Science, Irving K. Barber Faculty of (Okanagan)
Chemistry, Department of (Okanagan)
DSpace
UBCO
Bystrom, Liam
Master of Science - MSc
University of British Columbia
Fusobacterium nucleatum is a Gram-negative obligate anaerobe and a normal constituent of the human oral cavity. Additionally, F. nucleatum can disseminate through the bloodstream of its host and colonize distal sites including tumors of the colon where it can exacerbate disease. In the host, F. nucleatum encounters oxygen as well as derivative reactive oxygen and nitrogen species, but there is limited information on oxidative and nitrosative defense mechanisms present in F. nucleatum. Examination of F. nucleatum subsp. polymorphum’s genome revealed the presence of a putative flavodiiron protein (FDP) which is known to reduce O₂ and/or NO to H₂O or N₂O, respectively. Colocalized with the FDP gene is a gene encoding a multicomponent enzyme called butyryl-CoA reductase (BCR). BCR contains three distinct domains: butyryl-CoA dehydrogenase (BCD), the C-terminal domain of the α-subunit of an electron transfer flavoprotein (ETFα), and rubredoxin (Rd). This thesis details work which shows that BCR functions to oxidize butyryl-CoA and transfer the reducing equivalents to the FDP for reduction of O₂ to H₂O. Electron transfer is dependent on the formation of a neutral semiquinone of the FAD bound to BCD and an anionic semiquinone of the FAD bound to ETFα. This enzyme pair gives F. nucleatum the ability to use a common metabolite (butyryl-CoA) with a high reduction potential to reduce O₂ and prevent oxidative stress. By using butyryl-CoA as reductant, F. nucleatum can derive additional energy from its amino acid-based metabolism while simultaneously removing toxic O₂. The novel BCR-FDP is one of the only enzymes characterized from F. nucleatum to function in oxidative stress defense, making it a potentially critical enzyme for F. nucleatum’s survival and subsequent infection. Bioinformatic analysis revealed that BCR-FDP is prevalent in many species of anaerobic gut bacteria where it may function in maintaining anaerobiosis and therefore a healthy gut microbiome.
i COUPLING BUTYRYL-COENZYME A OXIDATION TO OXYGEN REDUCTION IN FUSOBACTERIUM NUCLEATUM By Liam Bystrom B.Sc., The University of British Columbia (Okanagan), 2022 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR MASTER OF SCIENCE in THE COLLEGE OF GRADUATE STUDIES (Biochemistry and Molecular Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Okanagan) November 2024 © Liam Bystrom, 2024 ii The following individuals certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis entitled: COUPLING BUTYRYL-COENYZME A OXIDATION TO OXYGEN REDUCTION IN FUSOBACTERIUM NUCLEATUM Submitted by Liam Bystrom in partial fulfillment of the requirements of the degree of Master of Science Dr. Kirsten Wolthers, Irving K. Barber Faculty of Science, University of British Columbia, Okanagan Supervisor Dr. Stephen McNeil, Irving K. Barber Faculty of Science, University of British Columbia, Okanagan Supervisory Committee Member Dr. Richard Plunkett, Irving K. Barber Faculty of Science, University of British Columbia, Okanagan Supervisory Committee Member Dr. Michael Deyholos, Irving K. Barber Faculty of Science, University of British Columbia, Okanagan University Examiner iii Abstract Fusobacterium nucleatum is a Gram-negative obligate anaerobe and a normal constituent of the human oral cavity. Additionally, F. nucleatum can disseminate through the bloodstream of its host and colonize distal sites including tumors of the colon where it can exacerbate disease. In the host, F. nucleatum encounters oxygen as well as derivative reactive oxygen and nitrogen species, but there is limited information on oxidative and nitrosative defense mechanisms present in F. nucleatum. Examination of F. nucleatum subsp. polymorphum’s genome revealed the presence of a putative flavodiiron protein (FDP) which is known to reduce O2 and/or NO to H2O or N2O, respectively. Colocalized with the FDP gene is a gene encoding a multicomponent enzyme called butyryl-CoA reductase (BCR). BCR contains three distinct domains: butyryl-CoA dehydrogenase (BCD), the C-terminal domain of the a-subunit of an electron transfer flavoprotein (ETFa), and rubredoxin (Rd). This thesis details work which shows that BCR functions to oxidize butyryl-CoA and transfer the reducing equivalents to the FDP for reduction of O2 to H2O. Electron transfer is dependent on the formation of a neutral semiquinone of the FAD bound to BCD and an anionic semiquinone of the FAD bound to ETFa. This enzyme pair gives F. nucleatum the ability to use a common metabolite (butyryl-CoA) with a high reduction potential to reduce O2 and prevent oxidative stress. By using butyryl-CoA as reductant, F. nucleatum can derive additional energy from its amino acid-based metabolism while simultaneously removing toxic O2. The novel BCR-FDP is one of the only enzymes characterized from F. nucleatum to function in oxidative stress defense, making it a potentially critical enzyme for F. nucleatum’s survival and subsequent infection. Bioinformatic analysis revealed that BCR-FDP is prevalent in many species of anaerobic gut bacteria where it may function in maintaining anaerobiosis and therefore a healthy gut microbiome. iv Lay Summary Molecular oxygen (O2) is toxic to many bacteria, including Fusobacterium nucleatum, a common inhabitant of the oral cavity. This organism has become the subject of recent research due to its ability to migrate through the bloodstream, colonize colorectal cancer tumors, and worsen patient prognosis. Little is known about how F. nucleatum mitigates against O2 toxicity in the oral cavity and throughout its human host. Herein, we discovered and characterized a new enzyme (biological catalyst) from F. nucleatum that reduces O2 to water (H2O), with the aid of a common metabolite. Thus, this new enzyme is likely central to F. nucleatum’s survival in the human host, including in tumors, where it can negatively impact human health. However, analysis of the genomes of other bacteria reveal that this enzyme is prevalent in many beneficial bacteria of the colon. This new enzyme may also help maintain an O2-free environment in the colon, which is beneficial to human health. v Preface This research project was conceived by Dr. Kirsten Wolthers. The experimental data and data analysis was performed by Liam Bystrom while under the supervision of Dr. Kirsten Wolthers. The results presented have been published in the following paper: Bystrom, L. T., and Wolthers, K. R. New Electron-Transfer Chain to a Flavodiiron Protein in Fusobacterium nucleatum Couples Butyryl-CoA Oxidation to O2 Reduction. Biochemistry, 63, 2352-2368. (2024). No generative, artificial intelligence was used in the research described herein or in the drafting and preparation of this thesis. vi Table of Contents Abstract .......................................................................................................................................... iii Lay Summary ................................................................................................................................ iv Preface ............................................................................................................................................. v Acknowledgements ........................................................................................................................ xv Chapter 1: Introduction ................................................................................................................. 1 1.1 Anaerobic bacteria and oxygen ....................................................................................... 1 1.1.1 The Great Oxidation Event ............................................................................................ 1 1.1.2 ROS/RNS interfere with anaerobic metabolism ............................................................ 2 1.1.3 Anaerobic bacteria and the oxygen response ................................................................. 5 1.2 Flavin cofactors ..................................................................................................................... 8 1.2.1 UV-visible spectroscopy properties of flavins ............................................................... 9 1.3 Fusobacterium nucleatum ................................................................................................... 10 1.3.1 Amino acid metabolism of F. nucleatum ..................................................................... 11 1.3.2 F. nucleatum in the oral cavity and dissemination ....................................................... 14 1.3.3 F. nucleatum and colorectal cancer .............................................................................. 15 1.4 Flavodiiron proteins ............................................................................................................ 18 1.4.1 Flavodiiron proteins as oxidative stress-clearing enzymes .......................................... 20 1.4.2 Mechanism of flavodiiron proteins .............................................................................. 21 1.4.3 Classification of flavodiiron proteins ........................................................................... 24 1.4.4 Diiron centre motif in proteins ..................................................................................... 25 1.6 The electron transfer partner of FDP in F. nucleatum ......................................................... 26 1.7 Butyryl-CoA dehydrogenase ............................................................................................... 27 1.8 Electron transfer flavoproteins ............................................................................................ 27 1.9 Rubredoxins ........................................................................................................................ 28 1.10 Research objectives ........................................................................................................... 29 Chapter 2: Coupling oxidation of butyryl CoA to oxygen reduction through BCR-FDP ......... 30 vii 2.1 Chapter summary ................................................................................................................ 30 2.2 Materials and methods ........................................................................................................ 30 2.2.1 Materials ...................................................................................................................... 30 2.2.2 Data analysis ................................................................................................................ 30 2.2.3 Cloning of BCR, BCR-FDP, BCD, and ETFa ............................................................ 31 2.2.4 Protein expression ........................................................................................................ 32 2.2.5 Aerobic protein purification ......................................................................................... 33 2.2.6 Anaerobic protein purification ..................................................................................... 34 2.2.7 SDS-PAGE ................................................................................................................... 34 2.2.8 Size exclusion chromatography ................................................................................... 34 2.2.9 UV-visible spectroscopy .............................................................................................. 35 2.2.10 Protein quantification and extinction coefficients ..................................................... 35 2.2.11 Flavin analysis and quantification .............................................................................. 35 2.2.12 Inductively coupled plasma mass spectrometry (ICP-MS) ....................................... 37 2.2.13 Reduction-oxidation (redox) titrations ....................................................................... 37 2.2.14 Butyryl-CoA synthesis ............................................................................................... 38 2.2.15 Stopped-flow measurements ...................................................................................... 38 2.2.16 Electron paramagnetic resonance (EPR) ................................................................... 39 2.2.17 Generation of nitric oxide .......................................................................................... 39 2.2.18 Measurement of O2, NO, and H2O2 activity .............................................................. 40 2.2.19 Bioinformatic analysis ............................................................................................... 41 2.3 Results ................................................................................................................................. 41 2.3.1 Analysis of cofactor iron-binding sites of BCR and FDP based on sequence homology and protein models ................................................................................................................ 41 2.3.2 Flavin binding sites of FDP, BCD, and ETFa ............................................................. 42 2.3.3 BCR and BCR-FDP purification and properties .......................................................... 45 2.3.4 BCR and BCR-FDP UV-vis spectroscopy ................................................................... 47 2.3.5 Cofactor incorporation in BCR and BCR-FDP ........................................................... 51 2.3.7 Anaerobic purification of ETFa .................................................................................. 54 2.3.8 Reduction oxidation titrations of BCD, ETFa, BCR, and BCR-FDP ......................... 57 viii 2.3.9 Pre-steady state kinetic analysis of butyryl-CoA’s reduction of BCD and BCR ......... 61 2.3.10 Catalytic activity of BCR-FDP with NO, O2, and H2O2 ............................................ 64 2.3.11 The prevalence of the BCR-FDP operon across bacterial species ............................. 68 2.4 Discussion ........................................................................................................................... 71 2.4.1 Cofactor binding sites and content of BCR-FDP ......................................................... 71 2.4.2 Characterization of the electron transfer pathway of BCR-FDP ................................. 73 2.4.3 Chemical modification of ETFa’s FAD ...................................................................... 74 2.4.4 Redox potentials of BCR-FDP’s cofactors .................................................................. 76 2.4.5 Importance of BCR-FDP to F. nucleatum .................................................................... 79 2.4.6 BCR-FDP in F. nucleatum’s metabolism ..................................................................... 80 2.4.7 Importance of butyrate to the gut microbiome ............................................................ 81 2.4.8 Anaerobicity of the gut microbiome ............................................................................ 83 Chapter 3: Conclusion ................................................................................................................. 85 3.1 Discovery and importance of BCR-FDP in F. nucleatum ................................................... 85 3.2 Proposed role of BCR-FDP in the gut microbiome ............................................................ 86 3.3 Future directions ................................................................................................................. 86 References .................................................................................................................................... 88 ix List of Tables Table 1: Primer sequences of each sequence of interest ............................................................... 32 Table 3: Ratios of cofactor to protein for BCR and BCR-FDP. Expected ratios are in parentheses................................................................................................................................................ 52 Table 3: Extinction coefficients corresponding to 280 nm and the maximum flavin peak for each protein ................................................................................................................................... 57 x Table of Figures Figure 1: ROS and RNS species form through reaction cascades…………………………. 3 Figure 2: Reaction mechanism of pyruvate formate lyase…………………………………. 5 Figure 3: Reaction mechanism of 2-hydroxyglutaryl-CoA dehydratase…………………... 6 Figure 4: An overview of the enzymatic oxidative stress response of anaerobic bacteria… 9 Figure 6: The different redox states of the flavin cofactor………………………………… 11 Figure 7: UV-vis spectra of free FAD and FMN…………………………………………... 13 Figure 8: Glutamate and lysine fermentation in F. nucleatum ……………………………. 16 Figure 9: Energy conservation using electron bifurcation and the Rnf complex………….. 17 Figure 10: F. nucleatum’s role in aggravating colorectal cancer…………………………... 19 Figure 11: A summary of known enzymatic oxidative stress defenses in F. nucleatum…... 22 Figure 12: Tertiary fold motifs of E. coli’s FDPs………………………………………….. 23 Figure 13: Head-to-tail dimer structure of FDPs…………………………………………... 24 Figure 14: The O2 and NO reduction mechanisms of FDPs……………………………….. 27 Figure 15: The classes of FDPs……………………………………………………………. 29 Figure 16: A model of BCR coupled to FDP. , …………………………………………..…31 Figure 17: Canonical vs bifurcating ETFs…………………………………………………. 33 Figure 18: The iron binding sites of BCR and FDP……………………………………….. 47 Figure 19: AlphaFold model of FDP compared with the flavodiiron protein from E. coli... 48 Figure 20: A comparison of the AlphaFold model of BCD with butyryl-CoA dehydrogenase from A. fermentans.................................................................................................................49 Figure 21: A comparison of the AlphaFold model of ETFa with the electron transfer flavoprotein from P. denitrificans………………………………………………………….. 50 Figure 22: SDS-PAGE analysis of BCR-FDP and BCR....................................................... 51 Figure 23: UV-visible spectra of BCR and BCR-FDP grown in LB and M9 minimal media......................................................................................................................................53 Figure 24: Optical spectra of BCR......................................................................................... 55 Figure 25: EPR spectrum of BCR purified aerobically and anaerobically............................ 56 Figure 27: SDS-PAGE analysis of BCD and ETFa.............................................................. 59 Figure 28: UV-vis spectra of BCD and ETFa showing bound and unbound flavin............. 62 Figure 29: UV-vis spectra of various redox states of anaerobically purified ETFa.............. 63 xi Figure 30: Comparison of EPR spectra of aerobic and anaerobic ETFa.............................. 64 Figure 31: Redox titrations of BCD, ETFa, BCR, and BCR-FDP........................................ 71 Figure 32: EPR spectra of BCR-FDP taken at two different points in a redox titration........ 73 Figure 33: Stopped-flow data from reaction of BCD domain with butyryl-CoA.................. 78 Figure 34: Multiwavelength stopped-flow data of BCR and butyryl-CoA........................... 79 Figure 35: NO activity of BCR-FDP..................................................................................... 81 Figure 36: O2 reductase activity of BCR-FDP....................................................................... 82 Figure 37: The reaction of BCR-FDP with butyryl-CoA does not produce H2O2................. 83 Figure 38: The reaction of butyryl-CoA and BCR-FDP does not consume H2O2................. 84 Figure 39: Histogram of homologous sequences to BCR...................................................... 86 Figure 40: Prevalence of the BCR-FDP dicistron across bacterial species........................... 87 Figure 41: Mechanism of oxygen-dependent flavin modification......................................... 92 Figure 42: Reduction potential diagram of BCR-FDP.......................................................... 96 Figure 43: Two known aerobic respiratory chains present in anaerobes............................... 99 Figure 44: Butyrate inhibits histone deacetylases to influence gene expression................... 101 xii List of Symbols and Abbreviations °C degrees Celsius Å angstrom bp BCR FDP ETF⍺ BCD Rd BLAST PCR DNA ATCC E. coli LB Amp Kan OD IPTG FAD FMN PMSF SDS PAGE CV UV BSA NO HCl TFA base pairs butyryl-CoA reductase flavodiiron protein Electron transfer flavoprotein subunit ⍺ butyryl-CoA dehydrogenase Rubredoxin basic local alignment search tool polymerase chain reaction deoxyribonucleic acid American Type Culture Collection Escherichia coli Luria-Bertani ampicillin kanamycin optical density isopropyl ß-D-1-thiogalactopyranoside flavin adenine dinucleotide flavin mononucleotide phenylmethylsulfonyl fluoride sodium dodecyl sulfate polyacrylamide gel electrophoresis column volume ultraviolet bovine serum albumin nitric oxide hydrochloric acid trifluoroacetic acid xiii PDA kobs Kd ICP-MS kDa Sc Y In Tb PDB SSN GND AhpCF GPx SOD SOR NOX MsrAB IBD photodiode array observed rate constant dissociation constant inductively-coupled plasma mass spectrometry kilodaltons Scandium Yttrium Indium Terbium Protein Data Bank Sequence Similarity Network Genome Neighbourhood Diagram Alkyl hydroperoxide reductase Glutathione peroxidase Superoxide dismutase Superoxide reductase NADH oxidase Methionine sulfoxide reductase Inflammatory bowel disease ADP AMP NOS NIR ONOO- TCA Fld Adenine diphosphate Adenine monophosphate Nitric oxide synthase Nitrite reductase Peroxynitrite Tricarboxylic acid Flavodoxin FldOR ROR Rnf ETF-QO Fd Flavin oxidoreductase Rubredoxin oxidoreductase Rhodobacter nitrogen fixation Electron transfer flavoprotein ubiquinone oxidoreductase Ferredoxin xiv NROR NADH rubredoxin oxidoreductase xv Acknowledgements I am forever grateful to Dr. Kirsten Wolthers and the support, advice, and guidance you have given me throughout my degree. Your mentorship has provided me with direction going forward. Thank you for teaching me. I am deeply lucky to have gone through my graduate program with an amazing group of lab mates who have mentored me, helped me, and created the perfect environment to do science in. Thank you Amanda, Alex, Long, and Lexi for being fellow lab rats and friends. I would like to thank my committee members Dr. Richard Plunkett and Dr. Stephen McNeil for their support during my degree. Thank you to Dr. Brendan D’Souza for encouraging me and helping me develop as a teacher. I am also grateful for the help I have received from Dr. Wesley Zandberg, Dr. Frederic Menard, Dr. Isaac Li, Dr. Thuy Dang, and Dr. Conor Pranckevicius, especially for the gracious use of their laboratory space and equipment. I also wish to thank all the friends I have made in the graduate program at UBCO. Your assistance in things both scientific and not, helped created this thesis. Thank you to my friends who helped keep me sane despite a wide diaspora. I couldn’t ask for a better group of guys. Finally, I wish to thank my family for their unwavering support and love that they have showed me. I am here because of you. xvi To my parents, who have always supported me. Thank you for being the best parents I could have. I will never say that enough. “It’s the kind of thing that makes you glad stopped and smelled the pine trees along the way, you know?” 1 Chapter 1: Introduction The obligate anaerobe, Fusobacterium nucleatum, is a common member of the oral microbiota, but the bacterium is also found at extra-oral sites (tumors, abscesses, and amniotic fluid) where it is associated with a range of pathologies1–3. In these niches, F. nucleatum is regularly exposed to atmospheric oxygen as well as oxidative stress generated by the human immune system. However, F. nucleatum survives despite this. Unfortunately for its human host, F. nucleatum’s persistence can exacerbate disease (e.g. cancer, periodontitis, adverse pregnancy outcomes). F. nucleatum must possess defense mechanisms to mitigate against oxidative stress, but currently our knowledge of these systems is limited. Discovering new enzymatic mechanisms will deepen our understanding of how such an obligate anaerobe subsists in habitats where it encounters O2 and other reactive oxygen species (ROS). In the introduction of this thesis, I will discuss different types of ROS and reactive nitrogen species (RNS) and the enzymatic systems used by anaerobic bacterium to counteract them. I will discuss the importance of flavin cofactors to these enzymatic systems. Looking specifically at F. nucleatum, I will outline its oxidative stress response and its role in disease. I will explain the function, mechanism, and classification of flavodiiron proteins (FDPs) as F. nucleatum encodes for a putative FDP gene which may help combat oxidative stress. Finally, this introduction will discuss the FDPs colocalization with a novel modular butyryl-CoA which contains three distinct domains. This prompts an explanation of each domain. 1.1 Anaerobic bacteria and oxygen 1.1.1 The Great Oxidation Event Ancient Earth was an anoxic environment which gave rise to life. Approximately 2.5 billion years ago, oxygen levels began to increase in the atmosphere primarily due to the development of photosynthesis by cyanobacteria4,5. As oxygen levels increased and eventually stabilized, early organisms faced an evolutionary challenge. Some embraced oxygen, developing respiratory chains that used oxygen as a terminal electron acceptor. Other microbes did not change their metabolism, maintaining their anaerobic lifestyle. The prevalence of oxygen relegated 2 anaerobes to anoxic niches such as soil, sediment, and deep oceans. With the evolution of multicellular organisms, other anoxic niches were created for anaerobic microorganisms to inhabit. Two of these anoxic niches are the mammalian gut and the subgingival plaque of the oral cavity. This separation of aerobes and anaerobes in distinct ecological niches was framed by early microbiologists as a failure by anaerobes to evolve in the presence of oxygen6. However, many obligate anaerobes acquired mechanisms by which to mitigate against oxidative stress. These defense mechanisms enable anaerobes to inhabit environments where they can be transiently exposed to O2 and other ROS. 1.1.2 ROS/RNS interfere with anaerobic metabolism Oxygen can react with intracellular radicals like flavins and quinones to form superoxide (O2-) (Figure 1). O2- can react with iron-sulfur clusters and mononuclear iron centers by one-electron oxidation7. This leads to dissociation of the iron atom from the protein scaffold, yielding inactive enzyme. Common enzymes affected are dehydratases in the tricarboxylic acid (TCA) cycle (aconitase and fumarase) and those in branched-chain amino acid synthesis (homoaconitase and isopropyl malate dehydratase)7,8. O2- can easily cascade to form other ROS like the hydroperoxyl radical (HOO•) and H2O2 which can damage all major biological macromolecules9,10. For example, H2O2 has a propensity to react with ferrous iron through the Fenton reaction. The product of the Fenton reaction is OH• which oxidizes both DNA bases and ribose7. In addition to ROS, RNS also contribute to cellular damage. Nitric oxide (NO) is generated by the immune system by nitric oxide synthase (NOS) and by denitrifying bacteria through nitrite reductase (NIR)11,12. NO can then directly react with O2- to form peroxynitrite (ONOO-). ONOO- can then cascade to form peroxynitrous acid (ONOOH), nitrogen dioxide (NO2), and nitrous acid (HNO2). ONOO- can directly react with lipids, proteins, and DNA forming covalent adducts which are detrimental to biological function13. For example, the reaction of ONOO- with tyrosyl groups of signaling proteins involved in cell cycle regulation prevents phosphorylation. This arrests the signaling pathway14. Downstream products like ONOOH and HNO2 also react with functional groups to form covalent adducts. ONOOH reacts with thiol groups while HNO2 targets amine groups15. 3 Figure 1: ROS and RNS species form through reaction cascades. Starting from O2 and NO, a host of ROS and RNS can form through subsequent reductions and reactions. In addition to ROS and RNS, obligate anaerobes cannot tolerate molecular oxygen due to their specialized metabolism that relies on radical chemistry and low-potential metal centers to perform challenging chemical reactions. Two examples of oxygen-sensitive enzymes found in obligate anaerobes are pyruvate formate lyase and 2-hydroxyacyl-CoA dehydratase. Pyruvate formate lyase converts pyruvate to acetyl-CoA and formic acid. This reaction is part of central metabolism in a wide variety of anaerobes (Escherichia coli, Klebsiella pneumoniae, Enterococcus faecalis, Clostridium difficile, etc.).16–19 Pyruvate formate lyase allows for conversion of pyruvate to acetyl-CoA without generation of NADH. In fact, pyruvate formate lyase generates more ATP per mole of pyruvate than pyruvate dehydrogenase4. The pyruvate formate lyase catalytic cycle is dependent on a glycyl radical formed on a backbone glycine residue by a radical S-adenosyl-L-methionine (SAM) dependent activating enzyme (Figure 2)17. The glycyl radical generates a transient thiyl radical on a nearby cysteine which then serves to trigger carbon-carbon bond cleavage. The glycyl radical can persist for several days in vitro under anaerobic conditions because of captodative stabilization from the amino and carbonyl groups of the glycine residue, but it is highly reactive with O217. Electron transfer from the glycyl radical to O2 leads to cleavage of the polypeptide backbone and irreversible enzyme inactivation. 4 Figure 2: Reaction mechanism of pyruvate formate lyase. Active site glycine and cysteine residues participate in radical chemistry converting pyruvate to acetyl-CoA. The glycyl radical is initially formed by a SAM-dependent activating enzyme and its AdoMet radical. The glycyl radical is restored for subsequent catalytic cycles. 2-hydroxyacyl-CoA dehydratases are a crucial part of anaerobic amino acid fermentation, converting 2-hydroxyacyl-CoAs to enoyl-CoAs in Fusobacterium and Clostridium spp.20,21. These enzymes must catalyze the syn-elimination of water where a non-acidic b-Si-proton must be removed (pKa ~ 40) (Figure 3)22. This energetic barrier is circumvented by the formation of a ketyl radical intermediate by electron transfer from a low-potential iron-sulfur cluster. This radical species acts as a nucleophile to remove the OH group, lowering the pKa of the b-proton to 1422. This allows for a basic protein residue to extract the b-Si-proton, restoring the ketyl radical. The radical is oxidized to form the product with the electron being reused by the next substrate AdoMet Gly 5 molecule, acting as a cofactor. Radical formation is essential to this reaction, as the use of nearby polar residues would be unable to lower the pKa to the extent required. These two examples highlight the importance of radicals in anaerobic metabolism, helping to explain why O2 is toxic to obligate anaerobes. Figure 3: Reaction mechanism of 2-hydroxyglutaryl-CoA dehydratase. The 2-hydroxyacyl-CoA dehydratase mechanism is shown with this example from glutamate metabolism. An iron-sulfur cluster generates a substrate ketyl radical species which allows from subsequent syn-elimination. The radical is conserved in consecutive cycles through transfer onto the next incoming substrate molecule. 1.1.3 Anaerobic bacteria and the oxygen response O2 and related ROS pose a threat to all cells, in particular anaerobes. To preserve their specialized metabolism, anaerobes possess similar, even identical, oxygen defense mechanisms to aerobes. Enzymes such as catalase, alkyl hydroperoxide reductase (AhpCF), glutathione 6 peroxidase (GPx), SOD, SOR, NADH oxidase (NOX), and methionine sulfoxide reductase (MsrAB) can be found across anaerobic bacteria species (Figure 4)23–28. This array of enzymes argues against the narrative that anaerobic bacteria lack defenses against oxygen as they use a wide variety of enzymes to prevent cellular damage. Catalase, the most well-known oxidative stress response enzyme, is often not associated with anaerobic bacteria, but exceptions include Desulfovibrio vulgaris, Desulfovibrio gigas, B, fragilis, Methanosarcina barkeri, Methanobrevibacter arboriphilu and E. coli29. Those anaerobes that lack catalase consume H2O2 through other mechanisms. For example, AhpCF and GPx are two alternative catalase-type enzymes. AhpCF is a heterodimeric enzyme complex which uses NAD(P)H to reduce peroxyl groups on an array of biological compounds26. These peroxyl groups form on lipids and proteins from the reaction of alcohol groups with oxygen and ROS30. Once formed, they inhibit natural function or can cause protein misfolding. AhpC functions as a peroxidase and AhpF as an NAD(P)H reductase, passing electrons to AhpC. AhpCF has also been shown to directly reduce H2O2 to water in E. coli strains lacking catalase31. The extent to which AhpCF is present across bacteria is unknown, but it is found in several anaerobic species30,32. Its ability to replace catalase activity makes it potentially a vital enzyme for anaerobic bacteria. GPx, like AhpCF, reduces peroxyl groups that form on cellular metabolites when the cell is exposed to oxidative stress27. GPx relies on an intracellular pool of glutathione. GPx reduces peroxyl groups by oxidizing glutathione to form glutathione disulfide27. The disulfide is reduced by glutathione reductase (GSR) using NADH, restoring glutathione for further catalytic cycles. SOD and SOR are the primary enzymes used to reduce superoxide. SOD catalyzes the dismutation of O2- to H2O2 and O2. SODs have been primarily studied in eukaryotes, with these enzymes using a copper-zinc active site33. However, iron-based SODs (FeSODs) have been isolated from prokaryotes, including anaerobic bacteria34. FeSODs have been found in E. coli, Desulfovibrio desulfuricans, Chlorobium thiosulfatophilum, and some Clostridium species35–37. Even more widespread in anaerobes than FeSOD are SORs. These enzymes reduce O2- to H2O2, performing only the reductive half of the SOD reaction28. SORs, like SODs, are metalloproteins, binding iron for catalysis. They ultimately use NADH as a source of electrons, typically receiving reducing equivalents from an intermediate electron transport protein like rubredoxin (Rd)28. NADH oxidases (NOX) are employed by bacteria to reduce O2 to either H2O2 or H2O. This reaction stops oxidative stress at the source, preventing dangerous cascades through O2- and H2O2. 7 There are two forms of the enzyme, NOX-1 and NOX-2. NOX-1 converts O2 to H2O2 and NOX-2 directly reduces O2 to H2O25. NOX-1 and NOX-2 have low sequence homology38. The electrons from NADH are passed to flavin cofactors, either flavin mononucleotide (FMN) or flavin adenine dinucleotide (FAD), and then used to reduce O2. NADH oxidases have been identified in anaerobes including F. nucleatum, Lactobacillus brevis, Thermotoga maritima, and several Streptococcus species39–42. The thiol ether of methionine residues is susceptible to oxidation by ROS, primarily H2O224. The resulting methionine sulfoxide can cause protein misfolding or malfunction. To counter this, organisms use MsrAB, a heterodimeric enzyme complex which specifically acts on methionine residues that have oxidized to methionine sulfoxide. MsrA and MsrB reduce S-methionine sulfoxide and R-methionine sulfoxide groups respectively24. While the majority of species use distinct proteins for each form of the enzyme, some bacteria encode both enzymes as a fusion protein: MsrAB26. This enzyme is quite ubiquitous across prokaryotes and in found in both aerobes and anaerobes43. In summary, obligate anaerobes use a number of enzymatic systems to mitigate against oxidative stress and the inactivation of O2 sensitive enzymes. Figure 4: An overview of the enzymatic oxidative stress response of anaerobic bacteria. Anaerobes possess a wide array of enzymes to reduce O2, O2-, and H2O2. 8 1.2 Flavin cofactors Many enzymes that function to reduce O2 or ROS use flavin cofactors44. FMN and FAD are the two main types of flavin cofactors (Figure Figure 5 5). FMN is composed of an isoalloxazine ring bound to a ribitol group at the N10 position. The ribitol terminates in a phosphate group yielding a nucleotide45. FAD adds a secondary nucleotide from the first phosphate group with a cyclic ribose and adenine base (ADP). Out of all these building blocks, it is the isoalloxazine ring that is redox active, alternating between the quinone (FADox, FMNox), semiquinone (FADH•, FMNH•), and hydroquinone states (FADH2, FMNH2) (Figure 6)44. The semiquinone state can be stable without a proton, giving a negative charged anionic semiquinone species (FAD•-, FMN•-). The negative charge is stabilized by positively charged side chains of nearby amino acids46. Figure 5: Structures of flavin adenine dinucleotide and flavin mononucleotide. Flavins can function with one or two electron transitions. This dual property is controlled by the neighbouring protein environment which can act to stabilize or destabilize the semiquinone species. The protein environment surrounding the isoalloxazine ring controls the midpoint FMN FAD 9 reduction potential of the oxidized/semiquinone transition (E¢0 ox/sq) and semiquinone/hydroquinone transition (E¢0 sq/hq)45. For free flavin in solution, E¢0 ox/sq is more negative than E¢0 sq/hq: thus, the semiquinone does not accumulate and the flavin only functions in two electron transitions. However, many protein scaffolds can stabilize the semiquinone species, which inverts the reduction potentials such that E¢0 ox/sq is more positive than E¢0 sq/hq. Inversion of the potentials enables the flavin to function in one electron transfer. Figure 6: The different redox states of the flavin cofactor. Subsequent reductions from the quinone yield the semiquinone and hydroquinone species. Incoming electrons enter the system at the N5 position. 1.2.1 UV-visible spectroscopy properties of flavins The isoalloxazine ring of flavins is detectable using UV-visible spectroscopy. Flavins in the fully oxidized quinone state exhibit two bands labeled as bands I and II (Figure 7)44,46. Band II for free FMN and FAD is centered at 375 nm44. However, band I for FMN is centered at 446 nm while FAD’s band I is positioned at 450 nm. This difference in the absorbance maxima (lmax) of band I can be exploited to determine the unknown identity of flavin bound to protein after it is released from the protein through heat denaturation. The amplitudes of band I and II and their lmax can also vary once bound to protein due to the electronic effects of the nearby residues on the isoalloxazine ring. Upon reduction to either the neutral or anionic semiquinone, the detectable UV-H+ e- H+ e- e- H+ e- 5 10 Quinone Semiquinone Anionic Semiquinone Hydroquinone 2 H+ 10 visible peaks also change45. The transition to the neutral semiquinone is characterized by a decrease in the absorbance in band I (~450 nm) with an increase at 580 nm. On the other hand, the anionic semiquinone produces the same decrease in band I with a shift in band II from 375 nm to 380 nm. Reduction to the hydroquinone species removes these newly formed peaks at 580 nm and 380 nm, resulting in a spectrum devoid of clear peaks. Using these characteristic peak changes, the type of flavin (FAD or FMN) and its redox state can be determined. Figure 7: UV-vis spectra of free FAD and FMN. FMN and FAD can be differentiated due to a difference in the absorbance maxima of band I. FMN maximally absorbs at 446 nm while FAD has a maximum at 450 nm. 1.3 Fusobacterium nucleatum F. nucleatum is a Gram-negative, obligate anaerobe and is a common constituent in the human oral microbiome. It is non-motile and non-spore forming with a long, fusiform shape47. F. nucleatum belongs to the family Fusobactericeae and the phylum Fusobacteriota. Early attempts to classify F. nucleatum placed it in the family Bacteroidaceae, due to its Gram-negative nature47. However, modern genetic analysis reveals that, despite being Gram-negative, F. nucleatum shares significant portions of its genome, particularly genes encoding for central metabolism, from Gram- 11 positive organisms such as Clostridria48. This has led to the view that F. nucleatum genetically acquired Gram-negative traits (outer cell membrane) through lateral gene transfer49. The late acquisition of an outer cell membrane may have provided F. nucleatum with an advantage in evading the immune system. The immune system produces different cytokines in response to Gram-negative and Gram-positive organisms50. Additionally, the O-chain of the lipopolysaccharides located on the outer-membrane of Gram-negative organisms lends protection against complement-mediated killing by the host immune system51. These high-molecular weight side chains sterically prevent the binding of complement proteins like the membrane attack complex, which forms pores in the membrane if allowed to attach52. Like Gram-positive Clostridria, F. nucleatum relies on amino acid fermentation for its primary source of energy53. It primarily metabolizes glutamate, lysine, histidine, and serine. The product of this fermentation is butyrate, labelling F. nucleatum as a butyrate producer. F. nucleatum tends to migrate to extra-oral sites such as the human gut to colonize tumor microenvironments54. Here, its role as a butyrate producer may come into play. 1.3.1 Amino acid metabolism of F. nucleatum Similar to the Gram-positive Clostridium, F. nucleatum is weakly saccharolytic and preferentially derives energy through the fermentation of amino acids. The organism uptakes glutamate, lysine, histidine, and serine and can also degrade oligopeptides for catabolism49. Glutamate, lysine, and histidine all feed into two parallel pathways and form a common catabolite: crotonyl-CoA (Figure 8)55,56. Crotonyl-CoA is then converted to butyryl-CoA using a butyryl-CoA dehydrogenase (BCD) and electron bifurcating electron transfer flavoprotein (ETF). Electrons for this reaction are sourced from NADH. The reduction of crotonyl-CoA by NADH is highly exergonic as the midpoint reduction of NADH/NAD+ is -320 mV while the midpoint reduction potential of butyryl-CoA/crotonyl-CoA is -10 mV versus the standard hydrogen electrode. To conserve energy, this NADH-dependent reduction of crotonyl-CoA by the BCD-ETF complex is coupled to NADH-dependent reduction of flavodoxin. This reaction is endergonic as the reduction potential of FMNsq/hq is -420 mV. The coupling of an endergonic and exergonic reaction in the BCD-ETF is referred to as electron bifurcation and is mediated by the flavin cofactors of the BCD-ETF complex. Electron transfer from reduced flavodoxin to the membrane-bound Rhodobacter nitrogen fixation (Rnf) complex leads to the reduction of NAD+ and transport of Na+ out of the cell. The 12 Na+ electrochemical gradient can in turn fuel ATP production from ATP synthase (Figure 9). Butyryl-CoA is converted to end-products acetate and butyrate which are exported from the cell. In sum, butyrate and ATP production in F. nucleatum is dependent on fermentation of lysine, histidine, and glutamate57,58. 13 Figure 8: Glutamate and lysine fermentation in F. nucleatum. Glutamate, histidine, and lysine metabolism leads to the common metabolite crotonyl-CoA. 1 glutamate dehydrogenase, 2 2-hydroxyglutarate dehydrogenase, 3 glutaconate CoA-transferase, 4 2-hydroxyglutaryl-CoA dehydratase, 5 glutaconyl-CoA decarboxylase, 6 lysine 2,3-aminomutase, 7 lysine 5,6-aminomutase, 8 3,5-diaminohexanoate dehydrogenase, 9 3-keto-5-aminohexanoate cleavage enzymes, 10 3-aminobutyryl-CoA ammonia lyase, 11 butyryl-CoA dehydrogenase + electron transfer flavoprotein, 12 butyryl-CoA reductase, 13 flavodiiron protein, 14 butyryl-CoA:acetoacetate-CoA transferase, 15 butyryl-CoA:acetate CoA transferase, and 16 3-hydroxybutyryl-CoA dehydrogenase (and several more). Histidine 14 Figure 9: Energy conservation using electron bifurcation and the Rnf complex. The BCD:ETF complex (ETFa and ETFb are ETF subunits) performs electron bifurcation using electrons from NAD(P)H to reduce both crotonyl-CoA and flavodoxin (Fld). The low reduction potential of flavodoxin is used to generate NADH at the Rnf complex as well as establish a Na+ gradient. 1.3.2 F. nucleatum in the oral cavity and dissemination In the oral cavity, F. nucleatum is considered a “bridging” organism as it adheres to early colonizers of the biofilm (Streptococcus and Actinomyces spp.) and late colonizers that include periopathogens (Porphyromonas gingivalis and Treponema denticola)54,59. The organism encodes multiple adhesion proteins that enable it to coaggregate with many oral bacteria. F. nucleatum in this state is commensal with its human host but can transition into a pathogenic organism60. This 15 transition begins with the invasion of gingival epithelial cells which triggers a pro-inflammatory response61. Prolonged inflammation results in periodontal disease. Further invasion of host cells allows F. nucleatum to enter the bloodstream enabling it to disseminate to extraoral sites62. These sites include amniotic fluid and tumors and adenomas of the breast, colon, pancreas, and esophagus62–64. It is F. nucleatum’s dissemination to the gut, and subsequent strong association to colorectal cancer, which has been the central driver for research into the bacterium. 1.3.3 F. nucleatum and colorectal cancer F. nucleatum’s association with colorectal cancer began with the discovery of high levels of the bacterium in tumors compared to normal adjacent healthy tissue65. Total RNA was extracted from tumors and healthy tissues and analyzed with RNA-seq. The only bacterial species to show a significant increase in RNA in tumor tissue was F. nucleatum subsp. nucleatum. Since this initial finding in 2012, further research has revealed that F. nucleatum enhances cancer development through a variety of mechanisms and leads to poorer patient prognosis64. Accordingly, F. nucleatum has been classified as an oncogenic bacterium54. To date, mechanisms include the promotion of inflammation, angiogenesis, and metastasis along with the inhibition of chemotherapy and the immune response (Figure 10)66–69. The initial immune system response to F. nucleatum infection is pro-inflammatory with the release of cytokines interleukin 10 (IL-10), tumor necrosis factor-a (TNF-a), and nuclear factor kappa-b (NF-kB)70. This creates a tumor microenvironment, accelerating mutation rate and cancer progression. F. nucleatum also promotes the expression of annexin A1 in cancer cells71. Annexin A1 is stimulated when the bacterium binds to cancer cells via FadA, an adhesin protein. Annexin A1 then signals for expression of the transcription factor b-catenin which regulates cell proliferation71. Overexpression of b-catenin causes unregulated cell growth and tumor progression through the b-catenin-Wnt signaling cascade. It should be noted that the b-catenin pathway is one of many discovered and proposed biochemical impacts of F. nucleatum on cancer cells. F. nucleatum likely gravitates towards developing tumors due to the outpouring of growth substrates from unchecked cancer cells. Once there, F. nucleatum’s presence in a tumor translates to shorter survival times for patients as observed in one study of >1000 colorectal cancer cases64. 16 Figure 10: F. nucleatum’s role in aggravating colorectal cancer. F. nucleatum has been shown to promote inflammation, angiogenesis, and metastasis while inhibiting natural killer cells and chemotherapy treatment69. 1.3.4 Oxidative stress response of F. nucleatum An aspect of F. nucleatum that has not received much attention is its response to oxidative stress. In both the oral biofilm and tumor microenvironments, F. nucleatum likely encounters O2 and ROS. With the knowledge that other anaerobic species rely on enzymes to remove oxygen and other ROS, it is conceivable that F. nucleatum employs similar mechanisms. F. nucleatum has been shown to be aerotolerant, surviving 90 minutes in aerated culture2 or up to a 40% pure O2 atmosphere39. Indeed, annotation of the F. nucleatum genome reveals that it encodes a few enzymes that function in oxidative stress response (Figure 11). These enzymes include NOX, MsrAB, and AhpCF. These three enzymes have been briefly studied by other researchers. NOX activity was found through assays of cell pellets extracted from aerated cultures of F. nucleatum. These assays showed that NOX activity increased as aeration increased from 100% N2 to a 50:50 mix of N2 and air39. However, no attempts at NOX purification and in vitro characterization of the isolated protein were performed. Analysis of F. nucleatum’s genome reveals two NOX candidates, both showing homology to NOX-2 which reduces O2 directly to H2O. 17 The best characterized oxidative stress defense enzyme from F. nucleatum is an MsrAB24. A single gene locus encoding for an MsrAB has been identified. Using RT-PCR, expression of MsrAB was shown to increase when F. nucleatum was exposed to H2O224. A genetic knockout of msrAB in F. nucleatum resulted in a strain that exhibited increased sensitivity to H2O224. Furthermore, the knockout strain showed decreased adhesion and invasion of cancer cells. Located downstream in the same operon are two genes: modR and modS. Deletion of modR decreased MsrAB expression and deletion of modS increased MsrAB expression. ModRS therefore acts as a two-component signal transduction system that responds to H2O2 exposure and regulates expression of MsrAB24. Lastly, AhpCF has been identified in F. nucleatum26. The gene encoding AhpCF has been cloned and recombinantly expressed. Once purified, the enzyme was used to generate antibodies which were used to construct a vaccine. The vaccine inhibited F. nucleatum’s growth, showing the significance of AhpCF26. However, enzyme activity was not measured. The limited studies on F. nucleatum’s oxidative stress response do not reveal much about how F. nucleatum copes with environmental oxygen and immune system-generated ROS/RNS. Adding to the mystery of F. nucleatum’s aerotolerance is that it lacks catalase, SOD, and SOR, considered vital enzymes to aid against oxidative stress. However, F. nucleatum does encode for an FDP which may serve to reduce O2 and/or NO. 18 Figure 11: A summary of known enzymatic oxidative stress defenses in F. nucleatum. F. nucleatum encounters a variety of ROS and RNS species. To date, an NOX-2, AhpCF, MsrAB, and now an FDP enzyme have been characterized. 1.4 Flavodiiron proteins FDPs are a family of enzymes that were discovered 30 years ago28. They derive their name from their two composite domains: an N-terminal b-lactamase-like domain that contains a diiron site and a flavodoxin domain on the C-terminus that contains an FMN cofactor. The FMN of the flavodoxin domain accepts incoming electrons from an external source and then passes these electrons to the catalytic diiron site for substrate reduction. The b-lactamase-like domain shares the same tertiary structure motif as true β-lactamases (enzymes which cleave β-lactam antibiotics). β-lactamases have a αβ/βα sandwich fold motif and a binuclear metal binding site (Figure 12A)72. In FDPs, this metal site is occupied by two iron atoms bridged by hydroxide (µ-OH). The iron atoms are coordinated by 3 carboxylate and 4 imidazole ligands73,74. Like the β-lactamase-like domain, the flavodoxin domain’s tertiary structural is well conserved with flavodoxins. Flavodoxins typically exhibit an αβ parallel fold motif, where a singular 5-stranded parallel β sheet is surrounded by α helices (Figure 12B)75. 19 Figure 12: Tertiary fold motifs of E. coli’s FDPs. A) The αβ/βα sandwich fold of the β-lactamase-like domain. The internal β sheets are coloured shades of blue with the exterior α helices in green. B) The αβα fold of the flavodoxin domain. The β sheet is coloured blue. (PBD: 5LLD) In FDP monomers, the diiron site and FMN are placed too far apart for electron transfer to occur (~ 40 Å). However, FDPs form a head-to-tail dimer with the FMN of one monomer placed near the diiron site of the second monomer (~ 6 Å) (Figure 13)76. The dimeric configuration allows for electron transfer from the reduced FMN cofactor to the diiron motif. Crystal structures reveal that some FDPs form tetramers from the association of two head-to-tail dimers77. The first FDP was discovered in anaerobe D. gigas. Enzymatic assays showed that the FDP reduced oxygen to water (Figure 12). The reaction was dependent on electrons transferred to the FMN through a reduced Rd78. Rd is in turn reduced by an NADH-dependent rubredoxin oxidoreductase (NROR), a flavin-containing enzyme. Other FDPs were then found in other bacteria, including E. coli. The FDP from E. coli was the first to exhibit the ability to reduce NO to N2O (Figure 13)79. FDPs characterized to date can reduce either O2 or NO, or both28. For example, E. coli’s FDP is able to reduce both O2 and NO, but exhibits a much higher catalytic efficiency for NO79. A singular FDP, from C. difficile, has also shown H2O2 reductase activity which remains to be seen in other enzymes80. A B 20 Figure 12: Overall reactions of FDPs with both O2 and NO. Figure 13: Head-to-tail dimer structure of FDPs. The crystal structure of the FDP from D. gigas adopts a characteristic head-to-tail dimer structure orienting the diiron site of one monomer close to the flavin site of the other. (PBD: 1E5D) One monomer is shown in green and the other is shown in blue. The FMN cofactor is shown as sticks with the iron atoms and bridging µ-OH shown as balls. 1.4.1 Flavodiiron proteins as oxidative stress-clearing enzymes FDPs have been identified mainly in bacteria, with several being found in Archaea, eukaryotes, and several photosynthetic organisms such as cyanobacteria, green algae, mosses, and gymnosperms76. The primary function in bacteria is to act as an oxidative and or nitrosative stress response, directly reducing O2 and NO before they can cause cellular damage and react to form 21 downstream ROS and RNS. It is no surprise, therefore, that FDPs are primarily found in obligate anaerobes where O2 is a toxic diatomic gas28,79,81. NO is also a toxic molecule that inhibits bacterial growth at sufficient cellular concentrations. NO arises in the mammalian gut from the breaking down of nitrate by gut microbes through the denitrification process82. NO is also generated by the human immune system to kill invading pathogens. NO is produced enzymatically by NOS, which is overexpressed in activated macrophages12,83. An FDP with the ability to reduce NO to N2O, like the FDP in E. coli, gives a bacterium a direct method to protect itself from both bacterial derived NO as well as NO produced by activated macrophages. In this capacity, FDPs may be considered a virulence factor. It remains elusive as to what determines a particular FDPs affinity for O2 vs NO. The primary coordination sphere for the diiron motif is the same for O2 and NO selective FDPs. It has been speculated that substrate specificity may be controlled by the second coordination shell, which does vary from one FDP to the next. 1.4.2 Mechanism of flavodiiron proteins While the structural reasons behind O2 and NO affinities are unknown, the reduction mechanisms for each substrate have been better studied. Through modeling, the binding and subsequent reduction of O2 has been analyzed as two separate processes: an oxidative part and a reductive part84. Here, oxidation and reduction are referring to the oxidation state of the FDPs’ cofactors. The oxidative part begins with the binding of O2 to a fully reduced active site (FeII-FeII- FMNH2) (Figure 14A). O2 binds and then forms a peroxide species. Next, the proton transfer occurs leading to the formation of a hydroperoxide group, bound to only one of the iron atoms84. This hydroperoxide favours O-O bond cleavage more so than the peroxide. The O-O bond is cleaved yielding hydroxo and oxyl radical ligands. The oxyl radical is rearranged to form an oxo-ferryl group84. This oxo-ferryl group is finally reduced to a second hydroxo ligand. This concludes the oxidative half of the reaction. The active site is now FeIII-FeIII- FMNox. The reductive part begins with the reduction of the FMN cofactor to FMNH2 by an external source (NADH)84. Each hydroxo ligand is then converted to water sequentially with proton transfer, with an intermediate of one hydroxo and one water ligand being formed. Once both water ligands have formed, the FMN cofactor is again reduced to FMNH2 and then water is released84. 22 The NO mechanism has been modeled and concretely identified using mechanistic studies. The mechanism again starts with both cofactors in the reduced state (FeII-FeII- FMNH2) (Figure 14B). A single NO binds forming {FeNO}7 (Enemark-Feltham notation) closely followed by a second giving [{FeNO}7]2, or a diferrous dinitroyl complex85. The bound NO is reduced with electrons from the iron atoms and released as N2O in the rate determining step of the reaction85. A second round of catalysis can occur by the regeneration of the diferrous state through reduction by FMNH2. This explanation still yields several unanswered questions. The exact route by which the N-O bond of NO is broken and the N-N bond of N2O is formed are unknown. One theory is that a hyponitrite intermediate is formed first (N-N bond formation) followed by subsequent N-O cleavage86. Additionally, the proton source used to generate the second product, H2O, is unknown. 23 Figure 14: The O2 and NO reduction mechanisms of FDPs. A) The O2 mechanism is shown first and split into the oxidative and reductive halves. B) The NO mechanism shows one cycle of catalytic activity. A second cycle can occur without an external supply of electrons from NAD(P)H. Reductive Half Oxidative Half A B 24 1.4.3 Classification of flavodiiron proteins Archetypal FDPs contain a b-lactamase-like domain and a flavodoxin domain. These FDPs rely on an external source of electrons for substrate reduction. Classically, NADH serves as this source. NADH is oxidized by secondary NADH oxidoreductase, and the electrons are passed to an intermediate electron transfer protein, typically Rd. The electrons are then passed onto the FDP. This canonical set up establishes that each FDP must have its own electron transport chain to perform catalysis. FDPs are divided into different classes (Class A- Class H) based on what parts of the electron transport chain are incorporated directly into the FDP polypeptide at the C-terminus (Figure 15)28,76. All classes contain the base unit of a b-lactamase-like domain and flavodoxin. Class A FDPs (D. gigas) contain only the base unit78. Class A is the type of FDP found in F. nucleatum. Class B FDPs (E. coli) and Class D FDPs contain a canonical Rd or short-type Rd domain respectively79. Class E adds an iron-sulfur cluster is place of this Rd (Fe3S4)87. Class C FDPs add a flavin oxidoreductase, able to directly oxidize NADPH without any secondary enzymes28. Class F fuses a rubredoxin oxidoreductase (ROR) with the FDP. Classes G and H combine both Rd and flavin oxidoreductase (FldOR) domains with Class H incorporating two Rd domains28. All these classes use NAD(P)H as a source of electrons. The only known exception are methanogens which instead use coenzyme F42088. Another common theme is the almost ubiquitous use of Rd as an electron donor to the FMN site. Except for Class C, all other classes use Rd. 25 Figure 15: The classes of FDPs. Each class contains the diiron (b-lactamase-like) and flavodoxin domains with additional domains attached to the C-terminus. Many contain Rb or RbSC (rubredoxin short type) domains. Several directly incorporate a reductase domain in the form of a FldOR or ROR. 1.4.4 Diiron centre motif in proteins The diiron motif found in FDPs is not unique to this enzyme family. Non-heme diiron centers are found in hemerthyrins, rubrerthyrins, methane monooxygenases, ribonucleotide reductases, and ferritins89. Hemerthyrins, unintuitively, do not contain a heme. They are named for their function: O2 transport in the blood of marine invertebrates. Rubrerthyrins can be thought of as a combination of hemerythrins and FDPs. They contain a N-terminus hemerythrin-like diiron site and a C-terminus Rd domain89. Like FDPs, rubrerthyrins function in oxidative stress, binding H2O2 at their diiron site, and using Rd to accept incoming electrons from NAD(P)H. They directly reduce H2O2 to H2O. The last of the diiron proteins to bind oxygen atoms are methane monooxygenases. Methane monooxygenases bind methane and oxygen to form methanol in the first step of methanotrophic metabolism90. They, like FDPs, use an FAD cofactor to accept incoming reducing equivalents89. Ribonucleotide reductases and ferritins are more ubiquitous 26 across domains of life than other diiron proteins. Ribonucleotides reductases are found in all organisms performing the essential conversion of ribonucleotides into deoxyribonucleotides. This reaction proceeds with formation of a tyrosyl radical which is partially stabilized by the diiron site91. Also essential, ferritins control the storage of iron atoms. They form large multimeric protein complexes, with each monomer containing a diiron site89. For instance, human ferritin uses 24 monomers to form a large interior pocket to store iron until needed. These examples show the versatility of the diiron site as a common motif, with a tendency to function in electron transfer and oxygen binding. 1.6 The electron transfer partner of FDP in F. nucleatum Examination of F. nucleatum’s genome sequence reveals that it encodes an FDP (FNP2073) in a dicistronic fashion with a multicomponent protein (FNP2074). BLAST sequence analysis reveals that FNP2074 has three domains with homology to a BCD, the C-terminal domain of the a subunit of ETF, and a Rd respectively (Figure 16). This unique trifecta of domains in a single polypeptide has not been observed or characterized before. As Class A FDPs require an external source of electrons and quite often use Rd as an electron transfer partner, it was hypothesized that this novel, modular enzyme oxidizes butyryl-CoA to reduce Rd, which subsequently reduces the FMN cofactor of the FDP. As such, FNP2074 was renamed butyryl-CoA reductase (BCR). Figure 16: A model of BCR coupled to FDP. Distinct domains of BCD, ETFa, and Rd are shown along with their expected cofactors. 27 1.7 Butyryl-CoA dehydrogenase BCDs are a part of the short chain fatty acyl-CoA dehydrogenase group of enzymes. BCDs bind FAD as a cofactor and are composed of an aba tertiary fold motif92. Across both mammalian and bacterial crystal structures, this fold is conserved92. In mammals, BCDs function in the mitochondria, oxidizing butyryl-CoA to form crotonyl-CoA. As described in further detail below, electrons from this reaction are transferred to an ETF (Figure 17). BCDs are also part of the BCD-ETF complex present in anaerobic bacteria, where they play an important role in energy conservation through electron bifurcation. 1.8 Electron transfer flavoproteins ETFs are widespread enzymes found in many different species. Current classification of ETFs divides them into two groups: canonical and bifurcating (Figure 17)93. ETFs are dimeric, comprising of an a and a b subunit. Together, these two subunits form three domains. Domain I is composed of the N-terminus of the a subunit, domain II is made up of the C-terminus of both subunits, and the remaining N-terminal region of the b subunit comprises domain III93. Canonical ETFs bind an AMP in the b subunit (domain III) and an FAD with the a subunit (domain II). These ETFs are located in the mitochondria where they act as central redox partners, accepting electrons from dehydrogenase enzymes involved in fatty acid oxidation and amino acid catabolism93. These enzymes include glutaryl-CoA, isobutyryl-CoA, isovaleryl-CoA, and other acyl-CoA dehydrogenases93. They then pass these electrons to electron transfer flavoprotein-ubiquinone oxidoreductase (ETF-QO) which then delivers them to coenzyme Q10 (ubiquinone). Coenzyme Q10 then passes electrons down the electron transport chain to drive the formation of a proton gradient in the intermembrane space. The bound FAD cofactor of canonical ETFs facilitates this single electron transfer, with the flavin cofactor cycling between the anionic semiquinone and hydroquinone93. The AMP in the b subunit is likely an evolutionary relic and does not have a functional role. In bifurcating ETFs, the AMP is replaced by a second FAD cofactor. The two FADs are designated a-FAD and b-FAD relative to the a and b subunits they respectively bind. This second b-FAD is the FAD that functions in electron bifurcation. The full ETFab binds to BCD forming the BCD-ETFab complex94. The catalytic cycle initiates with hydride transfer from NADH to the 28 b-FAD which leads to formation of the b-FAD hydroquinone (b-FADH2). Rapid electron transfer from the b-FADH2 to the anionic semiquinone of the a-FAD (a-FAD•-), generates the hydroquinone of a-FAD (a-FADH2) and the semiquinone form of b-FAD (b-FADH•) As the b-FADsq/b-FADox pair exhibits a very low reduction potential (-700 mV), b-FADH• is then able to transfer an electron to either a flavodoxin or ferredoxin (Fd)95. The hydroquinone of a-FAD then donates electrons to the FAD of BCD (d-FAD). A second round of NADH oxidation and electron bifurcation enables the reduction of crotonyl-CoA by BCD94. Figure 17: Canonical vs bifurcating ETFs. Canonical ETFs oxidize butyryl-CoA during b-oxidation and funnel those electrons to the ETC for ATP generation. They contain an AMP bound to their b subunit. Bifurcating ETFs instead contain a second FAD in their b subunit which bifurcates electrons from NAD(P)H, using one to reduce ferredoxin or flavodoxin and the other to reduce crotonyl-CoA. Given that the central ETF domain of BCR only shows homology to the FAD-containing C-terminal a-subunit (hereafter named ETFa), the domain is unlikely to interact with NADH and perform electron bifurcation. Instead, the FAD cofactor of the ETFa subunit is more likely to act in electron transfer between the attached BCD and Rd domains. 1.9 Rubredoxins Rds are small proteins (~6 kDa) that participate in electron transfer. They are unique to bacteria and archaea. They possess a single iron binding site where the iron is coordinated by four cysteine residues. At the iron site, Rds perform single electron transfer, alternating between ferrous 29 and ferric forms. Despite possessing no inorganic sulfur, Rds are occasionally referred to as having a Fe1S0 cluster96. The coordinating SCys ligands follow a conserved dual CxxC motif. The distance between the two CxxC motifs is used to classify Rds as either canonical or short chain. Canonical Rds have a ~30 amino acid spacer between each motif while short chain Rds have ~1287. Beside their role as FDP electron transfer partners, Rds also participate in fatty acid oxidation, carbon fixation, and superoxide reduction97,98. Rds exhibit varied reduction potentials, ranging from -50 to +50 mV99. 1.10 Research objectives The FDP from F. nucleatum presents a possible mechanism through which the bacteria might circumvent oxidative and nitrosative stress. Given FDPs’ dual activity with O2 and NO, this enzyme might help F. nucleatum in its oral microbiome niche where it is exposed to environmental oxygen and in its distal migrations where it feasibly encounters NO produced by our immune system. FDP’s colocalization with BCR suggests a role for BCR as an electron transfer partner for FDP’s reductase activity. The inclusion of a Rd domain in BCR further hints that it participates in electron transfer with the FDP. The objective of this research was to characterize this novel oxidative stress response system. The first goal was to recombinantly express and purify both FDP and BCR from F. nucleatum. UV-visible and EPR spectroscopy could then be used to determine cofactor identity and follow flavin transitions through quinone, semiquinone, and hydroquinone states. Next, the ability of BCR to oxidize butyryl-CoA and the properties of the electron transport system through the distinct domains (BCD, ETFa, and Rd) could be examined. The final objective was to test the catalytic activity of FDP on potential substrates: O2, NO, and H2O2. 30 Chapter 2: Coupling oxidation of butyryl CoA to oxygen reduction through BCR-FDP 2.1 Chapter summary BCR-FDP may be vital to the oxidative stress defense of F. nucleatum. To determine the specific role of BCR-FDP in F. nucleatum, both BCR’s and the FDP’s gene sequences were cloned into plasmids and transformed into E. coli. Each gene was expressed in culture media to produce recombinant protein for analysis. Separately, BCD and ETFa domains of BCR were also cloned and recombinantly expressed. Once purified, proteins were characterized using a variety of methods to determine the cofactor content, electron transfer properties, and active substrates. BCR-FDP was found to be an oxygen reductase, reducing oxygen to water, a reaction that was dependent on butyryl-CoA. This reaction enables F. nucleatum to survive oxidative stress using a high-potential electron donor and common metabolite butyryl-CoA. Due to the abundance of butyryl-CoA produced through amino acid fermentation by F. nucleatum, BCR-FDP activity also facilitates the recycling of crotonyl-CoA, enabling the organism to profit energetically from transient exposure to oxygen. 2.2 Materials and methods 2.2.1 Materials Oligonucleotides were purchased from Integrated DNA Technologies (IDT) (Coralville, IA, USA). Restriction enzymes (NdeI, NheI, and BamHI), Q5 DNA polymerase, antarctic phosphatase, and DNA ligase were all purchased from New England Biolabs (Ipswich, MA, USA). Plasmid extraction was performed using the E.Z.N.A Plasmid DNA Mini Kit from Omega Bio-tek (Norcross, GA, USA). Butyric anhydride was purchased from Tokyo Chemical Industry (Portland, OR, USA). All other chemicals used were purchased from Thermo Fisher Scientific. 2.2.2 Data analysis All data analysis along with data figure creation was done in the program OriginPro (OriginLab, v. b9.3.226). Protein structure figures were created and visualized with PyMOL (Schrödinger, v. 2.5.8). 31 2.2.3 Cloning of BCR, BCR-FDP, BCD, and ETFa Contiguous genes FNP2073 and FNP2074 from F. nucleatum subsp. polymorphum ATCC 10953 are annotated to encode putative flavodiiron protein and multidomain protein comprising an acyl-CoA dehydrogenase, ETF⍺, and Rd, respectively. Primers were designed to amplify FNP2074 as well as the dicistron FNP2073 and FNP2074 (Table 1) by PCR. Primers were also designed to amplify the individual BCD (residues 1-379 of BCR) and ETF⍺ (residues 369-560 of BCR) domains (Table 1). These domains were identified using BLAST analysis and an AlphaFold model of BCR that depicted the domains (Figure 16). All forward primers contained an NdeI restriction site and all reverse primers contained a BamHI site. The exception was the forward primer for the BCR-FDP dicistronic unit which used an NheI site. This was done as the DNA sequence encoding the FDP contains an internal NdeI site. PCR amplification was performed using Q5 DNA polymerase with F. nucleatum subsp. polymorphum ATCC 10953 genomic DNA as a template. PCR cycle conditions were as follows: initial denaturation at 95°C for 2 min followed by 30 cycles of heat denaturation at 95°C for 30 s, annealing at 63°C for 30s, and extension at 72°C for 30s/kbp followed by a final extension for 5 min at 72°C. The PCR products corresponding to BCR, BCD, and ETF⍺ were digested with NdeI and BamHI, while the PCR product corresponding to BCR-FDP was digested with NheI and BamHI. The cDNA corresponding to BCR was ligated into the NdeI and BamHI site of pET15b. The cDNA corresponding to BCR-FDP was ligated into the NheI and BamHI site of pET28a while the cDNA corresponding to BCD and ETF⍺ were ligated into the NdeI and BamHI site of pET28a. The subcloning generated four plasmid constructs: pET15b_BCR, pET28a_BCD, pET28a_ETF⍺, and pET28a_BCR-FDP. The ligation mixtures were then used to transform E. coli XL1-blue electrocompetent cells using electroporation. The cells were plated on Luria-Bertani (LB) agar plates containing the appropriate antibiotic (100 µg/mL ampicillin for pET15b and 50 µg/mL kanamycin for pET28a). This cloning approach gave constructs with a His6-tag fused to the N-termini of BCR, BCD, and ETF⍺. The cloning strategy for pET28a_BCR-FDP resulted in a His6-tag fused to the N-terminus of the BCR sequence. Plasmid constructs were extracted and purified from transformed XL1-blue cultures using the E.Z.N.A Plasmid DNA Mini Kit (Omega Bio-tek). Extracted plasmids were digested using the same restriction enzymes used for cloning and then the digest was run on a 0.5% agarose gel. The restriction fragment length verified the presence of 32 insert in extracted plasmids. For protein expression, the purified plasmid constructs were transformed into E. coli BL21(DE3) competent cells. Single colonies were used to inoculate 5 mL overnight LB cultures. Glycerol stocks of the transformed cells- prepared by adding 500 µL of 50% glycerol (v/v) to 500 µL of LB overnight culture- were stored at -70°C. Preliminary sequencing analysis was performed for plasmid constructs pET15b_BCR, pET28a_BCD, pET28a_ETF⍺, and pET28a_BCR-FDP. Sanger sequencing was performed by Fragment Analysis and DNA Sequencing Services (FADSS) at UBC Okanagan using an Applied Biosystems 3130xl Genetic Analyzer (ThermoFisher). Approximately the first 100-200 bps downstream of the T7 promoter were sequenced using the T7 forward primer. Analysis of the resulting chromatograms verified that the inserted sequences of BCR, BCD, ETF⍺, and BCR-FDP contained no errors over the initial ~100 bps. Table 1: Primer sequences of each sequence of interest Sequence Forward and Reverse Primers BCR 5¢-CTA GCT AGC ATG CTT TTT AAA ACT ACT G-3¢ 5¢-CCG GAT CCT TAT TTT GCA TCT ATA TAT CC-3¢ BCR-FDP 5¢-CTA GCT AGC ATG CTT TTT AAA ACT ACT G-3¢ 5¢-CGG GAT CCT TAT TTA TTT AAA GAT TCA ACT AAT AC-3¢ BCD 5¢ GGA ATT CCA TAT G CTT TTT AAA ACT ACT GAA GAA C-3¢ 5¢- CGG GAT CCT TAT CCT ATT AAG TGT GCA GCT ATA AC-3¢ ETF⍺ 5¢-GGA ATT CCA TAT GCA AAG AGT TGT TAT AGC-3¢ 5¢-CGG GAT CCT TAA TTG TCT AAT TCC TTA GTT G-3¢ 2.2.4 Protein expression Starter cultures were prepared by inoculating 100 mL of LB media containing the appropriate antibiotic with swabs of the glycerol stock of E. coli BL21(DE3) transformed with either pET28a_BCR-FDP, pET15b_BCR, pET28a_BCD, or pET28a_ETF⍺. The starter culture was allowed to grow overnight at 37°C with shaking at 180 rpm. Ten mL of starter culture was added to 1 L cultures of LB media containing the appropriate antibiotic. The cultures were grown 33 at 37°C until the optical density at 600 nm (OD600), reached 0.6. Isopropyl ß-D-1-thiogalactopyranoside (IPTG) was added to the 1 L culture to a final concentration of 0.2 mM. The cultures were then allowed to grow overnight (~16 hours) at 25°C. The cultures were then harvested via centrifugation (4355 x g for 15 min at 4°C). The resulting cell pellets were pooled and stored at -70°C. E. coli BL21(DE3) cells harbouring pET15b_BCR or pET28a_BCR-FDP were also expressed in M9 minimal media using the following protocol. Ten mL of a 100 mL LB started culture of E. coli BL21(DE3) was used to inoculate 1.5 L M9 cultures contained 1x M9 salts, 2 mM MgSO4, 100 µM CaCl2, 20 mM D-(+)-glucose, 0.2 mM Fe(NH4)2(SO4)2, and the appropriate antibiotic. The cultures were grown at 37°C until the OD600 reached 0.6. At this point, IPTG (0.2 mM) and Fe(NH4)2(SO4)2 (0.2 mM) were added to the cultures. The cultures were grown overnight at 18°C, harvested via centrifugation, and the cell pellets were stored at -70°C. 2.2.5 Aerobic protein purification All proteins were purified using the same protocol and all steps were performed on ice or at 4°C. Cell pellets (between 10-15 g) were resuspended in 50 mM phosphate buffer containing 0.15 M NaCl at pH 7.4 (Buffer A) with 10 mM imidazole. Two protease inhibitors, benzamidine (0.5 mM) and phenylmethylsulfonyl fluoride (PMSF, 10 µM), were added to the resuspended cell suspension. The cells were lysed for 30 min by sonication with the following settings: 25% power for 8 s at 40 s intervals. The lysed cells were centrifuged at 40,000 x g for 1 hour. The resulting supernatant was pooled and 200 µM of flavin adenine dinucleotide (FAD) was added. Flavin mononucleotide (FMN; 200 µM) was also added to the clarified cell extract for the purifications of BCR-FDP. The supernatant was applied to a 5 mL Ni-nitrilotriacetic (Ni-NTA) fast flow column which had been previously equilibrated with 25 mL of Buffer A. The column was then washed with 25 mL of Buffer A containing 10 mM imidazole and then a further 25 mL of Buffer A with 40 mM imidazole. The protein was eluted with Buffer A supplemented with 300 mM imidazole. 5-10 mL of eluent was collected and dialyzed overnight against 2 L of Buffer A. Protein not immediately used for analysis was flash frozen using liquid nitrogen and stored at -70°C. 34 2.2.6 Anaerobic protein purification BCR and ETF⍺ were also purified under anaerobic conditions. Anaerobicity was maintained by performing all steps in a glove box (Belle Technology, Weymouth, UK) with an O2 concentration < 5 ppm. All buffers used were purged with 99.998% N2 (1 h per 100 mL of buffer) prior to introduction into the glove box. A temperature of 4-10°C was maintained through use of an ice box and all buffers were pre-chilled in ice before use. The cell pellet was brought into the glove box and resuspended in Buffer A supplemented with benzamidine (0.5 mM) and PMSF (10 µM). The resuspended cells were lysed by sonication with the same settings as those described for aerobic purification. The lysed cells were sealed in centrifuge tubes and brought outside the glove box for centrifugation (50,000 x g for 1 hour). The supernatant was returned to the glove box and applied to a gravity flow Ni-NTA column (2.5 cm x 8 cm) equilibrated in Buffer A containing 10 mM imidazole. The column was washed with 50 mL of Buffer A containing 10 mM imidazole. The protein was then eluted from the column with Buffer A containing 300 mM imidazole. Protein not immediately used for analysis was flash frozen using liquid nitrogen and stored at -70°C. 2.2.7 SDS-PAGE Protein size and purity was assessed using SDS-PAGE. Protein (at approximately 1 mg/mL) was mixed with an equal volume of SDS loading dye (62 mM Tris-HCl, 10% v/v glycerol, 10% v/v SDS, 0.0025% v/v bromophenol blue, 5% v/v b-mercaptoethanol, pH 6.8) and heated to 95°C for 5 min. Following cooling on ice for 3mins, the protein was then applied to a 12% SDS polyacrylamide gel along with the PageRuler protein ladder (ThermoFisher), which was used to estimate molecular mass of BCR, BCD, ETF⍺, and FDP. Gels were run at 120 V until the dye front reached the bottom of the gel. Gels were stained with Coomassie blue and destained with destaining solution (20% v/v methanol, 10% v/v acetic acid) for visualization of protein. 2.2.8 Size exclusion chromatography Size exclusion chromatography was performed to assess oligomeric state of BCR and the BCR-FDP complex. A Superdex 200 16/200 column was equilibrated with 3 column volumes (CVs) of 50 mM phosphate buffer, 0.15 M NaCl at pH 7.4. Protein at 2 mg/mL in a total volume 35 of 5 mL was added to the column and the absorbance at 280 nm was monitored. Elution volume was calculated from absorbance peaks at 280 nm and compared to a standard curve to extract molecular weight. The standard curve was constructed using cytochrome C (12 kDa), carbonic anhydrase (29 kDa), alcohol dehydrogenase (150 kDa), and b-amylase (200 kDa). Blue dextran was used as a measure of the void volume. Equation 1 was used to calculate Kav for each protein and plotted against the log of the molecular weight to form the standard curve. Equation 1: 𝐾!" = #!$#"##$#" Where Kav is the fraction of pore volume accessible to the molecule, Ve is the elution volume of molecule, V0 is the void volume of the column, and Vt is the total volume of the column. 2.2.9 UV-visible spectroscopy A PerkinElmer Lambda 25 ultraviolet (UV)-visible spectrophotometer was used to generate absorbance spectra of protein under aerobic conditions while a PerkinElmer Lambda 265 UV-visible spectrophotometer housed in the glove box was used to acquire absorbance spectra under anaerobic conditions. 2.2.10 Protein quantification and extinction coefficients The protein concentration of all samples was measured through the use of the Pierce BCA Protein Assay Kit (ThermoFisher). A standard curve was generated using BSA. Protein concentration was tested in triplicate. The protein concentration was then used to calculated extinction coefficients for the lmax at the 280 nm peak. 2.2.11 Flavin analysis and quantification The identity of the flavin (FAD or FMN) present in each protein was determined by methods employing UV-visible spectroscopy or fluorescence spectroscopy. For the method involving UV-visible spectroscopy, a 1 mL sample of protein was heated to 95°C for 5 min to induce protein denaturation and release of the flavin. The precipitated protein was separated from the free flavin by centrifugation (16,000 x g for 5 min). The absorbance spectrum was taken of the 36 supernatant. A solution of FAD was identified by a lmax at 450 nm and a solution of FMN was identified by a lmax at 446 nm. A mixture of free FAD and FMN was identified by a lmax between 450 and 446 nm. Fluorescence spectroscopy was used to quantitate the individual concentrations of FAD and FMN within a protein sample100. The fluorescence method exploits the fact that FMN has a 10-fold higher quantum yield compared to FAD, and that snake venom phosphodiesterase converts FAD to FMN and adenine monophosphate. To quantitate the relative amounts of FAD and FMN in a sample, the fluorescence spectrum of the flavin solution is taken before and after addition of the snake venom phosphodiesterase. The relative increase in fluorescence of the flavin solution after the addition of snake venom phosphodiesterase can be used to quantitate the amount of FAD and FMN in the solution. The protocol initiated with heat denaturation of the protein (95°C for 5min) followed by removal of the precipitated protein through centrifugation (16,000 x g for 5 min). A PerkinElmer LS50-B spectrophotometer was then used to acquire fluorescence emission envelope from 500 to 600 nm following excitation at 450 nm (the flavin absorbance maxima) of the supernatant which contained the free flavin. Snake venom phosphodiesterase (Sigma-Aldrich, 20 U/mg) was added to the flavin solution. Following a 10 min incubation, the fluorescence spectra was acquired again using the same settings. If the protein contains only FMN, no fluorescence change is observed upon PDE treatment. Conversely, if the protein contains only FAD, a 10-fold fluorescence increase is expected. For solutions containing both flavin cofactors, the molar ratio (r) between FAD and FMN can be calculated from the fluorescence increase after PDE treatment using Equation 5 which is based on Equations 2-4. Equation 2: 𝐹% = 𝑓&'( ∙ [𝐹𝐴𝐷] + 𝑓&)* ∙ [𝐹𝑀𝑁] Equation 3: 𝐹+ = 𝑓&)*([𝐹𝐴𝐷] + [𝐹𝑀𝑁]) Equation 4: 𝑓&'( = 0.1𝑓&)* Equation 5: 𝑟 = (-%∙$%$"$-%)-%$$%$" 37 Where r is the molar ratio between FAD and FMN, F0 is the fluorescence of released flavin, Ff is the fluorescence after phosphodiesterase treatment, fFAD is the fluorescence of a 1 µM solution of FAD, and fFMN is the fluorescence of a 1 µM solution of FMN. 2.2.12 Inductively coupled plasma mass spectrometry (ICP-MS) Inductively coupled plasma mass spectrometry (ICP-MS) was used to analyze metal content after protein purification for BCR and BCR-FDP. Each protein was concentrated to approx. 100 µM using centricons (Vivaspin 20, Sigma-Aldrich) with a 30 kDa cutoff. The resulting flow-through from the centricons was used a blank for metal analysis. ICP-MS analysis was performed by the Fipke Laboratory for Trace Element Research using an 8900 ICP-QQQ-MS (Agilent, Canada) with an ASX520 autosampler (Cetac, AB, Canada). Samples were subjected to acid mineralization through addition of 0.2 mL HNO3 to 500 µL of protein sample. Samples were incubated in acid for 30 min at 25°C and then diluted to 5 mL with double-deionized water. The ICP-MS instrument was calibrated with a four-point curve using internal standards of Sc, Y, In, and Tb. Maximum sensitivity was maintained by keeping the oxide and divalent-ion ratios below 2%. The metal concentration of each corresponding blank was subtracted from the amount in each protein sample to give the final metal concentration harboured in each protein. 2.2.13 Reduction-oxidation (redox) titrations To observe the various redox states of the flavin cofactors, BCR, BCR-FDP, BCD, and ETF⍺ were subjected to redox titrations. All redox titrations were performed in the anaerobic glove box using anaerobic buffer and reagents. Protein stocks (1 mL) were made anaerobic through overnight exposure to the N2 atmosphere. Protein was then diluted with 50 mM HEPES, 0.15 M NaCl, pH 7.4 buffer until a final concentration and final volume of 50 µM and 1 mL were obtained. Dithionite was used as a reductant and potassium ferricyanide was used as an oxidant. 0.1 M solutions of dithionite and potassium ferricyanide were created and diluted to 1-5 mM. These dilute stocks were added in 1 µL aliquots. After each addition, a UV-visible spectrum was obtained. 38 2.2.14 Butyryl-CoA synthesis Butyryl-CoA was synthesized in a similar manner as previously described21. 25 µmol of coenzyme A (CoASH) was dissolved in 5 mL of 0.5 M potassium bicarbonate. Separately, 35 µmol of butyric anhydride was dissolved in 0.5 mL of acetonitrile. The butyric anhydride was then mixed with the coenzyme A solution and allowed to stir for 10 min. Free thiol tests conducted with 5,5’dithiobis(2-nitrobenzoic acid) following the 10 min incubation showed no free thiol. The mixture was then brought to pH 2 using 6 M HCl. A C18 column (gravity flow) was then used for purification. The column was pre-washed with methanol and then equilibrated with 3 CVs of 0.1% trifluoroacetic acid (v/v, TFA). The reaction mixture was then applied over the column. The column was washed with 3 further CVs of 0.1% TFA. The butyryl-CoA product was eluted from the column using 0.1% TFA containing 50% acetonitrile (v/v) into a small round bottom flask. To remove the organic solvent, the round bottom flask was attached to a rotary evaporator. The rotary evaporator was set to a pressure of 145 mbar and the water bath was set to 50°C. Acetonitrile evaporation occurred within 5 minutes. The remaining solution was aliquoted into 1.5 mL Eppendorf tubes and flash frozen using liquid nitrogen. The frozen tubes were placed into a lyophilizer (freeze dryer) and aqueous solvent was allowed to sublime overnight. The resulting white powder was then stored at -70°C until use. 2.2.15 Stopped-flow measurements All stopped-flow data were obtained from a SF-61DX2 stopped-flow (TgK Scientific, Bradford-on-Avon, UK) with the sample handling unit of the system located in the anaerobic glove box. All experiments were performed at 20°C in 50 mM HEPES, 0.15 M NaCl, pH 7.4 buffer that was made anaerobic through extensive purging with N2. Enzyme (BCD or BCR) at 10-15 µM was rapidly mixed against butyryl-CoA in 7 to 10-fold excess (100 µM) to create pseudo-first order conditions. Single-wavelength measurements at 450 nm were collected in triplicate using the photomultiplier tube. Multiwavelength scans were collected using the photodiode array (PDA) detector over a set time frame. Observed rate constants (kobs1 and kobs2) were extracted by fitting the single-wavelength data were fit to a double exponential equation (Equation 6) in the Kinetic Studio software (TgK Scientific, v.2.0.33.17111): 39 Equation 6: 𝑦 = −𝐶1 ∙ 𝑒($0&'()∙1) − 𝐶2 ∙ 𝑒($0234*∙1) + 𝐶 where y is the absorbance at 450 nm and C1 and C2 are the amplitude of absorbance change associated with each kinetic phase. The extracted rate constants were averaged and plotted against substrate concentration in an attempt to measure the dissociation constant (Kd). 2.2.16 Electron paramagnetic resonance (EPR) Electron paramagnetic resonance (EPR) experiments were carried out with a Magnettech ESR5000 spectrometer (Bruker, Billerica, MA, USA). Protein samples were prepared at concentrations of 100 µM and placed in 100 µL capillary tubes. All experiments carried out at 295 K. A microwave power of 80 milliwatts with a modulation amplitude of 0.9 mT and a modulation frequency of 100 kHz were used. A sweep time of 120s was used and 5 scans were collected. The magnetic field was scanned from 3330 to 3420 G. 2.2.17 Generation of nitric oxide NO was generated through the reaction of 6 M sulfuric acid and 8.7 M sodium nitrite (NaNO2)101. Due to the ability of NO to rapidly react with O2 to form NO2, the reaction was performed under anaerobic conditions. NaNO2 was placed in a pressure-equalizing dropping funnel and a double-necked round bottom flask containing the sulfuric acid was placed beneath the funnel. The top of the funnel was sealed using a rubber septum. A needle connected to a gas cannister containing 99.998% N2 was inserted into the septum. The second neck of the round bottom flask was connected via rubber tubing to a glass bubbler containing 1 M NaOH. NaOH reacts with NO2, and thus acts as a purification step by preventing NO2 contamination. A second glass bubbler containing deionized water was attached to the first serving to remove any traces of base from the gas. This bubbler was connected to a needle inserted into a second septum attached to a glass bottle containing 50 mM HEPES-NaOH, 0.15 M NaCl, pH 7.4 buffer. All connections within the system were wrapped in parafilm. All glass-on-glass connections were sealed with vacuum grease. Once assembled, pure N2 gas was bubbled through the system to purge oxygen and create anaerobicity. N2 gas was passed through the system for 3 hours. At this point, the dropping funnel was carefully opened to allow dropwise addition of NaNO2 to the sulfuric acid. The resulting NO generated was bubbled through the system and was solubilized in the terminal buffer solution (1.88 mM at 295 K). NO generation was conducted over 1 hour. 40 2.2.18 Measurement of O2, NO, and H2O2 activity O2, NO, and H2O2 activity was all measured through the use of Clark-type electrodes (ISO-OXY-2, ISO-NOP-2, and ISO-HPO-2). These electrodes were connected to a TBR Free Radical Analyzer which displayed real time current data to the LabScribe software (iWorx, Dover, NH, USA v.4.0). Both the electrodes and radical analyzer were purchased from World Precision Instruments (Sarasota, Florida, USA). The electrodes were housed in sensor probes which were immersed in solution for measurement. All activity assays were performed in 50 mM HEPES, 0.15 M NaCl, pH 7.4 buffer. Assays were performed in plastic vials with constant stirring, and sealed with a rubber cap. Each sensor was allowed to equilibrate in buffer solution for at least 1 hour prior to use. To measure O2 activity, the sensor (ISO-OXY-2) was allowed to reach baseline at 25°C in aerobic conditions. After a stable baseline was established, enzyme was added (BCR or BCR-FDP) to a final concentration of 2 µM based on the concentration of bound flavin. The mixture was again allowed to equilibrate (5 min). Butyryl-CoA was then added in 100 µM injections to start the reaction. The O2 consumption over time was then measured via the current output. To track NO activity, the sensor apparatus and freshly prepared NO-saturated buffer were moved to the anaerobic glove box. Anaerobic buffer was used to equilibrate the sensor (ISO-NOP-2). After equilibration, NO-saturated buffer was sequentially injected into the assay vial (approx. 35 µM NO). 2 µM enzyme was added. After 5 min, 100 µM butyryl-CoA was added to the solution. NO concentration was followed over time using the current output. H2O2 activity was monitored using the H2O2 sensor (ISO-HPO-2). The H2O2 sensor was used to check for H2O2 activity through initial injections of 5 µM H2O2. 2 µM enzyme and 100 µM butyryl-CoA were then added sequentially. H2O2 concentration was then tracked over time using the current output. The product formation of BCR-FDP’s reaction with O2 was also examined with this sensor. The reduction of O2 is known in flavodiiron proteins to produce water. However, to confirm that the enzyme was not producing H2O2, the same protocol used to measure O2 reduction was performed in the presence of the H2O2 sensor. 41 2.2.19 Bioinformatic analysis AlphaFold (Google, DeepMind, v.4.0) was used to evaluate the flavin binding sites of BCR and the diiron site of FDP102,103. The sequence corresponding to BCD, ETF⍺, and the FDP were inputted into AlphaFold to generate structures. The generated BCD and ETF⍺ structures were used to the search Protein Data Bank (PDB) using the DALI server104. The resulting top hits from the PDB were then overlayed on the BCD and ETF⍺ structures. The flavin binding sites were analyzed and residues forming polar contacts with the flavin were identified. The diiron site of FDP was also analyzed. The FDP from E. coli (PDB: 5LLD) was overlayed on the AlphaFold model of FDP from F. nucleatum. The protein residues binding to the F. nucleatum FDP diiron centre were identified from this comparison. The phylogenetic distribution of the BCR-FDP dicistron was evaluated using the Genome Neighbourhood Tool from the Enzyme Function Initiative. The sequence of BCR (FNP 2074) was queried against the UniRef90 database. An e-value of 10-5 was used and a maximum of 1000 return sequences was set. The resulting search showed two clusters of hits of retrieved protein sequences, one with a size of ~380 residues and another, smaller one with a size of ~640 residues. Given that the size of FNP 2074 (BCR) is 636 residues, the second cluster was chosen for further analysis. A sequence similarity network (SSN) was generated using sequences ranging from 630-660 residues with an alignment score of 120. A genome neighbourhood diagram (GND) was also generated to determine the co-occurrence of BCR (FNP2074) and FDP, and the distribution of BCR amongst different taxa. 2.3 Results 2.3.1 Analysis of cofactor iron-binding sites of BCR and FDP based on sequence homology and protein models AlphaFold models were generated of the FDP and each of the domains of BCR as preliminary structural investigation into the cofactor binding sites. The AlphaFold models were then used to query the Dali server to find published protein structures with the highest-level of structural similarity. The level of structural similarity was assessed by the Z-score (and RMS deviation). Z-score must be above 8. This approach proved challenging for the Rd domain due to 42 its small size (6 kDa) yielding a low model confidence score (<90%). Nevertheless, BCR is predicted to bind an iron atom through its Rd domain. Rds have a conserved iron binding motif made up of 4 cysteine residues located in two CxxC motifs, where x is any amino acid. This motif is conserved in BCR (Cys589/Cys592 and Cys621/Cys624) and is depicted in an AlphaFold model of the domain (Figure 18A). The two CxxC motifs are separated by 29 residues, which classifies the Rd domain of BCR as a canonical Rd96,105,106. The diiron site of FDP (FNP2073) is similarly conserved in comparison to structurally determined flavodiiron proteins77. As shown in Figure18B, an AlphaFold model of the FDP shows that the two iron atoms are coordinated by a mixture of carboxylate and histidine ligands which is consistent with flavodiiron proteins found in E. coli, D. gigas, and M. thermoacetica74,77,107. Three carboxylate ligands (Glu81, Asp83, and Asp168) and four histidine ligands (His79, His84, His149, and His231) bind to the iron (Figure18B; numbering from FNP2073). Figure 18: The iron binding sites of BCR and FDP. A) An AlphaFold model of the Rd domain of BCR. The 4 cysteine residues which coordinate the iron atom are shown. B) A superposition of an AlphaFold generated diiron coordination sphere of FDP overlayed with the FDP of E. coli (PDB: 5LLD). Residues from F. nucleatum’s FDP are shown in red and those from E. coli’s FDP are shown in grey. 2.3.2 Flavin binding sites of FDP, BCD, and ETFa Using the AlphaFold model of the FDP as a query, the Dali server was used to identify the flavodiiron protein from E. coli (FDPEC; PDB: 1YCF) as having high homology to FDP. The A B Cys589 Cys592 Cys621 Cys624 His231 His227 His149 His147 His80 His79 His85 His85 Asp84 Asp83 Glu82 Glu81 Asp168 Asp166 43 alignment had a Z-score of 47.9 and an RMS deviation of 2.2 Å over 401 residues. These two proteins share only 24% sequence identity. An alignment of FDP-FDPEC showed clear tertiary structure similarity (Figure 19A). The FMN binding site was investigated and residues which formed hydrogen bonds to the isoalloxazine ring were identified (Figure 19B). The residues Asn319, Asn320, Ala353, Trp352, and Ser351 all likely form hydrogen bonds. Trp352 is also in position for p stacking interactions with the isoalloxazine ring. Figure 19: AlphaFold model of FDP compared with the flavodiiron protein from E. coli. A) A full overlay of FDP (red) with FDPEC (grey). B) The FMN binding site of FDP. Five residues are shown to form hydrogen bonds with the isoalloxazine ring. Bold type shows residues of F. nucleatum’s FDP. The PDB entry with the highest structural homology to an AlphaFold model of the BCD domain was found to be butyryl-CoA hydrogenase from Acidaminococcus fermentans (BCDAF; PDB: 4L1F)108. The structural alignment had a Z-score of 62.9 and an RMS deviation of 0.8 Å of the Ca carbons over 379 residues. BCD shares 57% sequence identity with BCDAF. The AlphaFold model of BCD was overlayed with BCDAF (Figure 20A). As the PDB file of BCDAF included FAD in the crystal structure, the flavin binding site of BCD and proximal residues that form hydrogen bonds to the isoalloxazine ring were located (Figure 20B). The residues that coordinate the flavin of BCDAF were conserved in BCD. These residues include Phe123 Leu125, Thr126, and Thr158 which form H-bond interactions with the isoalloxazine ring. Furthermore, Glu363, which functions as a A B Ser351 Gly346 Asn320 Asn315 Asn319 Asn314 Trp352 Trp347 Ala353 Gly348 44 catalytic base in the oxidation of butyryl-CoA, is also conserved92,109. The flavin binding site of BCDAF is sandwiched between a beta sheet-rich region and alpha helix-rich region which are both conserved in BCD. Figure 20: A comparison of the AlphaFold model of BCD with butyryl-CoA dehydrogenase from A. fermentans. A) The AlphaFold model of BCD (blue) overlayed on the dimeric structure of BCDAF (grey). BCDAF forms a dimer while BCD does not. B) A depiction of the FAD binding site of BCD. Residue numbering is conserved across BCD (blue) and BCDAF (grey). The Dali server identified ETF from Paracoccus denitrificans (ETFPD; PDB: 1EFP) as having the highest structural homology to ETFa110. The Z-score of the alignment was 21.6 with an RMS deviation of 0.9 Å for 61% of the Ca carbons of ETFa. An overlay of ETFa and ETFPD is shown in Figure 21A. Upon comparison of the ETFa and ETFPD alignment, two distinct structural features were apparent. First, ETFa, as expected, lacked homology to the majority of the ETFb subunit (domain III) of ETFPD. However, it did show homology to a lone alpha helix from subunit b, the C-terminal arm of ETFb. This alpha helix is positioned close to subunit a in full-length ETF proteins making up domain II. Second, ETFa was only composed of domain II, and lacked the domain I of typical ETFs110. Domain II is known to bind FAD while domain I performs a stabilization role helping to ensure close association between domain II and III. An overlay of the cofactor binding sites shows that residues neighbouring the isoalloxazine ring are conserved in ETFa which include Val491, Ser494, Arg467, and His541 (Figure 21B). A B Thr158 Phe123 Leu125 Thr126 Tyr362 Glu363 45 Figure 21: A comparison of the AlphaFold model of ETFa with the electron transfer flavoprotein from P. denitrificans. A) An overlay of ETFPD with ETFa (blue). ETF subunit a (grey) and ETF subunit b (purple) of ETFPD are shown. The conserved alpha helix used by ETFa “borrowed” from ETF subunit b can be seen. B) The FAD binding site of ETFa with the coordinating residues shown. Residues shown in blue are from ETFa and those in grey are from ETFPD. Bold residue labeling corresponds to ETFa with the numbering being consistent with full length BCR. In summary, these results establish that both FAD coordination spheres of BCR and the FMN site of FDP are conserved when compared to structurally similar homologs. This adds confidence to predictions of their function. In particular, since the amino acid residues surrounding flavin cofactors directly affects their reduction potential, we can make reasonable predictions on the thermodynamic favourability of electron transfer steps. 2.3.3 BCR and BCR-FDP purification and properties BCR and BCR-FDP were purified to homogeneity from BL21(DE3) cell pellets transformed with plasmid constructs pET15b_BCR and pET28a_BCR-FDP using NiNTA affinity chromatography. Protein yield ranged from 4-6 mg/L of culture for both BCR and BCR-FDP. Purity and protein size were confirmed with SDS-PAGE analysis (Figure 22). Purified BCR showed a single band with molecular weight of 69 kDa which closely matched the calculated A B Val491 Val241 Ser491 Thr241 Arg467 Arg227 His541 His264 46 weight of 70.9 kDa based on amino acid sequence. BCR-FDP showed two bands with sizes 69 kDa and 37 kDa which corresponded to BCR and FDP (FDP had a calculated mass of 36.6. kDa). This indicated that BCR and FDP co-purified as the cloning strategy placed a single His6 tag on the BCR N-terminus. Therefore, BCR and FDP form a protein complex. Figure 22: SDS-PAGE analysis of BCR-FDP and BCR. A) A 12% SDS-PAGE gel of purified BCR-FDP. Lane 1: PageRuler protein ladder. Lane 2: BCR-FDP (~1 mg/mL). Two bands of sizes 69 and 37 kDa are visible which match the calculated sizes of BCR (70.9 kDa) and FDP (36.6 kDa). B) A 12% SDS-PAGE gel of purified BCR. Lane 1: PageRuler protein ladder. Lane 2: BCR (~1 mg/mL). To further investigate protein complex formation, size exclusion chromatography was performed. Both BCR and BCR-FDP were run over a size exclusion column. The elution volumes (Ve) of BCR and BCR-FDP were 57.4 and 52.2 mL. The Ve values were inputted into Equation 1 and converted to Kav values. A standard curve of Kav vs molecular weight was constructed with proteins of known weight. The Kav values of BCR and BCR-FDP were inputted into the equation of the line to calculate molecular weight. BCR had a molecular weight of 315 kDa and BCR-FDP had a molecular weight of 489 kDa. These values are much higher than the bands observed via SDS-PAGE analysis indicating that BCR and BCR-FDP form multimers. Specifically, both BCR and BCR-FDP form tetramers as their molecular weights calculated from the standard curve are approximately 4 times the size of their known monomeric weights. BCR likely forms a homotetramer (a4) and BCR-FDP likely forms a heterotetramer (a4b4). 1 2 1 2 69 kDa BCR 37 kDa FDP A B 69 kDa BCR 70 kDa 40 kDa 47 2.3.4 BCR and BCR-FDP UV-vis spectroscopy Both BCR and BCR-FDP were predicted to copurify with several flavin cofactors. BCR should contain two FAD molecules per polypeptide while BCR-FDP should contain two FAD and 1 FMN molecules per polypeptide. The BCR and BCR-FDP eluate from the Ni-NTA column had a distinct yellow colour which served as initial evidence for the presence of flavin cofactor. Purified protein extracts of BCR and BCR-FDP were subjected to UV-vis spectroscopy. The initial spectra of both proteins are seen in Figure 23. These spectra are dominated by bands II and I of the flavin cofactors known to absorb at ~375 and 450 nm, respectively, further confirming the presence of flavin moieties111. Protein concentration was determined through the Pierce BCA assay using BSA as a standard. After protein quantification, the extinction coefficients at 280 nm of BCR and BCR-FDP were calculated as 45.1 mM-1cm-1 and 103.4 mM-1cm-1, respectively. Initially, LB media was chosen as the growth media for the production of recombinant proteins. However, the UV-visible spectra of BCR and BCR-FDP indicated limited iron incorporation into the Rd domain. Rd with the iron in the ferric state is known to have peaks at 360, 490, & 580 nm112. However, as shown in Figure 23, there was no detectable presence of Rd peaks. Specifically, the spectra lacked a 580 nm peak which is not obscured by bands I and II of the flavins. These data indicated little to no iron incorporation occurred when transformed cells harbouring BCR or BCR-FDP were grown in LB media. It has been shown for Rd that zinc can bind and contaminate its active site74,113. Therefore, it was hypothesized that zinc was outcompeting iron. As iron is essential for catalysis, the expression medium was switched to M9 minimal media, a chemically defined media that has no added zinc. The use of M9 allowed for closer control of the metal ions present. To further promote iron incorporation, the M9 media was supplemented with 400 µM ferrous iron. As shown in Figure 24, this expression strategy resulted in incorporation of iron into the Rd motif, as evidenced by a broad band at ~580 nm in both BCR and BCR-FDP. Additionally, band I of the flavin peak (normally ~450 nm) showed a more prominent shoulder at 480 nm, likely arising from an overlapping signal from Rd which exhibits a 490 nm peak. Together, these two spectral features indicated higher iron incorporation into the Rd motif. 48 Figure 23 : UV-visible spectra of BCR and BCR-FDP grown in LB and M9 minimal media. A) The UV-vis spectra of BCR purified from LB media (blue) and M9 minimal media (black). Peaks are seen at 378, 383 and 448 nm. Rd UV-vis spectral properties are seen in M9 minimal media with a prominent shoulder at 480 nm and a distinct broad peak at 580 nm. B) The UV-vis spectra of BCR-FDP purified from LB media (blue) and M9 minimal media (black). Peaks are seen at 400 and 448 nm. Similar Rd UV-vis properties to BCR are observed. Purified BCR and BCR-FDP were dialyzed against 50 mM Tris-HCl, 0.15 M NaCl, pH 7.4 buffer following elution from the Ni-NTA column. The UV-visible spectra of BCR acquired after a 16-hour dialysis yielded a red-shift in the lmax of band II of the flavin cofactor (448 nm to 463 nm, Figure 24A). This longer wavelength absorbance maxima is indicative of modification at the 8-methyl group of the flavin isoalloxazine ring46,114. Previous studies have shown that the 8-methyl group of FAD coordinated to the alpha domain of ETF is known to react with dioxygen to form an 8-formyl group, which can further be modified to 8-hydroxy or 8-amino flavin46,115. The 433 nm peak also observed in the dialyzed BCR sample is diagnostic of the anionic semiquinone of 8-formyl flavin, which has been shown to have a higher reduction potential (E¢ox/asq -47 mV) compared to the unmodified FAD cofactor (E¢ox/asq -61 mV) bound to the alpha subunit of ETF46 In fact, the reduction potential of 8-formyl FAD (FAD/FAD-•) is elevated enough such that the one electron reduced form is observed in aerated buffer46,115. To validate the presence of modified flavin, BCR was heated to 95°C for 5 min to release free flavin. Free authentic FAD exhibits lmax at 370 and 450 nm while free authentic FMN has peaks at 370 and 446 nm100. The denatured BCR A B 49 sample released free flavin with lmax at 368 and 466 nm for bands I and II (Figure 24A, inset). The shift in the lmax values from that of authentic flavin wavelengths, confirms that the FAD cofactor was being chemically modified during dialysis46,115. EPR was also used to confirm that the FAD cofactor was modified. EPR is a widely used technique to identify and characterize lone electron radical species such as the semiquinone of flavin cofactors. As shown in Figure 25, the dialyzed BCR sample exhibited an EPR signal arising from the air-stable anionic semiquinone. The spectrum had a g-factor of 2.004 which confirmed the signal was due to an organic radical (flavin semiquinone)116. The spectrum also exhibited a peak-trough width of 13 G. Unmodified FAD of the ETF alpha subunit in the anionic semiquinone state has a diagnostic peak-trough width of 15 G while modified flavin has a peak-trough width of 13 G46,115. Thus, the EPR data further confirms the presence of modified flavin in BCR. To prevent flavin modification in BCR, the protein was purified in an anaerobic glove box. A UV-vis spectrum was taken directly after elution from the anaerobic column yielded slight peaks at 354 and 423 nm (Figure 24B). These absorbance maxima are indicative of reduced flavin (i.e. the cofactor is in the hydroquinone state). Oxidation of this sample with potassium ferricyanide gave lmax values at 376, 396, and 444 nm; these absorbance features arise from the spectral overlap of the oxidized, unmodified flavin and the Fe-Cys4 cofactor of the Rd domain. Heating anaerobic BCR to release the flavin cofactors yielded a band II lmax at 452 nm. This value is also closely associated with authentic FAD. EPR was used to further compare flavin modification. Anaerobic BCR was chemically reduced with dithionite to generate the anionic semiquinone. An EPR spectrum of anaerobic BCR in the semi-reduced state had a peak-trough width of 16 G with a g-factor of 2.004 (Figure 25). The peak-trough width closely matches authentic FAD and the g-factor confirms the presence of flavin semiquinone. The above results confirm that chemical modification of the a-FAD is dependent on O2. Accordingly, all further experiments of BCR used the anaerobically purified form of the protein. In contract to BCR, BCR-FDP did not show any flavin modification after dialysis as determined by UV-vis (Figure 26 ). The spectrum before dialysis gave lmax at 389 and 449 nm which remained constant after aerobic, overnight dialysis. These UV-vis peaks are comparable to unmodified band I and II lmax peaks. It is likely that in its full tetrameric form, BCR-FDP’s a-FAD is less solvent exposed compared with BCR, minimizing modification. 50 Figure 24: Optical spectra of BCR. A) UV-vis spectra of aerobically purified BCR before (black line) and after (red line) aerobic dialysis. Inset: UV-vis spectrum of free flavin released from denatured BCR. Modified flavin was present due to 368 and 466 nm peaks. B) UV-vis spectra of BCR purified anaerobically. The black spectrum was taken directly after column purification and the red spectrum was taken ~20 hours after purification. Inset: UV-vis spectrum of free flavin released from anaerobic BCR. Unmodified FAD was deemed present due to 444 nm and 452 nm peaks. Figure 25: EPR spectrum of BCR purified aerobically and anaerobically. A) EPR spectrum of BCR purified aerobically. The calculated g-factor and peak-trough width are 2.004 and 13 G. B) EPR spectrum of BCR purified anaerobically. The calculated g-factor and peak-trough width are 2.004 and 16 G. A B B A 51 Figure 26: Flavin bound to BCR-FDP is not modified in the presence of oxygen. A) The UV-vis spectra before (black line) and after dialysis (red line) are the same. Inset: UV-vis spectrum of flavin released from BCR-FDP. 2.3.5 Cofactor incorporation in BCR and BCR-FDP ICP-MS was used to quantitate metal content in BCR and BCR-FDP expressed in both LB and M9 minimal media. The absolute quantity of iron and zinc in each protein sample was converted to molar concentration and compared to the protein concentration. The calculated molar ratios of iron to protein are presented in Table 3. The expected ratios of iron to protein are 1:1 for BCR (Fe-Cys4 group of Rd domain) and 3:1 for BCR-FDP (Rd domain and diiron site). In LB media, the ratios of iron to protein were 0.04:1 and 0.23:1 for BCR and BCR-FDP respectively, well below expected values. The zinc ratios were high in LB media with ratios of 0.80:1 and 0.25:1 for BCR and BCR-FDP. The high levels of zinc, particularly in BCR, shows the propensity of the Rd motif to bind zinc. Previous work with Rd from Clostridium pasteurianum has also found contaminating zinc113. The authors found that the presence of zinc was irrespective of protein purification in a homologous or heterologous system. Iron incorporation was improved in both BCR and BCR-FDP upon switching to iron-supplemented M9 minimal media. This approach yielded expected iron to protein ratios of 0.91:1 52 and 2.2:1 for BCR and BCR-FDP, respectively. The switch to M9 media dropped the zinc incorporation to near zero in BCR (0.05:1). Zinc incorporation was still observed in BCR-FDP in M9 media (0.72:1). However, it was assumed that a fraction of the protein population had sufficient iron content to be catalytically active. Table 3: Ratios of cofactor to protein for BCR and BCR-FDP. Expected ratios are in parentheses. Protein BCR BCR-FDP Media LB M9 LB M9 [Iron]:[Protein] 0.04:1 (1:1) 0.91:1 (1:1) 0.23:1 (3:1) 2.2:1 (3:1) [Zinc]:[Protein] 0.80:1 0.05:1 0.25:1 0.72:1 [FAD]:[Protein] 0.25:1 (2:1) 0.24:1 (2:1) 0.83:1 (2:1) 0.51:1 (2:1) [FMN]:[Protein] - - 0.65:1 (1:1) 0.51:1 (1:1) An FAD is predicted to bind to each of the BCD and ETFa domains of BCR giving an expected ratio of 2 molecules of FAD per molecule of protein. FDPs are known to bind FMN with an expected ratio of 1 FMN per protein. Flavin content was determined through use of fluorescence spectroscopy. This technique takes advantages of the fact that a solution of FMN has a 10-fold greater quantum yield compared to an equimolar solution of FAD. Protein samples were heated to release bound flavin and an initial spectrum was taken. The sample was digested with phosphodiesterase which converts FAD to FMN. This method determines the concentration of FAD and FMN present in a protein sample. The levels of flavin incorporation were low across both media type and protein (Table 3). The ratios of FAD to BCR were 0.25:1 and 0.24:1 in LB and M9 media. In BCR-FDP, FAD had ratios of 0.83:1 and 0.51:1 in LB and M9 media with FMN staying consistent at 0.65:1 and 0.51:1 across media type. This was despite the addition of free flavin to crude protein extract prior to purification by Ni-NTA chromatography. Comparing BCR to BCR-FDP, higher levels of FAD incorporation were observed in BCR-FDP with at least double the amount of bound FAD. BCR-FDP also incorporated FMN easier than FAD. The ratio of FMN to protein more closely approached the expected ratio of 1:1 than FAD to protein approached the ratio of 2:1, indicating better retention of this cofactor in the flavodoxin motif of the FDP. With the flavin quantitated, the 53 extinction coefficients were calculated. FAD bound to BCR had an extinction coefficient of 11.3 mM-1cm-1 at 454 nm while BCR-FDP had an extinction coefficient of 9.3 mM-1cm-1 at 473 nm. 2.3.6 ETFa and BCD domain purification and properties To help disentangle the properties of the multidomain protein BCR, the BCD and ETFa domains were cloned separately into expression vectors. These vectors (pET28a_BCD and pET28a_ETFa) were transformed into E. coli BL21(DE3) cells. Following growth of the transformed cells in LB media, the recombinant proteins were purified using Ni-NTA chromatography. As shown in Figure 27, both BCD and ETFa were >95% pure as judged by SDS-PAGE. BCD and ETFa appeared as single bands with molecular weights of 40 and 20 kDa respectively. These matched the calculated molecular masses of 41.3 and 20.1 kDa. Purified BCD was yellow, and the UV-vis spectrum of the protein exhibited peaks at 370 and 446 nm, characteristics of a flavoprotein (Figure 28A). Free free flavin following heat denaturation of BCD showed an UV-vis spectrum that closely resembled authentic FAD with a lmax at 449 nm. Using an extinction coefficient of 12.3 mM-1cm-1 for free FAD, the extinction coefficient of FAD bound to the BCD domain was calculated to be 14.9 mM-1cm-1. The extinction coefficient at 280 nm was calculated to be 31.6 mM-1cm-1 for BCD. Figure 27: SDS-PAGE analysis of BCD and ETFa. A) A 12% SDS-PAGE gel of purified BCD. Lane 1: PageRuler protein ladder. Lane 2: BCD (~1 mg/mL). A single band of 41 kDa is observed. B) A 12% SDS-PAGE gel of purified ETFa. Lane 1: PageRuler protein ladder. Lane 2: ETFa (~1 mg/mL). A 20 kDa band is observed. 20 kDa 41 kDa BCD 2 2 1 1 A B 40 kDa 20 kDa ETFa 54 In contrast to BCD, purified ETFa was orange in colour and the optical spectrum of the protein exhibited a prominent peak at 482 nm and several smaller peaks clustered around where band I normally absorbs (Figure 28B). It was evident that the FAD bound to the ETFa domain was also being chemically modified. There is a literature precedent for modified flavin occurring in ETFs46,115. As only the ETFa domain, not the BCD domain, showed a modified FAD, the modified FAD previously observed in BCR was assigned to the a-FAD coordinated to the ETFa domain. Heating of the ETFa domain gave a lmax at 486 nm which is also indicative of modified FAD (Figure 28B). Figure 28: UV-vis spectra of BCD and ETFa showing bound and unbound flavin. A) The UV-vis spectra of BCD after purification (black line) and after boiling of the protein sample (red line). The boiling process releases the flavin cofactor. Upon boiling band II of the flavin shifts from 446 nm to 449 nm which is indicative of FAD. B) The UV-vis spectra of ETFa after purification (black line) and after boiling of the protein sample (red line). The lack of a canonical flavin spectrum after purification (a disparate collection of peaks) as well as the peak at 486 nm indicates the presence of modified flavin cofactor. 2.3.7 Anaerobic purification of ETFa As purification of BCR under anaerobic conditions prevented flavin modification, the same approach was used for ETFa. Elution of ETFa from the Ni-NTA column in the glove box B A 55 exhibited peaks 380 and 356 nm. The peak at 380 nm is indicative of the anionic semiquinone while the 356 nm band is indicative of the two-electron reduced hydroquinone. The prominent peak at 736 nm is likely a charge-transfer band between a reduced hydroquinone and quinone species of two juxtaposed a-FAD cofactors. This conclusion is based on previous work with EtfAB from Rhodopseudomonas palustris where a similar peak was attributed to a charge-transfer complex forming from the interaction of the flavins of ETFa and ETFb, one in the hydroquinone state and the other in the fully oxidized quinone state117. As only one flavin is present in ETFa, the charge-transfer band observed in Figure 29 is ascribed to the juxtaposition of two ETFa molecules, one in the quinoneox state and the other in the hydroquinonered. Oxidation of ETFa with ferricyanide caused the 736 nm peak to disappear. This establishes that the band is redox dependent. Oxidation also caused the appearance of 398 and 450 nm bands; these correspond to band II and band I of the flavin cofactor (ETFa showed a smaller peak-trough width compared to anaerobically purified ETFa. While the peak-trough width differs in magnitude from BCR, the trend of a modified FAD giving a lower width is conserved.). Here, band II had a larger absorbance amplitude than band I, a property unusual for flavoproteins. However, this has been observed in canonical ETFs73,93. Heat-denaturation of anaerobically purified ETFa yielded a free flavin spectrum very similar to authentic FAD, with lmax at 372 and 452 nm (Figure 29, inset). However, the presence of a shoulder at 528 nm, indicates that a small population of the flavin still being chemically modified. The calculated extinction coefficient of FAD bound to ETFa was 11.6 mM-1cm-1 and the protein extinction coefficient at 280 nm was calculated as 12.4 m mM-1cm-1 (Table 3). 56 Figure 29: UV-vis spectra of various redox states of anaerobically purified ETFa. The spectrum after purification (black line) and the spectrum after complete oxidation (red line) are shown. Inset: UV-vis spectrum of flavin released after heating of ETFa. Like for BCR, EPR spectra were taken of both aerobically and anaerobically purified samples of ETFa to confirm the presence/absence of modified flavin (Figure 30). ETFa was partially reduced with dithionite to generate the semiquinone (the EPR active species). Aerobic ETFa gave a peak-trough width of 15 G while anaerobic ETFa gave a value of 18 G. Both samples gave g-factors of 2.004 confirming the signal is due to semiquinone. These peak-trough width values are approximately 2-3 G higher than seen with full-length BCR; however, the results mirror that observed for BCR, where aerobically purified ETFa showed a smaller peak-trough width compared to anaerobically purified ETFa. While the peak-trough width differs in magnitude from BCR, the trend of a modified FAD giving a lower width is conserved. 57 Figure 30: Comparison of EPR spectra of aerobic and anaerobic ETFa. A) EPR spectrum of aerobic ETFa. The spectrum has a peak-trough width of 15 G and a g-factor of 2.004. B) EPR spectrum of anaerobic ETFa. The spectrum has a peak-trough width of 18 G and a g-factor of 2.004. Table 3: Extinction coefficients corresponding to 280 nm and the maximum flavin peak for each protein Protein BCR BCR-FDP BCD ETFa eprotein (mM-1cm-1) 45.1 103.4 31.6 12.4 ebound flavin (mM-1cm-1) 11.3 9.3 14.9 11.6 2.3.8 Reduction oxidation titrations of BCD, ETFa, BCR, and BCR-FDP Redox titrations were performed of BCD, ETFa, BCR, and BCR-FDP to investigate the redox states of the flavin cofactors. The reductant dithionite or the oxidant potassium ferricyanide were used to transition the flavin cofactors through quinone, semiquinone, and hydroquinone oxidation states. Sequential reduction of fully oxidized BCD resulted in a decrease in the lmax at 372 and 446 nm peak and an increase at 567 nm. This spectral transition corresponds to conversion A B 58 of the oxidized quinone state of the flavin to the neutral semiquinone (Figure 31A). The formation of the semiquinone is denoted by black spectra. Upon further reduction, this broad band centered at 567 nm decreases as do the lmax at 372 and 446 nm. This phase of the titration denotes conversion of the semiquinone to the hydroquinone. The loss of the semiquinone is marked by blue spectra (Figure 31A). Evidence of the d-FAD forming the blue semiquinone indicates that the cofactor is able to participate in single electron transfer. Given that the optical spectrum of ETFa following anaerobic purification indicated that the protein was partially reduced, titration of the protein began by fully reducing the protein using dithionite. Fully reduced ETFa was then slowly re-oxidized using ferricyanide. The UV-vis spectra showed formation of the red anionic semiquinone at 370-380 nm (Figure 31B) following the first equivalent of potassium ferricyanide. During this initial phase of the redox titration, a peak formed at 736 nm. Peak formation at 736 nm coincided with oxidation of the hydroquinone to the anionic semiquinone. The peak remained constant while the semiquinone fully formed and then finally disappeared with further addition of potassium ferricyanide and the complete oxidation of the flavin to the quinone. The spectra are coloured coded to follow the formation (grey), constant region (blue), and loss (red) of the 736 nm peak (Figure 31B). As previously described, this 736 nm peak is ascribed to a charge-transfer complex as a result of the interaction of two FAD species: one in the oxidized quinone state and the other in the reduced hydroquinone state46. The appearance and disappearance of this peak over the titration’s course as the FAD’s redox state changes fits this hypothesis. However, this charge-transfer complex is likely non-physiological as the a-FAD in ETFa is part of a larger protein complex in vivo, which would ostensibly prevent Van der Waals contact between two a-FADs. The titration data show that ETFa forms a stable anionic semiquinone and thus participates in single electron transfer. With the titrations of BCD and ETFa serving as a reference, the titration of BCR was conducted. Expectedly, the UV-vis spectra of full length BCR bare resemblance to the spectra of individual domains (Figure 31C). Addition of potassium ferricyanide resulted in the formation of a peak at 371 nm denoting formation of the red anionic semiquinone. This anionic semiquinone is ascribed to the a-FAD of ETFa. Further addition of ferricyanide led to full oxidation with the formation of the quinone state of the flavin cofactors. Peaks at 396 and 444 nm are indicative of quinone formation. Due to the level of baseline drift/noise in the spectra, no clear 580 nm peak was observed for the blue semiquinone of BCD was identified. The UV-vis spectrum of fully 59 oxidized BCR can be visualized as a composite spectrum of ETFa and BCD. ETFa presents as an atypical flavin spectra with band I having a higher absorbance than band II while BCD shows more a typical ratio with band II having the higher absorbance. The different amplitudes associated with band II of a-FAD and d-FAD result in a spectrum of BCR that is relatively flat across the top with peaks at 396 and 444 nm. Understanding the composite nature of BCR’s spectrum was aided through the purification and spectral analysis of the isolated ETFa and BCD domains. Finally, a redox titration was performed for BCR-FDP. The titration was split into three distinct phases, each denoted by a different colour (Figure 31D). The first phase (red) showed a reduction at wavelengths 390, 454, and 580 nm with the addition of dithionite. The peak 390 nm gradually shifted towards 380 nm during the first phase. During this phase of the titration, the oxidized flavin cofactors were reduced. In particular, the anionic semiquinone of ETFa was formed as shown by the formation of a peak at 380 nm. The Rd ferric iron centre was also reduced to the ferrous form as evidenced by the loss of all peak definition at 580 nm. The second phase (blue) was characterized by constant absorbance at 383 nm with a further decrease at 454 nm. Here, more anionic semiquinone was formed as all remaining oxidized flavin was reduced. At this stage, the electrons were primarily located on the ETFa FAD and Rd Fe(SCys)4 motif. The final phase (black) again showed a constant absorbance at 383 nm while the peak 454 peak continued to diminish. In this final phase, a second anionic semiquinone was formed and is attributed to the one-electron reduction of the FMN of the FDP74. The multiple phases associated with the titration suggest separate flavin semiquinone species are being formed. In an attempt to distinguish between the anionic semiquinone of ETFa’s FAD and the anionic semiquinone of the FMN of FDP, further EPR experiments were conducted. 60 Figure 31: Redox titrations of BCD, ETFa, BCR, and BCR-FDP. A) Redox titration of BCD. The formation (black) and loss (blue) of a 567 nm peak are highlighted. B) Redox titration of ETFa. This titration started from the fully reduced state (bottom grey spectrum). The colour coding marks the formation (grey), constant region (blue), and loss (red) of the 736 nm peak. C) Redox titration of BCR. The black spectra mark the formation of the anionic semiquinone. Blue spectra show the conversion of anionic semiquinone to oxidized quinone. D) Triphasic redox titration of BCR-FDP. The red spectra show conversion of oxidized flavin to ETFa’s anionic semiquinone and reduction of Rd. The blue spectra show complete conversion to anionic semiquinone of ETFa. The final phase (black) shows the formation and eventual reduction of the anionic semiquinone of the FMN of FDP. A B C D 61 EPR spectra were taken of BCR-FDP during phase 2 and 3 of the titration, in an effort to identify different semiquinone species, as the peak-trough width can vary depending on the flavin binding environment118. The spectrum of phase 2 was anticipated to capture the anionic semiquinone of ETFa while the phase 3 spectrum expected to report on the anionic semiquinone of the FMN in the FDP. An EPR spectrum taken during phase 2 yielded a peak-trough width of 17 G while phase 3 yielded a width of 15 G (Figure 32). A width of 17 G aligned closely with the previously taken EPR spectrum of ETFa (Figure 30). Therefore, the anionic semiquinone observed during phase 2 was assigned to the FAD of ETFa and the phase 3 anionic semiquinone was assigned to the FMN of FDP. These data indicate that the midpoint reduction potential of a-FADox/sq is more positive than FMNox/sq. Figure 32: EPR spectra of BCR-FDP taken at two different points in a redox titration. A) An EPR spectrum taken during phase 2 of BCR-FDP’s redox titration (see Figure31D, blue traces). The calculated g-factor and peak-trough width are 2.004 and 17 G. B) An EPR spectrum taken during phase 3 of BCR-FDP’s redox titration (see Figure31D, black traces). The calculated g-factor and peak-trough width are 2.004 and 15 G. 2.3.9 Pre-steady state kinetic analysis of butyryl-CoA’s reduction of BCD and BCR Stopped-flow spectroscopy was used to measure observed rate constants associated with butyryl-CoA reduction of the BCD domain and full length BCR. Figure 33A shows time-resolved A B 62 spectra of BCD following rapid-mixing with a 10-fold excess of butyryl-CoA acquired over 5s. Only a 30% reduction in the d-FAD absorbance maxima was observed indicating that butyryl-CoA is unable to fully reduce the cofactor. In contrast, full reduction was observed during the redox titration of the BCD domain using dithionite (Figure 31A). The incomplete reduction is attributed to the thermodynamically unfavourability of butyryl-CoA reduction of the BCD d-FAD cofactor. The reduction potential of butyryl-CoA/crotonyl-CoA is -10 mV (E¢°)119,120 while the potential of the FADox/sq couple (-70 mV; Megasphaera elsdenii) of BCD from has been shown to be more electronegative121. The observed rate constants for reduction of the BCD domain were determined by fitting single wavelength experiments acquired at 450 nm to a double exponential equation (Equation 6) (Figure 33B). The observed rate constants for butyryl-CoA’s reaction with the BCD domain were 7.1 ±0.1 and 1.1 ±0.1s-1. The amplitudes associated with each of these phases are ~0.015-0.011 AU and 0.011-0.008 AU respectively as shown in Figure 33B. The observed rate did not change significantly as a function of butyryl-CoA concentration (50-200 µM) indicating the dissociation constant (Kd) for BCD-butyryl-CoA is < 50 µM (Figure 33B; inset). Figure 33: Stopped-flow data from reaction of BCD domain with butyryl-CoA. A) Multiwavelength data from rapid mixing of 5 µM BCD with 100 µM butyryl-CoA. A decrease in absorbance at 446 nm is indicative of BCD reduction. Black trace represents mixing of 5 µM BCD with buffer. B) Single wavelength trace at 450 nm from rapid mixing of 5 µM BCD with 100 µM butyryl-CoA over 5 seconds. Inset: Observed rate constants are plotted against butyryl-CoA concentration in an attempt to determine Kd. A B 63 Stopped-flow spectroscopy was also used for pre-steady state kinetic analysis of BCR’s reaction with butyryl-CoA. As shown in Figure 34, rapid mixing of BCR with a 7-fold excess of butyryl-CoA resulted in conversion of a fraction of the oxidized flavin to the anionic semiquinone as evidenced by as partial reduction in the peak 444 nm and an increase at 380 nm. These time-resolved spectra indicated that butyryl-CoA reduction of d-FAD of the BCD domain is accompanied by single electron transfer to a-FAD to form the anionic semiquinone (denoted by the 380 nm peak). The absence of spectral data between the initial spectrum, acquired by mixing BCR with buffer (Figure 34, first black trace), and the first recorded trace (acquired at 1.6 ms) indicates that the observed rate constant for formation of the anionic semiquinone is >1000 s-1. This contrasts with the BCD domain where the observed rate constants for reduction of d-FAD are 7.1 and 1.1 s-1. It is likely the butyryl-CoA dependent reduction of BCR is considerably faster because it is tightly coupled to rapid interflavin electron transfer to the a-FAD. The reduction potential of the FADox/asq of ETFa is higher compared to other flavins, which favours rapid electron transfer122. Figure 34: Multiwavelength stopped-flow data of BCR and butyryl-CoA. Reduction occurs primarily within the dead time of the instrument as shown by the gap between the first trace (5 µM BCD with buffer) and the second. Build-up at 370 nm shows formation of an anionic semiquinone. 64 2.3.10 Catalytic activity of BCR-FDP with NO, O2, and H2O2 FDPs are known to react with two diatomic substrates: NO and O276. Some FDPs show higher specificity for NO or O2 or show near equal catalytic efficiency for both molecules. The diiron active sites for NO and O2 reductases are largely conserved, so the structural rationale for selectivity remains elusive. There has also been some recent evidence that FDPs can also reduce H2O2 with the FDP from Syntrophomonas wolfei showing the highest activity for H2O2 reduction ever recorded for an FDP 87. Given the differences in substrate scope amongst flavodiiron proteins, it was crucial to determine if BCR-FDP was able to reduce NO, O2, and/or H2O2. The rate of consumption in solution of all three substrates was monitored using Clark-type electrodes which measured current in real-time. Current was a measure of substrate concentration. Sequential additions of substrate, enzyme, and then butyryl-CoA were used for each substrate. The first substrate tested was NO. NO-saturated buffer was used as the source of NO and activity was tested under anaerobic conditions. NO-saturated buffer was added in three injections (Figure 35). Following addition of 2.5 µM BCR-FDP to the solution, there was no observable change in NO concentration. The subsequent addition of butyryl-CoA also showed no change in current. This experiment was repeated with from three separately purified samples of BCR-FDP and with commercially acquired butyryl-CoA (Sigma), and in all assays, no NO reductase activity was detected; This indicated that BCR-FDP was not able to reduce NO. 65 Figure 35: NO activity of BCR-FDP. NO concentration is monitored with an NO Clark-type electrode. Sequential additions of approximately 30 µM NO, 2.5 µM BCR-FDP, and 100 µM butyryl-CoA show no significant decrease in NO concentration (current). Next, the ability of BCR-FDP to reduce O2 was tested. The same experimental methodology to NO activity was used except the experiment was performed under aerobic conditions and with an O2-selective electrode. A baseline of O2 concentration was established by letting the Clark-type electrode equilibrate in aerobic buffer (50 mM HEPES-NaOH, 0.15 M NaCl, pH 7.4) for 1 hour at 20°C. Addition of 2.5 µM BCR-FDP to this buffer showed no change in this baseline (Figure 36A). Further addition of 100 µM butyryl-CoA initiated a steep decrease in O2 concentration. This decrease was reproducible upon repeated additions of 100 µM butyryl-CoA. Given that reduced flavin cofactors can also directly reduce O2 (forming H2O2), the assay was repeated with BCR instead of BCR-FDP46,114,115. The assay with BCR showed no change in O2 concentration upon addition of butyryl-CoA (Figure 36B). Therefore, the decrease in O2 concentrations seen with BCR-FDP is attributable to O2 reductase activity at the diiron site. 66 Figure 36: O2 reductase activity of BCR-FDP. A) O2 concentration is monitored through use of an O2 sensor. Addition of butyryl-CoA to an aerobic BCR-FDP solution shows a decrease in O2 concentration (current). B) Addition of butyryl-CoA to an aerobic solution of BCR shows no change in O2 concentration. This control shows that the decrease in O2 concentration in A) is not due to flavin cofactors directly reducing O2 FDPs that have been characterized to date have identified water as the product of O2 reduction73,123. However, O2 could also partially reduce to H2O2. To confirm the product of the BCR-FDP reaction, the O2 activity experiment was repeated with an H2O2 electrode. As shown in Figure 37, no changes in H2O2 levels were observed upon addition of 100 µM butyryl-CoA and BCR-FDP. Thus, the product of BCR-FDP’s catalyzed-reduction of O2 is water. A B 67 Figure 37: The reaction of BCR-FDP with butyryl-CoA does not produce H2O2. H2O2 concentration is monitored with an H2O2 sensor. Three additions of butyryl-CoA do not show an increase in H2O2 concentration. The H2O2 sensor was also used to test for the ability of BCR-FDP to catalyze the reduction of H2O2 with butyryl-CoA serving as the reductant. To perform this experiment, an H2O2 sensor was placed in buffer (50 mM HEPES-NaOH, 0.15 M NaCl, pH 7.4) and 5 µM injections of H2O2 were added sequentially. 2.5 µM BCR-FDP was added and a large decrease in H2O2 concentration was observed (Figure 38A). Addition of butyryl-CoA saw the same rate of decrease as just enzyme. As the addition of enzyme led to the decrease in H2O2 rather than butyryl-CoA, this observation is due to H2O2 directly reacting with protein rather than being reduced at the diiron site. This was confirmed by repeating this experiment with BCR. A similar decrease was seen (Figure 38B). Therefore, BCR-FDP is specific only for O2 and shows no NO or H2O2 reductase activity. 68 Figure 38: The reaction of butyryl-CoA and BCR-FDP does not consume H2O2. A) H2O2 concentration is monitored in real-time with an H2O2 sensor. Sequential additions of 5 µM H2O2 produce a current response. Addition of BCR-FDP sees a steep decrease in H2O2 concentration. Addition of butyryl-CoA sees no observable change in the rate of decrease. B) The same steep decrease in H2O2 concentration is seen with BCR. This steep decrease can be attributed to H2O2’s direct reaction with protein. 2.3.11 The prevalence of the BCR-FDP operon across bacterial species Bioinformatic tools were used to assess the frequency at which the genes encoding BCR and FDP appear in bacterial genomes as a dicistron. Tools available through the Enzyme Function Initiative (EFI) were used for this analysis (https://efi.igb.illinois.edu/)124,125. Option A of the EFI-EST tools was used in which the protein sequence of BCR (FNP2074) was used to retrieve homologs with an e-value <10-5 from the UniRef90 database. Only protein sequences from bacteria were considered, using the taxonomy filter. This returned 834 unique sequences which clustered in two different sequence lengths: 380-390 and 635 amino acids (Figure 39). Given that the size of FNP2074 is 636 amino acids, the smaller cluster was excluded from further analysis. This reduced the number of sequences from 834 to 416. The large cluster was used to generate a sequence similarity network (SSN) from which a genome neighbourhood diagram (GND) was constructed. The SSN had 513 unique nodes. A B 69 Figure 39: Histogram of homologous sequences to BCR. Number of sequences is plotted against sequence length. Two distinct clusters emerge: one centered at 380 residues and the other at 635 residues. The SSN had a single cluster of proteins with 513 nodes where each node is one homolog. The SSN was then used to generate a GND. Ten genes upstream and downstream of each node were used to construct the GND. Examination of the diagrams revealed that in each of 416 sequences a BCR homolog colocalized with an FDP homolog, which suggests that BCR evolved to serve as a redox partner for FDP and does not have additional cellular functions (Figure 40B). The dicistron appears to be primarily found in anaerobic organisms of the gut microbiota. These include butyrate-producers of the Firmicute/Bacillota phyla (Clostridium, Brachyspira, Instestimonas, Eubacteria, Anaerococcus, Peptoniphilus) and members of the Fusobacterium genus. Thus, it appears the BCR-FDP dicistron serves to maintain anaerobicity and/or remove oxidative molecules in the mammalian gut. Furthermore, these species are thought to contribute to gut health through butyrate production which only occurs under anaerobic conditions57,126,127. Because of this, BCR-FDP might be beneficial to gut health by helping lower O2 concentrations, preventing outgrowth of pathogenic facultative anaerobes (e.g. E. coli, Salmonella typhimurium). 70 Figure 40: Prevalence of the BCR-FDP dicistron across bacterial species. A) A taxonomy distribution diagram of the SSN generated for BCR. Clostridia and Bacilli classes make up the most hits. B) A sample of the GND generated from the SSN of BCR. The black rectangle highlights BCR homologs (red) and FDP homologs (purple/brown). A B 71 2.4 Discussion In this thesis, I characterized a new multicomponent and multi-subunit enzyme (BCR-FDP) from F. nucleatum that uses reducing equivalents derived from butyryl-CoA oxidation to selectively reduce O2 to H2O. Previous studies have shown that FDPs have a wide variety of electron transfer partners, spawning an extensive classification system. None of the classes previously identified show a combination of FDP and a multicomponent enzyme comprising of a BCD, ETFa, and Rd domain. This unique BCR-FDP complex is also among the first oxidative stress defense proteins characterized in F. nucleatum. F. nucleatum’s colonization of the oral microbiome makes it vulnerable to oxygen. BCR-FDP’s oxygen reductase activity could therefore be critical to its survival in its oral niche. Moreover, the bacterium’s ability to migrate through the bloodstream to infect distal sites would also be aided by BCR-FDP’s activity. In this discussion, I will detail BCR-FDP’s electron transfer pathway and predict the redox potentials of the cofactors involved using previously published work. I will discuss the importance of BCR-FDP to F. nucleatum in oxygen defense as well as how it fits into central metabolism. Using the bioinformatic work performed, I will expand on the important role BCR-FDP may perform in keeping a healthy human gut microbiome. 2.4.1 Cofactor binding sites and content of BCR-FDP Central to the function of BCR-FDP are the various cofactors (FAD, FMN, and iron centers) that perform electron transfer and catalysis. Bioinformatic and structural analysis of the binding sites reveal that each site shares structural similarity and sequence identity with respective homologs. The BCD domain is most structurally homologous to BCD from A. fermetans. Inspection of the overlayed active sites shows conservation of residues predicted to bind to the d-FAD. Crucially, the key catalytic residue for butyryl-CoA oxidation Glu363 is also conserved. Without a crystal structure of BCD, these comparisons help to predict the structure-function relationship of BCD’s residues. As the important flavin binding residues and catalytic base residue are conserved, it is reasonable to conclude that F. nucleatum’s BCD domain functions the same as BCDs from other organisms. The ETFa domain of BCR-FDP is smaller in size compared to other a subunit ETFs in that it only comprises domain II, and lacks domain I which helps oligomerize a and b subunits of 72 canonical and bifurcating ETFs108. However, given that the b subunit is not present in BCR, domain I would be considered superfluous. Domain II, on the other hand, is the binding site for a-FAD; thus ETFa domain is expected to bind an FAD and participate in electron transfer93,95. ETFa was structurally homologous to an ETF from P. denitrificans with two cationic residues being conserved. Residues His541 and Arg467 are in the FAD binding pocket of ETFa and function in stabilization of the anionic semiquinone. In particular, the Arg residue is known to be vital to anionic semiquinone formation in ETFs. Substitution of this residue for an Ala results in a severe loss of anionic semiquinone build-up over a redox titration128. Therefore, the conservation of this Arg residue is crucial for the function of ETFa as a single-electron transfer protein. AlphaFold models and sequence analysis of F. nucleatum’s FDP revealed conserved residues at both the FMN and diiron binding sites when compared to the FDP from E. coli. In particular, a Trp residue (Trp352) is conserved from E. coli. This residue, due to its coplanarity with the flavin isoalloxazine ring, is thought to participate in p stacking interactions with the FMN77. Its role in F. nucleatum is likely similar, with AlphaFold models placing it in a similar, coplanar position to the isoalloxazine ring. In F. nucleatum, the FDP binding site retains the well-conserved histidine and carboxylate ligand combination that is present in the FDP family. Upon successful recombinant expression and purification of BCD, ETFa, BCR, and BCR-FDP, the first experiments carried out examined which of the expected cofactors copurified with the proteins. BCR, along with ETFa and BCD, copurified with FAD as seen in previously characterized ETF and BCD domains in the literature92,93. BCR bound FAD at a 0.24:1 ratio when it was expressed and purified from M9 minimal media. There was little change in flavin levels when switching from LB to minimal media. However, the switch in bacterial growth conditions drastically improved iron incorporation moving from a ratio of 0.04:1 in LB to 0.91:1 in minimal media. Improved iron incorporation was due to lower concentrations of Zn in the M9-minimal media. This conclusion is based on prior studies which have shown Zn has the ability to bind to the Cys4 cofactor binding site of Rd129,130. To improve flavin incorporation into BCR, the crude extract was supplemented with 200 µM of FAD during purification. However, despite supplementation, sub-stoichiometric amounts of FAD were bound to BCR. This observation is consistent with purified ETF and BCD from other 73 organisms131,132. Sub-stoichiometric amounts of the flavins are likely attributed to their weak binding affinity to their cognate domains. A similar issue was observed with BCR-FDP. Both ratios of FAD and FMN to protein were low in M9 minimal media (0.51:1 for both). Like ETFs and BCDs, FDPs are known to be problematic with cofactor integration during recombinant expression and subsequent purification80,133. BCR-FDP also suffers from the same Zn contamination of the Rd as BCR. Additionally, Zn also binds to true b-lactamase enzymes which share high structural homology to the b-lactamase-like domain of FDPs134. This may represent a second route for possible Zn contamination in BCR-FDP. Switching from LB to M9 minimal media did improve iron levels (0.23:1 to 2.2:1), showing BCR-FDP is subjected to Zn contamination like BCR. The media switch had a negative to negligible effect on FAD and FMN incorporation. Due to weak binding of the flavins, measuring catalytic turnover based on protein concentration is challenging, as it is unclear what fraction of BCR-FDP is fully replete with cofactor. To obtain a more accurate measurement of catalytic turnover, enzyme concentration was based on the bound flavin concentration, rather than protein concentration (i.e. absorbance value at 280 nm). 2.4.2 Characterization of the electron transfer pathway of BCR-FDP To enable the reduction of O2 using electrons from butyryl-CoA, BCR-FDP must alternate between one and two-electron transfers. BCR-FDP achieves this employing flavin cofactors. The initial reduction of BCD with butyryl-CoA is a two-electron transfer with BCD’s flavin moving directly from FADox to FADhq. This was observed during the stopped-flow experiments reacting BCD and excess butyryl-CoA. A reduction at 450 nm signified that butyryl-CoA was oxidized, passing two electrons to the BCD domain. However, BCD must also participate in one-electron transfer to facilitate electron transfer to the ETFa domain. BCD was shown to form the neutral semiquinone upon reduction with dithionite with a peak forming at 567 nm. BCD can therefore participate in both one- and two-electron transfers. In contrast, ETFa only mediates single electron transfer. It accepts a single electron from BCD and passes a single electron to Rd, an obligate one-electron acceptor. Redox titrations of ETFa show a characteristic anionic semiquinone peak forming at 380 nm, which is observed in both canonical and bifurcating ETFs. The negative charge on the isoalloxazine ring of the a-FAD 74 is stabilized by nearby residues Arg447 and His541 in ETFa from F. nucleatum. The electron transfer between BCD and ETFa is similar to that observed between acyl-CoA dehydrogenases and canonical ETFs in the mitochondria, where ETF is an electrical conduit transferring electrons from fatty acid oxidation to the electron transport chain. Subsequent electron transfer from ETFa to Rd is harder to spectrally track due to the overlapping of diagnostic peaks between Rd and flavins. Only through O2-reductase activity can the presence of reduced Rd be inferred. Redox titrations of BCR-FDP showed two distinct ASQs. Both were observed in distinct phases of the titration as defined by periods of constant absorbance at 380 nm. The FMN in FDPs, like ETFa, is known to form the anionic semiquinone as opposed to the neutral semiquinone76. The diiron site has no UV-vis spectral properties and so reduction of the diiron site was not observed. 2.4.3 Chemical modification of ETFa’s FAD CDuring the analysis of the BCR domain, the presence of abnormal band I and II peaks of bound and unbound flavin was alarming. Investigation of this spectral phenomena showed a literature precedent for chemical modification of the 8’ position of the isoalloxazine ring in ETFa. ETFa was subject to chemical modification during aerobic purification both as a lone domain and when part of BCR. In BCR, this manifested as UV-vis spectral changes after overnight dialysis. These changes are diagnostic of the 8-formyl flavin in the anionic semiquinone state46. The EPR spectrum of BCR confirmed the presence of a modified radical species. The spectrum showed a peak-trough width of 13 G which is lower than expected for the anionic semiquinone (15 G)135. However, this smaller width is too small to be attributable to the 8-formyl flavin derivative which has a peak-trough width of 10 G46. 8-formyl flavin can further alter, given time, to other modified flavin species. As EPR data were collected at a later point than when UV-vis spectroscopy was performed, it is likely the EPR signal of 13 G of BCR is attributable to the presence of a further modified species. Aberrant flavin modification is known to directly affect the rate of electron transfer, lowering it by ~50%115. It is believed the modified flavin affects protein-protein interactions between the ETFa and its dehydrogenase transfer partner115. The solvent exposed nature of ETFa’s FAD means it would be able to interact with other proteins. Anaerobic purifications of BCR performed in a glove box eliminated the issue of modification. Normal 75 absorbance maxima of bands I and II were observed along with EPR peak-trough widths closer to true anionic semiquinone values (16 G). Prior studies have shown that amino, hydroxy, and formyl modifications of the flavin isoalloxazine ring are responsible for the change in UV-vis properties. These modifications arise due to the reaction of the 8’ methyl group with O2 (Figure 41). This begins with a negative charge on the N1, stabilized by the nearby Arg residue which results in delocalization of electron density from the methyl group to the N1 and O2 of the isoalloxazine ring46. The propensity for charge-delocalization allows for proton abstraction of the C8 methyl-group forming an anionic paraquinone methide intermediate which is subject to further oxidation. A basic residue is also proposed to participate in the reaction by acting as a nucleophile and extracting a proton from the methyl group46. Also crucial to chemical modification is the degree to which the FAD cofactor is exposed to the solvent. In the isolated BCR and ETFa, a-FAD may be more solvent exposed compared to BCR-FDP, which would promote reaction with O2 or another oxidizing agent. Figure 41: Mechanism of oxygen-dependent flavin modification. Electron delocalization of the methyl group favours formation of a methide group which can directly react with molecular O2. Here, 8-formyl flavin is formed as a result of the reaction. ETFa, purified as a lone domain, underwent modification much faster than BCR. Immediately after purification, a diverse range of peaks was observed with UV-vis spectroscopy with an absorbance maximum at 482 nm. This peak is diagnostic of 8-amino or 8-hydroxy modified flavin46,136. However, 8-amino modifications require an amine-based buffer. As protein 76 purification was performed in phosphate buffer, it is unlikely to be an amino modification. Furthermore, the presence of this peak has been previously attributed to the 8-hydroxy modification in ETF137. The increased rate of flavin modification in ETFa from BCR indicates that here, the FAD is also solvent exposed like previously characterized ETFs. Without the larger protein scaffold of BCR, chemical modification occurs immediately after purification, rather than after overnight dialysis. It is unlikely that these chemical modifications occur in vivo. No flavin modification was observed in preparations of BCR-FDP. Therefore, with a protein of sufficient size, the a-FAD is shielded from the solvent preventing O2 access. Moreover, F. nucleatum is an anaerobe, and O2 concentrations in the cell will be significantly less under normal atmospheric conditions. 2.4.4 Redox potentials of BCR-FDP’s cofactors The amino acid residues in the first coordination sphere surrounding a redox active cofactor directly affects its reduction potential. The electrostatic interactions between the cofactor and the protein scaffold modulate the reduction potentials of the cofactor. For example, as a flavin cofactor cycles between quinone, semiquinone, and hydroquinone states, the cofactor adopts different charges and hydrogen-bond donors/acceptors. The resulting interactions between the protein scaffold and flavin cofactor alter the Gibbs free energy for cofactor binding. This DG then impacts the reduction potential (∆𝐺 = −𝑛𝐹𝐸°). Thus, given that each flavin and iron binding site of BCR-FDP show high homology to previously characterized proteins, the reduction potentials of BCR-FDP can be predicted (Figure 42). The midpoint reduction potential of the butyryl-CoA to crotonyl-CoA couple is -10 mV and the standard reduction potential of oxygen is 1.2 V. Oxygen is an excellent terminal oxidant and ensures the overall reaction of BCR-FDP is favourable. Tracing the electron flow through each cofactor shows not every step is favourable. The midpoint reduction potential of BCD’s FAD from M. elsdenii has been calculated at -79 mV with mammalian BCD’s showing even lower midpoint potentials (-120 mV)121,138. The BCD domain in BCR-FDP is likely similar. Thus, hydride transfer from butyryl-CoA to d-FAD is not thermodynamically favourable. This is evidenced in the multiwavelength stopped-flow data following the reaction of excess butyryl-CoA and BCD where incomplete reduction of the d-FAD was observed. For comparison, full reduction was observed when BCD was titrated with dithionite. 77 The a-FAD of the ETFa domain exhibits higher reduction potentials. ETFa’s reduction potential for the FADox/sq couple range from -75 to 196 mV120,122,131. The FADsq/hq couple has been calculated at -50, 25, and 15 mV93,139,140. Therefore, electron transfer from d-FADH2 hq (obtained following hydride transfer from butyryl-CoA) to a-FAD is likely thermodynamically favourable. Indeed, pre-steady state analysis following reaction of BCR with butyryl-CoA shows rapid electron transfer to ETFa (>1000 s-1). The BCR reaction had a considerably faster rate constant when compared to isolated BCD (7.1 ±0.1and 1.1 ±0.1 s-1). The presence of ETFa with a higher reduction potential likely accelerates the rate of electron transfer to a-FAD. Rds have varied reduction potentials with most proteins ranging from -50 and 50 mV141. However, these values have been measured for isolated Rds without interactions from other proteins. In the case of rubrerthyrin, a Rd domain is fused to a larger protein: a H2O2 reductase with a diiron site. The reduction potential of this Rd domain has been measured at 280 mV142. This is due to the positioning of two basic residues (Asn and His) near the iron site of Rd which significantly raises the reduction potential due to stabilization of the reduced state143. In the case of BCR, without information concerning the tertiary structure of the protein, it is challenging to infer something similar. AlphaFold models of BCR do not provide concrete data about how each distinct domain (BCD, ETFa, and Rd) arrange in relation to each other due to low confidence in loop regions of the protein. It is still worth noting though that interaction of Rd with other protein domains can modulate and raise its reduction potential and may allow BCR to continue a favourable reduction through ETFa to Rd. Even if no interactions occur, it is conceivable that a high potential ETF could still perform electron transfer to a Rd (50 mV) especially when tied to the reaction of the FDP. The flavodoxin domain of FDP binds an FMN and has a varied reduction potential across species. Interestingly, there appears to be a correlation between higher potential and O2 activity. FDPs which display either dual or only NO activity have lower reduction potentials in the range of -40 to -117 mV FMNox/sq and -100 to -200 mV FMNsq/hq28. Those that display only O2 activity have inflated potentials of -63 mV to 25 mV FMNox/sq and -83 mV to 25 mV FMNsq/hq. Additionally, the two potentials of FMNox/sq and FMNsq/hq are quite close together in O2-specific FDPs. In Giardia intestinalis and Trichomonas vaginalis these pairs are -63/-83 and 25/25 mV respectively144,145. The similarity in reduction potentials results in a destabilization of the semiquinone state, which favours two electron reduction. The FMN and the diiron site must both 78 be fully reduced to perform the four-electron reduction of O2. Preventing semiquinone formation (and hence single electron transfer) may help ensure that O2 is not partially reduced leading to ROS formation. Given the O2-specific nature of F. nucleatum’s FDP, it can be postulated that its FMN cofactor has a high reduction potential. This may help to ensure electron transfer from BCR as well as prevent aberrant, incomplete reduction. The reduction potentials of the diiron site of FDP are generally quite high. Reduction potentials of the first and second reductions (Fe3+-Fe3+/Fe3+-Fe2+ and Fe3+-Fe2+/Fe2+-Fe2+) range from 163-190 mV and 2-132 mV in O2-specific FDPs144–146. High reduction potentials ensure thermodynamically favourable electron transfer from the reduced FMN of the flavodoxin domain whilst still being lower than O2 itself. Overall, BCR-FDP, with its variety of cofactors, has a similar variety in reduction potential. Due to the structural homology each domain shares with corresponding representative proteins, it is likely that BCR-FDP displays reduction potentials in the range of previously characterized proteins. While modulations arising from inter-domain interactions may affect the reduction potential, these are prone to improving the favourabilty of electron transfer through the domains. Future work will focus on determining reduction potentials of each of the cofactors, as these values would help elucidate the thermodynamics governing electron transfer through this novel multicomponent redox chain system. BCR-FDP does not exhibit NO or H2O2 activity. The lack of NO activity may be due to the structure of the enzyme, but it may also come down to reduction favourability. BCR-FDP uses a high-potential reductant, butyryl-CoA (-10 mV), as a source of electrons rather than NADH like other FDPs. This may limit its ability to reduce NO as NO has a much lower reduction potential than O2. (-0.68 V vs 1.2 V)147. Perhaps this difference favours O2 reduction. H2O2 does exhibit a favourable reduction potential (1.7 V) but only a single FDP has shown H2O2 activity148. Given this, nothing is known about what may select for H2O2 activity or indeed how widespread FDP-H2O2 activity is. 79 Figure 42: Reduction potential diagram of BCR-FDP. Each redox-active domain of BCR-FDP is plotted against their predicted reduction potential. Ranges are shown with grey arrows. 2.4.5 Importance of BCR-FDP to F. nucleatum BCR-FDP is the first enzyme from F. nucleatum that has been shown to reduce O2. While NOX-2 enzymes are known to exist in F. nucleatum, the in vitro activity of this enzyme has not been measured. The ability to reduce O2 to H2O can assist F. nucleatum as it travels though the various niches it can inhabit. In the oral cavity, environmental O2 is toxic for anaerobes. BCR-FDP’s O2 activity is could be vital for the bacterium’s survival. If F. nucleatum infects neighbouring tissues and enters the bloodstream, BCR-FDP could help lower O2 concentrations present in these environments. BCR-FDP may also improve F. nucleatum’s ability to colonize tumour sites, increasing tumorigenesis. The use of a common metabolite, butyryl-CoA, to power the reduction of O2 means this enzymatic system is also dual purpose. Not only could BCR-FDP function to remove harmful O2, preventing oxidative stress, but it could also increase ATP generation for F. nucleatum. 80 2.4.6 BCR-FDP in F. nucleatum’s metabolism F. nucleatum’s primary source of energy is amino acids. Three of these, lysine, glutamate, and histidine, funnel into two enzyme pathways which converge to produce crotonyl-CoA. Crotonyl-CoA is then converted to butyryl-CoA by a BCD:ETF complex (distinct from BCR). This reduction is fueled by NAD(P)H and is a bifurcating system, simultaneously reducing flavodoxin and crotonyl-CoA to butyryl-CoA. The reduced flavodoxin powers the Rnf membrane-bound complex to export Na+ and thus generate ATP synthase activity (Figure 9). This enzymatic sequence is central to F. nucleatum’s energy generation. Therefore, any way to further funnel metabolites into the BCD:ETF complex would be highly beneficial to the bacterium. Butyryl-CoA is produced at high concentrations in F. nucleatum. This is evidenced by F. nucleatum’s ability to produce butyrate, a downstream product of butyryl-CoA56,149. The CoA moiety of butyryl-CoA is transferred to acetate by butyrate-CoA transferase generating butyrate and acetyl-CoA55. Being able to recycle butyryl-CoA, before it is excreted as butyrate, would increase the efficiency of F. nucleatum’s amino acid metabolism. BCR-FDP offers this option. It converts butyryl-CoA back to crotonyl-CoA using O2 as a terminal oxidant. Crotonyl-CoA can then be used by the BCD-ETF system to generate more reduced flavodoxin for ATP production. In this biological context, BCR-FDP is in essence a self-contained, soluble respiratory pathway. O2 is acting as a terminal electron acceptor and the butyryl-CoA/crotonyl-CoA pair is performing the electron transfer role of a quinone pool. Oxidation of butyryl-CoA by O2 regenerates crotonyl-CoA for further use in ATP production. The presence of an aerobic respiratory pathway in strict anaerobes is not unheard of. There are two known examples of O2-dependent respiration in anaerobes (Figure 43). Bacteroides spp., Desulfovibrio spp., and M. thermoacetica employ membrane-bound respiratory chains that have the same basic components as aerobes150–152. A membrane-bound NADH dehydrogenase passes electrons from NADH to a quinone pool. This pool in turn feeds a terminal cytochrome bd oxidase. Cytochrome bd oxidases are distinct from their relative: heme-copper based cytochrome c oxidase. They use only heme to bind and reduce oxygen while generating a proton gradient150. These respiratory chains function at low oxygen levels which has reclassified some anaerobic species as nanoaerophiles. The second example of O2-dependent respiratory chains are FDPs. O2-specific FDPs have been proposed to function in, in addition to oxidative stress, energy generation through O2 reduction. The mechanism here is less clear than the first example. The chain consists of a NADH-rubredoxin 81 oxidoreductase coupled to a Rd terminating in the FDP (Class A FDPs). Cell extracts of Desulfovibrio spp. have been observed to produce ATP when exposed to oxygen but the cause is unknown153. It has been proposed that the consumption of O2 by FDPs limits the oxidation of key metabolic substrates allowing more substrate to enter ATP synthesis pathways4. The metabolic link in F. nucleatum is much clearer. BCR-FDP can assist in the recycling of butyryl-CoA/crotonyl-CoA and conserve energy through the Rnf. BCR-FDP is therefore not only an important enzyme for mitigating against oxidative stress, but also directly ties into central metabolism, enabling the organism to profit energetically from transient exposure to oxygen. Figure 43: Two known aerobic respiratory chains present in anaerobes. The first is a classical membrane-bound respiratory chain using NADH to feed into a quinone pool for proton gradient generation. The second is a soluble chain oxidizing NADH with a NADH rubredoxin oxidoreductase (NROR). These electrons pass through Rd ending up on an FDP for O2 reduction. 2.4.7 Importance of butyrate to the gut microbiome A brief bioinformatic probe of the prevalence of BCR was conducted using the EFI tools. This analysis revealed that BCR homologues are present in many bacterial species. Intriguingly, many of these species tended to be inhabitants of the gut microbiome (Clostridium, Brachyspira, 82 Instestimonas, Eubacteria, Anaerococcus, Peptoniphilus, and Fusobacteria)58. These species are also known butyrate producers. Butyrate production in the gut has been well-studied and is crucial to a healthy gut microbiome58. This short-chain fatty acid is an end-product of substrate fermentation in bacteria which is excreted and used in the surrounding gut tissues. It has been associated with inflammation regulation, oncogenic resistance, and oxidative stress defense. Butyrate mediates inflammation by downregulating cytokines IL-12 and IL-6 along with the antibody IgA154,155. It acts by inhibiting histone deacetylases, altering gene expression patterns, and decreasing transcription of those genes (Figure 44)154. This prevents overexpression and possible inflammatory damage of both human gut tissue and the gut microbiome. The same mechanism of histone deacetylase inhibition has also been shown to prevent cancer development by stopping several key oncogenic pathways. Butyrate inhibits Akt/ERK signaling156. These two transcription factors promote cell motility, which when overexpressed, can cause increase cell invasion and metastasis. Cancer cells also treated with butyrate show decreased motility and immunoblotting screens of treated cells show lower expression of these two transcription factors156. Butyrate also lowers glucose transport through inhibition of the GPR109A-Akt signaling cascade. This pathway culminates in expression of glucose transporters, which can turn oncogenic if overexpressed157. Again, through histone deacetylase inhibition, butyrate prevents expression of the transcription factors, stopping uncontrolled glucose uptake. Butyrate also functions to maintain anaerobicity in the gut microbiome. This not only helps mediate oxidative stress but also prevent the growth of facultative anaerobes. 83 Figure 44: Butyrate inhibits histone deacetylases to influence gene expression. Butyrate inhibits histone deacetylase (HDAC) leading to a change in gene expression pattern. This change arrests transcription of oncogenic genes. 2.4.8 Anaerobicity of the gut microbiome The human gut can be thought of as an oxygen gradient. The duodenum contains low levels of oxygen (32 mm Hg) while the large intestine contains virtually none (<1 mm Hg)158. This gradient defines the bacterial microbiome within the gut with more aerotolerant organisms closer to the stomach and obligate anaerobes in the colon. The gut microbiome is incredibly important for human health, as alterations in its composition (dysbiosis) are associated with many diseases including cancer, obesity, inflammatory bowel disease (IBD), and neurological conditions159–161. Maintaining anaerobicity in the gut is vital to a healthy microcosm and prevention of these adverse outcomes. The normal bacterial constituents of the gut, which lead to a healthy microbiome, are predominantly obligate anaerobes of the Firmicutes (class Clostridria) and Bacteroidetes (Bacteroidia) phlya162. The metabolism of Clostridia is key for maintaining gut health as it works synergistically with the host to maintain anaerobicity162,163. This example illustrates the role of butyrate in maintaining an anaerobic gut environment. Clostridia derive energy through the anaerobic 84 fermentation of carbohydrates and amino acids, producing butyrate as an end-product. Butyrate is taken up by colonocytes and stimulates b-oxidation and oxidative phosphorylation164,58. Stimulation of ATP synthase reduces oxygen, removing it from the gut. Very active oxidative phosphorylation leads to a depletion of O2 in the colonic lumen, generating a state of anaerobiosis, favouring growth of obligate anaerobes165. The mutualistic relationship between Clostridia and the human host is essential for good gut health. Gut dysbiosis erupts once anaerobicity is compromised leading to greater colonization from facultative anaerobes (primarily Enterobacteriaceae) which can use oxygen as a terminal electron acceptor163. The high redox potential associated with the O2/H2O couple results in more ATP production per mole of glucose, resulting in overgrowth of the facultative anaerobes. These include the pathogens Salmonellae, Streptococcus pneumoniae, Enterococcus faecium, Pseudomonoas aeruginosa, and E. coli166. The discovery of BCR in many gut anaerobes directly links to the importance of anaerobicity. By performing O2 reduction, the activity of BCR-FDP would contribute to the maintenance of an anaerobic gut environment, preventing pathogenic colonization. 85 Chapter 3: Conclusion The discovery and subsequent characterization of BCR-FDP is an important step towards understanding the oxidative stress response in F. nucleatum. It is one of the first enzymes involved in oxidative stress defense from F. nucleatum to be recombinantly expressed and have activity demonstrated in vitro. The O2 reductase activity observed in vitro likely translates to an in vivo mechanism for F. nucleatum to protect itself from O2, as well as downstream reactive molecules like O2- and H2O2. The activity of BCR-FDP is also directly connected to F. nucleatum’s metabolism through its use of butyryl-CoA as a reductant. The large availability of butyryl-CoA due to F. nucleatum’s amino acid-centered metabolism lends a second function to BCR-FDP, allowing it to recycle this common metabolite and improve energy efficiency. The prevalence of BCR-FDP in gut-based bacteria uncovered by bioinformatic analysis suggests that this enzyme system may play a role in maintaining anaerobiosis in the gut, a vital condition for a healthy colon. Future work with this unique dicistron could explore several different avenues. Several questions remain unanswered about F. nucleatum’s BCR-FDP. These include the molecular structure of the enzyme as well as the kinetic and thermodynamic properties associated with electron transfer and enzyme turnover. Future studies could examine BCR-FDP gene regulation in F. nucleatum and other anaerobic gut bacteria that harbor the gene. Understanding the biological role of the enzyme in terms of mitigating oxidative stress or adaptation to difference ecological niches could also be addressed through gene knockout studies in F. nucleatum or anaerobic gut bacteria. The latter approach could further our understanding of the role of BCR-FDP in maintenance a healthy gut microbiome, a topic of increasing scientific relevance and interest. 3.1 Discovery and importance of BCR-FDP in F. nucleatum BCR-FDP joins a handful of established enzymes which function in oxidative stress defense in F. nucleatum. These defense mechanisms are likely vital to F. nucleatum’s survival in the oral cavity during environmental O2 exposure. BCR-FDP could also be a part of F. nucleatum’s opportunistic behaviour, helping shield the bacterium from oxidative harm during its invasion of neighbouring cells and dissemination through blood vessels. A robust O2 defense mechanism could also help F. nucleatum colonize distal sites such as the amniotic fluid and tumors. While performing this protective role, BCR-FDP also contributes to metabolic efficiency through the 86 recycling of butyryl-CoA. Reduction of O2 by butyryl-CoA produces crotonyl-CoA. Crotonyl-CoA is then used in electron bifurcation to produce reduced flavodoxin. Reduced flavodoxin is then oxidized at the Rnf complex to power the generation of Na+ gradient for ATP production. The reduction of reduction of crotonyl-CoA regenerates butyryl-CoA, establishing a cycle. Thus, BCR-FPD (a simple, soluble respiratory chain) gives F. nucleatum the ability to profit energetically from exposure to O2. Moreover, BCR-FDP can couple oxidative stress defense to metabolic efficiency, giving F. nucleatum a powerful tool to withstand the presence of O2. 3.2 Proposed role of BCR-FDP in the gut microbiome F. nucleatum, upon entering the colon and colonizing tumor microenvironments, is an opportunistic pathogen. Its use of BCR-FDP may serve to help it survive these environments, perhaps acting as a virulence factor. However, BCR-FDP may function differently in other organisms. BCR-FDP homologues, through bioinformatic analysis, were discovered in other anaerobic gut bacteria. These species were members of the Firmicutes and Fusobacteriota phyla. Additionally, these species primarily are known butyrate producers which is important to maintaining anaerobiosis and a healthy gut. Allowing O2 to build-up in the gut enables pathogenic, facultative anaerobes to outcompete obligate anaerobes as they are able to use O2-based respiratory chains to produce ATP. BCR-FDP could be another mechanism for these healthy butyrate producers to remove O2 from the environment to prevent harmful infection. 3.3 Future directions Questions remain about the role of BCR-FDP in F. nucleatum and its role in the wider gut microbiome. The activity of BCR-FDP is reliant on electron transfer through its various iron and flavin cofactors. Given that BCR-FDP exhibits O2 reductase activity with the substrate butyryl-CoA, electron transfer through the cofactors occurs. Future work to determine the exact reduction potentials of each cofactor would establish the thermodynamics of the electron transfer path as well as allow for comparisons to existing published reduction potentials of BCD, ETF, Rd, and the FDP. By coupling redox titrations to potential voltammetry, these reduction potentials could be extracted. Future work should also examine the kinetics of O2 reduction. Previous work with FDPs have found kinetic parameters (KM, kcat, Vmax) for the reduction of both O2 and NO28. This kinetic 87 work relies on the absorbance of NADH at 340 nm. Given the lack of NADH used by BCR-FDP, the Clark-type electrodes used to detect O2 and NO activity could be used to measure the rate of substrate consumption to plot kinetic data. Obtaining kinetic parameters would give an idea of BCR-FDP’s role in F. nucleatum, particularly the catalytic efficiency (kcat/KM). The catalytic efficiency could be used to compare to other oxidative stress defense enzymes to gain a greater understanding of how efficiently BCR-FDP reduces O2. Also of interest is the structure of BCR-FDP. Solving the crystal structure of BCR-FDP would increase our understanding of the mechanism of catalysis and mode of electron transfer between redox cofactors. The discovery of BCR-FDP homologues in butyrate producers uncovers many potential research questions. This includes whether BCR-FDP exhibits any NO or H2O2 activity in these other anaerobic organisms. It would also be of interest to evaluate the role of BCR-FDP on bacterial survival through gene knock-out studies and what physiological conditions, if any, lead an upregulation of BCR-FDP gene expression in F. nucleatum or other gut microbes. Gene knockout has been performed in F. nucleatum with a plasmid-based system which allows for non-polar gene deletion167. DNA regions approximately 1 kb upstream and downstream of the target are amplified with PCR and cloned into a suicide plasmid. This is the transformed into F. nucleatum where it undergoes crossover events to excise the target gene. 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Thesis/Dissertation
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Biochemistry and Molecular Biology
Vancouver : University of British Columbia Library
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Wolthers, Kirsten
Coupling butyryl-coenzyme A oxidation to oxygen reduction in Fusobacterium nucleatum
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