during cell division. 3.3.9 Sequencing DNA samples for sequencing were sent to Eurofins Genomics. As per the specifications of the service providers, the final concentration of the pure recombinant plasmids was 100-150 ng/μl and that of purified PCR products (100-300 bp) was 10-20 ng/μl. Bioinformatic analysis of the sequencing result was done using Lasergene-DNAStar 7.0 (2.11).

3.4 Microbiological methodologies

3.4.1 Determination of the microbial growth The count of E. coli during culture growth was determined using a spectrophotometer by measuring the OD of 1 ml of bacterial culture at 600 nm. Fresh LB medium served as the blank. 8 An OD600 of 0.1 corresponds to ~1×10 cells/ml culture (Lech and Brent, 2001)

3.4.2 Preparation of competent cells

3.4.2.1 Preparation of E. coli competent cells by calcium chloride method This technique was first used by Mandel and Higa (1970) who presented the uptake of phage DNA by bacterial cells in the presence of calcium ion. A LB plate was streaked with frozen glycerol stock (3.4.4) of DH5α under sterile conditions and was incubated overnight at 37˚C. A single E.coli colony was inoculated in 5 ml LB media and was incubated overnight at 37˚C with continuous shaking (200 rpm). Overnight culture (1 ml) served as an inoculum for 50 ml LB media. The OD600 of the resulting preparation was ~0.2-0.3. The culture was grown at 37˚C under continuous shaking (200 rpm) conditions until an OD of 0.6-0.8 was reached. The culture was centrifuged for 5 min at 5000 rpm and 4˚C. The supernatant was discarded and the pellets hence obtained were washed with 50 ml pre-chilled calcium chloride solution (50 mM). Following another round of centrifugation at 5000 rpm for 5 min at 4˚C, the supernatant was discarded. The pellets were resuspended in 20 ml calcium chloride solution (50 mM) and incubated on ice for 20 min. Following incubation, the suspension was centrifuged, the supernatant discarded and the pellets were resuspended in 860 μl 50 mM calcium chloride. The suspension was transferred to a sterile pre-cooled 1.5 ml Eppendorf tube and incubated on ice for 20 min. Sterile glycerol (140 μl) was added to the suspension and mixed gently. The resulting suspension was transferred in pre-cooled 1.5 ml Eppendorf tubes as 50 μl aliquots. The Eppendorf tubes were subjected to snap freezing in liquid nitrogen and stored at -80˚C for future use in transformation protocol (3.4.3.1).

3.4.2.2 Preparation of A. tumefaciens electrocompetent cells A LB plate with rifampicin (100 μg/ml) and gentamycin (100 μg/ml) was streaked with frozen glycerol stock (3.4.4) of A. tumefaciens (C58C1/pMP90) under sterile conditions and was incubated for two days at 28˚C. A single colony of A. tumefaciens was inoculated in 10 ml YEB media containing rifampicin (100 μg/ml) and gentamycin (100 μg/ml) and was incubated

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Methods overnight at 28˚C with continuous shaking at 220 rpm. The overnight culture was transferred in 300 ml of fresh YEB medium supplemented with previously mentioned antibiotics and incubated at 28˚C with continuous shaking until an OD600 of 0.5-0.7 was obtained. The culture was then transferred to six sterile falcon tubes (50 ml) and centrifuged at 5000 rpm for 10 min. The supernatant was discarded and the pellets obtained were washed three times with 25 ml of ice- cold sterile-filtered glycerol (10% v/v). The content of each tube was divided in two sterile falcon tubes. After centrifugation for 10 min at 5000 rpm, the supernatant was discarded; the pellets were resuspended in 2 ml of ice-cold sterile-filtered glycerol (10%) and again centrifuged. After centrifugation the pellets were combined and incubated with 3 ml of 10% glycerol at 4˚C for 24 h. Following incubation, the suspension was divided into aliquots of 50 µl and stored at - 80˚C.

3.4.3 Transformation of competent cells

3.4.3.1 Transformation of E. coli competent cells by heat-shock The ligation (3.3.7.3), BP (3.3.8) and LR (3.3.8) reaction products (5 μl) were added independently to 50 μl aliquot of ice-thawed competent E. coli DH5α (3.4.2.1) and placed on ice for 20 min. Following ice-treatment, the mixtures were subjected to a heat-shock at 42˚C for 45 s. This step was followed by immediate cooling on ice for 5 min. 250 μl SOC medium was added to the transformation mixture and was incubated at 37˚C for 60 min with continuous shaking (220 rpm). After incubation, 250 μl of the DH5α mix was spread on LB plate supplemented with suitable antibiotics. For the selection of recombinants with pJET1.2 vector backbone, ampicillin (100 μg/ml) containing plates were used. Selection of recombinants with entry clone and expression clone was done on plates carrying ZeocinTM (50 μg/ml) and kanamycin (50 μg/ml), respectively. The plates were incubated at 37˚C overnight. For ZeocinTM-based selection, LB plates with half salt content (0.5% NaCl) were used.

For the transformation of competent cells with the recombinant expression plasmid (pRSETB), 1 μl of the plasmid (~50-100 ng) was transferred to 50 μl aliquot of competent E. coli BL21 cells (3.4.2.1) and incubated on ice for 20 min. The mix was subjected to a heat-shock treatment at 42˚C for 20 s followed by immediate cooling on ice for 5 min. 250 μl SOC medium was added to the mix and was incubated at 37˚C for 60 min with continuous shaking (220 rpm). An aliquot of 80 μl was spread on LB plate supplemented with ampicillin (100 μg/ml) and chloramphenicol (30 μg/ml). The plates were incubated at 37˚C overnight.

3.4.3.2 Transformation of A. tumefaciens by electroporation For electroporation 5 μl DNA (2-10 ng/μl) was added to 50 μl A. tumefaciens electrocompetent cells (3.4.2.2) placed on ice. The tube was gently tapped to mix the contents. The mix was transferred to a pre-cooled electroporation cuvette (1 mm cuvette, 2.2 kV; E. coli Pulser cuvette, Bio-Rad) such that the contents settled at the bottom of the cuvette. The cuvette was then placed in the chamber slide and pushed in the shocking chamber until it made contact with the electrodes. A pulse (2.5 kV, 400 Ώ, and 25 μF) was given using MicroPulser (Bio-Rad). The

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Methods cuvette was withdrawn from the chamber and mixed with 1 ml antibiotic-free YEB medium. The mix was subsequently transferred to 10 ml of YEB medium and incubated at 28˚C for 2 h with continuous shaking at 220 rpm. Various dilution of the Agrobacterium culture was then spread on YEB agar plates augmented with gentamycin, rifampicin and kanamycin at concentrations 100 μg/ml, 100 μg/ml and 50 μg/ml, respectively. The plates were then incubated at 28˚C for 2 days.

3.4.4 Preparation of stock cultures A single positive colony of transformed E. coli and A. tumefaciens was inoculated in 10 ml LB and YEB media containing suitable antibiotics, respectively. The overnight grown culture (750 µl) was mixed well with 250 μl of sterile LB:glycerol or YEB:glycerol (40: 60 v/v) and stored at -80˚C in 2 ml cryo tubes.

3.5 Biochemical methodologies

3.5.1 Heterologous expression in E. coli An E. coli BL21 colony containing the desired expression plasmid was inoculated in 10 ml of LB medium containing 100 μg/ml ampicillin and 30 μg/ml chloramphenicol and was incubated at 37˚C overnight with continuous shaking at 220 rpm. 4 ml of the overnight culture was inoculated into 100 ml of the LB medium and incubated at 37˚C with continuous shaking (220 rpm) until an OD600 0.6–0.8 was reached. 1 ml of the culture was retained and centrifuged at 5000 rpm for 5 min to collect the before-induction pellets for SDS-PAGE (3.5.4) analysis. 0.5 mM sterile-filtered IPTG was added to the remaining culture and was grown overnight at 25˚C with continuous shaking at 150 rpm. 1 ml culture was again retained to get the after-induction pellets for SDS-PAGE analysis. The cells were harvested by centrifugation at 5000 rpm for 5 min at 4˚C. The pellets could be used immediately for downstream processing or could be stored at -20˚C.

3.5.2 Extraction and purification of the hexahistidine (His6)-tagged recombinant protein The fresh or frozen pellets were resuspended in pre-cooled 6 ml lysis buffer (2.9.6) and were disrupted by sonification on ice for 6 min at 30% pulse using a Branson sonifier B15. The slurry thus obtained was centrifuged at 14000 rpm for 20 min at 4˚C. The supernatant containing the crude soluble proteins was collected in a 15 ml falcon tube for protein purification by affinity chromatography. An aliquot of 50 μl was retained for SDS-PAGE (3.5.4) analysis.

Purification of His6-tagged protein was done using the immobilized metal affinity chromatography (IMAC) employing the Ni-Nitrilotriacetic acid (NTA) agarose (Qiagen). Ni2+ ion has six complex ligand binding sites. Four sites are bound to NTA which in turn is linked to sepharose matrix. The remaining two sites interact with the histidine residues of the His6-tagged protein. The protein can be eluted either by lowering the pH of the elution buffer (~pH 5) as the protonated nitrogen in histidine residue can no longer bind to Ni2+ or by adding imidazole in the elution buffer which competes with histidine residue for Ni2+ due to structural

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2+ similarity. Imidazole (250 mM) displaces the His6-tagged protein, which was bound to Ni . Elution with imidazole is a better method for elution of protein as lowering the pH can affect protein stability and activity. A low concentration of imidazole is also added to lysis and washing buffer to prevent the binding of non-specific proteins.

Ni-NTA agarose (100 µl suspension) was added to 6 ml of the clear crude and soluble protein lysate. After shaking at 4˚C for 1 h, the mixture was loaded on an empty PD-10 column and the gel matrix was retained. Once the solution had passed through the column, the agarose with the bound protein was washed with 6 ml of washing buffer (2.9.6). The His6-tagged fusion protein was then eluted with 2.5 ml elution buffer (2.9.6). High imidazole concentration that was used for elution of tagged protein was removed from the purified protein by gel filtration (3.5.3). An aliquot of 50 μl pure protein was kept for SDS-PAGE (3.5.4) analysis.

3.5.3 Gel filtration chromatography In order to free the soluble protein fraction from low molecular weight molecules, it was subjected to gel filtration chromatography. This technique employs porous matrix and steric factors for the separation of proteins based upon their molecular size. Large proteins which cannot access the pores of the matrix are eluted first. Smaller molecules like salts which have a greater degree of access to the porous matrix are eluted later. A PD-10 (Sephadex G-25, Amersham Biosciences) column was equilibrated with 25 ml desalting buffer. The protein solution (2.5 ml) was loaded on the column. When the solution was completely immersed into the separation bed, 3.5 ml of desalting buffer (100 mM Tris-HCl pH 9 for HcBD and 100 mM KH2PO4, pH 7 for HcBZL) was added to elute the protein. For regeneration of the PD-10 column, 25 ml of 0.16 M NaOH was added followed by washing with distilled water until the pH of the eluate was neutral.

3.5.4 Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) It is a procedure used for the separation of proteins based solely on their molecular weight in a cross-linked matrix under the influence of an electric field. SDS is an anionic detergent that imparts a uniform negative charge to the protein there by linearizing and denaturing it. Charge and structure of the protein thus do not influence the rate of protein migration when an electric field is applied. PAGE is a discontinuous system for separation of proteins, comprising of a stacking gel (5%, pH 6.8) and a resolving gel (12%, pH 8.8). When power is supplied to the electrophoretic system the protein-SDS complex forms a crisp layer between the leading chloride ions and the trailing glycine thereby getting stacked in the stacking gel before separating in the resolving gel/separating gel. When in resolving gel, proteins migrate at different rates because of the sieving properties of the gel. Smaller protein-SDS complexes migrate faster than the larger protein-SDS complexes. The polyacrylamide gel is composed of acrylamide and bisacrylamide which copolymerize and make a 3D network of straight chain of acrylamide interconnected by bisacrylamide bridges. Copolymerization occurs by a free radical mechanism where APS is the source of the sulfate free radicals and TEMED acts as a catalyst for the formation of sulfate free radicals.

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For gel preparation, all the components of resolving gel/separating gel (2.9.5) were mixed together in a flask, TEMED being the last component to be added. The contents were gently mixed, poured in between the glass plates of the setup, covered with a layer of water to avoid contact with air and allowed to set. After solidification of the resolving gel, water was removed and the components of the stacking gel (2.9.5) were mixed. The stacking gel was poured over the resolving gel, the comb was inserted and the gel was allowed to solidify for at least 20 min. The protein samples to be analyzed by SDS-PAGE were mixed with protein loading dye (2.9.5) and denatured at 95˚C for 10 min. Samples were loaded on the gel and run at 200 V. The run was tracked by visualizing bromophenol blue dye and was ended once the dye front reached the front of the gel. The gel was then stained by immersing in the staining solution (2.9.5) for 30 min with gentle shaking, followed by destaining overnight in a destaining solution (2.9.5).

3.5.5 Determination of protein concentration using the method of Bradford (1976) The Bradford assay is a dye-based assay used for the determination of the total protein concentration in a sample. When an acidic solution of Coomassie Brilliant Blue G-250 binds to the protein, there is a shift in its absorption maximum from 465 nm to 595 nm. The increase in absorbance at 595 nm is recorded using a UV/VIS spectrophotometer. For the determination of protein concentration, two different assays were set up as mentioned in Table 3.15.

Table 3.15: Set up for a Bradford assay

Components Blank assay Test assay Test sample 5 μl (Buffer in which the protein 5 μl protein was eluted) Bradford solution 900 μl 900 μl (2.9.7) Deionized water 95 μl 95 μl Final volume 1000 µl

Addition of components, gentle vortex and incubation at room temperature for 5 min was followed by the measurement of the optical density of the assays at 595 nm. A blank assay lacking the test protein was set up to control the changes in absorbance caused by a change in the pH of the solution or because of the presence of salts.

The equation used for the calculation of the protein concentration is as follows:

where C is the concentration of the test protein (μg/μl)

OD595 is the absorbance of the test assay recorded at 595 nm after normalization with respect to the blank assay,

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Methods

V is the volume of the test protein (μl) used in the test assay,

S is the slope of the calibration curve (1/μg) that was prepared by plotting the varying amount of BSA in μg (X-axis) against the corresponding optical density measured at 595 nm (Y-axis).

3.5.6 ALDH assay The components of the standard ALDH assay are shown in Table 3.16.

Table 3.16: Components of an ALDH assay for HPLC and spectrophotometric analyses

Component Stock Volume in 200 μl assay Volume in 1000 μl assay Aldehyde 100 mM 1 5 NAD+ 50 mM 4 20 Protein X (1.5 µg) X (7.5 µg) Buffer 100 mM Tris-HCl pH 8 for substrate specificity determination and enzyme characterization. 100 mM Tris-HCl pH 9.5 for enzyme kinetic study

3.5.7 Analysis of ALDH assay incubation 3.5.7.1 Incubations to be analyzed by HPLC

Addition of all components to the assay was followed by short vortex and initial incubation at 37˚C for 15 min. After optimization of the assay conditions, incubation was done at 50˚C which led to maximum enzyme activity. The reaction was terminated by the addition of 20 μl ice-cold stop solution (acetic acid: methanol; 4:1) and vortexing. The enzymatic product was mixed with 200 μl ethyl acetate, vortexed and centrifuged at 14000 rpm for 5 min. The organic phase (150 μl) was collected and the procedure was repeated. In the second extraction procedure, 200 μl of the organic phase was collected. The total organic phase was pooled and air-dried. The residue was dissolved in 50 μl methanol (HPLC grade), 20 μl of which was injected for product analysis by HPLC. For preliminary HPLC-based substrate specificity assay, 6 μg of protein was used in a 200 μl assay. For each experiment conducted with HcBD protein, both a control assay with denatured protein and a standard assay with active protein were prepared.

3.5.7.2 Incubations to be analyzed by spectrophotometer

All the reaction components except the protein were added in a 1 ml cuvette, mixed well and then blanked at 340 nm. The reaction was initiated by the addition of the protein. The change in absorbance was recorded over 5 min and represented the enzymatic conversion of NAD+ to NADH as the reaction proceeded.

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3.5.8 CoA ligase assay A standard CoA ligase assay consisted of the components mentioned in Table 3.17.

Table 3.17: Components of a CoA ligase assay for HPLC- and spectrophotometer-based analyses

Component Stock (mM) Volume in a 125 μl assay Volume in a 1000 μl assay Acid 10 5 40 MgCl2 125 2.5 20 ATP 125 2.5 20 CoA 20 2.5 20 Protein X (2.5 μg) X (10 µg) Buffer 100 mM KH2PO4 buffer pH 6.5

3.5.8.1 Incubations to be analyzed by HPLC Addition of the standard components of the assay mix was followed by gentle vortex and incubation at 30˚C for 20 min. The reaction was terminated by the addition of ice-cold stop solution (12.5 μl; acetic acid: methanol; 4:1), vortexed and centrifuged at 14000 rpm for 10 min. The aqueous phase was collected and 20 μl of the same was injected for product analysis by HPLC.

3.5.8.2 Incubations to be analyzed by spectrophotometer The enzyme activity was measured spectrophotometrically at room temperature. All the components of the assay were added except for CoA, mixed well and blanked. The assay was initiated by the addition of CoA. The formation of CoA esters was recorded as change in the absorption monitored for 10 min at the wavelengths of 311, 333, 345, 346 and 352 nm. The wavelengths correspond to the absorption maxima of cinnamoyl-CoA, 4-coumaroyl-CoA, feruloyl-CoA, caffeoyl-CoA, and sinapoyl-CoA, respectively (Stöckigt and Zenk, 1975; Lee et al., 1997).

3.5.9 Biochemical characterization of HcBD and HcBZL For biochemical characterization of HcBD and HcBZL enzymes, pure recombinant proteins were utilized. Various factors (pH, temperature, time, protein amount, cofactors, etc.) affect the enzyme activity and conditions required for obtaining maximum product formation in the assay must be identified. pH affects the protonation and deprotonation of ionisable groups in the of the enzyme and the cofactors in the assay affect enzyme activity. Extreme pH causes irreversible denaturation of enzymes. Temperature affects the velocity of the reaction as well as the native structure of the protein. Optimum temperature should be the one where high enzyme activity can be achieved without promoting enzyme denaturation. In most cases the optimum temperatures are 25˚C, 30˚C or 37˚C. Higher temperature not only adversely affects enzyme conformation and stability by breaking weak hydrogen and ionic bonds but also affect the stability of other components in the assay mix. Amount of enzyme and time influence product

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Methods accumulation, however, for studying kinetic parameters of the enzyme these factors should be chosen wisely to obtain a linear correlation with reaction velocity (Bisswanger, 2014).

The various conditions tested to determine the optimum parameters for the enzyme activity of HcBD and HcBZL are mentioned in Table 3.18.

Table 3.18: Various parameters tested for optimization of enzyme activity

HcBD Parameter Conditions pH 6.5 to 12 100 mM potassium phosphate buffer pH 6.5 to 7.5 100 mM Tris-HCl buffer pH 7 to 12 Temperature 25˚C to 60˚C Protein amount 1 to 45 μg Incubation time 0 to 20 min Substrate specificity Tested substrates included various aromatic aldehydes and the aliphatic aldehyde acetaldehyde Cofactor NAD+, NADP+, FAD (Final concentration 1 mM) Divalent cation Mn2+, Mg2+, Co2+, Zn2+, Ni2+, Ca2+, Fe2+, Cu2+ (Final concentration 1mM) + + + Univalent cation K , Na , NH4 (Final concentration 1 mM) DTT 0 to 1000 µM Storage condition Storage temperature -4˚C, -20˚C and -80˚C (1 day) Storage time (-80˚C) 80 days HcBZL Parameters Conditions pH 5 to 10 100 mM potassium phosphate buffer for the pH range 5 to 7.5 100 mM Tris-HCl buffer for the pH range 7.5 to 10 Protein amount 0.5 to 100 μg Incubation time 0 to 180 min Substrate specificity A range of substrates was tested that included benzoic acid and its hydroxylated and amino derivatives, trans-cinnamic acid and its hydroxylated and methoxylated derivatives, fatty acids and miscellaneous substrates Temperature 20 to 60 ˚C Divalent cation Mn2+, Mg2+, Co2+, Ni2+, Ca2+, Fe2+, Cu2+ (Final concentration 2 mM) The optimum amount of Mn2+ and Mg2+ required in the assay was tested over a range of 1μM to 10 mM + + + + Univalent cation K , Na , NH4 , Li (Final concentration 2 mM) DTT 0 to 1000 μM Storage condition Storage temperature -4˚C, -20˚C and -80˚C (1 day) Storage time (-80˚C) 30 days

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3.5.10 Assay for optimization of luciferase protein amount A 200 µl assay containing 100 mM Tris-HCl buffer (pH 7.8), 0.15 mM D-luciferin, 500 nM ATP, 2.5 µM MgCl2 and varying amounts of recombinant luciferase was used to determine the optimum luciferase amount required for luciferase-based substrate specificity assay. In parallel an assay with the commercial luciferase (Sigma) was also conducted. Photon emission was measured for 10 s by the luminometer (Victor2 multilabel counter; Wallac 1420) and expressed as counts per second.

3.5.11 Luciferase-based substrate specificity assay for CoA ligases The luciferase-based substrate specificity assay is a versatile and sensitive assay which is suitable for screening various CoA ligases of unknown catalytic efficiency and activity optimum with a vast array of potential substrates. The sensitivity of the luciferase-based assay can be easily modulated by changing the duration of incubation with the CoA ligase under study (Schneider et al., 2005). The components of a luciferase-based substrate specificity assay are mentioned in Table 3.19.

Table 3.19: Components of the luciferase-based substrate specificity assay

Substrates Stock solutions Volume in 100 µl assay (mM) Aromatic acid 10 2 ATP 10 0.5 MgCl2 10 2.5 CoA 10 1 DTT 130 0.75 Protein (HcBZL) X μl (1 μg) Buffer 100 mM KH2PO4 buffer pH 6.5

After adding all components, the reaction mixture was incubated at 37˚C for 120 min. For ATP determination, 2 µl of the aforementioned reaction mixture was taken, diluted to 100 µl with 100 mM Tris-HCl buffer (pH 7.8) and pipetted onto a 96-well flat-bottom microtiter plate (Eppendorf). The plate was then placed at the measurement position in a luminometer. A second reaction mixture (100 µl) containing 0.5 µg of P. pyralis luciferase, D-luciferin (4.6 µg) and 100 mM Tris-HCl (pH 7.8) was injected by the instrument into each sample in the microtitre plate. Photon emission was measured for 10 s by the luminometer. The data (counts per second) obtained on analysis was represented as relative luciferase activity. ATP determination for all samples was normalized with respect to an assay mix containing 4-coumaric acid and Rhodopseudomonas palustris BZL (RpBZL; Geissler et al., 1988; Egland et al., 1995), in which no ATP depletion occurred and thus the relative luciferase activity was set to 100%.

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3.5.12 Agroinfiltration and transient expression in N. benthamiana The two consecutive steps were carried out for studying the sub-cellular localization of HcBD and HcBZL in the plant cell. Transformed agrobacteria containing the HcBD and HcBZL expression clones were grown for 2 days at 28˚C on YEB-agar medium supplemented with kanamycin (50 μg/ml), rifampicin (100 μg/ml) and gentamycin (100 μg/ml). A single colony was inoculated in 10 ml YEB medium supplemented with the same antibiotics as mentioned earlier and grown ~30-36 h at 28°C with continuous shaking (220 rpm). The cultures were later centrifuged at 5000 rpm for 15 min at room temperature. The supernatant was discarded and the cells were resuspended in the activation medium (2.9.3) such that the OD600 of the resulting suspension was 1. A. tumefaciens helper strain (Voinnet et al., 2003) was also grown under similar conditions except that the selection was made on kanamycin (50 μg/ml) and rifampicin (100 μg/ml). After centrifugation, the pellets were resuspended in activation medium such that the final OD600 was 1. Prior to agroinfiltration, agrobacteria cultures carrying HcBD and HcBZL fusion constructs were mixed with helper strain in a 1:1 ratio (v:v) to improve the efficiency of transformation. The mixed suspensions were then incubated at 28˚C for 2 h with continuous shaking (100 rpm). The suspensions were then infiltrated on the lower surface of 4-6-week-old N. benthamiana leaves using a 1 ml needle-free syringe. 3-4 leaves per plant were infiltrated with the desired suspension and analyzed 2-4 days post-infiltration. Leaves expressing HcBD fusion protein were analyzed 3 days post-infiltration.

For co-localization experiments, full-length HcBZL and truncated-HcBZL constructs were cultivated in 10 ml YEB medium supplemented with kanamycin (50 μg/ml), rifampicin (100 μg/ml) and gentamycin (100 μg/ml) at 28˚C for 30-36 h with continuous shaking (220 rpm). A. tumefaciens harboring the peroxisomal marker (Nowak et al., 2004) was cultivated in YEB medium with kanamycin (50 μg/ml) and rifampicin (100 μg/ml). A. tumefaciens harboring eqFP611 (peroxisomal and cytoplasmic marker) construct was cultivated in YEB medium with kanamycin (50 μg/ml), rifampicin (100 μg/ml) and streptomycin (100 µg/µl). All the suspensions were incubated till a final OD600 of 1 was reached. Agrobacteria carrying HcBZL expression clones and those with the marker constructs were mixed in varying ratios, incubated and infiltrated in N. bentamiana leaves as mentioned earlier.

3.6 Analytical methods

3.6.1 High-performance liquid chromatography

3.6.1.1 Method used for the analysis of xanthone-rich extracts The methanolic extracts of H. calycinum cells and the media were analyzed by the method in which the mobile phase consisted of two solvents: water acidified with formic acid (1 mM; A) and methanol (B). The gradient program started with 50% B for 2 min, 50% to 75% B in 13 min, 75% to 90% B in 20 min, and ended with 90% to 100% B in 1 min. The flow rate was 0.5 ml/min. The detection wavelengths were set at 254 and 292 nm for the detection of hyperxanthone E- and patulone-like products, respectively.

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3.6.1.2 Method used for the analysis of product formation catalyzed by HcBD Different gradient methods were used for the analysis of products whose formation was catalyzed by HcBD. The information on the solvents used, gradients applied and the detection wavelengths are mentioned in Table 3.20.

Table 3.20: HPLC-DAD methods and detection wavelengths used for the quantification of various enzymatically formed acids

Gradient 1 Gradient 2 Gradient 3 Gradient 4 Time A B Time A C Time A B Time A C 0 60 40 0 75 25 0 80 20 0 98 2 5 55 45 5 75 25 5 80 20 5 98 2 17 55 45 10 55 45 8 70 30 8 60 40 20 0 100 20 0 100 15 70 30 15 40 60 20 0 100 20 0 100 A, Acidified water (1 mM formic acid); B, Methanol; C, Acetonitrile. Time is expressed in minutes. Flow rate = 0.7 ml/min Gradients Detected aromatic acids Detection wavelengths (nm) 1 trans-Cinnamic acid 280 2-Hydroxybenzoic acid (salicylic acid) 295 2 Benzoic acid 230 4-Hydroxy-3-methoxycinnamic acid (ferulic acid) 322 3 4-Hydroxybenzoic acid 254 2-Methoxybenzoic acid (2-anisic acid) 295 3,4-Dihydroxybenzoic acid (protocatechuic acid) 254 4-Hydroxy-3-methoxybenzoic acid (vanillic acid) 254 4 3-Hydroxybenzoic acid 295

3.6.1.3 Method used for the analysis of product formation catalyzed by HcBZL Product formation catalyzed by HcBZL was analyzed by HPLC using a method in which the mobile phase consisted of 10 mM ammonium acetate in water (pH 5.6; solvent A) and 100% acetonitrile (solvent B). The gradient was at a flow rate of 0.5 ml/min and started with 5% B for 2 min, 5% to 10% B in 13 min and ended with a gradual increase to 90% B in 10 min. The detection wavelength was 261 nm for benzoyl-CoA, propanoyl-CoA, butyryl-CoA, isobutyryl- CoA, hexanoyl-CoA, heptanoyl-CoA, and octanoyl-CoA.

3.6.1.4 Calibration curves For quantification of enzymatic reaction products formed by HcBD, standard curves were prepared for authentic references such as benzoic, 2-hydroxybenzoic, 3-hydroxybenzoic, 4- hydroxybenzoic, 3,4-dihydroxybenzoic, 4-hydroxy-3-methoxybenzoic, 2-methoxybenzoic, trans-cinnamic and 4-hydroxy-3-methoxycinnamic acids. Calibration curves were generated by

57

Methods injecting the compounds dissolved in methanol (HPLC grade) in quantities ranging from 5 to 500 ng at the HPLC and documenting the area under the curve (AUC) at the detection wavelengths mentioned in Table 3.20. Each point on the calibration curve is the mean of 2-3 repeats. The coefficient of determination (R2) was > 0.99 (Appendix Fig. A3).

Similarly, the calibration curves for xanthones and CoA thioesters were prepared. In order to quantify hyperxanthone E- and patulone-like xanthones, the standard references were dissolved in methanol, injected at the HPLC as dilutions ranging from 200-1400 ng and the AUC was reported at 254 nm for hyperxanthone E-like xanthones and at 292 nm for patulone-like xanthones. Each data point on the calibration curve is the mean of two repeats and the R2 values were > 0.99 (Appendix Fig. A2). Benzoyl-CoA and isobutyryl-CoA dissolved in methanol were injected as dilutions ranging from 10-1500 ng. Each data point is presented as the mean of the AUC from 2-3 repeats observed at 261 nm. The R2 values of the linear equations were > 0.99 (Appendix Fig. A4).

3.6.2 Liquid chromatography-Mass spectrometry (LC-MS)

3.6.2.1 Sample preparation A large scale (5 ml) CoA ligase assay was carried out for ESI-MS analysis of the formed benzoyl-CoA. The final concentration of the enzyme, main substrates, and the co-substrates was kept the same as in the standard assay. Tris-HCl buffer pH 7.5 was used and the incubation time was increased to 120 min. The product formed was purified using solid-phase extraction cartridges (1000 mg, C18 polypropylene column, Chromabond Macherey-Nagel) as described previously (Beuerle and Pichersky, 2002a). Fractions containing the unreacted free CoA and benzoyl-CoA were identified spectrophotometrically by the high UV absorption at 261 nm on rinsing the cartridges with 4% ammonium acetate and water, respectively. The benzoyl-CoA-rich fractions eluted with water were pooled and lyophilized overnight. The lyophilized product was dissolved in 1 ml methanol (LC-MS grade). For product analysis of non-aromatic CoA esters, the standard enzymatic incubations were stopped and centrifuged at 14000 rpm for 10 min. The supernatants were collected and injected at the HPLC for the semi-preparative purification of the CoA ester product.

3.6.2.2 Sample analysis ESI-MS analysis of purified enzymatic products and xanthones was done with the help of Dr. Till Beuerle, Institute of Pharmaceutical Biology. The samples (~10 µg/ml) were analysed by 3200 QTrap massspectrometer (Applied Biosystems/MDS SCIEX, Darmstadt, Germany), equipped with an electrospray ionization (ESI) interface (Turbo V). The samples were injected with the integrated 3200 QTrap syringe pump (Syringe; 1,000 ml, i.d. 2.3 mm; Hamilton, Nevada, USA) at a flow rate of 5 or 10 µl/min. The mass spectrophotometer was operated in negative mode with declustering potential being -20 to -80 V and -30 V for analyses of thioesters and xanthones, respectively. Nitrogen gas was used for nebulization, with the curtain gas, gas 1, and gas 2 settings ranging from 10-20, 10-20 and 0-20, respectively for the analysis of thioesters.

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Methods

For xanthone analysis the curtain gas, gas 1, and gas 2 settings were 20. The molecular ion peaks [M-H]- of the products were further analyzed by MS/MS experiments in the enhanced product ion (EPI) mode of the instrument using nitrogen gas for collision-induced dissociation at high- level setting. The collision energy was -20 to -55 eV for thioesters and -10 to -25 eV for xanthones. Data procurement and assessment were done with the Analyst Software (Version 1.6.2; Applied Biosystems/ MDS SCIEX).

3.6.3 Gas chromatography-Mass spectrometry (GC-MS) Assays containing recombinant HcBD enzymes were analyzed via GC-MS with the help of Dr. Till Beuerle for the presence of benzoic and trans-cinnamic acids. Incubations (1 ml) were prepared to contain the same final concentration of substrate as in the standard assay (3.5.6). Accordingly, the protein concentration was 30 μg/ml of incubation volume. The enzymatic reaction was stopped by the addition of 100 μl of 2 M hydrochloric acid followed by extraction with two volumes of dichloromethane. The organic phase was collected in HPLC glass vials and evaporated to dryness. The residue was subjected to trimethylsilyl (TMS) derivatization of the acidic protons. MSTFA (40 µl) was added to the residue and incubated at 70˚C for 45 min. Before starting the GC-MS analysis, the concentration of the samples was adjusted to 0.1 mg/ml assuming a 50% conversion of the substrate. The GC-MS analysis was carried out on an Agilent 6890 gas chromatograph equipped with a ZB5MS column (30 m long, 0.25 mm i.d., 0.25 µm ft). Injector and transfer line were set at 270˚C and 290˚C, respectively. The thermal program used was 70˚C for 3 min followed by an increase in temperature from 70˚C to 310˚C over a period of 24 min at the rate 10˚C/min and finally 310˚C for 5 min. The split ratio was 1:10, the injection volume was 1 µl and the carrier gas (helium) flow was 1 ml/min via Agilent 5975B MSD (Agilent, Waldbronn, Germany).

3.6.4 Phylogenetic analyses The amino acid sequences of various functional plant ALDH family 2 (ALDH2) members and acyl-activating enzymes (AAEs) were retrieved from NCBI (https://www.ncbi.nlm.nih.gov/). Their accession numbers are listed in Appendix Tables B 1 and B 2, respectively. Multiple sequence alignment was done using the MUSCLE tool integrated into MEGA 7 (Kumar et al., 2016). The phylogenetic trees (Cladograms) were generated by the neighbor-joining method (Saitou and Nei, 1987) using MEGA 7. Rattus norvegicus ALDH2 (Accession number: P11884.1) and P. pyralis luciferase (Accession number: AAA29795.1) served as out-groups for rooting the ALDH and CoA ligase phylogenetic trees, respectively. Statistical robustness was ensured via the bootstrap test with 1000 replications (Felsenstein, 1985). The evolutionary distances were calculated using the Poisson correction method (Zuckerkandl and Pauling; 1965) and were represented in the units of the number of amino acid substitutions per site. The pairwise deletion method was used for managing missing data and nullifying gaps (Gaid et al., 2012).

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Methods

3.6.5 Laser scanning microscopy Infiltrated N. benthamiana leaf discs were analyzed for YFP, CFP, or eqFP611 expressions by confocal laser scanning microscopy (cLSM-510META; Carl Zeiss) 2-4 days after infiltration. The samples were examined using the Plan-Neofluar 10x/0.3 for an overview and the C- Apochromat 40x/1.2 water-immersion objective for detailed analysis. Excitation of fluorophores was accomplished using argon laser (458 nm). The emitted light passed the primary beam splitting mirror UV/488/543/633 nm, followed by separation with a secondary beam splitter at 545 nm. The emitted fluorescences were detected with band pass filters at 475-525, 505-530 and 560-615 nm for CFP, YFP, and eqFP611, respectively. The autofluorescence of chlorophyll was detected at 650 nm via a long pass filter (Gehl et al., 2009). The spectral signatures were examined with optimal detection at 485, 528 and 611 nm of the aforesaid fluorophores in the lambda mode, respectively. The images were analyzed with Zen 2.3 (Carl Zeiss).

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Results

4 Results

4.1 Changes in xanthone content of H. calycinum cell cultures upon elicitor treatment H. calycinum cell cultures propagated in LS medium respond to elicitor treatment by the accumulation of a prenylated xanthone, hyperxanthone E (4 mg/g dry weight; Gaid et al., 2012). Similar to previous reports (Klingauf et al., 2005; Fiesel et al., 2015), cell suspension cultures were treated with yeast extract on the 4th day after transfer to fresh medium when the cells were at the end of the lag phase (Appendix Fig. A1). For the detection and quantification of xanthones, methanolic extracts of yeast extract-treated and water-treated (control) cell cultures were prepared as mentioned in section 3.2. The media were also extracted. Xanthones were detected only in yeast extract-treated cultured cells. No xanthones were observed in the medium of yeast extract-treated cultures and in water-treated cell suspension cultures. The total xanthone content of the yeast extract-treated cultured cells was determined by HPLC-DAD analysis (3.6.1.1) and is represented by the existence of three xanthone peaks in Fig. 4.1a. Compound 1 corresponding to peak 1 was hyperxanthone E. The identity of the compound was confirmed by the comparison of its UV absorption spectrum (Fig. 4.1b) and retention time with that of authentic reference. However, as can be seen in Fig. 4.1a the peak is broad and there may be another co-eluting compound. The MS/MS fragmentation data of the compound was analyzed in negative ion mode and is shown in appendix Fig. A8. Compound 2 corresponding to peak 2 in Fig. 4.1a had the same UV absorption maximum (Fig. 4.1c) and retention time as the patulone reference. It was purified by HPLC and analyzed by LC-MS. The fragmentation pattern of compound 2 in negative ion mode matched that of authentic patulone (Appendix Fig. A9). Compound 3 corresponding to peak 3 in Fig. 4.1a corresponds to an unknown xanthone with the UV absorption spectrum depicted in Fig. 4.1d. It was purified by HPLC and studied by MS/MS analysis. Upon collision-induced dissociation, a molecular ion peak with m/z 395 (Appendix Fig. A10) in negative ion mode was identified. The unknown compound had the same molecular ion peak as patulone, albeit with different fragmentation pattern. The presence of similar diprenylated xanthone or possibly a monogeranylated xanthone in yeast extract-treated cell cultures of H. calycinum was first observed by Müller (2013).

Xanthone accumulation started 8 h post-elicitation followed by a steady increase until 36 h (4.7 mg/g dry weight) after which it remained stable up to 48 h (Fig. 4.2). A slow and steady decrease in xanthone content was observed after 48 h. On day 6 after treatment, the xanthone content was almost half of the maximum value. Since no xanthones were detected in the medium the compounds did not leak out but may have been either degraded to HPLC-DAD-undetectable compounds, converted to methanol-unextractable conjugates or polymers or cross-linked with the cell wall.

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Results

Yeast-extract-treated cells; 24 h after treatment Yeast-extract-treated cells; 0 h after treatment mAU

100

90

80

70 1 2 60 3

50

40

30

20 0 5 10 15 20 25 30 35 40 min (a) mAU mAU mAU 16 25 16 14 22.5 14 12 20 12 244 260 10 17.5 254 10 318 8 15 292 238 236 8 12.5 6 312 6 4 10 364 4 364 7.5 2 2 5 0 0 200 250 300 350 nm 200 250 300 350 nm 200 250 300 350 nm (b) (c) (d) Fig. 4.1: HPLC analysis of methanolic extracts from elicitor-treated H. calycinum cell cultures 24 h (a, red) and 0 h (a, blue) post-treatment. The detection wavelength was 292 nm. UV absorption spectra are shown for hyperxanthone E (b), patulone (c) and unidentified xanthone (d) corresponding to peaks 1, 2 and 3 in a, respectively. mAU, milliabsorbance units.

6

5 weight) 4

(mg/g dry 3

2 xanthones xanthones

1 Total Total 0 0 8 16 24 32 40 48 56 64 72 80 88 96 104 112 120 128 136 144 Time post-elicitation (h)

Fig. 4.2: Elicitor-induced changes in the total xanthone content of H. calycinum cells. Data are means ± SD of three biological repeats.

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Results

4.2 H. calycinum benzaldehyde dehydrogenase (HcBD)

4.2.1 Cloning a full-length HcBD cDNA BD activity was previously reported in untreated and MJ-treated H. androsaemum cell suspension cultures (Abd El-Mawla and Beerhues, 2002). Based on this information, the publicly available Hypericum databases were screened for putative BD sequences. The nucleotide sequence encoding the functionally characterized coniferylaldehyde dehydrogenase (A. thaliana reduced epidermal fluorescence 1, AtREF1; accession number: NM_113359.4), predicted to be a cytosolic protein, served as the template for performing blastx in the MPGR database. A full- length sequence encoding a putative aldehyde dehydrogenase (sequence ID: hpa_locus_1862) with 70.2% amino acid identity with AtREF1 was retrieved. Sequence information of hpa_locus_1862 was used to clone a 1743 bp full-length HcBD cDNA from yeast extract-treated H. calycinum cell suspension cultures. The 1506 bp HcBD ORF shared 93.2% identity with the database sequence and was flanked by a 103 bp 5' UTR and a 128 bp 3′ UTR with a 6 bp poly(A) tail. The proofread HcBD ORF was digested with NheI/KpnI and ligated in the NheI/KpnI linearized pRSETB vector (Fig. 4.3; This work was done by Dr. Mariam Gaid).

Transformation of E.coli BL21 cells with the recombinant plasmid was followed by heterologous expression (3.5.1) and Ni-NTA His6-tagged protein purification (3.5.2). The CDS encoded an aldehyde dehydrogenase consisting of 501 amino acids with a predicted molecular mass of 54.27 kDa and a pI of 6.2. It contained the invariant catalytic glutamate residue (E268) involved in acylation and deacylation during ALDH catalysis, the invariant cysteine residue (C302) which acts as the active site nucleophile and the coenzyme-binding GXXXXG fingerprint sequence (G245STEVG250). It shared the highest identity (78.04%) with a predicted ALDH2 member from Hevea brasiliensis (NCBI accession: XP_021643083.1) and 69.8% identity with the aforesaid AtREF1.

The purity and the subunit molecular mass of the protein were determined by SDS-PAGE (Fig. 4.4). Samples for SDS-PAGE were prepared as mentioned in section 3.5.4. The molecular mass of HcBD observed on the gel after SDS-PAGE agreed with the predicted molecular mass of the amino acid sequence.

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Results

Digestion with NheI/KpnI

+ Ligation

NheI KpnI HcBD 1506 bp

Fig. 4.3: Schematic representation for ligating the NheI/KpnI digested HcBD ORF with the NheI/KpnI linearized pRSETB vector.

1 2 3 4 L

116.0

66.2 ~54kDa 45

35

25

Fig. 4.4: SDS-PAGE of affinity-purified recombinant HcBD protein. Protein bands were visualized through Coomassie Brilliant Blue staining. Lane 1, before induction; lane 2, after induction; lane 3, soluble fraction of crude protein; lane 4, ~10 µg pure protein (~54 kDa); lane 5, ladder. 4.2.2 Biochemical characterization of HcBD Initially Tris-HCl buffer pH 8 was selected for determination of substrate specificity. The amount of protein used was 6 μg in a 200 μl assay mix and the incubation time was 15 min. All

64

Results the tested substrates are shown in appendix Fig. A16. For all substrates product formation was determined by HPLC-DAD analysis, except for acetaldehyde where the acetic acid formation was spectrophotometrically determined at 340 nm in terms of NAD+ to NADH conversion. The identities of the enzymatically formed acid products were established by comparison of the retention times (Fig. 4.5) and UV absoption spectra (Fig. 4.6) with those of authentic acid reference compounds.

Benzoic acid reference trans-Cinnamic acid reference Standard assay Standard assay R

R

P P

S S

min min 5 10 15 20 25 5 10 15 20 25 Fig. 4.5: Stacked HPLC-DAD chromatograms showing HcBD activities. R, reference compound; S, substrate (benzaldehyde or trans-cinnamaldehyde); P, enzymatically formed product. Formation of benzoic and trans-cinnamic acids (P) was recorded at 227 and 280 nm, respectively, and confirmed by the UV spectra depicted in Fig. 4.6. Inserts represent the colour code for the corresponding sample.

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Results

278 mAU mAU mAU 100 1000 120

80 100 800

60 80 600 236 60 40 230 400 40 296 20 200 274 20 0 0 0 200 250 300 350 nm 200 250 300 350 nm 200 250 300 350 nm Benzoic acid trans-Cinnamic acid 3-Hydroxybenzoic acid

mAU mAU mAU 322

298 40 300 20 254 250 236 30 15 200

20 150 10 234 100 5 10 298 50

0 0 0 nm 250 300 350 200 250 300 350 nm 200 250 300 350 nm 4-hydroxy-3-methoxycinnamic acid 2-Hydroxybenzoic acid 4-Hydroxybenzoic acid

mAU mAU mAU 140 30 12 120 25 10 100 260 20 8 260 80

15 6 60 294 4 294 40 236 10 296 20 5 2

0 0 0

200 250 300 350 nm 225 250 275 300 325 350 375 nm 200 250 300 350 nm 2-Methoxybenzoic acid 3,4-Dihydroxybenzoic acid 4-Hydroxy-3-methoxybenzoic acid

Fig. 4.6: UV absorption spectra of the enzymatically formed products which were identical to those of authentic acid references. 3,4-Dihydroxybenzoic acid is represented by reference absorption spectrum as HcBD does not accept the corresponding substrate to form the product. As determined by HPLC-DAD analysis, HcBD preferred the substrate trans-cinnamaldehyde, for which the product formation was set to 100%. Relative activities with benzaldehyde and 3- hydroxybenzaldehyde were 73.7% and 38.2%, respectively (Fig. 4.7). The relative activity with coniferaldehyde, a substituted trans-cinnamaldehyde, was 16.9%. Less than 10% of relative activities were observed with hydroxylated and methoxylated benzaldehyde derivtives. The spectrophotometric analysis revealed trace activity with the aliphatic substrate acetaldehyde relative to an identical assay done with trans-cinnamaldehyde.

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Results

120

100

80

60 activity (%) activity

BD 40 Hc 20

0

Acetaldehyde

Benzaldehyde

4-Hydroxy-3-

4-Hydroxy-3-

trans-Cinnamaldehyde

methoxybenzaldehyde

3-Hydroxybenzaldehyde 2-Hydroxybenzaldehyde 4-Hydroxybenzaldehyde

2-Methoxybenzaldehyde

methoxycinnamaldehyde 3,4-Dihydroxybenzaldehyde Substrate (0.5 mM)

Fig. 4.7: Substrate specificity of the recombinant HcBD. Activities are presented relative to the maximum substrate preference with trans-cinnamaldehyde (100%). The enzymatic assays were measured using HPLC-DAD analysis except for the activity with acetaldehyde, which was analyzed spectrophotometrically and presented relative to an equivalent measurement with trans- cinnamaldehyde. Data are means ± SD of three independent biological replicates. HcBD catalyzed enzymatic conversions of benzaldehyde and trans-cinnamaldehyde to benzoic acid and trans-cinnamic acid, respectively, as confirmed by GC-MS analysis. The fragmentation patterns and the retention times of the silylated enzymatic products matched those of the silylated authentic references (Fig. 4.8 and Fig. 4.9). Because of solvent delay, benzaldehyde peak cannot be seen in the GC-MS chromatograms shown below.

67

Results

10.49 100 90 179.1 2e+007 80 70 105.1 60 77.1 50 135.1

40 ABUNDANCE 30 1e+007 20 51.1 10 89.6 194.1

Total Ion Current Ion Total 0 60 80 100 120 140 160 180 200 m/z

7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 Time (min) (a)

10.47 100 1e+007 90 179.0 80 105.0 70 77.1 60 135.0 50

40 ABUNDANCE 30 20 51.0 194.0 10 59.0 89.5 0 m/z

Total Ion Current Ion Total 60 80 100 120 140 160 180 200

7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 Time (min) (b)

1e+007 Total Ion Current Ion Total

7.00 8.00 9.00 10.00 11.00 12.00 13.00 14.00 15.00 16.00 Time (min) (c)

Fig. 4.8: GC-MS chromatograms for HcBD incubations with the substrate benzaldehyde. Benzoic acid reference (a), benzaldehyde incubation with active HcBD showing enzymatically formed benzoic acid (b), benzaldehyde incubation with denatured HcBD, lacking product formation (c). The mass spectra of benzoic acid reference and enzymatically formed benzoic acid after derivatization with MSTFA are shown as inserts in (a) and (b), respectively.

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Results

100 3e+007 14.56 90 131.0 205.1 80 70 103.0 161.1 60 2e+007 77.1 50

40 220.1 ABUNDANCE 30 145.0 45.0 1e+007 20

Total Ion Current Ion Total 10 59.1 192 0 40 60 80 100 120 140 160 180 200 220 m/z

10.00 11.00 12.00 13.00 14.00 15.00 16.00 17.00 18.00 19.00 20.00 Time (min) (a)

100 14.53 90 205.1 80 131.1 70 103.1 161.1 60 50 77.1 1e+007

40 220.1 ABUNDANCE 30 145.0 20 45.1 10 59.1 192 0

Total Ion Current Ion Total 40 60 80 100 120 140 160 180 200 220 m/z

10.00 11.00 12.00 13.00 14.00 15.00 16.00 17.00 18.00 19.00 20.00 Time (min) (b)

1e+007 Total Ion Current Ion Total

10.00 11.00 12.00 13.00 14.00 15.00 16.00 17.00 18.00 19.00 20.00

Time (min) (c)

Fig. 4.9: GC-MS analysis of HcBD incubations with trans-cinnamaldehyde. trans-Cinnamic acid reference (a), standard incubation with trans-cinnamaldehyde leading to formation of trans- cinnamic acid (b), control assay with heat-denatured protein (c). The mass spectra of trans- cinnamic acid reference and enzymatically formed trans-cinnamic acid after derivatization with MSTFA are shown as inserts in (a) and (b), respectively. Based upon the data on substrate specificity, the optimum reaction conditions required for HcBD activity were determined with respect to the physiological substrate benzaldehyde. HcBD characterization was done spectrophotometrically. When required HPLC-DAD was used to confirm the spectrophotometric results.

69

Results

With increasing HcBD protein amount and incubation time, the product accumulation was linear over the tested ranges (Fig. 4.10a and Fig. 4.10b). A concentration of 7.5 µg/ml assay and a time of 5 min were considered as suitable parameters and used for all subsequent experiments. Maximum enzyme activity was observed at 50˚C (100%) and ~21% and ~29% reductions in the enzyme activity were observed at 40˚C and 60˚C, respectively (Fig. 4.10c.). Only 60% relative activity was observed at 35˚C which reduced further to 40% at 30˚C. HcBD was active over a broad alkaline pH range. The enzyme activity was highest over the pH range 9.5 to 11. Relative activity of ~93% was observed at pH 9 (Appendix Fig. A5). Since the enzyme showed high activity at pH values where most enzymes are inactive, another method was used to confirm the spectrophotometric results. HPLC-DAD analysis of the incubations was done to detect benzoic acid accumulation at 230 nm rather than to measure NAD+ to NADH conversion at 340 nm, as in spectrophotometric analysis. In this second analysis a similar observation was made, with the enzyme activity being high in the alkaline pH range (8-11). Only trace activity was observed at pH 12 (Fig. 4.10d). Based on the observed results and previously published data on P. pyrifolia BD and S. aucuparia BD, pH 9.5 was considered as the optimum pH for HcBD activity.

120 120 100 100 80 80

60 60 activity activity (%)

40 40

BD

BD activity BD activity (%) Hc 20 Hc 20 0 0 0 5 10 15 20 25 0 2.5 5 7.5 10 12.5 15 17.5 20 Protein (μg/ml incubation) Time (min) (a) (b) 120 125 100 100 80 75 60

activity activity (%) 50

40 BD BD activity BD activity (%) 25

Hc 20 Hc 0 0 25 30 35 40 50 60 6.5 7 7.5 8 8.5 9 9.5 10 11 12 Temperature (˚C) pH (c) (d)

Fig. 4.10: Effect of variation in incubation parameters on the activity of HcBD. Protein amount (a); time (b); temperature (c); pH (d). Data are means ± SD of three biological repeats. Supplementing the assay with chloride salts of univalent cations did not affect the enzyme activity (Fig. 4.11a), while supplementing with divalent cations had either no influence or inhibitory effect on the activity (Fig. 4.11b). Addition of Cu2+, Mn2+ and Zn2+ lead to ~22%,

70

Results

~39% and ~82% reductions in enzyme activity, respectively. HcBD activity was strictly dependent on the presence of a coenzyme, with the preferred coenzyme being NAD+ (100%). The relative activity in the presence of NADP+ was ~20%. Trace activity (< 1%) was detected in the presence of FAD (Fig. 4.11c). Addition of the reducing agent DTT did not cause pronounced increase in enzyme activity. Maximum increase in the activity (10%) was observed at 1 mM DTT compared to the control incubation without DTT (Fig. 4.11d).

120 120 100 100 80 80 60 60

40 activity (%) 40 BD activity BD activity (%) 20 BD

Hc 20 Hc 0 0 Control NH4+ Na+ K+

Univalent cation (1 mM) Divalent cation (1 mM) (a) (b) 120 120 100 100 Increase in activity on 80 80 adding DTT 60 60

activity activity (%) DTT-independent

40 40 activity BD activity BD activity (%)

BD 20 20

Hc Hc 0 0 NAD+ NADP+ FAD+ Control Coenzyme (1 mM) DTT (μM) (c) (d)

Fig. 4.11: Effect of addition of supplements on HcBD activity in the standard assay. Univalent cation (a); divalent cation (b); coenzyme (c); DTT (d). Control in (a), (b) and (d) represents standard assay without supplements. Control in (c) is a standard assay without the coenzyme. Data are means ± SD of three biological repeats. The effect of various storage temperatures and storage durations were also recorded. Non- appreciable loss of activity occurred on storage of the protein at -20˚C and -80˚C for 24 h, while ~35% decrease occurred upon storage at 4˚C (Fig. 4.12a). The protein could also be stored for 80 days at -80˚C, however, ~45% loss of activity was observed (Fig. 4.12b).

The capability of the enzyme to catalyze a reversible reaction in the presence of NADH and benzoic acid was additionally examined. No benzaldehyde production was detected upon HPLC- DAD analysis.

71

Results

120 120 100 100 80 80 60 60

40 40

BD activity (%) BD activity

BD activity (%) BD activity Hc 20 Hc 20 0 0 Control -80 -20 4 Control 1 80 Storage temperature (˚C) Duration of protein storage (days) (a) (b)

Fig. 4.12: Effect of storage conditions on the activity of HcBD. Storage temperature for 24 h (a); duration of the storage at -80˚C (b). Control represents standard assay done with freshly purified protein. Data are means ± SD of three biological repeats. Based on the characterization experiments, the optimum conditions selected for the determination of kinetic parameters of the HcBD enzyme are listed in Table 4.1.

Table 4.1: Optimum conditions selected for kinetic characterization of HcBD

Parameter Optimum condition Protein amount 1.5 μg/200 μl incubation volume Time 5 min Cofactor NAD+ Buffer Tris HCl pH 9.5 Temperature 50˚C

Centered on the substrate-specificity data and cofactor data, benzaldehyde, trans- cinnamaldehyde and NAD+ were selected for determination of kinetic parameters of HcBD. Enzymatic incubations were evaluated by HPLC-DAD analysis. The Hyperbolic regression curves for the determination of the Michaelis-Menten constant (Km) and maximum velocity (Vmax) of HcBD are presented in Fig. 4.13. The kinetic parameters are summarized in Table 4.2.

The Km values for benzaldehyde (185.93 ± 38.16 µM) and trans-cinnamaldehyde (167.60 ± + 37.26 µM) were comparable. The Km values for NAD determined in the presence of benzaldehyde and trans-cinnamaldehyde were also comparable (256.85 ± 33.58 and 286.2 ± 67.41, respectively). However, the Kcat value for trans-cinnamaldehyde was 1.6-fold higher than that for benzaldehyde, resulting in a 1.8-fold higher catalytic efficiency of HcBD with trans- cinnamaldehyde than with benzaldehyde.

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Results

Table 4.2: Steady-state kinetic parameters of HcBD. Data are means ± SD of at least three biological replicates.

-1 -1 -1 4 Substrate Km (μM) Kcat (sec ) Kcat/Km (M sec ) x10 Benzaldehyde 185.93 ± 38.16 86.68 ± 3.32 46.62 ± 9.50 NAD+ 256.85 ± 33.58 77.50 ± 5.90 30.17 ± 3.40 trans-Cinnamaldehyde 167.60 ± 37.26 142.74 ± 25.27 85.17 ± 18.56 NAD+ 286.20 ± 67.41 144.00 ± 36.25 50.31 ± 7.10

400 400 350 350 300 300 250 250 200 200 150 150 100 100

50 50 Specific activity Specific activity (nkat/mg protein) 0 Specific activity (nkat/mg protein) 0 0 200 400 600 800 1000 0 300 600 900 1200 1500 Benzaldehyde (µM) NAD+ (µM) (a) (b) 700 700

600 600

500 500 /mg /mg protein)

400 400 nkat 300 300

200 200

100 100

Specific activity (nkat/mg protein) 0

Specific activity Specific activity ( 0 0 200 400 600 800 1000 1200 0 300 600 900 1200 1500 trans-Cinnamaldehyde (µM) NAD+ (µM) (c) (d)

Fig. 4.13: Determination of Michaelis-Menten constants (Km) and maximum velocities (Vmax) of HcBD with various substrates. Hyperbolic regression curves are presented for benzaldehyde (a); NAD+ in the presence of benzaldehyde (b); trans-cinnamaldehyde (c); NAD+ in the presence of trans-cinnamaldehyde (d). 4.2.3 Phylogenetic analysis of HcBD Functional plant ALDH2 protein sequences were retrieved from the NCBI databank. The accession numbers of protein sequences employed for phylogenetic reconstruction are mentioned in appendix Table B 1. R. norvegicus ALDH2 (Accession number: P11884.1) served as the outgroup to root the neighbor-joining tree (Fig. 4.14). Two distinct clusters comprising mitochondrial and cytoplasmic ALDH2 were formed as was previously observed by Skibbe et al.

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Results

(2002) and Končitíková et al. (2015). HcBD grouped together with the cytoplasmic ALDH2 members.

100 B. napus REF1 I 100 B. napus REF1 II 97 A. thaliana REF1

100 H. calycinum BD ALDH2C subfamily Z. mays RF2C Cytoplasmic ALDH2

49 Z. mays RF2F

70 Z. mays RF2D 100 Z. mays RF2E

99 N. tabacum ALDH A. majus BALDH ALDH2B subfamily 100 Z. mays RF2A Mitochondrial ALDH2 58 Z. mays RF2B R. norvegicus ALDH Outgroup

0.050 Fig. 4.14: Neighbor-joining tree presenting the phylogenetic relationships between HcBD and other members of the ALDH2 family. Numbers at the nodes are bootstrap values arising from 1,000 replicates and Poisson correction. The scale bar indicates 0.05 amino acid substitutions per site. Functionally characterized R. norvegicus ALDH was used to root the tree. Accession numbers of all the sequences are listed in appendix Table B 1. 4.3 H. calycinum benzoate-CoA ligase (HcBZL)

4.3.1 Bioinformatic analysis of H. perforatum transcriptomes to garner putative BZL sequences Using H. calycinum CNL as query (Accession number: AFS60176.1), a tBLASTn search was carried out using the default settings in the Onekp and MPGR databases to garner 106 sequences with unique identifiers. Thirty six sequences which shared 50-60% identity to HcCNL were selected for further analysis. The rationale behind this screening was that phylogenetic analysis of various CoA ligases placed CNLs and putative BZLs in the same clade while 4CLs and long chain fatty-acyl CoA synthetases (LACS) belonged to separate clades (Shockey and Browse, 2011; Gaid et al., 2012; Klempien et al., 2012, Park et al., 2017; Gonda et al., 2018). Five sequences in the Onekp database with full-length ORFs were finally selected for cloning. The closest NCBI and MPGR database hits of the selected sequences and their corresponding identities are indicated in Table 4.3. The selected sequences might show high expression in the roots of H. perforatum, as represented by the FPKM values of the closest homologs (> 90% identity) present in MPGR database (Fig. 4.15). Roots are the site of xanthone biosynthesis and accumulation in Hypericum species (Tocci et al., 2013b; 2018) making these sequences promising candidates for BZL cloning from H. calycinum. Upon sequence analysis it was

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Results observed that the sequences Onekp:BNDE_scaffold_2018166 and Onekp:BNDE_scaffold_2018167 were assembled to the same cDNA. Thus, four sequences were used for primer design and cloning.

Table 4.3: Closest NCBI and MPGR hits of selected CoA ligases

Sequence Closest NCBI hit and Identity Closest MPGR Identity accession nmber database hit PREDICTED: Manihot 76 % hpa_locus_4529_iso 91.67 % esculenta probable _2_len_1541_ver_2 onekp:BNDE_scaffold acyl-activating enzyme _2026015 1

XM_021758090.1 Populus tomentosa 73 % hpa_locus_13827_is 93.75 % Acyl activating enzyme o_2_len_1072_ver_ onekp:BNDE_scaffold 5 (AAE5) 2 _2009069

MF463696.1 acetate/butyrate-CoA 85% hpa_locus_6408_iso 97.53 % ligase AAE7, _6_len_1880_ver_2 onekp:BNDE_scaffold peroxisomal [Ricinus _2019579 communis]

XP_002519195.1 PREDICTED: 80 % hpa_locus_733_iso_ 98.80 % Jatropha curcas 2_len_1456_ver_2 onekp:BNDE_scaffold probable acyl- _2018167 activating enzyme 2

XM_012209972.2 PREDICTED: 80 % hpa_locus_733_iso_ 98.80 % Jatropha curcas 2_len_1456_ver_2 onekp:BNDE_scaffold probable acyl- _2018166 activating enzyme 2

XM_012209972.2

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Results

100 90 80 70 hpa_locus_4529_iso_1_len_1557_ver_ 60 2 50 hpa_locus_13827_iso_3_len_1294_ver _2 FPKM FPKM (%) 40 30 hpa_locus_733_iso_1_len_1485_ver_2 20 hpa_locus_6408_iso_7_len_1989_ver_ 10 2

0

flower petals

whole old leaves

oldest part of root

flower-whole buds

whole young leaves

flower-whole mature

whole mid aged leaves middle aged part of root Tissue

Fig. 4.15: Tissue-specific expression of various putative BZL transcripts. The data was procured from the MPGR database (http://medicinalplantgenomics.msu.edu/). FPKM, Fragments per kilobase of transcript per million reads mapped. Values are represented as percent relative to the tissue with highest distribution for each transcript. 4.3.2 Cloning of HcBZL RNA was extracted from yeastextract-treated H. calycinum cell cultures 8 h after the onset of the treatment as mentioned in section 3.3.1.1. The purity of the isolated RNA samples was determined by measuring the absorbance ratio at 260/280 nm using a Nanodrop. It was ~2.1 indicating pure RNA. RNA integrity was analyzed by running the sample (1 µg) on a denaturing agarose gel (1%). The observed bands corresponding to the two ribosomal subunits, 28s rRNA and 18s rRNA, had an intensity ratio of ~2:1 which pointed towards the intactness of the RNA (Fig. 4.16).

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1 2

Fig. 4.16: Agarose gel electrophoresis of a RNA sample extracted 8 h after yeast extract- treatment of H. calycinum cultured cells. The two bands in lane 2 correspond to 18S (below) and 28S (above) ribosomal subunits; lane 1, DNA ladder. Gene-specific primers mentioned in section 2.3 were designed for the above sequences. From reverse-transcribed RNA of yeast extract-treated H. calycinum cells (8 h post-induction) two ORFs could be amplified and were later referred to as HcBZL and HcAAE1. Transcripts were not detected for onekp:BNDE_scaffold_2026015 and onekp:BNDE_scaffold_2009069 after standard and touchdown PCR (Table 4.4).

Table 4.4: Sequences selected from the Onekp database for amplification of putative HcBZL CDS

Amino acid sequence of Percent identity Amplified Sequence the peroxisomal to HcCNL product targeting signal

onekp:BNDE_scaffold Transcript not 53.721 SKL* _2026015 detected

onekp:BNDE_scaffold Transcript not 51.812 SRL* _2009069 detected

onekp:BNDE_scaffold 1692 bp, referred 50.562 SKL* _2019579 to as HcAAE1

onekp:BNDE_scaffold 53.442 _2018166 RLx5HL** 1647 bp, referred onekp:BNDE_scaffold to as HcBZL 53.442 _2018167

* PTS1; **PTS2

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Results

HcBZL cDNA consisted of a 1647 bp ORF (Fig. 4.17 a), 136 bp 5' UTR and a 110 bp 3' UTR with a 26 bp poly(A) tail. The 5' UTR was obtained using 5'-CDS primed-cDNA along with UTR-specific forward (UTRF) and gene-specific reverse (GSPR1) primers. The 3' UTR was obtained using a 3' RACE-ready cDNA along with an internal gene-specific primer (HcEndF) and RACE Long. A second round of PCR was performed using the product of the first reaction as template along with HcEndF and RACE Short. Similarly, HcAAE1 was represented by a 1692 bp PCR product (Fig. 4.17 b).

2000 ~1647 bp

1500 2000 ~ 1692 bp 1500

1000 1000

(a) (b) Fig. 4.17: Agarose gel electrophoresis of HcBZL (a) and HcAAE1 (b) as proofread PCR products. NheI/NcoI digested HcBZL ORF was ligated in NheI/NcoI linearized pRSETB expression vector (Fig. 4.18) and expressed as an N-terminally His6-tagged protein. Similarly, NheI/KpnI digested HcAAE1 ORF was ligated in NheI/KpnI linearized pRSETB expression vector (Fig. 4.19) to be also expressed as an N-terminally His6-tagged protein.

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Results

Digestion with NheI/NcoI + Ligation

NheI NcoI HcBZL 1647 bp

Fig. 4.18: Schematic representation for constructing the recombinant HcBZL expression plasmid.

Digestion with NheI/KpnI + Ligation

NheI KpnI HcAAE1 1692 bp

Fig. 4.19: Schematic representation for constructing the recombinant HcAAE1 expression plasmid.

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Results

1 2 3 4 5 200 150 120

100 85

70 60 ~60 kDa

50

40

30

Fig. 4.20: SDS-PAGE of the affinity-purified recombinant HcBZL protein. Protein bands are visualized by staining with Coomassie Brilliant Blue. Lane 1, ladder; lane 2, before induction; lane 3, after induction; lane 4, soluble fraction of the crude protein; lane 5, ~5 μg pure protein (~60 kDa).

1 2 3 4 5

116.0

66.2 ~62 kDa

45

Fig. 4.21: SDS-PAGE of the affinity-purified recombinant HcAAE1 protein. Protein bands are visualized by staining with Coomassie Brilliant Blue. Lane 1, before induction; lane 2, after induction; lane 3, soluble fraction of crude protein; lane 4, ~8 μg pure protein (~62 kDa); lane 5, ladder. The HcBZL ORF encoded a 548 amino acid protein with a predicted molecular mass of 59.98 kDa and a pI of 6.63. The size of the protein band on the SDS-PAGE gel agreed with that of the predicted molecular mass (Fig. 4.20). The protein contained the ATP/AMP-binding BOX I (T194SGTTSRPKGV204) and the BOX II (G393EIMFRG399) motifs specific to the adenylate- forming enzyme superfamily. Online prediction tools anticipated the presence of a prototypical

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Results

PTS2 (R51LASALSHL60) in the protein. The HcAAE1 ORF encoded a 563 amino acid protein with a predicted molecular mass of 61.49 kDa and a pI of 7.23. The predicted molecular mass was in agreement with the mass of the band observed on the SDS-PAGE gel (Fig. 4.21). It also contained the BOX I (T199SGTTASPKGV209) and BOX II (G399EIVMRG405) motifs and a canonical PTS1 (–S561KL563).

HcBZL and HcAAE1 shared identities ranging from 51-55% to plant CNLs, 50-70% to plant short-, branched short- and medium-chain fatty acid CoA ligases, with the exception of 83% identity of HcAAE1 with Humulus lupulus carboxy CoA ligase 3 (HlCCL3) and less than 30% identities to plant 4CLs and LACS. HcBZL and HcAAE1 shared 54% identity.

4.3.3 Preliminary screening for the determination of the substrate specificities of HcBZL and HcAAE1 HcBZL and HcAAE1 were incubated initially with ten substrates for preliminary screening. HPLC-DAD analysis of the incubations showed that HcBZL accepted benzoic acid and short- and medium-chain fatty acids (C3-C7). However, 3-hydroxybenzoic, trans-cinnamic and 4- coumaric acids were not accepted by the enzyme. On the other hand HcAAE1 activated short- chain fatty acids (C3 and C4) but did not accept benzoic acid or any other aromatic acid (Fig. 4.22).

120 HcBZL 100 HcAAE1 80

60

40

Product formed (%) formed Product 20

0

Butyric acid Butyric

Benzoic acid Benzoic

Octanoic acid Octanoic

Hexanoic acid Hexanoic

Isobutyric acid Isobutyric

Propanoic acid Propanoic

Heptanoic acid Heptanoic

4-Coumaric acid 4-Coumaric

trans-Cinnamic acid trans-Cinnamic 3-Hydroxybenzoic acid 3-Hydroxybenzoic Substrate (0.4 mM)

Fig. 4.22: Substrate-utilization profiles of HcBZL and HcAAE1 as determined by HPLC-DAD analysis. Identities of the enzymatic products (benzoyl-CoA and isobutyryl-CoA) were established by comparison of the UV absorption spectra (Fig. 4.23) and retention times with those of the authentic the CoA thioester references. Additionally, the enzymatic products were purified for

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Results

ESI-MS analysis and the fragmentation patterns of the enzymatically formed products (benzoyl- CoA and isobutyryl-CoA) matched those of the authentic CoA thioester references (Fig. 4.24 and appendix Fig. A13, respectively). The UV absorption spectra of all the enzymatically formed aliphatic-CoA thioesters were the same (Fig. 4.23 b) but the retention time of the products varied (Fig. 4.25). The products were purified for ESI-MS analysis. The m/z of the molecular ion peaks [M-H]- and the fragmentation patterns were in good agreement with the theoretical masses of the aliphatic-CoA thioesters. The ESI-MS/MS analyzed fragmentation patterns of the aliphatic-CoA thioesters are summarized in the appendix (Fig. A11-A15).

mAU 261 mAU 261 140 40 250

35 120

30 100

25 80 20 60 15 40 10

5 20

0 0

220 240 260 280 300 320 340 360 380 nm 220 240 260 280 300 320 340 360 380 nm (a) (b) Fig. 4.23: UV absorption spectra of HcBZL catalyzed thioester products. Benzoyl-CoA (a); aliphatic-CoA thioesters (b).

3.1e6 407.9 3.0e6

2.8e6

2.6e6

2.4e6

2.2e6 523.1 2.0e6 Molecular Weight: 871.64

1.8e6 Benzoyl-CoA cps

1.6e6

1.4e6 Intensity, 425.9 1.2e6 790.4 1.0e6 158.9 -- 8.0e5 [M-H] 134.0 443.3 870.4 272.9 328.0 6.0e5 541.0

4.0e5 487.9 461.0 2.0e5 79.0

0.0 50 100 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 m/z, Da Fig. 4.24: ESI-MS/MS analysis of HcBZL catalyzed benzoyl-CoA formation, represented by the molecular ion peak [M-H]– at m/z 870.4. Characteristic fragments of the phosphoadenosine-

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Results containing moiety at m/z 408 and 426 agreed with previously published reports (Zirrolli et al., 1994).

CoA-ester products

Substrates Butyric acid Hexanoic acid Propanoic acid Benzoic acid 0 5 10 15 20 25 30 min Isobutyric acid Heptanoic acid

0 5 10 15 20 25 30 min Fig. 4.25: HPLC analysis of HcBZL activity with various substrates. Standard assay with native protein (top panel), control assay with heat-denatured protein (bottom panel). The chromatograms were monitored at 261 nm. The insert shows the substrates' color codes. Different from HcAAE1, the preference of HcBZL for benzoic acid as xanthone precursor, prompted its further investigation. Spectrophotometric analysis of the HcBZL incubations with (hydroxy)cinnamic acids supported its inability to activate trans-cinnamic, 4-coumaric, ferulic, caffeic, and sinapic acids hence ruling out the possibility of the enzyme being a bona fide CNL or 4CL.

4.3.4 Luciferase-based determination of the substrate-specificity of HcBZL

4.3.4.1 Cloning, heterologous expression and His6-tagged luciferase purification For the luciferase-dependent substrate specificity assay, in-house luciferase production was done. A DH5α harbouring pRS413-GAL1-luc*(-SKL) plasmid which comprised a luciferase ORF was ordered from the Addgene repository as agar stab. A bacterial colony was grown in LB medium containing ampicillin. The pRS413-GAL1-luc*(-SKL) plasmid containing the luciferase ORF was isolated from bacterial cells as mentioned in section 3.3.1.2. The plasmid served as template for amplification of the 1650 bp luciferase ORF. The proofread amplified product was subjected to NheI/KpnI restriction digestion followed by its ligation in a NheI/KpnI linearised pRSERTB expression vector (Fig. 4.26).

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Results

Amplifying Luciferase ORF NheI/KpnI digestion ~1650 bp Ligation in NheI/KpnI linearized pRSETB

Luciferase

pRS413-GAL1-luc*(-SKL) pRSETB-Luciferase

Fig. 4.26: Schematic representation of the subcloning of the luciferase ORF from pRS413-GAL- luc*(-SKL) into pRSETB expression vector. Following DH5α transformation, plasmid isolation and plasmid screening through restriction digestion, a positive clone was selected and sent for sequencing on both strands. The protein sequence encoded was 100% identical to Photinus pyralis luciferase (NCBI accession: AAA29795.1), except for the presence of a point mutation in the PTS1 (Appendix Fig. C1). E. coli BL21 cells were transformed with the recombinant plasmid. Following heterologous expression and His6-tagged protein purification, the ~60.7 kDa luciferase protein was obtained (Fig. 4.27).

1 2 3 4 5

116.0

66.2 ~60.7 kDa

45

35

Fig. 4.27: SDS-PAGE of affinity-purified recombinant P. pyralis luciferase. Protein bands were visualized by staining with Coomassie Brilliant Blue. Lane 1, before induction; lane 2, after induction; lane 3, soluble fraction of crude protein; lane 4, ~7 µg pure protein (~60.7 kDa); lane 5, ladder.

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Results

4.3.4.2 Optimization of the luciferase amount for the luciferase-based substrate-specificity assay Once pure luciferase protein was obtained, the luciferase-dependent increase in chemiluminescence was analyzed by a luminometer. Chemiluminescence is presented as counts per second (cps) and increased linearly with increasing protein amount up to 25 μg. A luciferase amount of 0.5 µg was selected for subsequent assays. The cps produced by 0.5 µg in-house luciferase was comparable with the cps produced by 1 µg of commercial luciferase (Sigma; appendix Fig. A6).

800 25

20 3 15 10 600 CPSx10 5

) 0 3 0 0.5 1

400 Amount of Luciferase (μg) CPS(x10 200

0 0 5 10 15 20 25 Amount of Luciferase (μg)

Fig. 4.28: Optimization of purified recombinant luciferase for its application in the luciferase- based substrate-specificity assay. The insert shows the linear increase in luminescence between 0 and 1 µg pure recombinant luciferase. A quantity of 0.5 µg of in-house prepared recombinant protein was selected for the assays. CPS, counts per second. 4.3.4.3 Substrate-specificity determination of HcBZL Following the screening with 10 substrates in section 4.3.3, wider arrays of substrates (Appendix Fig. A17) were tested using the luciferase-based assay. The assay takes advantage of the adenylate-intermediate-forming properties of luciferase and CoA ligase under study (Fig. 4.29). Relative luciferase activity is inversely correlated to the ATP utilization of the CoA ligase under study.

HcBZL Luciferase Recorded Left over ATP Luminescence Substrate by Luminometer ATP + MgCl2 + CoASH + CoA ester D-Luciferin + O2 + Oxyluciferin

Fig. 4.29: Principle underlying the luciferase-based substrate specificity assay (Schneider et al., 2005). RpBZL does not accept 4-coumaric acid to form 4-coumaroyl-CoA. Thus, incubation with 4- coumaric acid served as a control. In this control, unutilized ATP in the first step is utilized by luciferase in the second step to produce luminescence, which is recorded by a luminometer and is

85

Results set as 100% relative luciferase activity. Relative luciferase activity with butyric, benzoic, propanoic, hexanoic and isobutyric acids were 2.6%, 16%, 16.7%, 16.7% and 20.9%, respectively. Low relative luciferase activity is indicative of better substrate and ATP utilization by the CoA ligase. Hence, butyric, benzoic, propanoic, hexanoic and isobutyric acids were the preferred substrates for HcBZL and were chosen for determination of kinetic parameters of HcBZL. Relative luciferase activities of 100% or more were observed with trans-cinnamic acid and its hydroxylated derivatives as well as with hydroxylated derivatives of benzoic acid, which indicated that they were not accepted by HcBZL. Similar results were observed for acetic and

malic acids (Fig. 4.30).

6 6 6

acid

C C C

- - -

3 1 2

C C C

Dicarboxylic

Control Fatty acids 200

180

160

140

120

100

80

60

40

Relative luciferase activity % activity luciferase Relative * * * * 20 *

0

Malic Malic acid

Acetic acid Acetic

Ferulic Ferulic acid

Caffeic acid Caffeic

Butyric acid Butyric

Sinapic acid Sinapic

Benzoic acid Benzoic

Shikimic Shikimic acid

Octanoic acid Octanoic

Hexanoic acid Hexanoic

Mandelic acid Mandelic

Phenylalanine

Isobutyric Isobutyric acid

Propanoic Propanoic acid

4-Coumaric 4-Coumaric acid

trans-Cinnamic acid

2-Aminobenzoic acid 2-Aminobenzoic

4-Hydroxybenzoic acid 4-Hydroxybenzoic 3-Hydroxybenzoic 3-Hydroxybenzoic acid

2-Hydroxybenzoic acid 2-Hydroxybenzoic

4-Coumaric acid 4-Coumaric (RpBZL) acid

2,3-Dihydroxybenzoic acid 2,3-Dihydroxybenzoic 3,5-Dihydroxybenzoic 3,5-Dihydroxybenzoic acid

3,4,5-Trihydroxybenzoic acid 3,4,5-Trihydroxybenzoic 4-Hydroxy-3-methoxybenzoic acid 4-Hydroxy-3-methoxybenzoic Substrate (0.2 mM)

Fig. 4.30: Luciferase-based substrate utilization profile of HcBZL. ATP left over in the BZL assay was measured via the luciferase-coupled bioluminescence. Normalization of the data was done with respect to an assay mix containing RpBZL and its non-substrate 4-coumaric acid where no ATP utilization occurred and the relative luciferase activity was set to 100%. The best utilized substrates are depicted by asterisks.

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Results

4.3.5 Biochemical characterization of HcBZL

As mentioned in section 3.5.9, it is important to optimize various parameters associated with an enzymatic incubation in order to obtain maximum product formation and to determine the kinetic properties of an enzyme correctly. Taking this into consideration the optimum reaction conditions for HcBZL were determined with respect to benzoic acid, which is the physiological substrate. Analysis was done by HPLC-DAD.

With the increase in the amount of protein and incubation time, product accumulation in 250 µl assays was linear up to 10 μg protein and 30 min, respectively. A slow but continuous increase in benzoyl-CoA formation was observed up to 3 hours (Fig. 4.31a and Fig. 4.31b). The temperature optimum was 30˚C. With 10˚C decrease/increase in temperature ~22/30% decrease in enzyme activity occurred (Fig. 4.31c). Upon testing buffers with various pH it was determined that maximum enzyme activity occurred at pH 6.5, which was the optimum pH. More than 90% of enzyme activity was observed at a pH range of 6 to 7.5 (Fig. 4.31d).

120 120

100 100

(%) (%) 80 80

60 60

activity activity activity activity

40 40

BZL

BZL Hc 20 Hc 20 0 0 0 10 20 30 40 50 60 70 80 90 100 0 15 30 45 60 75 90 105 120 135 150 165 180 Protein (µg) Time (minutes) (a) (b)

120 120 100 100 80 80

60 60 activity activity (%)

activity activity (%) 40 40

BZL BZL

20 c 20 H Hc 0 0 20 30 40 50 60 5 5.5 6 6.5 7 7.5 8 8.5 9 9.5 10 Temperature (˚C) pH (c) (d)

Fig. 4.31: Effect of the variation in various incubation parameters on the activity of HcBZL. Protein amount (a); incubation time (b); temperature (c); pH (d). Data are represented as means ± SD of three biological repeats. HcBZL activity was strictly dependent on the presence of a divalent cation. Among the various divalent cations tested, Mn2+ led to maximal enzyme activity followed by Mg2+ and Co2+ (84%

87

Results and 83.6%, respectively), whereas the addition of Cu2+ abolished the enzyme activity (Fig. 4.32a). The optimum concentration of Mn2+ or Mg2+ required in the enzymatic incubation was ~63 µM. Around 90% relative activity was observed for both Mn2+ and Mg2+ at 30 μM final concentration (Fig 4.32c). Addition of the chloride salts of univalent cations to the incubation led to an increase in enzyme activity. Potassium at a final concentration of 2 mM augmented the enzyme activity by 20% and therefore potassium phosphate buffer was selected for future experiments (Fig. 4.32b). Around 20% increase in the enzyme activity was also observed after adding 250-500 µM DTT (Fig. 4.32d).

120 140 120 100 Increase in

(%) 100 activity on 80 univalent 80 cation addition

60 activity activity (%) activity activity 60 Activity

40 BZL

BZL 40 independent Hc Hc of univalent 20 20 cation addition 0 0 Mn2+ Mg2+ Co2+ Ni2+ Ca2+ Fe2+ Cu2+ K+ Na+ NH4+ Li+ Control Divalent cation (2.5 mM) Univalent cation (2 mM) + Mg2+ (2.5 mM) (a) (b) 120

100

80

60 Manganese ion activity activity (%)

40 Magnesium ion BZL

Hc 20

0

1 5

10

0.5

0.03 0.25

0.001 0.125 0.0625 Divalent cation (mM) (c) 140 120 100 Increase in 80 activity upon DTT addition activity activity (%) 60

BZL 40 DTT- Hc 20 independent activity 0 0 50 62.5 125 250 500 750 1000 DTT (µM) (d) Fig. 4.32: Effect of addition of supplements to the incubation of HcBZL. Divalent cation (a); univalent cation (b); effect of increasing concentrations of Mn2+ and Mg2+ (c); DTT (d). Control in (b) represents a standard assay with Tris-HCl buffer. Data are represented as means ± SD of three biological repeats.

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Results

The effect of varying storage temperature and storage duration on the enzyme activity was also recorded. As HcBD, HcBZL was stored at 4˚C, -20˚C and -80˚C for 24 h. No significant loss of activity was observed after storage at 4˚C, -20˚C and -80˚C for 24 h (Fig. 4.33a). A loss of 32% of enzyme activity occurred upon storage of the protein at -80˚C for a month (Fig. 4.33b).

120 120 100 100 80 80 60 60

40 40 BZl activity BZl activity (%)

BZL activity BZL activity (%) 20 20

Hc Hc 0 0 Control 4 -20 -80 Control 1 30 Storage temperature (˚C) Storage period (days) (a) (b)

Fig. 4.33: Effect of storage conditions on the activity of HcBZL. Storage temperature for 24 h (a); duration of the storage at -80˚C (b). Control represents standard assay done with freshly purified protein. Data are means ± SD of three biological repeats. Table 4.5: Optimum incubation conditions for HcBZL activity with benzoic acid as substrate, used to determine Michaelis-Menten kinetics.

Factor Optimum condition pH 6.5 Time 20 min Temperature 30˚C Divalent cation Mg2+ Buffer Potassium phosphate Protein concentration 5 μg/ 250 μl

Using the above-mentioned optimum conditions (Table 4.5), the kinetic properties of HcBZL were determined as mentioned in Table 4.6. The hyperbolic regression curves are shown in Fig. 4.34 and Fig. 4.35. The catalytic efficiencies (Kcat/Km) of HcBZL for benzoic acid and isobutyric acid were comparable. However, they were ~3.5-fold lesser than those for propanoic acid and hexanoic acid and ~6-fold lesser than that for butyric acid. The Km values for the co-substrates ATP and CoA in the presence of benzoic acid were 14.12 ± 2.75 µM and 42.29 ± 7.70 µM, respectively, and were comparable to the Km values for ATP and CoA in the presence of isobutyric acid (13.58 ± 2.85 µM and 33.80 ± 1.84 µM, respectively).

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Results

Table 4.6: Steady-state kinetic parameters of HcBZL. Data are represented as means ± SD of at least three independent biological repeats.

-1 -1 -1 Substrate Km (μM) Kcat (s ) Kcat/Km (M s )

Benzoic acid 99.76 ± 14.45 0.079 ± 0.006 791.95 ± 161.77

Propanoic acid 103.03 ± 26.87 0.29 ± 0.059 2821.52 ± 698.67

Butyric acid 112.07 ± 11.19 0.53 ± 0.029 4744.79 ± 207.05

Isobutyric acid 65.93 ± 5.30 0.059 ± 0.011 845.46 ± 136.52

Hexanoic acid 112.77 ± 10.34 0.32 ± 0.021 2847.51 ± 304.31

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Results

1.4 1 1.2 0.9 0.8

1 protein) 0.7 0.8 /mg 0.6

nkat 0.5 0.6 0.4 0.4 0.3 0.2 0.2

Specific activity (nkat/mg protein) (nkat/mg activitySpecific 0.1 Specific activity ( activitySpecific 0 0 0 100 200 300 400 500 0 100 200 300 400 500 Benzoic acid (µM) Isobutyric acid (µM) (a) (d) 2 1.6 1.8 1.4 1.6 1.4 1.2 1.2 1 1 0.8 0.8 0.6 0.6 0.4 0.4

0.2 0.2

Specific activity (nkat/mg protein) (nkat/mg activitySpecific Specific activity (nkat/mg protein) (nkat/mg activitySpecific 0 0 0 50 100 150 200 250 300 350 400 0 100 200 300 400 CoA (µM) CoA (µM) (b) (e) 1.8 1 1.6 1.4 0.8

1.2 /mg protein) /mg

1 0.6 nkat 0.8 0.4 0.6 0.4 0.2

0.2

Specific activity ( activitySpecific Specific activity (nkat/mg protein) (nkat/mg activitySpecific 0 0 0 50 100 150 200 0 25 50 75 100 125 150 175 200 ATP (µM) ATP (µM) (c) (f)

Fig. 4.34: Determination of the Michaelis-Menten constants (Km) and maximum velocities (Vmax) of HcBZL with various substrates and co-substrates. Hyperbolic regression curves are depicted for benzoic acid (a), CoA in the presence of benzoic acid (b), ATP in the presence of benzoic acid (c), isobutyric acid (d), CoA in the presence of isobutyric acid (e), and ATP in the presence of isobutyric acid (f). Data are represented as means ± SD of at least three independent biological repeats.

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Results

5 6 4.5 5

4 protein) 3.5 4 /mg /mg 3

nkat 2.5 3 2 2 1.5 1 1

0.5

Specific activity ( activitySpecific Specific activity (nkat/mg protein) (nkat/mg activitySpecific 0 0 0 100 200 300 400 500 0 100 200 300 400 500 600 700 Propanoic acid (µM) Hexanoic acid (µM) (a) (b)

9 8 7 6 5 4 3 2

1 Specific activity (nkat/mg protein) (nkat/mg activitySpecific 0 0 100 200 300 400 500 600 700 Butyric acid (µM) (c)

Fig. 4.35: Determination of the Michaelis-Menten constants (Km) and maximum velocities (Vmax) of HcBZL with various substrates. Hyperbolic regression curves are depicted for propanoic acid (a), hexanoic acid (b), and butyric acid (c). Data are represented as means ± SD of at least three independent biological repeats. 4.3.6 Phylogenetic reconstitution of HcBZL and HcAAE1 Functional plant AAE sequences were retrieved from the NCBI databank. The accession numbers of the protein sequences employed for phylogenetic reconstruction are indicated in appendix Table B 2. P. pyralis as a functionally characterized luciferase (NCBI accession: AAA29795.1) was used to root the neighbor-joining tree (Fig. 4.36). It was observed that the CNLs and the 4CLs belonged to different clades as was observed previously (Shockey and Browse, 2011; Gaid et al., 2012; Klempien et al., 2012, Park et al., 2017; Gonda et al., 2018). HcBZL and HcAAE1 belonged to the large clade comprising the CNLs and other short-chain AAEs referred to here as benzenoid/short-chain acyl-CoA ligases.

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100 HlCCL13 Humulus lupulus 95 HlCCL2 Humulus lupulus 100 HcBZL Hypericum calycinum AtAAE1 Arabidopsis thaliana 98 75 onekp:BNDE scaffold 2026015 AtAAE7 Arabidopsis thaliana Benzenoid/Short-chain HlCCL3 Humulus lupulus 56 100 acyl–CoA ligases 99 HcAAE1 Hypericum calycinum 100 AtAAE11 Arabidopsis thaliana AtBZO1/CNL Arabidopsis thaliana 100 99 CbCNL/BZL Clarkia breweri 76 PhCNL Petunia hybrida 89 HcCNL Hypericum calycinum AtAAE2 Arabidopsis thaliana

99 CmCNL Cucumis melo 94 HlCCL4 Humulus lupulus 60 onekp:BNDE scaffold 2009069

95 Sa4CL3 Sorbus aucuparia 69 Pt4CL2 Populus tremuloides 99 Rg4CL1 Ruta graveolans At4CL3 Arabidopsis thaliana 100 At4CL2 Arabidopsis thaliana 100 At4CL1 Arabidopsis thaliana 4-Coumarate-CoA ligases 100 Rg4CL2 Ruta graveolens 92 Sa4CL1 Sorbus aucuparia 68 Pt4CL1 Populus tremuloides 50 Ph4CL1 Petunia hybrida 62 Sa4CL2 Sorbus aucuparia 98 HlCCL1 Humulus lupulus PpLuciferase Photinus pyralis Outgroup

0.10 Fig. 4.36: Neighbor-joining tree representing the phylogenetic relationships between HcBZL, HcAAE1 and other plant AAEs. Numbers at the nodes are bootstrap values arising from 1,000 replicates and Poisson correction. The scale bar indicates 0.1 amino acid substitutions per site. Functionally characterized luciferase from P. pyralis luciferase was used to root the tree. Accession numbers of the sequences used are listed in appendix Table B 2. 4.4 Elicitor-induced accumulation of HcBD and HcBZL transcripts as well as xanthones in H. calycinum cell cultures As mentioned in section 3.1.1, H. calycinum cells and medium were harvested at various time points. A part of the harvested cells was snap frozen in liquid nitrogen and stored at -80˚C for RNA extraction. The remainder of the cells and the culture medium were used for total xanthone analysis. RNA was isolated from untreated (0 h) and yeast extract-treated (1, 2, 4, 8, 12, 16, 20, 24, 36 and 48 h) H. calycinum cell cultures as mentioned in section 3.3.1.1 and treated as mentioned in section 3.3.5.4. The purity of the isolated RNA samples was examined by measuring the absorbance ratio at 260/280 using a Nanodrop. All samples showed a ratio of ~2.1 indicating pure RNA. RNA integrity was analyzed by running samples on a denaturing agarose gel with the observed bands corresponding to the two ribosomal subunits, 28s rRNA and 18s rRNA.

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4.4.1 RT-PCR analysis of HcBD expression For determination of HcBD expression by RT-PCR, 18s rDNA and H2A genes served as the reference genes for normalization of the expression data. HcBD, 18s rDNA and H2A primers were designed to fetch 537 bp, 485 bp and 527 bp fragments, respectively.

1.4

1.2

1

0.8

transcript 0.6 BD

Hc 0.4 of of

0.2 Relative integrated intensity intensity Relativeintegrated 0 0 1 2 4 8 12 16 20 24 36 Post-elicitation time (h)

Fig. 4.37: Elicitor-induced changes in HcBD expression. HcBD transcript levels were normalized with respect to H2A (•••) and 18s rRNA (---) transcripts. Data are presented as means ± SD of three biological replicates. RT-PCR analysis indicated stable expression of the 18s rDNA and H2A genes in all samples. However, expression of HcBD varied in H. calycinum cells, with an up-regulation of HcBD expression soon after yeast extract-treatment. Maximum transcript accumulation occurred at 8- 12 h post-elicitation and then the level started to decrease (Fig. 4.37).

4.4.2 RT-qPCR analysis of HcBZL expression HcBZL expression was studied by RT-qPCR. H2A and actin genes were used as reference genes for normalization of expression data. These reference genes were previously used in RT-qPCR analysis of genes associated with the xanthone biosynthetic pathway in H. calycinum cells and H. perforatum roots. The primers for the reference genes were designed as stated previously (El Awaad et al., 2016; Tocci et al., 2018; Nagia et al., 2019) and the amplicon sizes were 72 and 77 bp, respectively. Primers for HcBZL were designed using Primer3 software (2.10). The primers had 55% GC content. The Tm was ~60˚C and fetched a 160 bp amplicon.

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Initially, the amplification efficiency (E) and specificity of each primer pair for its corresponding gene was checked. The cDNAs from three independent biological repeats were analyzed (each in triplicate). An undiluted pool of cDNA was prepared which was subjected to four five-fold serial dilutions. For each sample a 10 µl PCR assay (Table 3.8) was performed using the thermal profile mentioned in Table 3.9. No template controls (NTC) and no reverse-transcriptase controls (NRC) were included to check for primer dimer and genomic DNA contamination, respectively.

A standard curve was generated using the Bio-Rad software for each primer pair and probe set at the end of the run which was based on the quantitative cycle (Cq) values for undiluted and various dilutions of the pooled cDNA. The slope of standard curve is indicative of E with E = 10[-1/slope] and amplification efficiency percentage E% = (E-1)*100. An E value of 2 is indicative of perfect doubling of the amplicon with each cycle in the exponential phase with E% being 100%. The slope, correlation coefficient (r2) and E% are indicated in Table 4.7.

Table 4.7: Slope of the standard curves, correlation coefficient (r2) and amplification efficiency percentage generated by the software used for each primer pair and probe set

Bio_1 Bio_2 Bio_3 Gene Slope r2 E % Slope r2 E % Slope r2 E % HcBZL -3.15 0.994 107.7 -3.26 0.993 102.6 -2.908 0.998 120.8 HcActin -2.981 0.992 116.5 -3.136 0.998 108.4 -3.167 0.992 106.9 HcH2A -3.311 0.996 100.5 -3.361 0.999 98.4 -3.205 0.998 105.1

Melting curve analysis indicated the specificity of the primers. As seen in appendix Fig. A7, the melt curves are indicative of amplification of a single gene-specific amplicon. Product- specificity was also confirmed by sequencing. No primer-dimer formation occurred. No amplification was observed in NTC and NRC. For the differential expression study of untreated versus treated samples, 1 ng cDNA per 10 μl reaction (1:25 dilution) was selected. Zero-time- point/untreated sample data served as the calibrator/control. First, relative expression/ relative quantity (RQ) was calculated using the Pfaffl (2001) method with equation 1, and then the reference genes were used to normalize the gene expression according to equation 2 giving normalized relative expression/normalized relative quantity (NRQ).

RQ = E-ΔCq (Equation 1) where ΔCq = Cqcontrol-Cqtreated

NRQ = RQTarget/(RQref 1*RQref 2*RQref n)1/n (Equation 2) where n is the number of reference genes

RT-qPCR analysis indicated that the HcBZL up-regulation began 4 h post-elicitation and increased until 16 h after which the expression levels started to decrease slowly (Fig. 4.38).

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3.5 3 2.5 2 1.5 1 0.5

Normalized relative expression relative Normalized 0 0 1 2 4 8 12 16 20 24 36 48 Post-elicitation time (h) Fig. 4.38: Changes in the normalized relative expression of HcBZL in H. calycinum cell cultures upon treatment with yeast extract. Expression in control (0 h) is set as 1. Data are represented as mean ± SD of three biological replicates with three technical repeats each. Both HcBD and HcBZL transcript accumulation preceded the accumulation of the xanthones (Section 4.1; Fig. 4.2) suggesting the in vivo involvement of the HcBD and HcBZL enzymes in xanthone biosynthesis.

4.5 Subcellular localization of HcBD and HcBZL

4.5.1 Preparation of HcBD and HcBZL YFP fusion constructs for subcellular localization Various online subcellular localization prediction tools anticipated the cytoplasmic and peroxisomal locations of HcBD and HcBZL in the plant cell, respectively (Table 4.8). However, there were some outliers and experimental data was required to determine the correct location of the proteins in the plant cell.

Table 4.8: Predicted localization of HcBD and HcBZL

Prediction software Predicted location of HcBD Predicted location of HcBZL Plant-mPLoc Chloroplast Peroxisome PPero No PTS1 No PTS1 DeepLoc-1.0 Cytoplasm Peroxisome (Likelihood: 0.6323) (Likelihood: 0.5099) LocTree3 Mitochondrion Peroxisome (Expected accuracy: 89%) (Expected accuracy: 89%)

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To determine the HcBD and HcBZL locations in the plant cell, N-terminal and C-terminal YFP fusions were constructed. Gateway cloning technology was used for the preparation of expression clones encoding the fusion proteins. Initially, the respective attB PCR products were amplified. Templates for amplification were the HcBD-pRSETB and HcBZL-pRSETB plasmids. Primers used are stated in section 2.3. Amplification of the attB PCR product for construction of the YFP-ORF expression clone for N-terminal fusion was straight forward, however, the natural stop codon in the ORF was removed when amplifying the attB product for the construction of the ORF-YFP expression clone giving C-terminal fusion. Agarose gel images indicating bands corresponding to attB PCR products of HcBD and HcBZL are shown in Fig. 4.39.

1700 bp 1500 bp

HcBD-DEL HcBD attB HcBZL attB HcBZL-DEL attB with stop 1200 bp with stop attB No stop codon codon No stop codon codon

(a) (b)

Fig. 4.39: Agarose gel images indicating bands corresponding to attB PCR products of HcBD (a) and HcBZL (b). Once attB PCR products were produced, respective entry clones were generated using BP cloning as mentioned in section 3.3.8. The entry clones were verified by setting up a simple PCR with the entry clone as the template and gene-specific forward and M13 reverse primers. The amplified product is represented by a band of the correct size on agarose gel. Schematic representations for generation of entry clones of HcBD and HcBZL are shown in Fig. 4.40 and Fig. 4.41, respectively.

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BP cloning + attB1 attB2 HcBD attB product (with stop codon)

(a)

1500 bp HcBD HcBD-DEL Entry clone Entry clone

(c)

BP cloning +

attB1 attB2 HcBD-DEL attB product (no stop codon)

(b)

Fig. 4.40: Schematic representation of construction of HcBD entry clones. HcBD entry clone (a); HcBD-DEL entry clone (b); Agarose gel image indicating bands corresponding to HcBD ORF and HcBD-ORF-DEL amplified from the entry clones (c). DEL, deletion of the stop codon.

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BP cloning +

attB1 attB2 HcBZL attB product (with stop codon) (a)

1700bp 1500bp HcBZL HcBZL-DEL Entry Entry clone clone

(c) BP cloning +

attB1 attB2 HcBZL-DEL attB product (without stop codon)

(b)

Fig. 4.41: Schematic presentation of the construction of HcBZL entry clones. HcBZL entry clone (a); HcBZL-DEL entry clone (b); Agarose gel image indicating bands corresponding to HcBZL ORF and HcBZL-ORF-DEL amplified from the entry clones (c). DEL, deletion of the stop codon. Following isolation of entry clones, LR cloning was carried out as mentioned in section 3.3.8. LR cloning led to generation of expression clones (Fig. 4.42 and Fig. 4.43). The expression clones were verified by setting up a simple PCR with the expression clone as the template and

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Results gene-specific forward and reverse primers. The amplified product was represented by a band of the correct size on agarose gel (Fig. 4.42c and Fig. 4.43c).

+ LR cloning

(a)

HcBD-YFP YFP-HcBD Expression Expression clone clone

(c)

LR cloning +

(b)

Fig. 4.42: Schematic representation of construction of HcBD expression clones. YFP-HcBD (a); HcBD-YFP (b); Agarose gel image indicating bands corresponding to the HcBD ORF amplified from the expression clones (c).

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+ LR cloning

(a)

1700 bp 1500 bp YFP-HcBZL HcBZL-YFP Expression Expression clone clone

(c)

LR cloning +

(b) Fig. 4.43: Schematic representation of construction of HcBZL expression clones. YFP-HcBZL (a); HcBZL-YFP (b); Agarose gel image indicating bands corresponding to HcBZL ORF amplified from the expression clones (c). Expression of the fusion constructs was controlled by the constitutive cauliflower mosaic virus 35S promoter and the synthase terminator sequence. The expression clones were sequenced using YFP-specific primers (2.3) to ensure that the fusion occurred and the correct

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Results reading frame was maintained. The positive expression clones were then transferred to A. tumefaciens C58C1 (pMP90 helper plasmid) via electroporation as mentioned in section 3.4.3.2. Growth, preparation of various concoctions of agrobacteria transformed with different expression clones and agrofiltration of N. benthamiana leaves were done as mentioned in section 3.5.12.

4.5.2 Subcellular localization of HcBD For analysis of HcBD fusion constructs, agroinfiltrated N. benthamiana leaf discs were analyzed by laser scanning microscopy 3 days post-agroinfiltration. Upon transient expression both fusion proteins, YFP-HcBD and HcBD-YFP, were detected in the cytoplasm of the cell (Fig. 4.44).

35S HcBD YFP S

a b c

35S YFP HcBD S

d e f

Fig. 4.44: Subcellular localization of HcBD fused to YFP in N. benthamiana leaf epidermis cells. Transient expression of HcBD-YFP (a, b, c) and YFP-HcBD (d, e, f). Both constructs resulted in cytoplasmic proteins (a, d). Red indicates the chlorophyll autofluorescence (b, e).The corresponding images were superimposed (c, f). 35S, promoter from cauliflower mosaic virus; YFP, yellow fluorescent protein; S, stop codon. Bars = 10 μm.

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4.5.3 Subcellular localization of HcBZL The HcBZL sequence contains the N-terminal PTS2, R51LASALSHL60. To determine experimentally the role of the N-terminal PTS2 in subcellular localization of HcBZL and to exclude the presence of a C-terminal PTS1 in HcBZL, C-terminal (HcBZL-YFP) and N-terminal (YFP-HcBZL) fusions were constructed, respectively. For co-localization, the coding sequence of the CFP fused to a PTS1 tripeptide was used as a peroxisomal marker (Nowak et al., 2004). In addition, co-localization was studied using the dually localized peroxisomal and cytoplasmic marker eqFP611 (Forner and Binder, 2007; Gehl et al., 2009). The fusion products encoded by the various constructs were detected by cLSM.

Upon transient expression in N. benthamiana leaf epidermal cells, YFP-HcBZL was exclusively localized in the cytoplasm, indicating the absence of a PTS1 (Fig. 4.45). By contrast, the protein encoded by the HcBZL-YFP construct was dually localized, with YFP fluorescence being strongly detected in the cytoplasm and weakly found in the peroxisomes in punctate form (Fig. 4.46).

35S YFP HcBZL S

PTS2 SLS

Fig. 4.45: Cytoplasmic localization of YFP-HcBZL upon transient expression of the construct in N. benthamiana leaf epidermis cell. 35S, promoter from cauliflower mosaic virus; PTS2, type 2 peroxisomal targeting signal; SLS, terminal tripeptide of the HcBZL protein; YFP, yellow fluorescent protein; S, stop codon; Bars = 10 μm. The punctate fluorescence resulting from transient expression of HcBZL-YFP (Fig. 4.46a), overlapped with that of the peroxisomal marker (Fig. 4.46b), leading to white fluorescence (Fig. 4.46c), which did not overlap with the autofluorescence of chlorophyll (Fig. 4.46d). N. benthamiana leaves were infiltrated with transformed A. tumefaciens C58C1 concoctions with varying ratios of HcBZL-YFP and CFP-PTS1 contructs (1:1, 2:1, 5:1, 10:1; v:v). Analysis was undertaken between 2 days after agroinfiltration when the expression of fusion proteins was weak and 4 days after agroinfiltration when the expression was high. The peroxisomal localization of HcBZL-YFP was extremely weak and could be seldom found on repeated analyses. Upon increasing A. tumefaciens C58C1 habouring HcBZL-YFP in the concoctions to be infiltrated, an increase in cytoplasmic fluorescence was observed rather than in punctate fluorescence in N. benthamina epidermal cells.

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To validate peroxisomal and cytoplasmic localization of HcBZL, N. benthamiana leaves were cotransformed with a mixture of A. tumefaciens C58C1 harboring HcBZL-YFP and the far-red fluorescent protein eqFP611. The fusion proteins encoded by both the constructs were co- localized in the cytoplasm and the peroxisomes (Fig. 4.46e and f). Upon overlap they produced pale blue fluorescence (Fig. 4.46g), which did not superimpose with the autofluorescence of chlorophyll (Fig. 4.46h).

35S HcBZL YFP S 35S CFP PTS1 S PTS2 SLS +

a b c d

35S HcBZL YFP S 35S eqFP611 PTS1 S PTS2 SLS + e f g h

Fig. 4.46: Co-transformation of N. benthamiana leaf epidermis cells with HcBZL-YFP and either the peroxisomal marker CFP-PTS1 (a-d) or the dually localized peroxisomal and cytoplasmic marker eqFP611 (e-h). Yellow YFP fluorescence (a, e), cyan CFP fluorescence (b), blue eqFP611 fluorescence (f). White areas indicate the perfect overlap of yellow and cyan fluorescent proteins (c). Pale blue areas indicate the perfect overlap of yellow and far red fluorescent (eqFP611) proteins (g). Red color indicates chlorophyll autofluorescence (d, h). 35S, promoter from cauliflower mosaic virus; PTS1, type 1 peroxisomal targeting signal; PTS2, type 2 peroxisomal targeting signal; SLS, terminal tripeptide of HcBZL; YFP, yellow fluorescent protein; CFP, cyan fluorescent protein; eqFP611, Entacmaea quadricolor far red fluorescent protein was manually selected to appear blue in order to differentiate it from the autofluorescence of chlorophyll; S, stop codon. Bars =10 µm. Furthermore, to experimentally confirm that the peroxisomal localization of HcBZL was due to the presence of a PTS2, three truncated HcBZL constructs were prepared (Fig. 4.48a). attB PCR products were amplified for Fragment 1 (246 bp), Fragment 2 (1419 bp) and Fragment 3 (108 bp) of the HcBZL ORF. Preparation of entry and expression clones was done using the Gateway

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Results cloning protocol, following a similar scheme as was mentioned previously for the full-length sequences. Expression clones with truncated HcBZL sequences are shown in Fig. 4.47.

(a) (b)

(c)

Fig. 4.47: Expression clones generated using three truncated HcBZL attB PCR products (b,c,d) as presented in Fig. 4.48a. The products of both the transiently expressed Fragment 1-YFP construct containing a PTS2 and the Fragment 2-YFP construct without any detectable PTS were localized to the peroxisomes (Fig. 4.48b and c, respectively). Localization of the Fragment 2-YFP fusion to the peroxisomes suggested the presence of an embedded PTS. YFP-Fragment 3 was localized to the cytoplasm indicating the inability of SLS to function as PTS1 (Fig. 4.48d).

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HcBZL S (a) PTS2 SLS

Truncation S PTS2 SLS

35S Fragment 1 YFP S 35S Fragment 2 YFP S 35S YFP Fragment 3 S PTS2 SLS SLS

b c d

Fig. 4.48: Subcellular localization of truncated HcBZL proteins fused to YFP. Schematic diagram of the fusion constructs (a) and the outcome of their transient expression in N. benthamiana leaf epidermis cells (b-d). Fragment 1-YFP and Fragment 2-YFP were targeted to peroxisomes (b and c, respectively). YFP-Fragment 3 was a cytoplasmic protein (d). Bars =10 μm. Upon co-localization, Fragment 1-YFP and Fragment 2-YFP present in peroxisomes (Fig. 4.49a and e) overlapped with the punctate cyan fluorescence of the peroxisomal marker (Fig. 4.49b and f). Superimposition resulted in white fluorescent dots (Fig. 4.49c and g), which were not superimposable with the autofluorescence of chlorophyll as individually shown in Fig. 4.49d and h.

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35S Fragment 1 YFP S 35S CFP PTS1 S PTS2 +

a b c d

35S Fragment 2 YFP S 35S CFP PTS1 S SLS +

e f g h

Fig. 4.49: Co-transformation of N. benthamina leaf epidermis cells with truncated HcBZL proteins fused to YFP and the CFP-PTS1 peroxisomal marker. Fragment 1-YFP (a-d); and Fragment 2-YFP (e-h). Schematic diagrams of the truncated HcBZL-YFP fusion and marker constructs are shown on the top panels. Yellow YFP fluorescence (a, e) and cyan CFP fluorescence (b, f). Co-localization of Fragment 1-YFP as well as Fragment 2-YFP with the peroxisomal marker protein led to overlapping punctate white fluorescence in the peroxisomes (c, g). Red indicates chlorophyll autofluorescence (d, h). Bars = 10 µm.

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5 Discussion

5.1 Xanthones in Hypericum cell suspension cultures Plants respond to biotic and abiotic stresses of the natural environment by the production of secondary metabolites as a part of their defense mechanism arsenal. This stress situation is mimicked in the case of in vitro cultures to either induce or enhance the formation of valuable secondary metabolites by the addition of elicitors (Namdeo, 2007; Shakya et al., 2019). Species of the genus Hypericum accumulate an array of pharmacologically significant secondary metabolites, mainly hyperforins, hypericins, flavonoids, and prenylated xanthones (Crockett et al., 2011; Beerhues, 2011; Tocci et al., 2018). Xanthones are polyketide scaffolds which play a dual role in plants subjected to biotic stress. They act as antioxidants protecting against oxidative damage and as phytoalexins impairing pathogen growth (Franklin et al., 2009). Lately, an orchestra of trials have been performed primarily on the root cultures derived from Hypericum species to evaluate their total xanthone content before and after elicitation following application of biotic and abiotic stressors or changing physical stimuli (Tocci et al., 2011; 2013a; Tusevski et al., 2013a, 2013b; Valletta et al., 2016; Badiali et al., 2018). Hairy roots of H. perforatum have emerged as a promising system for xanthone production by accumulating a total of 28 xanthones (Tusevski et al., 2013a). Different from xanthone producing root cultures, reports on the xanthone content of Hypericum cell suspension cultures are scarce.

H. perforatum sub-species angustifolium cell suspension cultures accumulate paxanthone constitutively (0.13 mg/g dry weight). Upon chitosan treatment (200 mg/l), an increase in the total xanthone content occurred (0.61 mg/g dry weight; Tocci et al., 2010). H. calycinum cell suspension cultures, on the other hand, did not show any constitutive production of xanthones prior to elicitor treatment (Gaid et al., 2012; current study). The total xanthone content observed in the present study was 4.7 mg/g dry weight after treatment, which is comparable to the previous report (4 mg/g dry weight; Gaid et al., 2012) and is 7.7-fold higher than that observed in chitosan-treated H. perforatum sub-species angustifolium cell suspension cultures (Tocci et al., 2010). A comparable xanthone content (~4 mg/g dry weight) was observed when H. perforatum cells were treated with A. tumefaceins and Colletotrichum gloeosporioides (0.25 g/l). The latter is the causative agent of anthracnose disease in Hypericum (Conceição et al, 2006; Franklin et al., 2009). Another report indicates that the highest total xanthone content in H. perforatum cell suspension cultures after treatment with salicylic acid (50 µM) was ~0.5 mg/g dry weight (Zubrická et al., 2015). A similar xanthone content was observed in H. androsaemum cells when cultivated in modified B5 medium (Schmidt et al., 2000a). Methyl jasmonate treatment of H. androsaemum cell suspension cultures led to improved xanthone content (~3.5 mg/g dry weight; Abd El-Mawla and Beerhues, 2002). Interestingly, cocultivation of A. tumefaciens or A. rhizogenes with H. perforatum cell suspension cultures for 21 days led to ~30-fold higher xanthone accumulation in the plant cells than reported for H. calycinum cells and 21 xanthones accumulated in the cultured cells (Tusevski et al., 2015). Based on the current study and previous

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Discussion reports it can be suggested that H. calycinum cell suspension cultures represent a promising system for enhanced xanthone production. It is suggested that cocultivation with Agrobacterium may improve xanthone formation both quantitatively and qualitatively.

The acetone extract of yeast extract-treated H. calycinum cells cultivated in LS medium was reported to contain hyperxanthone E (Gaid et al., 2012). According to a recent report yeast extract-treated H. calycium cells cultivated in a modified MS medium accumulated both hyperxanthone E and patulone (Nagia et al., 2019). The genes encoding the enzymes which catalyze the in vitro monopenylation and gem-diprenylation reaction leading to the formation of 1,3,6,7-tetrahydroxy-8-prenylxanthone and patulone (8,8-diprenyl-1,3,6,7- tetrahydroxyxanthone), respectively, were cloned from H. calycinum and H. sampsonii (Nagia et al., 2019). In the current study, methanolic extracts of yeast extract-treated H. calycinum cells contained three prenylated xanthones. The first compound was the monoprenylated xanthone phytoalexin hyperxanthone E and the second compound was the gem-diprenylated xanthone patulone, as was reported previously for H. calycinum cultured cells. Hyperxanthone E was first identified in the aerial parts of H. scabrum and also in twigs of Garcinia esculenta (Tanaka et al., 2004; Zhang et al., 2014). Patulone was first identified in H. patulum (Ishiguro et al., 1997) and later in H. sampsonii (Li et al., 2004). The third compound corresponding to peak 3 in Fig. 4.1a is a yet unidentified diprenylated xanthone. Putatively, the third peak can refer to γ-mangostin (1,3,6,7-tetrahydroxy-2,8-diprenylxanthone). However, sharing the same molecular ion m/z [M- H]- of 395 and major MS/MS fragments at m/z 351, 339, 283 does not guarantee identical structures. The same is true for their close UV absorption maxima. γ-Mangostin isomers have also been identified in H. perforatum hairy roots and in cell suspension cultures cocultivated with Agrobacterium (Tusevski et al., 2013a, 2013b, 2015). The possibility of the unidentified xanthone being monogeranylated cannot be excluded.

The biosynthesis of xanthones is catalyzed by a sequence of enzymes, two of which have been studied in the current work. HcBD and HcBZL belong to the superfamilies of aldehyde dehydrogenases and adenylate-forming enzymes, respectively.

5.2 ALDH [aldehyde: NAD(P)+ oxidoreductase]/ HcBD The majority of aldehydes found in a biological system are either derived from the metabolism of amino acids, lipids, carbohydrates, and vitamins or they are derivatives of xenobiotic metabolism. Biogenic aldehydes are relatively toxic molecules in the biological system. The electrophilic nature of the carbonyl group of an aldehyde is responsible for its high reactivity and interaction with cellular nucleophiles and therefore their level in the biological system must be regulated. NAD(P)+-dependent irreversible oxidation of endogenous and exogenous aromatic and aliphatic aldehydes to acids by ALDH is one of the most efficient ways of detoxifying biological system (Lindahl, 1992; Vasiliou et al., 2000; 2004).

The ALDH superfamily is evolutionarily ancient and members of this superfamily are present in archaea, eubacteria and eukarya emphasizing its importance throughout evolutionary history. As

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Discussion of the year 2002, 555 distinct ALDH genes were identified (Sophos and Vasiliou, 2003). Based on primary sequence identity ALDHs have been classified into different families and subfamilies (Vasiliou et al., 1999). The ALDH gene superfamily in eukaryotes has been classified into 24 families. Plants and animals share many ALDH families but also contain unique genes and families. Plant ALDHs are represented in 14 ALDH families namely, ALDH2, ALDH3, ALDH5, ALDH6, ALDH7, ALDH10, ALDH11, ALDH12, ALDH18, ALDH19, ALDH21, ALDH22, ALDH23 and ALDH24 with families ALDH2, ALDH3, ALDH5, ALDH6, ALDH7 and ALDH18 having mammalian orthologues (Zhang et al., 2012). ALDH19 is unique to S. lycopersicum, its orthologous genes have not been reported in any of the other studied plant genomes (Jimenez-Lopez et al., 2016). ALDH21 and ALDH23 are unique to mossess and ALDH 24 is unique to Chlamydomonas reinhardtii (Zhang et al., 2012; Brocker et al., 2013). While the model plant A. thaliana contained 16 ALDH genes (Kirch et al., 2004; Brocker et al., 2013), the P. triocarpa genome has 26 ALDH genes (Brocker et al., 2013; Tian et al., 2015). The human genome contains 19 ALDH genes in 11 ALDH families (Vasiliou and Nebert, 2005; Jackson et al., 2011). A possible explanation for the presence of so many ALDH genes is the necessity to provide ALDH activity in various subcellular compartments. While some small aldehydes (e.g. acetaldehyde) can move from one subcellular compartment to another, others may not as their molecular sizes and LogP values preclude passive diffusion across membranes. Presence of ALDH activity in various compartments containing aldehyde-generating pathways will protect the organelles from aldehyde-induced damages.

Perozich et al. (1999) aligned 145 ALDH sequences and presented 10 most conserved sequence motifs in the ALDH superfamily (Fig. 5.1; yellow). The 10 motif residues were located at or near the active site of the enzyme. It was observed that 16 amino acid residues were conserved in at least 95% of the ALDH sequences analyzed, out of which the following 4 amino acid residues were invariant: Gly187, Gly240, Glu333 and Phe335 (numbered as per rat class 3 ALDH/ALDH3). Fifteen of these amino acids including the 4 invariant residues are also conserved in HcBD, the enzyme studied here (Fig. 5.1, marked by asterisks). Instead of Arg25, HcBD contains Asn. The Asn residue appears to be conserved in all plant ALDH2 members mentioned in the alignment except for Z. mays RF2C (ZmRF2C) which contains an Asp residue. Of the 16 residues, 12 are clustered in 7 motifs as shown in Fig. 5.1. Despite the differences in sequence, substrate specificity and mode of oligomerization, all ALDHs have similar structural folds (Liu et al., 1997). HcBD shares ~52%, ~53% and ~28% identities with bovine ALDH2, human mitochondrial ALDH2 and rat ALDH3.

110

Discussion

BD BD BD BD BD BD BD BD. RF2C BD RF2C BD RF2C BD RF2C BD RF2C BD RF2C BD RF2C REF1 REF1 REF1 REF1 REF1 REF1 REF1 Hc At Zm Am Hc At Zm Am Hc At Zm Am Hc At Zm Am Hc At Zm Am Hc At Zm Am Hc At Zm Am R R R R E D D E T T T T P P P P E E E E A A A A

P P P P V A R V Q Q Q Q S S S S F F Y F L L L L

the the ten

D D D D M Y A Q E E E E K K K K Q Q Q Q G G G G 1. I I R L K K K A A A A A G G G G K R D D Y Y Y Y

T T T T T G G C P P P T G G G G K K K A K K R S L M L L B B * * E E E P F L V Q K K K K L L L L T D D D S T T S W W W W Motif 9 F F F F A Q A E V V V L E E E E V V V V N N N N P P P A T T T T F F F Y V V V V L L L L Q Q Q Q N N N N S S T P

K K K K L L L P M M V V S S S T P P P P A A A A N N N N It depicts G G G G K K K K T T A S V V V V G G G G R C K R Y H Y K

Motif 2

S S S S G G G G C C C N P K P P L Q Q Q K K Q R I L L L domain GXXXXG

L A V A A G A S * G G G G K K K K R R S E I I I I P P P A es es were introduced to

S A A A N D D D A A A C L L L L S A V M A G V V T M T T BD. D D D D I V V W A A A A N N N N R T S G E E E E V V V V

V I V V A A T A L M L L S S S S P S P A E E E D V V V V Hc F F F F D D D E A A A A L A E K D D N K I V V L T S T A E Q K Q L L L L P P P P A A A A F F F F T T T E K K K K

G G G G A K A A A A G G A A A S P P P P K K K K V T T V

binding - * N N D N A A A A V V V V A A A L D D D D F F F F Q Q Q Q I I I I I L L V K K K K Q Q R E G G G G K K K K L L L L F F F L E E E E V T F W M M M L T V V V M M M L Y Y Y Y L L L L E E E D F A F G I I V V V T V T L L L I K N K N K K K K A I V N F F F F K K L I W W S R S S A T Y D D K T T T D H N H H M M M M R R R K G D K K M M M Q L L L L Motif 8

F F F Y E E E K M I T V G G G G K K S L V V V V A A A S

family family including the invariant catalytic ne ne residue at the position corresponding K K R Q D E D E A S T L V V V T A A A A P P P P D D D Y

* was was suggested to be important for protein I I V V I I I V P P P P E D E E K K L R G G G G M M M E * E E E S L L L L F F Y F T T T T K E E A F F F F G G G G * Motif 4 * - P V V D D D D N N N N S S S S A V A K I I I I A S M K - L K P A A A A W W W W G G G G L L A A E E E E D E D E * *

P K P P F F F F P P P P T T T T T K K K D D D D R R K R

glutami

L V V K K K R R I I V I F F F F Q E K E Q Q Q K G C G G a V T E I M N H L I I I I S S S A V V V V A Y A A F N F I

Motif 1 and and the coenzyme L T F P M I L M H N H Q V V V L F V F F I I I I G G G G S A S E I L I I G G G G K K K K E K E E A K T L S S S S S G G E R K R K V V V A D D D D D D D D M M M M M M M M Motif 10 D N K L G A G Q V V V V I V V V Y Y Y Y D D D D K K K K * T C S A R R R R G G G G D D D D I I I V D E D D Y Y Y Y H K S A A E E E F I V I M M M M G G G K K T K K G G G G * G G G A S F Y Y P P P P H H H H E E E E V V V V G G G G

#

sidue marked sidue by marked # , , green) N N N A G G G A E E E E S S A R Q Q Q H D D D D Y Y F F

T - A A S T T P L K K H A A A C V V V V T A T S P P P P 302 - - - T M M M M L L L L I I V L F F Y F F F F F C C A I - - - G R R R K T T T T A A A A V V I T V I I V D D D A - - - Y P P P P Y Y Y Q A A A A R R R R T T T T D L P A

- - - R W W W W G G G V G G G G S S T S P P P P S D D D 16 amino acid residues which are conserved in at least 95% of the

- - - Y P P P P H F H H A A A A G S G G E Q E Q F F F F

- - - V G G G G F L M H T T T T A A A A I I I I A G A T rs of the aligned sequences sequences listed Appendix of rs are the in aligned Table the - - - G H H N E - S - P P S P P M V V C Y F Y Y L F F D Motif 7 * - - - R D D D D E Q R G G G G G C C C C Y Y Y Y Y Y Y F - - - G F F F F R R Q D Y F F F I I I C G G G G C C C C *

- - - L A A A A S T A A G G G G E E E Q K K K K N N N N isks are * - - - Y H Y E K M M M - N T P S G G G G D D D T I V I I - - - S R R R R K K K P I V V V K K K Q G G G G W W W W

- - - Q A A A A L L L I I I V I N N N N S I C L I I I V ALDH2. ALDH2. The alignment was done by CLUSTAL W. Dash

- - - K A A A A V T T T N N N N F Y T F A A P R M I A T

aster

Motif 6 * - - - G K N K A E E A L L L L L C F Y F K K K D G G G G - - - G V V V V G G G G V V V V V C T L G G G G S A A A - - - R A A A A H H H H G G G G A G A A G G G G R K R R - - - S L L L R I I V I D D A E L L F F T T T T I I I L Motif 3 - - - L D D D N K K K K P P P P L L N H L L V E S S S A

- - - L V V V I D D D D V I V L A A V A V L L L R R R R , , blue) and cysteine (C

- - - P D D D D A A A A G G G G L L L L T T T T S S S M 268 - - - V E E A E A A A W A A A A D D N E A A A A V V V M - - - S K K K A G G G G Q E E E V A V V G G G G T T T T - - - R D D D D A A A A K K R H A A A A E E E S N N N N - - - S G G G G Y N Y Y A S A F A K M K K N R E A I A A - - - L E E E E Y Y Y Y L L L L D D D D K K K A V L V T

- - - L A A A A R R R R H H H K V I L V G G G G N D D D ; ; red). Marked by - - - S V I I V L F L F A A A S D D D D L H I S L I I L

- - - S K T S H T H L L Y Y Y V A A A A E E D R D D N N 250

- - - F A A A A N G H R F F F L D D D D I I I I K Q K Q ences ences (Perozich et al. 1999). The histidine re

- - - R S I I I A A A V L L L L D N D E Y Y Y Y T S T T

glutamate glutamate (E in in human mitochondrial

H - - H V V V V V T A F A A A A F F F C A S R K V L V F

equ

D - T A E E E E S A G M S S S S V I V V L L L L I I I V

: : Multiple sequence alignment of selected ALDH2 family members including

Motif 5

A E A A G G G G P P P P L L L L I L I I V I V I G G G G

1

487 .

M M M M N N T S I I I I P S P P V L V F R K K K A A A A

STEVG

5 245

1 1 1 1 45 44 46 78 122 121 123 155 198 198 199 231 275 275 276 308 352 352 353 385 429 429 430 462

characteristic characteristic motifs (yellow) of the aldehyde dehydrogenase (ALDH) super residues (G ALDH s folding and/or assembly of monomeric subunits. Marked in grey is to Glu numbe accession The alignment. the optimize Fig.

111

Discussion

The ALDH superfamily consists of three catalytically important, diagnostic amino acids or domains, which are located within motif 4, 5 and 6. The first diagnostic domain of the ALDH superfamily is the coenzyme-binding signature, GXXXXG, of the Rossmann fold located in motif 4 (Fig. 5.1, red). It interacts with the nicotinamide moiety of the coenzyme (Hempel et al., 1993; Liu et al., 1997; Steinmetz et al., 1997). The second diagnostic amino acid residue in ALDH superfamily is the glutamic acid residue, which acts as a general base important for the activation of the catalytic cysteine residue, the acylation and deacylation steps (Wang and Weiner, 1995; Steinmetz et al., 1997; Tsybovsky and Krupenko, 2011). It is located in motif 5 (Fig. 5.1, blue). The highly conserved H235 residue of rat liver mitochondrial ALDH was suggested to be required for the proper folding and/or assembly of the newly synthesized polypeptides into the native conformation (Zheng and Weiner, 1993). This histidine residue is conserved in HcBD and other plant ALDHs shown in Fig. 5.1 (marked by #). The third diagnostic residue is the strictly conserved cysteine residue that acts as the active-site nucleophile, which forms a covalent bond with the aldehyde (Farrés et al., 1995; Tsybovsky and Krupenko, 2011). It is located in motif 6 (Fig. 5.1, green).

The general mechanism of ALDH catalysis involves two main steps. The first is acylation and the second is deacylation. In the first step, the invariant cysteine residue of the active site makes a nucleophilic attack on the carbonyl carbon of the aldehyde to form a thiohemiacetal intermediate. This is followed by hydride transfer from the aldehyde to the C-4 atom of the nicotinamide ring of NAD(P)+ leading to the conversion of the thiohemiacetal intermediate to a thioester intermediate and formation of NAD(P)H. In the second step of the reaction, an activated water molecule hydrolyzes the thioester releasing the acid product (Fig. 5.2). Steinmetz et al. (1997) reported the crystal structure of bovine mitochondrial ALDH2 in which the coenzyme is stabilized in the binding pocket through a combination of van der Waals interactions and hydrogen bonding. Lys192 interacts with 2' and 3' hydroxyl oxygens of the adenosine ribose of NAD+ and holds them in position. Similarly, Glu399 stabilizes the position of nicotinamide ribose through the formation of hydrogen bonds with the 2' and 3' hydroxyl groups. These two residues were crucial for the hydride transfer step (Ni et al., 1997; Sheikh et al., 1997). The side chain amide nitrogen of Asn169 and the peptide nitrogen of Cys302 stabilize the developing oxyanion in the thiohemiacetal transition state and orient the thiohemiacetal for optimal hydride transfer to the C-4 atom of NAD+ (Steinmetz et al., 1997). Both Lys192 and Glu399 are conserved in HcBD and other plant ALDH2 and are located in motif 2 and motif 8, respectively (Fig. 5.1).

In ~50% of the oriental population, the E487K point mutation in human mitochondrial ALDH2 is associated with impaired alcohol metabolism and acute alcohol intoxication (Yoshida et al., 487 1984). The E K mutation led to an increase in Km and reduction in Kcat of mitochondrial ALDH2 for NAD+. The E487K mutation occurred in the oligomerization domain and was located at a key interface between subunits immediately below the active site of another monomer, affecting catalytic activity and stability of the enzyme (Farrés et al., 1994; Xiao et al., 1996;

112

Discussion

Steinmetz et al., 1997). In planta, HcBD and other plant ALDH2 contain a glutamine residue (grey) at the position corresponding to Glu487 (Fig. 5.1).

Fig. 5.2: The mechanism of ALDH catalysis. 1, activation of the catalytic Cys302 through abstraction of a proton by Glu268 and an ordered water molecule. 2, nucleophilic attack on the carbonyl carbon of the aldehyde substrate by the thiolate functional group of the catalytic Cys302 . 3, formation of the thiohemiacetal intermediate stabilized by Asn169 and Cys302, followed by hydride transfer from the thiohemiacetal intermediate to the C-4 atom of the nicotinamide ring of NAD(P)+ held in proper position by Lys192 and Glu399. 4, hydrolysis of the resulting thioester intermediate mediated by proton abstraction by Glu268 and followed by acid product release. 5, dissociation of the reduced coenzyme followed by regeneration of the enzyme by NAD(P)+ binding (Adapted from Koppaka et al., 2012). In mammalian tissue, at least three major classes of ALDH have been identified. Class 1 ALDHs are commonly cytosolic, class 2 ALDHs are mitochondrial and class 3 ALDHs are found in both the cytosolic and microsomal fractions. Class 1 and 2 ALDHs prefer small aliphatic aldehydes while the physiological role of the class 3 ALDHs is oxidation of medium-chain saturated and unsaturated aldehydes produced during cellular lipid-peroxidation (Lihndahl and Petersen, 1991; Lihndahl, 1992).

Plant ALDHs are localized in various subcellular compartments including the cytoplasm (Li et al., 2000), mitochondria (Nakazono et al., 2000; Liu et al., 2001; Liu and Schnable, 2002; Long et al., 2009), peroxisomes (Mitsuya et al., 2009; Missihoun et al., 2011) and the chloroplasts (Stiti et al., 2011). The differential localization of ALDH enzymes hints at functional specialization and suggests that different compartments may require ALDH proteins with specific biochemical properties. Multiple plant ALDHs are involved in stress-regulated

113

Discussion detoxification pathways (Stiti et al., 2011). For example, A. thaliana ALDH10A8 and ALDH10A9 are stress-responsive genes which encode detoxification enzymes controlling aminoaldehyde levels (Missihoun et al., 2011; Zarei et al, 2016). N. tabacum ALDH2a (TobALDH2a) is suggested to carry out acetaldehyde detoxification and to participate in energy production during pollen development (op den Camp and Kuhlemeier, 1997) while the Oryza sativa ALDH2a is probably essential for the detoxification of acetaldehyde during reaeration after submergence (Nakazono et al., 2000). O. sativa betaine aldehyde dehydrogenase (BADH1) was suggested to be involved in acetaldehyde detoxification in the peroxisomes and O. sativa ALDH7B6 plays an important role in the maintenance of seed viability by detoxifying aldehydes generated by lipid peroxidation (Mitsuya et al., 2009; Shin et al., 2009).

Some ALDHs participate in biosynthetic pathways of secondary metabolites like A. thaliana REF1 and Brassica napus REF orthologs contributing to the formation of ferulate and sinapate soluble and wall-bound esters (Nair et al., 2004; Mittasch et al., 2013). The BD activity of crude protein preparations from H. androsaemum in vitro cultures contributes to xanthone biosynthesis (Abd El-Mawla and Beerhues, 2002). Similarly, the detected BD activities in crude protein preparations from P. pyrifolia and S. aucuparia in vitro cultures confirmed their participation in the biosynthesis of biphenyls (Gaid et al., 2009; Saini et al., 2017). The cDNA encoding a mitochondrial BD protein was cloned from A. majus (AmBD) which contributes to methyl benzoate production in vivo (Long et al., 2009).

The catalytically active forms of ALDH are homodimer and homotetramer (dimer of dimer; Yoshida et al., 1998; González-Segura et al., 2005). Nevertheless, a broad-range aldehyde dehydrogenase from Geobacillus thermodenitrificans appeared as a major band of 220 kDa and a minor band of 440 kDa on a native PAGE gel, indicating that the purified ALDH was present mainly as a tetramer while the minor band might be formed because of dimerization of tetramers. However, the dimerization of tetramer was not further investigated (Li et al., 2010). Plant ALDH2 family members are tetrameric enzymes. Accordingly, identified ALDH2 members Z. mays fertiliity restorer 2A (ZmRF2A), ZmRF2B, ZmRF2C, ZmRF2F and AmBD existed as tetramers in the native state as deduced by gel filtration experiments (Liu and Schnable, 2002; Long et al., 2009; Končitíková et al., 2015). A. thaliana REF1 was also suggested to be a tetramer (Nair et al., 2004). HcBD in its native state also exists as a tetramer as previously reported (Awadalah, 2015). Each monomeric subunit of an oligomer consists of three domains, namely the catalytic domain, the coenzyme-binding domain and the oligomerization domain (Liu et al, 1997; Steinmetz et al., 1997). The structural basis explaining the dimeric and tetrameric forms of ALDHs is not known. Previously it was thought that the additional amino acids present at the C-terminal end of dimeric ALDH3 but absent in tetrameric ALDH1 and ALDH2 might play a role in influencing subunit assembly. However, removal and addition of amino acid residues from the C-terminus of ALDH3 and ALDH1 or ALDH2, respectively affected only the enzyme stability but not the oligomeric state. It was shown that the number of hydrophobic residues is higher in the dimer-dimer contact area of ALDH1 compared to the equivalent region

114

Discussion of ALDH3 and is possibly the force that drives assembly (Rodriguez-Zavala and Weiner, 2002). Also, in Pseudomonas aeruginosa betaine aldehyde dehydrogenase (PaBADH) a cysteine residue at the dimer interface was found to be involved in holding the dimer and consequently the tetramer together (González-Segura et al., 2005). The dissociation of the tetrameric PaBADH into monomers led to a loss of activity suggesting that the minimum functional oligomer amongst ALDH superfamily members is probably a dimer.

ALDH activity may or may not be affected by the presence of univalent or divalent cations. BD activity detected in crude extracts of S. aucuparia was reduced by the addition of the chloride salt of divalent cations (Gaid et al., 2009). A similar phenomenon was observed for HcBD activity where the presence of a divalent cation in an incubation had a partial inhibitory effect on the enzyme activity. Addition of Zn2+, Mn2+ and Cu2+ (1 mM) in incubation caused a reduction in HcBD activity by 82%, 39% and 22%, respectively. Addition of univalent cations had no effect on the enzyme activity. Similarly, studies conducted on Xenopus laevis ALDH1A1 indicated that Zn2+, Mn2+, Mg2+ and Ca2+ ions are uncompetitive inhibitors for the enzyme- substrate complex with Zn2+ being the most potent inhibitor (Rahman and Yamauchi, 2006). However, the contrasting effect of Mg2+ on Saccharomyces cerevisiae ALDH1 and human ALDH2 were also presented (Wang et al., 1998). Possibly an inhibition mechanism as observed for X. laevis ALDH1A1 can explain the effect of divalent cations on HcBD activity. The majority of ALDHs including HcBD are active in the absence of univalent cation except for a handful (Valenzuela-Soto et al., 2003; Garza-Ramos et al., 2013). Although the enhancing role of the monovalent ion with few ALDHs is not clearly understood, it possibly stabilizes, through its coordination, the position of a loop involved in coenzyme binding and thus contributes to the optimal active site conformation for the coenzyme binding (Tylichová et al., 2010).

ALDHs show activity over a broad pH range. Similar to HcBD, the optimum pH observed for Z. mays RF2A and RF2C were 9 and 8, respectively (Liu and Schnable, 2002). A. majus BD was active over a broad pH range (pH 6 to 9) with optimum pH being 8 (Long et al., 2009). Recently, A. thaliana aminoaldehyde dehydrogenases (AMADH) exhibited high pH optima at 10.5 and 9.5-9.7 for AtALDH10A8 and AtALDH10A9, respectively (Zarei et al., 2016). Pisum sativum AMADH1 and AMADH2 had the highest 3-aminopropionadehyde conversion rate at pH 9.7 and 10.2, respectively. The catalytic cysteine (pKa = 8.0) is suggested to exist in thiolate form when the pH is significantly above pH 8, hence facilitating an efficient nucleophilic attack on the substrate (Tylichová et al., 2010). The pH and temperature optima for BD activities from the crude extracts of P. pyrifolia and S. aucuparia were 9.5 and 40˚C, respectively (Gaid et al., 2009; Saini et al., 2017). Vanillin dehydrogenase from Streptomyces sp. NL15-2K which exhibited 79% relative activity with benzaldehyde also displayed similar pH and temperature optima (9.5, 45˚C; Nishimura et al., 2018). Similarly, HcBD showed the highest activity at pH 9.5 and 50˚C.

Members of the ALDH2 family are NAD+-specific. The specificity is due to the presence of a conserved glutamic acid residue in motif 2 (Fig. 5.1) that forms hydrogen bonds with the 2' and 3' hydroxyl groups of adenine ribose of NAD+. However, it prevents the binding of the 2'-

115

Discussion phosphate group of NADP+ through electrostatic repulsion, thus making NAD+ the preferred and NADP+ the poorly accepted coenzyme (Stiti et al., 2014). Correspondingly, a glutamic acid residue was observed in motif 2 of HcBD, which explains its low activity with NADP+ (~20%) when the activity with NAD+ was set to 100%. This indicates that HcBD is also NAD+-specific. Similarly, Z. mays RF2C, RF2D, RF2E and RF2F possessing the conserved glutamic acid residue showed 0.5%-9% relative activity when NADP+ was used as a cofactor (Končitíková et al., 2015). Acceptance of NADP+ as cofactor was suggested to occur when there was a concomitant effect of weakening steric hindrance and elimination of the electrostatic repulsion force in the region surrounding the 2′-phosphate group of the adenine moiety (Stiti et al., 2014).

The plant ALDH superfamily consists of 14 families (Zhang et al., 2012). Phylogenetic analysis and functional characterization showed that HcBD is a NAD+-dependent aromatic ALDH belonging to ALDH2 family (Fig. 4.14). Multiple ALDH2 family members have been identified in a number of plant species. For example, 4 members were identified in P. trichocarpa (Brocker et al., 2013; Tian et al., 2015), 13 in M. domestica (Li et al., 2013), 3 in A. thaliana (Kirch et al., 2004), 6 in Z. mays (Jimenez-Lopez et al., 2010), and 5 in V. vinifera (Zhang et al., 2012; Brocker et al., 2013). The ALDH2 family consists of four subfamilies, namely ALDH2B, ALDH2C, ALDH2D and ALDH2E (Brocker et al., 2013). AmBD along with the benzaldehyde accepting mitochondrial fertility restorer from Z. mays (ZmRF2A; Liu and Schnable, 2002) belongs to the subfamily ALDH2B which comprises mitochondrial ALDHs (Fig. 4.14; Skibbe et al., 2002; Končitíková et al., 2015). AmBD was shown to contribute to methyl benzoate production in vivo. Even though the affinity of AmBD to benzaldehyde was high (Km = 1.37 µM), its catalytic efficiency was 4.8 times lesser than that with acetaldehyde (Long et al., 2009). Similarily, the catalytic efficiency of HcBD for the non-physiological substrate trans- cinnamaldehyde was 1.8 times higher than that with benzaldehyde. Despite its physiological association with normal anther development and restoration of cytoplasmic male fertility, the mitochondrial ZmRF2A was found to accept benzaldehyde in in vitro assays (Liu et al., 2001; Liu and Schnable, 2002). HcBD, on the other hand, belongs to the ALDH2C subfamily including the cytoplasmic ALDHs (Fig. 4.14). HcBD shares ~55% and ~70% identities with mitochondrial and cytoplasmic members of ALDH2, respectively. Z. mays RF2C, RF2D, RF2E and RF2F are other cytoplasmic ALDHs which accept benzaldehyde with Km being 83, 22, 12, and 146 µM, respectively. However, the catalytic efficiency of HcBD with respect to benzaldehyde was ~31, ~7, ~6 and ~233 times greater than that of RF2C, RF2D and RF2E and RF2F, respectively (Končitíková et al., 2015). Interestingly, the catalytic efficiency of HcBD was also 2.1 times greater compared to AmBD. The high conversion rate of benzaldehyde to benzoic acid might reflect the high demand of benzoic acid precursor for the biosynthesis of xanthones in Hypericum. RF2C, RF2D, RF2E and RF2F also accepted trans-cinnamaldehyde with Km values of 10, 69, 52 and 116 µM, respectively. Similar to HcBD, a distinct in vivo function of trans- cinnamaldehyde oxidation has not been assigned to these proteins (Končitíková et al., 2015). Earlier, AtREF1 and REF1 orthologs from B. napus were reported to oxidize sinapaldehyde and coniferaldehyde in vivo and contributed to the biosynthesis of cell wall-linked and soluble

116

Discussion sinapate and ferulate esters (Nair et al., 2004; Mittasch et al., 2013). Interestingly, HcBD shares 70% identity with these proteins and is phylogenetically close. However, it poorly catalyzes the oxidation of coniferaldehyde to ferulic acid (Fig. 4.7). Also, many ALDH2 are suggested to be involved in acetaldehyde detoxification pathways (op den Camp and Kuhlemeier, 1997; Nakazono et al., 2000). Nonetheless, the poor utilization of acetaldehyde by HcBD excludes its participation in such a pathway (Fig. 4.7).

5.3 Adenylate-forming enzymes Adenylating enzymes catalyze a reaction in which an otherwise unreactive carboxylic acid is condensed with ATP to form a reactive carboxylate adenylate intermediate and a pyrophosphate leaving group. This step is referred to as adenylation and it proceeds through a negatively charged pentavalent transition state. The same enzyme then catalyzes a second reaction in which a nucleophile reacts with the carboxylate adenylate to generate the desired product upon AMP release (Fig. 5.3; Schmelz and Naismith, 2009). In the case of CoA ligases, the attacking nucleophile is the thiol group of CoA and the second step is thioesterification. Formation of CoA thioesters from carboxylic acids activates the acid for further biotransformation reactions and also facilitates enzyme recognition. Previously, based upon the sequence analysis, the superfamily of adenylate-forming enzymes was divided into three major subfamilies including the adenylation domains of nonribosomal peptide synthetases (NRPS), acyl-CoA synthetases/ aryl-CoA synthetases and luciferase (Fulda et al., 1994). According to a recent classification based on the common adenylation chemistry, the adenylate-forming enzymes were classified into 3 classes. Class I comprises the former subfamilies and is also referred to as the ANL superfamily of adenylating enzymes (the acyl/aryl-CoA synthetases, the adenylation domain of NRPS, and the luciferases; Gulick, 2009), class II comprises aminoacyl-tRNA synthetases and class III comprises NPRS-independent siderophores (NIS). All three classes have different structures (Schmelz and Naismith, 2009). The class I adenylate-forming enzymes have a large N-terminal domain and a small C-terminal domain connected via a flexible hinge, the active site being located at the interface of the two domains (Conti et al., 1997; Bains and Boulanger, 2007; Wu et al., 2009; Hu et al., 2010). As per reports on 4-chlorobenzoate-CoA ligase, 4-chlorobenzoate and ATP bind entirely in the N-terminal domain. The nucleotide part of the CoA binds to the surface of the C-terminal domain and the pantetheine arm enters a tunnel formed by residues of both N- and C-terminal domains which leads to the adenylate intermediate binding site in the N-terminal domain. Flexibility at the hinge region is vital for alternation from adenylate-forming conformation to thioester-forming conformation (Wu et al., 2009).

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Discussion

Fig. 5.3: Two-step reaction mechanism of adenylate-forming enzymes. A, adenosine; AMP, adenosine monophosphate; ATP, adenosine triphosphate; PPi, inorganic pyrophosphate. Adapted from Schmelz and Naismith, 2009. Among plants, 4CL is a well studied representative of adenylate-forming enzymes. It is the last enzyme of the general phenylpropanoid pathway working in consort with phenylalanine ammonia lyase (PAL) and cinnamate 4-hydroxylase (C4H) to channel the carbon flux from primary metabolism to various pathways in secondary metabolism. It activates hydroxycinnamic acids into their corresponding thiol esters, which are then used in the biosynthesis of various phenylpropanoid-derived compounds (C3-C6) such as lignin, flavonoids, coumarins and wall- bound phenolics (Dixon and Paiva, 1995; Douglas, 1996). The affinity of bonafide 4CLs for trans-cinnamic acid is usually low (Ehlting et al., 1999). Another enzyme competing for trans- cinnamic acid is CNL. It is the first enzyme of the benzenoid pathway and channels trans- cinnamic acid to the biosynthesis of benzenoid compounds (C1-C6; Klempien et al., 2012; Gaid et al., 2012). Recently, a 4CL-like enzyme has been cloned from Scutellaria baicalensis which activates trans-cinnamic acid in vivo and contributes to flavone biosynthesis (Zhao et al., 2016).

5.3.1 Acyl/Aryl activating enzymes from H. calycinum cell cultures Sequence alignment of various adenylate-forming enzymes of class I, which also includes the aforementioned 4CLs and CNLs, revealed that the BOX I motif is the most conserved, being represented in all the members of ANL with a high degree of similarity. It is the ATP/AMP- binding signature motif (Fulda et al., 1994; Gulick, 2009). Mutation of the lysine residue in BOX I led to almost complete loss of activity in A. thaliana 4CL isoform 2 (At4CL2) and E. coli fatty acyl CoA synthetase FadD (Stuible et al., 2000; Weimar et al., 2002). BOX II is another conserved motif in which the cysteine residue was suggested to stabilize the protein structure or indirectly support the catalytic activity of 4CL and related enzymes. Removal of BOX II led to complete loss of activity in the mutant enzyme; however it was not clear if the loss of activity was due to loss of function associated with BOX II or because of structural perturbation (Becker- Andre et al., 1991; Stuible et al., 2000). BOX II is completely conserved in all plant 4CLs. HcBZL and HcAAE1 both contain the conserved BOX I and BOX II motifs (Fig. 5.4).

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Discussion

Twelve amino acids lining the substrate binding pocket of At4CL2 form a signature motif that determines the substrate specificity code for 4CLs (Schneider et al., 2003). Only three of the twelve residues were conserved in HcCNL, HcBZL and HcAAE1 (Fig. 5.4). A recent in silico analysis of 374 amino acid sequences of class I adenylate-forming enzymes revealed that the sequences contain five invariant residues and twenty-two and fifty-six residues were conserved in at least 80% and 60% of the aligned sequences, respectively (Clark et al., 2018).

BZL activates benzoic acid to benzoyl-CoA in an ATP-dependent two-step reaction (Fig. 5.5). RpBZL was previously cloned from benzoate degrading microbes and its reaction mechanism is well-studied. In microbes, the apparent Km value for benzoic acid ranged from 4.4 µM to 160 µM. Microbial BZLs function at an optimum temperature of 30˚C or 37˚C. The pH optimum ranged between 8 and 9. They are reported to exist as monomers as well as dimers (Geissler et al., 1988; Egland et al., 1995; Schühle et al., 2003; Kawaguchi et al., 2006; Bains and Boulanger, 2007; Thornburg et al., 2015). In planta, crude protein preparations exhibiting BZL activity have been reported though no BZL encoding cDNA has been isolated from these plants. Nevertheless, partially purified monomeric BZL from C. breweri flowers could convert benzoic acid (100%) and ortho-aminobenzoic acid (50%) to their CoA esters (Beurle and Pichersky, 2002b). The optimum temperature and pH for the BZL activity ranged from 30-40˚C and 7.2-8.4, respectively. Similarly, protein preparations of H. androsaemum displayed 42% activity with 3- hydroxybenzoic acid when the benzoic acid conversion was set to 100% (Abd El-Mawla and Beerhues, 2002). The temperature and pH used for the assay were 30˚C and 7.5, respectively. Also, crude protein from C. erythraea cell cultures possessed a 3BZL activity where 3- hydroxybenzoic acid was the preferred substrate (100%) and the relative activity with benzoic acid was 18%. Temperature and pH optima were 25-30˚C and 7, respectively (Barillas and Beerhues, 1997). Parially purified 3BZL from C. erythraea cell cultures had a temperature and pH optima of 35˚C and 7.5, respectively (Barillas and Beerhues, 2000). The optimum temperature for HcBZL activity (30˚C) is in the range previously reported for microbial and plant BZL activities. More than 90% HcBZL activity was observed in the pH range of 6-7.5 which is also in the range previously reported for BZL activities from plant crude extracts. Since both monomeric and dimeric BZLs have been reported earlier, it would be interesting to determine the native molecular mass and the oligomeric state of HcBZL.

Benzoate activation has also been detected as a side activity with some plant CNLs. Very recently, C. melo CNL (CmCNL) has been reported to activate benzoic acid with a relative activity of 35% of that with trans-cinnamic acid (Gonda et al., 2018). Likewise, the relative activity of A. thaliana benzoyloxyglucosinolate 1/CNL (AtBZO1/CNL) with benzoic acid was 27% compared to that with trans-cinnamic (Lee et al., 2012).

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Discussion

CoASH BZL BZL

AMP Benzoic acid ATP PPi Benzoyl-AMP Benzoyl-CoA

Fig. 5.5: Two-step reaction catalyzed by benzoate-CoA ligase (BZL). CoASH/CoA, coenzyme A; AMP, adenosine monophosphate; PPi, inorganic pyrophosphate. Adenylate-forming enzymes are strictly dependent on the presence of a divalent cation for enzyme activity and the majority of the enzymes preferred Mg2+. In adenylate-forming enzymes with bound ATP, the divalent cation neutralizes the charge on the ATP, stabilizes the negatively charged transition state and also neutralizes the inorganic pyrophosphate leaving group (Schmelz and Naismith, 2009). HcBZL activity is dependent on the presence of a divalent cation and is highest in the presence of Mn2+ followed by Mg2+ and Co2+. C. breweri BZL activity was also mainly dependent on Mg2+ and Mn2+. Co2+ could partially restore the enzyme activity (Beuerle and Pichersky, 2002b). Similarly, C. erythraea 3BZL activity was strictly dependent on Mg2+ and Mn2+ (Barillas and Beerhues, 2000).

HcBZL accepts benzoic acid, the substrate of interest and short-chain fatty acids to produce activated products. Substituted benzoic acids, trans-cinnamic acid and its substituted forms were not accepted by the enzyme as determined by the luciferase-, HPLC-DAD-, LC-MS- and spectrophotometry-based assays. Kinetic characterization of HcBZL indicates that the Km for benzoic acid is within the range that has been previously observed for microbial BZLs. The uniqueness of HcBZL lies in the fact that its Km values with benzoic acid, ATP and CoA are low (99.76, 14.12, and 42.29 μM, respectively) compared to all previously reported plant CNL- mediated benzoate activation. The catalytic efficiency of HcBZL with benzoic acid was 2.8 times greater than that of AtBZO1/CNL (Lee et al., 2012). Except for P. hydrida 4CL (Ph4CL), which was found to accept benzoic acid at a very low affinity (Km = 9008 μM; Klempien et al., 2012), no plant 4CL accepted or was tested with benzoic acid. The non-utilization of trans- cinnamic acid and its derivatives by HcBZL suggests that the enzyme neither participates in the general phenylpropanoid pathway like the 4CLs (Vogt, 2010) nor acts as CNL which activates trans-cinnamic acid and constitutes the first step of the CoA-dependent route for benzoic acid biosynthesis in plants (Klempien et al., 2012; Gaid et al., 2012; Lee et al., 2012). In H. perforatum, activated benzoic acid that is benzoyl-CoA is the precursor of xanthones (Schmidt and Beerhues, 1997), the recently isolated biphenyls (Bréard et al., 2018) and the volatile benzylbenzoate (Bertoli et al., 2011) however, the latter two group of compounds have not been reported in H. calycinum cell cultures. Xanthones were the sole benzoate-derived polyketides that were detected after yeast extract-treatment of these cell cultures (Gaid et al., 2012). Isobutyryl-CoA, the product of HcBZL incubation with isobutyric acid in vitro, is the precursor for hyperforin biosynthesis (Beerhues, 2006; Gaid et al., 2018). H. calycinum cell cultures have

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Discussion been previously tested for the presence of hyperforin, however, only traces of the compound were reported in the cells (~0.03 mg/g dry weight; Klingauf et al., 2005). A recent analysis of the aerial parts of wild H. calycinum plants reports 0.01 mg/g dry weight accumulation of hyperforin in the flowers while this compound could not be detected in the leaves and the stem (Cirak et al., 2016). Hyperforin accumulation is reported to occur in aerial parts of the intact plants, while xanthones are predominantly localized in the roots (Soelberg et al., 2007; Tocci et al., 2013b; 2018). Accordingly, FPKM data for the sequence fragment hpa_locus_733_iso_2_len_1456_ver_2 (MPGR database) which shares ~99% identity with HcBZL is highest in the roots (100%) and ~20-40% in all other tissues (Fig. 4.15).

Phylogenetic reconstruction clusters HcBZL and HcAAE1 in the same clade as CNLs and short- chain acyl-activating CoA ligases (Fig. 4.36). Humulus lupulus carboxyl CoA ligase 2 and 4 (HlCCL2 and HlCCL4) preferred the branched short-chain fatty acids isovaleric acid and isobutyric acid, respectively and were involved in bitter acid biosynthesis. HlCCL13 shared 97.9% sequence identity with HlCCL2. HlCCL3 accepted various short-chain fatty acids; however, the preferred substrates were propanoic and butyric acids. Similarly, A. thaliana AAE1 and 2 (AtAAE1 and AtAAE2) accepted short- and medium-chain fatty acids (C4-C6). The preferred substrate for AtAAE1 and AtAAE2 were butyric acid and isovaleric acid, respectively (Xu et al., 2013). AtAAE7 also preferred butyric acid (Shockey et al., 2003). AtAAE11, previously known to prefer hexanoic acid (Shockey et al., 2003), was clustered with the CNLs. AtAAE11 was tested only with straight-chain acids (C2-C14) and it is likely a trans-cinnamic acid-activating enzyme, whose function has not been identified yet. Additionally, AtAAE11 shared high homology with P. hybrida AAE, which was involved in floral volatile production through its possible trans-cinnamic acid activating function (Colquhoun et al., 2012). The aforesaid data along with the predicted peroxisomal localization suggest AtAAE11 to be a CNL. Possibly multiple gene duplication events followed by refunctionalization of genes have taken place over time leading to the functional diversity of proteins observed in the large clade consisting of CNLs, BZL and the short-chain acyl-activating CoA ligases. However, little is known about the in vivo function of the majority of the aforementioned AAEs.

5.3.2 Validation of HcBZL substrate-specificity through homology modelling and docking of substrates and intermediates To interpret the substrate preference of HcBZL, a series of tested substrates were docked in the HcBZL homology model (Fig. 5.6a), which was generated using the programs Chimera and Modeller. This work was done by Dr. Lutz Preu from the Institute of Medicinal and Pharmaceutical Chemistry, TU Braunschweig. The template for HcBZL modelling was RpBZL (PDB ID: 4EAT) in complex with the substrate benzoic acid. The crystal structure was determined by Thornburg et al. (2015). The template was chosen based upon the functional relatedness of the enzymes, as both enzymes activate benzoic acid while trans-cinnamic or 4- coumaric acids were not utilized. A similar approach was followed by Schneider et al. (2003) when the first homology model of plant 4CL (At4CL2) was generated.

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Discussion

Following HcBZL homology modelling, benzoic acid was docked in the substrate-binding pocket (SBP) of the model using the software ArgusLab 4.0. A reasonable pose comparable to that in the RpBZL SBP could not be achieved when benzoic acid was docked in HcBZL SBP (Fig. 5.6b). Interestingly, a new SBP in the vicinity seemed to be tailor-made for docking of benzoic acid (Fig. 5.6c). The following four hydrophobic residues lined the HcBZL SBP and interacted with the phenyl ring of the docked benzoic acid: Leu367, Ile395, Phe397, and Phe422 (Fig. 5.6d). A pi-stacking interaction appeared to exist between the phenyl rings of benzoic acid and the side chain of Phe397. The interplanar distance between the two phenyl rings was ~4.0 Å (Fig. 5.6d). A similar interplanar distance was observed in the crystal structure of P. tomentosa 4CL isoform 1 (PDB ID: 3NI2) between the parallelly arranged 4-hydroxyphenyl rings of 4-coumaric acid and the side chain of Tyr236 (Hu et al., 2010).

Fig. 5.6: 3D model of HcBZL (grey) generated by using RpBZL (PDB ID: 4EAT_A, blue) as a template. The 3D structure was calculated and visualized using the programs Modeller and Chimera, respectively (a). Molecular surface maps (mesh) highlight the favorable accommodation of benzoic acid in the hydrophobic surfaces of the HcBZL (b) and RpBZL (c) SBPs. 3D model of the HcBZL SBP showing the main hydrophobic amino acid residues lining the SBP (d). Both oxygen atoms of the carboxylate functional group of benzoic acid interacted through hydrogen bonds with the side chain OH group of Thr344. One of the oxygen atoms of the carboxylate functional group exhibited a further hydrogen bond to the side chain of Thr341, while the other oxygen atom formed a hydrogen bond with the backbone NH of Glu348 (Fig. 5.7a). The

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Discussion aforementioned amino acid residues also interacted with the benzoyl-AMP intermediate when it was docked in the HcBZL SBP (Fig. 5.7b, grey). In this case, the carbonyl of the ester moiety formed hydrogen bonding with the backbone of Glu348 and Trp349, whereas the α-phosphate of AMP interacted with the side chain of Thr344 and the side chain and the backbone of Thr341. Earlier, a comparable Glu residue in E. coli FadA was also suggested to be involved in ATP binding. Also, a Thr residue was suggested to play a role in AMP intermediate stabilization. Its mutation led to a major loss of enzyme activity (Weimar et al., 2002). Interestingly, in the case of Bacillus brevis PheA two conserved Thr residues interacted with the α-phosphate of AMP and coordinated binding of Mg2+-AMP together with an invariant Glu residue (Conti et al., 1997).

The 3-hydroxy group of the ribose moiety in benzoyl-AMP interacted with the side chains of Asp426 and Arg441 through hydrogen bonds (Fig. 5.7b). Previously, a comparable binding mode was observed in the RpBZL crystal structure complexed with benzoyl-AMP (PDB ID: 4ZJZ), where the ribose moiety interacted with Asp406 and Arg421 (Thornburg et al., 2015). A similar interaction was also made by an Asp residue in B. brevis PheA (Conti et al., 1997). Hexanoic acid, when docked in HcBZL SBP, oriented itself in a similar manner as benzoic acid by the formation of hydrogen bonds between the carboxylate functional group and the side chain OH group of Thr344 and the backbone of Glu348 and Trp349 (Fig. 5.7a). Docking of hexanoyl-AMP exhibited the equivalent binding pattern for the hexanoyl part of the molecule as was reported for the benzoyl part of benzoyl-AMP. The α-phosphate group is bound to side chain OH groups of Thr341 and Thr344, whereas the carbonyl group was hydrogen-bound to the backbones of Glu348 349 and Trp . The shape and the position of the C2-C6 carbons of the hexanoyl moiety of the AMP intermediate resembled the benzene ring of benzoyl-AMP in the SBP, however, the adenyl part was located in a slightly different manner with respect to benzoyl-AMP (Fig. 5.7c).

a b c

Fig. 5.7: Docking models of the HcBZL SBP with benzoic acid (pink, a) and hexanoic acid (yellow, a) as well as the intermediates benzoyl-AMP (grey, b) and hexanoyl-AMP (green, c). Docking was done using ArgusLab. Blue lines represent hydrogen bonds. Docking of hydroxylated derivatives of benzoyl-AMP (3-hydroxy-, 4-hydroxy-, 3,5-dihydroxy and 3,4,5-trihydroxybenzoyl-AMP; Fig. 5.8) revealed a specific binding pattern, which also comprised Thr341, Thr344, Glu348 and Trp349. In the case of 3-hydroxybenzoyl-AMP, Pro339 was

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Discussion additionally involved. Depending on the position of the hydroxyl group(s) on the benzene ring at least two of the aforementioned amino acid residues were involved in the formation of hydrogen bonds. This was also true for the 4-OH group of 4-coumaroyl-AMP.

Initially, it was assumed that the origin of the substrate specificity of HcBZL was based on the principle of size exclusion. It seemed reasonable that small ligands like propanoic, butyric, isobutyric or hexanoic acids might fit properly in the small binding pocket envisaged for benzoic acid, whereas the sterically more demanding ligands such as trans-cinnamic and 4-coumaric acids lead to unfavorable steric clashes. However, this notion was questioned upon the docking of 3-hydroxybenzoic acid in the SBP. It exhibited a similar binding pose as benzoic acid with an additional hydrogen bond between the 3-OH group and the side chain of Asp426, which should have favoured its binding. Since none of the substituted hydroxybenzoic acids were accepted by HcBZL in the luciferase-based activity assay, a second assumption was made according to which the acyl/aryl-adenylate governed the formation of thioesters products. As exemplified below (Fig. 5.8) the docking of several hydroxylated-AMP congeners including 4-coumaroyl-AMP exhibited a typical binding pattern based on the four residues Thr341, Thr344, Glu348 and Trp349. Interestingly, these residues are also essential for the binding of benzoic acid (excluding Trp349) and benzoyl-AMP. These four residues seem to form the fundamental docking motif for the binding and the correct positioning of the acids as well as their acyl/aryl-AMP derivatives in the SBP. Also, the docking results for the acyl/aryl-AMP derivatives suggested that this motif acts as a conformational switch. For the AMP-congeners of well-accepted substrates such as benzoyl- AMP or hexanoyl-AMP, it was found that the α-phosphate moiety was hydrogen-bound to the side chains of Thr341 and Thr344, while the carbonyl group of the ester moiety was bound to the backbone of Glu348 and Trp349. On the contrary, the hydroxylated AMP-congeners were bound with their OH groups to Glu348 and Trp349, which led to a change in the molecule's conformation in which the α-phosphate group was bent away from Thr341 and Thr344. As suggested previously the interaction of Thr with the α-phosphate of AMP is essential for stabilization of the AMP intermediate and for subsequent product formation (Conti et al., 1997; Weimar et al., 2002). So, it is proposed that the contact of the α-phosphate moiety of AMP with Thr341 and Thr344 of HcBZL is essential for the stabilization of the AMP-intermediate and the corresponding conformation is a prerequisite for the thioesterification. This proposition could also be corroborated through the docking of cinnamoyl-AMP. Since trans-cinnamic acid is not preferred by HcBZL it was expected that cinnamoyl-AMP would adopt a conformation similar to other hydroxylated-AMP congeners. Indeed cinnamoyl-AMP showed the typical inactive conformation like the hydroxylated congeners in which the α-phosphate moiety was bent away from Thr341 and Thr344.

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Discussion

a b

c d

Fig. 5.8: Docking models of the HcBZL SBP docked with 3-hydroxybenzoyl-AMP (pink, a); 3,5-dihydroxybenzoyl-AMP (grey, a); 4-hydroxybenzoyl-AMP (grey, b); 3,4,5- trihydroxybenzoyl-AMP (grey, c); cinnamoyl-AMP (pink, d) and 4-coumaroyl-AMP (green, d) intermediates. Docking was done with ArgusLab. Blue lines represent hydrogen bonds. 5.3.3 Dual localization of HcBZL The phenomenon that the protein product (or products) of a single gene are targeted to more than one location within the cell is known as dual targeting. Dual targeting can be achieved by the use of a single translation product or two translation products. Single translation products can be dual targeted either by using two separate targeting signals on different ends of the same protein or through the use of an ambiguous targeting signal, which directs the protein to two organelles. The use of two translation products is usually controlled at the mRNA level. This can either be the result of alternative transcription initiation or differential splicing resulting in two separate mRNAs encoding different targeting signals. Alternative translation initiation can also be utilized to produce two different proteins with different targeting signals (Carrie and Whelan, 2013). Other factors causing dual-targeting are gene duplication, leaky ribosomal read-through of stop codons, post-translational modifications of the targeting signals, insufficient targeting signals and the nature of protein folding (Freitag et al., 2012; Ast et al., 2013, Kalderon and Pines, 2014).

For peroxisomes, most known matrix proteins are targeted to the peroxisomal matrix by the presence of the tripeptide PTS1 located at the C-terminus of the proteins, while other matrix proteins carry a PTS2, which is a conserved nonapeptide embedded in the N-terminal domain (Gould et al., 1989; Swinkels et al., 1991). A study conducted by Hooks et al. (2012) indicated that multiple AAEs from A. thaliana contain both PTS1 and a chloroplast transit peptide (cTP) and are located in peroxisomes and plastids upon construction of N- and C-terminal GFP fusions,

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Discussion respectively. This work indicated that a clear distinction should be made between dually targeted proteins, which have the capability of being sent to two compartments but reside only in one, and dually localized proteins that truly reside in two compartments (Hooks et al., 2012). Dual localization in the cytoplasm and the peroxisomes has been formerly reported for A. thaliana glutathione reductase isoform 1 which is the first plant protein to be dually localized to the cytoplasm and the peroxisome. It contains an insufficient C-terminal PTS1 (TNL tripeptide) that causes weak peroxisomal targeting in vivo in onion epidermal cells and only cytoplasmic targeting in A. thaliana protoplasts (Kataya and Reumann, 2010). HcBZL is also dually localized in N. benthamiana leaf epidermal cells with high abundance in the cytoplasm and weak abundance in the peroxisomes. Upon stringent screening, peroxisomal localization of HcBZL was observed and it coincided with the peroxisomal marker protein. The peroxisomal targeting of HcBZL is attributed to the presence of a prototypical major PTS2 (RLx5HL; R51LASALSHL59) near the N terminus. The PTS2 domain of HcBZL follows all the criteria of a characteristic PTS2 domain (Reumann, 2004). The motif RLx5HL is reported to be sufficient for peroxisomal localization of a protein (Kato et al., 1996; 1998). Thus, it would be safe to say that the strength of the PTS2 is not the factor that determines the cytoplasmic localization of HcBZL. Post-translationally, peroxisomal matrix proteins are transported in the organelle in their native state. In the native configuration, HcBZL PTS2 may be present in a protein fold that is sparsely accessible to the PTS2 signal receptor, peroxin-7 (PEX-7; Braverman et al., 1997), leading to poor peroxisomal transport efficiency and HcBZL abundance in the cytoplasm. Alternatively, binding of an accessory protein that regulates the cytoplasmic and peroxisomal distribution of the protein may make the PTS2 signal inaccessible for recognition by PEX-7. It was postulated that truncation of HcBZL could improve the peroxisomal transport efficiency by making the PTS2 accessible to PEX-7. Indeed, the truncated HcBZL YFP fusion proteins displayed a better peroxisomal transport efficiency, which was reflected by the improvement in the number of the fluorescently labeled peroxisomes that could be visualized by cLSM. The results suggested that the PTS was accessible for recognition by the receptor after truncation. X-ray crystallographic analysis can shed light on the structure of the native HcBZL and the position and accessibility of the PTS2.

HcBZL Fragment 1 encompassing PTS2 (Fig. 4.48b) and Fragment 2 (Fig. 4.48c) containing an embedded PTS were localized to the peroxisomes. Fragment 2 consisting of 472 amino acids was predicted to be located in the peroxisomes by Cell-PLoc 2.0/ Plant-mPLoc and PSORT II 113 121 subcellular localization prediction tools. The nonapeptide K IIFVDHQL (KIx5QL) was identified as a targeting signal by PSORT II in the truncated protein. Indeed, KIx5QL fits in the PTS2 consensus reported for eukaryotic proteins (Petriv et al., 2004). However, it has not been included in elaborated reports on the plant PTS2 consensus (two major R[LI]x5HL and nine minor R[QTMAV]x5HL, RLx5H[IF], R[IA]x5HI; Reumann, 2004; Reumann et al., 2004). Thus, it is not clear if KIx5QL is the element that leads to the peroxisomal localization of Fragment 2. Formerly, few plant proteins without any predictable PTS1/PTS2 were also localized to the peroxisomes (Goyer et al., 2004; Reumann et al., 2009; Quan et al., 2010). Fragment 3 which is

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Discussion composed of 35 amino acids and terminates with a non PTS1 tripeptide (SLS, Fig. 4.48d) was localized in the cytoplasm and the nucleus of the cell. However, the nuclear localization of YFP- Fragment 3 fusion protein can be explained by its small size which facilitated its passive diffusion into the nucleus (Wang and Brattain, 2007).

5.4 In vivo participation of HcBD and HcBZL in the CoA-dependent non-β-oxidative route to synthesize xanthones in Hypericum Expression analysis indicated that the accumulation of HcBD transcript began 2-4 h after elicitation and was highest between 8-12 h, after which it gradually decreased until 36 h (Fig. 4.37). Similarly, HcBZL transcript up-regulation began 4 h post-elicitation and increased until 16 h after which a gradual decrease in the transcript level was observed until 48 h (Fig. 4.38). The increases in the HcBD and HcBZL transcript levels coordinated with the previously reported elicitor-induced transcript maxima of HcPAL (8 h), HcCNL (8 h), HcBPS (8 h), HcXS (8 h) and HcPTs (8-16 h; Gaid et al., 2012; Fiesel et al., 2015; El-Awaad et al., 2016; Nagia et al., 2019). Interestingly, the increases in HcBD and HcBZL transcripts preceded xanthone accumulation, which was highest at 36-48 h after elicitation (Fig. 4.2). This suggests that together with the HcPAL and HcCNL enzymes, HcBD and HcBZL direct the carbon flux from the CoA- dependent non-β-oxidative route towards the biosynthesis of prenylated xanthones. The results also support the view that in vivo formation and activation of benzoic acid and its subsequent utilization in the xanthone biosynthetic route occur in part, if not completely, via the activity of functional HcBD and HcBZL enzymes. RNA interference and down-regulation of HcBD and HcBZL can provide insight into the contributions that the two genes and their corresponding enzymes make towards the formation and activation of benzoic acid and subsequently xanthone formation.

5.5 Proposed subcelullar trafficking of benzoic acid from the CoA-dependent non-β- oxidative route to xanthone biosynthesis H. androsaemum cell cultures were reported to form benzoyl-CoA via the CoA-dependent non- β-oxidative pathway, unlike P. hybrida, which uses both the CoA-dependent β-oxidative and the CoA-independent non-β-oxidative pathways (Abd El-Mawla et al., 2002; Boatright et al., 2004). The CoA-dependent non-β-oxidative route in Hypericum begins with HcCNL which was previously identified to be located in the peroxisomes of N. benthamiana leaf epidermis cells (Gaid et al., 2012). The molecular identification of the enzyme catalyzing the next step that is the intermediate conversion of cinnamoyl-CoA to benzaldehyde is still to be accomplished, leaving its subcellular localization an open question. On conducting subcellular localization of HcBD, this enzyme catalyzing the conversion of benzaldehyde to benzoic acid was localized to the cytoplasm. Subcellular localization experiments with fluorescent HcBZL fusion proteins indicated that the enzyme was predominantly located in the cytoplasm of N. benthamiana epidermis cells. The current results suggest that benzaldehyde, which is possibly formed in the peroxisome by CHL activity, moves across the peroxisomal membrane by diffusion. The small size and the logP value of ~1.5 of benzaldehyde support this hypothesis. Once in the cytoplasm,

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Discussion benzaldehyde is oxidized by the cytoplasmic HcBD into benzoic acid, which is then activated by cytoplasmic HcBZL to benzoyl-CoA. BPS, a cytoplasmic enzyme, later accepts benzoyl-CoA to form benzophenone, which is the direct precursor for xanthone biosynthesis (Fig. 5.9; El-Awaad et al., 2016). Movement of cinnamoyl-CoA out from the peroxisomes into the cytoplasm of the cell would require a transporter as the presence of the polar CoA moiety precludes its passive diffusion across the membrane. Also, it will be an energetically wasteful process as cytoplasm already possesses 4CLs that can activate trans-cinnamic acid albeit at small percentages. This again suggests that CHL is possibly a peroxisomal enzyme acting on cinnamoyl-CoA formed in the peroxisome to yield benzaldehyde.

Cytoplasm Peroxisome

CHL CNL

Benzaldehyde Cinnamoyl-CoA Cinnamic acid

BD BPS BZL + 3X Benzaldehyde Benzoic acid Benzoyl-CoA Malonyl-CoA 2,4,6-Trihydroxybenzophenone

1,3,7-TXS

1,3,6,7-Tetrahydroxy 1,3,7-Trihydroxy PTs X6H Prenylated xanthone xanthone xanthone Chloroplast Endoplasmic reticulum

Fig. 5.9: Proposed biosynthetic pathway depicting the contributions of HcBD and HcBZL to xanthone formation proceeding via the CoA-dependent non-β-oxidative pathway route of benzoic acid biosynthesis. Trafficking of intermediates and localization of reactions among subcellular compartments are indicated. BD, benzaldehyde dehydrogenase; BPS, benzophenone synthase; BZL, benzoate-CoA ligase; CHL, cinnamoyl-CoA hydratase-lyase; CNL, cinnamate- CoA ligase; 1,3,7-TXS, 1,3,7-trihydroxyxanthone synthase; X6H, xanthone 6-hydroxylase; PTs, prenyltransferases. The broken grey arrow represents steps that have not yet been reported at the molecular level.

129

Discussion

5.6 Perspectives H. calycinum cell cultures treated with yeast extract accumulate three prenylated xanthones which yield a total xanthone content of 4.7 mg/g dry weight after two days of treatment. Prenylated xanthones possess antibacterial and antifungal activities against pathogens which either attack plants or cause infections in humans (Larcher et al., 2004; Franklin et al., 2009; Dharmaratne et al., 2013; Tocci et al., 2013a; 2013b). H. calycinum cell suspension cultures can serve as a substitute of the widely used H. perforatum plant for enhanced production of prenylated xanthones in an upscaled bioreactor system evading bottlenecks associated with the chemical synthesis of prenylated xanthones. Previously, cell suspension cultures of Panax ginseng were cultivated in bioreactors to increase the ginsenoside content (Langhansová et al., 2005) and cell suspension cultures of Orostachys cartilaginous, a plant with medicinal value have recently been cultivated in a bioreactor system to enhance the production of bioactive phenolic compounds (Piao et al., 2017).

The biosynthesis of the xanthone core has been completely elucidated at the molecular level in Hypericum. However, little genetic information was available on the upstream pathway i.e. CoA- dependent non-β-oxidative route supplying the starter substrate benzoyl-CoA. Molecular information assembled from the current study on HcBD and HcBZL along with sequence information on other enzymes of the xanthone biosynthetic pathway will allow for future engineering of the whole pathway in heterologous hosts to create a bioplatform for producing structurally complex xanthones.

HcBD and HcBZL can be downregulated in Hypericum in order to understand their in vivo contribution and the underlying regulation of the two enzymes in xanthone biosynthesis. Accordingly, other related or more distant members of the ALDH and AAE families contributing to xanthone biosynthesis could be identified. Using a homology-based cloning approach sequence information of HcBD and HcBZL can also be used to identify similar sequences from other Hypericum species for which transcriptomics data are not available.

Homology modelling of HcBZL has identified amino acids that may affect the enzyme activity. Site-directed mutagenesis can be used to ascertain the function of these amino acids and to design sequences that can activate different acids for the production of novel pharmacologically uncharted polyketide scaffolds. Furthermore, the crystal structure of HcBD and HcBZL will help in understanding their molecular architectures and the evolution of their catalytic mechanisms in planta.

130

Summary

6 Summary • Xanthones are polyphenolic compounds found in fungi, lichens and higher plants. They may contribute either individually or through synergism with other secondary metabolites to the overall medicinal effect useful for the treatment of diseases. In plants, they provide protection against oxidative damage and pathogen attack. They are mainly distributed in the Clusiaceae, Hypericaceae, and Gentianaceae families. • A model system for studying xanthone biosynthesis is provided by Hypericum in vitro cultures. H. calycinum cell suspension cultures accumulate prenylated xanthones upon elicitor-treatment. The majority of the molecular players involved in the biosynthesis of prenylated xanthones have been identified from H. calycinum, making it a suitable system for studying the biosynthesis of benzoyl-CoA, the starter molecule required for xanthone biosynthesis. • In Hypericum, benzoyl-CoA formation occurs through the CoA-dependent non-β- oxidative route established at the biochemical level. It involves the sequential activity of four enzymes, namely CNL, CHL, BD and BZL. H. calycinum CNL is located in the peroxisomes. The molecular information on and the location of the remaining enzymes in the plant cell remain elusive. • Metabolic profiling of H. calycinum cultures before and after yeast extract treatment indicated the accumulation of three prenylated xanthones in the treated cultures only. Apart from the previously reported hyperxanthone E and patulone, an unidentified diprenylated xanthone was detected in the methanolic extract of the treated cultures. The total xanthone content was highest between 36 and 48 h of treatment. • Following the screening of H. perforatum transcriptomes, two sequences were successfully cloned and heterologously expressed in E. coli BL21 cells. His6-tag protein purification on Ni-NTA yielded the pure recombinant proteins which were referred to as HcBZL and HcAAE1 with molecular masses of ~60 and ~62 kDa, respectively. A BD sequence previously cloned by Dr. Mariam Gaid was also heterologously expressed, yielding a recombinant protein of ~54 kDa in SDS-PAGE. • Determination of the substrate specificity of HcBD by HPLC-DAD analysis indicated its preference for trans-cinnamaldehyde and benzaldehyde. Substituted trans- cinnamaldehyde and benzaldehyde derivatives were poorly converted by the enzyme. Acetaldehyde was also poorly accepted by HcBD as determined by a spectrophotometric assay. The preference for trans-cinnamaldehyde and benzaldehyde and the identities of the respective products were confirmed by GC-MS analysis. • Preliminary determination of the substrate specificities of HcBZL and HcAAE1 by HPLC-DAD and spectrophotometric analysis revealed that HcBZL preferred benzoic acid and C3-C7 fatty acids. HcBZL did not activate substituted benzoic and (hydroxy)cinnamic acid derivatives. HcAAE1, on the other hand, did not activate any aromatic acid and preferred C3-C4 fatty acids.

131

Summary

• HcBZL accepted the physiological starter substrate for xanthone biosynthesis and was tested with 24 potential substrates including benzoic acid and its substituted derivatives, trans-cinnamic acid and its substituted derivatives, fatty acids and other miscellaneous acid substrates using a luciferase-based substrate specificity assay. Production of recombinant luciferase protein and optimization of its utilization in the assay were done in-house. Of the 24 tested substrates, benzoic, propanoic, butyric, isobutyric and hexanoic acids were preferred by the enzyme. • The luciferase-based substrate specificity results were validated by docking various substrates and reaction intermediates in a HcBZL homology model established by Dr. Lutz Preu, Institute of Pharmaceutical and Medicinal Chemistry. The results indicated two different conformations of AMP-intermediates in the active site and suggested that the contact of the α-phosphate of the AMP-intermediate with Thr341 and Thr344 of HcBZL is essential for the stabilization of the intermediate and thereby for the next conversion of the two-step reaction, that is thioesterification. • The reaction conditions for the two enzymes were optimized with respect to various reaction parameters. Kinetic characterization of the two enzymes with respect to various substrates and co-substrates was done using the Hyperbolic regression analysis. • The affinities of HcBD for benzaldehyde and trans-cinnamaldehyde were similar. + Similarly, the Km values for NAD measured in the presence of benzaldehyde and trans- cinnamaldehyde were comparable. However, the turnover rate of HcBD for trans- cinnamaldehyde was 1.6 times higher than that for benzaldehyde, resulting in a 1.8-fold difference in the catalytic efficiencies. • The catalytic efficiencies of HcBZL for benzoic and isobutyric acids were similar. However, they were ~3.5-fold lesser than those for propanoic and hexanoic acids and ~6- fold lesser than that for butyric acid. The Km values for the co-substrates ATP and CoA in the presence of either benzoic acid or isobutyric acid were similar. • An up-regulated and coordinated increase in the expression of HcBD and HcBZL was observed between 8-12 and 12-16 h after yeast extract treatment, respectively, which preceded the maximum xanthone accumulation in the cells (36-48 h), strongly suggesting the in vivo involvement of HcBD and HcBZL in xanthone biosynthesis. • Subcellular localization experiments using fluorescent protein fusions revealed the cytoplasmic localizations of HcBD and the dual cytoplasmic and peroxisomal localization of HcBZL. HcBZL was mainly found in the cytoplasm. Owing to the presence of a functional prototypical PTS2, it was also weakly detected in the peroxisomes. No PTS1 was present in HcBZL. Using the previously published data and aforementioned results, it was possible to depict the contributions of HcBD and HcBZL to xanthone biosynthesis including the possible distribution of the reactions among various subcellular compartments.

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Appendix

8 Appendix

A Figures

0.6 0.5 0.4 0.3 0.2

Dry biomass (g) biomass Dry 0.1 0 0 2 4 6 8 10 12 14 16 Time (day)

Fig. A1: Growth curve of H. calycinum cell suspension cultures. The arrow marks the end of log phase and the time point selected for the treatment with yeast extract. Data are means ± SD of three biological repeats.

500 2500 y = 0.3019x y = 1.299x 400 R² = 0.9953 2000 R² = 0.9965 300 1500

200 1000 AUC at 292 nm 292 at AUC AUC at 254 nm 254 at AUC 100 500 0 0 0 500 1000 1500 0 500 1000 1500 Hyperxanthone E (ng) Patulone (ng)

Fig. A2: Calibration curves for xanthone references in HPLC analysis.

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1200 1200 400 y = 8.4483x y = 9.2479x y = 2.6653x 1000 1000 R² = 0.9999 R² = 1 R² = 0.9989 nm 300 800 800

600 280 600 200 400 400

100

AUC at at AUC nm 295 at AUC AUC at 230 nm 230 at AUC 200 200 0 0 0 0 20 40 60 80 100 0 20 40 60 80 100 0 20 40 60 80 100 Benzoic acid (ng) trans-Cinnamic acid (ng) 3-Hydroxybenzoic acid (ng)

2500 1200 600 y = 2.091x y = 10.211x y = 10.622x 1000 500 2000 R² = 0.9972 R² = 0.9999 R² = 1 800 400 1500 600 300 1000 400 200

500

AUC at 295 nm 295 at AUC nm 254 at AUC AUC at 322 nm 322at AUC 200 100 0 0 0 0 50 100 150 200 0 100 200 300 400 500 0 10 20 30 40 50 4-Hydroxy-3-methoxycinnamic acid 2-Hydroxybenzoic acid (ng) 4-Hydroxybenzoic acid (ng) (ng) 600 200 1000 y = 4.7548x y = 2.7867x y = 6.6632x 500 800 R² = 0.9977 R² = 0.9984 150 R² = 0.9999 400 600 100 300 400 200 50

200 nm 254at AUC AUC at 254 nm 254 at AUC AUC at 295 nm 295 at AUC 100 0 0 0 0 10 20 30 40 50 0 20 40 60 80 100 0 20 40 60 80 100 2-Methoxybenzoic acid (ng) 3,4-Dihydroxybenzoic acid (ng) 4-Hydroxy-3-methoxybenzoic acid (ng)

Fig. A3: Calibration curves for acid references in HPLC analysis.

2500 4000 y = 2.3105x y = 2.0429x 2000 R² = 0.9984 3000 R² = 0.9998 1500 2000 1000 1000

AUC at 261 nm 261at AUC 500 AUC at 261 nm 261at AUC 0 0 0 200 400 600 800 1000 0 500 1000 1500 Benzoyl-CoA (ng) Isobutyryl-CoA (ng)

Fig. A4: Calibration curves for CoA thioester references in HPLC analysis.

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120 100 80 60

40 BD activity BD activity (%)

Hc 20 0 6.5 7 7.5 8 8.5 9 9.5 10 10.5 11 pH

Fig. A5: Effect of pH on HcBD activity determined by spectrophotometric assay. Data are means ± SD of three biological repeats.

25 In-house Luciferase

20 Sigma Luciferase 3 15

10 CPS x 10x CPS 5

0 0 0.2 0.4 0.6 0.8 1

Amount of luciferase (µg) Fig. A6: Efficiencies of purified in-house recombinant luciferase and commercial luciferase (Sigma) for application in the luciferase-based substrate-specificity assay. The cps level produced by 0.5 µg in-house luciferase was comparable with that produced by 1 µg of commercial luciferase, as previously utilized in assays (Schneider et al., 2005; Gaid et al., 2012).

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(a)

(b)

(c)

Fig. A7: Melt curves for various amplified products. Histone 2A, Tm = 78˚C (a); Actin, Tm = 80˚C (b); HcBZL, Tm = 83˚C (c).

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Appendix

283.1 6.7e6 6.5e6

6.0e6

5.5e6

5.0e6 [M-H]- 327.2 4.5e6

4.0e6

3.5e6

3.0e6 Intensity, cps 2.5e6

2.0e6

1.5e6 258.1 272.2 1.0e6 242.9 256.2 230.1 295.2 5.0e5 311.2

0.0 60 80 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 400 m/z, Da Fig. A8: ESI-MS/MS analysis of hyperxanthone E with a molecular ion [M-H]– at m/z 327.2.

326.2

5.5e6

5.0e6

4.5e6

4.0e6

3.5e6

3.0e6

Intensity, cps Intensity, 2.5e6 [M-H]- 395.3 2.0e6

1.5e6

283.1 1.0e6

309.2 5.0e5 271.0

0.0 60 80 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 400 m/z, Da Fig. A9: ESI-MS/MS analysis of patulone with a molecular ion [M-H]– at m/z 395.3.

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283.1 1.10e7 1.05e7 1.00e7 9.50e6 9.00e6 8.50e6 8.00e6 7.50e6

7.00e6 351.2 6.50e6 6.00e6 5.50e6 5.00e6

Intensity, cps Intensity, 4.50e6 4.00e6 3.50e6 [M-H]- 3.00e6 271.1 395.2 2.50e6 297.2 309.2 2.00e6 270.1 1.50e6 281.1 311.2 339.2 307.2 1.00e6 254.9 242.2 325.3 177.1 363.2 379.3 5.00e5 226.9 0.00 60 80 100 120 140 160 180 200 220 240 260 280 300 320 340 360 380 400 m/z, Da Fig. A10: ESI-MS/MS analysis of an unidentified xanthone with a molecular ion [M-H]– at m/z 395.2

408.3 1.20e6 1.15e6 1.10e6 1.05e6 1.00e6 9.50e5 9.00e5 8.50e5 Molecular Weight: 823.59 8.00e5 Propanoyl-CoA 7.50e5 426.2 475.5 7.00e5 6.50e5 6.00e5 5.50e5

Intensity, cps 5.00e5 - 4.50e5 742.8 [M-H] 4.00e5 822.4 3.50e5 3.00e5 159.1 395.3 2.50e5 493.4 2.00e5 328.4 134.1 273.1 1.50e5 1.00e5 79.0 687.4 5.00e4 0.00 50 100 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 m/z, Da Fig. A11: ESI-MS/MS of HcBZL-formed propanoyl-CoA with a molecular ion [M-H]- at m/z 822.4.

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408.4 4.6e6

4.4e6

4.2e6

4.0e6

3.8e6

3.6e6

3.4e6

3.2e6 Molecular Weight: 837.62 Butyryl-CoA 3.0e6 489.4 2.8e6 426.4

2.6e6 - 2.4e6 [M-H] 836.3 2.2e6

2.0e6 Intensity, cps 1.8e6

1.6e6 756.7 1.4e6

1.2e6 159.0 507.5 1.0e6

8.0e5 134.0 328.3

6.0e5 273.1 4.0e5 701.6 738.5 818.8 2.0e5

0.0 50 100 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 m/z, Da Fig. A12: ESI-MS/MS of HcBZL-formed butyryl-CoA with a molecular ion [M-H]- at m/z 836.3.

407.9 1.6e6

1.5e6

1.4e6

1.3e6

1.2e6

1.1e6

1.0e6 Molecular Weight: 837.62 489.2

cps 9.0e5 Isobutyryl-CoA , ,

8.0e5 425.9

Intensity 7.0e5 158.9

6.0e5 756.1

5.0e5 134.0 272.9 328.0 - 4.0e5 [M-H] 507.2 3.0e5 836.0

2.0e5 79.0 346.0 738.4 1.0e5

0.0 50 100 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 m/z, Da Fig. A13: ESI-MS/MS of HcBZL-formed isobutyryl-CoA with a molecular ion [M-H]- at m/z 836.0

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Appendix

408.1 3.2e6

3.0e6 [M-H]- 2.8e6 864.3 2.6e6

2.4e6 Molecular Weight: 865.67 Hexanoyl-CoA 2.2e6 426.2

2.0e6 517.1

1.8e6 , , cps 1.6e6 784.4

Intensity 1.4e6

1.2e6

1.0e6 535.3

8.0e5

158.9 6.0e5 437.3 133.9 328.0 4.0e5 273.1 729.3 488.0 766.4 846.5 2.0e5

0.0 50 100 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 950 m/z, Da Fig. A14: ESI-MS of HcBZL-formed hexanoyl-CoA with a molecular ion [M-H]- at m/z 864.3.

408.2 3.0e6

2.8e6

2.6e6

2.4e6

2.2e6 Molecular Weight: 879.70 426.1 [M-H]- 2.0e6 Heptanoyl-CoA 531.5 878.4 1.8e6

1.6e6 , , cps

1.4e6 Intensity

1.2e6 798.9

1.0e6

8.0e5 549.4 158.9

6.0e5

328.1 4.0e5 134.0 451.5 273.1 618.4 743.7 2.0e5

0.0 50 100 150 200 250 300 350 400 450 500 550 600 650 700 750 800 850 900 m/z, Da Fig. A15: ESI-MS of HcBZL-formed heptanoyl-CoA with a molecular ion [M-H]- at m/z 878.4.

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Appendix

Benzaldehyde 2-Hydroxy- 3-Hydroxy- 4-Hydroxy- 3,4-Dihydroxy- benzaldehyde benzaldehyde benzaldehyde benzaldehyde (protocatechuic aldehyde)

4-Hydroxy- trans- 4-Hydroxy- 2-Methoxy- Acetaldehyde 3-methoxy- Cinnamaldehyde 3-methoxy- benzaldehyde benzaldehyde cinnamaldehyde (2-Anisaldehyde) (Vanillin) (Coniferaldehyde)

Fig. A16: Substrates used for the HcBD substrate specificity determination.

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Appendix

Benzoic acid and its substituted forms (C6-C1)

Benzoic acid 2-Hydroxybenzoic 3-Hydroxybenzoic 4-Hydroxybenzoic 2,3-Dihydroxybenzoic acid acid acid acid

3,5-Dihydroxybenzoic 3,4,5-Trihydroxybenzoic 4-Hydroxy-3-methoxybenzoic 2-Aminobenzoic acid acid acid acid

trans-Cinnamic acid and its substituted forms (C6-C3)

trans-Cinnamic 4-coumaric acid Caffeic acid Ferulic acid Sinapic acid acid Fatty acids

Acetic acid Propanoic acid Butyric acid Isobutyric acid Hexanoic acid Octanoic acid

Miscellaneous substrates

Shikimic acid (C6-C1) L-Phenylalanine (C6-C3) Malic acid (Dicarboxylic Mandelic acid (C6-C2) acid)

Fig. A17: List of substrates used in the luciferase-based substrate-specificity assay for HcBZL

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Appendix

B Tables

Table B 1: Accession numbers and names of functional ALDH2 family members used for the phylogenetic reconstruction of HcBD.

Annotation Alternative Source Accession number name NtALDH TobALDH2A/ Nicotiana tabacum CAA71003.1 ALDH2B2 AmBD BALDH Antirrhinum majus ACM89738.1 AtREF1 ALDH2C4 Arabidopsis thaliana Q56YU0.2 ZmRF2A ZmALDH2- AHM26656.1 1/ALDH2B2 ZmRF2B ZmALDH2- AHM26657.1 2/ALDH2B5 ZmRF2C ZmALDH2- AHM26658.1 3/ALDH2C1 Zea mays

ZmRF2D ZmALDH2- AIV00510.1 4/ALDH2C2 ZmRF2E ZmALDH2- AIV00511.1 5/ALDH2C4 ZmRF2F ZmALDH2- AHM26659.1 6/ALDH2C5 HcBD* - Hypericum MK988622 calycinum BnREF1_I - Brassica napus CBP94209.1

BnREF1_II - CBP94210.1 RnALDH2# ALDH-E2 Rattus norvegicus P11884.1 ALDH, aldehyde dehydrogenase; BD, BALDH, benzaldehyde dehydrogenase; REF1, reduced epidermal fluorescence 1; RF, fertility- restorer. * Sequence is represented by its nucleotide accession number # Mammalian sequence served as an outgroup

166

Appendix

Table B 2: Functional AAEs used for phylogenetic reconstruction of HcBZL and HcAAE1.

Sequence name Species Accession no. AtAAE1 Arabidopsis thaliana OAP16550.1 AtAAE2 OAP16179.1 AtAAE7 Q8VZF1.1 AtAAE11 Q9C8D4.1 HlCCL1 Humulus lupulus AGA17918.1 HlCCL2 AGA17919.1 HlCCL3 AGA17920.1 HlCCL4 AGA17921.1 HlCCL13 AGA17930.1 At4CL1 Arabidopsis thaliana Q42524.1 At4CL2 AAD47193.1 At4CL3 AAD47194.1 Sa4CL1 Sorbus aucuparia ADF30254.1 Sa4CL2 ADE96996.1 Sa4CL3 ADE96997.1 HcCNL Hypericum calycinum AFS60176.1 PhCNL Petunia hybrida AEO52693.1 Ph4CL1 AEO52694.1 AtBZO1/CNL Arabidopsis thaliana Q9SS01.1 CbCNL/BZLa Clarkia breweri AEO52695.1 CmCNL Cucumis melo XP_008463174.1 Rg4CL1 Ruta graveolans ABY60842.1 Rg4CL2 ABY60843.1 HcBZLb Hypericum calycinum MK990612 HcAAE1b MK990613 onekp:BNDE_scaffold_2026015c Hypericum perforatum Current study onekp:BNDE_scaffold_2009069c Pt4CL1 Populus tremuloides AAC24503.1 Pt4CL2 AAC24504.1 PpLuciferased Photinus pyralis AAA29795.1 AAE, Acyl-activating enzyme; BZL, benzoate-CoA ligase; BZO1, benzoyloxyglucosinolate 1; CCL, carboxyl CoA ligase; CNL, cinnamate-CoA ligase; 4CL, 4-coumarate-CoA ligase. a Function is not reported b Sequence is represented by its nucleotide accession number c Sequence obtained from Onekp database d Insect sequence

167

Appendix

C Alignment

NCBI_Luciferase MEDAKNIKKGPAPFYPLEDGTAGEQLHKAMKRYALVPGTIAFTDAHIEVNITYAEYFEMS 60 Cloned_Luciferase MEDAKNIKKGPAPFYPLEDGTAGEQLHKAMKRYALVPGTIAFTDAHIEVNITYAEYFEMS 60

NCBI_Luciferase VRLAEAMKRYGLNTNHRIVVCSENSLQFFMPVLGALFIGVAVAPANDIYNERELLNSMNI 120 Cloned_Luciferase VRLAEAMKRYGLNTNHRIVVCSENSLQFFMPVLGALFIGVAVAPANDIYNERELLNSMNI 120

NCBI_Luciferase SQPTVVFVSKKGLQKILNVQKKLPIIQKIIIMDSKTDYQGFQSMYTFVTSHLPPGFNEYD 180 Cloned_Luciferase SQPTVVFVSKKGLQKILNVQKKLPIIQKIIIMDSKTDYQGFQSMYTFVTSHLPPGFNEYD 180

NCBI_Luciferase FVPESFDRDKTIALIMNSSGSTGLPKGVALPHRTACVRFSHARDPIFGNQIIPDTAILSV 240 Cloned_Luciferase FVPESFDRDKTIALIMNSSGSTGLPKGVALPHRTACVRFSHARDPIFGNQIIPDTAILSV 240

NCBI_Luciferase VPFHHGFGMFTTLGYLICGFRVVLMYRFEEELFLRSLQDYKIQSALLVPTLFSFFAKSTL 300 Cloned_Luciferase VPFHHGFGMFTTLGYLICGFRVVLMYRFEEELFLRSLQDYKIQSALLVPTLFSFFAKSTL 300

NCBI_Luciferase IDKYDLSNLHEIASGGAPLSKEVGEAVAKRFHLPGIRQGYGLTETTSAILITPEGDDKPG 360 Cloned_Luciferase IDKYDLSNLHEIASGGAPLSKEVGEAVAKRFHLPGIRQGYGLTETTSAILITPEGDDKPG 360

NCBI_Luciferase AVGKVVPFFEAKVVDLDTGKTLGVNQRGELCVRGPMIMSGYVNNPEATNALIDKDGWLHS 420 Cloned_Luciferase AVGKVVPFFEAKVVDLDTGKTLGVNQRGELCVRGPMIMSGYVNNPEATNALIDKDGWLHS 420

NCBI_Luciferase GDIAYWDEDEHFFIVDRLKSLIKYKGYQVAPAELESILLQHPNIFDAGVAGLPDDDAGEL 480 Cloned_Luciferase GDIAYWDEDEHFFIVDRLKSLIKYKGYQVAPAELESILLQHPNIFDAGVAGLPDDDAGEL 480

NCBI_Luciferase PAAVVVLEHGKTMTEKEIVDYVASQVTTAKKLRGGVVFVDEVPKGLTGKLDARKIREILI 540 Cloned_Luciferase PAAVVVLEHGKTMTEKEIVDYVASQVTTAKKLRGGVVFVDEVPKGLTGKLDARKIREILI 540

NCBI_Luciferase KAKKGGKSKL 550 Cloned_Luciferase KAKKGGK-KL 549 Fig. C1: Alignment of the luciferase amino acid sequences obtained from the NCBI databank and encoded by the luciferase ORF of the pRS413-GAL1-luc*(-SKL) plasmid. Red rectangle marks the PTS1 which is absent from the in-house expressed luciferase.

168

Appendix

Poonam Singh 26-2, Mendelssohnstraße 38106, Braunschweig Born on 22.09.1989, Bijpur, U.P., India Nationality: Indian

Education/ Work experience Oct 2015 - ongoing Doctoral candidate at Institute für Pharmazeutische Biologie, TU Braunschweig, Germany Title: Biochemical and molecular investigations of benzoic acid metabolism in Hypericum calycinum cell cultures 2012 - 2014 Master of Technology (M. Tech.) from the Department of Agricultural and Food Engineering, Indian Institute of Technology Kharagpur, India Oct 2013 - March 2014 Masters project at Institute für Pharmazeutische Biologie, TU Braunschweig, Germany Title: Molecular cloning of 3-hydroxybenzoate: CoA ligase from Gentianaceae 2008 - 2012 Bachelor of Technology (B. Tech.) from Amity Institute of Biotechnology, Amity University, Uttar Pradesh, India Mar 2012 - May 2012 Project work done on the topic “Analysis of active pharmaceutical ingredient by Chromatography” at Teva API India Ltd., Gajraula, India 2007 Senior secondary (10+2), D.A.V. Public School, N.T.P.C. Rihandnagar, Uttar Pradesh, India 2005 High School (10th Standard), St. Josephs school, N.T.P.C Rihandnagar, Uttar Pradesh, India

Scholarships • DAAD PhD Scholarship for the session 2015 • DAAD IIT-TU9 Masters Sandwich Scholarship for the session 2013-2014 to conduct 6 months project work at Institute für Pharmazeutische Biologie, TU Braunschweig, Germany • 100% scholarship for the session 2008-2012 to study B.Tech. (Biotechnology) from Amity University, Uttar Pradesh, India

Awards • Best scientific poster prize by Phytochemical Society of Europe at the Green for Good IV- Biotechnology of Plant Products conference (19.06.2017 - 22.06.2017, Olomouc, Czech Republic) • Institute silver medal by Indian Institute of Technology Kharagpur for the academic year 2013-2014

Publications • Ali, R., Agarwal, P., Singh, P., Beuerle, T., Ernst, L., Mitra, A., Gaid, M., Liu, B. & Beerhues, L. Gentianaceae benzophenone synthases prefer 3-hydroxybenzoate CoA esters as starters. (Poster) Botanikertagung, Kiel, Germany (17.09.2017 - 21.09.2017).

169