Science, Faculty of
Chemistry, Department of
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UBCV
Daniel-Ivad, Phillip
Doctor of Philosophy - PhD
University of British Columbia
Studying individual biosynthetic transformations in the creation of natural products often reveals surprising and powerful chemical reactivities and molecular handling strategies. Indolocarbazole bisindoles have been widely tested in clinical studies and arise from oxidative dimerization of L-tryptophan. Among bisindoles, reductasporine bears an unusual dimethylpyrrolinium structure. Its biosynthesis differs from other indolocarbazole pathways by two tailoring enzymes: the imine reductase RedE and N,N-dimethyltransferase RedM. Here I reconstitute this pathway in vitro and show that RedE prevents unstable indolocarbazole intermediates from becoming oxidized and provides reduced didemethylreductasporine substrate to RedM. Employing X-ray crystallography, I solved two ternary complexes of RedE co-crystallized with the substrate-mimic arcyriaflavin A, revealing an extended active site cleft with distinct secondary indolocarbazole binding site. Site-directed mutagenesis confirms the conserved active site aspartate (D168) is essential for activity and anchors the substrate via hydrogen-bonding. Variants targeting the secondary binding site reduce catalytic efficiency, suggesting this site protects the substrate from autooxidation. I solved the 1.7 Å structure of RedM demonstrating it adopts distinct open and closed conformations with either SAH or SAM cofactor, respectively. Site-directed mutagenesis, docking and sequence bioinformatics identify conserved substrate-recognizing residues and suggest dimethyltransferase catalytic activity likely arises from precise orientation and desolvation of the substrate. Recently, Cap15 has been shown to be an oxygen- and pyridoxal phosphate (PLP)-dependent enzyme and is the first example of this activity in the L-seryl-tRNA(Sec) selenium transferase enzyme family. It catalyzes oxidative-decarboxylation of 5ʹ-glycyl uridine to the corresponding 5ʹ-carboxamide uridine. Solving for the 2.40 Å resolution crystal structure of Cap15 shows PLP bonds to K230 and a phosphate anion in the active site bridges N- and C-terminal domains. Sequence analysis reveals the loop proximal to the internal aldimine and hydrogen bonding to the active site phosphate is strictly conserved among Cap15 homologues present in capuramycin-type gene clusters. The crystal structures provide a basis for further investigations into the sequence-determinants of secondary binding site formation in RedE, dimethylation activity in RedM and oxygen-consuming activity in Cap15. RedE in particular can serve as a starting-point in the engineering of imine reductases to accommodate large substrates in the production of industry-relevant chiral amines.
CRYSTALLOGRAPHIC AND BIOCHEMICAL CHARACTERIZATION OF KEY STEPS IN REDUCTASPORINE AND CAPURAMYCIN BIOSYNTHESIS by Phillip Daniel-Ivad B.Sc., McMaster University, 2014 A DISSERTATION SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Chemistry) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) February 2024 © Phillip Daniel-Ivad, 2024 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Crystallographic and biochemical characterization of key steps in reductasporine and capuramycin biosynthesis submitted by Phillip Daniel-Ivad in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Chemistry Examining Committee: Katherine Ryan, Professor, Chemistry, UBC Supervisor Martin Tanner, Professor, Chemistry, UBC Supervisory Committee Member Harry Brumer, Professor, Chemistry, UBC Supervisory Committee Member David Perrin, Professor, Chemistry, UBC University Examiner Filip van Petegem, Professor, Biochemistry & Molecular Biology, UBC University Examiner Additional Supervisory Committee Members: Marco Ciufolini, Professor Emeritus, Chemistry, UBC Supervisory Committee Member iii Abstract Studying individual biosynthetic transformations in the creation of natural products often reveals surprising and powerful chemical reactivities and molecular handling strategies. Indolocarbazole bisindoles have been widely tested in clinical studies and arise from oxidative dimerization of L-tryptophan. Among bisindoles, reductasporine bears an unusual dimethylpyrrolinium structure. Its biosynthesis differs from other indolocarbazole pathways by two tailoring enzymes: the imine reductase RedE and N,N-dimethyltransferase RedM. Here I reconstitute this pathway in vitro and show that RedE prevents unstable indolocarbazole intermediates from becoming oxidized and provides reduced didemethylreductasporine substrate to RedM. Employing X-ray crystallography, I solved two ternary complexes of RedE co-crystallized with the substrate-mimic arcyriaflavin A, revealing an extended active site cleft with distinct secondary indolocarbazole binding site. Site-directed mutagenesis confirms the conserved active site aspartate (D168) is essential for activity and anchors the substrate via hydrogen-bonding. Variants targeting the secondary binding site reduce catalytic efficiency, suggesting this site protects the substrate from autooxidation. I solved the 1.7 Å structure of RedM demonstrating it adopts distinct open and closed conformations with either SAH or SAM cofactor, respectively. Site-directed mutagenesis, docking and sequence bioinformatics identify conserved substrate-recognizing residues and suggest dimethyltransferase catalytic activity likely arises from precise orientation and desolvation of the substrate. Recently, Cap15 has been shown to be an oxygen- and pyridoxal phosphate (PLP)-dependent enzyme and is the first example of this activity in the L-seryl-tRNA(Sec) selenium transferase enzyme family. It catalyzes oxidative-decarboxylation of 5ʹ-glycyl uridine to the corresponding 5ʹ-carboxamide uridine. Solving for the 2.40 Å resolution crystal structure of Cap15 shows PLP bonds to K230 and a iv phosphate anion in the active site bridges N- and C-terminal domains. Sequence analysis reveals the loop proximal to the internal aldimine and hydrogen bonding to the active site phosphate is strictly conserved among Cap15 homologues present in capuramycin-type gene clusters. The crystal structures provide a basis for further investigations into the sequence-determinants of secondary binding site formation in RedE, dimethylation activity in RedM and oxygen-consuming activity in Cap15. RedE in particular can serve as a starting-point in the engineering of imine reductases to accommodate large substrates in the production of industry-relevant chiral amines. v Lay Summary Biosynthesis is the protein-directed process of assembling new and complex molecules from simpler starting materials in living organisms. These proteins, called enzymes, effectively form microscopic assembly line pathways where each chemically modifies the molecule passed to it before releasing it for the next enzyme in the line. In this work I study the chemistry of and determine the three-dimensional structures of enzymes in the reductasporine and capuramycin biosynthetic pathways, both compounds with antibiotic properties. Knowing the chemistries and structures of these enzymes is essential in our understanding of how enzymes modify small molecules. It allows us to test designs and make comparisons between enzymes. Ultimately, it can allow us to engineer sets of enzymes to have powerful chemistries into living microbes that efficiently and cheaply produce pharmaceuticals, effectively creating custom biosynthetic pathways. vi Preface I performed all experiments and analyzed all data collected here under the mentorship of Dr. K. S. Ryan. Chapters 1 and 5 were written by me with Dr. K. S. Ryan providing thoughtful editing. A version of Chapters 2 has been published in the Journal of Biological Chemistry and a version of Chapter 3 has been accepted for publication at the Journal of Biological Chemistry: Daniel-Ivad, P., and Ryan, K. S. (2024) Structure of methyltransferase RedM that forms the dimethylpyrrolinium of the bisindole reductasporine. J. Biol. Chem. 300, 105520. Daniel-Ivad, P., and Ryan, K. S. (2024) An imine reductase that captures reactive intermediates in the biosynthesis of the indolocarbazole reductasporine. J. Biol. Chem. in press. Dr. K. S. Ryan and I are the authors for both manuscripts. I wrote the manuscripts and revised the texts per edits by Dr. K. S. Ryan. Dr. C. L. Drennan at the Massachusetts Institute of Technology provided pET28a-staP plasmid. A version of Chapter 4 has been published in collaboration with the Dr. S. Van Lanen lab at the University of Kentucky: Daniel-Ivad, P. G., Van Lanen, S., and Ryan, K. S. (2023) Structure of the Oxygen, Pyridoxal Phosphate-Dependent Capuramycin Biosynthetic Protein Cap15. Biochemistry 62, 2611–2621. I wrote the manuscript and revised the text per edits by Dr. K. S. Ryan and Dr. S. Van Lanen. Dr. S. Van Lanen provided pET28a-cap15 plasmids alongside genuine synthetic samples of CarU and synthetic precursor. vii Table of Contents Abstract ........................................................................................................................................ iii Lay Summary ................................................................................................................................. v Preface ........................................................................................................................................... vi Table of Contents ......................................................................................................................... vii List of Tables ................................................................................................................................ xii List of Figures ............................................................................................................................ xiii List of Abbreviations ................................................................................................................. xvii Acknowledgements .................................................................................................................. xviii Dedication .................................................................................................................................... xix Chapter 1: Oxygen-dependent chemical diversity in bisindole, amino acid and nucleoside natural products ............................................................................................................................ 1 1.1 Natural products, pharmaceuticals and biosynthesis ....................................................... 1 1.2 Bisindole alkaloid natural products ................................................................................. 5 1.3 The conserved bacterial bisindole biosynthetic pathway ................................................ 9 1.3.1 Violacein ................................................................................................................ 12 1.3.2 Chromopyrrolic acid derivatives and bisindolylmaleimides ................................. 14 1.3.3 Indolocarbazoles .................................................................................................... 15 1.3.4 Indolotryptolines .................................................................................................... 16 1.4 Other amino acid oxidative entries into biosynthetic pathways .................................... 18 1.4.1 L-Enduracididine ................................................................................................... 21 1.4.2 Azomycin ............................................................................................................... 21 1.4.3 Indolmycin ............................................................................................................. 23 viii 1.4.4 Capuramycin .......................................................................................................... 24 1.5 Objective of Dissertation ............................................................................................... 26 Chapter 2: Structure of methyltransferase RedM that forms the dimethylpyrrolinium of the bisindole reductasporine ....................................................................................................... 28 2.1 Synopsis ......................................................................................................................... 28 2.2 Introduction ................................................................................................................... 28 2.3 Results ........................................................................................................................... 30 2.3.1 In vitro reconstitution of reductasporine biosynthesis ........................................... 30 2.3.2 Crystal structure of RedM ..................................................................................... 34 2.3.3 Modelling the RedM-substrate complex ............................................................... 39 2.3.4 Mutagenesis of the RedM active site ..................................................................... 42 2.3.5 Sequence Similarity Network of COMT-type Methyltransferases ....................... 47 2.4 Discussion ...................................................................................................................... 50 2.5 Experimental Procedures ............................................................................................... 60 2.5.1 General methods .................................................................................................... 60 2.5.2 Cloning and expression of redE and redM ............................................................ 60 2.5.3 Cloning, expression of rebO and rebD, and production of chromopyrrolic acid .. 61 2.5.4 Expression and purification of StaP, ferredoxin and ferredoxin reductase ........... 62 2.5.5 In vitro production of reductasporine and pathway intermediates ........................ 62 2.5.6 Synthesis of arcyriarubin A ................................................................................... 63 2.5.7 Synthesis of arcyriaflavin A .................................................................................. 63 2.5.8 Synthesis of didemethylreductasporine ................................................................. 64 2.5.9 Enzymatic methylation of didemethylreductasporine ........................................... 64 ix 2.5.10 Selenomethionine incorporation into RedM .......................................................... 64 2.5.11 Crystallization of RedM ........................................................................................ 65 2.5.12 Data collection, structure determination and refinement ....................................... 66 2.5.13 Substrate Docking .................................................................................................. 67 2.5.14 Site directed mutagenesis of RedM ....................................................................... 67 2.5.15 Visualization and analysis software ...................................................................... 68 2.5.16 Sequence similarity networks ................................................................................ 68 Chapter 3: An imine reductase that captures reactive intermediates in the biosynthesis of the indolocarbazole reductasporine ........................................................................................... 69 3.1 Synopsis ......................................................................................................................... 69 3.2 Introduction ................................................................................................................... 69 3.3 Results ........................................................................................................................... 73 3.3.1 RedE dataset phasing, twinning and initial model building .................................. 73 3.3.2 Structure of RedE .................................................................................................. 76 3.3.3 Resemblance to other NAD(P)H-dependent reductases ........................................ 82 3.3.4 Domain contacts among IRED and RedAm enzymes. .......................................... 84 3.3.5 Single residue mutations in RedE .......................................................................... 88 3.3.6 Inspection of StaP products ................................................................................... 91 3.4 Discussion ...................................................................................................................... 92 3.5 Experimental procedures ............................................................................................... 98 3.5.1 Cloning, expression and purification of redE and redM ....................................... 98 3.5.2 Selenomethionine incorporation into RedE ........................................................... 99 3.5.3 RedE crystallization ............................................................................................... 99 x 3.5.4 Data collection, structure determination, and refinement .................................... 100 3.5.5 Site-directed mutagenesis of RedE ...................................................................... 101 3.5.6 Cloning and expression of rebO and rebD, and production of CPA ................... 101 3.5.7 Expression and purification of StaP, ferredoxin and ferredoxin reductase ......... 101 3.5.8 Analysis of RedE variants ................................................................................... 102 3.5.9 Time-course analysis of StaP reaction products .................................................. 102 3.5.10 Visualization and analysis software .................................................................... 103 Chapter 4: The structure of the oxygen, pyridoxal phosphate-dependent capuramycin biosynthetic protein Cap15 ....................................................................................................... 104 4.1 Synopsis ....................................................................................................................... 104 4.2 Introduction ................................................................................................................. 104 4.3 Results ......................................................................................................................... 107 4.3.1 Overall structure of Cap15 .................................................................................. 108 4.3.2 Active site ............................................................................................................ 113 4.3.3 External aldimine docking ................................................................................... 115 4.3.4 Conservation of active site residues .................................................................... 120 4.4 Discussion .................................................................................................................... 126 4.5 Experimental procedures ............................................................................................. 131 4.5.1 Purification of Cap15 ........................................................................................... 131 4.5.2 Size exclusion chromatography ........................................................................... 132 4.5.3 Crystallization ...................................................................................................... 132 4.5.4 Data collection, structure determination and refinement ..................................... 133 4.5.5 Molecular model preparation and docking .......................................................... 133 xi 4.5.6 Sequence analysis ................................................................................................ 134 Chapter 5: Conclusions ............................................................................................................. 135 5.1 Detours in bacterial bisindole biosynthesis ................................................................. 135 5.2 Structural characterization of Cap15, a key enzyme in capuramycin biosynthesis ..... 141 References ................................................................................................................................... 144 xii List of Tables Table 1.1 Bisindole natural products with associated biosynthetic gene clusters found in the MIBiG database. .............................................................................................................................. 6 Table 2.1 Data collection and refinement statistics. ...................................................................... 35 Table 2.2 COMT-type methyltransferase family enzymes with distinct open and closed conformations. ............................................................................................................................... 52 Table 2.3 COMT-type family enzymes capable of furnishing at least two methyltransfers to a substrate. ........................................................................................................................................ 54 Table 3.1 Crystallographic data collection and refinement statistics ............................................ 75 Table 3.2 Sixteen IRED and RedAm hits from secondary structure matching of RedE using PDBeFOLD. .................................................................................................................................. 85 Table 4.1 Data collection and refinement statistics. .................................................................... 107 Table 4.2 Secondary structural alignments of Cap15 to representative aspartate aminotransferase family enzymes and arginine oxidases. ....................................................................................... 110 Table 4.3 Autodock Vina benchmarking of conformational searches of medium to large sized external aldimines found in the PDB. .......................................................................................... 118 Table 4.4 Multiple sequence alignment of the top N blastp hits querying the Cap15 sequence against the clustered nr NCBI database ....................................................................................... 121 Table 4.5 DGL/Cap15-like cluster active site sequence motifs. ................................................. 125 xiii List of Figures Figure 1.1 Examples of molecules from each natural product class. .............................................. 2 Figure 1.2 The staurosporine pathway CPA synthase, StaD, is proposed to couple two 3-indolepyruvic acid enamine tautomer molecules via a radical mechanism. .................................. 10 Figure 1.3 Shared biosynthetic pathway of bacterial bisindoles. .................................................. 11 Figure 1.4 Proposed biosynthetic pathway to violacein after divergence from shared bacterial bisindole biosynthesis. ................................................................................................................... 13 Figure 1.5 Structures of synthetic bisindolylmaleimides entered into clinical trials. .................... 14 Figure 1.6 Proposed biosynthetic steps towards indolotryptoline cladoniamide C. ...................... 17 Figure 1.7 Biosynthetic pathways to enduracididine, azomycin and indolmycin initiated by arginine oxidases MppP, RohP and Ind4, respectively. ................................................................ 20 Figure 1.8 Selection of liposidomycin-type and capuramycin-type NPs with their defining GlyU and CarU groups highlighted in red and blue, respectively. ......................................................... 25 Figure 1.9 Uridine 5ʹ-monophosphate (UMP) is transformed into CarU in at least four steps. .... 26 Figure 2.1 Indolocarbazoles, their gene clusters and shared biosynthetic enzymes. ..................... 30 Figure 2.2 In vitro reconstitution of reductasporine biosynthesis. ................................................ 31 Figure 2.3 Representative SDS-PAGE gels of recombinant RedE and RedM Ni-IMAC and size-exclusion chromatography fractions. ............................................................................................. 32 Figure 2.4 1H-NMR (600 MHz, DMSO-d6) of reductasporine isolated from a large-scale in vitro reconstitution of biosynthetic pathway from CPA. ....................................................................... 33 Figure 2.5 Structure of RedM. ....................................................................................................... 34 Figure 2.6 Size-exclusion chromatography traces for RedM (red) and molecular weight standards. ....................................................................................................................................... 37 xiv Figure 2.7 RedM active site residue contacts. ............................................................................... 38 Figure 2.8 Docking poses of didemethylreductasporine (3) into the open and closed conformations of RedM. ................................................................................................................ 41 Figure 2.9 Sequence alignment of RedM against representative structural homologues available in the PDB grouped by presence or lack of the catalytic histidine found as serine at position 253 in RedM. ........................................................................................................................................ 44 Figure 2.10 In vitro assays of RedM variants and active site comparisons to COMT-type homologues. ................................................................................................................................... 46 Figure 2.11 Sequence similarity network of the COMT-like enzyme family (IPR016461) generated using the Uniref50 database. ......................................................................................... 49 Figure 2.12 View of the crystal packing of apo-RedM and the RedM-SAM binary complex. .... 51 Figure 3.1 Shared steps and examples of indolocarbazole biosynthesis. ...................................... 71 Figure 3.2 Proposed biosynthetic pathway to reductasporine (1). ................................................ 72 Figure 3.3 Asymmetric unit of the selenomethionine-substituted RedE structure. ....................... 74 Figure 3.4 Cartoon representations of the overall RedE structure. ............................................... 77 Figure 3.5 Ligand mFo-DFc omit maps contoured at 3𝜎. .............................................................. 79 Figure 3.6 Key ligand interactions in the in the extended RedE active site. ................................. 80 Figure 3.7 Ninety degree separated views of representative dehydrogenases structurally related to RedE. ............................................................................................................................................. 82 Figure 3.8 Multiple sequence alignments of RedE homologues. .................................................. 86 Figure 3.9 Multiple sequence alignment of residues in the contours of the secondary binding cleft in RedE. ......................................................................................................................................... 87 Figure 3.10 LC-MS traces of RedE variant reaction products. ..................................................... 90 xv Figure 3.11 Extracted ion chromatograms for m/z = 296, the mass consistent with indolocarbazole 3, of StaP reaction products over 2.5 h. .............................................................. 92 Figure 3.12 Residue H249 in the secondary binding site potentially selects for the imine tautomer of 3 that lines up for reduction upon direct transfer to the primary active site ............................. 97 Figure 4.1 Enzymatic transformations catalyzed by Cap15, DGL and SelA. ............................. 105 Figure 4.2 Overview of the structure of Cap15. .......................................................................... 109 Figure 4.3 Topology diagram, interdimer interface, and secondary structures with residue ranges. ..................................................................................................................................................... 110 Figure 4.4 Side by side comparisons of representative structures of PLP-dependent enzyme families. ....................................................................................................................................... 111 Figure 4.5 Size-exclusion chromatography traces for Cap15 octamer and molecular weight standards. ..................................................................................................................................... 113 Figure 4.6 View of the Cap15 active site residues and UV-Vis spectrum of Cap15 solution and crystal. .......................................................................................................................................... 115 Figure 4.7 Representative poses of external aldimines for the docking of water and phosphate anion using the flexible side chain method in Autodock Vina. ................................................... 117 Figure 4.8 Visualization of Autodock Vina flexible side chain method conformational searches benchmarking. ............................................................................................................................. 119 Figure 4.9 Alignment of genes in the vicinity of Cap15 homologues as clustered by the SSN of the SelA-like family (AST = 110). .............................................................................................. 122 Figure 4.10 Sequence similarity network of the L-seryl-tRNA selenium transferase-like family (IPR018319). ............................................................................................................................... 124 Figure 4.11 Proposed Cap15 mechanism after structural considerations. ................................... 130 xvi Figure 5.1 A proposed biosynthetic pathway to erdasporines. .................................................... 137 Figure 5.2 Select spiroindimicins and indimicins isolated from marine actinomycetes. ............ 138 Figure 5.3 Proposed biosynthetic transformation required to yield spiroindimicins. ................. 140 Figure 5.4 Select tjipanazoles isolated from cyanobacteria Tolypothrix tjipanasensis. .............. 141 xvii List of Abbreviations AAS aromatic aldehyde synthase AAT aspartate aminotransferase AFA arcyriaflavin A AO arginine oxidase Arg L-arginine AST alignment score threshold BME β-mercaptoethanol CarU uridine-5ʹ-carboxamide COMT caffeic acid O-methyltransferase CPA chromopyrrolic acid CPAS chromopyrrolic acid synthases eDNA environmental DNA FDR ferredoxin-NADP oxidoreductase FDX ferredoxin I GlyU uridine-5ʹ-glycine ICZ indolocarbazole IPTG β-D-1-thiogalactopyranoside IRED imine reductase NP natural product PLP pyridoxal-5ʹ-phosphate PVA protodeoxyviolaceinic acid RedAm reductive aminase SSN sequence similarity network TCEP tris(2-carboxyethyl)phosphine Trp L-tryptophan U5ʹA uridine-5ʹ-aldehyde UMP uridine monophosphate xviii Acknowledgements I thank Dr. Melanie Higgins for insight into X-ray crystallographic experiments and modelling, Dr. Hai-Yan He for helpful discussions on enzymatic assays, Dr. Jason Hedges for their keen eye on literature, as well as my fellow lab mates who kindly look out for my best interests. I thank Dr. Katherine Ryan for immense patience in light of my diving into scientific rabbit holes and her guidance in returning me from them. Special thanks are owed to my family, who have supported me throughout my years of education Funding for this work came from a Four-Year Doctoral Fellowship from the University of British Columbia, the Michael Smith Foundation for Health Research, and the Natural Sciences and Engineering Research Council of Canada. Thanks to the Canadian Light Source and Stanford Synchrotron Radiation Lightsource for access to beamtime. xix Dedication To prattling conversations 1 Chapter 1: Oxygen-dependent chemical diversity in bisindole, amino acid and nucleoside natural products 1.1 Natural products, pharmaceuticals and biosynthesis Natural products (NPs) are molecules naturally produced in an organism and include sugars, lipids, amino acids, nucleic acids and their derivatives, though working definitions of NPs focus on the distinct and often complex molecules produced via organism-specific secondary metabolism. Where primary metabolism covers all essential processes and molecules for the survival and proliferation of an organism, secondary metabolism covers all non-essential processes and molecules that nonetheless confer conditional fitness benefits that are critical to the success of an organism in its native environment. For example, microbes compete amongst each other via secreted NPs involved in cell-to-cell communication (quorum sensing), chemical warfare (antibiotics), and nutrient uptake (siderophores).1 An estimated 50-70 % of approved pharmaceutical drugs are NP molecules, are similar to NPs or are NP derivatives.2 This wide use of NPs and related compounds originates from the success of the industrial production and application of early NP antibiotics, such as penicillin and streptomycin in the 1940s and 1950s, and following NP screening campaigns initiated at pharmaceutical companies.3 As more of these molecules were characterized it became clear that they possessed incredible structural and chemical diversity and complexity difficult to access with organic synthetic approaches.4,5 2 Figure 1.1 Examples of molecules from each natural product class. Polyketides, erythromycin; alkaloids, lysergic acid; phenylpropanoids, cyanidin; peptides, enterobactin; nucleosides, sinefungin; glycosides, kanamycin; and isoprenoids, retinal. Each molecule is labelled with its general function or activity. NPs can be organized into seven broad classes based on their primary metabolite precursors and common themes in enzymatic assembly: peptides, polyketides, isoprenoid/terpenoid, phenylpropanoids, alkaloids, nucleosides and glycosides (Figure 1.1).6 Study of the biosynthetic steps from similar precursors out into the immense carbon skeleton 3 diversity observed in each class has led to the identification of ‘gatekeeper enzymes’ which act on primary metabolites to produce products irreversibly committed to secondary metabolism. Often, these are the simple coupling of a metabolite to a growing biosynthetic intermediate, such as the incorporation of malonyl-CoA via polyketide synthases or amino acids via non-ribosomal peptide synthetases. Alternatively, primary metabolites are directly modified or functionalized, such as the prenylation of tryptophan in the biosynthesis of alkaloid lysergic acid, the condensation of dopamine and 4-hydroxyphenylacetaldehyde in the biosynthesis of alkaloid (S)-reticuline, or the deamination of phenylalanine to cinnamate in the biosynthesis phenylpropanoids. Numerous tailoring enzymes then follow, which typically increase skeletal complexity, tune molecular properties and otherwise create variations.7 Investigations into these gatekeeper and tailoring enzymes have revealed sophisticated and powerful chemical transformations of carbon-carbon, carbon-hydrogen and carbon-heteroatom bonds including radical SAM enzymes, heme-dependent cytochrome P450s, flavin-dependent oxidoreductases, epoxidases, halogenases, Pictet-Spenglerases, Diels-Alderases (plus other electrocyclization-ases), and aldolases. Such enzymes can be the starting point for development of biocatalysts, useful to the pharmaceutical and other industries.8 More broadly, the field of NP biosynthesis is the study of how organisms themselves construct diverse NPs. Utilizing biosynthetic pathways via fermentation of NP-producing organisms offers an alternative to purely synthetic routes from industrial starting materials.9 Understanding the step-by-step biosynthetic pathways towards complex NPs also informs on ways to mimic nature’s approach and create more efficient syntheses.10 Synthetic and biological approaches may also be combined into semi-synthetic strategies where one isolates an abundant 4 pharmaceutical precursor from an organism before synthetically derivatizing it into its drug form.11 The ever-lowering price of DNA sequencing in the past several decades meant producing nucleotide data became widely accessible, and public databases of these sequences made the data widely available. Researchers spent this time developing strategies to gather genomic material from lab cultured strains, from the microbiomes of plants and animals, as well as general environmental samples like marine water and desert soils. Spreading the search for DNA to all imaginable areas of the Earth has been motivated by the success of metagenomics in revealing the depth of microbial diversity and symbiosis and in providing access to gene products otherwise not possible via traditional methods.12 Additions to characterized NP biosynthesis assisted via genomics has allowed for functional annotations of gene databases and enables either manual or automated predictions on whether an organism may produce a certain NP or related, novel NPs.13 While isolating DNA from environmental samples provides less complete genomes, it is less affected by the constraints of culturing bacteria in a lab setting making microbial biosynthetic “dark matter” accessible for investigation. For example, the insight that variation in the primary sequence of key biosynthetic enzymes in a pathway correlate with the carbon scaffold diversity of the consequent NP coded for in the gene cluster allows for directed searches of otherwise intractable DNA libraries. In this context searching for highly dissimilar, but still related, enzymes enable the discovery of NPs with novel structural features with hitherto uncharacterized biosynthetic chemistry. In this thesis, my work is focused on the characterization of key distinguishing tailoring enzymes in the biosynthesis of the bacterial bisindole NP reductasporine, and an unusual oxygen- and pyridoxal phosphate-dependent enzyme, Cap15, that catalyzes the formation of the 5 core carbon skeleton of capuramycin-type nucleoside NPs. Each of these topics are introduced below. 1.2 Bisindole alkaloid natural products L-Tryptophan (Trp) is the most complex and least abundant of the proteinogenic amino acids, with its characteristic indole group serving as a jumping off point for chemical complexity and diversity for many alkaloid NPs.14 This nucleophilic aromatic ring system can be modified enzymatically at almost every atom including nitration, chlorination, oxidation, and alkylation. Trp is known to undergo spontaneous, non-enzymatic chemistries, as well. Bisindole NPs are characterized by the presence of two indole moieties in NP molecules, usually excluding those with the trivial inclusion of two tryptophan units in peptide NPs. Hundreds of these bisindole molecules have been isolated from natural sources, many with desirable effects in pharmaceutical applications.15 Perhaps the most widely recognized bisindole compound is the natural dye indigo used to give denim jeans their distinctive colour, and this NP has been used as a colourant for hundreds of years.16 Natural indigo is produced from the spontaneous oxidative dimerization of 3-hydroxyindole hydrolyzed from the corresponding indole glycoside, indican.17 By 2009, the biosynthetic construction of only six bisindole molecules were known and a search of the MIBiG database records (as of this publication) identifies biosynthetic gene clusters for over twenty bisindole NPs fitting at least three groups: indolocarbazoles, indolotryptolines and diketopiperazine-related. (Table 1.1).15,18 The diketopiperazine-related bisindole compounds are typically of fungal origin, whereas indolocarbazole and indolotryptoline NPs are typically made by bacteria from genera like Chromobacterium, Streptomyces, Lechevalieria, Salinispora, and Nonomuraea.19 Exploration of 6 these pathways has revealed novel enzymatic mechanisms and motivated manipulating these enzymes in combinatorial syntheses of new bisindole molecules.15 Table 1.1 Bisindole natural products with associated biosynthetic gene clusters found in the MIBiG database. Natural Product MIBiG Accession Source organism Group Representative structure Indolocarbazoles AT2433-A1 BGC0000809 Actinomadura melliaura Bacteria K252a BGC0000814 Nonomuraea longicatena Bacteria Rebeccamycin BGC0000821 Lechevalieria aerocolonigenes Bacteria Staurosporine BGC0000825 Streptomyces sp. TP-A0274, Streptomyces clavuligerus, Salinispora arenicola CNS-205 Bacteria Hydroxysporine BGC0001223 uncultured bacterium Bacteria Reductasporine BGC0001224 uncultured bacterium Bacteria Erdasporine BGC0001336 uncultured bacterium Bacteria NHOOHOONClNHONOHOHOOON NHNOOOMeHOMeOOOHOHNClClNHOHN OHOHNOOMeNHNONHNHHNONHOHHONHN+NHOOHNNHNHHO7 Indimicin BGC0001782 Streptomyces sp. SCSIO 03032 Bacteria Loonamycin BGC0002460 Nocardiopsis flavescens Bacteria Tjipanazole BGC0002482 Symphyonema bifilamentata 97.28 Cyanobacteria Indolotryptoline BE-54017 BGC0001333 uncultured bacterium AB1650 Bacteria Cladoniamide BGC0001334 Streptomyces uncialis Bacteria Borregomycin BGC0001335 uncultured bacterium Bacteria Lazaramide BGC0001337 uncultured bacterium Bacteria Diketopiperazine-related Terrequinone A BGC0000442 Aspergillus nidulans FGSC A4 Fungi Melinacidin IV BGC0001585 Escovopsis weberi Fungi HNClNNHClMeOHOONClONOOHON OHOClNHOHOHClNHNHClONOHONOHONClONOHONOHONHClOOMeNMeONOCl NH HOClOOOHNHONOClNH OOHClNHOHOHNOH HNOHSSHONOONOHSOHNOONHNHS8 Ochrindole A BGC0001712 Aspergillus steynii IBT 23096 Fungi Okaramine BGC0001717 BGC0001718 Penicillium simplicissimum Aspergillus aculeatus Fungi Naseseazine C BGC0001948 Streptomyces sp. Bacteria (-)-ditryptophenaline BGC0002157 Aspergillus flavus Fungi Waikikiamide BGC0002254 Aspergillus sp. FM242 Fungi Other Methylarcyriarubin BGC0000817 uncultured bacterium Bacteria Violacein BGC0000829 Chromobacterium violaceum, Duganella sp. B2 Bacteria Marinacarboline D BGC0001137 Marinactinospora thermotolerans Bacteria Aetokthonotoxin BGC0002643 Aetokthonos hydrillicola Thurmond2011 Cyanobacteria Bisindoles are well-known to have bioactivity, and several have been studied as potential clinical drugs.20 For example, the fungal bisindole terrequinone A has anti-cancer and anti-viral properties.21 Staurosporine is widely used as a protein kinase C inhibitor, whereas rebeccamycin NHHNMeOHO OMeMeOHONONHNONHOONNHHNONONH ONHNNONONHONONNONHN OONHONOO O HONOMeOOONNHONHO NHNHNOHOHOHN NOHNNHBrNNHBrBrN Br Br9 is a topoisomerase inhibitor.22,23 Indolotryptolines inhibit the proteolipid subunit of vacuolar ATPase.24,25 In addition to these bioactivities, synthetic indolocarbazole isomers are promising for their use in OLEDs and transistor devices due to their low-lying HOMOs and consequent large band gap when included in polymer materials.20 1.3 The conserved bacterial bisindole biosynthetic pathway Despite the diversity in bacterial bisindole structures most have share the two initial biosynthetic steps. These are (1) the 𝛼-amine oxidation of Trp (or Trp derivative) to the imine by a flavin-dependent oxidase, and (2) oxidative C𝛽-C𝛽 dimerization between 𝛼-enamine-Trp derivatives by a heme-containing oxidase producing the 3-indolepyruvic acid imine dimer (Figure 1.2).19 After this reaction each intermediate is the basis for the final carbon skeleton for a distinct group of bacterial bisindole NP (Figure 1.3). Violacein is derived from the rearrangement of the 3-indolepyruvic acid imine dimer structure, intercepted before its spontaneous cyclization to chromopyrrolic acid (CPA).26 Lynamicins and bisindolylmaleimides are formed directly from CPA or the corresponding chlorinated derivatives.27,28 Indolocarbazoles arise from the oxidative bridging of each indole C2 carbon via the heme-dependent cytochrome P450 indolocarbazole synthases.29,30 This cyclization is proposed to produce 5,7-dicarboxy-indolocarbazole, which has not been directly observed in experiments, instead several decarboxylated and oxidized decomposition products are seen.19 However, the presence of downstream enzymes finely controls the ultimate outcome yielding many combinations of oxidation and decarboxylation as seen in erdasporines, indimicins, reductasporine, staurosporine and rebeccamycin.19,29 Lastly, indolotryptolines derive from maleimide indolocarbazoles (such as arcyriaflavin A) and feature a rearranged indolocarbazole scaffold with an apparent indole 10 ‘ring-flip’. This is accomplished in two oxidative steps with an intermediate maleimide N-methylation step which stabilizes an oxidized intermediate.31 Figure 1.2 The staurosporine pathway CPA synthase, StaD, is proposed to couple two 3-indolepyruvic acid enamine tautomer molecules via a radical mechanism. Because the core enzymes involved in constructing these types of bisindoles are conserved across different biosynthetic pathways, it is possible to identify new bisindole biosynthetic enzymes by searching for conserved features in the corresponding genes. In 2011, Chang and Brady screened a library of environmental DNA (eDNA) for the key C𝛽-C𝛽 𝛼-imine-Trp dimerizing enzyme (CPA synthase), which lead to the discovery of the BE-54017 indolotryptoline gene cluster32 and successively the gene clusters for indolocarbazoles erdasporine, hydroxysporine, and reductasporine; the indolotryptoline borregomycin; and bisindolylmaleimide methylarcyriarubin.14,19 11 Figure 1.3 Shared biosynthetic pathway of bacterial bisindoles. The common set of transformations performed by the flavoenzyme amino acid oxidases, CPA synthases, and the P450 enzyme indolocarbazole synthases provide a rich collection of active intermediates branching into a large variety of bacterial bisindoles. Shared intermediates are labelled according for R groups corresponding to hydrogen rather than alternate patterns of chlorine or hydroxylation. 12 1.3.1 Violacein Violacein is a purple pigment first isolated from Chromobacterium violaceum in 1882, and later isolated from Janthinobacterium lividum.33,34 It is proposed to protect these bacteria from damaging UV light, but also has clinical activities including antibiotic, antiviral and cytotoxicity against several cancer cell lines. In the biosynthetic pathway to violacein, VioA and VioB catalyze 3-indolepyruvate imine formation and C𝛽-C𝛽 bond formation, respectively.35 The 3-indolepyruvate imine dimer is labile and has not been isolated experimentally; however, the VioB homologue, StaD, is capable of joining 3-indolepyruvate into the more stable, non-imine 3-indolepyruvate dimer.36 In the next biosynthetic step, VioE captures the unstable 3-indolepyruvate imine dimer and catalyzes decarboxylative C𝛽 to C𝛼 indole migration (Figure 1.4).26,35 Subsequent, spontaneous cyclization produces protodeoxyviolaceinic acid (PVA). VioE accomplishes this 1,2-indole shift without the use of cofactors, while other aryl shifts, such as the phenyl shift from flavanone to isoflavone in plants, requires Fe(II) as a cofactor.37 PVA is a short-lived compound and is converted to violacein via FAD-dependent monooxygenases VioD and VioC.35 PVA or products produced by VioD and VioC may autooxidize to prodeoxyviolacein, proviolacein or deoxyviolacein. 13 Figure 1.4 Proposed biosynthetic pathway to violacein after divergence from shared bacterial bisindole biosynthesis. VioE is the key step distinguishing violacein from its related bisindole NPs. X-ray crystallographic structural elucidation of VioE in complex with 3-phenylpyruvate allowed for the proposal of a mechanism whereby the enzyme chaperones the unstable IPA imine dimer 14 intermediate into a more stable conformation in its active site, preventing cyclization to CPA, and highlighted residues essential to catalysis.26 There is a competing proposal that the substrate for VioE is the cyclized 3-indolepyruvate imine dimer before ammonia elimination, and cyclopropane ring formation is driven by the release of ammonia.35 1.3.2 Chromopyrrolic acid derivatives and bisindolylmaleimides CPA derives from the spontaneous cyclization of the C𝛽-C𝛽 dimerized 3-indolepyruvate imine produced by CPA synthases and is the biosynthetic precursor to lynamicins and bisindolylmaleimides (Figure 1.3).19,27,28 Lynamicins are variably halogenated, decarboxylated and methylester CPA derivatives with Gram-positive and Gram-negative antimicrobial activity.27 Lynamicin B is a chitinase inhibitor with potential use as an insecticide selective for crop pests, but not the natural pest predators.38 While a potential gene cluster for lynamicins have been identified, the conserved tryptophan oxidase is non-essential for lynamicin biosynthesis, implying complementation by an amino acid oxidase encoded elsewhere in the genome.39,40 In addition to the lynamicins, CPA is also presumed to be related to the biosynthesis of C-methylated indolocarbazole derivatives indimicins and the related spiroindimicin with substantially rearranged carbon skeletons.39,40 Figure 1.5 Structures of synthetic bisindolylmaleimides entered into clinical trials. 15 Several bisindolylmaleimides have entered clinical trials for their efficacy against various cancers and treatment of various outcomes of diabetes, including bisindolylmaleimide-I, rubroxistaurin, tivantinib and enzastaurin (Figure 1.5).20 These inhibit several isoforms of protein kinase C, and enzastaurin has been extensively studied in over 50 clinical trials as it selectively inhibits protein kinase C𝛽 and the PI3K/AKT pathway causing apoptosis in several cancer cell lines and inhibits angiogenesis mediated by vascular endothelial growth factor. Two phase III trials assessing enzastaurin efficacy in the treatment of glioblastoma and vascular Ehlers-Danlos syndrome are ongoing.41,42 Bisindolylmaleimides were originally isolated in the early 2000s from the fruiting bodies of slime molds,20 such as Dictydiaethalium plumbeum and Arcyria denudate, though the first associated biosynthetic gene cluster was found from eDNA in 2014 for methylarcyriarubin (mar).28 The gene cluster contains a CPA synthase (MarB), Rieske-type dioxygenase (MarC) and SAM-dependent methyltransferase MarM. 3-indolepyruvic acid imine is presumed to be generated by an enzyme outside the gene cluster before dimerization via MarB to CPA before oxidative-decarboxylation to arcyriarubin A by MarC. Maleimide N-methylation follows, catalyzed by MarM (Figure 1.3). It has yet to be shown whether the logic of bacterial bisindolylmaleimide biosynthesis applies to the eukaryotic slime molds. 1.3.3 Indolocarbazoles The indolocarbazoles are derived from CPA or its corresponding halogenated derivatives. As in the pathways discussed above, CPA is formed from the sequential action of the flavoprotein L-tryptophan oxidase (e.g. StaO, RebO, EspO) and a heme-dependent enzyme (e.g. StaD, RebD, EspD) dimerizes the resultant 3-indolpyruvate imine.19,43 This dimer then spontaneously cyclizes, eliminating ammonia, forming CPA. Next, a heme-dependent cytochrome P450 (e.g. StaP, RebP, EspP) couples the C2–C2ʹ indole-indole bond of CPA 16 producing the proposed dicarboxyindolocarbazole (Figure 1.3). This cytochrome P450 product is highly labile and has not been directly detected. It quickly decarboxylates and is susceptible to oxidation, spontaneously decomposing into primarily three products over time: K252c, 7-OH-K252c and arcyriaflavin A (or their corresponding halogenated derivatives).30 However, in the presence of downstream enzymes, such as RebC, StaC and EspM, decarboxylation and oxidation are effectively guided towards a particular outcome. In the erdasporine pathway (esp), EspM intercepts a non-oxidized, decarboxylated intermediate and methylates the remaining carboxylic acid to the methyl ester forming erdasporine B.44 In the rebeccamycin and staurosprine pathways, RebC and StaC are proposed to intercept 7-hydroxy-K252c, where RebC catalyzes a further decarboxylation-oxidation to the maleimide and StaC catalyzes decarboxylation to the pyrrolinone.45,46 Single amino acid mutations in both RebC and StaC can interchange their activities,47,48 and a RebC variant with 10 mutated residues (RebC-10x) shows improved StaC-like activity.46 Crystallographic trapping of indolocarbazole intermediates in RebC and RebC-10x isolates 7-carboxy-K252c in each active site; however, the StaC-like RebC-10x traps the keto-tautomer, amenable to decarboxylation,46 and RebC traps the enol-tautomer, stabilized for specific hydroxylation followed by decarboxylation.45 Additionally, the FAD cofactor in the StaC-like enzyme is displaced from the conformation poised for activity as observed in RebC.46 Thus, the cofactor may serve a different role in StaC, perhaps driving decarboxylation. 1.3.4 Indolotryptolines The cladoniamides were the first natural product indolotryptolines reported in scientific literature isolated in 2008.49 This group of bacterial bisindoles are analogous to the indolocarbazoles with an apparent flipped indole ring. The compound BE-54017 (abe) was 17 found via searching soil environmental DNA samples for CPA synthases.32 Heterologous reconstitution showed indolotryptoline biosynthesis proceeds through a maleimide indolocarbazole intermediate, where flavoenzyme AbeX1 catalyzes the first committed step to indolotryptolines and requires follow up by flavoenzyme AbeX2 to complete the indole ring flip. Biosynthetic studies on the cladoniamide pathway (cla) shows that ClaX1 produces the C4c-C7a cis-diol, followed by N-methylation by ClaM1 proposed to protect against intermediate degradation, and oxidative scaffold rearrangement by ClaX2 (Figure 1.6).31 Cladoniamide and BE-54017 differ only by a methyl group found on the hydroxylated tryptoline moiety. Figure 1.6 Proposed biosynthetic steps towards indolotryptoline cladoniamide C. 18 BE-54017 inhibits vacuolar H+-ATPase, and borregomycin is a strong CaMKIIδ inhibitor.25,32 The authors speculated that the non-planar structure limits its general cytotoxicity as a DNA intercalator, as seen in similar kinase inhibitors of fascaplysin and other indolocarbazoles. While the key steps in indolotryptoline biosynthesis have not yet been characterized in vitro, in vivo expression studies and experimental outcomes show a similar pattern in the creation of unstable intermediates, followed by enzymatic capture and chemical stabilization. 1.4 Other amino acid oxidative entries into biosynthetic pathways PLP-dependent enzymes are well known for their ability to catalyze reverse aldol reactions, decarboxylations and aminotransfers via the cofactor acting as an electron sink and stabilizing an effective carbanion intermediate.50 Molecular oxygen’s electron configuration is in a triplet ground state, necessitating reactivity pathways involving single electron transfers rather than the typical two-electron transfers of reactions involving the PLP cofactor. However, any enzyme with that sufficiently stabilizes an intermediate with carbanion resonance character may react with electrophiles, including oxygen.51 Throughout the 2000s it was demonstrated that PLP-dependent decarboxylases are able to not only react with oxygen, but seemingly catalyze non-native oxidation-decarboxylations either via the introduction of non-natural substrates or single-residue mutations.52 The first PLP-dependent enzymes discovered to have dedicated oxygen reactivity were the plant aromatic aldehyde synthases (AASs).50,53 These catalyze oxidative-decarboxylations consuming oxygen and producing stoichiometric ammonia, hydrogen peroxide and carbon dioxide and are involved in creating plant scent compounds such as phenylacetaldehyde. This is similar to the non-natural reactions described previously, but producing a singular product, rather 19 than a mixture of decarboxylation, oxidative deamination and abortive transamination products. Arginine oxidases (AOs) followed as the second example of PLP-dependent enzymes with dedicated oxygen reactivity.50,54 These catalyze the oxidation of Arg to (S)-4-hydroxy-2-iminoarginine or 4,5-dehydro-2-iminoarginine using two equivalents of oxygen, producing two equivalents of hydrogen peroxide. This oxidative manipulation of L-arginine (Arg) serves a key step in the initiation of at least three biosynthetic pathways to natural products: L-enduracididine, azomycin, and indolmycin (Figure 1.7). Both AASs and AOs originate from distinct PLP-dependent enzyme families, suggesting dedicated oxygen-dependent activity evolved convergently. Given the ubiquity of the cofactor more variations on the oxidative capability of PLP are likely to exist. 20 Figure 1.7 Biosynthetic pathways to enduracididine, azomycin and indolmycin initiated by arginine oxidases MppP, RohP and Ind4, respectively. Work in the area of capuramycin-type NP biosynthesis led to the in vitro characterization of the PLP-dependent oxygenase enzyme Cap15.55 It catalyzes the oxidation-decarboxylation of an amino acid-nucleoside intermediate to the corresponding amide, incorporating an oxygen atom into the amide while liberating carbon dioxide and not producing hydrogen peroxide. While sharing the core PLP-binding domain sequence features the enzyme is otherwise unrelated to 21 AASs and AOs and is more closely related to the L-seryl-tRNA selenium transferase, SelA, responsible for transforming tRNA(Sec)-bound L-serine into the corresponding L-selenocysteine.56 I briefly review the biosynthesis pathways involving AOs to provide context for the study of Cap15 responsible for a branch-point in the capuramycin-type NP biosynthesis. 1.4.1 L-Enduracididine L-Enduracididine is a non-proteinogenic amino acid found in several natural products derived from L-arginine.50 Molecules that contain an enduracididine residue include teixobactin, enduracidin, mannopeptimycin.57 Silvaggi and coworkers characterized the three-gene pathway to this amino acid coding for the AO, MppP, the cyclase, MppR, and aminotransferase, MppQ (Figure 1.7, left).54 MppP first converts L-arginine to 4-hydroxy-2-ketoarginine, which has now been shown to be a 4-electron oxidation utilizing two molecules of oxygen and producing two molecules of hydrogen peroxide. MppP was surprisingly found to be both PLP- and oxygen-dependent; the second example of dedicated oxygen-consuming activity in PLP-dependent enzymes. Then, MppR dehydrates and cyclizes 4-hydroxy-2-ketoarginine, and MppQ is a proposed aminotransferase. The MppP crystal structure allowed for the stereochemical assignment of the reaction product as (S)-4-hydroxy-2-ketoarginine. 1.4.2 Azomycin Azomycin or 2-nitroimidazole is a common core in various synthetic antimicrobial drugs.58,59 Studies from 1966-1970 established 2-aminoimidazole is a key intermediate on route to azomycin. A 1977 study found 4-hydroxy-2-ketoarginine to be a key precursor to 2-aminoimidazole, and the logical proposition was that that 2-aminoimidazole forms from a retroaldol-type reaction on 4-hydroxy-2-ketoarginine yielding pyruvate and 2-22 guanidinoacetaldehyde which subsequently cyclizes to 2-aminoimidazole.60 Furthermore, they showed that 4-hydroxy-2-ketoarginine formation was dependent on oxygen and PLP. Decades later, our research group explored gene clusters encoding sequence-related enzymes to Ind4 and MppP. A detailed biochemical and structural exploration of the encoded arginine oxidases from one such 5-gene cluster (rohPQRST) led to the identification of RohP – the enzyme responsible for 4-hydroxy-2-ketoarginine synthesis from arginine.61 Subsequent studies led to the in vitro reconstitution of the rohPQRST gene cluster, linking it to azomycin biosynthesis, thereby solving the mystery of the identity of the azomycin gene cluster.62 In these studies, RohR was shown to be the retroaldolase creating 2-guanidinoacetaldehyde from 4-hydroxy-2-ketoarginine (Figure 1.7, centre). Furthermore, RohQ is a cyclase catalyzing ring closure forming 2-aminoimidazole, and RohS oxidizes the amino group up to a nitro moiety yielding azomycin. The PLP-dependent enzyme RohP operates identically to MppP, hydroxylating L-arginine and consuming oxygen.54,61 Two pathways are possible: either a net 4-electron oxidation consuming two oxygen molecules, producing 4-hydroxy-2-ketoarginine, or a net 2-electron oxidation consuming a single oxygen molecule and producing 2-ketoarginine. In either case, oxygen is stoichiometrically converted to hydrogen peroxide. Using 50 % H218O showed the hydroxyl group is sourced from solvent water as opposed to hydrogen peroxide. Crystal structures of RohP show the enzyme traps the internal aldimine, quinonoid I and II of the external aldimine and product bound holo-enzyme, depending on the length of incubation with the substrate arginine and oxygen.61 (S)-4-hydroxy-2-ketoarginine is the configuration of the product as modelled in the crystal structure, similar to the MppP product. At least one RohP-23 quinonoid intermediate complex shows a solvent water molecule pre-arranged for addition to the arginine intermediate. 1.4.3 Indolmycin Indolmycin is an antibiotic discovered in 1960 inhibiting tryptophanyl-tRNA synthetase.50 It is constructed via a branched biosynthetic pathway where L-tryptophan is converted to indolmycenic acid through three steps: aminotransfer, C-methylation and 𝛼-ketone reduction (Figure 1.7, right).63 L-arginine is converted to D-4,5-dehydroarginine in two steps and subsequently combined with indolmycenic acid to form indolmycin A and B. The desaturation reaction of Ind4, which produces 4,5-dehydro-2-iminoarginine from L-arginine, is the critical transformation enabling oxazolinone ring formation in indolmycin.64 The Ind4 reaction is a 4-electron oxidation utilizing two molecules of oxygen and producing two molecules of hydrogen peroxide. Additionally, 2-electron oxidation intermediates can be isolated from reaction mixtures originating from 2-iminoarginine. The reaction proceeds without additional PLP, PMP or 2-ketoarginine. The related enzyme Plu4 from Pseudoalteromonas luteoviolacea indolmycin biosynthesis catalyzes the same reaction as Ind4, albeit with a different pH optimum and some in vitro accumulation of hydroxylated ketoarginine.65 Site-directed mutagenesis of key residues in the active site were not able to convert Plu4 from a desaturase to a hydroxylase, and a more holistic approach was taken to compare the structure of the Plu4 active site and to that of hydroxylase RohP via molecular dynamics simulations. These show the hydroperoxyl-external aldimine intermediate in RohP hydrogen bonds to a water molecule, positioning it for enantioselective C4 substitution; whereas, in Plu4 alternate conformations are preferred where water is not selectively organized for substitution, implying primarily desaturating activity. Plu4 active site 24 conformations exist that do pre-organize a water similarly to RohP and may explain the small amount of hydroxylating activity. The simulations point to much more difficult to characterize multiple-residue interactions resulting in conformational shifts in the active site may distinguish hydroxylation and desaturation activity between arginine oxidases. 1.4.4 Capuramycin Capuramycin-type nucleoside antibiotics are typically potent inhibitors of target bacterial translocase I.66,67 A number of capuramycin-type nucleoside antibiotics have been isolated. For example, capuramycin was isolated in 1986;68 the A-500359s were isolated from Streptomyces griseus SANK 60196, where A-500359 B is identical to capuramycin;66 the A-503083s came from Streptomyces sp. SANK 62799;69 and A-102395 came from Amycolotopsis sp. SANK 60206.70 The capuramycin-type nucleoside antibiotics are composed of a uridine-5ʹ-carboxamide (CarU) core and an unsaturated hexuronic acid attached to CarU via a 5ʹ-O-glycosidic bond. The hexuronic acid is joined via an amide to an L-aminocaprolactam in A-500359s and A-503083s or an arylamine-containing polyamide in A-102395. Liposidomycin-type molecules are related to capuramycin-type NPs in that instead of a 5ʹ-carboxamide functionalization of the core nucleoside they instead have a 5ʹ-glycyl extension (GlyU). These include the liposidomycins, A-90289s, caprazamycins, muraminomicins, muraymycins, and polyoxins, and vary in their glycosyl, fatty acid and amino acid substitution patterns (Figure 1.8).71,72 25 Figure 1.8 Selection of liposidomycin-type and capuramycin-type NPs with their defining GlyU and CarU groups highlighted in red and blue, respectively. Biosynthetic gene clusters for three capuramycin-type have been characterized and each has 18 homologous genes involved in biosynthesis, regulation and resistance.73–75 In these pathways, CarU is synthesized from L-threonine and uridine monophosphate (UMP) in at least four steps. First, CapA, an Fe(II)- and 𝛼-ketoglutarate-dependent oxygenase, oxidatively dephosphorylates UMP to uridine-5ʹ-aldehyde (U5ʹA). Second, CapH, related to PLP-dependent serine hydroxymethyltransferases, acts on L-threonine and U5ʹA, catalyzing the transfer of a glycyl group to U5ʹA forming GlyU with release of acetaldehyde.76 Third, Cap15 decarboxylates and oxidizes GlyU to CarU, incorporating an oxygen atom from molecular oxygen into CarU (Figure 1.9).55 In vitro assays show that Cap15 is not active on the CapA product ((5ʹS, 6ʹS)-GlyU) necessitating an epimerization step to (5ʹS, 6ʹR)-GlyU; however, the enzyme responsible 26 has not yet been identified. Cap15 is the key enzymatic step distinguishing capuramycin-type NP biosynthesis from liposidomycin-type. Figure 1.9 Uridine 5ʹ-monophosphate (UMP) is transformed into CarU in at least four steps. In capuramycin biosynthesis UMP is oxidatively desphosphorlyated to uridine 5ʹ-aldehyde (U5A) by the iron:𝛼-ketoglutarate-dependent CapA followed by transaldolation of L-threonine to U5A via CapH transferring a glycyl unit yielding (5ʹS, 6ʹS)-glycyluridine (GlyU) and releasing acetaldehyde. In vitro assays show Cap15 catalyzes O2-, PLP-dependent oxidative-decarboxylation of (5ʹS, 6ʹR)-GlyU to CarU, necessitating an intermediate 𝛼-epimerization step. 1.5 Objective of Dissertation X-ray crystallography has played a major role in the characterization of biosynthetic enzymes in bisindole biosynthetic pathways. For example, the mechanism of VioE from violacein biosynthesis was partially elucidated by data from X-ray crystallography, and, similarly, major insights came from the structures of the indolocarbazole biosynthetic enzymes StaP, RebC, and StaC. In the field of oxygen- and pyridoxal phosphate-dependent enzymology, X-ray crystallography has also provided major insights into the arginine oxidases. 27 In this thesis, I similarly sought to use enzymatic and structural characterization to provide insight into the reductasporine biosynthetic enzymes RedE and RedM, along with the capuramycin biosynthetic enzyme Cap15, an O2-, PLP-dependent enzyme. Chapter 2 discusses the in vitro reconstitution of reductasporine biosynthesis along with structural characterization of the reductasporine biosynthetic enzyme RedM. Chapter 3 is focused on the structural characterization of the reducasporine biosynthetic enzyme RedE. Chapter 4 is the structural characterization of the capuramycin biosynthetic enzyme Cap15. Finally, Chapter 5 discusses these results together and indicates future areas for further investigation. 28 Chapter 2: Structure of methyltransferase RedM that forms the dimethylpyrrolinium of the bisindole reductasporine 2.1 Synopsis Bisindoles are biologically active natural products that arise from the oxidative dimerization of two molecules of L-tryptophan. In bacterial bisindole pathways, a core set of transformations is followed by the action of diverse tailoring enzymes that catalyze reactions that lead to diverse bisindole products. Among bisindoles, reductasporine is unique due to its dimethylpyrrolinium structure. Its previously reported biosynthetic gene cluster encodes two unique tailoring enzymes, the imine reductase RedE and the dimethyltransferase RedM, which were shown to assemble reductasporine through heterologous expression. To gain more insight into the unique tailoring enzymes in reductasporine assembly, I reconstituted the biosynthetic pathway to reductasporine in vitro and then solved the 1.7 Å resolution structure of RedM. Our work reveals key conformational changes that occur in RedM and identifies key active site residues. Our work sets the stage for understanding how RedM evolved to react with a pyrrolinium scaffold and may enable development of new dimethyltransferase catalysts. 2.2 Introduction Bacterial bisindoles are a group of alkaloid natural products that arise from the oxidative dimerization of L-tryptophan.77,78 The first bisindole isolated was the purple pigment violacein in 1882,79 and the discovery of the potent kinase inhibitor staurosporine in 197780 led to huge interest in the large class of indolocarbazole natural products as drug leads starting in the late 1980s81 (Figure 2.1A). Biosynthetic studies beginning in the 2000s revealed that bacterial indolocarbazoles arise from the oxidative dimerization of L-tryptophan through three key steps: (1) oxidation to indole pyruvate enamine; (2) oxidative dimerization to give chromopyrrolic acid, 29 and (3) aryl-aryl coupling to give the indolocarbazole structure (Figure 2.1C).82–84 A combination of enzyme-mediated and non-enzymatic oxidation then gives the core aglycone structure observed in diverse bisindoles84,85 To target new bacterial bisindole gene clusters and their encoded metabolites, Brady and co-workers developed degenerate primers specific to the chromopyrrolic acid synthase genes to enable the targeted isolation of bisindole biosynthetic gene clusters from environmental DNA.86 Using these primers, they isolated novel biosynthetic gene clusters, and through various reconstitution techniques, reported a number of new bisindole structures, including erdasporine and reductasporine (Figure 2.1A).87 Reductasporine is an unusual dimethylpyrrolinium structure whose synthesis is linked to the six-gene cluster, redODPEMT (Figure 2.1B).87 While the intact or refactored gene cluster failed to give any metabolites when expressed in E. coli, the Brady group was able to express the core indolocarbazole genes espODP from the erdasporine gene cluster88 to generate the conserved bisindole intermediates in E. coli and addition of redE and redM genes gave reductasporine, thus linking redE and redM to reductasporine production. Our lab became interested in both RedE and RedM, as they are unusual biosynthetic enzymes among the arsenal of indolocarbazole enzymes. In particular, RedM is a rare N,N-dimethyltransferase. Here I report the in vitro activity of RedE and RedM and solve a 1.7 Å resolution structure of RedM in complex with each SAM and SAH. Our work reveals a large conformational change in RedM that may be relevant to catalysis, along with identifying key active site residues. 30 Figure 2.1 Indolocarbazoles, their gene clusters and shared biosynthetic enzymes. a) Natural product indolocarbazoles. b) Gene clusters for reductasporine (red), erdasporine (esp), staurosporine (sta), and rebeccamycin (reb). Indolocarbazole is abbreviated ICZ. c) Shared biosynthetic assembly of indolocarbazoles. 2.3 Results 2.3.1 In vitro reconstitution of reductasporine biosynthesis Indolocarbazoles are formed from L-tryptophan through a three-enzyme pathway (Figure 2.1C). Previous work by Chang and coworkers showed production of reductasporine in E. coli + espODP + redEM, and removal of either redE or redM gave only indolocarbazole byproducts. Chang and coworkers put forward a reasonable biosynthetic proposal where RedE intercepts an unstable intermediate product, 12,13-dihydro-6H-indolo[2,3-a]pyrrolo[3,4-c]carbazole (4), from the reaction of the cytochrome P450 enzyme. RedE reduces compound 4 to didemthylreductasporine (3) and then RedM catalyzes two methylations to give reductasporine (Figure 2.2A). 31 Figure 2.2 In vitro reconstitution of reductasporine biosynthesis. A, Proposed pathway for formation of reductasporine from chromopyrrolic acid. B, HPLC-MS traces (base peak count) of 300 µM CPA, 10 mM NADPH, 3 mM SAM, 5 µM StaP 10 µM Synechococcus elongatus ferredoxin (FDX), 1 µM Synechococcus elongatus ferredoxin reductase (FDR) and 50 mM HEPES pH 8 in the presence of active and/or boiled (denoted as b) RedE and RedM (20 µM each). C, Methylation of synthetic 3 to reductasporine by RedM (2 µM RedM 2 mM SAM and 500 µM 3 over 25 min). 32 To provide in vitro evidence for the accumulation of proposed intermediates and the functions of enzymes RedE and RedM, I sought to reconstitute the pathway with purified enzymes (Figure 2.3). I isolated CPA from the cleared E. coli lysates containing RebO and RebD incubated with tryptophan. I purified RedE, RedM, cytochrome P450 StaP, Synechococcus elongatus ferredoxin (UniProt ID P0A3D2) and ferredoxin reductase (UniProt ID Q31PL1), each recombinantly expressed in E. coli.89 In control reactions, StaP together with ferredoxin, ferredoxin reductase, and NADPH consumes chromopyrrolic acid, as expected. Inclusion of RedE results in the accumulation of a new product ([M+H] = 298), with a mass corresponding to 3. Inclusion of both RedE and RedM yields reductasporine as verified via NMR (Figure 2.2B, Figure 2.4). To further interrogate the reaction, I directly incubated synthetic 3 with RedM with SAM and observe via LC-MS analysis the accumulation and disappearance of a peak consistent with the monomethylated intermediate (2) over 25 minutes (Figure 2.2C), providing initial evidence that the dimethylation occurs over discrete steps. Figure 2.3 Representative SDS-PAGE gels of recombinant RedE and RedM Ni-IMAC and size-exclusion chromatography fractions. P, cell debris pellet; S, supernatant; FT, column flow-through. 33 Figure 2.4 1H-NMR (600 MHz, DMSO-d6) of reductasporine isolated from a large-scale in vitro reconstitution of biosynthetic pathway from CPA. 34 2.3.2 Crystal structure of RedM To provide insight into the unusual N,N-dimethylation catalyzed by RedM, I solved the structure of selenomethionine-labelled RedM, and used this model to solve crystal structures of the apo enzyme, as well as a co-crystal structure with SAH and a co-crystal structure with SAM, all to 1.7-1.85 Å resolution (Table 2.1). Attempts to soak in or co-crystallize arcyriaflavin A, reductasporine or indolocarbazole precursor chromopyrrolic acid did not yield ligand bound structures. Figure 2.5 Structure of RedM. A, Topology diagram with labelled secondary structures. N-terminal dimerization domain highlighted in pink and cyan, and C-terminal Rossman-like SAM-binding domain coloured orange and red. Helix 𝛼10 is omitted from the diagram as it is not found on all protomers and borders the gap between the N-terminal domain and interdomain helix. B, Overall fold of RedM with SAH bound (black). The domain bridging 𝛼11 helix is indicated with a triangle. C, RedM-SAH and -SAM binary complexes superposed by their dimerization domains. One protomer of each dimer in the RedM-SAM crystal structure asymmetric unit shows a distinct deviation from the more open conformation seen in the RedM-SAH complex. The more constricted, closed conformation arises from the Rossman-like domain’s rotation towards the dimerization domain (dashed arrow) and a substantial deflection of the interdomain 𝛼11 helix (triangle). 35 Table 2.1 Data collection and refinement statistics. a apo RedM RedM-SAH RedM-SAM PDB Accession 8TJI 8TJK 8TJJ Wavelength (nm) 0.97946 0.97946 0.97946 Resolution range (Å) 72.5–1.73 (1.76–1.73) 38.1–1.85 (1.89–1.85) 89.2–1.82 (1.85–1.82) Space group P 21 21 21 P 21 21 21 P 21 Unit cell 61.95 71.61 145.00 61.83 71.90 144.84 113.86 58.78 113.92 90 103.11 90 Total reflections 778150 (21218) 729957 (44012) 823931 (41491) Unique reflections 67710 (3483) 55851 (3356) 125933 (6228) Multiplicity 11.5 (6.1) 13.1 (13.1) 6.5 (6.7) Completeness (%) 99.4 (95.0) 99.9 (98.4) 95.9 (96.7) Mean I/sigma(I) 23.6 (5.8) 15.4 (1.2) 9.5 (1.6) Wilson B-factor 17.89 38.1 21.01 Rmerge 0.065 (0.241) 0.068 (1.703) 0.146 (1.215) Rmeas 0.067 (0.263) 0.070 (1.772) 0.159 (1.316) Rpim 0.019 (0.101) 0.019 (0.485) 0.061 (0.501) CC1/2 0.999 (0.960) 0.999 (0.730) 0.997 (0.367) Reflections used in refinement 67168 (6471) 55630 (5406) 125905 (12596) Reflections used for Rfree 3390 (337) 1993 (193) 6531 (622) Rwork 0.1669 (0.2045) 0.1969 (0.3635) 0.1753 (0.2946) Rfree 0.1992 (0.2339) 0.2285 (0.3896) 0.2175 (0.3301) Total non-hydrogen atoms 5726 5084 11226 macromolecules 4835 4728 9814 ligands 36 11 179 solvent 855 345 1233 Protein residues 631 631 1297 RMS (bonds) 0.004 0.005 0.005 RMS (angles) 0.68 0.75 0.77 Ramachandran favored (%) 98.39 98.23 97.58 Ramachandran allowed (%) 1.61 1.77 2.42 Ramachandran outliers (%) 0.00 0.00 0.00 Rotamer outliers (%) 0.21 0.68 0.73 Clashscore 1.86 1.50 2.68 Average B-factor 21.98 45.45 30.81 macromolecules 19.93 45.17 29.98 ligands 31.15 57.83 37.63 solvent 33.24 48.87 36.47 Number of TLS groups 14 4 17 a Statistics for the highest-resolution shell are shown in parentheses. 36 RedM is a homodimeric protein with monomers consisting of a largely 𝛼-helical N-terminal domain (𝛼1-10, 𝛽1-3) and a Rossman-like fold C-terminal domain (𝛼12-16, 𝛽4-10) bridged by a long interdomain 𝛼-helix (𝛼11) (Figure 2.5A,B). Size-exclusion chromatography supports that RedM exists as a homodimer in solution (Figure 2.6). The N-terminal domains of the homodimer form the majority of the dimer interface with the only contact with the C-terminal domain occurring between 𝛼5, 𝛼8, 𝛼9 (residues 62-73, 112-125) and 𝛼15ʹ (residues 292-304). The extensive dimer interface is approximately 3150 Å2, accounting for nearly one-fifth of the total monomer surface area, as determined by PISA.90 These buried residues in the interface are primarily hydrophobic, and the few polar residues are small serine or threonine forming hydrogen bonds to nearby peptide bonds. The two domains of each monomer are separated by a gap of ~12 amino acids between residues ~132 and ~144 ahead of the long interdomain 𝛼-helix (𝛼11), which lacks clear electron density in each structure. In RedM-SAM cocrystal structures 𝛼11 adopts a pronounced kink at either M157 or M162 in several protomers. The C-terminal domain possesses a Rossman-like fold typical of Class I SAM-dependent methyltransferases, with a 𝛽-sheet of five parallel strands terminated with two antiparallel strands each interspersed with 𝛼-helices.91–93 SAM binds in the structurally conserved cleft between the sharp turn after 𝛽4 and the end of the 𝛽5 strand. 37 Figure 2.6 Size-exclusion chromatography traces for RedM (red) and molecular weight standards. Absorbance read at 280 nm. RedM dimer, 76 kDa; RedM monomer, 38 kDa; sweet potato 𝛽-amylase, 200 kDa; yeast alcohol dehydrogenase, 150 kDa; bovine serum albumin, 66.5 kDa; bovine carbonic anhydrase, 29 kDa; equine cytochrome c, 12 kDa Secondary structure matching searches using PDBeFOLD94 ranks RedM as most related to other SAM-dependent enzymes acting on small molecules, all of which are part of the caffeic acid O-methyltransferase-type family (COMT-type, IPR016461). The full-length monomer of RedM bears the greatest structural similarity to the human N-acetylserotonin O-methyltransferase (ASMT, PDB: 4A6E, 29 % identity, rmsdC𝛼 = 1.6 Å, 291 C𝛼). Other highly ranked structural homologues include: PigF (7CLF), an O-methyltransferase involved in prodigiosin biosynthesis;95 PdxI (7BQK), a pericyclase related to the LepI pericyclase in leporin biosynthesis;96,97 Ca9OMT (4E70); an O-methyltransferase modifying the lignin precursor coniferyl alcohol;98 BchU (1X19), a C-methyltransferase involved in bacteriochlorophyll C 02040608010012014040 50 60 70 80 90 100 110Absorbance (mAU)Volume (mL)66 kDa150 kDa200 kDaBlue Dextran(2000 kDa)12 kDa29 kDa38 biosynthesis;99 and PhzM (2IP2), an N-methyltransferase part of the pyocyanin biosynthetic pathway.100 Each retains the glycine rich sequence along with the neighbouring acidic residue indicative of the SAM binding cleft.91–93 Figure 2.7 RedM active site residue contacts. A, Residue contacts with SAH in the open conformation and B, SAM in the closed conformation. The conserved SAM-binding motifs I-II (D182-G184, D207) are highlighted and form key hydrogen bonds to the 𝛼-amine and ribosyl moiety. The closed conformation brings residues F153 and M157 to within 4 Å of the active methylsulphur moiety. The homodimers of both apo-RedM and RedM-SAH structures superpose well on each other (rmsdC𝛼 = 0.175, 553 C𝛼) and comparatively poorly upon the RedM-SAM homodimer (rmsdC𝛼 = 1.244, 593 C𝛼) due to a substantial displacement of the C-terminal Rossman-like SAM-binding domain (Figure 2.5C). The movement is an approximate 12° rotation towards 𝛼11 about an axis aligned with 𝛼15, where the loop regions around 𝛼15 act as hinges. Residues furthest from 𝛼15 are displaced up to 7 Å. Helix 𝛼11 accommodates this conformational change by developing a kink at M157 or S161 with 31–49° changes in the Phi and Psi angles at these residues. The overall length of the monomer is reduced 2.5 Å and the central binding cavity 39 volume is reduced by ~60 % (estimated via POCASA101), and likely represents a closed conformation of RedM. The cofactor binding pocket is outlined by the C-terminal ends of 𝛽4-8 and clear electron density shows the cofactor is held in an extended conformation similar to related small molecule methyltransferases.91–93 These Class I methyltransferases bear the conserved E/DxGxGxG SAM-binding motif and is found as 182-DIGCAEG-188 in RedM. The main chains of C185, A186 and G188 form the foundation to the cavity between the ribosyl and methionine moieties, while D182 hydrogen bonds to two conserved water positions creating the basis for a network of hydrogen bonds bridging the 𝛼-amine to I183, Y280 and G249 via its amide nitrogen (Figure 2.7). The carbonyl groups of G184 and G249 receive hydrogen bonds from the 𝛼-amine, while H250 and Y280 donate hydrogen bonds to the cofactor carboxylate group. The ribosyl hydroxyl groups interact closely with D207, representing the second conserved binding motif in SAM-dependent methyltransferases.93 N254 and W255 hydrogen bond a third conserved water position which bridges to the furan oxygen. The adenine ring is clamped between F235 and L208 and capped with a hydrogen bond to D234. Upon closure of the active site, residues F153 and M157 from the interdomain helix 𝛼11 are brought to within 4 Å of the SAM methylsulfur, displacing solvent (Figure 2.7B). 2.3.3 Modelling the RedM-substrate complex Despite efforts to co-crystallize RedM with reductasporine or substrate analogues, no clear electron density for additional ligands was resolved. I therefore focused on modelling the RedM-substrate complexes via docking experiments, noting that the rigidity of the indolocarbazole skeleton might limit the substrate to a single conformation. 40 Docking search with 3 on the open conformation places moderately favorable poses at the junction of helices 𝛼7, 𝛼11, 𝛼15 and 𝛼ʹ1 and 𝛼ʹ2 of the dimer partner, an average of 20 Å away from the bound SAH cofactor (Figure 2.8A). This junction is lined with mainly hydrophobic residues (L18, M25, L104, M112, P293, L296, M297, P336), and several small polar residues (D15 Q19, T22, S108, T300). D15 may be engaged in a salt bridge to R163 joining the N-terminus to the interdomain helix (𝛼11); however, this interaction only has unambiguous electron density in one monomer of the RedM-SAH complex. In docking the substrate to the closed conformation several poses remain in this distal area, but higher-scoring poses occur near SAM (Figure 2.8B). These poses have strained geometry between the donor methylsulfur and acceptor nitrogen for methyl transfer. Allowing free rotation of the N254 side chain in the docking procedure yields binding conformations with near ideal geometry (Figure 2.8C). Docking 2 under identical settings results in poses with close proximity to enable methyl transfer, but non-ideal geometry. Introducing free rotation in neighbouring residues S253, M297 and M301 yielded more poses in the vicinity of SAM with similar geometry. 41 Figure 2.8 Docking poses of didemethylreductasporine (3) into the open and closed conformations of RedM. A, The open conformation shows no favourable binding in the vicinity of SAH and poses aggregate among the hydrophobic residues of the N-terminal dimerization domain. B, Docked poses in the closed conformation show that constriction of the binding cavity forms a pocket for indolocarbazole binding. Several poses have favourable 42 geometry for methyl transfer. C, Active site residues surrounding a docked 3 pose with pyrroline nitrogen positioned in line with and 3.6 Å away from the SAM methylsulfur group. 3 is docked to the closed conformation of RedM and the N254 sidechain was allowed to rotate freely during docking. Five methionine residues and F153 pack closely with the indolocarbazole aromatic ring system. Docking search space spans the entire internal cavity, and distances are shown in units of Angstrom. A putative indolocarbazole binding site can be described using these docking results as a guide (Figure 2.8C). The cavity is located at the junction of helices 𝛼7, 𝛼8, 𝛼11, 𝛼15 and the loop between 𝛽7 and 𝛼14, forming a roughly 7 Å diameter hole through the center of the protomer in the open conformation. The area is lined with hydrophobic residues and are mostly methionine residues (L104, M107, M112, F153, M157, L160, M297, M301). Methionine sulfur-𝜋 interactions are noted as stabilizing contacts with aromatic groups.102,103 The closure of this cavity brings residues L160 and M297 within 4 Å of each other and forms the hydrophobic floor to the active binding pocket. R111 and E304 form a salt bridge overtop the cavity and joins to 𝛼15 to the N-terminal helical bundle. N254 is in close proximity to the pyrroline nitrogen of the indolocarbazole substrate and may hydrogen bond in some conformations. The complete active site may include the residues with unresolved electron density in the range from 133-149, which form two 𝛼-helices capping the SAM-binding cavity in similar methyltransferase structures.100,104–106 2.3.4 Mutagenesis of the RedM active site In structural homologues, the residue at position S253 in RedM is typically a histidine and has been shown to be involved in deprotonation, either deprotonating the substrate before methylation, or deprotonating the resulting product (Figure 2.9).104,107–109 Coniferyl alcohol 9-O-methyltransferase instead has a cysteine at this position, and this cysteine is implicated by 43 mutagenesis as catalytically important, possibly taking part in acid-base chemistry.98 These residues catalyzing proton transfer are also associated with one or two neighbouring aspartate or glutamate residues thought to position the active residue for proton abstraction or may participate as a catalytic partner.104,107,110–114 In RedM, these residues are E281 and E310, with E281 positioned ideally for H-bonding to S253, and with E310 more than 4 Å away from S253. I generated the S253A and S253C variants, neither of which altered product formation; however, S253H lost activity (Figure 2.10A). Variant E281Q did not affect product formation. Several other structurally characterized methyltransferases do not conserve the catalytic histidine, including carminomycin 4-O-MT (DnrK),115 3-hydroxykynurenine C-MT (SibL),116 dimethylsulfoniopropionate S-MT (DsyB),117 8-demethyl-8-amino-D-riboflavin N-MT (RosA)105 and phenazine N-MT (PhzM).100 Mutational studies in SibL, DnrK and DsyB indicate a tyrosine in the 𝛼-helix capping the active site is essential for activity in SibL and significant for activity in DnrK and DsyB.115–117 Aligning these enzymes with PhzM, RosA and RedM shows that each conserves this tyrosine, and variant RedM Y140F results in substantial loss in product formation (Figure 2.10A). 44 Figure 2.9 Sequence alignment of RedM against representative structural homologues available in the PDB grouped by presence or lack of the catalytic histidine found as serine at position 253 in RedM. 45 Residues conserved above 80 % identity are highlighted blue, and positions of catalytically important positions highlighted in pink. RedM substrate binding residues are marked with orange circles, SAM-binding residues with green squares, and residues mutated in RedM with arrows. Alignment was generated using COBALT.118 UniProt accessions numbers: RedM (A0A0F7G196), RosA (K4RFM2), PhzM (Q9HWH2), SibL (C0LTM6), DnrK (Q06528), TnmH (A0A125SA05), MmcR (Q9X5T6), NcsB1 (Q84HC8), COMT-S (A0A4P8DY91), MsCOMT (P28002), PsSOMT1 (I3V6A7). NCBI accession numbers: DsyB (WP_028467200.1). Next, I sought to perturb the N,N-dimethylating activity of RedM. Docking indicates the pocket opposite the SAM cofactor may accommodate a methyl group during the second methyl transfer. It is lined by residues H250, S253, N254, E281, T282 M297, M301 and E310, and most of these residues are associated with substrate recognition or with potentially catalytic residues. I generated H250W, despite the association of the histidine with SAM-binding, as many plant O-methyltransferases conserve a tryptophan at this position, and the extra bulk may occlude a methyl from the adjacent pocket. The H250W variant had a complete loss of activity. Docking results suggest the area around 𝛼1 and 𝛼2 may provide a pre-catalysis staging area for indolocarbazoles before moving through the central tunnel to the active site (Figure 2.8A,B). T22, in the bend between 𝛼1 and 𝛼2, points directly into this area and mutation to a larger residue may interfere with substrate access. However, the T22Q variant has no impact on product formation (Figure 2.10A). 46 Figure 2.10 In vitro assays of RedM variants and active site comparisons to COMT-type homologues. A, Base peak LC-MS traces of CPA+StaP+RedE+RedM variant reaction mixes incubated at room temperature. Complete conversion to reductasporine (m/z = 326, blue) is seen after one hour alongside a background peak (m/z = 310, orange). Complete loss in product accumulation is seen in RedM-S253H and -H250W and partial loss in RedM-Y140F. B, Active site docking of didemethylreductasporine (3) with residues targeted for mutation rendered as light green sticks. Equivalent residues and cofactors in DnrK (pink, 1TW2), CA9OMT (cyan, 4EVI) and SibL (orange, 4U1Q) are overlayed for comparison. Y140 is not resolved in the structural models of RedM; however, its position may be inferred from these related methyltransferases. Structures are aligned by the Rossman-like domain (DnrK, RMSD = 0.99 Å, 133 C𝛼; CA9OMT, 1.32 Å, 155 C𝛼; SibL, 1.04 Å, 154 C𝛼) and Y142, Y162 and Y134 from each are an average of 4.3 Å from the cofactor sulfur atom. 47 2.3.5 Sequence Similarity Network of COMT-type Methyltransferases In order to visualize the distribution of active site motifs I created a sequence similarity network (SSN) of the COMT-type enzyme family (IPR016461) to which RedM belongs. The initial SSN at a permissive alignment score threshold (AST ≥ 29) divides the family into one main cluster and four small clusters (Figure 2.11A). One includes MppJ, which binds iron in its active site in order to catalyze the unusual benzylic C-methylation of phenylpyruvate.119 This iron binding site is unique in the COMT-type enzyme family. At a more restrictive alignment score threshold (≥ 50) the central cluster divides into four large clusters with more than 70 nodes. Most of the structurally characterized enzymes (19 of 41) are in the central, highly connected cluster and includes the first structurally characterized plant small molecule methyltransferases chalcone O-methyltransferase (ChOMT) and isoflavone-7-O-methyltransferase (IOMT)104, and many methyltransferases participating in secondary metabolism (phenazines, bergaptol, scoulerine, flavonoids and many others). This encompasses at least five methyltransferases that are capable of furnishing two methyltransfers: RosA105, xanthohumol 4-O-methyltransferase120, trans-resveratrol di-O-methyltransferase121, myricetin O-methyltransferase (CrOMT2)122, and 3-aminomethylindole N-methyltransferase (ANMT).123 𝛽-alanine N-methyltransferase (BANMT) is capable of methylating 𝛽-alanine up to three times.124 RedM occupies a more sparsely characterized cluster where the only structurally characterized enzymes are the C-methyltransferase SibL (catalyzing the C4 methylation of 3-hydroxykynurenine), O-methyltransferase PigF (part of prodigiosin biosynthesis), and human acetylserotonin O-methyltransferase.95,116,125 Other characterized proteins include the O-methyltransferase AviB2 (catalyzing methyl transfer to 3-hydroxy-5-methyl-1-naphthoate) and the O-methyltransferase TcmO (catalyzing methyl transfer to the 8-hydroxy in tetracenomycin biosynthesis).126,127 The 48 remaining two large clusters of the network contain the majority of the eukaryotic COMT-type proteins primarily from fungal organisms. The first has 18 characterized natural product methyltransferases, which includes enzymes involved in the biosynthesis of fumagillin, oxepinamides, phomasetin and kotanin, as well as the structurally characterized OxaC from oxaline biosynthesis128 and the pericyclase-dehydratase LepI from leporin biosynthesis.129 The second cluster has the structurally characterized Alder-ene pericyclases PdxI, EpiI and AdxI (from the putative pyridoxatin and fusaricide biosynthetic gene clusters)97 along with 31 characterized natural product methyltransferases and a surprising 14 characterized transcriptional coactivators similar to the aflatoxin pathway regulator AflS (formerly AflJ).130,131 Overall, the characterized methyltransferases in this family are largely O-methyltransferases (78 %) with N-methyltransferases the next most common (9 %), and the remainder characterized as transcriptional coactivators, pericyclases, C-methyltransferases and a single S-methyltransferase. Examining the RedM subcluster at an AST of 50 or above shows some discrete separation in chemical activity (Figure 2.11B-C). For example, the C-methyltransferase SibL (4U1Q) and human acetylserotonin O-methyltransferase (4A6E) partition out into distinct clusters. The O-methyltransferase in prodigiosin biosynthesis, PigF, is not well connected and bridges several clusters near ASMT. Multiple sequence alignment and searches for the breadth of active site motifs show that it is common for methionine residues to be present in the active site, specifically at positions 157, 297 and 301 (RedM numbering). This includes the common FxxxM motif found in the interdomain helix, and the MxxxM motif found in 𝛼15 in RedM (Figure 2.11C). Missing the catalytic histidine at position 253 is uncommon, and when not present it is often supplemented with an alternative active site base, such as a tyrosine in the capping helix of the active site (Y140 in RedM). This tyrosine is nearly ubiquitous in the SibL subcluster, a 49 different entirely uncharacterized subcluster nearby, and several proteins related to RedM (Figure 2.11B). Figure 2.11 Sequence similarity network of the COMT-like enzyme family (IPR016461) generated using the Uniref50132 database. A, Visualization of the network with minimum edge alignment score of 29 and coloured according to the subclusters retained at an alignment score threshold of 50, inclusive. Most crystal structures are found in the largest subcluster alongside chalcone OMT and isoflavone-7-OMT the founding members of the family (red nodes). The second largest subcluster (green nodes) contains fungal pericyclases such as PdxI (PDB: 7BQJ), transcriptional coactivators and other methyltransferases. The third largest subcluster (orange nodes) is made up of mostly secondary metabolism methyltransferses such as OxaC (5W7P) and one example of a pericyclase in LepI (6IX5). RedM is 50 found alongside human acetylserotonin MT (ASMT, 4A6E), SibL (4U1Q), demethylspheroidene O-methyltransferase (5I2H) and PigF (7CLF) (blue nodes). B, C, The cluster containing RedM isolated at an alignment score threshold of 50. B, Purple and orange nodes have no catalytic histidine and orange nodes have a tyrosine residue corresponding to Y140 in RedM. This is a common catalytic motif in sequences related to SibL and is widely present in a nearby uncharacterized subcluster. This motif is only sporadically present in sequences related to RedM. C, Nodes coloured according to putative substrate recognizing positions 153-FxxxM-157 / 297-MxxxM-301 (RedM numbering) from least stringent (purple) to most stringent (yellow). Selections: [FL]xxx[ML] / xxxx[FML], purple; [FL]xxx[ML] / [FYML]xxx[FML], maroon; FxxxM / [FYML]xxxM, orange; FxxxM / MxxxM, yellow. 2.4 Discussion In vitro reconstitution of the last steps in reductasporine biosynthesis revealed that RedE and RedM are independently functioning enzymes. RedE captures and reduces an unstable indolocarbazole intermediate, forming didemethylreductasporine (3), which is then a substrate for RedM. A time-course analysis with synthetic 3 reveals that RedM catalyzes the stepwise methylation of the RedE product, giving reductasporine. To provide insight into the structural basis for catalysis, I solved the structures of apo, SAH-, and SAM-bound RedM. Structural comparisons shows RedM to be a typical member of natural product methyltransferases, classified in the InterPro database under the COMT-type methyltransferase family (IPR016461), possessing the characteristic N-terminal dimerization and C-terminal Rossman-like domains.91,92 Comparing the crystal structures of complexes RedM-SAH and RedM-SAM shows a distinct conformational change from open to closed, respectively. In the closed conformation, residues F153 and M157 are within 4 Å of the methylsulphur cation of the SAM cofactor, and the resulting cation–𝜋 and hydrophobic interactions may stabilize the closed conformation (Figure 2.7). Chains found in the closed conformation in the RedM-SAM model have higher than average B-factors for the model and less energetically favourable crystal contacts between 51 asymmetric units. Chains found in the open conformation have average B-factors and extensive crystal contacts in all models (Figure 2.12). Poorer crystal contacts in the RedM-SAM model may result from the different crystal form of the RedM-SAM binary complex or may represent the Rossman-like domain pulling away from contact, instead being stabilized in a closed conformation by interactions with the methylsulphur cation of SAM. Nonetheless, these types of conformational changes are common in this protein family, with 11 of the 41 structurally characterized COMT-type methyltransferases displaying unambiguous open and closed conformations (Table 2.2). Given the abundance of open, closed and intermediate conformations of COMT-type methyltransferases found in the PDB it is clear that these conformational changes likely play a role in enzyme catalysis in RedM and related enzymes. apo-RedM RedM-SAM Figure 2.12 View of the crystal packing of apo-RedM and the RedM-SAM binary complex. 52 The protein backbone is rendered as C𝛼 traces and qualitative coloring by B-factor from blue to green to red. The apo-RedM structure shows low average B-factors with high values only seen in less organized loop regions. The RedM-SAM binary model shows that the crystal forms planes of well packed open conformation chains with low B-factors alternating with poorly packed closed conformation chains with higher B-factors. Table 2.2 COMT-type methyltransferase family enzymes with distinct open and closed conformations. The COMT-type methyltransferase family is characterized with an N-terminal 𝛼-helical dimerization bundle and C-terminal Rossman-like domain which binds an S-adenosyl methionine cofactor. Name Source Function Open Closed RdmB133 Streptomyces purpurascens aclacinomycin 10-hydroxylase 1xdu 1qzz 1r00 1xds NcsB1134 Streptomyces carzinostaticus neocarzinostatin O-methyltransferase 3i58 3i5u 3i64 3i53 LpCOMT111 Lolium perenne (perennial ryegrass) caffeic acid O-methyltransferase 3p9c 3p9i 3p9k MppJ119 Streptomyces hygroscopicus phenylpyruvate C(3)-methyltransferase 4kif 4kig 4kib 4kic SibL116 Streptosporangium sibiricum 3-hydroxykynurenine C-methyltransferase 4qvg 4x3q 4u1q 6OMT114 Thalictrum flavum subsp. glaucum (Yellow meadow rue) (RS)-norcoclaurine 6-O-methyltransferase 5icg 5icc 5ice 5icf PsSOMT1107 Papaver somniferum (Opium poppy) scoulerine 9-O-methyltransferase 1 6i5q 6i5z 6i6k 6i6l 6i6m 6i6n FaOMT1a Fragaria ananassa (Strawberry) furaneol O-methyltransferase 6i70 6i71 6yjw 6i72 6i73 TfSOMTa Thalictrum flavum subsp. glaucum (Yellow meadow rue) (S)-scoulerine 9-O-methyltransferase 6nei 6nej PigF95 Serratia marcescens 4-hydroxy-2,20-bipyrrole-5-carbaldehyde O-methyltransferase 7clu 7clf DsyB117 Nisaea denitrificans 4-methylthio-2-hydroxybutyrate S-methyltransferase 7wdq 7wdw a structure deposited to the PDB Docking the indolocarbazole substrate into the active site closed conformation highlights several key residues likely important to substrate recognition and catalytic activity. I wished to see if these residues are conserved in the wider COMT-type enzyme family, and whether sequence-based analyses could reveal any determinants to heteroatom methylation or dimethylating activity in general. N-methylating enzymes are found throughout the COMT-type 53 family as a relative minority representing less than 9 % of Swissprot annotated sequences. These enzymes methylate a variety of nitrogen functional groups including amines, amides, lactams, anilines and pyridines. COMT-type enzymes capable of catalyzing multiple methyl transfers have so far only been characterized in plants and bacteria and include RedM, OxyT135,136, RosA, LaPhzM, BenF, CrOMT2, ROMT, AMNT and BANMT (Table 2.3). Checking the substrates methylated by many COMT-type enzymes shows that all contain at least one aromatic ring with example substrates being monolignols,110,111 hydroxyisoflavones,104,137 (S)-norcoclaurine,114 and scoulerine107 in plants, and carminomycin,115 bacteriochlorophyllide,99 tiancimycin109 and phenazines100,106 in bacteria, and some small aromatic substrates such as acetylserotonin in animals.125 The binding cavities of these enzymes isolated in ternary complexes often contain 𝜋-𝜋 and S-𝜋 interactions between residues and the aromatic systems of the bound ligand. It has been observed that methionine residues play a role in stabilizing 𝜋 systems.102,103 This is represented in RedM where the active site is lined with five methionine residues, with M157, M297 and M301 forming major contacts with the docked indolocarbazole substrate. The first is found in a 153-FxxxM-157 motif in the interdomain helix, which is common throughout plant and bacterial COMT-type enzymes (Figure 2.9). In RedM this motif moves to within 4 Å of the methylsulfur moiety of SAM, displacing solvent and likely contacting the indolocarbazole substrate forming stabilizing edge on 𝜋-stacking and 𝜋-sulfur interactions. The opposite face of the substrate is supported by residues 297-MxxxM-301 in RedM. This motif is less widespread, found in 9.1 % of sequences in the RedM SSN subcluster; however, generalizing the position 297 to residues that interact favourably with aromatic substrates ([FYML]xxxM) covers 58% of sequences (Figure 2.11C). These four positions are the active site cavity facing residues of their respective 𝛼-helices near the SAM cofactor and are necessarily key interaction points to orient 54 substrates for methyl transfer in the typical COMT-type fold. Sequence conservation in 153-FxxxM-157 suggests the motif is a common strategy, whereas positions 297 and 301 may be more substrate specific. Overall, no clear residue conservation among N-methyltransferases indicating a preference for nitrogen atoms over oxygen atoms was observed. Table 2.3 COMT-type family enzymes capable of furnishing at least two methyltransfers to a substrate. Origin Name UniProt Involved in the biosynthesis of: Methylated moiety PDB bacteria RedM A0A0F7G196 reductasporine aminoalkane 8TJJ OxyT135,136 Q3S8P6 oxytetracyclin aminoalkane None RosA105 K4RFM2 roseoflavin aniline 4D7K LaPhzM106 A0A172J1V3 myxin phenol 6C5B BenF138 B1GSM8 benastatin anthrone none plant TaOMT2 a,139 Q38J50 tricetin phenol none CrOMT2122 Q8GSN1 syringetin phenol none ROMT121 B6VJS4 resveratrol phenol none AMNT123 Q96565 gramine aminoalkane none BANMT a,124 Q84N56 𝛽-alanine betaine 𝛽-amine none a BANMT and TaOMT2 are able to transfer three methyl groups to their base substrates. While the structure of RedM closely matches that of the COMT-type enzyme family it is missing the characteristic catalytic histidine found in most other members of the family, instead having a serine at this position (S253). Mutation of this serine to alanine or cysteine does not impact catalytic activity, while mutation to histidine eliminates it. Thus, S253 does not play an active role in catalysis, but productive catalysis is sensitive to the steric bulk or side chain charge at this position. The nearby N254 is shown in docking to be in close proximity to the pyrroline nitrogen of the indolocarbazole substrate and may play a role in substrate recognition and orientation for methyl transfer. This is a common proposal among other small molecule methyltransferases with asparagine or aspartate at this position.110,115,114,107,95 55 Mutational studies in COMT-type C-methyltransferase SibL, O-methyltransferase DnrK and S-methyltransferase DsyB indicate a tyrosine in the two 𝛼-helices capping the active site between the N- and C-terminal domains affects catalytic activity. In SibL, Y134F eliminates catalytic activity and the residue is proposed to deprotonate the methyl acceptor carbon atom after methylation and reforming an aromatic ring.116 In DnrK, the Y142W mutation reduces catalytic activity by 52 %,115 while in DsyB, the Y129F mutation reduces catalytic activity by 60 %.117 Each RedM structure has a disordered region around residues 133-149, which includes the analogous position (Y140). The Y140F variant has greatly reduced reductasporine accumulation, suggesting the region participates in completion of the active site and the residue promotes turnover. This loop may be transiently stable in order to enable the replacement of SAH with SAM after the first round of methyl transfer. DnrK and DsyB are stated to be ‘entropic enzymes’ where the catalytic rate increase is attributed to desolvation and precise positioning of the substrate.93,115,117 RedM may operate under a similar schema given that the presence of tyrosine is not necessary for catalysis unlike the equivalent residue in SibL and no other active site residues are good candidates for direct acid-base chemistry. A similar entropic enzyme explanation is given for the transmethylation mechanism of bacterial N-methyltransferase PhzM, as the enzyme lacks any apparent catalytic base and the low pKa of the phenazine nitrogen is amenable to methyl transfer without extra deprotonation at neutral pH.100 Searching the literature, Swissprot database and BLAST results suggest that to date nine COMT-type family enzymes have characterized dimethylating activity (Table 2.3). Multiple sequence alignment and SSN distribution of these does not reveal any clear sequence-based indications of dimethylating activity. The addition of steric bulk to the RedM active site opposite the S-methyl of the SAM cofactor via H250W and S253H extinguished methylation activity entirely. H250W 56 may have interfered with effective SAM binding, as H250 interacts with the SAM carboxylate, despite tryptophan being tolerated at this position in several COMT-type enzymes. S253H may be too large of a residue and excludes adequate alignment of the pyrroline nitrogen with SAM. However, this strategy has been effective in altering monomethylating enzymes into dimethylating enzymes and vice-versa. For example, SxtA is a PKS iron-dependent C-methyltransferase structurally similar to MppJ, which methylates malonyl-ACP to methylmalonyl-ACP.140 Mutation of an active site isoleucine to threonine removes steric bulk from the active site and allows for the binding of the monomethylated product in a flipped orientation followed by a second methylation event. The mutation emulates the active site of the closely related dimethylating methyltransferase AprA.141 Due to the warping of the active site in order to accommodate the iron cation and subsequent altered substrate binding there is no equivalent mutation in RedM. The COMT-S enzyme, responsible for the monomethylation of various hydroxylated coumarins, was converted into a dimethylating enzyme via a single mutation N325A.108 N325 is predicted to mediate substrate recognition through hydrogen bonding and removal of that recognition element combined with the reduction of the side chain size allows for the nonselective methylation of both hydroxyl groups in the substrate esculetin. The equivalent position in RedM is T305, which as a small residue may enable multiple methyl transfers. However, the geometry of docked indolocarbazole substrate suggests that substitution of T305 with a larger residue would have little impact on substrate binding. Additionally, other dimethylating COMT-type enzymes have a variety of small (Ala, Thr, Val) and large (Phe, Tyr, Arg, Leu) residues at this position. Lastly, resveratrol O-methyltransferase (ROMT) non-selectively dimethylates resveratrol to pterostilbene, but the mutant L117F, a position far from the site of methyl transfer, is able to primarily accumulate the monomethylated pinostilbene 57 product.142 This demonstrates that subtler manipulations of the binding cavity shape can determine the extent of methylation. Characterized COMT-type family dimethyltransferases have substrates that are either achiral or highly symmetric. For example, the N-MTs RedM, OxyT135,136, RosA105, AMNT123 and BANMT124 each only require an amine inversion or C-N bond rotation for a second methyl transfer to occur. TaOMT2139, CrOMT2122 and ROMT121 operate on achiral tricetin, myricetin and resveratrol, respectively, and require a simple ring flip via C-C bond rotation in order to methylate the second phenol position. TaOMT2 is unique in that it can also methylate a third, nonequivalent hydroxyl position in tricetin. LaPhzM methylates the highly symmetrical iodinin twice and requires a 180° flip of the entire molecule before the second methyl transfer.106 Lastly, BenF of the benastatin biosynthetic gene cluster methylates an anthrone carbon position twice.138 The second methyl transfer likely occurs as the planar nucleophilic tautomer of the monomethylated intermediate forms and attacks the SAM cofactor. Observed multi-methyltransferase activity in COMT-type family enzymes seems to arise from the natural symmetry between nucleophilic positions within a substrate paired with limited or no exclusion of the nearly equivalent monomethylated intermediate from the active site. Notably, the remaining characterized N-methyltransferases each catalyze a single methyl transfer to a nitrogen atom part of a substituted amide or aromatic heterocyclic ring. These are nucleophilic positions that can only support a single methyl group without additional chemical changes. Multi-methylating activity is less tied to specific trends in sequence across methyltransferases and is more dependent on the nature of the substrate and subtle changes in the shape and surface of the active site, as seen in COMT-S, ROMT, AprA and SxtA. This is also observed in other SAM-dependent enzymes outside the COMT-type family catalyzing sequential 58 methyl transfers, such as glycine/sarcosine N-methyltransferase143 (GSMT), lysine methyltransferase SET7/9144 and rubisco large subunit methyltransferase145 (LSMT). GSMT is able to catalyze two sequential methylations to glycine to form N-dimethylglycine as part of a biosynthetic pathway to the osmoprotectant betaine.143 When comparing GSMT against the related glycine N-methyltransferase (29 % identity; RMSD = 2.1 Å, 230 C𝛼), which only catalyzes a single methyl transfer to glycine, the active site pocket has a marked smaller volume incapable of accommodating N-methylglycine in a conformation for productive methylation. Similarly, the active site mutations Y305F and Y245A in SET7/9 expands accessibility to methylated substrates shifting catalytic activity from monomethylation to dimethylation or trimethylation, respectively.144 LSMT forms a more open active site compared to the similar enzyme SET7/9, and cocrystal structures with N𝜀-methyl-L-lysine show that the N-methyl preferentially points away from the SAM cofactor leaving an unobstructed path for further methylation events.145 Furthermore, the ability of the enzyme to cycle between open and closed conformations easily would allow for the substrate and SAM to be captured in the open conformation, transmethylated upon closure of the enzyme, followed by release of SAH and replacement with SAM in the open conformation. Potentially, the singly methylated intermediate can be retained in the open active site where the necessary rotations or inversions can occur before a second methylation with freshly acquired SAM cofactor. Docking the indolocarbazole substrate to the open conformation of RedM seems to indicate a preference for binding far away from the SAM binding site near 𝛼1, 𝛼2 and the loop between 𝛽9 and 𝛽10. (Figure 2.8A) The indolocarbazole substrate may access the active site cavity from this position and be temporarily held there between methyltransfers. However, mutating the position immediately between 𝛼1 and 𝛼2 59 (T22Q) in order to disrupt any such interaction does not alter catalytic activity noticeably. In LaPhzM, this end of the active site is proposed to control substrate access to the enzyme, where the residues Y158 and F290 form a junction limiting the maximal size of a molecule able to access the active site.106 L160 and M297 in RedM form a similar gatekeeping junction where upon closure of the enzyme they contact the other forming a ‘floor’ to the active site. The N,N-dimethyltransferase RedM of the reductasporine biosynthetic pathway is the third example of a COMT-type family N-methyltransferase to have its structure characterized, and the first to be captured in open and closed conformations. Docking didemethylreductasporine to the closed conformation indicated key amino acid residues in substrate recognition. Sequence analysis shows that these residues are widely conserved as an FxxxM motif in the 𝛼-helix joining the N- and C-terminal domains and a reciprocal [FYML]xxxM motif on the opposite wall of the active site. The second motif is less strictly conserved but is preferentially found as large hydrophobic residues. Mutations to the active site indicate no residues are likely to participate directly in catalysis, suggesting RedM operates as an entropic enzyme leveraging desolvation, orientation and proximity effects to promote methyl transfer, as seen in other COMT-type family enzymes.100,115,117 Finally, it seems that dimethylating activity in COMT-type enzymes are a natural result of processing largely symmetric, planar substrates where the enzyme cannot distinguish between nearly identical positions after amine inversions, bond rotation or tautomerization. These crystal structures give insight into the final steps of reductasporine biosynthesis and highlights the importance of domain movements in catalysis for this family of enzymes. 60 2.5 Experimental Procedures 2.5.1 General methods Primers were synthesized by Integrated DNA Technologies. Q5 DNA polymerase, T4 DNA ligase, and restriction endonucleases were purchased from New England Biolabs. DNA sequencing was carried out by either NAPS Unit DNA Sequencing Facility or CMMT/BCCHR DNA Sequencing Core Facility (University the British Columbia). Nickel sepharose protein purification resin and HiLoad 26/60 Superdex 200pg size exclusion column was purchased from GE Healthcare. ISP4 and TSB media were purchased from BD. Other general reagents were purchased from Anatrace, Bio Basic Inc., Hampton Research, Sigma-Aldrich, Thermo Fisher Scientific Canada, and VWR International as necessary. HPLC analysis was carried out on an Agilent 1260 HPLC apparatus, and LC-MS analysis on an Agilent 6120 Quadruple LC/MS system in positive and negative ion mode. 2.5.2 Cloning and expression of redE and redM The gene redE was assembled from 36 oligonucleotides designed to exclude NdeI and XhoI restriction sites using the DNAWorks software.146 The gene redM was codon optimized, and cloned into pUC17 by Bio Basic Incorporated. Both redE and redM were amplified and ligated into the NdeI/XhoI sites of the pET28a plasmid. Plasmids pET28a-redE or pET28a-redM were transformed into chemically competent E. coli BL21(DE3) and grown in lysogen broth (LB; 10 g NaCl, 10 g tryptone, 5 g yeast extract per 1 L water) (37℃, 180 rpm) with kanamycin (50 𝜇g/mL) to an OD600 of 0.6 before cooling to 16 ℃ (30 min). Cultures were induced with 100 𝜇M β-D-1-thiogalactopyranoside (IPTG) and grown for a further 18 h. 61 Cells were harvested by centrifugation (4 ℃, 10 min, 3000 × g), and resuspended in 50 mM Tris (pH 8), 500 mM NaCl, 5 mM imidazole and 1.5 mM tris(2-carboxyethyl)phosphine (TCEP). Bacteria were lysed by sonication (25% amplitude, 4 s on / 8 s off cycles, 6 min), and cell debris removed via centrifugation (45 min, 15000 × g). The supernatant was loaded onto Ni-IDA affinity resin, and then eluted using a 20-500 mM imidazole step gradient in 50 mM Tris (pH 8), 500 mM NaCl, 1.5 mM TCEP. Elution fractions containing recombinant protein were filtered through a 0.22 𝜇m syringe filter and purified via size-exclusion chromatography in 20 mM Tris (pH 8), 50 mM NaCl and 1.5 mM TCEP. SDS-PAGE was used to assess purity of resulting fractions. 2.5.3 Cloning, expression of rebO and rebD, and production of chromopyrrolic acid The genes rebO and rebD were amplified from gDNA (Lechevalieria aerocolonigenes DSM 44217). These were ligated into pET28a and pET22b, respectively. E. coli BL21(DE3) cells overexpressing RebO and RebD were grown according to the previously published procedure.82 Cells were harvested by centrifugation (4℃, 10 min, 3000 × g), and resuspended in 20 mM Tris (pH 8), 500 mM NaCl, 5 mM imidazole. Bacteria were lysed by sonication (25% amplitude, cycles of 4 s on and 8 s off for 6 min), and cell debris removed via centrifugation (45 min, 15000 × g). The cleared supernatants containing RebO and RebD were added to a solution of L-Trp (100 mg, 490 𝜇mol) buffered in 25 mM HEPES (pH 8, 250 mL) and stirred for 18 h at room temperature. The reaction was quenched via the addition of HCl until pH < 2 before removing precipitate via centrifugation. The supernatant was passed through a Waters 20cc C18 Sep-Pak® cartridge and eluted by a H2O : MeOH step gradient (0-100% MeOH). The fractions containing 62 chromopyrrolic acid were combined, concentrated and purified using a semi-preparative C18 column (Phenomenex Luna C18(2), 100 Å, 250×10 mm, 5 𝜇m) 5-100% ACN in H2O. 2.5.4 Expression and purification of StaP, ferredoxin and ferredoxin reductase Plasmid pET28a-staP was kindly provided by Catherine Drennan (Massachusetts Institute of Technology), and plasmid stocks of ferredoxin-NADP oxidoreductase (FDR) and ferredoxin I (FDX) from Synechococcus elongatus PCC 7942 were received from Dr. Hai-Yan He. E. coli BL21(DE3) harbouring pET28a-staP or pET28a-fdx were grown in LB medium with kanamycin (50 𝜇g/mL) to OD600 of 0.5 before moving to 16 ℃ for 30 min. 1 mM 𝛿-aminolevulinic acid and 100 𝜇M IPTG were added to cultures before growing for a further 18 h. E. coli BL21(DE3) transformed with pET28a-fdr were grown similarly with the omission of 𝛿-aminolevulinic acid. Cells were harvested by centrifugation (4℃, 10 min, 3000 × g), and resuspended in 50 mM Tris (pH 8), 500 mM NaCl, 5 mM imidazole. Bacteria were lysed via sonication (25% amplitude, cycles of 4 s on and 8 s off for 6 min), and cleared via centrifugation (45 min, 15 000 × g). Soluble 6×His tagged StaP, FDX and FDR were purified via Ni affinity column and imidazole step gradient (5-300 mM). Dialyzed protein fractions were screened for activity against CPA before adding glycerol to 10 % (v/v) and storing at −70 ℃. 2.5.5 In vitro production of reductasporine and pathway intermediates Dicarboxyindolocarbazole was generated in situ from CPA and StaP in a tandem reaction containing 250 𝜇M CPA, 5 mM NADPH, 50 mM HEPES pH 8, 1 𝜇M StaP, 20 𝜇M FDX, 1 𝜇M FDR and 20 𝜇M RedE. Reductasporine production used the same conditions supplemented with 5 mM SAM and 20 𝜇M RedM. Reactions were quenched with two volumes of MeOH and denatured protein removed by centrifugation. HPLC assays were run on an Agilent 1260 HPLC 63 apparatus with a Phenomenex Luna C18(2), 100 Å, 250 × 4.6mm, 5 𝜇m column at 1 mL min-1 using a gradient of 2-100 % ACN in 0.1 % formic acid. Elution was monitored at 260, 280, 300 and 325 nm or by mass. Large-scale preparation of reductasporine used the same conditions as above at a total volume of 3 mL and the reaction was supplemented with extra NADPH, StaP, FDX and FDR after two and four hours. 1H-NMR (600 MHz, DMSO-d6) δ 3.53 (6H, s, H3-14, H3-14a), 5.52 (4H, s, H2-5, H2-7), 7.29 (2H, t, H-3, H-9), 7.48 (2H, t, H-2, H-10), 7.78 (2H, d, H-1, H-11), 7.96 (2H, d, H-4, H-8), 11.43 (2H, brs, NH-12, NH-13) 2.5.6 Synthesis of arcyriarubin A Based on procedures published by Faul et al. 1995 and Mei et al. 2017.147,148 To a N2 purged suspension of 52 mg magnesium (2.1 mmol) in 2 mL THF, 170 𝜇L of ethyl bromide (2.3 mmol) was added and stirred at 40 ℃ for 1 h. Indole solution (260 mg, 2.2 mmol, 2 mL toluene) was added dropwise before heating at 60 ℃ for 1 h. Dibromomaleimide (78 mg, 0.31 mmol, 1 mL THF) was added slowly before refluxing for 15 h. Quench proceeded with 1 M HCl and was partitioned into ethyl acetate and water. Organic crude was washed with water and brine and the aqueous wash was extracted with ethyl acetate. The combined organic phase was dried with sodium sulphate and solvent removed. The crude was purified via silica column (1:1 hexanes:ethyl acetate), yielding 50 mg of arcyriarubin A as a red oil (50 % yield). 2.5.7 Synthesis of arcyriaflavin A Based on the procedure published by Reddy et al. 2003.149 Iodine (8 mg, 32 𝜇mol) was added to an arcyriarubin A solution (50 mg, 150 𝜇mol, 24 mL 1:1 THF:ACN), and stirred under a mercury lamp (400 W) for 23 hours. Solvent was removed and dissolved in 2 mL THF before 64 precipitating product with 15-20 mL hexane. The filtered solids were dried under vacuum and used without further purification (45 mg, 91 % yield). 2.5.8 Synthesis of didemethylreductasporine Based on previously published amide,150 lactam151 and isoindoline-1,3-dione reductions.152,153 A suspension of NaBH4 in THF (28 mg, 0.5 mL) was purged with N2, before adding arcyriaflavin A (10 mg, 31 𝜇mol, 1 mL THF). The solution was cooled to 0 ℃ and 250 𝜇L of a 370 mM iodine solution in THF was added dropwise over 10 min. The reaction was stirred for 2.5 h before bringing it to 65 ℃ for 16 h. Quench proceeded with 400 𝜇L methanol followed by 1 M HCl and partitioning between ethyl acetate and water before washing with 1 M NaOH and brine. Semi-preparative HPLC at 30 % ACN yields didemethylreductasporine (5 mg, 50 % yield) with K252c identified as the major side product alongside unreacted arcyriaflavin A. 2.5.9 Enzymatic methylation of didemethylreductasporine RedM activity was assayed independently from RedE by including 2 𝜇M RedM, 2 mM SAM, 500 𝜇M didemethylreductasporine, 50 mM HEPES pH 8 in a reaction mixture. Aliquots of the reaction mixture were taken at 5, 10 15, 20 and 25 minutes and quenched with two volumes of acetonitrile. Reaction products were measured via HPLC on a Phenomenex Luna C18(2), 100 Å, 250 × 4.6mm, 5 𝜇m column at 0.8 mL min-1 using a gradient of 5-90 % ACN in 0.1 % formic acid over 11 min. Elution was monitored by UV-Vis absorbance at 325 nm and by mass. 2.5.10 Selenomethionine incorporation into RedM Plasmid pET28a(+)-redM was transformed into E. coli BL21(DE3) expression hosts and plated on kanamycin supplemented LB agar for overnight growth. Single colonies were grown for 8 h in 5 mL LB with kanamycin before inoculating 30 µL into 30 mL of M9 minimal 65 medium and grown overnight. This starter culture (10 mL) was added to 1 L of M9 medium with kanamycin and grown to an OD600 of 0.7 AU before cooling to 16 ℃. L-configured lysine, threonine and phenylalanine (100 mg each), and leucine, isoleucine, valine and selenomethionine (50 mg each) were combined and added to each 1 L culture.154 After 25 min of mixing, cultures were induced with 100 µM IPTG and grown for 16-18 h. Selenomethionine RedM derivatives were purified as described previously with buffers freshly supplemented with 5 mM β-mercaptoethanol. 2.5.11 Crystallization of RedM Initial conditions amenable to RedM crystallization were determined via commercially available trial screens (Index, Hampton, MCSG) in a 96-well plate, sitting-drop format (1 µL protein : 1 µL crystallization solution adjacent to 50 µL of mother liquor). In 0.2 M (NH4)2SO4, 0.1 M MES pH 6.5, 30% w/v polyethylene glycol monomethylether 5000, RedM at 7 mg/mL crystallized as plates within 3 weeks or in 0.05 M (NH4)2SO4, 0.05 M BisTris-HCl pH 6.5, pentaerythritol ethoxylate (15/4_EO/OH) at 17 mg/mL RedM as prisms. Initial hits were optimized in hanging-drop screens set up in 24-well plates with drops of 1-2 µL of protein (13-30 mg/mL) and 1-2 µL crystallization solution. Optimal growth of RedM selenomethionine derivative crystals was found to be 0.3 M (NH4)2SO4, 0.1 M Tris pH 8.0, 30% PEG MME 5000 at 12.5 mg/mL protein. Diffraction quality native RedM crystals grew under 0.2-0.3 M (NH4)2SO4, 0.1 M MES pH 6.5, 18-24 % polyethyleneglycol monomethyl ether 5000 at 13-16 mg/mL RedM, as well as 0.09 M (NH4)2SO4, 0.05 M BisTris-HCl pH 6.5, 30 % pentaerythritol ethoxylate (15/4_EO/OH) at 30 mg/mL RedM. 66 The RedM-SAH crystal was retrieved from a pentaerythritol ethoxylate condition before soaking for 3 h in freshly made well solution supplemented with 1 mM S-adenosylhomocysteine. This crystal was moved to a cryoprotectant solution consisting of the well solution made with 10 % (v/v) ethylene glycol for less than 30 s before flash freezing under a gaseous stream of nitrogen held at 100 K. The RedM-SAM crystal originated from a polyethylene glycol condition and was soaked in freshly mixed well solution supplemented with 10 mM S-adenosylmethionine for 5 minutes. apo-RedM and RedM-SAM crystals were moved into cryoprotectant consisting of the mother liquor and 20 % (v/v) ethylene glycol for less than 30 s, before flash-freezing in nitrogen. These crystals were tested locally for diffraction using a Rigaku MicroMaxTM-007HF (1.54 Å, rotating copper anode) with Saturn 944+ CCD detector before storing for further data collection at a synchrotron facility. 2.5.12 Data collection, structure determination and refinement Datasets for RedM were collected at the Canadian Light Source (CLS, Saskatoon, Canada) on beamline 08B1-1 (Saskatoon, Canada) using a Rayonix MX300-HE detector, and at the Stanford Synchrotron Radiation Lightsource (SSRL, Menlo Park, United States) on beamline BL9-2 with a Pilatus 6M PAD detector. Diffraction images were processed and integrated using iMOSFLM155,156 or XDS.157 Data were scaled and merged in AIMLESS158, and phases determined by single anomalous dispersion of selenium atoms or via molecular replacement of the partially solved selenomethionine derived RedM model using the PHENIX software package.159 An initial model was created and refined using the PHENIX Autobuild software.160 This initial model was improved upon through cycles of manual modelling with Coot161 and refinement with REFMAC162 and phenix.refine.163 67 2.5.13 Substrate Docking Molecular docking of didemethylreductasporine, demethylreductasporine and reductasporine with open and closed conformations of RedM was performed using AutoDock VINA.164 Coordinates of K252c were retrieved from the Protein Data Bank (PDB ligand code: K2C) and modified in PyMOL165 to each reductasporine derivative before optimizing geometries using eLBOW.166 RedM protein models bound with SAM and SAH were prepared for docking by removing water, adding exchangeable hydrogen atoms and designating flexible residues via MGLTools for AutoDock. Docking was bounded to a 40 × 40 × 40 Å box centered in the cavity adjacent to SAH or SAM and contained the majority of the interior surface of both open and closed conformations of the RedM monomer. The exhaustiveness parameter was set to 24 after no noticeable changes to docking outcomes were observed at higher values. Analysis and visualization of docking results were done using PyMOL.165 2.5.14 Site directed mutagenesis of RedM Primers were designed with an overlapping region containing the mutation and 5ʹ non-overlapping arms. Overlapping areas were chosen with melting temperatures approximately 55 ℃ and non-overlapping regions had melting temperatures about 7 ℃ higher (calculated against the complementary DNA strand).167 Standard PCR was carried out using the pET28a(+)-redM plasmid with an annealing step set to the higher melting temperature and ended with a 10 min elongation step. The PCR product mixture was incubated with DpnI before transforming the digest solution as described previously into E. coli DH5𝛼. Mutations were confirmed by sequencing. 68 2.5.15 Visualization and analysis software Crystal structures are visualized using Pymol.165 Estimates of internal cavity volume were generated with POCASA (grid size = 0.5 Å, probe radius = 3 Å, single point flag = 11, protein depth flag = 10).101 Surface areas and dimer interaction surfaces were calculated via PISA.90 Bending residues and hinge axes were found through DynDom.168 Protein sequence alignments were generated using Clustal Omega169 or COBALT118 and visualized with Jalview.170 2.5.16 Sequence similarity networks Initial SSN was created using the EFI-EST service171 the two annotated dimerization domains (plant methyltransferase dimerization, IPR012967; acetylserotonin O-methyltransferase dimerization, IPR031725); however, the network covered approximately half of the known structural homologues to RedM. The query was expanded to include the associated methyltransferase domain (IPR001077), while reducing the granularity of sequence space by using the UniRef50 database. The resulting network was missing only two RedM structural homologues from the 41 represented in the PDB (MppJ and Rv2258c). Edges were removed incrementally until distinct clusters formed between enzymes from plants, animals/fungus and bacteria (an alignment score > 54). All sequences from the cluster containing RedM were retrieved and reanalyzed via EFI-EST for a more granular SSN of related enzymes. The EFI-GNT tool was used to look up local gene arrangements in order to identify candidate biosynthetic gene clusters as an indication of isofunctionality within a cluster. 69 Chapter 3: An imine reductase that captures reactive intermediates in the biosynthesis of the indolocarbazole reductasporine 3.1 Synopsis Imine reductases (IREDs) and reductive aminases (RedAms) have been used in the synthesis of chiral amine products for drug manufacturing; however, little is known about their biological contexts. Here I employ structural studies and site-directed mutagenesis to interrogate the mechanism of the IRED RedE from the biosynthetic pathway to the indolocarbazole natural product reductasporine. Co-crystal structures with the substrate-mimic arcyriaflavin A reveals an extended active site cleft capable of binding two indolocarbazole molecules. Analysis of the structure and site-directed mutagenesis of a conserved aspartate in the primary binding site reveals a new role for this residue in anchoring the indolocarbazole substrate above the NADPH cofactor. Additionally, variants targeting the secondary binding site greatly reduce catalytic efficiency. As isoindole compounds are susceptible to spontaneous oxidation, I propose the secondary site acts to protect against autooxidation and the primary site drives catalysis through precise substrate orientation and desolvation effects. The structure of RedE with its extended active site can be the starting point as a new scaffold for engineering IREDs and RedAms to intercept large substrates relevant to industrial applications. 3.2 Introduction Imine reductases172,173 have drawn considerable interest for their application as biocatalysts in the synthesis of pharmaceuticals.174–177 IREDs can be used in chiral amine syntheses starting from imines178,179 or amine-ketone pairs.180 They are valued for their ability to create secondary and tertiary chiral amines, as well as catalyzing reductive amination.177,181 Recently, two 70 reductive aminases have been engineered to produce industrially useful amounts of drug candidate.182,183 However, with the focus on industrial applications and engineering of IREDs, the study of these enzymes in their natural context has been neglected. RedE, an IRED, serves as a biosynthetic enzyme in the formation of the indolocarbazole natural product reductasporine.184 Indolocarbazoles have been a rich source of potential chemotherapeutics with many entering clinical trials for activity against cancers,20 and the semi-synthetic staurosporine derivative, midostaurin, was approved in 2017 as an anticancer treatment.185 In the field of biosynthesis, a number of studies have highlighted the shared and divergent features of indolocarbazole assembly, with initial biosynthetic steps catalyzed by a core set of oxidative enzymes, followed by the action of diverse tailoring enzymes.19 Each indolocarbazole gene cluster preserves the three genes coding for a FAD-dependent tryptophan oxidase (redO, rebO, staO), a heme-containing chromopyrrolic acid synthase (redD, rebD, staD) and a cytochrome P450 indolocarbazole synthase (redP, rebP, staP) (Figure 3.1). The tryptophan oxidase first converts two L-tryptophan molecules to the corresponding enamines before chromopyrrolic acid synthase couples the two at the C𝛽 position.43 The indole-3-pyruvate imine dimer spontaneously cyclizes into CPA.26,36 The indolocarbazole synthase oxidatively bridges the indole C2 positions yielding a proposed dicarboxyindolocarbazole intermediate (4).30,186 This indolocarbazole is prone to decarboxylation and oxidation, and has not yet been directly detected. Several indolocarbazole biosynthetic gene clusters encode unique enzymes that capture and stabilize an indolocarbazole decomposition product before promoting the formation of a more stable compound for further tailoring.29,44–47 For example, the rebeccamycin biosynthetic enzyme RebC and the homologous staurosporine biosynthetic enzyme StaC capture reactive intermediates and funnel them toward the 8- and 4-electron oxidation products, 71 respectively (Figure 3.1B). The trapping of distinct tautomers of 7-carboxy-K252c in crystals of RebC and in RebC-10x, an enzyme with StaC-like activity, suggest that careful control of the hybridization at the C7 position can enable simple decarboxylation in the keto tautomer to staurosporine aglycone K252c (in the case of StaC) or decarboxylation after hydroxylation of the enol tautomer to arcyriaflavin A (in the case of RebC).45,46 A B Figure 3.1 Shared steps and examples of indolocarbazole biosynthesis. A, The conserved catalytic steps to produce chromopyrrolic acid. B, Formation of the indolocarbazole core from CPA and the proposed tailoring steps catalyzed by RebC (rebeccamycin biosynthesis), StaC (staurosporine biosynthesis) and RedE (reductasporine biosynthesis). NH𝛽NHOHO HN𝛽ʹNHOOHNH2NHO OHindole-3-pyruvic acid enamineL-tryptophan chromopyrrolic acid (CPA)StaDRebDRedDStaORebORedONH3 NHNHHNOHOHOONH2NHO OHspont.NHNHHNOHOHOONHNHHN OHOONHNHHN OHHOONHNHHNO ONHNHHN ONHNHHNRebC StaCRebC-10xNHNHHNNHNHN+RedERedMchromopyrrolic acid (CPA)StaPRebPRedPNHNHHNOHOHOOReductasporine (1)234Indolocarbazole core7-carboxy-K252c (keto tautomer)(enol tautomer)Arcyriaflavin A K252c72 The reductasporine gene cluster was discovered while screening the metagenomes of soil samples for chromopyrrolic acid synthase-like genes.184 Chang et al. reconstituted this pathway in E. coli and proposed RedE reduces the 5,7-didehydro-N,N-didemethylreductasporine (3) to didemethylreductasporine (2) (Figure 3.2). This proposal is based on the propensity of the dicarboxyindolocarbazole intermediate 4 to spontaneously decarboxylate at physiological conditions, although 3 was not detected directly.30,184 Recently, our group reconstituted reductasporine biosynthesis in vitro, and our biochemical results confirm the pathway proposed by Chang et al. In analogy to the role of the StaC/RebC enzymes, RedE captures a reactive intermediate from the indolocarbazole biosynthetic pathway; however, it catalyzes a reduction rather than an oxidation. Based on these biochemical data, RedE would have to intercept an intermediate while not allowing for oxidation, indicating a novel mechanism. Figure 3.2 Proposed biosynthetic pathway to reductasporine (1). RedE acts to intercept spontaneous decarboxylation products after cytochrome P450, StaP, catalyzes the bridging of the C2 positions from each indole in CPA. The simplest substrate for RedE is 5,7-didehydro-N,N-didemethylreductasporine (3), which is proposed to be reduced to didemethylreductasporine (2) before methylation to 1 via SAM-dependent methyltransferase, RedM. Here, I present crystal structures of RedE, including apo, NADP+-bound and two ternary complexes with arcyriaflavin A (AFA) and NADP(H). These crystal structures reveal an unexpected secondary indolocarbazole binding site adjoined to the primary active site. Our site-directed mutagenesis of residues at each site then shows both sites are critical for catalysis. Our NHNHHNNHNHHNNHNHHNOHOOOHNHNHN-2 × CO2 RedE RedMCPA 2 13NHNHHNOHOOOH StaP473 biochemical characterization of StaP reaction products reveals the accumulation and decomposition of 3, and I propose that RedE intercepts and reduces 3 from solution. The secondary active site may protect the indolocarbazole from oxidation and select for the isoindole imine tautomer before direct transfer to the primary site for reduction. Close inspection of the secondary indolocarbazole binding site and comparison to other structurally characterized IREDs suggest potential sequence determinants that would favour the formation of this cleft, indicating wider implications on the malleability of IRED active sites. 3.3 Results 3.3.1 RedE dataset phasing, twinning and initial model building Twinning was detected in selenomethionine-substituted RedE and apo-RedE crystal datasets. The second moment of intensity (⟨I2⟩/⟨I⟩2) reaching 1.78 (untwinned: 2.0, perfect twin: 1.5) and ⟨|L|⟩, 0.43 (untwinned: 0.5, perfect twin: 0.375). The L-test for twinning identifies a twin fraction as high as 23 %. When a monoclinic unit cell fulfills any condition 𝛽 ≈ 90°, a ≈ c, or 2c･cos 𝛽 ≈ −a it is possible to form a pseudomerohedral twin.187–189 The cell dimensions of the P 21 space group crystals of SeMet-RedE and apo-RedE meet the third condition, and a model can be solved for using the appropriate twin law (h, −k, −h−l). The resulting SeMet-RedE model arranges as a pair of dimers in the asymmetric unit. In several chains, 𝛼11 (residues 256-278) has too poor electron density to model completely, and another 16 residues in several loop regions are also missing (Figure 3.3). Nonetheless, this model is sufficient to solve for phases via molecular replacement in other RedE diffraction datasets. RedE co-crystals with NADP+ and AFA were not affected by twinning. 74 Figure 3.3 Asymmetric unit of the selenomethionine-substituted RedE structure. The model is missing residues in several loops (A24, G25, K127-Y130, P194-N197, Y230-F235) and 𝛼11 (residues 256-278) due to poor electron density. 75 Table 3.1 Crystallographic data collection and refinement statistics.a apo-RedE RedE-NADP+ RedE-AFA-NADP+ RedE-AFA PDB Accession 8U04 8U05 8U06 8U07 Wavelength (Å) 0.9796 0.9796 0.9796 0.9796 Resolution (Å) 93.36-1.62 (1.65-1.62) 68.88-1.62 (1.65-1.62) 37.75-1.60 (1.63-1.60) 37.94-1.63 (1.67-1.63) Space group P 21 P 21 P 21 P 21 Cell dimensions (Å) 51.78 186.72 71.69 𝛽 = 111.12° 44.74 111.31 72.04 𝛽 = 107.04° 54.11 110.77 66.99 𝛽 = 107.6° 54.59 111.53 67.69 𝛽 = 108.47° Rmerge 0.095 (1.425) 0.080 (0.414) 0.073 (0.887) 0.071 (0.665) Rmeas 0.103 (1.546) 0.087 (0.449) 0.078 (0.957) 0.077 (0.722) Rpim 0.040 (0.593) 0.034 (0.172) 0.029 (0.355) 0.030 (0.276) Total number of reflections 999547 (50013) 493577 (27473) 677620 (32657) 644294 (34386) Total number of unique reflections 154438 (7659) 75988 (4141) 95462 (4628) 93667 (5126) ⟨I/𝜎(I)⟩ 11.9 (1.6) 12.9 (4.2) 14.4 (2.2) 12.8 (2.3) CC1/2 0.998 (0.448) 0.996 (0.945) 0.999 (0.845) 0.998 (0.948) Completeness 96.3 (97.6) 88.9 (97.9) 96.6 (95.1) 98.7 (97.9) Multiplicity 6.5 (6.5) 6.5 (6.6) 7.1 (7.1) 6.9 (6.7) Wilson B-factor 20.26 16.61 17.91 22.98 Reflections used in refinement 154374 (7727) 75899 (2967) 95375 (6731) 93448 (7688) Reflections used for Rfree 7991 (408) 3872 (151) 2000 (141) 1708 (141) Rwork 0.1633 (0.2777) 0.1715 (0.2139) 0.1553 (0.2574) 0.1929 (0.3384) Rfree 0.2076 (0.2817) 0.2050 (0.2792) 0.1889 (0.2682) 0.2191 (0.3900) Number of non-hydrogen atoms 9463 5005 5191 4894 macromolecules 8280 4160 4275 4180 ligands 33 140 154 128 solvent 1150 705 762 586 Protein residues 1152 578 581 566 RMS (bonds) 0.002 0.005 0.01 0.005 RMS (angles) 0.52 0.77 1.08 0.71 Ramachandran favored (%) 97.63 98.26 98.09 98.03 Ramachandran allowed (%) 2.28 1.74 1.91 1.97 Ramachandran outliers (%) 0.09 0 0 0 Rotamer outliers (%) 0.62 0.5 0.48 0.25 Clashscore 3.43 1.54 2.43 2.4 Average B-factor 23.46 22.89 26.33 34.41 macromolecules 21.1 20.45 23.77 32.36 ligands 36.51 28.54 28.48 43.59 solvent 40.07 36.17 40.27 47.01 Number of TLS groups 14 8 6 6 a Data in parentheses are for the high-resolution shell. 76 3.3.2 Structure of RedE I solved the apo, binary complex and two ternary complex structures of RedE with NADP+ and AFA to resolutions of 1.60-1.63 Å (Table 3.1, Figure 3.4). RedE resembles the canonical IRED fold with an N-terminal NADPH-binding Rossmann domain and C-terminal helical bundle mediating homodimerization.179–181,190 The N-terminal Rossmann domain (residues 2-161) consists of an eight-stranded 𝛽-sheet with the first six strands (𝛽1-6) running parallel and the following two strands (𝛽7-8) antiparallel. Each parallel strand is bounded by an 𝛼 helix (𝛼1-7) forming 𝛽𝛼𝛽 motifs and a 310-helix forms in the loop between the antiparallel 𝛽6 and 𝛽7 strands (Figure 3.4A). The common Rossmann domain GxGxxG nucleotide binding motif is preserved at residues 9-14 (GLGPMG).191 The C-terminal domain (residues 193-288) is a five-membered helical bundle where the latter four 𝛼-helices (𝛼9-12) wrap around the interdomain helix of its dimer partner (𝛼8ʹ) forming an extensive 4020 Å2 interface (27 % of total monomer surface area) between both N- and C-terminal domains as calculated by PISA.90 A total of 22 and 79 residues are involved in polar and hydrophobic contacts, respectively (Figure 3.4B). The majority of these interactions are within the dimerization domains with only 17 residues interfacing between N- and C-terminal domains. Hydrogen bonding and electrostatic interactions occur between E153 to Rʹ209, D160 to Gʹ195, and T121 and S123 to Eʹ234. The N-terminal end of 𝛼6 is associated with an iodide ion which is in turn stabilized by Rʹ254. The majority of the hydrophobic contacts between the two domains are found between 𝛽6-8 and 𝛼ʹ9 where M119, Y130, L131, L133, M155 and L157 interlock with Vʹ206, Rʹ209, Wʹ210 and Tʹ213. Lastly, residues at the N-terminal ends of 𝛼6 and the interdomain helix (P96, V97, A162 and M163) stack against the loop between the C-terminal end of the dimer partner’s interdomain helix (𝛼ʹ8) and 𝛼ʹ9, specifically Pʹ195 and Gʹ196. 77 Figure 3.4 Cartoon representations of the overall RedE structure. A, Protomer with highlighted secondary structures. A 310-helix forms between 𝛽6 and 𝛽7. B, Polar contacts between protomers of the RedE dimer. Asymmetric units of the C, RedE-NADP+ binary complex, D, RedE-AFA complex copurified with NADP(H) with alternate location of arcyriaflavin A in the primary binding site (grey), and E, RedE-AFA-NADP+ ternary complex. Ligand molecules shown as ball-and-stick models in black, and iodide ions as purple spheres. Datasets from crystals of RedE grown with 10 mM NADP+ produce Fo–Fc electron density omit maps that can be unambiguously modelled as NADP+ (Figure 3.5A). They bind broad clefts formed from the confluence of 𝛼1, 𝛼2 and 𝛽1-5 with the characteristic GxGxxG NADPH 78 binding sequence found at the centre between 𝛽1 and 𝛼1 (9-GLGPMG-14) with hydrogen bonds forming between the C5ʹ-phosphate and M13 amide N-H and an ordered water held between the amide nitrogen atoms of G11 and G14 and C65 amide carbonyl (Figure 3.6A). The adenosine C2ʹ-phosphate is held in place by a salt bridge with R33 alongside hydrogen bonds with Q32, T34 and K37. The adenine ring rests against the A71 side chain and is parallel stacked with the R33 guanidino group. E74 hydrogen bonds to the adenine C6-amine. The backbone N-H of A67 hydrogen bonds with the adenosine furan oxygen, while the S83 side chain and L66 backbone carbonyl and interact with the C2ʹ- and C3ʹ-hydroxyl of the nicotinamide riboside, respectively. The re-face of the nicotinamide C4 is positioned overtop the side chain of M13 with the si-face facing into a deep pocket formed at the dimer interface of the C-terminal helical bundle. Comparison of the apo and binary structures show that R33 moves out of the adenine binding area in order to stack against the purine ring while forming a salt bridge to the nearby 2ʹ-phosphate. As the substrate and products of RedE are transiently stable, I elected to employ AFA as a mimic, because it shares the planar indolocarbazole core of 3. Processing diffraction data of RedE crystallized from solutions with 1.2 mM AFA and 10 mM NADP+ show electron density consistent with both AFA and NADP+ in each protomer (Figure 3.5A,B). In this structure, the positioning of NADP+ is similar to that observed in the binary complex. AFA sits above the si-face of the nicotinamide C4 with the maleimide carbonyl situated 3 Å away in a geometry ideal for hydride transfer. AFA is enveloped by a highly hydrophobic pocket lined with M119, T121, L172, M175, W210ʹ, F217ʹ, F235ʹ, L239ʹ, T243ʹ and I246ʹ, and anchored within it via interactions to D168 with each indole nitrogen (Figure 3.6C). Lastly, the guanidine group of R242 hydrogen bonds to the maleimide carbonyl. 79 Figure 3.5 Ligand mFo-DFc omit maps contoured at 3𝜎. A, NADP+ cofactor, B, primary site arcyriaflavin A (AFA) and C, secondary site AFA. NADP(H) cofactor co-purified with the RedE-AFA co-crystal is modelled as NADP+. 80 Figure 3.6 Key ligand interactions in the in the extended RedE active site. A, The NADPH binding cleft of RedE with bound NADP+. The conserved GxGxxG Rossmann domain binding motif is coloured pale green (9-GLGPMG-14). B, The secondary indolocarbazole binding site. Residues from the neighbouring protomer are coloured orange. C, The primary indolocarbazole binding site occupied by NADP+ and arcyriaflavin A. The maleimide carbonyl is positioned 3 Å above the si-face of the nicotinamide ring in an ideal geometry for hydride transfer. Residues from the neighbouring protomer are coloured cyan. Hydrogen bonds are indicated by dashed lines with distances labelled in Angstrom. In structures solved from RedE crystals formed in a solution with only 1.2 mM AFA, there is electron density consistent with AFA binding the cleft adjoined to the hydrophobic active 81 site pocket (Figure 3.5C). This secondary binding cleft forms between 𝛽5, 𝛼6, and 𝛼8 of the Rossmann domain and 𝛼ʹ10 from the C-terminal helical bundle. AFA forms hydrogen bonds from each indole N-H to T169 and a water held between the S165 carbonyl and the D168 sidechain (Figure 3.6B). In one protomer H249 may form a weak hydrogen bond to the nearby maleimide carbonyl (3.5 Å). AFA makes hydrophobic contacts with G94, S95, P96, I246, A250 and L253, and the G94 peptide carbonyl is held in place with a hydrogen bond to the S165 side chain. An iodide ion bounded by the positive dipole N-terminal end of 𝛼4 and R254 completes the outer boundary of the cleft. Despite not including NADP+ in the preparation of these crystals, electron density is consistent with the presence of NADP(H) cofactor retained during purification. The electron density has poorer definition than that of the RedE-NADP+ or RedE-NADP+-AFA structures and can be modelled by partial occupancy of a NADP+ cofactor (Figure 3.5A). Additionally, there is sparse electron density above the nicotinamide moiety resembling partial occupancy of AFA in the same pose as characterized in the RedE-NADP+-AFA ternary structure (Figure 3.5B). The density is a poor fit and modelling either AFA or water in this area produces similar Rfree values. Furthermore, placing an AFA molecule in the active site clashes with the clearly defined position of the AFA molecule in the secondary binding cleft. Placing both indolocarbazoles in the active site simultaneously forces a contact between indole rings of less than 2 Å, which is too small to represent simultaneous binding. Modelling this relation as alternate conformations results in occupancy of 55-60 % in the distal cleft. This feature may result from heterogeneity in binding configurations throughout the crystal. 82 3.3.3 Resemblance to other NAD(P)H-dependent reductases Secondary structure searching via PDBeFOLD169 shows RedE’s closest structural matches are other imine reductases and reductive aminases (Figure 3.7). These are defined by the reciprocal domain swapping of each monomer’s C-terminal helical bundle. However, compared to all other imine reductases and reductive aminases RedE has a pronounced crescent shape imparted by the secondary indolocarbazole binding cleft formed between 𝛼ʹ10 of the dimerization domain and 𝛼6 of the Rossmann domain. In other IREDs this is a tighter binding interface between protomers yet buried surface area calculations indicate similar contact areas between 3900 and 4600 Å2 (4020 Å2 in RedE). The loss of close contacts between 𝛼ʹ10 and 𝛼6 in RedE are supplemented with dimer interactions found at the loop between 𝛼8 and 𝛼9 set against the N-terminal ends of 𝛼ʹ6 and 𝛼ʹ8 of its dimer partner in addition to closer interactions between the 𝛽-sheet in the Rossmann domain and 𝛼ʹ9 in the dimerization domain. This extra surface area may provide support to maintain the separation between the domains. Figure 3.7 Ninety degree separated views of representative dehydrogenases structurally related to RedE. 6-PGDH is shown as a monomer with the last 𝛼-helix involved in dimerization hidden and the C-terminal extension mimicking the 𝛽HAD fold homodimer interface coloured grey. HIBADH; 3pdu IRED; 3zhb RedE PYCR; 5bse PDH; 7mqv90°90°6-PGDH; 7cb083 The next closest structural matches are from the 𝛽-hydroxy acid dehydrogenase (𝛽HAD) family, which include 3-hydroxyisobutyrate dehydrogenase192 and L-serine dehydrogenase193 among several other characterized enzymes (Figure 3.7).194,195 These share an N-terminal Rossmann NAD(P)H binding domain and C-terminal helical bundle mediating dimerization; however, a sharp turn in the helix following the interdomain helix (𝛼9 in RedE) redirects the chain back towards the Rossmann domain resulting in no reciprocal domain swapping.192 Despite this large deviation, the backbone traces of the overall quaternary structure of the dimer is maintained between IREDs and 𝛽HADs. The related 6-phosphogluconate dehydrogenases (6-PGDH) shares a similar fold to 𝛽HADs, where the helical bundle of a 6-PGDH protomer has a circa 130 amino acid C-terminal extension which replicates the fold of an interacting 𝛽HAD dimer.196 RedE has sparing, but notable, structural similarity with pyrroline-5-carboxylate reductases197 and prephenate dehydrogenases (Figure 3.7).198 These enzymes share a similar layout to IREDs and 𝛽HADs with an N-terminal Rossmann binding domain and C-terminal helical bundle mediating dimerization. They even share a similar presence or lack of reciprocal domain swapping resulting from a sharp turn in an alpha helix in the dimerization domain. However, the organization of the helical dimerization bundle is substantially different with several helices following the long axis of the enzyme rather than being arranged perpendicularly to it. 84 3.3.4 Domain contacts among IRED and RedAm enzymes. Of the 16 unique IRED and RedAm hits from secondary structure matching using the RedE protomer, the median RMSD of C𝛼 atoms is 2.76 Å covering 93 % of residues in RedE (Table 3.2). Among these enzymes RedE is the only one to have an extended loop between 𝛼8 and 𝛼9 which interacts with the Rossmann domain of its partner monomer (Figure 3.8A). Searching the PDB for all IREDs and RedAms shows that this is a unique feature to RedE and is an apparent insertion of four residues (194-PGGN-197). AtRedAm has a two residue insertion in this loop; however, the area does not interact with the neighbouring Rossmann domain.199 Extending the search to primary sequence matches in the UniProt database shows that this ‘PGGN’ loop is found in ten other IRED-like enzymes, though the insertion may be more accurately described as residues 193-GPGG-196 (Figure 3.8B). This loop interacts closely with P96, V97, D160, A162 and M163 in the Rossmann domain where P96 and D160 are conserved positions among structurally characterized IREDs (Figure 3.9). Among more closely related sequences to RedE, the V97 position is found as a hydrophobic residue when the ‘GPGG’ insertion is present and as a glutamate when it is not. AlphaFold predicts these loops adopt a similar conformation to RedE and each predicted assembly possess a cleft between 𝛼6 and 𝛼ʹ10.200 The sequence alignment of helix 𝛼10 indicates no residues are strongly conserved among all IREDs, while the loop spanning 𝛽5 and 𝛼6 is often conserved as [ST][ST]x[ST]P (Figure 3.9). Among structurally characterized IREDs and RedAms a histidine or asparagine is often found at I246 in RedE and is involved in cross-domain contacts (Figure 3.8B). This position is exclusively found as a hydrophobic residue in other IRED-like sequences with a ‘GPGG’ insertion between 𝛼8 and 𝛼9. 85 Table 3.2 Sixteen IRED and RedAm hits from secondary structure matching of RedE using PDBeFOLD. Representative PDB ID UniProt ID RMSD (Å) Percent aligned C𝛼 Percent identity 5A9S R4SNK4 2.00 91.7% 43.7% 5OCM D2B7Z8 2.15 95.7% 51.1% 3ZHB Q1EQE0 2.17 94.4% 46.8% 3L6D Q88J51 2.33 93.6% 23.5% 6GRL I0UYD6 2.60 93.4% 36.3% 5OJL Q0CCT3 2.66 92.4% 35.8% 4OQZ S3Z901 2.69 93.4% 28.9% 3QHA A0A0H2ZSR3 2.71 78.9% 16.0% 4D3F Not available 2.81 94.9% 31.5% 6SLE Q4WDZ8 2.84 92.3% 32.9% 4OQY M4ZS15 2.94 88.6% 32.2% 4D3S A0A0J9X1X6 3.02 90.9% 33.7% 6TO4 L7U9F5 3.13 92.7% 38.2% 6SMT A0R5X0 3.16 94.8% 33.1% 6JIT D3Q3R0 3.18 90.0% 33.0% 7OSN A0A1S8Y2S4 3.74 76.1% 43.1% Median 2.76 92.5% 33.4% Average 2.76 90.9% 35.0% 86 Figure 3.8 Multiple sequence alignments of RedE homologues. Featuring the conserved Rossmann domain nucleotide binding motif (GxGxxG, left) the PGGN loop unique to the RedE structure (centre) and residues of the 𝛼10 helix (right). A, Nine of sixteen unique IRED or RedAm hits from secondary structure matching search of the RedE protomer. Sequences are labelled with their name, if applicable, and PDB ID. B, The top 70 sequence similarity BLAST search hits of the UniProt database find only ten other IRED-like proteins with a PGGN-like loop, and each retains a hydrophobic residue at position I246 rather than the common histidine found in other IRED-like sequences (arrow). After the top 10 sequences only every fifth sequence is shown in the alignment. AoIRED (PDB: 5FWN) is the 69th search hit. Sequences are labelled by their UniProt ID. 87 Figure 3.9 Multiple sequence alignment of residues in the contours of the secondary binding cleft in RedE. Featuring the loop between 𝛽5 and 𝛼6 (left), and the N-terminal region of the interdomain helix (𝛼8, right). A, Nine of sixteen unique IRED or RedAm search hits from secondary structure matching of the RedE protomer. B, Sequence similarity searches of the UniProt database find ten other IRED-like proteins with a PGGN-like loop (yellow). Each does not have a conserved glutamate residue at V97 (RedE numbering) and preserves a valine residue at V166 rather than a leucine (arrow). 88 3.3.5 Single residue mutations in RedE Based on the ternary co-crystal structures of RedE, nine single residue variants were generated to investigate the potential role of each binding site for catalytic activity. In the primary site, the D168A variant targets the position often found to be sensitive to mutation in many IREDs and 𝛽HADs.190 The role of the residue at this position is not fully understood, and mutation can eliminate or reduce catalytic activity or alter stereoselectivity. To study the role of this variant, I employed a one-pot reaction where the RedE substrate is created in situ from CPA by the indolocarbazole synthase StaP, and RedM is included to methylate any RedE products (Figure 3.10). This tandem reaction setup is required due to the short lifespan of StaP and RedE reaction products. Under these conditions an ion of 310 m/z, consistent with the [M+H]+ parent ion of methylated 3 discussed below, elutes over one minute when wild-type RedE and RedM is included in solution. For the D168A variant, there is no accumulation of this ion or reductasporine production, suggesting a critical role for this residue. Next, I sought to make mutants that would perturb the hydrophobic lining of the primary site. L239 presses directly on the maleimide ring of AFA forcing it to pack closely to the nicotinamide ring of the cofactor, while the more distant M175 packs with L172, Wʹ210, Fʹ217 and Fʹ235 and together form half of the active site pocket. Assays of the L239A variant result in reduced reductasporine formation, whereas M175A has activity similar to the wild-type RedE. Lastly, residue R242 hovers above the active site near the maleimide ring of AFA and may participate in substrate recognition. It is also a rare large residue to find in this position (Figure 3.8B) and may act to prevent a close fit between domains. However, the R242A variant has identical activity to wild-type RedE. 89 In the secondary site, H249 is positioned to hydrogen bond with indolocarbazole substrates and may stabilize 3 via its pyrrolic nitrogen or may interact with a carboxy group of a carboxylated precursor (such as 4). The H249A variant has markedly reduced reductasporine accumulation, and no [M+H]+ 310 m/z compound is observed, suggesting an important role for this histidine. Next, I created variant A250L to obstruct ligand binding to the secondary site. The A250L variant has normal reductasporine production; however, the accumulation of the [M+H]+ 310 m/z compound is eliminated. Sequence analysis of IREDs found in the PDB and enzymes most similar to RedE suggest there are several residues involved in domain-domain contacts that are not preserved in RedE. For example, IREDs will often have a glutamate or aspartate in place of L253 in RedE and a histidine or asparagine in place of I246, which interact with residues of the nearby 𝛼6 helix and loop of the Rossmann domain. However, replacing these residues in RedE, as in variants L253E and I246H, results in no change from WT activity. Lastly, R254 forms the back wall of the secondary binding cleft and interacts with the unique PGGN loop. Changes to this residue could destabilize the loop and widen the secondary side, but RedE-R254A mutants behave similarly to the wild-type enzyme. 90 Figure 3.10 LC-MS traces of RedE variant reaction products. One pot reactions include 500 𝜇M CPA, 1 mM NADPH, 1 𝜇M StaP, 5 𝜇M FDR, 20 𝜇M FDX, 20 𝜇M RedM and 20 𝜇M RedE variant held at pH 8 with 20 mM HEPES. Production of reductasporine is verified via its unique UV-vis spectrum and mass (pink, 9.2 min, 326 Da). The removal of aspartate from the primary active site via D168A eliminates any reductasporine production and shifts its metabolite profile to look more like inactivated RedE. L239A in the primary site and H249A in the secondary site both reduce reductasporine accumulation and shifts the metabolite profile to appear like boiled RedE (bRedE). A250L does not greatly affect reductasporine production, but no longer accrues the m/z = 310 compound eluting at 13.8 min. Tailing and shoulders of the m/z = 326 peak in each trace is caused by a compound of that mass from enzyme purification. Peaks are coloured according to mass ([M+H]+, m/z): red, 310; light blue, 312; pink, 326; purple, 328; yellow, 342. The peak associated with arcyriaflavin A ([M+H]+ = 326 m/z) is coloured dark blue. 91 3.3.6 Inspection of StaP products Elution of an unknown compound with mass [M+H]+ 310 m/z as a broad peak in RedE variant tandem assays prompted us to inspect StaP reaction products more closely. Previous characterization of StaP reaction products assayed mixtures after 3 hours,29 and given the known propensity for these compounds to decompose quickly I surmised short-lived compounds may have been overlooked. Incubating StaP with appropriate cofactors and CPA substrate, followed by denaturation with acetonitrile after 0, 4, 8, 15, 30 and 160 minutes, the accumulation and depletion of a new compound with [M+H]+ = 296 m/z is detected. This mass is consistent with 3, the proposed RedE substrate (Figure 3.11A). Supplementing reactions with 1 mg/mL catalase in an effort to reduce non-specific oxidation due to hydrogen peroxide released during incomplete turnover of the StaP cofactor system does not alter the compound’s lifespan (Figure 3.11B). 92 Figure 3.11 Extracted ion chromatograms for m/z = 296, the mass consistent with indolocarbazole 3, of StaP reaction products over 2.5 h. A, Reaction includes 500 𝜇M CPA, 1 mM NADPH, 1 𝜇M StaP, 5 𝜇M FDR, 20 𝜇M FDX, 20 mM HEPES pH 8, and B, supplemented with 1 mg/ml catalase. 3.4 Discussion Structures of RedE bound with the substrate mimic arcyriaflavin A unambiguously shows two indolocarbazole binding sites. The primary binding site positions the C5 atom immediately above the nicotinamide ring of the NADPH cofactor for hydride transfer, and the secondary binding site places the indolocarbazole in an adjoining cleft formed between the dimerization domain and Rossmann domain. This secondary binding site is unprecedented among IREDs and RedAms; however, a putative oxidoreductase from Pseudomonas putida KT2440 (PDB: 3L6D, 24 % identity) features a similar cleft separating these domains. Given this unique structural 93 feature is important to catalysis in RedE as determined by mutations I sought out structural and sequence features distinguishing RedE from other IREDs and RedAms which may be involved in the creation of this cleft. With the increased publication of IRED structures over the past ten years, it has become well-established that the Rossmann domain swivels relative to the dimerization domain into distinct open and closed conformations with at least seven IRED and RedAm enzymes captured in either state.199,201–206 These domain movements are often seen when comparing apo structures to binary or ternary complexes. Superposing open and closed conformation protomers onto RedE shows that RedE resembles an exaggerated open conformation, with all atom RMSD values on average 0.3 Å lower for open conformations as to closed conformations. RedE shows no conformational changes upon cofactor or ligand binding, aside from the reorganization of the loop after 𝛽2 upon NADP+ binding positioning R33 to interact with the adenine ring and 2ʹ-phosphate of the cofactor. Given the ternary complexes of RedE have little to no room for the Rossmann domain to move towards the dimerization domain due to the tightly packed AFA in each binding site, the structure of RedE may represent a maximally closed conformation despite resemblance to other IRED open conformations. The open to closed transition in these other IREDs hinge at the N-terminal end of the interdomain helix (RedE: 𝛼8) just below the contact point between 𝛼6 and 𝛼ʹ10. A prominent domain-domain bridging interaction is often found here as a glutamate residue at position 250 or 253 (RedE numbering) whose side chain associates with the N-terminal dipole moment of 𝛼6179,181,201–204,207–211 while hydrogen bonding, or as a salt bridge between an arginine from 𝛼ʹ8 to an aspartate or glutamate on helix 𝛼6.205,206 RedE does not preserve either contact point, instead P194 and G195 from the ‘PGGN’ insertion loop connecting 𝛼ʹ8 to 𝛼ʹ9 (Figure 3.8) extends out to residues P96, V97 and R100 of 𝛼6, and D160, 94 A162, M163 and V166 of the N-terminal of 𝛼8. This is the only interdomain contact to 𝛼6 aside from Rʹ254 in 𝛼ʹ10 via an iodide ion associated with the helix’s N-terminal positive dipole moment. However, removing this interaction in the R254A variant leaves RedE with wildtype activity levels, and likely does not play a significant role in stabilizing the enzyme’s structure. The second series of contacts between domains of IRED and RedAm enzymes occur between 𝛼ʹ9 and 𝛽6-8, which are largely hydrophobic. Many enzymes have a series of large and medium sized residues arranged at this interface, such as tryptophan, phenylalanine, tyrosine and histidine.179,180,201–203,205–207,209,211 Other IRED and RedAm enzymes which do not have large residues at these sites simply do not have intimate interdomain contacts between 𝛼ʹ9 and the 𝛽-sheet of the Rossmann domain, and the majority of domain-domain interactions occur at 𝛼ʹ10.181,199,204,210,212,213 RedE has a series of small residues in 𝛼ʹ9 (A202, V206, T213) which interlock with 𝛽-sheet residues (L131, L133, M155, L157), resulting in a tight interdomain interface. The combination of loose interactions between 𝛼ʹ10 and 𝛼6 alongside a tight interface between 𝛼ʹ9 and 𝛽6-8 results in the Rossmann domain being pulled down towards 𝛼ʹ9 and away from 𝛼ʹ10 relative to other IREDs, creating the exaggerated open conformation and secondary indolocarbazole binding cleft. The other IRED-like enzyme from Pseudomonas putida KT2440 (PDB: 3L6D) also has a similar interdomain cleft between helices 𝛼ʹ10 and 𝛼6. There is a similar domain-domain contact pattern as RedE with loose to no direct connections between 𝛼ʹ10 and 𝛼6, and a tight interface between 𝛼ʹ9 and 𝛽6-8 with mainly small interlocking residues. Notably, the N-terminal of the interdomain helix tucks W169 underneath 𝛼6, imparting a large rotation to the Rossmann domain away from the interdomain helix relative to RedE. Also, 𝛼ʹ9 adopts a kink not seen in 95 other IRED structures between residues 214-218 allowing the relatively large F217 to pack parallel to the Rossmann domain B-sheet rather than push against it. In IRED and RedAm enzymes an active site aspartate or tyrosine is typically found to be essential for catalysis.190 𝛽HAD enzymes are found to use lysine residues at this position in order to catalyze ketone reductions. RedE maintains this active site aspartate (D168); however, it is solely used to anchor the indolocarbazole substrate in the primary active site via hydrogen bonding to the two indole N-H moieties. Inspection of the active site reveals there are no candidates for a catalytic base in RedE, with the only protic residue within a 5 Å sphere of the AFA maleimide group being R242, an unlikely proton donor. Furthermore, the R242A mutant has no impact on enzymatic reaction products. Additionally, StaP is capable of producing an unstable product consistent with 3, the proposed isoindole substrate for RedE. Isoindole readily tautomerizes in solution despite its 10𝜋 electron aromatic ring system.214 The planar, aromatic 2H-isoindole tautomer dominates in polar aprotic solvents, whereas the imine 1H-isoindole tautomer dominates in nonpolar or protic solvents.214,215 Therefore, if the 26𝜋 electron system of 3 acts analogously to isoindole, the imine form should dominate in the primary active site either due to water exposed isoindole nitrogen or pre-equilibration to the imine form from solution or secondary binding site. Without a dedicated catalytic base and with the reactive imine form of the substrate potentially favoured in solution the subsequent reduction via NADPH appears to be catalyzed in RedE by simple substrate orientation and desolvation effects. When the close packing of the primary site around the isoindole nitrogen is disturbed, as in L239A, the overall production of products is greatly reduced. This may be due to the increased mobility of the indolocarbazole and consequent loss in precision alignment of the substrate for efficient hydride transfer. 96 The unexpected distal secondary indolocarbazole binding site formed at the gap between the C-terminal helical dimerization domain and N-terminal Rossmann domain implies a mechanism for unstable substrate management. Compound 3 is found to be unstable, mostly disappearing after 2 hours in vitro (Figure 3.11). This is consistent with other isoindole compounds that are known to oxidize readily when exposed to aerated solutions215 and form compounds analogous to known StaP oxidation products.30 The secondary binding site may sequester indolocarbazole substrate from solution, protecting it from autooxidation and enabling subsequent reduction. Disruption of the secondary site asserts that the site is essential to efficient catalysis with mutant H249A losing substantial activity and A250L no longer accumulating methyl-3. By the structure, A250L sterically excludes indolocarbazoles from binding, whereas the effect of the H249A mutation is less trivial to explain. H249 could play a role in substrate recognition where it would hydrogen bond to the isoindole nitrogen locking the substrate into place within the secondary site. This hydrogen bond may act to lessen the nucleophilic character at C5 and C7 similar to the effect of N-substitution on isoindole molecules, which tend to favour the planar, aromatic tautomer.214,215 Alternatively, H249 may play an active role in selecting for the particular imine tautomer that lines up with the NADPH cofactor for reduction (Figure 3.12). This would imply a direct transfer of the imine substrate from the secondary site to the primary site. Such a transfer could be guided by D168, as an indole nitrogen is available for hydrogen bonding from the secondary site to D168 if the side chain rotates and displaces the nearby water molecule (Figure 3.6B). With this hydrogen bond in place the side chain can swing back down and guide the indolocarbazole from the secondary site to the primary site. These ideas of substrate protection and tautomer selection determining the flux of indolocarbazole products down a productive biosynthetic route mirrors the proposed mechanisms distinguishing RebC and 97 StaC activity in the generation of the aglycone cores of rebeccamycin and staurosporine, respectively.29,45,46 RebC and the StaC-like RebC mutant, RebC-10x, were each crystallized with 7-carboxy-K252c in their active sites, but in alternate tautomeric forms.45,46 RebC traps the enol form requiring decarboxylation-oxidation in order to form arcyriaflavin A,45 whereas RebC-10x traps the keto form, making the carboxylate group available for decarboxylation directly to K252c, the staurosporine aglycone (Figure 3.1B).46 The secondary active site I report here for RedE represents a novel approach to the similar issue of handling short-lived substrates in indolocarbazole biosynthesis. Figure 3.12 Residue H249 in the secondary binding site potentially selects for the imine tautomer of 3 that lines up for reduction upon direct transfer to the primary active site Over the past ten years IREDs and RedAms have drawn increased attention over their potential application to the synthesis of chiral amine products for use in drug manufacturing.174–177 RedAm substrate scopes have previously been limited to smaller ketone/amine combinations, with recent efforts directed at identifying enzymes capable of accommodating larger molecules and engineering them do so more effectively.204,205,211,216,217 Tantalizingly, the RedE structure shows that the combined indolocarbazole binding cleft is some 24 Å wide providing ample space for engineering efforts towards effecting arbitrary reductive aminations. Alternatively, RedE may 98 be used to identify the sequence determinants of an extended binding cleft in order to create them in proven RedAms for expanded substrate scope. In summary, I report the structure of the reductasporine biosynthetic IRED RedE co-crystallized with AFA revealing an unprecedented extended active site cleft capable of binding two indolocarbazole molecules. The structure of and site-directed mutagenesis in the primary binding site suggests reduction is catalyzed via precise substrate orientation and desolvation effects, and the secondary binding site may serve to capture and protect indolocarbazole substrates from spontaneous oxidation. This secondary site may even select for the imine tautomer readily reduced in the primary site when transferring directly from secondary to primary active site. Approaching IREDs from their context in biosynthesis can serve us unique insight into the family via unique structural variations and natural substrate scope, and RedE provides a promising new protein scaffold for reference in iterative engineering. 3.5 Experimental procedures 3.5.1 Cloning, expression and purification of redE and redM Genes redE and redM, codon optimized for expression in E. coli, were cloned into pET28a at the NdeI and XhoI restriction sites with a stop codon was included before the XhoI cut site. Plasmids pET28a-redE or pET28a-redM were transformed into calcium-competent E. coli BL21(DE3) by heat-shock and grown in lysogen broth (LB; 10 g/L NaCl, 10 g/L tryptone, 5 g/L yeast extract) (37 ℃, 180 rpm) with kanamycin (50 𝜇g/mL) to an OD600 of 0.6 before cooling to 16 ℃. Expression was induced by addition of IPTG (to 100 𝜇M) and incubated at 16 ℃ for a further 18 h. Cells were pelleted by centrifugation (4 ℃, 10 min, 3000 × g), and resuspended in 50 mM Tris (pH 8), 500 mM NaCl, 5 mM imidazole supplemented with either 1.5 mM TCEP or 5 mM β-mercaptoethanol (BME) reducing agent. Cell suspensions were lysed by sonication (25 99 % amplitude, 4 s on / 8 s off cycles, 6 min), and cell debris removed via centrifugation (45 min, 15000 × g). The supernatant was loaded onto Ni-IDA affinity resin and eluted with a 20-500 mM imidazole step gradient in 50 mM Tris (pH 8), 500 mM NaCl, and either 1.5 mM TCEP or 5 mM BME. Elution fractions with recombinant protein were passed through a 0.22 𝜇m syringe filter and purified via size-exclusion chromatography (20 mM Tris (pH 8), 50 mM NaCl with corresponding reducing agent. SDS-PAGE was used to track the purity of resulting fractions for further use in crystallography screens. 3.5.2 Selenomethionine incorporation into RedE E. coli BL21(DE3) transformed with pET28a-redE were grown in LB medium with kanamycin (50 𝜇g/mL) overnight. From this culture, a 30 𝜇L aliquot was inoculated into 30 mL of antibiotic supplemented M9 medium for growth overnight. Starter cultures (10 mL) were each added to 1 L of M9 medium with kanamycin and grown at 37 ℃ (180 rpm) for 4 h to an OD600 of 0.7 before cooling to 16 ℃ over 30 min. Lysine, threonine and phenylalanine (100 mg each) followed by leucine, isoleucine, valine and selenomethionine (50 mg each) were added to 1 L of medium.154 Cultures were then induced with 100 𝜇M IPTG and grown for 18 h. Selenomethionine-substituted RedE were purified as described previously with buffers supplemented with 5 mM BME. 3.5.3 RedE crystallization Purified protein was screened on a variety of commercially available screens (Hampton, Index, Top96, MCSG) in a 96-well sitting drop format with 2 𝜇L drops (1:1 ratio of protein to precipitant solution). Preliminary hits were optimized in a hanging-drop format with drops mixed in a 1:1 ratio of well to protein solution. Diffraction quality RedE crystals were grown over wells containing 0.33 M NaI, 0.1 M Tris pH 8.5, 21-23% PEG 3350 and 1.5 mM TCEP or 5 mM BME 100 using up to 42 mg mL-1 of protein. Under these conditions RedE co-crystallized with NADP+ (100 mM in water) and/or arcyriaflavin A (100 mM in DMSO) when 10 mM and 1.2 mM of each, respectively, were included in the protein solution. Crystals of apo-RedE grew at 0.35 M NaI, 0.1 M bicine pH 9, 25 % PEG 3350 and 5 mM BME. Selenomethionine-substituted RedE crystals were prepared similarly with mother liquor at 0.3 M NaI, 0.1 M Tris pH 8, 25% PEG 3350 and 10 mg mL-1 protein. All crystals were soaked for 1-10 min in mother liquor prepared to 10-15% ethylene glycol before flash-freezing in liquid nitrogen. Crystals were tested for diffraction using a Rigaku MicroMaxTM-007HF (1.54 Å, rotating copper anode) with Saturn 944+ CCD detector, and those diffracting to at least 3 Å were retained for data collection at a synchrotron. 3.5.4 Data collection, structure determination, and refinement Datasets for the selenomethionine-substituted RedE were collected on beamline 08ID-1 at the Canadian Light Source (Saskatoon, Canada) using MX300-HE and Pilatus 6 M detectors. Datasets for RedE co-crystals were collected on beamline BL9-2 at the Stanford Synchrotron Radiation Lightsource (Menlo Park, United States). Data sets were integrated using iMOSFLM155 or XDS218 and merged in AIMLESS.158 Selenomethionine structures were phased via single anomalous dispersion of selenium and iodine atoms via the Autosol tool in the Phenix software package.219 The SAD phasing results were then input into Autobuild160 for initial model building followed by several rounds of manual inspection and building in COOT161 and refinement via phenix.refine.163 Native protein datasets were processed similarly with phases solved for via molecular replacement using the SeMet-RedE model through PHASER-MR.220 Ligand restraints for arcyriaflavin A were generated using eLBOW.166 B-factors and partial occupancy for alternate conformations of arcyriaflavin A were calculated through the automated 101 phenix.refine procedure to optimize stereochemical and atomic displacement parameter weights to minimize the Rfree value. 3.5.5 Site-directed mutagenesis of RedE Primers were designed so that the overlap region containing the mutation had a Tm ~ 55 ℃ and the non-overlapping region had a Tm about 7 ℃ higher (calculated against the complementary DNA strand).167 Where possible mutations were included that removed or added restriction sites for preliminary screening via digests. Standard PCR was carried out using an annealing step set to the higher Tm and ended with a 10 min elongation step. The PCR product mixture was incubated with DpnI before transforming the digest solution as described previously into E. coli DH5𝛼. Purified plasmids were screened for restriction fragment polymorphism and mutations confirmed by sequencing. 3.5.6 Cloning and expression of rebO and rebD, and production of CPA Genes rebO and rebD were amplified from gDNA (Lechevalieria aerocolonigenes DSM 44217), and ligated into pET28a and pET22b, respectively. These plasmids were transformed into chemically competent E. coli BL21(DE3) cells and used to prepare CPA according to the previously published procedure.43 3.5.7 Expression and purification of StaP, ferredoxin and ferredoxin reductase Plasmid pET28a-staP was kindly provided by Catherine Drennan (Massachusetts Institute of Technology), and plasmids pET28a-fdr and pET28a-fdx coding for Synechococcus elongatus PCC 7942 ferredoxin-NADP oxidoreductase (FDR) and ferredoxin I (FDX), respectively, were received from Dr. Hai-Yan He (Institute of Medicinal Biotechnology, Chinese Academy of Medical Sciences). 102 E. coli BL21(DE3) transformed with pET28a-staP, pET28a-fdr or pET28a-fdx were grown in LB medium with kanamycin (50 𝜇g/mL) to an OD600 of 0.5 before cooling to 16 ℃ over 30 min. Each culture was then supplemented with 100 𝜇M IPTG and cells with plasmids coding for heme proteins (StaP and FDX) were further supplemented with 1 mM 𝛿-aminolevulinic acid before incubating for 18 h. Media was removed by centrifugation (4℃, 10 min, 3000 × g), and cells resuspended in 50 mM Tris (pH 8), 500 mM NaCl, 5 mM imidazole. The bacterial suspension was lysed via sonication (25% amplitude, cycles of 4 s on and 8 s off for 6 min), and cell debris cleared via centrifugation (45 min, 15 000 × g). Supernatant was passed through a Ni-affinity column and enzyme harvested over an imidazole step gradient (5-300 mM). Fractions containing recombinant enzyme were dialyzed to remove excess imidazole and salt before screening for activity against CPA followed by adding glycerol to 10 % (v/v) and storing at −70 ℃. 3.5.8 Analysis of RedE variants A typical reaction includes 500 𝜇M CPA incubated with 1 𝜇M StaP, 20 𝜇M RedM and 20 𝜇M RedE variant in the presence of 1 mM NADPH, 5 𝜇M FDR and 20 𝜇M FDX cofactors held at pH 8 with 20 mM HEPES. CPA, FDR and FDX. Reactions are quenched with the addition of an equal volume of ACN and analyzed on a reverse phase HPLC column (1 mL min-1, 98 % H2O to 98 % ACN gradient over 30 min, each solvent supplemented with 0.1 % formic acid). Elution was monitored using a diode array from 220-500 nm or by mass. 3.5.9 Time-course analysis of StaP reaction products StaP reactions include 500 𝜇M CPA, 1 mM NADPH, 1 𝜇M StaP, 5 𝜇M FDR, 20 𝜇M FDX, held at pH 8 with 20 mM HEPES at room temperature. Reactions were quenched with equal volumes of ACN and analyzed on a reverse phase HPLC column (1 mL min-1, 98 % H2O to 98 103 % ACN gradient over 18 min, each solvent supplemented with 0.1 % formic acid). Elution was monitored at 280 nm and by mass. 3.5.10 Visualization and analysis software Pymol was used for secondary structure matching and creating protein visualizations.165 Surface areas and dimer interaction surfaces were calculated via PISA.90 Protein sequence alignments were generated using Clustal Omega169 or COBALT118 and visualized with Jalview.170 104 Chapter 4: The structure of the oxygen, pyridoxal phosphate-dependent capuramycin biosynthetic protein Cap15 4.1 Synopsis Pyridoxal phosphate-dependent enzymes able to use oxygen as a co-substrate have emerged in multiple protein families. Here I use crystallography to solve the 2.40 Å resolution crystal structure of Cap15, a nucleoside biosynthetic enzyme that catalyzes the oxidative decarboxylation of glycyl uridine. Our structural study captures the internal aldimine, pinpointing the active site lysine as K230 and showing the site of phosphate binding. Our docking studies reveal how Cap15 is able to catalyze a stereoselective deprotonation reaction, and bioinformatic analysis reveals active site residues that distinguish Cap15 from the structurally related D-glucosaminate-6-phosphate ammonia lyase and L-seryl-tRNA(Sec) selenium transferase (SelA). Our work provides the structural basis for further mechanistic investigation of a unique biosynthetic enzyme and provides a blueprint for understanding how oxygen reactivity emerged in the SelA-like protein family. 4.2 Introduction Pyridoxal-5ʹ-phosphate (PLP)-dependent enzymes rarely react with oxygen. However, the cofactor is theoretically capable of reacting with oxygen, because it transitions through a stabilized carbanion, and all carbanions are subject to reaction with electrophiles, including oxygen.221,222 In recent years, a number of other PLP-dependent natural product biosynthetic enzymes have been reported to use oxygen in their catalytic cycles.223–237 These enzymes include Cap15 (UniProtKB: E1CG36), a key enzyme from the biosynthetic pathway to capuramycin, a nucleoside antibiotic from Streptomyces griseus.238 Capuramycin and its derivatives are antimycobacterial agents that inhibit bacterial translocase I.239–242 Cloning and characterization 105 of the biosynthetic gene clusters of capuramycin-like molecules243–245 shows that the capuramycins’ distinctive uridine-5ʹ-carboxamide (CarU) moiety is assembled in at least three steps starting from uridine-5ʹ-monophosphate.238,245,246 First, the Fe(II)/𝛼-ketoglutarate-dependent dioxygenase CapA oxidatively dephosphorylates uridine monophosphate to the aldehyde. Second, the PLP-dependent transaldolase CapH condenses the aldehyde with threonine to form 5ʹ-C-glycyl uridine (GlyU). Third, Cap15 catalyzes the oxidative-decarboxylation of this unusual 𝛽-hydroxyamino acid intermediate to give the CarU product (Figure 4.1). Figure 4.1 Enzymatic transformations catalyzed by Cap15, DGL and SelA. D-glucosaminate-6-phosphate ammonia-lyase (DGL) and L-seryl-tRNA(Sec) selenium transferase (SelA) are the only two characterized enzymes closely related to Cap15 Huang and co-workers carried out detailed in vitro analysis of Cap15 to reveal that Cap15 is an O2- and phosphate-dependent protein, unable to execute the reaction either in anaerobic conditions or in non-phosphate buffers.238 Incorporation of 18O2 in the product demonstrated that the enzyme is a monooxygenase, in contrast to enzymes like the arginine oxidases RohP234 and MppP,235 or aromatic acetaldehyde synthases223,247 where water is the source of oxygen in the final product. While each of these enzymes, including Cap15, come from the aspartate O N NHOHHOHOH2NOOOHOHO N NHOHHOHOOOOHH2NCap15CO2H2PO4-O25ʹ-C-glycyl uridine (GlyU) 5ʹ-C-carboxamide Uridine (CarU)+H3NOHHOOHO-OPOOO--OOHOOHO-OPOOO--ODGLNH4+H2NHOOtRNASecSelAPOSe-O--O POO-O--OH2N-SeOtRNASec106 aminotransferase (AAT)-like class of PLP-dependent enzymes, Cap15 is the first oxygen-dependent enzyme found in the L-seryl-tRNA(Sec) selenium transferase-like family (IPR018319). The only two other characterized enzymatic activities in this family are the eponymous L-seryl-tRNA(Sec) selenium transferase (SelA), which catalyzes selenophosphate dependent selenium transfer to L-seryl-tRNA(Sec),56,248 and D-glucosaminate-6-phosphate ammonia lyase (DGL), which catalyzes 𝛽-hydroxyl elimination and 𝛼-deamination of D-glucosaminate-6-phosphate. (Figure 4.1).249 Notably, both Cap15 and DGL act on D-amino acid substrates, which is unusual for AAT-like enzymes and are typically substrates of structurally distinct PLP-dependent racemases and transaminases.250,251 To interrogate the unique monooxygenase activity of Cap15, determine its relationship to other members of its enzyme family, and provide more general insight into PLP-dependent enzymes that employ O2 for catalysis, I set out to solve its three-dimensional structure. Here I report the homooctameric crystal structure model of Cap15 solved at 2.40 Å with phosphate bound in the active site near the PLP internal aldimine, providing insight into substrate specificity and the utilization of molecular oxygen during catalysis. 107 4.3 Results Table 4.1 Data collection and refinement statistics. holo-Cap15a PDB Accession 8T7J Wavelength (Å) 0.97946 Resolution range (Å) 38.90 – 2.40 (2.49 – 2.40) Space group C 2 2 21 Unit cell 142.59 220.986 96.735 90 90 90 Total reflections 812679 (53569) Unique reflections 60031 (4126) Multiplicity 13.5 (13.0) Completeness (%) 99.51 (99.38) Mean I/𝜎(I) 19.3 (1.8) Wilson B-factor 50.89 Rmerge 0.090 (1.501) Rmeas 0.093 (1.562) Rpim 0.025 (0.462) CC1/2 1.000 (0.881) Reflections used in refinement 59710 (5889) Reflections used for Rfree 1989 (195) Rwork 0.2218 (0.3129) Rfree 0.2676 (0.3829) Number of non-hydrogen atoms 10019 Macromolecules 9768 Ligands 28 Solvent 223 Protein residues 1438 RMS (bonds) (Å) 0.003 RMS (angles) (°) 0.59 Ramachandran favored (%) 96.65 Ramachandran allowed (%) 2.99 Ramachandran outliers (%) 0.36 Rotamer outliers (%) 0.24 Clashscore 2.08 Average B-factor (Å2) 73.15 Macromolecules 73.51 Ligands 74.97 Solvent 57.00 Number of TLS groups 25 a Values in parenthesis are for the highest resolution shell 108 4.3.1 Overall structure of Cap15 The refined Cap15 structure (Table 4.1) exhibits the fold of type I PLP-dependent enzymes, typified by AAT.252,253 In such structures, the monomer consists of a large, rigid PLP-binding major domain, and a smaller, more mobile minor domain formed from a short stretch of N-terminal amino acids and approximately 130 amino acids from the C-terminal. This fold persists even at very low sequence identities.251,254 In Cap15, residues 45-280 form the PLP-binding major domain with a core of a seven-strand mixed 𝛽-sheet (𝛽2-8), composed of five parallel and two antiparallel strands sandwiched between seven 𝛼-helices (𝛼1-10) (Figure 4.2, 4.3A). Each parallel strand is arranged as part of a 𝛽𝛼𝛽 motif. Residues 17-44 and 281-384 form the minor domain, consisting of two small adjacent 𝛽-sheets (𝛽9-10, 12 and 𝛽1, 11) stacked against three 𝛼-helices (𝛼11-13). The B-factors of the minor domain (C𝛼 average of 79 Å2) are much higher than that of the major domain (C𝛼 average of 45 Å2). High B-factors are also observed in the minor domains of SelA248 and DGL.249 In the case of DGL, the high B-factors in the minor domain have been attributed to its relatively open conformation.249 The folds of the major and minor domains are widely conserved in the AAT family. A comparison of Cap15 with structurally related members of the AAT family show that the minor domain can have several large insertions with additional secondary structures when compared against the Cap15 structure (Table 4.2, Figure 4.4). 109 Figure 4.2 Overview of the structure of Cap15. A, Secondary structures of the Cap15 monomer with conserved active site residues shown as green sticks. B, View down the 2-fold crystallographic symmetry axis of the Cap15 octameric assembly. C, View of residues S255-K265 between 𝛼9 and 𝛼10 (blue). This loop is buried at the center of homodimer interface and contacts the equivalent loop in its dimer partner (violet), the phosphate moiety of the internal aldimine, several N-terminal residues (beige), the N-terminal end of 𝛼3 (yellow), as well as residues in 𝛼5 (orange). D, Hydrogen bonds at the dimer-dimer interface. 110 Figure 4.3 Topology diagram, interdimer interface, and secondary structures with residue ranges. A, Topology diagram of Cap15, highlighted by the major (orange/red) and minor (cyan/purple) domains. The internal aldimine (K230) is indicated with a hexagon and conserved active site residues are marked in green circles (H180, D206, R318, R322, R353, R367). B, Focus on a single Cap15 homodimer with hydrogen bonding residues at the interdimer interface highlighted as sticks. C, Cap15 secondary structures with residue ranges labelled. Table 4.2 Secondary structural alignments of Cap15 to representative aspartate aminotransferase family enzymes and arginine oxidases. Major Domain (236 C𝛼) Minor Domain (127 C𝛼) Enzyme PDB RMSD (Å) N (C𝛼) RMSD (Å) N (C𝛼) SelA 248 3WCO 1.3 152 1.2 66 Kynureninase 255 2HZP 4.1 149 3.4 76 Tryptophanase 256 2C44 3.5 188 3.0 66 GAD65 257,258 2OKK 3.6 192 3.2 72 Cystalysin 259 1C7N 3.5 138 3.9 82 Serine hydroxymethyltransferase 260 1KKJ 4.2 187 3.4 71 Ornithine decarboxylase 261 1C4K 3.3 159 3.1 78 MppP 228 5DJ1 4.7 183 3.1 80 RohP 234 6C3B 5.1 200 3.3 87 Plu4 262 7N79 5.4 195 3.2 85 111 Figure 4.4 Side by side comparisons of representative structures of PLP-dependent enzyme families. Cap15 belongs to the L-seryl-tRNA selenium transferase-like family (IPR018319) and bears greatest structural resemblance to SelA (RMSD = 1.3 Å, n = 152) among these examples; however, the fold for the large domain (blue) is largely conserved among PLP-dependent enzymes with RMSDs from 3.3 Å to 4.2 Å. The small domain (red) has more structural variability, but the core of three 𝛼-helices and three strand 𝛽-sheet is retained among all examples with alignment RMSDs of these structural elements ranging from 1.2 Å up to 3.9 Å. The N-terminal residues (green) show more variety and can fold into independent domains as in ornithine decarboxylase or other secondary structures that interact with the C-terminal domain. Cap15 forms a homodimer with an interface spanning approximately 3200 Å2 (19 % of the monomer surface area) involving 168 residues and forming 36 hydrogen bonds and two salt bridges (Figure 4.3B). Major contributors to the interface are the N-terminal residues wrapping around the dimer partner between the 𝛼1, 𝛼2, and 𝛼10 helices. The N-terminus of 𝛼3 has several dimer-stabilizing interactions with the loop between 𝛼9 and 𝛼10, while the C-terminus of 𝛼4 interacts with residues 135-142 from the 𝛼5 helix. Lastly, the loop between 𝛼9 and 𝛼10 forms the deepest and largest portion of the dimer interface juxtaposed against the equivalent loop from the dimer partner in addition to the internal aldimine loop at K230 (Figure 4.2C). R259 and Cap15 SelA(3wco)Ornithine Decarboxylase(1c4k)Tryptophanase(2c44)Cystalysin(1c7n)GAD65(2okk)Kynureninase(2hzp)Serine Hydroxymethyltransferase(2hzp)112 K262 span the active site interface to form a salt bridge and hydrogen bond with the phosphate moiety of the nearby PLP. Four small 520 Å2 interfaces between homodimer subunits build out an octameric assembly (Figure 4.2B) similar to the one observed in DGL.249 Each interface involves 40-50 residues forming eight hydrogen bonds and two salt bridges (Figure 4.2D). The interdimer interface occurs between the major domain 𝛽-sheets where loops C-terminal to strands 𝛽3-5 form the bulk of the dimer-dimer interactions. Hydrogen bonds occur between V153 and Sʹ128, N154 and Sʹ130, E155 and Gʹ133 and the corresponding reciprocal interactions at the symmetric interface. K125 to Eʹ149 and the reciprocal E149 to Kʹ125 form the two salt bridges at the centre of the dimer-dimer interface. This configuration is reminiscent of the decameric SelA assembly, which is a pentamer of homodimers.56 The interdimer interface occurs at the same area of each protomer, but an extra loop of amino acids forms a two-strand antiparallel 𝛽-sheet between SelA dimers resulting in roughly double the contact surface area (ca. 1180 Å2). Size exclusion chromatography analysis of Cap15 in a gel filtration column rated for effective separation of molecular weights between 10-600 kDa shows a broad peak at an elution volume corresponding to a complex of approximately 300 kDa, consistent with the 336 kDa octamer (Figure 4.5). 113 Figure 4.5 Size-exclusion chromatography traces for Cap15 octamer and molecular weight standards. Absorbance read at 280 nm. Cap15 octamer, 336 kDa; Cap15 dimer, 84 kDa; sweet potato 𝛽-amylase, 200 kDa; yeast alcohol dehydrogenase, 150 kDa; bovine serum albumin, 66.5 kDa 4.3.2 Active site Pyridoxal 5-phosphate binds to Cap15 at K230 as a Schiff base (Figure 4.6A). The PLP phosphate is bound by a network of hydrogen bonds including the A88 (3.0 Å) and A89 (3.2 Å) amide nitrogens, the T37 (3.0 Å) and S227 (2.7 Å) side chain hydroxyls, Rʹ259 (2.8 Å) guanidino group and Kʹ262 (2.8 Å) amine. Rʹ259 and Kʹ262 reach across the dimer interface to the PLP phosphate forming salt bridges. The PLP is positioned at the N-terminal end of 𝛼3, a typical feature in the AAT family.252 D206 provides the negative charge and hydrogen bond receptor to maintain the protonated state of the PLP pyridinium ring necessary for activity. This protonated state is visible in the solution state UV-Vis spectrum as a peak at 415 nm (Figure 4.6B). This corresponds with the expected 𝜆max for internal aldimines ranging from 0102030405060708040 50 60 70 80Absorbance (mAU)Volume (mL)66 kDa150 kDa200 kDaBlue Dextran(2000 kDa)Cap15114 410-430 nm.263–267 The peak around 325-330 nm may arise from a substituted internal aldimine intermediate similar to cystalysin.264,267 Most AAT family enzymes have a hydrogen bond between the PLP 3-hydroxyl and a nearby residue, as well as a 𝜋-stacking interaction between the pyridinium ring and an aromatic residue. These interactions are not present in Cap15, as the PLP pyridinium ring is shifted away from stacking with the nearby Y131 and is instead cradled between Y131 and H180 with less specific interactions. Plu4 also has no 𝜋-stacking interaction with the PLP pyridinium ring.262 These key interactions are largely identical to DGL and SelA, with different residues at positions 88 and 89 in DGL and SelA, though the main chain hydrogen bonding is maintained. SelA also replaces Kʹ262 with a second arginine at that position. Members of the AAT family typically have an arginine in the small domain that recognizes the 𝛼-carboxylate in their substrates. Cap15, DGL and SelA lack an arginine at this position in 𝛽9 (R386 in chicken AAT), and instead have several other arginine residues in close proximity directed towards the active site. In particular, near 𝛽11 in Cap15 and DGL has R353 and R332, respectively, whose guanidino group extends into the same space as the side chain of the conserved arginine in AAT. In Cap15 the other arginine residues include R318, R322, and R367, and this group of four residues orient around a large volume of electron density which is well modelled by a phosphate anion. The side-chain hydroxyl of T179 forms a hydrogen bond to this phosphate anion as well. Initial crystallization attempts used HEPES and Tris buffers in order to avoid phosphate salt crystals; however, potassium phosphate buffers were required for consistent Cap15 crystallization. Furthermore, Huang et al. find that Cap15 only has measurable in vitro activity while phosphate anions are present.238 Each chain in the model has a phosphate anion at this position, though with high B-factors (average of 69 Å2), consistent with the high B-factors of the C-terminal minor domain. 115 Figure 4.6 View of the Cap15 active site residues and UV-Vis spectrum of Cap15 solution and crystal. A, Cap15 binds PLP at K230 in the active site with a nearby phosphate anion and each is shown with mFo – DFc map contoured at 3𝜎. Hydrogen bonds and salt bridges are drawn as dashed lines, and residues R259 and K262 from the dimer partner are shown in blue. B, UV-Vis absorbance spectra of Cap15 solution and crystal. The crystal UV-Vis spectrum is scaled down to match the solution spectrum absorbance at 325 nm. 4.3.3 External aldimine docking Attempts to co-crystallize Cap15 with uridine, CarU and acetal-protected CarU yielded no observable electron density for any of these molecules in any resulting structure. Therefore, in order to provide insight into the role of nearby active site residues, I sought to dock the Cap15 substrate (5ʹS, 6ʹR)-GlyU into the active site cleft of the internal aldimine form. Since the role of the phosphate anion cannot be determined, it was both excluded and included during the substrate docking. Rigid enzyme docking using Autodock Vina produces several substrate poses 116 with geometry unlikely to form external aldimines. Allowing T37, S177 and H180 to move freely using Autodock Vina’s flexible residues option does not yield substantially different substrate poses. Next, I sought to dock the entire GlyU-PLP external aldimine; however, no poses could reproduce the appropriate PLP-binding, regardless of the exhaustiveness of searches or flexible residue settings. However, aligning the GlyU-PLP external aldimine model with the PLP of the internal aldimine and setting the external aldimine as a flexible residue under Autodock Vina’s settings works around this limitation. In this scenario, when a molecule is docked to the enzyme Vina will perform a conformational search of the external aldimine to find a favourable conformer. Docking either the phosphate anion or water into the active site produces two predominant binding modes. One places the uracil ring at the junction of R318, R322, R353 and R367 and hydrogen bonds to S177 (Figure 4.7A). The other orients the uracil ring into the gap between R353 and Q33 where the ribose hydroxyls are free to hydrogen bond with the docked phosphate anion, which is held in a similar fashion to the experimentally determined position (Figure 4.7B). When H180 is allowed to move freely during docking it orients to make a hydrogen bond with a ribose hydroxyl. K230 in this state must be set to be a flexible residue as it will otherwise occlude sensible conformations of the external aldimine, and it was modelled in its protonated state. It is found to extend to either the PLP phosphate group or the docked phosphate anion if it is placed nearby. Since docking water under these conditions produces a conformer that could accommodate a phosphate anion bound to the nearby cluster of arginine residues, I repeated the docking run again with the phosphate anion held rigidly at the experimentally determined position and reproduced the conformation, including residue positioning. 117 Figure 4.7 Representative poses of external aldimines for the docking of water and phosphate anion using the flexible side chain method in Autodock Vina. Residues allowed to move freely in a conformational search are coloured yellow and hydrogen bonds are indicated with dashed lines. A, Representative poses of the (5ʹS, 6ʹR)-GlyU-PLP external aldimine where the uracil ring occupies the phosphate anion binding site, and B, where the position of the phosphate anion is reproduced in the active site. C, Representative poses for the 6ʹ-epimer, (5ʹS, 6ʹS)-GlyU-PLP, external aldimine without docked active site phosphate and D, with docked active site phosphate. Both display poor geometry for catalysis with the imine bond rotated out of plane from the pyridinium ring system or the 𝛼-hydrogen pointing away from the presumed catalytic base K230 which initiates catalysis, respectively. Water and hydrogens omitted for clarity. Unfortunately, this method of restraining the docking search space is limited by the scoring algorithm being tied to the docked molecule rather than the GlyU external aldimine or both the external aldimine and phosphate anion. However, this strategy has been applied to the docking of covalent inhibitors, and is found to reliably reproduce experimental binding modes.268 Benchmarking this strategy with medium to large sized external aldimines found in the PDB, 118 with a focus on nucleoside sugar substrates, reproduces the experimentally observed binding mode with low RMSD values (Table 4.3, Figure 4.8). If this Cap15 model represents the catalytically active conformation, then it is likely that one of the binding modes produced using this flexible side chain method is a reasonable approximation of the binding mode of the GlyU external aldimine. This indicates that residues S177 and H180 can play a role in substrate recognition. Table 4.3 Autodock Vina benchmarking of conformational searches of medium to large sized external aldimines found in the PDB. The pyridinium ring and phosphate moieties of the external aldimine were fixed in place at their experimental coordinates, and the remainder of the molecule is allowed to move freely. In each case Autodock Vina reproduced the observed binding mode at low RMSD values. 1DJ9 3B8X 3DR4 4K2M 4OCA 4ZTC 7N63 8E77 Number of rotatable bonds 10 15 14 9 15 15 13 16 Number of poses matching experimental result 14 6 6 20 7 5 6 4 Total number of poses 20 20 11 20 20 15 20 20 Ratio 0.70 0.30 0.55 1.00 0.35 0.33 0.30 0.20 RMSD over poses matching experimental results (Å) Average 0.750 1.180 0.517 1.472 0.608 0.293 1.473 1.643 Maximum 1.102 1.660 0.579 2.006 1.637 0.301 1.748 1.737 Median 0.830 1.130 0.506 1.855 0.534 0.294 1.695 1.724 Minimum 0.525 1.006 0.499 0.237 0.292 0.277 0.303 1.389 119 Figure 4.8 Visualization of Autodock Vina flexible side chain method conformational searches benchmarking. The experimental coordinates of each external aldimine are rendered as grey ball and stick models and docked poses as light blue lines. While the scoring algorithm is not tied explicitly to the external aldimine conformation, the method reliably produces docking poses with low RMSD values to the experimental model. Atoms in the PLP moieties are excluded from RMSD calculations. Next, I expanded our conformational searches to the (5ʹS, 6ʹS)-GlyU epimer, as it is reported that Cap15 exclusively transforms the (5ʹS, 6ʹR)-GlyU epimer. When the phosphate anion is held rigidly in the active site, there are no calculated poses that have the imine bond in plane with the PLP pyridinium ring. Similar poses are observed when an identical search is performed without the phosphate anion in the active site (Figure 4.7C). Alternatively, when the 120 phosphate anion is docked into the active site the imine bond only becomes planar to the ring system when it adopts a conformation similar to the (5ʹS, 6ʹR)-GlyU epimer (Figure 4.7D). This points the 𝛼-hydrogen away from K230 and the nearest residue capable of acting as a base, H180, is over 5 Å away and oriented improperly for hydrogen abstraction. Therefore, this docking approach reproduces the logical expectation that the active site cannot accommodate 𝛼-hydrogen abstraction from the (5ʹS, 6ʹR)-GlyU epimer. 4.3.4 Conservation of active site residues To provide more information on active site residues, multiple sequence alignments of Cap15 BLAST269,270 search results on the NCBI clustered nr database271 were carried out. The top 1000 hits have strongly conserved PLP-binding residues (T37, Y131, D206, A208, S227, K230, Rʹ259, Kʹ262), phosphate coordinating residues (R318, R322, R353, R367), as well as two residues after 𝛽5 (S177, H180). (Table 4.4) Other residues forming the boundary to the active site cavity (Q33, S36) are not conserved, even amongst the most similar 250 sequences. Notably, T179, which hydrogen bonds to the active site phosphate anion, is present in only ten other sequences as 177–SxTH–180, and the most conserved sequence here is 161–SHH–163 (DGL numbering) with one fewer residue in the loop. Superposing DGL onto Cap15 shows that T179 best matches to DGL-H162’s position, and R178 represents a single residue insertion. Exploring the genomic contexts of the ten other sequences that share this 177–SxTH–180 motif show that each is in a gene cluster possessing a CapA and CapH homologue; two enzymes necessary to produce the Cap15 substrate. (Figure 4.9) Expanding the BLAST search scope to the top 5000 hits shows that the conservation in the phosphate anion binding region and SHH motif is diminished, while the residues involved in PLP-binding remain well conserved. 121 Table 4.4 Multiple sequence alignment of the top N blastp hits querying the Cap15 sequence against the clustered nr NCBI database.271 Only sequences with more than 85% coverage, and consequently possessing both large and small domains of the type I fold PLP-dependent enzymes were retained for sequence alignment. N represents the maximum number of target sequence hits returned. The number of sequences remaining after filtering low coverage hits is reported in brackets. Conservation rates (percent ratio of sequences residue(s) is/are found in) are reported in parentheses. Most conserved residue and conservation rate N = 250 [248] N = 500 [500] N = 1000 [930] N = 5000 [2300] PLP binding residues T37 T (100 %) T or S (98 %) T or S (99 %) T or S (99 %) Y131 F (91 %) Y or F (100 %) Y or F (99 %) Y or F (76 %) D206 D (100 %) D (100 %) D (100 %) D (100 %) A208 A (100 %) A (99 %) A (99 %) A (96 %) S227 S (100 %) S (99 %) S (99 %) S (94 %) K230 K (100 %) K (100 %) K (100 %) K (99 %) R259 R (100 %) R (100 %) R (99 %) R (99 %) K262 K (100 %) K (100 %) K (99 %) K (96 %) PO43- binding residues R318 R (100 %) R (98 %) R (74 %) R (29 %) R322 R (100 %) R (100 %) R (99 %) R (50 %) R353 R (100 %) R (100 %) R (98 %) R (47 %) R367 R (100 %) R (99 %) R (75 %) R (29 %) Other active site residues Q33 S or C (88 %) S or C (86 %) S or C (86 %) S or C (30 %) S36 M (92 %) M (92 %) M (92 %) M (36 %) S177 S (100 %) S (100 %) S (97 %) S (39 %) T179 H (98 %) H (98 %) H (96 %) H (39 %) H180 H (100 %) H (100 %) H (98 %) H (39 %) In order to visualize the distribution of active site motifs I created a sequence similarity network (SSN)171 of the SelA-like enzyme family (IPR018319). This family includes all homologues of Cap15, and has only three other characterized enzymes: SelA56,248 (Aquifex aeolicus), putative archaeal SelA272 (Methanocaldococcus jannaschii) and DGL249 (Salmonella enterica). The initial SSN (alignment score threshold (AST) ≥ 40) splits the family into three main clusters representing these three characterized enzymes, where Cap15 resides in the same cluster as DGL (Figure 4.10). All clusters retain the key PLP-binding residues. Raising the AST to 76 splits the DGL/Cap15-like cluster in two, and Cap15 remains clustered with DGL. 122 Comparing these two clusters via multiple sequence alignments in Clustal Omega shows that the DGL/Cap15-like cluster widely preserves the arginine rich phosphate-binding motif and SHH loop, while the other cluster has variable sequences in these two regions. Figure 4.9 Alignment of genes in the vicinity of Cap15 homologues as clustered by the SSN of the SelA-like family (AST = 110). Genes are coloured per their proposed function or their similarity to those enzymes. In every example there are homologues of capA and capH responsible for the biosynthesis of GlyU. Each Cap15 homologue bears an active site SxTH motif rather than the more common SHH motif found throughout the SSN. The A-503083 (cap, accession: AB538860) and A-102395 (cpr, accession: KP995196) gene clusters are represented in the cluster, while the A-500359 (accession: AB476988) cluster was not. The corresponding open reading frame (orf12ʹ) product is not present in the Uniprot database used to generate the SSN. Remaining gene clusters originate in (i) Kitasatospora viridis, accession: NZ_VIWT01000005 (ii) Actinokineospora spheciospongiae, accession: NZ_AYXG01000004 (iii) Actinocrispum wychmicini, accession: NZ_SLWS01000004 (iv) Streptomyces sp. Ncost-T6T-1, accession: NZ_LACR01000019 (v) Streptomyces sp. Alain-F2R5, accession: NZ_NQOS01000102 (vi) Streptomyces sp. alain-838, accession: NZ_NQNW01000033. All accession numbers are either GenBank or NCBI reference sequence identifiers. viviviiiiiicpr (A-102395)(A-500359)cap (A-503083)5 kbpcap15GlyUamide transferself-resistancepyranosepolyamideaminocaprolactamribose modificationcapQ and capRtransporter123 The Cap15 subcluster separates from the main DGL cluster at an AST of 109 producing a total of 11 subclusters with five or more members (Figure 4.10). Multiple sequence alignment of members of these subclusters shows that the Cap15 subcluster is one of three that do not preserve the SHH active site motif found in DGL (Table 4.5). Of these three, the Cap15 subcluster is the only one to retain the arginine rich phosphate-binding motif and histidine at position 180 (Cap15 numbering) present in 98 % of sequences in the SSN. Of the remaining eight subclusters each retains an equivalent R322 and R353, while one retains an equivalent R367 in addition, and five retain all four phosphate-binding arginine residues. 124 Figure 4.10 Sequence similarity network of the L-seryl-tRNA selenium transferase-like family (IPR018319). Each node represents groups of sequences with minimum sequence identities of 50 % over more than 80 % of their sequence and edges represent alignment scores between representative sequences for each node. The network groups into four main clusters at an alignment score threshold of 40 (AST = 40), which are defined by the characterized enzymes in this family: SelA (red), putative archaeal SelA (purple), DGL/Cap15 (orange) and one cluster that has no characterized members (dark blue). Raising the AST to 76 separates the nodes in the DGL/Cap15 cluster into two new subclusters, alongside several small groups of nodes (not shown). Cap15 (hexagon) remains part of this new subcluster up to an AST of 108. Nodes in the DGL/Cap15 subcluster are coloured according to the groups that remain connected at an AST greater than 109. The nodes most closely related to Cap15 (light blue) each represent homologues part of capuramycin-like gene clusters. Each gene cluster encodes a CapA and CapH homologue, two necessary enzymes involved in the biosynthesis of the Cap15 substrate. 125 Table 4.5 DGL/Cap15-like cluster active site sequence motifs. Each sequence logo depicts the percent conservation by residue and consensus residues of a multiple sequence alignment between sequences of the subcluster or the entire cluster. The height of each character column is determined by the maximum percent conservation of the most conserved residue. Subcluster Nseq 177-SxTH-180a 314-DxxGRxIxR-318 R353 365-DPR-367 All sequences 631 1 215 DGL-like 81 3 126 4 9 5 8 Cap15-like 9 7 143 8 5 9 18 10 7 11 6 a The R178 position is omitted from the Cap15 subcluster sequence logo for visual clarity. 126 4.4 Discussion Consistent with the relatively high sequence identity to DGL (41 %), the Cap15 structure superposes well onto DGL with RMSD values of 0.73 Å (NC𝛼 = 228) and 0.86 Å (NC𝛼 = 82) for the major and minor domains, respectively. Superposing the major domains of the DGL and Cap15 dimer shows that the Cap15 minor domain is rotated towards the active site by a median of 4.5 Å per residue compared to the DGL minor domain. The smallest relative displacements are around residues contacting the major domain (residues 344-351, 2.1 Å average) and the largest displacement at distal residues (residues 294-310, 6.4 Å average). While the B-factors of the minor domain in Cap15 are high, the position of the domain may represent a more closed conformation of the enzyme. Outside of the well conserved PLP-binding residues, there are two additional groups of conserved residues that may be involved in substrate binding. The short loop region after 𝛽5 (residues 177-SRTH-180), and the group of four arginine residues (R318, R322, R353, R367) involved in binding the phosphate anion are in the active site cleft and widely conserved in the DGL-Cap15 cluster identified in the sequence similarity network (AST=76). S177, H180, D314, R322, R353, and D365 are all conserved at rates greater than 95 % among all sequences in the cluster. D314 and D365 flank R322 and may serve to balance the densely packed positive charges of the nearby arginine residues. R318 and R367 are conserved in at least 71 % of sequences and are mainly found in six of the eleven subclusters. Phillips et al propose that the equivalent phosphate-binding arginine residues in DGL coordinate its D-glucosaminate-6-phosphate substrate phosphate moiety.249 A similar two residue phosphate-binding motif can be observed in Salmonella enterica L-threonine-O-3-phosphate decarboxylase (CobD, 1LC8);273 however, it incorporates the conserved active site arginine 127 residue which typically recognizes the substrate 𝛼-carboxylate in other AAT family enzymes not found in Cap15.274 Other AAT family enzymes missing this conserved arginine residue are found to have an arginine at the equivalent position to Cap15-R353. For example, in DesI (2PO3),275 GDP-perosamine synthase (3DR7)276 and ColD (3B8X),277 R327, R315 and R331, respectively, interact with their substrate’s pyrophosphoryl groups. ArnB (4OCA)278 and PseC (2FNU)279 have a histidine at this position which also binds their substrate’s pyrophosphoryl group. Our docking results suggest two possible modes of external aldimine substrate orientation: one where these phosphate-binding arginine residues along with S177 recognize the GlyU uracil ring and another where they bind a phosphate anion which interacts with the GlyU ribose hydroxyls. It is likely that these residues are necessary either for substrate recognition or for phosphate binding in the active site. In the case of phosphate binding the anion may be required to stabilize the minor domain’s position above the active site, as in vitro activity is only observed in phosphate buffers.238 H180, part of the short sequence of residues after 𝛽5 (177-SRTH-180) is well conserved among all members of DGL/Cap15-like enzymes. In several other AAT-like enzymes this loop is not part of the active site, as seen in SelA (3WCN),248 DesI (2PO3),275 PseC (2FNU),279 GDP-perosamine synthase (3DR7),276 ColD (3B8X),277 and ArnB (4OCA).278 Several other enzymes incorporate this loop into their active site, such as cystathionine gamma synthase (1QGN),280 cysteine desulfurase (6MR2),281 O-acetylhomoserine sulfhydrylase (3RI6),282 Plu4 (7N79)262 and MppP (5DJ1).228 In these enzymes an asparagine aligns with H180 structurally and is positioned in the active site for substrate interactions and/or hydrogen bonding to the PLP hydroxyl moiety. However, this asparagine is bookended by two proline residues and as a result the loop takes a much different path through space compared to Cap15 bringing it much closer to the PLP 128 moiety. In phosphoserine aminotransferase (5YB0)283 the equivalent loop is part of the active site and enters far further than the Cap15 loop. T148 structurally aligns with H180 in this case and hydrogen bonds to the PLP hydroxyl. Ultimately, H180 may serve similar roles in the active site of Cap15. However, recent spectroscopic experiments on mutants of the equivalent position in DGL (H163A) have suggested that the residue is a stereospecific proton donor.284 While H180 does not seem positioned to act as an acid or base based on our docking results, it may be favourably positioned to interact with an oxidized PLP intermediate based on its proximity to the predicted external aldimine C4ʹ and C𝛼 atoms; two likely oxidized, electron-rich positions.221 With the addition of Cap15, native oxygen- and PLP-dependent enzymatic activity has now been observed in four enzyme families: PLP-dependent decarboxylases (IPR002129), Class I/II or V aminotransferases (IPR004839, IPR000192), and SelA-like enzymes (IPR018319). Each are a subgroup of the AAT-like enzyme family and have unique active site arrangements making it difficult to determine a common mechanism of oxygen activation. However, a common theme in aromatic aldehyde synthases (AAS) mechanisms when comparing to the related decarboxylases is that residues implicated in proton transfer to the 𝛼-carbon are disrupted.221 Maintaining the imine at the 𝛼-carbon, rather than immediately reprotonating at that position, may render the intermediate susceptible to spontaneous oxidation. This would be consistent with the layout of the Cap15 active site, where there is no available protic side chain outside of K230 which likely initiates the catalytic cycle with the deprotonation of the 𝛼-hydrogen. Additionally, a protonated K230 may not be readily available for the immediate re-formation of the starting external aldimine as it will be attracted towards either the PLP phosphate moiety or the phosphate anion sequestered by residues R318, R322, R353 and R367. Based on in vivo gene knockouts and metabolic profiling, PvdN has been proposed to catalyze 129 decarboxylation-oxidation similar to Cap15, transforming a glutamate moiety to the corresponding succinimide in pyoverdine biosynthesis. However, without in vitro characterization, several issues remain mysterious: the origin of the oxygen atom of the amide, the timing of CO2 release, and whether hydrogen peroxide is produced. However, inspection of the published PvdN structure285 shows it bears greater resemblance to amino acid decarboxylases and their closely related oxygen- and PLP-dependent AASs. With the determined Cap15 structure and relevant active site residues identified through docking and sequence conservation I propose the following mechanism. GlyU displaces K230 and forms the external aldimine with the 𝛼-carboxyl group held out of plane via a hydrogen bond to T37. K230 deprotonates the 𝛼-hydrogen and may be ‘discouraged’ in re-protonating the substrate via its attraction to either of the nearby phosphate moieties. Oxygen diffuses into the active site and a single electron is transferred to its triplet state and may be protonated by H180 or stabilized by its positive charge. The peroxide radical then rebounds, forming a bond to the 𝛼-carbon. The consequent 𝛼-peroxyl intermediate decarboxylates forming a carbonyl in a fashion similar to hydrogen peroxide induced decarboxylation of 𝛽-keto acids.286–289 The release of hydroxide may be facilitated by the proximate H180 as well. Hydrolysis or reformation of the internal aldimine releases CarU from the active site (Figure 4.11). 130 Figure 4.11 Proposed Cap15 mechanism after structural considerations. HN-O O P O--OO+HN-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262K230T37HOOO-HOOHNH2NHNOOHOHOHN-O O P O--OO+HN-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262K230T37HOOO-HONH3+RO2H180NHHNHN-O O P OOO+HN-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262K230T37HOOO-HONH3+RH180NHHNH180NHHNO OHCO2HN-O O P O--OONH2-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262K230T37HOHONH3+RH180NHHNOOHN-O O P O--OO-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262K230T37HONH3+H180NHHN+H2NOOOHR COO-H+HN-O O P O--OO+HN-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262K230T37HOOO-HONH3+RH180NHHNOHOHN-O O P O--OO+HN-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262K230T37HOOHONH3+RH180NHHNO2electron transferC𝛼 rebound C4ʹ reboundH+CO2H2OHN-O O P O--OONH2-O OD206 HOY131Rʹ259H2N+H2NNH3+Kʹ262T37HOHORH180NHHNO+HNK230reforminternalaldiminereforminternalaldimineH2O131 In summary, I report the holo structure of the PLP-dependent monooxygenase-decarboxylase, Cap15, responsible for forming the characteristic carboxamide moiety of the capuramycin nucleoside core. The presence of a phosphate anion bridging the flexible C- and N-terminal domains provides an explanation for the unusual phosphate dependence previously observed, and sequence comparisons show the residues contacting the phosphate are widely conserved in hundreds of SelA-like family enzymes including the structurally related DGL. Flexible docking of the substrate (5ʹS, 6ʹR)-GlyU external aldimine versus the non-substrate 6ʹ epimer convincingly explain the observed stereospecificity. Cap15 joins a handful of other oxygen and PLP-dependent with experimentally determined structures and will serve as part of the foundation for future biochemical investigations into the residue determinants of oxygen consumption, hydrogen peroxide formation and substrate decarboxylation in these enzymes. 4.5 Experimental procedures 4.5.1 Purification of Cap15 Plasmids of pET28a-cap15 were transformed into chemical competent E. coli BL21(DE3) and grown in LB medium (37℃, 180 rpm) with kanamycin (50 𝜇g/mL) to an OD600 of 0.6 before cooling to 16 ℃ (30 min). Cultures were induced with 100 𝜇M β-D-1-thiogalactopyranoside and grown for a further 18 h. Cells were pelleted by centrifugation (4 ℃, 10 min, 3000 × g), and resuspended in 50 mM potassium phosphate (pH 8), 500 mM NaCl, 5 mM imidazole, 10 % glycerol and 0.5 mM dithiothreitol binding buffer. The suspension was lysed via sonication (20 % amplitude, 4 s on / 8 s off cycles, 6 min), and cell debris removed with centrifugation (45 min, 15000 × g). The supernatant was loaded onto Ni-IDA resin and eluted using a 20-500 mM imidazole step gradient in, and elution fractions containing recombinant Cap15 (200-300 mM) were combined and dialyzed against a 1 L solution of 20 mM 132 potassium phosphate, 50 mM NaCl and 0.5 mM PLP for 16 h at 4℃. Cap15 was further purified via size exclusion chromatography (20 mM potassium phosphate pH 8, 50 mM NaCl), and pure fractions as assessed by SDS-PAGE were combined for use in crystallography trials. 4.5.2 Size exclusion chromatography Samples were concentrated to 2 or 5 mL before loading onto a HiLoad® 16/600 Superdex® (Cytiva) gel filtration column equilibrated with appropriate buffer (typically 20 mM Tris, HEPES or potassium phosphate pH 7-8 containing 50 mM NaCl), and eluted at a flow rate of 1 mL/min. Analytical samples of blue dextran (2000 kDa), sweet potato β-amylase (200 kDa), yeast alcohol dehydrogenase (150 kDa), bovine serum albumin (66 kDa), bovine carbonic anhydrase (29 kDa), and horse cytochrome c (12.4 kDa) were used to calibrate the column for the estimation of molecular weights. 4.5.3 Crystallization Cap15 protein solutions were concentrated via centrifugation using 10 kDa filters to concentrations ranging from 10-15 mg/mL and screened with commercially available conditions (Index, Hampton, MCSG) in a 96-well sitting-drop format (1 µL protein : 1 µL crystallization solution, 50 µL reservoir solution). Initial conditions were optimized in hanging drop screens with drops of 1-2 µL of protein and 1-2 µL crystallization solution. Diffraction quality crystals grew at 0.2 M sodium acetate and 17 % polyethylene glycol 3350 at 10-15 mg/mL Cap15 in a 2:1 protein solution : crystallization solution drop ratio. Uridine, CarU and acetal-protected CarU were co-crystallized with Cap15 by added each to protein solutions as necessary to concentrations of 1-10 mM. CarU and related synthetic intermediates were prepared as previously described (18). Cap15 crystals were soaked with CarU and acetal-protected CarU up to five days at 0.7-1 mM or 10-90 minutes at 4-10 mM. Prepped 133 crystals were then washed through a cryoprotectant solution consisting of the well solution made with 16 % (v/v) ethylene glycol for 1-5 seconds before flash freezing in liquid nitrogen or 100 K gaseous nitrogen and stored for data collection at a synchrotron facility. 4.5.4 Data collection, structure determination and refinement X-ray diffraction dataset were collected at the Stanford Synchrotron Radiation Lightsource (SSRL, Menlo Park, United States) on beamline BL9-2 with a Pilatus 6M PAD detector. Before and after data collection UV-Vis spectra of crystals were recorded using a microspectrophotometer. Diffraction images were processed and integrated using XDS.157 Data were scaled and merged in AIMLESS,158 and phases determined by molecular replacement via Phaser220 using separate major and minor domains of DGL as search models (PDB: 7LCE). Chain A of DGL was split at the T266 nitrogen to produce each search model after removing water, ions, ligands and side chains and setting B-factors to 30 Å2. The top molecular replacement solution was improved upon through cycles of manual modelling with Coot161 and refinement using phenix.refine.163 4.5.5 Molecular model preparation and docking The Cap15 dimer (chains A and C) had all solvent removed and polar hydrogens added using AutoDock Tools.290 Similarly, (5ʹS, 6ʹR)-GlyU and (5ʹS, 6ʹS)-GlyU were created based on the uridine ideal ligand coordinates deposited in the PDB (URI). These ligand models’ geometry was adjusted using eLBOW and its simple geometry optimization setting.166 The external aldimines were built from joining these GlyU models with the pyridoxal 5-phosphate moiety from the internal aldimine modelled into the Cap15 active site. (5ʹS, 6ʹR)-GlyU was docked into the active site using AutoDock Vina164 with and without the phosphate anion. Up to six residues 134 (Q33, S36, T37, S177, T179 and H180) were allowed to move freely in the active site during docking. The (5ʹS, 6ʹR)-GlyU and (5ʹS, 6ʹS)-GlyU external aldimines were modelled as flexible residues where the PLP pyridinium ring and phosphate moieties were fixed in place where they are found in the internal aldimine. Either a water molecule or phosphate anion was docked into the active site to allow Vina to perform a conformational search on the external aldimine. This does not allow for direct scoring of the external aldimine and serves as a qualitative assessment of possible conformers under these restraints. K230 was always allowed to move freely during docking and up to three additional residues (S177, T179 and H180) were allowed to move freely during docking. 4.5.6 Sequence analysis Sequence similarity networks were made using EFI-EST291 using the Uniref50292 representation of the SelA-like enzyme family (IPR018319). An initial alignment score threshold of 40 was selected to visualize the basic clustering of the network. The threshold was increased to 76 to visualize sequences most related to Cap15 and DGL, and Clustal Omega169 was used to generate a multiple sequence alignment of these sequences. Sequence logos were generated from this alignment via Jalview293 per the clusters at an alignment score threshold of 110. Networks were analyzed and visualized using Cytoscape.294 135 Chapter 5: Conclusions 5.1 Detours in bacterial bisindole biosynthesis Bacterial bisindole NPs have attracted much attention for their anti-cancer properties, exemplified by the indolocarbazoles staurosporine and rebeccamycin.20 Screening campaigns for bioactive molecules led to the discovery of a wide array of indolocarbazoles, featuring varying halogenation, hydroxylation, oxidation, methylation and glycosylation patterns, launching many clinical trials, ultimately culminating in the approval of the semi-synthetic midostaurin for the treatment of neurological cancers.185 Investigations of the biosynthetic pathways to these indolocarbazoles found that they share common steps.19 Most bacterial bisindole assembly pathways proceed through the C𝛽 dimerization of two 3-indolepyruvic acid imines, catalyzed by iron-dependent chromopyrrolic acid synthases (CPAS). In the early 2010s, Brady and coworkers used this insight to identify many new bacterial bisindole scaffolds by developing degenerate primers to probe for CPASs in environmental DNA.32 They validated a connection between CPAS primary sequence diversity and bacterial bisindole scaffold diversity, leading to the discovery of gene clusters for new bisindolylmaleimides, indolotryptolines and indolocarbazoles.32,44,28,184 The small biosynthetic gene cluster of reductasporine (red), isolated from degenerate primer screening of Sonoran Desert eDNA samples by Brady and coworkers, is a two-gene deviation from the typical indolocarbazole gene cluster.184 The two additional genes, redE and redM, encode enzymes that catalyze formation of a reduced and dimethylated pyrrole core, a pathway unlike any other previously characterized indolocarbazole NP. In my work, the reconstitution of reductasporine biosynthesis in vitro demonstrated that the dehydrogenase RedE 136 is an NADPH-dependent imine reductase (IRED), and its X-ray crystal structure reveals that it has two distinct indolocarbazole binding sites. Site-directed mutagenesis at each site demonstrates that both active sites are essential for production of reductasporine, as demonstrated through tandem assays with RedE variants, where the enzymes StaP and RedM are employed to provide the RedE substrate and enable detection of the reductasporine product, respectively. Furthermore, my analysis of StaP reaction products on time scales less than 30 minutes reveal a short-lived indolocarbazole decomposition product consistent with indolo[2,3-a]pyrrolo[3,4-c]carbazole, lending credence to the proposal that this molecule is the substrate to RedE. The secondary binding site seems to have evolved to bind this unstable intermediate and shelter it from oxidation and possibly control tautomerism, before shuttling the substrate to the primary active site for reduction. Future work on RedE can be directed toward testing for the sequence determinants that form the secondary active site via site-directed mutagenesis and engineering those insights into other IREDs and reductive aminases in order to expand their substrate scopes to larger, more complex molecules with importance for pharmaceutical applications. Characterization of the SAM-dependent methyltransferase RedM, the other unique enzyme encoded by the reductasporine biosynthetic cluster, via X-ray crystallography shows it belongs to Class I COMT-type methyltransferases. It adopts clear open and closed conformations when bound with SAH and SAM, respectively. Site-directed mutagenesis of the active site demonstrates that several residues are responsible for successful dimethylation of the RedM substrate. Sequence analysis in the COMT-type enzyme family shows commonalities in these residues and likely correspond to substrate binding residues. These patterns are hydrophobic amino acid motifs including methionine, phenylalanine and tyrosine associated with binding 137 aromatic substrates. Fittingly, most substrates of the COMT-type enzyme family have at least one aromatic moiety. Site-directed mutagenesis does not identify a basic residue in the active site. Such a residue would be responsible for direct substrate deprotonation ahead of either methylation event, initiating . However, mutation of Y142 does reduce activity substantially. Thus, I suggest RedM catalyzes methylation primarily through substrate orientation, proximity and desolvation effects, exploiting the moderate pKa of pyrroline. Future work can focus on introducing single residue mutations that subtly shift the steric environment around the substrate in order to enrich for a singly methylated product. This would inform engineering efforts that want to introduce multi-methylating activity in related enzymes. Figure 5.1 A proposed biosynthetic pathway to erdasporines. EspM is presumed to intercept a singly carboxylated indolocarbazole intermediate that results from spontaneous decarboxylation of the indolocarbazole synthase (EspP) product. 138 Biosynthetic studies on indolocarbazoles have been ongoing for more than 20 years, but there are still many open questions with regard to bacterial bisindoles that could be the starting point for future studies in this area. In particular, several indolocarbazoles have unusual, uncharacterized biosynthetic steps. For example, erdasporine is an indolocarbazole with a methyl ester at the maleimide ring.44 The erdasporine gene cluster (esp) encodes the methyltransferase EspM. This enzyme presumably intercepts a non-oxidized singly decarboxylated indolocarbazole synthase product and generates a methyl ester, preventing further decarboxylation (Figure 5.1). Thus, EspM may play a similar role as RedE, StaC and RebC in that it intercepts a reactive bisindole intermediate as a substrate, stabilizing it for downstream tailoring. Further structural and mechanistic studies on EspM may reveal any unusual strategies that have evolved to accomplish this unwieldy transformation cleanly. Figure 5.2 Select spiroindimicins and indimicins isolated from marine actinomycetes. The C2-C3 bond in the indimicins are coloured red to highlight the apparent C-methylation and reduction of the indole compared to the potential biosynthetic precursors CPA and carboxyl-indolocarbazole. Another interesting set of bisindoles are the spiroindimicins295,296 and the indimicins,297 which are produced by Streptomyces marincola and related marine actinomycetes (Figure 5.2).40 Both groups of NPs are isolated from single species cultures, and although limited biosynthetic studies have been performed on this strain, it is conceivable that the pathways may have a direct 139 biosynthetic connection. Indeed, the indimicins and spiroindimicins have an identified biosynthetic gene cluster (spm) associated with them; however, attempts to characterize the individual steps of the biosynthesis yielded no candidates for key transformations in producing these molecules.39 For these pathways, the authors propose that the gene cluster may be split into a second operon at a different location in the genome. The search for responsible genes may be challenging as the enzymes involved may be unique compared to characterized bisindole gene clusters. For indimicins A–D, there is an apparent reductive C-methyltransfer that occurs across the electron-rich indolocarbazole system (Figure 5.2). Indimicin E, the related bisindolylpyrrole, has a similar reduced and C-methylated indole and may arise from the same enzyme. C3-indole methylation or prenylation is an uncommon transformation14 and expanding the repertoire of indole derivatization via the discover and study of this potential methyltransferase and/or reductase pair is of interest. Spiroindimicins A and B are two distinct bisindole molecules with substantially rearranged carbon skeletons featuring spiro-carbons likely arising from CPA.39 The proposed biosynthetic pathway relies on oxidative indole-indole coupling traverse via methylated CPA derivatives (Figure 5.3). This proposed transformation is unique though with intriguing parallels to the C2ʹ-C2 coupling catalyzed by indolocarbazole synthases. It is yet to be determined whether a single enzyme produces the scaffolds for spiroindimicin A and B non-specifically or whether each has a bespoke enzyme. 140 Figure 5.3 Proposed biosynthetic transformation required to yield spiroindimicins.39 A final series of compounds that may be interesting to study are the cyanobacterial indolocarbazoles, tjipanazoles.298 The tjipanazoles differ from other indolocarbazoles discussed above in that they lack a pyrrole or maleimide moiety (Figure 5.4). Though cyanobacterial biosynthetic pathways can be distinct from actinobacterial pathways, the analysis of the genome of Fischerella ambigua revealed a candidate bioynthetic gene cluster with rebO, rebD, rebP, and rebC gene homologues corresponding to indolocarbazole aglycone biosynthesis in rebeccamycin-producing strains.299 This suggests that tjipanazoles may traverse through a typical indolocarbazole biosynthetic pathway before disassembling a pyrrole or maleimide moiety. The cluster encodes the unique genes tjp3-6, which may be necessary for this disassembly process. Altogether, further study of bisindole biosynthesis is likely to continue to reveal unique biosynthetic enzymes for years to come. 141 Figure 5.4 Select tjipanazoles isolated from cyanobacteria Tolypothrix tjipanasensis. 5.2 Structural characterization of Cap15, a key enzyme in capuramycin biosynthesis Cap15 is an oxygen- and PLP-dependent enzyme, and differs from others in this functional class of enzymes, the arginine oxidases and aromatic aldehyde synthases, in that it incorporates oxygen into its product and does not produce hydrogen peroxide.55 In other words, it is a monooxygenase, not an oxidase. I have solved the crystal structure of Cap15 responsible for the oxidative-decarboxylation of amino acid nucleoside GlyU. The PLP-bound structure shows a conspicuous area of electron density bracketed by several arginine residues that is well modelled by a phosphate anion. Phosphate is necessary to observe measurable in vitro activity, and its presence in the active site suggests the anion plays a structural role in completing active site. Modelling the external aldimine into the active site highlights a four-residue loop with sequence SRTH that may be intimately involved in substrate binding and/or catalysis. Wider sequence analysis of this motif in the SelA enzyme family show that it is exclusively found in gene products produced in capuramycin-like biosynthetic gene clusters. Future work should focus on synthesizing the GlyU substrate in order to proceed with assaying single residue mutants, with 142 special attention on residues from the SRTH motif and phosphate-binding arginine residues. These should be accompanied by experiments that test the necessity of the phosphate ion via substitution with similar anions (e.g. sulphate) and assessing the stoichiometry of the interaction. Naturally, Cap15 crystals should again be screened for co-crystallization with substrate GlyU. There are several key open questions in families of PLP-dependent enzymes with dedicated oxygen-consuming activity that could be studied once these proposed studies on Cap15 with the GlyU substrate are carried out. First, the most similar enzyme to Cap15, the homologue DGL249 (41 % sequence identity), catalyzes a water elimination reaction of a phosphorylated sugar substrate and characterizing the chemistry any intermediate homologues may inform on the constituents of oxygen-consuming activity in Cap15. Perhaps even allowing for the engineered interconversion between Cap15-like and DGL-like activity. Second, I note that the pyoverdine biosynthesis protein PvdN300 catalyzes a similar decarboxylation-oxidation reaction in the conversion of a 𝛾-glutamyl ester in pyoverdine biosynthesis to a succinamyl ester; however, its crystal structure is more similar to decarboxylases related to aromatic aldehyde synthases.301 A detailed in vitro characterization of the PvdN enzyme would establish where it lies in the context of oxygen-consuming and PLP-dependent enzymes. Indeed, it would need to include establishing whether its activity depends on oxygen at all, then measure the stoichiometry of oxygen consumption and corresponding production of products (including carbon dioxide, hydrogen peroxide or otherwise), as well running isotope experiments to establish the origin of the oxygen incorporated into the amide. Altogether, these types of experiments may establish a connecting paradigm between oxidase and monooxygenase activity in this class of enzymes. 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Thesis/Dissertation
eng
Chemistry
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Ryan, Katherine S.
Crystallographic and biochemical characterization of key steps in reductasporine and capuramycin biosynthesis
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