Science, Irving K. Barber Faculty of (Okanagan)
DSpace
UBCO
Mothersole, Robert Geoffrey
Doctor of Philosophy - PhD
University of British Columbia
Fusobacterium nucleatum, a Gram-negative oral bacterium, is linked to periodontal disease, colorectal cancer and adverse pregnancy outcomes. The research herein uncovers the kinetic, structural and mechanistic properties of three F. nucleatum proteins that are potential therapeutic targets. Chapter 2 focuses on the characterisation of flavodoxin (FnFld), an FMN-containing electron transfer protein. FnFld was shown to have the highest recorded reduction potentials amongst characterised long-chain flavodoxins. Fluorescence binding studies revealed that the elevated potentials are due to preferred binding of the FMN hydroquinone anion over the oxidised FMN quinone. The structural rationale for the electropositive reduction potentials was provided through the x-ray crystal structure of FnFld. Chapter 3 investigates the thermodynamic and biophysical effects of four naturally occurring substitutions present in FnFld. A glycine to lysine substitution in a highly conserved phosphate-binding loop does not elicit a dramatic effect on FMN binding or the redox properties of the flavin cofactor, but it does contribute to a positively charged binding cleft that may serve as a redox partner recognition site. Three other residues, Y56, N89 and F93 were shown to modulate the electropositive reduction potentials. Chapter 4 investigates lanthionine synthase (LS), which belongs to the fold II family of PLP-dependent enzymes. Although LS is catalytically promiscuous like other family members, it shows catalytic preference toward the condensation of two molecules of L-cysteine to produce lanthionine and HâS. Inspection of the LS crystal structure revealed a novel loop region containing S234 that forms electrostatic interactions with lanthionine. The catalytic importance of this residue was confirmed through mutagenesis. Chapter 5 focuses on the characterisation of a bifunctional ornithine decarboxylase/arginase (ODA). Sequence alignment revealed that ODA contains two naturally occurring amino acid substitutions in the conserved arginase binuclear Mn²+ binding site. ICP-MS analysis showed that wild type ODA is devoid of Mn²+ . Restoration to the conserved Mn²+ binding site improved Mn²+ binding, but not catalytic activity. We confirmed that H119, D199 and K230 are vital residues in the ornithine decarboxylase active site. Finally, known inhibitors of ornithine decarboxylase and arginase had no effect on ODA activity, suggesting that the enzyme contains distinct active sites.
Characterisation of Three Cofactor-containing Proteins from the Oral Bacterium Fusobacterium nucleatum by Robert Geoffrey Mothersole M.Chem., The University of York, 2016 A DISSERTATION SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY In THE COLLEGE OF GRADUATE STUDIES (Biochemistry and Molecular Biology) THE UNIVERSITY OF BRITISH COLUMBIA (Okanagan) July 2020 © Robert Geoffrey Mothersole ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Characterisation of Three Cofactor-containing Proteins from the Oral Bacterium Fusobacterium nucleatum submitted by Robert Geoffrey Mothersole in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Biochemistry and Molecular Biology Examining Committee: Dr. Kirsten Wolthers (Irving K. Barber Faculty of Science) Supervisor Dr. Kevin Smith (Irving K. Barber Faculty of Science) Supervisory Committee Member Dr. Isaac Li (Irving K. Barber Faculty of Science) Supervisory Committee Member Dr. Filip Van Petegem (Faculty of Medicine) University Examiner Dr. Todd Holyoak (University of Waterloo) External Examiner iii Abstract Fusobacterium nucleatum, a Gram-negative oral bacterium, is linked to periodontal disease, colorectal cancer and adverse pregnancy outcomes. The research herein uncovers the kinetic, structural and mechanistic properties of three F. nucleatum proteins that are potential therapeutic targets. Chapter 2 focuses on the characterisation of flavodoxin (FnFld), an FMN-containing electron transfer protein. FnFld was shown to have the highest recorded reduction potentials amongst characterised long-chain flavodoxins. Fluorescence binding studies revealed that the elevated potentials are due to preferred binding of the FMN hydroquinone anion over the oxidised FMN quinone. The structural rationale for the electropositive reduction potentials was provided through the x-ray crystal structure of FnFld. Chapter 3 investigates the thermodynamic and biophysical effects of four naturally occurring substitutions present in FnFld. A glycine to lysine substitution in a highly conserved phosphate-binding loop does not elicit a dramatic effect on FMN binding or the redox properties of the flavin cofactor, but it does contribute to a positively charged binding cleft that may serve as a redox partner recognition site. Three other residues, Y56, N89 and F93 were shown to modulate the electropositive reduction potentials. Chapter 4 investigates lanthionine synthase (LS), which belongs to the fold II family of PLP-dependent enzymes. Although LS is catalytically promiscuous like other family members, it shows catalytic preference toward the condensation of two molecules of L-cysteine to produce lanthionine and H2S. Inspection of the LS crystal structure revealed a novel loop region containing S234 that forms electrostatic interactions with lanthionine. The catalytic importance of this residue was confirmed through mutagenesis. iv Chapter 5 focuses on the characterisation of a bifunctional ornithine decarboxylase/arginase (ODA). Sequence alignment revealed that ODA contains two naturally occurring amino acid substitutions in the conserved arginase binuclear Mn2+ binding site. ICP-MS analysis showed that wild type ODA is devoid of Mn2+. Restoration to the conserved Mn2+ binding site improved Mn2+ binding, but not catalytic activity. We confirmed that H119, D199 and K230 are vital residues in the ornithine decarboxylase active site. Finally, known inhibitors of ornithine decarboxylase and arginase had no effect on ODA activity, suggesting that the enzyme contains distinct active sites. v Lay Summary Fusobacterium nucleatum is a bacterium that is ubiquitous to human oral cavity. It contributes to the development of periodontal diseases, and an emerging body of research links the bacterium to colorectal cancer and preterm births. Therefore, finding a method to slow or stop the growth of the bacteria will help improve the health outcomes for people with F. nucleatum infections. This can be done by selectively targeting and disabling proteins involved in vital processes for the survival of the organism. We have identified three unique proteins that may act as potential drug targets. In this thesis, we have investigated these proteins to determine their viability to be used as therapeutic targets. A number of techniques have been used to ascertain the structure and mechanism of these proteins. We were able to conclude that all three proteins showed unique properties that may enable them to serve as drug targets. vi Preface Some of the results presented in Chapters 2 and 3 have been published and some figures reproduced with permission: Mothersole, R. G.; Macdonald, M.; Kolesnikov, M.; Murphy, M. E. P.; Wolthers, K. R., Structural insight into the high reduction potentials observed for Fusobacterium nucleatum flavodoxin. Protein Science 2019, 28 (8), 1460-1472. The crystal structures were determined in collaboration with Maxim Kolesnikov and Dr. Michael Murphy at the University of British Columbia Vancouver. Marta Macdonald, an undergraduate researcher in the Wolthers group conducted some of the fluorescence stopped flow experiments for wild type Fld and the K13G variant presented in section 2.2.5. Some of the results presented in Chapter 4 have been published and some figures reproduced with permission: Mothersole, R. G.; Wolthers, K. R., Structural and Kinetic Insight into the Biosynthesis of H2S and L-Lanthionine from L-Cysteine by a Pyridoxal L-Phosphate-Dependent Enzyme from Fusobacterium nucleatum. Biochemistry 2019, 58 (34), 3592-3603. Brody Geoghegan, an undergraduate researcher in the Wolthers group, produced the K230M, H119Q and D199A variants via site directed mutagenesis outlined in section 5.2.4. ICP-MS experiments, outlined in section 5.3.10, were performed by Dr. Mark Button at the FiLTER Laboratory at the University of British Columbia Okanagan. vii Table of Contents Abstract ........................................................................................................................................ iii Lay Summary ................................................................................................................................ v Preface ........................................................................................................................................... vi Table of Contents ........................................................................................................................ vii List of Tables .............................................................................................................................. xiii List of Figures .............................................................................................................................. xv List of Abbreviations ............................................................................................................... xxiii Acknowledgements ................................................................................................................... xxv 1. Introduction ....................................................................................................................... 1 1.1 Fusobacterium nucleatum: an emerging pathogen .................................................. 1 1.1.1 F. nucleatum is associated with periodontal diseases ................................. 1 1.1.2 A link between F. nucleatum and abnormal pregnancy outcomes ............. 2 1.1.3 Recent links to colorectal cancer ................................................................ 3 1.1.4 Three proteins from F. nucleatum have been studied ................................. 4 1.2 Flavodoxin ................................................................................................................ 5 1.2.1 Role of flavodoxins in anaerobic microorganisms ..................................... 6 1.2.2 General structure of flavodoxins ................................................................. 8 1.2.3 FMN enables electron shuttling .................................................................. 9 1.2.4 Comparing flavodoxins from multiple species ......................................... 10 1.2.5 Preliminary success with flavodoxins as a drug target ............................. 12 viii 1.3 Lanthionine synthase .............................................................................................. 13 1.3.1 Meso-lanthionine is a component of the F. nucleatum peptidoglycan layer ................................................................................................................... 14 1.3.2 Importance of H2S in bacteria ................................................................... 14 1.3.3 PLP-dependent enzymes ........................................................................... 15 1.3.4 Sequence similarity with O-acetyl-L-serine sulfhydrylase and cystathionine β-synthase ................................................................................................. 17 1.3.5 Structural insight gained from crystal structures of lanthionine synthase 22 1.4 Ornithine decarboxylase/ arginase ......................................................................... 24 1.4.1 The ornithine decarboxylase component .................................................. 25 1.4.2 Catalytic mechanism of ornithine decarboxylase ..................................... 26 1.4.3 Treatments targeting ornithine decarboxylase .......................................... 27 1.4.4 Sequence alignment of the ODC domain .................................................. 29 1.4.5 The arginase component ........................................................................... 30 1.4.6 Catalytic mechanism of arginase .............................................................. 31 1.4.7 Inhibition of arginase ................................................................................ 32 1.4.8 Sequence alignment of the arginase domain ............................................. 33 1.5 Research Objectives ............................................................................................... 34 2. Flavodoxin, Part 1 ................................................................................................................. 36 2.1 Chapter Summary ................................................................................................... 36 2.2 Materials and Methods ........................................................................................... 37 2.2.1 Materials ................................................................................................... 37 2.2.2 Cloning petFld into pET41a plasmid ........................................................ 37 ix 2.2.3 Purification and expression of wild type FnFld ........................................ 38 2.2.4 Fluorescence titrations .............................................................................. 40 2.2.5 Stopped Flow Spectrophotometry ............................................................. 40 2.2.6 Redox Potentiometry ................................................................................ 41 2.2.7 FnFld crystallisation and structure determination ..................................... 43 2.3 Results .................................................................................................................... 44 2.3.1 Properties of purified FnFld ...................................................................... 44 2.3.2 FMN binding to apo-flavodoxin ............................................................... 46 2.3.3 Redox potentials ........................................................................................ 50 2.3.4 Three-dimensional structure ..................................................................... 52 2.4 Discussion .............................................................................................................. 59 2.4.1 Analysis of the FnFld crystal structure ..................................................... 59 2.4.2 FnFld elicits relatively weak FMN binding .............................................. 59 2.4.3 Increasing ionic strength results in decreased binding affinity towards FMN ................................................................................................................... 62 2.4.4 Perturbation of the redox potentials .......................................................... 64 2.5 Concluding remarks ............................................................................................... 67 3. Flavodoxin, Part 2 ................................................................................................................. 69 3.1 Chapter Summary ................................................................................................... 69 3.2 Materials and Methods ........................................................................................... 70 3.2.1 Materials ................................................................................................... 70 3.2.2 Generation of the single site substitutions ................................................ 70 3.2.3 Determination of the full crystal structure of K13G ................................. 71 x 3.2.4 Binding and redox analysis with flavodoxin variants ............................... 72 3.3 Results .................................................................................................................... 72 3.3.1 Three-dimensional structure ..................................................................... 72 3.3.2 FMN binding to apo-flavodoxin variants.................................................. 75 3.3.3 Redox potentials of flavodoxins variants .................................................. 77 3.4 Discussion .............................................................................................................. 80 3.4.1 K13 accounts for slight decrease in FMN binding affinity ....................... 80 3.4.2 Y56 increases FMN binding affinity ........................................................ 81 3.4.3 N89 results in a more electropositive Esq/hq ............................................. 83 3.4.4 Role of F93 ............................................................................................... 84 3.5 Concluding remarks ............................................................................................... 85 4. Lanthionine Synthase ........................................................................................................... 87 4.1 Chapter Summary ................................................................................................... 86 4.2 Material and Methods............................................................................................. 88 4.2.1 Materials ................................................................................................... 88 4.2.2 Cloning petFld into pET15b plasmid ........................................................ 88 4.2.3 Purification and expression of wild type lanthionine synthase ................. 89 4.2.4 Variants produced by site-directed mutagenesis ....................................... 91 4.2.5 Steady-state kinetic assays measuring H2S production ............................ 91 4.2.6 Steady-state kinetic assays measuring L-cysteine production .................. 92 4.2.7 High-performance liquid chromatography (HPLC) .................................. 92 4.2.8 Stopped-flow spectroscopy ....................................................................... 93 4.2.9 Fluorescence titrations .............................................................................. 94 xi 4.3 Results .................................................................................................................... 95 4.3.1 Properties of purified lanthionine synthase ............................................... 95 4.3.2 Steady-state kinetics analysis of H2S production ...................................... 98 4.3.3 Steady-state kinetic analysis of L-cysteine production ........................... 100 4.3.4 HPLC analysis of products ..................................................................... 101 4.3.5 Pre-steady-state kinetic analysis with L-cysteine ................................... 106 4.3.6 Pre-steady-state kinetic analysis with D,L-lanthionine........................... 113 4.3.7 Pre-steady-state kinetic analysis with O-acetyl-L-serine ........................ 119 4.3.8 LS binding to products and product analogues ....................................... 124 4.4 Discussion ............................................................................................................ 127 4.4.1 Preferred reactions of wild type LS ........................................................ 127 4.4.2 Binding of potential inhibitors ................................................................ 132 4.4.3 Structural insight into the role of S234 ................................................... 132 4.5 Concluding remarks ............................................................................................. 134 5. Ornithine Decarboxylase/Arginase ................................................................................... 136 5.1 Chapter Summary ................................................................................................. 136 5.2 Materials and Methods ......................................................................................... 137 5.2.1 Materials ................................................................................................. 137 5.2.2 Cloning gene for ODA into pET15b plasmid ......................................... 138 5.2.3 Purification and expression of wild-type ODA....................................... 138 5.2.4 Variants produced by site-directed mutagenesis ..................................... 140 5.2.5 Sub-cloning ODC and arginase component parts ................................... 141 5.2.6 Measuring the rate of CO2 formation with L-arginine and L-ornithine .. 142 xii 5.2.7 Rate of urea formation by ODA and the isolated arginase component .. 144 5.2.8 Determining quaternary structure via gel filtration chromatography ..... 144 5.2.9 Divalent metal content of ODA by ICP-MS ........................................... 145 5.3 Results .................................................................................................................. 146 5.3.1 Properties of purified ODA and the isolated ODC and arginase components ................................................................................................................. 146 5.3.2 Steady state kinetic analysis of wild type ODA and the isolated ODC component with L-ornithine as the substrate .......................................... 151 5.3.3 Kinetic analysis of wild type ODA and the isolated arginase component with L-arginine as the substrate .............................................................. 152 5.3.4 No inhibition observed of wild type ODA .............................................. 155 5.3.5 Metal Analysis of ODA and the S609D/N713D variant ........................ 155 5.4 Discussion ............................................................................................................ 155 5.4.1 Advantages of bifunctional enzyme to F. nucleatum ............................. 155 5.4.2 Comparison of kinetic activity ................................................................ 157 5.4.3 Comparison of quaternary structures ...................................................... 157 5.4.4 Ornithine decarboxylase active site residues successfully identified ..... 159 5.4.5 ODA may function via a unique arginase mechanism ............................ 159 5.4.6 Substrate channelling from the arginase domain to the ODC domain .... 160 5.5 Concluding remarks ............................................................................................. 160 6. Conclusion ........................................................................................................................... 162 References .................................................................................................................................. 168 xiii List of Tables Table 1.1 Distribution of acidic and basic residues in flavodoxins from various organisms ...... 12 Table 2.1 Binding constants for FMN binding to apo-FnFld ....................................................... 48 Table 2.2 Data collection and refinement statistics for wild type FnFld ..................................... 53 Table 2.3 Binding constants for FMN binding to apo-Fld from various organisms .................... 60 Table 2.4 Comparison of the redox potentials and free energies associated with FMN binding to apo-Fld from various organisms ................................................................................................... 65 Table 3.1 List of primers used in site directed mutagenesis to produce the FnFld variants ........ 70 Table 3.2 Data collection and refinement statistics for K13G ..................................................... 74 Table 3.3 Binding constants for FMN binding to variants of apo-FnFld ..................................... 76 Table 3.4 Comparison of the redox potentials and free energies associated with FMN binding to the different apo-Fld variants ........................................................................................................ 79 Table 4.1 Steady-state kinetic parameters for formation of H2S by LS ....................................... 99 Table 4.2 Steady-state kinetic parameters for formation of L-cysteine by LS .......................... 100 Table 4.3 Pre-steady state kinetic parameters for LS and S234A with L-cysteine .................... 109 Table 4.4 Pre-steady state kinetic parameters for LS and S234A with D,L-lanthionine ........... 116 Table 4.5 Pre-steady state kinetic parameters for LS and S234A with O-acetyl-L-serine ........ 119 Table 4.6 The equilibrium dissociation constants for the ten ligands with wild type LS .......... 126 xiv Table 4.7 Comparison of steady-state kinetic parameters for formation of H2S by LS in the presence of L-cysteine with CBS from other organisms ............................................................ 128 Table 4.8 Comparison of steady-state kinetic parameters for formation of L-cysteine in the presence of O-acetyl-L-serine by LS with OASS from other organisms ................................... 131 Table 5.1 Sequence of oligonucleotides used to produce the ODA variants ............................. 141 Table 5.2 Number of subunits formed by ODA and the isolated arginase and ornithine decarboxylase components determined by size-exclusion chromatography .............................. 148 Table 5.3 Kinetic parameters for the ODA and ODC/ODA in the presence of L-ornithine ...... 152 Table 5.4: Steady-state kinetic parameters for the ODA variants in the presence of L-arginine..................................................................................................................................................... 154 xv List of Figures Figure 1.1 Structure of the FMN cofactor...................................................................................... 5 Figure 1.2 Reversible process of electron shuttle from PFOR to FqrB and NADP+ via Fld ......... 7 Figure 1.3 Cartoon of the crystal structure of Fld from Anabaena ................................................ 8 Figure 1.4 The three oxidation state of the FMN cofactor ............................................................. 9 Figure 1.5 Sequence alignment a selection of flavodoxins from different species ...................... 11 Figure 1.6 A selection of compounds that have inhibitory effects on H. pylori flavodoxin........ 13 Figure 1.7 Production of lanthionine from two molecules of L-cysteine catalysed by LS .......... 14 Figure 1.8 Mechanism of the formation of L-cysteine O-acetyl-L-serine via an α-aminoacrylate intermediate in M. tuberculosis OASS ......................................................................................... 18 Figure 1.9 Mechanism of the formation of L-lanthionine from two molecules of L-cysteine via an α-aminoacrylate intermediate in S. typhimurium OASS ............................................................... 19 Figure 1.10 Mechanism of the formation of L-cystathionine from L-cysteine and L-homocysteine via an α-aminoacrylate intermediate in human CBS .................................................................... 20 Figure 1.11 Four reactions catalysed by human CBS .................................................................. 21 Figure 1.12 Cartoon depiction of the homodimer of LS .............................................................. 23 Figure 1.13 Crystal structure of the internal aldimine of LS ....................................................... 23 Figure 1.14 Crystal structure showing the 69TSGNT73 loop in LS .............................................. 24 xvi Figure 1.15 Reactions catalysed by ODA, the conversion of L-arginine to L-ornithine and urea followed by conversion of L-ornithine to putrescine and CO2 ..................................................... 25 Figure 1.16 ODC catalytic mechanism for the production of putrescine and CO2 from L-ornithine via a quinonoid intermediate mediated by the ODC component of ODA .................................... 27 Figure 1.17 Mechanism of irreversible inhibition of mouse ODC by DFMO ............................. 28 Figure 1.18 Sequence alignment of the ODC component of F. nucleatum ODA with ornithine decarboxylases and arginine decarboxylases ................................................................................ 29 Figure 1.19 Homology model of the predicted active site of the ODC component of ODA ...... 30 Figure 1.20 Catalytic mechanism for the hydrolysis of L-arginine to urea and L-ornithine by arginase ......................................................................................................................................... 31 Figure 1.21 Structures of the arginase inhibitors ABH and BEC ................................................ 33 Figure 1.22 Sequence alignment of the arginase component of F. nucleatum ODA with other arginases ........................................................................................................................................ 33 Figure 1.23 Crystal structure of the active site of human arginase I overlaid with a homology model of the arginase component of ODA ................................................................................... 34 Figure 2.1 A thermodynamic loop that connects the reduction potentials of FMN in solution with the reduction potentials and the free energies associated with FMN binding .............................. 43 Figure 2.2 SDS 14% polyacrylamide gel showing both apo and holo-flavodoxin purified by ammonium sulfate precipitation and anion exchange chromatography ....................................... 45 Figure 2.3 UV/visible absorbance spectrum of purified holo-FnFld ........................................... 45 xvii Figure 2.4 Change in fluorescence emission of FMN following the addition of apo-Fld ........... 46 Figure 2.5 General decrease in Kd for the FnFld-FMNox complex .............................................. 47 Figure 2.6 A) Fluorescent traces following the binding of an excess of 50 µM FMN to 1 µM apo-Fld. B) Linear regression of recorded kobs plotted against FMN concentration ............................ 49 Figure 2.7 Plot of the association constants for FMN binding to apo-Fld against ionic strength 50 Figure 2.8 Overlaid absorption spectra of holo-Fld acquired during chemical reduction of the flavin ............................................................................................................................................. 51 Figure 2.9 Plot of the absorbance at 580 nm of holo-Fld against the reduction potential ........... 52 Figure 2.10 A cartoon of the crystal structure of wild type FnFld............................................... 54 Figure 2.11 A electron density map of the FMN cofactor bound to wild type Fld with a loop comprised of residues Q57 to G59................................................................................................ 54 Figure 2.12 Close up showing the residues that make up the P-loop and residues G59, N89, S96 and C98 surrounding the FMN cofactor ....................................................................................... 55 Figure 2.13 A close-up of Y56, F93 and E145 residues surrounded the FMN cofactor ............. 56 Figure 2.14 Close-up showing the distance between the FMN N5 and the closest residue on the amino acid backbone (V58) .......................................................................................................... 57 Figure 2.15 Charge distribution map of the crystal structure of FnFld........................................ 58 Figure 2.16 Overlaid crystal structures of holo-FnFld and apo-FnFld ........................................ 58 Figure 2.17 Comparison of the interactions between NH of residue 59 (58 in case of FnFld) and the FMN N5 in four different long chain flavodoxins .................................................................. 61 xviii Figure 2.18 Conformation change undergone by residues W57 and Y94 following binding of FMN in Fld from Anabaena ......................................................................................................... 63 Figure 2.19 Conformation change undergone by residues Y56 and F93 following binding of FMN in FnFld ......................................................................................................................................... 63 Figure 2.20 Peptide flip of N58 and V59 observed in Fld from S. elongatus.............................. 65 Figure 2.21 Structures of various flavodoxins with known reduction potentials coloured by electrostatic potential .................................................................................................................... 66 Figure 3.1 A electron density map of the FMN cofactor bound to K13G with a loop comprised of residues Q57 to G59...................................................................................................................... 73 Figure 3.2 Overlay of the crystal structures of wild-type FnFld and K13G comparing the phosphate binding loop ................................................................................................................. 73 Figure 3.3 A) Change in fluorescence emission of FMN following the addition of apo-forms of Fld B) Linear regression of recorded kobs plotted against FMN concentration for apo-forms of Fld....................................................................................................................................................... 76 Figure 3.4 Overlaid absorption spectra of A) K13G, C) Y56W, E) F93Y and G) N89D acquired during chemical reduction of the flavin cofactor. Plot of the absorbance at 580 nm of B) K13G, D) Y56W, F) F93Y and H) N89D against the reduction potential normalised to the standard hydrogen electrode ........................................................................................................................................ 79 Figure 3.5 Overlay of the crystal structures of wild-type FnFld and K13G comparing the position of the FMN cofactor relative to residues Y56 and F93 ................................................................ 81 Figure 3.6 Overlay of the crystal structures of wild-type FnFld and Anabaena Fld ................... 82 xix Figure 3.7 A) Position of D90 relative to FMN N1 in Fld from S. elongates B) Position of D95 relative to FMN N1 in Fld from D. vulgaris C) Position of N89 relative to FMN N1 in FnFld .. 83 Figure 4.1 SDS 10% polyacrylamide gel showing the purification of wild type LS ................... 96 Figure 4.2 Absorption spectra of the PLP cofactor from wild type LS ....................................... 97 Figure 4.3 Absorption spectra of the PLP cofactor from S234A ................................................. 98 Figure 4.4 Michaelis-Menten plot of the production of H2S in the reaction of L-cysteine catalysed by A) wild type LS and B) S234A ................................................................................................ 99 Figure 4.5 Michaelis-Menten plots of the production of L-cysteine in the reaction of O-acetyl-L-serine and Na2S catalysed by A) wild type LS and B) S234A ................................................... 101 Figure 4.6 Four reactions catalysed by human CBS .................................................................. 102 Figure 4.7 Analysis of dansylated products using reverse phase HPLC ................................... 103 Figure 4.8 Analysis of dansylated products using reverse phase HPLC ................................... 104 Figure 4.9 Analysis of dansylated products using reverse phase HPLC ................................... 105 Figure 4.10 Time resolved spectra following rapid mixing of L-cysteine and wild type LS .... 107 Figure 4.11 The conversion of the PLP cofactor from the internal aldimine in LS to the α-aminoacrylate intermediate and then to the external aldimine with lanthionine ........................ 108 Figure 4.12 Single wavelength absorbance trances acquired at A) 320 nm and B) 406 nm following rapid mixing of wild type LS with varying concentration of L-cysteine. C) Linear dependence of kobs1 with wild type LS on L-cysteine concentration. D) Hyperbolic dependence of kobs2 on L-cysteine concentration ................................................................................................................. 110 xx Figure 4.13 Averaged single wavelength absorbance trace at 428 nm obtained after mixing wild type LS with L-cysteine .............................................................................................................. 111 Figure 4.14 Time resolved spectra following rapid mixing of L-cysteine and S234A .............. 112 Figure 4.15 A) Single wavelength absorbance trances acquired at 406 nm following rapid mixing of S234A with varying concentration of L-cysteine B) Linear dependence on L-cysteine concentration of kobs1 with S234A. C) Hyperbolic dependence on L-cysteine concentration of kobs2 with S234A ................................................................................................................................. 113 Figure 4.16 Time-resolved UV-vis absorption spectra following rapid mixing of wild type LS with D,L-lanthionine ................................................................................................................... 114 Figure 4.17 Formation of the gem-diamine intermediate and external aldimine following addition of lanthionine to LS .................................................................................................................... 115 Figure 4.18 Single wavelength stopped-flow absorbance changes at A) 320 nm, C) 406 nm and E) 428 nm. The resulting observed rate constants were fitted to either linear or hyperbolic equations for B) 320 nm, D) 406 nm and F) 428 nm ................................................................. 116 Figure 4.19 Time resolved spectra following rapid mixing of D,L-lanthionine and S234A ..... 118 Figure 4.20 A) Single wavelength absorbance trances acquired at 406 nm following rapid mixing of S234A with varying concentration of D,L-lanthionine. B) Linear dependence on D,L-lanthionine concentration of kobs with S234A ............................................................................. 118 Figure 4.21 A subset of the time resolved spectra following rapid mixing of O-acetyl-L-serine with wild type LS ........................................................................................................................ 120 xxi Figure 4.22 Single wavelength absorbance traces following rapid mixing of 64 µM LS with varying concentrations of O-acetyl-L-serine .............................................................................. 121 Figure 4.23 The formation of the two tautomeric states of the internal aldimine, external aldimine and α-aminoacrylate between LS and O-acetyl-L-serine ........................................................... 122 Figure 4.24 A subset of the time resolved spectra following rapid mixing of O-acetyl-L-serine S234A ......................................................................................................................................... 123 Figure 4.25 Single wavelength absorbance traces following rapid mixing of S234A with varying concentrations of O-acetyl-L-serine at A) 406 nm and B) 460 nm over a log time scale. C) Linear dependence of kobs1 on O-acetyl-L-serine concentration. D) Hyperbolic dependence of kobs2 on O-acetyl-L-serine concentration...................................................................................................... 124 Figure 4.26 Change in fluorescence emission of wild type LS following the addition of A) D,L-lanthionine, B) D,L-2,6-diaminopimelic acid and C) meso-2,6-diaminopimelic acid ............... 127 Figure 4.27 Crystal structure of yeast CBS in the A) PLP-serine external aldimine state and B) α-aminoacrylate state...................................................................................................................... 129 Figure 4.28 Crystal structure of LS bound to serine .................................................................. 130 Figure 4.29 The bacterial biosynthetic pathway for the production of L-cysteine from L-serine utilising SAT and OASS ............................................................................................................. 131 Figure 4.30 Crystal structure of the active site of wild type LS with A) no ligand and B) L-lanthionine................................................................................................................................... 133 Figure 4.31 Sequence alignment between F. nucleatum LS and various OASS enzymes ........ 134 xxii Figure 5.1 A) SDS 8% polyacrylamide gel showing the purification of wild type ODA. B) SDS 10% polyacrylamide gel showing wild type ODA, the isolated ODC component and the isolated arginase component following purification by affinity chromatography and anionic exchange chromatography .......................................................................................................................... 147 Figure 5.2 Calibration curve determined from gel filtration ...................................................... 148 Figure 5.3 UV/visible spectra of wild type ODA the isolated ODC component and the isolated arginase component .................................................................................................................... 150 Figure 5.4 UV/visible spectra of wild type ODA in the presence of various substrates ........... 150 Figure 5.5 UV/visible spectra of wild type ODA, D119A, H119Q and K230M....................... 151 Figure 5.6 Michaelis-Menten plot of the production of CO2 in the presence of L-ornithine and wild type ODA and ODC/ODA .................................................................................................. 152 Figure 5.7: Michaelis-Menten plot of the production of CO2 in the presence of L-arginine by ODA variants ........................................................................................................................................ 154 Figure 5.8 The crystal structure of the dimerised from of ODC from Lactobacillus sp. 30a .... 158 xxiii List of Abbreviations ABH 2(S)-amino-6-boronohexanoic acid Arg/ODA isolated arginase component of ODA ATCC American Type Culture Collection BEC S-(2-boronoethyl)-L-cysteine CBS cystathionine β-synthase cDNA complementary deoxyribonucleic acid DEAE diethylaminoethyl DFMO difluoromethylornithine DNA deoxyribonucleic acid dNTP deoxynucleoside triphosphate FAD flavin adenine dinucleotide FiLTER Fipke Laboratory for Trace Element Research Fld flavodoxin FMN flavin mononucleotide FMNhq flavin mononucleotide hydroquinone FMNox flavin mononucleotide oxidised FMNsq flavin mononucleotide semiquinone FnFld flavodoxin from F. nucleatum FNR ferredoxin-NADP+ reductase FqrB flavodoxin-quinone reductase Gal-GalNAc galactose- and N-acetyl-d-galactosamine HPLC high performance liquid chromatography ICP-MS inductively coupled plasma mass spectrometry ITPG isopropyl β- d-1-thiogalactopyranoside LS lanthionine synthase miRNA micro ribonucleic acid NAD+ nicotinamide adenine dinucleotide NADH nicotinamide adenine dinucleotide reduced NADP+ nicotinamide adenine dinucleotide phosphate NADPH nicotinamide adenine dinucleotide phosphate reduced NCBI National Center for Biotechnology Information Ni-NTA Ni2+ nitrilotriacetic acid nor-NORA Nâhydroxyânorâarginine OASS O-acetyl-L-serine sulfhydrylase ODA ornithine decarboxylase/ arginase ODC ornithine decarboxylase ODC/ODA ornithine decarboxylase component of ODA PAGE polyacrylamide gel electrophoresis PCR polymer chain reaction PDB protein data bank xxiv PFOR pyruvate-flavodoxin oxidoreductase PLP pyridoxal phosphate PSI photosystem I Q-ToF quadrupole time-of-flight R.M.S root mean square RNA ribonucleic acid SAT serine acetyltransferase TIGIT T cell immunoreceptor with Ig and ITIM domains xxv Acknowledgements I would like to thank my fantastic supervisor, Dr. Kirsten Wolthers, for support, guidance and patience over the past four years. Her invaluable experience and mentorship have made this possible. I would also like to thank my committee, Dr. Kevin Smith and Dr. Isaac Li, for their support. I owe thanks to all the members of the Wolthers Lab over the last few years, including a special thank you to Dr. Osei Fordwour, who has been a good source of guidance and a great friend. I must thank the Chemistry admin staff, Karen Krysko and Susan Black, for making sure all my paperwork always got filed on time. I thank Dr. Mark Button for running ICP-MS experiments and Dr. Eric Dennis for assistance running HPLC experiments. I must also be grateful to our collaborators at UBC Vancouver, Maxim Kolesnikov, Angela Lin and Dr. Michael Murphy for their help and expertise with protein crystallography. I give thanks to my parents, Hugh and Deyo Mothersole, and my brother, Dr. David Mothersole, as well as all my friends for their amazing support over the years. Finally, I thank my incredible wife, Mina, who has helped me both outside the lab and, on occasion, inside the lab, and has been my best friend and greatest supporter over the last few years.1 1. Introduction 1.1 Fusobacterium nucleatum: an emerging pathogen Fusobacterium nucleatum is a gram-negative bacterium belonging to the Bacteriaceae family. It is commonly found in the human oral cavity.1 First identified in the early twentieth century, F. nucleatum forms small spindle-shaped rods that are 5 â 10 µm long with sharp pointed ends. 2, 3 It is an obligate anaerobe, but can survive conditions of up to 6% O2. F. nucleatum is split into four subspecies: F. nucleatum, F. polymorphum, F. fusiforme and F. animalis. This thesis will focus on the first of these subspecies, F. nucleatum subsp. nucleatum, with the type strain ATCC 25586. 1.1.1 F. nucleatum is associated with periodontal diseases A growing body of research over the past few decades has uncovered the link between F. nucleatum and several diseases. F. nucleatum was initially shown to play a role in periodontal disease, a group of diseases that affect the gingiva, alveolar bone and surrounding tissue.4 Periodontal disease often causes gum degradation, tooth pain and, in more serious cases, tooth loss.5 The disease is very common with some estimates suggesting around one third of the US population between 30â 90 years of age have either a mild, moderate or a severe form of the disease.6 Fortunately, despite being a fairly common disease, it is rarely fatal. F. nucleatum is able to co-aggregate with bacteria that populate dental plaque, a microbial biofilm occupying surfaces within the mouth.3 The formation of this biofilm takes place through several stages. In the initial stages, primary colonisers consisting of aerobic and facultative anaerobes such as Streptococcus species adhere to the tooth surface.7 F. nucleatum is able to adhere 2 to these primary colonisers through expressed adhesion proteins. For example, F. nucleatum expresses the adhesion proteins RadD, Aid1 and CmpA, which can bind to the streptococcal adhesion SpaP6.8 As the biofilm forms, F. nucleatum can aggerate with secondary colonisers such as the anaerobes Porphyomonas gingivalis and Treponema denticola using RadD, Fap2 and FomA, fusobacterial adhesin proteins. RadD also mediates adhesive interactions between F. nucleatum and other pathogenic bacteria such as Acinomyces naeslundii and fungi such as Candida albicans. F. nucleatum is often referred to as a âbridgingâ organism based on its ability to adhere to both early and late colonising pathogenic microbes.9 F. nucleatum also supports the biofilm microbial biofilm community through cross-feeding and metabolic interactions. For example, F. nucleatum accepts L-ornithine excreted by Streptococcus gordonii through the arginine-ornithine antiporter.10 In addition, F. nucleatum can adhere to and invade human epithelial cells. F. nucleatum enters epithelial cells via a zipping mechanism that involves the bacterium remaining in close contact with the host membrane during penetration.9 It is highly uncommon for oral bacteria to be able to penetrate host cells in this manner. Once inside the cell, F. nucleatum can mediate invasion of the cell by other bacteria associated with periodontal diseases, such as P. gingivalis.11 Moreover, F. nucleatum can induce the expression of β-defensin 2, an antimicrobial peptide, and IL-6 and IL-8, pro-inflammatory cytokines, in oral epithelium cells.12, 13 This results in inflammation of the cells which can promote tumour progression and increase the infectivity of other pathogens. 1.1.2 A link between F. nucleatum and abnormal pregnancy outcomes F. nucleatum is also associated with an increased risk of abnormal pregnancy outcomes, such as preterm and still births.14, 15 Preterm births are one of the most prominent causes of infant 3 mortality in developed countries.16 Risk factors commonly associated with pre-term births include the following: high or low maternal age, proximity in time to a previous delivery, ethnicity and race, low body mass-index and smoking or alcohol consumption.17 An additional risk factor for preterm births is periodontal disease. There is a 10-fold increase in preterm delivery before 28 weeks for women suffering from periodontal disease; this is believed to be due to the presence of F. nucleatum that has crossed into the placenta.18 Studies involving mice injected with F. nucleatum have also routinely shown an increased chance for preterm and still births during pregnancy.14, 19 An investigation by Han et al (2004) demonstrated that as the infected dose of F. nucleatum was increased, the likelihood of foetal death also increased. They showed that the bacterium did not persist in the liver or spleen, and that counts decreased over time such that it was eliminated within 24 h. However, F. nucleatum was shown to colonise the placenta and spread to the amniotic fluid and foetuses within 48 to 72 h, during which time foetal death was recorded. A similar investigation by Stockham et al (2015) identified that of the four subspecies, only F. nucleatum subsp. nucleatum was able to successfully penetrate the placenta. Only this strain was able to induce foetal miscarriage and, once again, F. nucleatum was not detected in the blood or spleen. These studies suggest that some strains of F. nucleatum can cross haematologically from the mother to the placenta, enabling access to the foetal circulation. 1.1.3 Recent links to colorectal cancer More recently F. nucleatum has been linked to colorectal carcinoma.20 Colorectal carcinoma is the third most commonly diagnosed form of cancer worldwide and is the fourth highest cause of cancer death. 16, There are over 130,000 new cases and an estimated 50,000 deaths from colorectal cancer each year in the United States.21, 22 Whole genome sequencing of the microbiota located in and around colorectal carcinomas identified Fusobacterium as the most 4 abundant taxon.23 In another study, F. nucleatum genomic DNA was shown to be 400 times more abundant in the colorectal carcinoma compared to surrounding healthy tissue, suggesting that there may be a link between the bacteria and tumorigenesis.24 A study by Mima et al (2015) which followed over 1000 colorectal cancer patients showed that the abundance of F. nucleatum DNA in colorectal cancer tissue correlated with reduced survival rate.25 While the exact mechanism by which F. nucleatum is involved in tumorigenesis is not completely understood, there are a number of properties of F. nucleatum that can provide insight into the relationship between the bacterium and tumour progression.26 F. nucleatum is able to adhere to E-cadherin to activate β-catenin signalling via the FadA adhesion, a virulence factor expressed by the bacterium.27 The activation of β-catenin signalling leads to increased expression of transcription factors, oncogenes, inflammatory genes, and stimulates growth of colorectal cancer cells. Furthermore, the Fap2 protein produced by F. nucleatum has been shown to interact with TIGIT, an immune receptor present on T Cells and Natural Killer Cells. This results in the inhibition of immune cell activities against colorectal tumours.28 ApcMin/+ mice exposed to F. nucleatum exhibit a proinflammatory expression signature, but do not exacerbate colitis, enteritis, or inflammation-associated intestinal carcinogenesis that is commonly associated with other bacteria linked to colorectal carcinoma.29 In addition, the fusobacterial adhesin Fap2 can recognise the specific sugar residue, Gal-GalNAc, which is overexpressed in colorectal tumorigenesis, enabling further accumulation of F. nucleatum within the tumour.30 F. nucleatum can suppress specific miRNAs involved in modulation of autophagy, this can result in the development of chemoresistance on the colorectal cancer cells.31 1.1.4 Three proteins from F. nucleatum have been studied 5 Given the strong association of F. nucleatum with colorectal cancers, preterm births and other conditions, it is important to find methods to selectively target the growth of F. nucleatum. One method is to inhibit specific proteins that perform vital functions for the survival of the organism. By disabling these proteins, growth of the bacteria can be slowed or stopped and patients with F. nucleatum infections may have better health outcomes. In this thesis, we have investigated and characterised three proteins encoded in the F. nucleatum genome that are potential drug targets. These three proteins are: an electron transfer flavodoxin (Fld), lanthionine synthase (LS) and a bifunctional ornithine decarboxylase/ arginase (ODA). 1.2 Flavodoxin Flavodoxins (Fld) are small electron transfer proteins involved in both photosynthetic and non-photosynthetic reactions in many bacteria and some species of algae.32 Flds contain a non-covalently bound flavin mononucleotide (FMN, Figure 1.1) cofactor that enables the protein to serve as a low potential cellular reductant, connecting a multitude of oxidation-reduction pathways. Flds were first discovered in the mid-1960s in blue-green algae (also known as cyanobacteria) and the nitrogen fixing bacterium, Clostridium pasteurianum.33, 34 In these organisms, Fld was found to substitute for ferredoxin, an iron-sulfur containing electron transfer protein, in low-iron conditions. 6 Figure 1.1 Structure of the FMN cofactor. 1.2.1 Role of flavodoxins in anaerobic microorganisms In nitrogen fixing bacteria, ferredoxin and Fld are iso-functional as both transfer electrons from NADPH-ferredoxin reductase to nitrogenase, thereby coupling NADPH oxidation to N2 reduction. In cyanobacteria such as Anabaena and Synechococcus, ferredoxin and Fld shuttle electrons from photosystem I (PSI) to a wide range of metabolic processes. Most of the photosynthetic derived reducing power is directed to ferredoxin-NADP+ reductase (FNR), which supplies NADPH for CO2 fixation in the Calvin-Benson-Bassham cycle along with other anabolic reactions.35, 36 Another reaction involving flavodoxin in enteric bacteria is the reductive activation of cobalamin-dependent methionine synthase. In this reaction Fld donates an electron to the cobalamin cofactor of methionine synthase which enables transfer of a methyl group from S-adenosylmethionine to homocysteine to produce methionine. F. nucleatum encodes several other enzymes that rely on electron shuttling from flavodoxin including the electron bifurcating butryl-acyl CoA dehydrogenase electron transfer flavoprotein (FN0783/84/85), biotin synthase (FN1000), ribonucleotide reductase (FN0102/FN0103), pyruvate-formate lyase activating enzyme (FN0621) and the Rnf-Na+ pump (FN1591-6).32 The F. nucleatum genome also encodes for 7 pyruvate-flavodoxin oxidoreductase (PFOR; FN1170) and flavodoxin-quinone reductase (FqrB; FN1984). PFOR catalyses the oxidative decarboxylation of pyruvate in the presence of coenzyme A to form acetyl-coenzyme A, which is a vital component of the citric acid cycle.37, 38 The electron generated from the oxidative decarboxylation is transferred from Fld to flavodoxin-quinone reductase (FqrB), which then reduces NADP+ to NADPH (Figure 1.2). Figure 1.2 Reversible process of electron shuttle from PFOR to FqrB and NADP+ via Fld. Bacteria such as Helicobacter pylori cannot grow without Fld, due to its participation in this specific pathway.39 Furthermore, the gene coding for Fld has been proven to be vital for the survival and growth of other pathogenic bacteria including Streptococcus pneumonia, Haemophilus influenza, E. coli, Salmonella enterica and Vibrio cholera.40-44 F. nucleatum is a flavodoxin specialist, in that its genome encodes for seven distinct flavodoxin and no genes for ferredoxin. A bioinformatic study of over 7000 organisms of Archaea, Bacteria and Eukarya, revealed that ~2% of these organisms are flavodoxin specialists, and belong to either Fusobacteria or Lactobacillus genera.45 Becoming an Fld specialist may be a protective strategy that F. nucleatum has adopted over time to cope with reactive oxygen species produced 8 by streptococcal species also found in dental plaque.46 Iron availability is also predicted to be limited in this environmental niche, so it may also be an approach to compensate for this low iron environment.47 The diversification of the flavodoxin pool in F. nucleatum suggests that the organism differentially controls the flux of electrons across distinct metabolic pathways. 1.2.2 General structure of flavodoxins Flavodoxins have a characteristic motif consisting of a five-stranded parallel β-sheet flanked by a pair of α-helices (Figure 1.3).48 FMN binds at the C-terminal end of the β-sheet. There are two main subsets of flavodoxins, long-chain and short-chain. Long chain flavodoxins contains an addition lobe of ~20 amino acids in the middle of the β5-strand. This lobe has no direct interaction with the FMN cofactor and no known biological function. 9 Figure 1.3 Cartoon of the crystal structure of Fld from Anabaena (PDB entry code 1FLV). The FMN cofactor is shown in grey, α-helices are shown in red and β-sheets in yellow. FMN binds to Fld via three loops: a phosphate binding loop (residues 10 â 15, FnFld numbering), which forms hydrogen bond interactions with the FMN 5â-ribityl phosphate group, a 50âs binding loop (residues 56 â 62) and a 90âs binding loop (residues 90 â 99).49 The 50âs and 90âs loops contain aromatic side chains that sandwich the isoalloxazine ring of FMN, forming extensive Ï-Ï interactions.50 Typically, a tryptophan residue in the 50âs loop sits on the re-face of the FMN isoalloxazine ring, while a conserved tyrosine residue in the 90âs loop is positioned on the si-face of the cofactor. 1.2.3 FMN enables electron shuttling FMN and flavin adenine dinucleotide (FAD) are the biologically active forms of riboflavin (vitamin B2), proteins that bind these coenzymes are referred to as flavoproteins. Flavoproteins are found in all domains of life.51 FAD contains an additional adenine diphosphate group.52 Flavins are organic molecules with a distinctive tricyclic isoalloxazine ring system that can interconvert between three oxidation states (Figure 1.4): oxidised (quinone), one-electron reduced (semiquinone) and two-electron reduced (hydroquinone).53 Over 90% of flavin-dependent proteins function as oxidoreductases in which the flavin cofactor cycles between two or three of these oxidation states.52 10 Figure 1.4 The three oxidation state of the FMN cofactor. The presence of the FMN cofactor enables Fld to shuttle single electrons to a variety of redox partner proteins. The binding of FMN to apo-protein dramatically perturbs the FMN reduction potentials, enabling it to participate in one electron transfer processes.54, 55 For free FMN in solution, only the hydroquinone and quinone forms of the cofactor form as the semiquinone/hydroquinone midpoint potential (Esq/hq, -172 mV), is lower than the oxidised/semiquinone midpoint potential (Eox/sq, -238 mV).56 However, binding of FMN to the apo-protein inverts these potentials, through stabilisation of the FMN semiquinone and destabilisation of the negatively charged FMN hydroquinone. Consequently, Eox/sq of Fld bound FMN increases to around -90 to -150 mV and Esq/hq decreases to -390 to -450 mV.54-56 The latter potential enables Fld to effectively substitute for ferredoxin as a low potential cellular reductant. In vivo, Fld primarily cycles between the semiquinone and hydroquinone states. For the majority of flavodoxins characterised to date, the FMNsq apo-protein complex is secured through the formation of an additional hydrogen bond between the N5(H) of the FMN semiquinone and a residue on the 50âs binding loop, I59 in Anabaena Fld.57 The negatively charged FMNhq is destabilised by neighbouring negatively charged amino acids residues surrounding the FMN binding pocket.58 The hydrophobic residues that sandwich the FMN isoalloxazine ring also contribute to destabilisation of FMNhq by increasing the electrostatic repulsion between the cofactor and protein through desolvation. 1.2.4 Comparing flavodoxins from multiple species FnFld is a long-chain flavodoxin as it contains an additional 20 residues within the β5-strand. Sequence alignment of a selection of flavodoxins from different species with FnFld using 11 MUSCLE (Figure 1.5) identified substitutions of four conserved residues within the amino acid sequence FnFld (K13, Y56, N89 and F93).59 K13 replaces a conserved glycine residue found in the phosphate binding loop with the T/S-X-T/S-G-X-T/S sequence. The serine/threonine residues within this loop interact with the 5â-ribityl phosphate group by forming hydrogen bonds.60 Glycine is important in the region as it enables the phosphate binding loop to remain flexible, enabling peptide backbone the adopt Ï and É¸ torsion angles of -11 °and Φ = 103°, normally disallowed for chiral amino acids. F93 and Y56 correlate with W94 and W57 in Anabaena Fld, residues that provide stabilising interactions with the isoalloxazine ring.61 The final residue, N89 replaces an aspartic acid that is known to have a significant effect on Esq/hq in other flavodoxins.62 In addition, the number of acidic residues in FnFld is much lower than other flavodoxins and the number of basic residues much higher (Table 1.1). The theoretical isoelectric point (pI) was determined to be 5.3, higher than that calculated for generic flavodoxins.63 The acidic qualities imparted on flavodoxin enable it to promiscuously interact with multiple redox partners which typically have basic binding clefts. It also enables flavodoxin to effectively substitute for ferredoxin, which is also an acidic protein. 12 Figure 1.5 Sequence alignment a selection of flavodoxins from different species aligned with MUSCLE: FUSON (Fusobacterium nucleatum strain ATCC 25586), PSEUD (Pseudomonas aeruginosa), CLOSB (Clostridium beijerinckii), DESUV (Desulfovibrio vulgaris), CHOND (Chondrus crispus), HELPI (Helicobacter pylori), ECOLI (Escherichia coli), ANABA (Anabaena), SYNEL (Synechococcus elongates), AZBVI (Azotobacter vinelandii), and RHODO (Rhodobacter capsulatus). Table 1.1 Distribution of acidic and basic residues in flavodoxins from various organisms. These were determined using the ProtParam tool from ExPASy.64 Organism # of acidic residues # of basic residues Net Charge Theoretical pI F. nucleatum 24 20 -4 5.3 D. vulgaris 30 11 -19 4.1 C. beijerinckii 28 12 -16 4.2 E. coli (FldA) 37 13 -24 4.2 Anabaena 31 15 -16 4.1 S. elongates 27 10 -17 4.0 D. desulfuricans 29 10 -19 4.1 FUSON -MKTIGIFYATLTKTTVGIVDEIEFFLKKD--DFKTFNVKNGVKE--IENFâ-ENLILVTPTYQVGEA--------HAAWMNN-LKKLEEIâ-DFTGKVVGLPSEUD --MKVAILSGSVYGTAEEVARHAQK-LLSAA-GLEASHLPRASLDE-LKAFAPEAFLVVTSTTGMGEL--------PDNLQPL-YYAIRDQLPAWHGLPGGVDESUV -MPKALIVYGSTTGNTEYTAETIARELADAGYEVDSRDAASVEAGGLFEGFâ-DLVLLGCSTWGDDSI------ELQDDFIPL-FDSLEET--GAQGRKVACCHOND ---KIGIFFSTSTGNTTEVADFIGKTLGAKA-DAPIDVDDVTDPQA-LKDYâ-DLLFLGAPTWNTGAD----TERSGTSWDEFLYDKLPEVâ-DMKDLPVAIHELPI -MGKIGIFFGTDSGNAEAIAEKISKAIGNA----EVIDVAKASKDQ-FNSFâ-TKVILVAPTAGAGDL--------QTDWEDF-LGTLEASâ-DFANKTIGLECOLI -MAITGIFFGSDTGNTENIAKMIQKQLGKD--VADVHDIAKSSKED-LEAYâ-DILLLGIPTWYYGEA--------QCDWDDF-FPTLEEIâ-DFNGKLVALANABA MSKKIGLFYGTQTGKTESVAEIIRDEFGND--VVTLHDVSQAEVTD-LNDYâ-QYLIIGCPTWNIGEL--------QSDWEGL-YSELDDVâ-DFNGKLVAYSYNEL -MAKIGLFYGTQTGVTQTIAESIQQEFGGES-IVDLNDIANADASD-LNAYâ-DYLIIGCPTWNVGEL--------QSDWEGI-YDDLDSVâ-NFQGKKVAYAZBVI -MAKIGLFFGSNTGKTRKVAKSIKKRFDDET-MSDALNVNRVSAED-FAQYâ-QFLILGTPTLGEGELPGLSSDCENESWEEF-LPKIEGLâ-DFSGKTVALRHODO -MAKIGLFFGSDTGTTRKIAKQIKDMFDDEV-MAKPLNVNRADVAD-FMAYâ DFLILGTPTLGDGQLPGLSANAASESWEEF-LPRIADQâ-DFSGKTIALFUSON VGLGNQFAFGESFCGGIRYLYDIVVKKGGKVVGFTSTDGYHYEETSIIENGK-FIGLALDEENQPNLTPKRIGDWITEIKKEFK-------PSEUD IGLGDS-SYGDTFCGGGEQVRELFGELGVREV---------------------LPMLRLDAS-ETVTPETDAEPWLAEFAAALKG------DESUV FGCGDS-SY-EYFCGAVDAIEEKLKNLGAEIV---------------------QDGLRIDGD-âPRAARDDIVGWAHDVRGAI--------CHOND FGLGDAEGYPDNFCDAIEEIHDCFAKQGAKPVGFSNPDDYDYEESKSVRDGK-FLGLPLDMVNDQIPMEKRVAGWVEAVVSETGV------HELPI VGLGDQDTYSETFAEGIFHIYE--KAKAGKVVGQTPTDGYHFEASKAVEGGK-FVGLVIDEDNQDDLTDGRISKWVEQIKGSFA-------ECOLI FGCGDQEDYAEYFCDALGTIRDIIEPRGATIVGHWPTAGYHFEASKGLADDDHFVGLAIDEDRQPELTAERVEKWVKQISEELHLDEILNAANABA FGTGDQIGYADNFQDAIGILEEKISQRGGKTVGYWSTDGYDFNDSKALRNGK-FVGLALDEDNQSDLTDDRIKSWVAQLKSEFGL------SYNEL FGAGDQVGYSDNFQDAMGILEEKISSLGSQTVGYWPIEGYDFNESKAVRNNQ-FVGLAIDEDNQPDLTKNRIKTWVSQLKSEFGL------AZBVI FGLGDQVGYPENYLDALGELYSFFKDRGAKIVGSWSTDGYEFESSEAVVDGK-FVGLALDLDNQSGKTDERVAAWLAQIAPEFGL--SLâRHODO FGLGDQVTYPLEFVNALFFLHEFFSDRGAKLVGRWPAKGYGFEDSLAVVEGE-FLGLALDQDNQAALTPERLKGWLSLIAADFGL--VLPAβ1 β2 β3 β4β5 β6α1 α2 α3α4 Î±550âs loop90âs loop long loopβ413 1.2.5 Preliminary success with flavodoxins as a drug target H. pylori is one of the main causes of stomach ulcers and currently the most common treatment consists of broad-spectrum antibiotics. However, in an attempt to reduce reliance on antibiotics investigators have looked into treating the disease using Fld as a therapeutic target.65 High-throughput screening of over 10,000 chemicals led to the identification of 29 compounds that could bind to Fld in a pocket near the active site and interrupt in vitro electron transfer between Fld and its redox partner proteins.66 Four of these compounds were shown to be bactericidal against H. pylori and none of the compounds were toxic to mice. The therapeutic index of three of the four inhibitors was improved through the design and synthesis of analogues.67 Five of these second generation inhibitors are shown in Figure 1.6. The most effective derivatives were found to contain a nitro group, such as nitroethylene and 7-nitrobenzoxadiazol.68 Once again these compounds were shown to be non-toxic in mouse models. These studies demonstrate the feasibility of using Fld as a drug target to treat bacterial infections, including those caused by F. nucleatum. 14 Figure 1.6 A selection of compounds that have inhibitory effects on H. pylori flavodoxin electron transfer. (I) 6,8-dichloro-2-(4-fluorophenyl)-3-nitro-2H-chromene, (II) 1-(2-nitrovinyl)-3,5-di(trifluoromethyl), (III) 4-[(4-methoxybenzy)thio]-7-nitro-2,1,3-benzoxadizole, (IV) 4-fluoro-β-nitrostyrene and (V) N,Ndimethyl-7-nitro-2,1,3-benzoxadiazol-4-amine. 1.3 Lanthionine synthase The second enzyme that will be examined in this thesis is lanthionine synthase (LS) encoded by the FN1220 gene. In the literature, LS is also annotated as Fn1220 and cysteine synthase. The latter annotation stems from its sequence similarity to cysteine synthase (also referred to as O-acetylserine sulfhydrylase). LS is a pyridoxal 5â-phosphate (PLP) dependent enzyme that catalyses the formation of lanthionine and H2S from two molecules of cysteine via a β-replacement reaction (Figure 1.7).69 Figure 1.7 Production of lanthionine from two molecules of L-cysteine catalysed by lanthionine synthase. 1.3.1 Meso-lanthionine is a component of the F. nucleatum peptidoglycan layer Meso-lanthionine is vital to the survival of F. nucleatum as it is a constituent of the peptidoglycan layer.70 In gram-negative bacteria, such as F. nucleatum, the peptidoglycan layer forms a thin cell wall that is flanked by an outer and inner membrane.71, 72 The peptidoglycan layer consists of a heteropolysaccharide composed of alternating N-acetylglucosamine and N-acetylmuramic acid. The repeated layers of the heteropolysaccharide are linked by a peptide cross-15 link. This peptide cross-link normally consists of L-alanine, D-glutamic acid, meso-diaminopimelic acid and D-alanine.73 The biosynthesis of meso-diaminopimelic acid is performed in other bacteria over the course of eight steps from L-aspartate.74 However, F. nucleatum lacks this biosynthetic pathway. Consequently, in the F. nucleatum peptidoglycan layer, meso-diaminopimelic acid is replaced with its analogue, meso-lanthionine.70 The only known synthetic route for lanthionine is LS. This potentially makes LS a viable therapeutic target for treating F. nucleatum infections. 1.3.2 Importance of H2S in bacteria The genome of F. nucleatum harbours four genes (FN1220, FN0625, FN1055 and FN1419) that encode for distinct enzymes that catalyse the production of H2S from L-cysteine.75 LS is the major producer of H2S in F. nucleatum, accounting for over 87% of H2S production by the bacterium.75 H2S has a distinctive ârotten eggâ smell and is toxic at concentration > 50 ppm. However, it is produced endogenously at low levels in different organisms, where it serves as a gasotransmitter. In mammals, H2S is involved in blood vessel dilation, inflammation, cardiac reactions to ischemia, regulation of the nervous system and secretion of insulin.76-78 In humans, high levels of H2S are associated with respiratory conditions, such as chronic bronchitis, emphysema and pneumonia, as well as cardiovascular diseases such as hypertension.77 In bacteria, H2S can provide defence against reactive oxygen species and has been associated with bacterial resistance to antibiotics.79, 80 In addition, H2S is used as a major cytoprotectant and redox modulator in bacteria.75, 81 1.3.3 PLP-dependent enzymes 16 PLP is the biologically active form of vitamin B6. It is associated with a variety of enzymes that catalyse a number of different reactions on amino acids.82 For example, PLP-dependent enzymes can catalyse decarboxylations, transaminations, racemisations and elimination or replacement reactions at the β-or γ-positions of amino acids.83 PLP-dependent enzymes account for 1.5% of all genes encoded for by prokaryotes.84 All PLP-dependent enzymes bind to the PLP cofactor through the formation of an imine linkage between a lysine residue in the enzyme active site and the aldehyde group of PLP. This forms a Schiff base, also known as the internal aldimine. This lysine is displaced upon formation of the external aldimine with the amino acid substrate. The mechanistic pathway of PLP-dependent enzymes diverges at this point, which is largely governed by active site residues that coordinate to PLP and substrates. Generally, there is heterolytic cleavage of a bond to the Cα (i.e. Cα-H or Cα-COO-) leading to formation of the carbanionic species, which is resonance stabilised through electronic delocalisation through the Schiff base and PLP pyridine ring.83 The structure of PLP-dependent enzymes are divided into five different folds: Fold Type I-V.85 Fold Type I (aspartate aminotransferase family) is the most structurally and functionally diverse family of PLP-dependent enzymes. These enzymes normally function as homodimers or higher-order oligomers. The homodimeric enzymes contain two actives site per dimer, which are situated on the dimer interface.86 Both monomers contribute residues to both active sites. The PLP cofactor binds such that the si-face is solvent exposed. Fold Type I typically contains a conserved aspartate residue that interacts with the PLP ring N atom, which helps stabilise PLP intermediates. The subunit of Fold Type I contains a large domain, consisting of a 7-stranded β-sheet at the N-terminus, and a small domain, consisting of 3- or 4-stranded β-sheet with α-helices on one side at the C-terminus. 17 Structures of Fold Type II enzymes (tryptophan synthase family) are similar to those from Fold Type I, but have distinct evolutionary routes.83 The most significant difference between Fold Type I and Fold Type II, is that each the active sites are composed solely of residues from one monomer. In addition, the Schiff base lysine residue resides closer to the N-terminus. The small domain at the C-terminus consists of a 2-stranded β-sheet surrounded by α-helices.85 Other PLP-dependent enzymes that belong to Fold Type II include serine dehydratase, threonine synthase, O-acetyl-L-serine sulfhydrylase and cystathionine β-synthase.86 LS is a Fold Type II PLP-dependent enzyme.87 Fold Type III (alanine racemase family) have the Schiff base lysine close to the N-terminus similar to Fold Type II. However, this lysine is preceded by a hydrophobic β-strand and there are two β-strands at the C-terminus.88 Fold Type IV (D-amino acid aminotransferase family) also function as homodimers. They differ as the PLP cofactor binds with the re-face solvent exposed. The N-terminal domains consists of a six-stranded antiparallel β-sheet and two α-helices. The final group, Fold Type V (glycogen phosphorylase family), is the most distinct group. The PLP phosphate group is involved in proton transfer during the enzymatic reaction for this group of enzymes. 1.3.4 Sequence similarity with O-acetyl-L-serine sulfhydrylase and cystathionine β-synthase A sequence homology search performed by NCBI BLAST shows that LS shares 48% sequence similarity with O-acetyl-L-serine sulfhydrylase (OASS) (EC 2.5.1.47) from Mycobacterium tuberculosis.89 The physiological reaction of OASS is the conversion of O-acetyl-L-serine and sulfide to L-cysteine and acetate. This reaction is the final step in the cysteine biosynthetic pathway. The precursor step in this pathway is serine acetyltransferase (SAT), which 18 transfers an acetyl group from acetyl-CoA to serine to form O-acetyl-L-serine.90 In bacteria such as M. tuberculosis, L-cysteine is an important building block for de novo protein synthesis.91 As with other PLP-dependent enzymes, the resting form of the enzyme contains a PLP cofactor that is bound to the enzyme via K44 (M. tuberculosis OASS numbering) to form the internal aldimine (Figure 1.8).90, 92 The reaction is initiated by the displacement of K44 by O-acetyl-L-serine to form an external aldimine with O-acetyl-L-serine via a gem-diamine intermediate. This is succeeded by abstraction of the Cα proton by K44. This leads to the β-elimination of acetate and formation of an α-aminoacrylate intermediate. The next step involves nucleophilic attack of HS- on the β-carbon of the α-aminoacrylate intermediate. Reprotonation of the Cα leads to formation of an external aldimine with L-cysteine, which is then released as the internal aldimine is regenerated. 19 Figure 1.8 Reaction mechanism of the formation of L-cysteine O-acetyl-L-serine via an α-aminoacrylate intermediate in M. tuberculosis OASS.92 However, OASS displays substrate promiscuity in that L-cysteine can also form an external aldimine with the enzyme in the absence of O-acetyl-L-serine (Figure 1.9) (Steps 1 and 2).93-95 Abstraction of the Cα proton by K41 leads to β-elimination of H2S and the formation of an α-aminoacrylate intermediate (Step 3).96 Nucleophilic attack by the thiolate from a second cysteine molecule on the Cβ and reprotonation of the Cα leads to the generation an external aldimine with L-lanthionine (Step 4). Product release leads to reformation of the internal aldimine (Steps 5 and 6). This is the mechanism by which LS produces L-lanthionine and H2S from two equivalent of L-cysteine. 20 Figure 1.9 Mechanism of the formation of L-lanthionine from two molecules of L-cysteine via an α-aminoacrylate intermediate in S. typhimurium OASS.93 Sequence homology performed by NCBI BLAST show that LS shares 37% sequence similarity with human cystathionine β-synthase (CBS) (EC 4.2.1.22).89 CBSâs canonical reaction involves the condensation of L-serine and L-homocysteine via a β-replacement reaction to form L-cystathionine.97 This is part of the biosynthetic route for L-cysteine from L-methionine (i.e. the reverse transsulfuration pathway). A second PLP-dependent enzyme, cystathionine-γ-lyase, catalyses the γ-elimination of cystathionine to produce ammonia, α-ketobutyrate and L-cysteine. In humans, L-cysteine is used in the synthesis of glutathione, an antioxidant. Production of L-cysteine also provides a catabolic route for the removal of excess sulfur containing amino acids from the body, as L-cysteine can undergo oxidation to produce CO2, pyruvate and SO42-. The 21 reaction begins with the displacement of K129 (human CBS numbering) by the amino group of L-serine to form the external aldimine via a gem-diamine intermediate (Figure 1.10).98 This is followed by abstraction of the Cα proton by K129, leading to β-elimination of H2O and the formation of an α-aminoacrylate intermediate.96 Nucleophilic attack by the thiolate from a molecule of L-homocysteine on the Cβ and reprotonation of the Cα leads to the generation an external aldimine with L-cystathionine. Product release leads to reformation of the internal aldimine. Figure 1.10 Mechanism of the formation of L-cystathionine from L-cysteine and L-homocysteine via an α-aminoacrylate intermediate in human CBS.98 However, CBS too displays substrate and catalytic promiscuity and can catalyse three additional side reactions that lead to the production of H2S. These reactions include the reaction 22 between two equivalents of L-cysteine to produce L-lanthionine and H2S, the condensation of L-cysteine and L-homocysteine forming cystathionine and H2S, and the β-elimination of L-cysteine producing L-serine and H2S (Figure 1.11).99 These side reactions of CBS, along with phycological reaction of cystathionine-γ-lyase, are a source of H2S in humans. While large quantities of H2S are toxic to humans, studies indicate that smaller concentrations can be beneficial. Physiological roles of H2S include stimulation and inhibition of the mitochondrial electron transfer, regulation of L and T type calcium channels and reducing the production of reactive oxygen species.100, 101 In the cardiovascular system, H2S can act as a vasodilator and an antihypertensive.102 In addition, H2S is utilised in various ways by the respiratory system, the central nervous system, the liver and the kidneys.103 H2S is also involved in a post translational modification called persulfidation, where the thiol groups on reactive cysteines residues are modified to persulfide groups.104 Figure 1.11 Four reactions catalysed by human CBS. Reaction 1 is the physiological reaction, the β-replacement of L-serine with L-homocysteine to produce cystathionine and H2O. Reaction 2 is the β-replacement of L-cysteine with L-homocysteine to produce cystathionine and H2S. Reaction 23 3 is the β-elimination of L-cysteine producing L-serine and H2S. Reaction 4 is the formation of L-lanthionine from the β-replacement L-cysteine by a second molecule of L-cysteine. 1.3.4 Structural insight gained from crystal structures of lanthionine synthase The crystal structures of the internal aldimine of LS (PDB entry code 5XEO) and the external aldimine with serine (PDB entry code 5XEN) and L-lanthionine (PDB entry code 5XEM) have been solved and are available on the PDB database.105 The overall structure of LS is typical of a Fold Type II PLP-dependent enzyme (Figure 1.12).87 There is a 6-stranded β-sheet flanked by α-helices at the N-terminal end with the PLP factor at the end of these α-helices. The C-terminal end contains a three stranded β-sheet surrounded by α-helices. LS forms a dimer, but the active sites of each monomer are independent of one another. These crystal structures depict several residues important for PLP and substrate binding that are conserved between LS, CBS and OASS. In the internal aldimine conformation (Figure 1.13) K42 forms a Schiff base with PLP, S276 forms a hydrogen bond with the PLP pyridine nitrogen and N72 forms a hydrogen bond with the 3â oxygen.106 A glycine rich loop (residues 186GTGGS190) creates an α-helix that forms a hydrogen bond network with the PLP phosphate.107 CBS and OASS both undergo a conformational change that is triggered by the binding of the substrate α-carboxylate to a conserved loop within the active site.108, 109 This conserved loop is also present in LS (79TSGNT83); thus, it is likely that LS also undergoes the same substrate-induced conformational change (Figure 1.14). 24 Figure 1.12 Cartoon depiction of the homodimer of LS (PDB entry code 5XEO). One monomer is coloured in grey and the other is coloured according to the secondary structure. α-helices are in red, β-sheet in yellow and loops in green. The PLP cofactor is shown in grey. Figure 1.13 Crystal structure of the internal aldimine of LS (PDB entry code 5XEO) showing covalent imine linkage with K52 and hydrogen bond interactions between S276, N82 and the PLP cofactor. 25 Figure 1.14 Crystal structure showing the 69TSGNT73 loop in LS A) With no substrate (PDB entry code 5XEO) and B) With L-lanthionine (shown in cyan) (PDB entry code 5XEM). The PLP cofactor is shown in grey. 1.4 Ornithine decarboxylase/ arginase Ornithine decarboxylase/arginase is a bifunctional enzyme unique to F. nucleatum. The N-terminal half of the protein (residues 1 â 500) contains the ornithine decarboxylase (ODC) component while the C-terminal half (residues 500 â 780), contains the arginase component.110 The arginase component is responsible for the hydrolysis of L-arginine to L-ornithine and urea while the decarboxylation of L-ornithine to putrescine and CO2 is catalysed by the ODC component (Figure 1.15).111 26 Figure 1.15 Reactions catalysed by ODA, the conversion of L-arginine to L-ornithine and urea followed by conversion of L-ornithine to putrescine and CO2. 1.4.1 The ornithine decarboxylase component Ornithine decarboxylase (ODC, EC 4.1.1.17) is a PLP-dependent enzyme found throughout all domains of life.112 ODC falls into one of two PLP-fold types, dependent on their source. Prokaryotic ornithine decarboxylases, such as that from Lactobacillus 30a., belong to Fold Type I, whereas eukaryotic ornithine decarboxylases, such as that from mice, belong to Fold Type III.88 Eukaryotic and prokaryotic forms of ODC are structurally unrelated as they developed via different evolutionary routes.113 The ODC component of ODA is homologous to the group I PLP-dependent decarboxylases. A handful of X-ray structures from the family of group I PLP-dependent decarboxylases has been published. They include arginine decarboxylase from E. coli (PDB entry code 2VYC), ornithine decarboxylase from Lactobacillus sp. 30a (PDB entry code 1ORD), lysine decarboxylase from E. coli (PDB ID: 3N75) and arginine decarboxylase from Salmonella typhimurium (PDB entry code 5XX1).114-116 All three of these proteins form quaternary structures consisting of 5 or 6 sets of homodimers, as such they exist as either decamers or dodecamers. Each dimer has two active sites that lie at the interface of two of the monomers.117 Lactobacillus sp. 30a ODC contains 27 domains that resembles the large and small domains of Fold Type I PLP-dependent enzymes. This includes a 7-stranded β-sheet flanked by α-helices. 1.4.2 Catalytic mechanism of ornithine decarboxylase While there are significant differences between the structural motifs of prokaryotic and eukaryotic ODC, their catalytic mechanism is similar. In the resting form of the enzyme, an internal aldimine is formed between PLP and an active site lysine residue. The initial step of the mechanism involves transimination, or formation of an imine linkage between the α-amine of L-ornithine and PLP, referred to as the external aldimine (Figure 1.16).118 This step is followed by α-decarboxylation to form a quinonoid intermediate, which is stabilised by electron delocalisation through the Schiff base and pyridine ring. The Cα is then protonated, forming the external aldimine with putrescine. Finally, transimination by the active site lysine releases the products and restores the enzyme to the internal aldimine state for another round of the catalytic cycle. Putrescine is a polyamine and a precursor to other polyamines such as spermidine and spermine.119 Polyamines are highly regulated aliphatic cations that can bind to DNA and modulate DNA-protein interactions.120 These polyamines are essential for cell proliferation and differentiation, two functions necessary for the survival and growth of an organism.121 Mutants of E. coli deficient in polyamine production have a 3-fold lower growth rate compared with wild type strains; a similar pattern has also been observed in mammalian cells deprived of polyamines.122, 123 Thus, it is conceivable that the ODC component of ODA is a good potential drug target for F. nucleatum. 28 Figure 1.16 ODC catalytic mechanism for the production of putrescine and CO2 from L-ornithine via a quinonoid intermediate mediated by the ODC component of ODA. 1.4.3 Treatments targeting ornithine decarboxylase Targeting ODC as a method to treat disease is not a novel idea. African trypanosomiasis, also known as African sleeping sickness, is a potentially fatal disease that affects rural populations in sub-Saharan Africa. The disease is caused by the Trypanosoma brucei parasite.124 It is transmitted by the bite of tsetse flies and since the beginning of the 20th century it has been responsible for the deaths of millions of people. In the late 20th century a treatment for this disease focussed on targeting ODC from the T. brucei parasite. It was discovered that treatment with difluoromethylornithine (DFMO), an analogue of ornithine, could selectively and irreversibly inhibit ODC and kill off the parasite.125 This treatment dramatically increased the survival rate 29 amongst patients suffering from African trypanosimiasis.124 DMFO acts through mechanism based inactivation of ODC (Figure 1.17). It forms a covalent adduct with a cysteine and a lysine residue found in the active site. 126 However, DFMO is only effective against eukaryotic ODC and not prokaryotic ODC as the latter does not contain an active site cysteine.113 Therefore it is unlikely that DFMO will inhibit the ODC component of ODA. Figure 1.17 Mechanism of irreversible inhibition of mouse ODC by DFMO. There are two routes that can be taken from the carbonic anion. The first (a) involves the elimination of the F- leaving group to form a conjugated imine. The second (b) involves the formation of a tautomeric imine. These both ultimately end in the same final product with DFMO covalently bound to the enzyme via a nucleophilic cysteine residue.126, 127 30 1.4.4 Sequence alignment of the ODC domain To identify conserved residues in the ODC domain of ODA, the primary sequence of the ODC component was aligned with other group I PLP-dependent decarboxylases from several bacteria using NCBI BLAST (Figure 1.18).89 Three conserved residues (H119, D199 and K230) were identified. The putative location of these residues in the active site was visualised by constructing a homology model of the ODC/ODA active site through Phyre2 (Figure 1.19).128 All three residues have been shown to play a vital role in catalysis in Lactobacillus sp. 30a H119 aligns with H223 in Lactobacillus sp. 30a.129 In the Lactobacillus sp. 30a crystal structure (PDB entry code 1ORD) H223 is positioned at the N-terminus of an α-helix and its imidazole ring interacts via Ï-Ï stacking with the PLP pyridine ring. The imidazole side chain may act as a proton donor to the Cα during the catalytic cycle.117, 130 ODA D199 corresponds to D316 in Lactobacillus 30a. The pyridinium N on PLP forms a salt-bridge to the D316 carboxylate side chain to help stabilise the quinonoid intermediate.117 Finally, K230 aligns with K355 when compared with Lactobacillus 30a. This residue forms an imine link with PLP to form the internal aldimine.85 Figure 1.18 Sequence alignment performed using NCBI BLAST between the ODC component of F. nucleatum ODA, Lactobacillus sp. 30a ornithine decarboxylase (UniProtKB P43099), Escherichia coli lysine decarboxylase (UniProtKB P0A9H3) and Salmonella enterica arginine F. nucleatum ODC 114 VPRNVHKSV-SA ...... IVH--SYDIPLI ...... TQSTHKILGAM 235 Lactobacillus ODC 218 FDRNNHKSVYNS ...... LCDYISYDSAWV ...... VQSVHKQQAGF 362 E. coli LDC 239 IDRNCHKSL-AH ...... --Q--TLDVPSI ...... TQSTHKlLAAL 372 S. enterica ADC 250 LDRNCHKSIEQ ...... LA--KTSDRIHF ...... THSTHKLLNAL 392 31 decarboxylase (UniProtKB A0A0T9WS57). In red are the three conserved residues: H119, D199 and K230. Figure 1.19 Homology model of the predicted active site of the ODC component of ODA based upon the crystal structure of Lactobacillus sp. 30a (PDB entry code 1ORD) using Phyre2.128 From this it was possible to identify three residues of interest: H119, D199 and K230. 1.4.5 The arginase component Arginase (EC 3.5.3.1) catalyses the hydrolysis of L-arginine to urea and L-ornithine, a transformation that is part of the urea cycle.131, 132 Arginases are found throughout the biosphere, in bacteria, yeasts, fungi, plants, and animals. Unlike ODC, bacterial and mammalian arginases have similar structural motifs and likely share a common ancestor.133 32 1.4.6 Catalytic mechanism of arginase In all arginases that have been characterised to date, including both eukaryotic and prokaryotic variants, two Mn2+ cations (labelled Mn2+A and Mn2+B) are required for activity.134 These Mn2+ cations help orientate a metal-bridging hydroxide ion which acts as the nucleophile in the hydrolysis reaction.135 The first step of the catalytic cycle involves nucleophilic attack of the hydroxide ion at the guanidinium carbon of L-arginine (Figure 1.20). This results in the formation of a tetrahedral intermediate. This intermediate collapses upon proton transfer to the ε-amino group of L-ornithine, which is mediated by D124 (human arginase numbering). Proton shuffling from the bulk solvent to the ε-amino group of ornithine releases the product.134 Finally, a water molecule displaces urea coordinated to the Mn2+ cations, a catalytic step that regenerates the nucleophilic hydroxide ion. Figure 1.20 Catalytic mechanism for the hydrolysis of L-arginine to urea and L-ornithine by arginase.135 33 While Mn2+ is the physiological activator of arginase, it has been reported that other divalent metal cations, including Co2+, Ni2+and Fe2+, can also participate in activation of the enzyme.136, 137 In human type II arginase, the replacement of Mn2+ with Co2+ led to a 2-fold increase in activity, while replacement with Ni2+ resulted in a 25% increase in activity.138 In contrast, replacement of Mn2+ with Fe2+ caused a 10% decrease in activity. Replacement of Mn2+ with other divalent metals (Zn2+, Mg2+, Cd2+ and Cu2+) all resulted in inactivity of the enzyme. 1.4.7 Inhibition of arginase Structural and mechanistic similarity between bacterial and human arginases have made it hard to design selective inhibitors to combat bacterial infections. However, arginase inhibitors have been used or proposed as treatments of a wide range of diseases including cancers, cardiovascular dysfunction in sickle cell, diabetic nephropathy, atherosclerosis and others.139-142 There are several known inhibitors of arginase.143 One of the most potent groups of arginase inhibitors are boronic acid derivatives of L-arginine such as S-(2-boronoethyl)-L-cysteine (BEC) and 2(S)-amino-6-boronohexanoic acid (ABH) (Figure 1.21).144 These act as competitive inhibitors and function by forming a boronate ion during the nucleophilic attack of the metal-bridging hydroxide at the guanidinium carbon.145 These structures mimic the tetrahedral intermediate preventing the release of product. Another potent group of arginase inhibitors are hydroxyguanidine derivatives such as Nâhydroxyânorâarginine (nor-NOHA).146 These too act as competitive inhibitors, with the N-OH moiety of the inhibitor displacing the metal-bridging hydroxide ion. 34 Figure 1.21 Structures of the arginase inhibitors ABH and BEC. 1.4.8 Sequence alignment of the arginase domain Sequence alignment of the arginase component of ODA with microbial and mammalian arginases revealed the substitution of two conserved active site residues in the ODA primary sequence (Figures 1.22). These are residues S609 and N713. The conserved residues in these positions are aspartic acids; they form part of the first coordination shell for the Mn2+ cations.134 The distinct arginase/ODA active is visualised by overlaying a homology model of the enzyme (generated with Phyre2 using human arginase as a template) with that of human arginase (Figure 1.23). The presence of S609 in the place of D124 (human arginase model) is of particular note, as this residue was shown in the human enzyme to be critical for turnover.147 Figure 1.22 Sequence alignment performed using NCBI BLAST of the arginase component of F. nucleatum ODA with arginase from Homo sapiens (UniProtKB P05089), Heliobacter pylori (UniProtKB A3R4G1), Plasmodium falciparum (UniProtKB Q8I384) and Leishmania mexicana (UniProtKB Q6TUJ5). Conserved residues that form the first coordination shell of the enzyme are highlighted in red. F. nucleatum 582 GGDHSISLGSVSG-VSLEKE--IGVLWISAHGDM ...... SIDMNVFD 716 H. sapiens 94 GGDHSLAIGSISGHARVHPD--LGVIWVDAHTDI ...... SFDVDGLD 230 H. pylori 87 SSEHANMFGIFQAFRSVHKDKKIGILYLDAHADI ...... SLDLDIMD 240 P. falciparum 192 GGDHGVAFSSILSSLQMYQN--LRVIWIDAHGDI ...... SLDIDSVD 331 L. mexicana 112 GGDHSIALGTVAGVLSVHPD--AGVIWVDAHADI ...... SYDVDTID 249 35 Figure 1.23 Crystal structure of the active site of human arginase I (blue; PDB entry 2PHA) overlaid with a homology model of the arginase component of ODA (yellow) generated using Phyre2. 1.5 Research Objectives The objective of this research is to perform structural and functional analysis of three proteins from F. nucleatum that may serve as potential drug targets to treat diseases associated with the organism, such as colorectal cancer. Chapter 2 of this thesis focuses on the characterisation of wild-type flavodoxin from F. nucleatum, which appears to be structurally distinct from other bacterial flavodoxins. Stopped-flow spectroscopy, fluorescence titration experiments and redox potentiometry were performed to determine the reduction potentials of the flavin cofactors and binding affinity of the various redox states of the FMN cofactor for the apoprotein. The structural basis for distinct FMN binding affinities was revealed by solving the crystal structure of the 36 enzyme in collaboration with Michael Murphy at UBC Vancouver. Chapter 3 looks at the effect of naturally occurring amino acid substitutions that appear in FnFld on the reduction potentials and binding affinities of the FMN cofactor. Chapter 4 focuses on the initial characterisation of lanthionine synthase. A variety of methods (steady-state kinetic analysis, stopped-flow spectroscopy and HPLC) were used to reveal the scope of reactions catalysed by LS, the nature of the rate determining step and the identification of catalytic intermediates, including their rates of formation and decay. The molecular structures of the enzyme were used to rationalise the unique catalytic properties of the enzyme. Finally, Chapter 5 focuses on the characterisation of ODA. Size exclusion chromatography was used to determine the quaternary structure of ODA and the isolated ODC and arginase components. The presence of Mn2+ in the arginase domain was determined by ICP-MS. Site-directed mutagenesis and steady-state kinetic analysis were used to evaluate the role of the conserved residues in the ODC active site and unusual substitutions within the arginase active site. These experiments helped reveal the mechanism of L-arginine turnover facilitated by ODA. 37 2. Flavodoxin, Part 1 2.1 Chapter Summary Flavodoxin from F. nucleatum (FnFld) is an electron transfer protein that utilises a non-covalently bound flavin mononucleotide (FMN) to shuttle single electrons to a variety of redox partner proteins. The protein family contains a number of conserved residues that play an important role in tuning the FMN reduction potentials by modulating the binding affinity for different redox forms of the flavin cofactor. Notably, natural substitutions of some of these conserved residues appear in the primary sequence of flavodoxin from F. nucleatum (FnFld). For example, an invariant glycine residue in the conserved phosphate binding loop is replaced by a lysine (K13). The conserved tryptophan and tyrosine residues which sandwich the FMN isoalloxazine ring are replaced by Y56 and F93 respectively in FnFld. Finally, an aspartic acid residue downstream of the 90âs loop is replaced by N89. Given that these interactions likely impact FMN binding, we sought to determine the FMN reduction potentials, the binding affinity of FMN to the apo-protein and the effect of ionic strength on FMN binding affinity. We also determined the crystal structure of FnFld in holo- and apo-form in collaboration with the Michael Murphy Lab at UBC Vancouver to provide structural insight into these naturally occurring substitutions. Binding studies performed by stopped-flow spectrophotometry and fluorescence titration experiments established that the binding affinity of FMN to apo-FnFld was lower compared to other related flavodoxins. In addition, the FMN binding affinity increased with an increase in ionic strength. Weaker binding at higher salt concentrations may be attributed to a large conformational shift of the 50âs and 90âs binding loops. This shift rotates the 90âs binding loop ~90o following 38 FMN binding. This brings the two loops closer together and enables the aromatic rings of Y56 and F93 to sandwich the FMN isoalloxazine ring. Consequently, the hydrogen bonds between the ribityl phosphate and the phosphate loop play a more important role in FMN binding. Redox potentiometry revealed that the Eox/sq (-43 mV) and Esq/hq (-256 mV) in FnFld were significantly more electropositive compared to other long-chain flavodoxins. The electropositive Eox/sq is likely attributed to the absence of a hydrogen bond between the FMN N5 and the peptide backbone in the FnFld-FMNox complex. The electropositive Esq/hq is attributed to the near absence of negatively charged acidic residues within 12 Ã of the FMN cofactor that would otherwise destabilise the binding of the negatively charged FMNhq. 2.2 Materials and Methods 2.2.1 Materials. Q5® High Fidelity DNA polymerase and restriction enzymes were from New England Biolabs (Ipswich, MA, USA). Oligonucleotides were purchased from Integrated DNA Technologies (Coralville, IA, USA). F. nucleatum genomic DNA was from the American Tissue Culture Collection (Manassas, VA, USA). XL1-Blue Competent cells were from Agilent, (Santa Clara, CA, USA). The DCâ¢ Protein Assay was from Bio-Rad (Hercules, CA, USA) and the E.Z.N.A plasmid mini kit was from VWR (Radnor, PA, USA). 2.2.2 Cloning petFld into pET41a plasmid The gene coding for FnFld (Fn0724) was amplified from the genomic DNA of F. nucleatum strain ATCC 25586 with Q5® High Fidelity DNA polymerase using primers 5â²âGGA ATT CCA TAT GAA GAC TAT TGG TAT TTT TTA TGCâ3â² (forward primer) and 5â²âCGG GAT CCT TAT TTG AAT TCT TTT TTT ATT TCâ3â² (reverse primer) that included NdeI and BamHI restriction sites (shown in bold).111 The amplified PCR product was inserted into the NdeI 39 and BamHI sites of the pET41a vector. The PCR reaction mixture contained Q5® High Fidelity DNA polymerase (0.02 U/μL), Q5® Reaction Buffer, 0.2 mM of each dNTP, 1.0 ng/mL F. nucleatum genomic DNA, 2.5 ng/mL of forward primer and 2.5 ng/mL of reverse primer in a total volume of 50 µL. The gene was amplified by cycling the reaction mixture 30 time through the following temperatures: 98 oC (10 s), 55 oC (30 s) and 72 oC (45 s) following incubation at 98 oC for 2 minutes. The PCR product, cleaved with NdeI and BamH1, was ligated into a pET41a plasmid digested with the same restriction enzymes. The ligated product was transformed into the XL1-Blue competent cells, and the cell mixture was plated onto LB-agar plates supplemented with ampicillin (100 µg/mL). Plasmid from a transformed cell was isolated using an E.Z.N.A plasmid mini kit.148 DNA sequencing by the Sequencing and Bioinformatics Consortium at UBC Vancouver confirmed the presence of the correct FnFld coding sequence in the pET41a plasmid. 2.2.3 Purification and expression of wild type FnFld Plasmid harbouring the cDNA for FnFld (named petFld) was transformed in E. coli strain Rosetta(DE3)pLysS for protein expression. A starter culture was prepared by inoculating 200 mL LB media, supplemented with ampicillin (100 µg/mL) and chloramphenicol (34 µg/mL), with a single colony generated from a transformed cell. This was incubated for approximately 16 hours at 37 oC. Aliquots (10 mL) of the starter culture were added to 500 mL of Terrific Broth (TB) containing ampicillin (100 µg/mL) and chloramphenicol (34 µg/mL). The culture was incubated at 30 oC at 200 rpm until an OD at 600 nm of ~0.6. Isopropyl β-D-1-thiogalactopyranoside (ITPG) was added to the cell cultures to a final concentration of 0.2 mM. The cultures continued to incubate for approximately 16 hours at 25 oC at 200 rpm. Cell cultures were centrifuged at 6371g for 15 min, and cell pellets were pooled together and stored at -80 oC until protein isolation. 40 The protein was purified following a published protocol.149 Cells (30 g) were resuspended in 200 mL of 50 mM Tris-HCl, pH 7.5 containing the protease inhibitors phenylmethylsulfonyl fluoride (1.0 mM) and benzamidine (0.1 mM). The cell suspension was sonicated on ice (32 % power for 8 s pulse at 1 min intervals) for 1 hour using a Qsonica Q500 sonicator. The suspension was centrifuged at 39191g for 60 min at 4 oC. Ammonium sulfate was gradually added to the pooled supernatant to a final concentration of 65% (w/v) and the solution was centrifuged at 39191g for 60 min at 4 oC. The supernatant was then loaded onto a diethylaminoethyl (DEAE) Sephacel column (50 â¨ 60 mm) equilibrated with 50âmM Tris-HCl pH 8.0 with 65% ammonium sulfate (Buffer A). The column was washed with two column volumes (CV) of Buffer A. FnFld was eluted by applying a linear gradient from 100 % Buffer A to 100 % 50âmM Tris-HCl, pH 8.0 (Buffer B) over four column volumes. Eluted protein was pooled and dialysed against a 4 L solution of 50 mM Tris-HCl, pH 7.5 for 16 hours. The dialysate was concentrated to ~10 mL using a Vivaspin centricon with a 30 kDa MW cut-off filter. The protein was then applied to a Q-sepharose HP column (26 â¨ 140 mm) equilibrated with 50 mM Tris-HCl, pH 7.5. Separate fractions of the holo and apo-protein were eluted from the Q-sepharose column using a linear gradient from 100 % Buffer C (50 mM Tris-HCl, pH 7.5) to 100 % Buffer D (50 mM Tris-HCl, pH 7.5, 1 M NaCl) over 8 column volumes. Fractions containing both forms of the protein were pooled separately and concentrated to 10 â 15 mL. SDS-PAGE was performed to assess purity of the protein. Homogenous stocks of the purified protein were then stored at -80 oC. DCâ¢ Protein Assay (based on the Lowry assay) was used to determine the concentration of both the apo- and holo-forms of the protein.150 For subsequent batches of purified protein, the protein concentration was determined using an extinction coefficient (ε280 nm) for holo-Fld (41.6 mM-1 cm-1) and apo-Fld (18.0 mM-1 cm-1). 41 2.2.4 Fluorescence titrations The equilibrium dissociation constant (Kd) for the apo-Fld-FMNox complex was determined through fluorescence titrations following a protocol published by Lostao et al (2000).61 Unbound FMN fluoresces with an emission maxima at 525 nm following excitation at 446 nm. Binding of the FMN to the apo-protein quenches the flavin fluorescence. All experiments were performed in 10 mM HEPES-NaOH, pH 7.5 at 25 oC. A 3 mL solution of 0.1 µM of FMN was placed in a fluorescence cuvette and inserted in LS50âB fluorescence spectrophotometer (Perkin-Elmer, MA, USA). Aliquots (1 µL) of apo-FnFld (150 µM) were added to the cuvette until a final protein concentration of 1 µM. Quenching of the flavin fluorescence was followed at 525ânm following excitation at 446ânm. The Kd was determined by fitting the change in fluorescence to a quadratic binding isotherm (Equation 2.1). Equation 2.1 ð¥ð¹ = (ð¥ð¹ððð¥2ð0)(ð0 + ð¿0 + ð¾ð â [(ð0 + ð¿0 + ð¾ð)2 â 4ð0ð¿0]12) where ÎF is the change in fluorescence emission, ÎFmax is the maximum change in fluorescence emission, Po is the total protein concentration, Lo is the total FMN concentration, and Kd is the equilibrium dissociation constant for the apo-protein-FMN complex. The titrations were performed at different ionic strengths (20 mM, 50 mM, 100 mM, 200 mM and 500 mM) by adding increasing concentration of NaCl to the titration buffer. 2.2.5 Stopped Flow Spectrophotometry Binding of FMN to apo-FnFld was monitored using a SFâ61DX2 stoppedâflow spectrophotometer (TgK Scientific, Bradford-on-Avon, UK) following a protocol detailed in 42 Lostao et al (2003).151 All experiments were performed in 10 mM HEPES-NaOH, pH 7.5 at 25 oC. A 1 µM solution of apo-FnFld was rapidly mixed in the stopped-flow apparatus with an excess of FMN (7.5-50 µM) in order to maintain pseudo-first order conditions. The excitation wavelength was set at 446 nm. The emitting light from the flavin was monitored by placing a 495 nm cut-off filter in the photomultiplier orthogonal to the incoming light beam. The pseudoâfirstâorder rate constant (kobs) was obtained by fitting an average of five traces to a standard single exponential equation. These kobs values were then plotted against the FMN concentration to determine kon. Stopped flow spectrophotometry was performed at different ionic strengths (20 mM, 50 mM, 100 mM, 200 mM and 500 mM) through the addition of NaCl to the buffer. Stopped flow spectrophotometry was also performed using riboflavin in the place of FMN, with the other reaction conditions kept the same. 2.2.6 Redox Potentiometry The reduction potentials of FnFld were determined through a series of redox titrations performed in a glove box (Belle Technology, Weymouth, UK) maintained under nitrogen atmosphere at 25 oC (O2 < 5 ppm). A titration buffer of 50 mM sodium phosphate pH 7.0 was made anaerobic by bubbling with N2 gas for 2 hours. The buffer was then allowed to equilibrate in the glove box for 16 hrs. A concentrated solution of holo-Fld was introduced to the glove box and gel filtered over a 10 mL size-exclusion column (EconoâPac 10 DG column, Bio-Rad, Hercules, CA, USA)) equilibrated with the anaerobic phosphate buffer. Holo-FnFld was diluted with the titration buffer to produce a 7 mL solution of ~50 µM FnFld. Three redox mediators, benzyl viologen (1 μM), phenazine methosulfate (1 μM), and 2âhydroxyâ1â4ânapthoquinone (1 μM) were added to the FnFld solution. The protein solution was gradually reduced with the addition of 1-10 µL aliquots of sodium dithionite. Following each addition of sodium dithionite the protein solution 43 was allowed to equilibrate for 5 mins and the potential was measured using a pH/ORP meter (Orion 3âStar Benchtop Meter, Thermo Scientificâ¢) equipped with an Ag/AgCl ORP electrode (Orion 9179 BNMD, Thermo Scientificâ¢, MA, USA) The full flavin absorbance spectrum was recorded with a Lambda 265 UV/visible spectrophotometer (Perkin Elmer, MA, USA) at each recorded potential. The volume of sodium dithionite added to the protein solution was varied in order to obtain a change of 20 â 40 mV between each data point. The experiment was stopped after there were three consecutive dithionite additions that produced no further absorbance change at 580 nm. The observed potentials were normalised to a standard hydrogen electrode by the addition of 204 mV to the recorded values. The observed absorbance values at 580 nm were plotted against the normalised potentials. The data were then fitted to a 2-electron process (Equation 2.2) derived from the extension of the BeerâLambert Law and the Nernst equation.152, 153 Equation 2.2 ð´ = ð10(ð¸âð¸1â²)/59 + ð + ð10(ð¸2â² âð¸)/591 + 10(ð¸âð¸1â²)/59 + ð + ð10(ð¸2â²âð¸)/59 where A is the total absorbance at 580ânm, E is the observed normalised potential and E1â² and E2â² are the FMNox/sq and FMNsq/hq, respectively. The component absorbance values contributed by the flavin in the oxidised, semiquinone, and hydroquinone states are represented by a, b, and c, respectively. The free energies (ÎG) associated with binding of the semiquinone (ÎGsq) and hydroquinone (ÎGhq) forms of FMN to apo-FnFld were determined by linking the redox potential of free FMN with the redox potentials of FnFld through a thermodynamic loop (Figure 2.1). This was performed using equations 2.3 and 2.4. 44 Equation 2.3 âð®ðð = âð®ðð â ð­(ð¬ðð/ðð â ð¬ðð/ðððððð) Equation 2.4 âð®ðð = âð®ðð â ð­(ð¬ðð/ðð + ð¬ðð/ðð â ð¬ðð/ððððððâ ð¬ðð/ðððððð) where F is the Faraday constant, Eox/sqfree and Esq/hqfree are the midpoint potentials of FMN in solution. Figure 2.1 A thermodynamic loop that connects the reduction potentials of FMN in solution (Eox/sqfree and Esq/hqfree) with the reduction potentials of the FnFld-FMN complex (Eox/sq and Esq/hq) and the free energies (ÎG) associated with binding FMN in the three different oxidised states. 2.2.7 FnFld crystallisation and structure determination Crystal structures of the apo- and holo-form of FnFld of were determined in collaboration with Michael Murphy lab in the Department of Microbiology and Immunology at UBC Vancouver. Crystals of wild-type Fld were obtained using a sitting-drop vapour diffusion method 45 in 96-well trays at 4 oC with a mixture containing 1 µL of reservoir solution (42%â46% PEG 600, 0.16â0.18 M calcium acetate, and 0.1 M sodium cacodylate) and 1 µL apo-FnFld or holo-FnFld (21 mg/mL). The crystals were soaked in a reservoir solution supplemented with 20% glycerol for 30 s as a cryoprotectant. The crystals were immersed in a stream of liquid nitrogen during acquisition of diffraction data. Diffraction data for wild-type Fld were collected on a Rigaku MicroMax 007-HF generator, VariMax HR optics, and Saturn CCD 944+ detector and data processed using MOSFLM.154 The phase problem was solved for the FnFld by molecular replacement using the structure of flavodoxin from Helicobacter pylori (PDB entry code 1FUE) using the AutoSol program from PHENIX.155 The model was built and refined using PHENIX and visualised with COOT.156 2.3 Results 2.3.1 Properties of purified FnFld Fld was recombinantly expressed and purified by ammonium sulfate precipitation, followed by DEAE-sephacel and Q-sepharose anionic exchange chromatography. The last chromatographic step separated both the holo and apo forms of flavodoxin. SDS-PAGE confirmed that the holo and apo-protein were homogenous with an apparent molecular weight of 19 kDa similar to the expected molecular weight of 18,829 Da (Figure 2.2). Holo-FnFld has a typical flavin absorbance spectrum with absorbance maxima at 380 and 450 nm (Figure 2.3). 46 Figure 2.2 SDS 14% polyacrylamide gel showing both apo and holo-flavodoxin purified by ammonium sulfate precipitation and anion exchange chromatography. Lane 1, protein molecular weight marker; Lane 2, purified holo-flavodoxin; Lane 3 purified apo-flavodoxin. Figure 2.3 UV/visible absorbance spectrum of purified holo-FnFld showing the absorbance peaks at 380 and 450 nm. Lane 1 Lane 2 Lane 3 47 2.3.2 FMN binding to apo-flavodoxin Dissociation constants (Kd) for the FMN binding to apo-FnFld was determined through a fluorescence titration experiment. The change in FMN fluorescence emission was plotted against the total amount of apo-FnFld added to the solution. The data were then fitted to a quadratic binding isotherm (Figure 2.4, Equation 2.1). The Kd for FMN binding to apo-Fld was 34 ± 6 nM at an IS of 2 mM (Table 2.1). There was no significant change in binding affinity at an IS > 100 mM; however, as the IS was increased to >200 mM, there was a gradual increase in FMNox binding affinity. At ionic strength of 500 mM, the Kd was 10 nM (Table 2.1, Figure 2.5). Figure 2.4 Change in fluorescence emission of FMN following the addition of apo-Fld. The titration was performed in 10 mM HEPES-NaOH, pH 7.5 at 25 oC with an IS 2 mM. To determine the equilibrium dissociation constant, data points were fitted to a quadratic binding isotherm (Equation 2.1). 48 Figure 2.5 General decrease in Kd for the FnFld-FMNox complex. The titrations were performed in 10 mM HEPES-NaOH pH 7.5 at 25 oC. The ionic strength was increased by addition of NaCl to the buffer. The association rate constant (kon) was determined by stopped-flow spectrofluorimetry under pseudo first order conditions. For this experiment apo-FnFld was rapidly mixed with a 7.5 â 50-fold molar excess of FMN in the stopped flow. The change in flavin fluorescence was monitored following a 1.6 ms mixing deadtime. The change in emission was monophasic and consequently was fitted to a single exponential equation to give a pseudo first order rate constant (kobs) (Figure 2.6A). The value of kon was determined by plotting kobs against the concentration of FMN (7.5 µM to 50 µM) (Figure 2.6B). The kon for FMN binding to apo-FnFld was 21 ± 1 mM-1 49 s-1 in 10 mM HEPES-NaOH pH 7.0, at 25 oC IS of 2 mM (Table 2.1). An increase in IS from 2 to 500 mM resulted in modest changes in the association rate constant (Table 2.1, Figure 2.7). The dissociation rate constant (koff) was determined through equation 2.5. For FMN binding to apo-FnFld in 10 mM HEPES-NaOH, pH 7.0 at 25 oC the koff was 7.2 â¨ 10-4 s-1 at an IS of 2 mM (Table 2.1). There were no significant changes in koff up to an ionic strength of 50 mM. However, there was a 5-fold slower off rate at an ionic strength of 500 mM. Flavin fluorescence was not quenched when riboflavin was rapidly mixed with the apoprotein, indicating that the ribityl phosphate is important for securing the FMN cofactor to the protein Equation 2.5 ð²ð =ððððððð Table 2.1 Association rate constants (kon) determined via stopped flow, equilibrium dissociation constant (Kd) determined via fluorescence titration and calculated dissociation rate constants (koff) for FMN binding to apo-FnFld in 10 mM HEPES-NaOH, pH 7.0 at 25 oC. Ionic Strength (mM) kon (mM-1 s-1) Kd (nM) koff (10-4 s-1) 2 21 ± 1 34 ± 6 7.2 ± 1.3 20 26 ± 1 37 ± 6 9.6 ± 1.7 50 28 ± 1 34 ± 2 9.4 ± 0.9 100 22 ± 1 30 ± 4 6.5 ± 0.9 200 20 ± 1 21 ± 2 4.1 ± 0.7 500 15 ± 1 10 ± 1 1.5 ± 0.2 50 Figure 2.6 A) An average of five fluorescent traces following the binding of an excess of 50 µM FMN to 1 µM apo-Fld. The horizontal line shows the recorded emission after mixing 50 µM FMN with buffer. The pseudo-first order rate constant (kobs) was determined by fitting the averaged trace A B 51 to a single exponential equation. The box below shows a plot of the residues determined when fitting to the single exponential equation. B) Linear regression of recorded kobs plotted against FMN concentration. The slope of this plot was used to determine the association rate constant (kon). The experiment was performed in 10 mM HEPES-NaOH, pH 7.5 at 25 oC. Figure 2.7 Plot of the association constants for FMN binding to apo-Fld against ionic strength. The experiment was performed in 10 mM HEPES-NaOH, pH 7.5 at 25 oC. 2.3.3 Redox potentials The redox potentials of FnFld were determined by titrating the oxidised protein with aliquots of sodium dithionite under an N2 atmosphere (O2 <5 ppm). Following the addition of each aliquot of dithionite the potential of the protein solution was measured and a UV/visible absorbance spectrum was recorded (Figure 2.8). The addition of the first reducing equivalent 52 causes a reduction in the λmax at 450 nm and the appearance of a broad absorbance band with a peak at 580 nm. These initial absorbance changes correspond to conversion of the oxidised FMN to the blue neutral semiquinone (FMNsq). The addition of the second reducing equivalent causes a further decrease in the absorbance maxima at 450 and 580 nm as FMNsq is converted to FMNhq. The change in absorption at 580 nm was plotted against the reduction potential normalised to the hydrogen electrode (Figure 2.9). The data were then fitted to equation 2.2 to determine Eox/sq and Esq/hq. These values were used with equations 2.3 and 2.4 to find the free energies associated with the binding of FMNsq (ÎGsq) and FMNhq (ÎGhq) to the apo-protein. FnFld was found to have an Eox/sq of -43 mV and Esq/hq of -256 mV. ÎGox, ÎGsq and ÎGhq were calculated to be -43, -61 and -53 kJ/mol respectively. These free energy values for apo-FnFld-FMNsq and apo-FnFld-FMNhq complexes correspond to Kd values of 0.17 nM and 0.42 nM. Figure 2.8 Overlaid absorption spectra of holo-Fld acquired during chemical reduction of the flavin. Solid lines denote spectra associated with FMNox to FMNsq transition and dotted lines represent the transition from FMNsq to FMNhq. 53 Figure 2.9 Plot of the absorbance at 580 nm of holo-Fld against the reduction potential normalised to the standard hydrogen electrode. The data were fitted to equation 2.2 to determine for Eox/sq and Esq/hq. 2.3.4 Three-dimensional structure The crystal structure of wild-type FnFld (PDB entry code 6OHL) was solved to 1.85 Ã resolution using molecular replacement (Figure 2.10 and 2.11). The data collection and refinement statistics are presented in Table 2.2. 54 Table 2.2 Data collection and refinement statistics for wild type FnFld. *Values in parentheses are for the highest resolution shell. Data Collection Wavelength (â«) 1.54 Resolution Range (â«) 25.07 â 1.85 Unique Reflections 23667 Data Completeness (%) Completeness (%) 98.3 (76.6)* Multiplicity 9.9 R-meas (%) 9.7 CC1/2 (%) 99.2 (79.9) I/Ï(I) 22.7 (1.7) Space group H32 Unit cell parameters a, b, c 118.9 â« , 118.9 â«, 103.4 â« âº, β, É£ 90°, 90°, 120° Refinement statistics R, Rfree 0.168, 0.191 r.m.s.d. bond lengths (â«) 0.012 r.m.s.d. bond angels (°) 1.02 Average B-factor (â«2) 22.0 Ramachandran plot: outliers ( %) Preferred 157 (98%) Allowed 3 (2%) Outliers 0 (0%) 55 Figure 2.10 A cartoon of the crystal structure of wild type FnFld (PDB entry code 6OHL) determined using a molecular replacement. The FMN cofactor is shown in stick form and loops involved in FMN binding are highlighted in blue. Figure 2.11 A electron density map of the FMN cofactor (grey) bound to wild type Fld with a loop comprised of residues Q57 to G59 (pink). A weighted 2Fo-Fc map contoured at 1 Ï is drawn in blue. The Fo-Fc difference map is contoured at 3 Ï and coloured green and red to indicate positive and negative peaks, respectively. N-terminus C-terminus 56 The overall structure of FnFld closely aligns with the α/β-fold that is prototypical of the flavodoxin family. The protein consists of five parallel β-strands flanked by two α-helices on one side and three α-helices on the other side. The FMN binding site is located at the C-terminal end of the β-sheet and is secured through three loops, the phosphate binding loop (or P-loop), the 50âs loop and 90âs loop. The 5â-ribityl phosphate group on FMN forms O-HO hydrogen bonds to the side chains of Y56, T10, T12, T14 and T15, it also forms NH-O hydrogen bonds with the amide backbone of residues that comprise the phosphate binding loop (L11, T12, T14 and T15) (Figure 2.12). The FMN isoalloxazine ring interacts with the backbone atoms of G59, N89, S96 and C98. Figure 2.12 Close up showing the residues that make up the P-loop and residues G59, N89, S96 and C98 surrounding the FMN cofactor. The FMN cofactor is shown in grey. The ribityl hydroxyl groups interact with the solvent and the side chain residues of T55 and E145 (Figure 2.13). The isoalloxazine ring is secured to the protein through van der Waals interactions with F93 on the si-face and Y56 on the re-face. F93 is nearly parallel to the ring, 57 enabling the formation of extensive Ï-Ï stacking interactions. The phenolic group of Y56 is rotated 30o with respect to the FMN isoalloxazine ring. Figure 2.13 A close-up of Y56, F93 and E145 residues surrounding the FMN cofactor. Y56 and F93 sandwich the FMN cofactor between the two aromatic side chains. E145 is the only negatively charged residue within 12 Ã of the FMN N1. In FnFld, the NH group on V58 is 4.9 Ã from the FMN N5 and N-H-N geometry does not favour formation of a hydrogen bond (Figure 2.14). The FMN N5 instead forms a hydrogen bond with a water molecule interacting with the O4 of the isoalloxazine ring. This O4 forms a hydrogen bond with the backbone amide of G59 and the hydroxyl group on S96. 58 Figure 2.14 Close-up showing the distance between the FMN N5 and the closest residue on the amino acid backbone (V58). Analysis of the electrostatic potential surface of Fld reveals a positive region surrounding the FMN cofactor, with two negative patches above and below the cofactor on the surface of the protein (Figure 2.15). There is a small solvent-filled basic cleft that comprises the side chain of K13, K37, N34 and H62. The structure of apo-FnFld (unpublished structure) is mostly the same as holo-FnFld. There is no significant change is the position of any of the α-helices or β-sheets. However, there is a notable conformational shift in the positions of the 50âs and 90âs binding loops (Figure 2.16). Much of the 50âs binding loop does not show up in the crystal structure of the apo-FnFld, this suggests that the region does not maintain a constant electron density and is much more flexible before FMN binding. In apo-FnFld the 90âs binding loop is twisted ~90o away from its position in holo-FnFld and the aromatic group on F93 sticks out into the solvent. 59 Figure 2.15 Charge distribution map of the crystal structure of FnFld showing the change in charged surface from -5 kT/e (red) to +5 kT/e (blue). The FMN cofactor is shown in stick form in grey. Figure 2.16 Overlaid crystal structures of holo-FnFld (pink) and apo-FnFld (blue). The FMN cofactor from holo-FnFld is shown in grey. 60 2.4 Discussion 2.4.1 Analysis of the FnFld crystal structure The tertiary structure of FnFld conforms to the typical α/β-fold of the flavodoxin family. The long-chain flavodoxin from Nostoc sp PCC 7120 (Anabaena, PDB entry code 1FLV) shows the highest structural similarity with FnFld with an R.M.S deviation 1.3 Ã for the of the Câº atoms.48 The conserved phosphate binding loop (T/S-X-T-G-X-T) in flavodoxin is unusual in that it does not contain any positively charged residues to stabilise and neutralise the negatively charged phosphate dianion. The ribityl phosphate is instead secured to the protein via a series of NH-O and OH-O hydrogen bonds provided by the backbone NH atoms and threonine side chains. While this is a-typical of the majority of phosphate binding loops found in nature, the P-loop is considered a fingerprint motif for members of the flavodoxin family.60 Despite the unusual presence of lysine in the place of glycine in this loop (K13), the number of hydrogen bonds between the P-loop and the phosphate in FnFld is the same as in other flavodoxins. 2.4.2 FnFld elicits relatively weak FMN binding FnFld, like some other flavodoxin (such as short chain flavodoxins Fld from C. beijerinckii and M. elsdenii) is unable to bind riboflavin.157, 158 In contrast, flavodoxins from Anabaena Fld, A. vinelandii and D. vulgaris can bind riboflavin, but with 6-orders of magnitude lower affinity compared to FMN.151, 159 In Fld from C. beijerinckii, the 90âs loop is approximately 3-5 residues smaller and the aromatic side chain W90 is rotated ~20o away from the isoalloxazine ring, instead of being nearly co-planar to the flavin as observed in Fld from Anabaena Fld. As a consequence, the pyrimidine ring of FMN is more exposed to solvent in Fld from C. beijerinckii. These structural 61 variations may account for the absence of riboflavin binding to Fld from C. beijerinckii. However, in FnFld, neither of these structural variations are observed; the 90âs loop is approximately the same length as Anabaena Fld and F93 sits planar to the isoalloxazine ring. Therefore, there must be a different explanation for the lack of riboflavin binding to FnFld. Fluorescence titration experiments and stopped-flow spectrophotometry revealed that FnFld elicits a higher Kd for the binding of FMNox to apo-FnFld compared to other studied flavodoxins (Table 2.3). The higher Kd appears to originate primarily from a slower association rate constant. Weaker binding of FMNox to apo-FnFld may be attributed to structural variations present in the 50âs loop, for example the presence of Y56 instead of the conserved tryptophan, which would decrease the van der Waals interactions between the protein and the FMN isoalloxazine ring.158 The importance of these interactions were demonstrated in Fld from Anabaena as substitution of the FMN-stacking W57 to a tyrosine weakened FMN binding.61, 151 Table 2.3 Comparison of the binding association (kon), dissociation rate constants (koff) and equilibrium dissociation constant (Kd) for FMN binding to apo-Fld between different organisms. Organism kon (mM-1 s-1) Kd (nM) koff (s-1) F. nucleatum 21 34 7.2 â¨ 10-4 D. vulgaris160 700 12.5 8.8 â¨ 10-3 M. elsdenii161 260 0.4 1.0 â¨ 10-4 Anabaena151 40 0.2 7.6 â¨ 10-6 P. elsdenii162 320 280 8.8 â¨ 10-2 A. vinelandii163 950 0.09 8.6 â¨ 10-5 62 The absence of a hydrogen bond between the FMN N5 and the peptide backbone in the 50âs loop of FnFld may also structurally account for weaker FMN binding. In long-chain flavodoxins, such as those from Anabaena, E. coli and S. elongates, the FMN N5 forms a hydrogen bond with the NH group of a residue within the 50âs binding loop (Figure 2.17).57, 164 The distance between the two heteroatoms is ~3.5 Ã to 3.7 Ã ; thus, it is considered a weak hydrogen bond interaction.60 In FnFld there is no analogous hydrogen bond, as the distance between the N5 of FMN and the closest hydrogen bond donor (NH on V58) is 4.9 Ã and the N-H-N angle is too far from the ideal of 180o. The hydrogen bond is also absent in short-chain Flds. Figure 2.17 Comparison of the interactions between NH of residue 59 (58 in case of FnFld) and the FMN N5 in four different long chain flavodoxins. FnFld (PDB entry code 6OHL), Fld from Nostoc sp PCC 7120 (Anabaena) (PDB entry code 1FLV), Fld from E. coli (PDB entry code 1AG9), and Fld from S. elongates with FMN in the oxidised and semiquinone forms (PDB entry codes 1OFV and 1CZL respectively). 63 2.4.3 Increasing ionic strength results in decreased binding affinity towards FMN The 3-fold decrease in Kd for the FnFld-FMNox complex at high ionic strength (500 mM) is largely attributed to a 5-fold slower dissociation rate constant. These results contrast with that observed for flavodoxins from Anabaena.151, 165 Ionic strength dependence studies for Fld from Anabaena revealed that FMNox bound more weakly to the apo-protein at high ionic strengths (The Kd was approximately 3-fold higher at an ionic strength of 500 mM compared to an ionic strength of 2 mM. Weaker binding was attributed to a 90-fold increase in koff that was partially compensated for with a 33-fold increase in kon. The authors conclude that the binding of FMNox was driven through Ï-Ï interactions between the isoalloxazine ring and residues Y94 and W57. Higher salt would favour formation of these initial Ï-Ï interactions leading to an increase in kon. Electrostatic interactions between the phosphate and the protein secured the cofactor to the protein. Higher salt would disrupt these interactions leading to a 90-fold increase in koff. The difference in ionic strength dependence on FMN binding to apo-Fld may be linked to the differences in the structural shift between holo- and apo-FnFld (unpublished) compared with holo- and apo-Fld from Anabaena (PDB entry code 1FTG). In Anabaena apo-Fld W57 and Y94 are close together within 5 Ã of each other (Figure 2.18). When FMN binds there is a subtle conformational shift and these two residues are forced apart to >7 Ã to enable the FMN cofactor to sandwich between them. W57 undergoes a ~45o rotation away from the FMN binding pocket and Y94 is pushed back so that it fits planar to the FMN cofactor. In the structure of apo-FnFld, these are much further apart, Y56 is >15 Ã away from F93 (Figure 2.19). As FMN binds, F93 undergoes a ~90o rotation to align planar to the to the isoalloxazine ring and Y56 undergoes a small shift towards the FMN binding pocket. These structural difference in the apo-forms of the two 64 flavodoxins likely account for the differences in FMN binding affinity as a function of ionic strength. Figure 2.18 Conformation change of W57 and Y94 following binding of FMN in Fld from Anabaena. Residues from apo-Fld (PDB entry code 1FTG) are in blue and residues from holo-Fld (PDB entry code 1FLV) are in pink. The FMN cofactor from holo-Fld is shown in grey. Figure 2.19 Conformation change undergone by residues Y56 and F93 following binding of FMN in FnFld. Residues from apo-FnFld are shown in blue and residues from holo-FnFld are shown in pink. The FMN cofactor from holo-FnFld is shown in grey. 65 2.4.4 Perturbation of the redox potentials Table 2.4 shows that Eox/sq for FnFld is more electropositive compared to other studied long-chain flavodoxins, but it is comparable to short-chain Flds from C. beijerinkii and D. desulfuricans.164, 166 Crystallographic studies of Fld in the semiquinone state from S. elongatus reveal that reduction of FMN to the semiquinone induces a peptide flip within the 50âs loop (Figure 2.20).57 Reduction of FMNox to FMNsq leads to protonation of the FMN N5, converting this atom from a hydrogen bond acceptor to a hydrogen bond donor. To neutralise the partial positive hydrogen bond on this atom in FMNsq, the peptide backbone of N58 and V59 in the 50âs loop rotates.57 Following this peptide flip, the distance between N58 and the FMN N5 is shortened from 3.5 Ã to 2.8 Ã , increasing the strength of the hydrogen bond formed. In long-chain flavodoxins, this now stronger hydrogen bond secures the FMNsq to the protein. Short-chain Flds also undergo a peptide flip to form a hydrogen bond with FMN N5(H). However, like FnFld they do not form hydrogen bond interactions with FMN N5 in the Fld-FMNox state. Thus, the difference in binding energy between the binding of the FMNox and FMNsq (âGox - âGsq) is larger in short-chain flavodoxins (10 â 20 kJ/mol) due to the formation of the new hydrogen bond between cofactor and protein. In long-chain flavodoxins the corresponding energy difference is much smaller (1 â 3 kJ/mol) because the transition substitutes a weak hydrogen bond for a stronger hydrogen bond. Consequently, short-chain Flds and FnFld have a more electropositive Eox/sq. 66 Figure 2.20 Peptide flip of N58 and V59 observed in Fld from S. elongatus as FMN (grey) is reduced from an oxidised form (pink) (PDB entry code 1CZU) to semiquinone form (blue) (PDB entry code 1CZL). Table 2.4 Comparison of the redox potentials and free energies associated with FMN binding in different redox states to both long-chain and short-chain apo-flavodoxin from different organisms. Organism Eox/sq (mV) Esq/hq (mV) ÎGox (kJ/mol) ÎGsq (kJ/mol) ÎGhq (kJ/mol) F. nucleatum -43 -256 -43 -61 -53 S. elongates*57 -221 -442 -51 -52 -26 Anabaena*167 -207 -439 -51 -54 -28 C. beijerinkiiâ¦164 -92 -399 -44 -58 -36 D. desulfuricansâ¦166 -58 -387 -36 -53 -32 D. vulgarisâ¦168 -143 -440 -55 -64 -38 E. coli (FldA)*49 -260 -452 -51 -49 -22 FMN free56 -238 -172 - - - * = Long chain apo-flavodoxin â¦ = Short-chain apo-flavodoxin 67 The Esq/hq for FnFld is also more electropositive than any of the other studied short-chain or long-chain flavodoxin (Table 2.4). The higher Esq/hq is attributed to unusually tight binding of the negatively charged FMNhq. The electrostatic potential surface of most flavodoxins contain a concentration of negatively charged amino acids surrounding the FMN cofactor (Figure 2.21).58 These negative charges in combination with the hydrophobic residues act to destabilise the binding of FMNhq anion, which lowers Esq/hq. Figure 2.21 Structures of various flavodoxins with known reduction potentials coloured by electrostatic potential from -5 kT/e (red) to +5 kT/e (blue). The PDB entry codes are shown in parentheses below the structures, FMN cofactors are shown in stick form in green. 68 In contrast, the electrostatic potential surface surrounding FnFld is not negatively charged. Tighter binding of FMNhq may be attributed to the near absence of negatively charged residues in close proximity to the FMN N1, the locus of the negative charge of the FMNhq anion. Anabaena Fld contains five negatively charge carboxyl group <12 Ã from the FMN N1, whereas FnFld only contains one residue (E145) (Figure 2.13).48 A second contributing factor may be an increase in solvent accessibility around the FMN, which was calculated to 127 Ã 2 for FnFld. This is higher than the solvent accessible surface areas determined for flavodoxin from Anabaena (111 Ã 2) (PDB entry code 1FL), S. elongatus (96 Ã 2) (PDB entry code 1CXK) and E. coli (101 Ã 2) (PDB entry code 1AHN). 2.5 Concluding remarks The binding affinity of FMN to apo-FnFld was found to be reduced compared with other related flavodoxins. The FMN binding affinity increased as the ionic strength was increased. These phenomena may be attributed to a significant conformation shift of the F93 residue that takes place during FMN binding. These results suggest that FMN binding is initially driven via binding of the ribityl phosphate to the P-loop. Redox potentiometry revealed that FnFld had a more electropositive Eox/sq and Esq/hq in comparison with other long-chain flavodoxins. The shift in Eox/sq may be caused by the absence of a hydrogen bond between FMN N5 and the 50âs loop. This was confirmed by the crystal structure which revealed that the distance between FMN N5 and the closest hydrogen bond donor (NH on V58) was 4.9 Ã , too far apart to form a hydrogen bond. The shift in Esq/hq may be attributed to unusually tight binding of the negatively charged FMNhq. This tighter binding is likely caused by a near absence of destabilising negatively charged residues near the FMN N1. 69 Several unusual amino acid substitutions have been identified in important conserved regions in the protein sequence. The impact of these naturally occurring substitutions will be the focus of the next chapter of this thesis. 70 3. Flavodoxin, Part 2 3.1 Chapter Summary Four unusual substitutions in the amino acid sequence of FnFld were found via sequence alignment with other members of the flavodoxin family. The first is the substitution of a normally invariant glycine present in the phosphate bind loop with a lysine residue (K13). Y56 and F93, which sandwich the FMN cofactor in FnFld are normally a tryptophan and tyrosine, respectively. Finally, a highly conserved aspartic acid residue in the 90âs binding loop shown to modulate the Esq/hq is N89 in FnFld. The impact of these naturally occurring substitutions on the biophysical and thermodynamic properties of FnFld was investigated through site directed mutagenesis. The mutations (K13G, Y56W, N89D and F93Y) reverted each naturally occurring substitution in FnFld to the conserved residue. The effects of each of these mutations on FMN binding and the redox properties of the protein were determined. In addition, the crystal structure of K13G was solved in collaboration with the Michael Murphy Lab at UBC Vancouver to provide additional structural insight. K13G was found to have a 4-fold increase in FMNox binding affinity and a modest increase in Eox/sq compared to wild-type FnFld. Y56W caused a 20-fold decrease in binding affinity, which was entirely attributed to an increased koff. N89D had a more electronegative Esq/hq value and slightly increased FMNox binding affinity. Finally, F93Y had a more electronegative Eox/sq value and slightly increased FMN binding affinity. The structural rationale for perturbations in the FMN binding affinity and reduction potentials induced by each of these substitutions are detailed in the Discussion. 71 3.2 Materials and Methods 3.2.1 Materials Oligonucleotides were purchased from Integrated DNA Technologies (Coralville, IA, USA). PfuTubo DNA polymerase and XL1-Blue Competent cells were from Agilent, (Santa Clara, CA, USA). The E.Z.N.A plasmid mini kit was from VWR (Radnor, PA, USA). 3.2.2 Generation of the single site substitutions Four variants (K13G, Y56W, N89D and F93Y) of FnFld were produced via site directed mutagenesis using PfuTurbo DNA polymerase (Aligent, CA, USA). 169 The PCR reaction mixture contained PfuTurbo DNA polymerase (0.05 U/μL), PfuTurbo reaction buffer, 0.2 mM of each dNTP, 1.0 ng/mL petFld, 2.5 ng/mL forward primer and 2.5 ng/mL reverse primer in a total volume of 50 µL. The entire plasmid was amplified by cycling the reaction mixture 18 time through the following temperatures: 95 oC (30 s), 55 oC (60 s) and 68 oC (360 s) following incubation at 95 oC for 30 seconds. The list of the primers used for generating each mutant are shown in Table 3.1. Table 3.1 List of primers used in site directed mutagenesis to produce the FnFld variants. Name Sequence Forward K13G 5â²-TTTTATGCAACTCTTACAGGAACAACAGTAGGAATTGTTGAT Reverse K13G 5â²-ATCAACAATTCCTACTGTTGTTCCTGTAAGAGTTGCATAAAA Forward Y56W 5â²-CTTATTTTAGTTACACCCACTTGGCAAGTTGGTGAGGCTCATGCAG Reverse Y56W 5â²-CTGCATGAGCCTCACCAACTTGCCAAGTGGGTGTAACTAAAAT AAG Forward N89D 5â²-GGGCTTGTTGGACTTGGAGATCAATTTGCTTTTGGAGAATCTTTC Reverse N89D 5â²-GAAAGATTCTCCAAAAGCAAACTGATTTCCAAGTCCAACAAGCCC Forward F93Y 5â²-GGACTTGGAAATCAATTTGCTTATGGAGAATCTTTCTGTGGTG Reverse F93Y 5â²-CACCACAGAAAGATTCTCCATAAGCAAATTGATTTCCAAGTCC 72 The PCR product was transformed into the E. coli XL1-Blue competent cells, and the cell mixture was plated onto LB-agar plates supplemented with ampicillin (100 µg/mL). Plasmid from a transformed cell was isolated using an E.Z.N.A plasmid mini kit.148 DNA sequencing by the Sequencing and Bioinformatics Consortium at UBC Vancouver confirmed the presence of the correctly inserted mutations. These variants were expressed and isolated following the same protocol for wild-type FnFld outlined in section 2.2.3. 3.2.3 Determination of the full crystal structure of K13G Crystal structures of the holo-form of K13G was determined in collaboration with the Michael Murphy lab in the Department of Microbiology and Immunology at UBC Vancouver. Crystals of K13G were obtained using a sitting-drop vapour diffusion method in 96-well trays at 4 oC with a mixture containing 1 µL of reservoir solution (21% PEG 3350, 0.23 M MgCl2, and 0.1 M HEPES-NaOH pH 7.5) and 1 µL K13G (8.3 mg/mL). The crystals were soaked in a reservoir solution supplemented with 15% glycerol for 30 s as a cryoprotectant. The crystals were immersed in a stream of liquid nitrogen during data collection. Diffraction data for K13G were collected at the Canadian Light Source beamline 08B1-1 and processed using XDS.170 The phase problem was solved for the K13G variant by molecular replacement using the structure of flavodoxin from Helicobacter pylori (PDB entry 1FUE) using the AutoSol program from PHENIX.155 The model was built and refined using PHENIX and visualised with COOT.156 73 3.2.4 Binding and redox analysis with flavodoxin variants The four FnFld variants underwent the same fluorescence titrations, stopped flow kinetic analysis and redox potentiometry as performed for wild-type FnFld, which are outlined in Chapter 2 (sections 2.2.4, 2.2.5 and 2.2.6, respectively). 3.3 Results 3.3.1 Three-dimensional structure The crystal structures of the K13G variant (PDB entry code 6OHK) was solved to 1.20 Ã resolution respectively using molecular replacement (Figure 3.1). The data collection and refinement statistics are presented in Table 3.2. The crystal structure for K13G and wild-type FnFld are nearly superimposable with a R.M.S deviation of 0.14 Ã for the Cα atoms. The FMN 5â-ribityl phosphate forms O-HO hydrogen bonds with Y56, T10, T12, T14 and T15, it also forms NH-O hydrogen bonds within the phosphate binding loop residues L11, T12, T14 and T15. The amide group of K13 (wild-type FnFld) and G13 (K13G) forms a hydrogen bond to the side chain of T10 (OH-N distance of 2.8 Ã ) (Figure 3.2). There are slight differences in the Ψ and Ï angles of the backbone of K13 (-4o and 79o) compared with those of G13 (-11o and 101o). Overall there is no change in the number of hydrogen bonds formed to ribityl phosphate between K13G and wild-type FnFld. 74 Figure 3.1 A electron density map of the FMN cofactor (grey) bound to K13G with a loop comprised of residues Q57 to G59 (pink). A weighted 2Fo-Fc map contoured at 1 Ï is drawn in blue. The Fo-Fc difference map is contoured at 3 Ï and coloured green and red to indicate positive and negative peaks, respectively. Figure 3.2 Overlay of the crystal structures of wild-type FnFld (pink) and K13G (blue) comparing the phosphate binding loop. The FMN cofactors are show in grey (wild-type FnFld) and yellow (K13G). 75 Table 3.2 Data collection and refinement statistics for K13G. *Values in parentheses are for the highest resolution shell. Data Collection Wavelength (â«) 0.980 Resolution Range (â«) 36.44 â 1.20 Unique Reflections 55815 Data Completeness (%) Completeness (%) 99.2 (92.0)* Multiplicity 6.6 R-meas (%) 6.1 CC1/2 (%) 99.9 (84.3) I/Ï(I) 17.1 (2.6) Space group P212121 Unit cell parameters a, b, c 36.09 â« , 67.25 â«, 72.88 â« âº, β, É£ 90°, 90°, 90° Refinement statistics R, Rfree 0.151, 0.164 r.m.s.d. bond lengths (â«) 0.013 r.m.s.d. bond angels (°) 1.20 Average B-factor (â«2) 17.0 Ramachandran plot: outliers ( %) Preferred 163 (98%) Allowed 3 (2%) Outliers 0 (0%) 76 3.3.2 FMN binding to apo-flavodoxin variants Dissociation constants (Kd) for the FMN binding to apo-FnFld was determined through a fluorescence titration experiment. The change in FMN fluorescence emission was plotted against the total amount of apo-FnFld added to the solution. The data were then fitted to a quadratic binding isotherm (Figure 3.2A, Equation 2.1). The Kd for K13G (9.0 ± 0.8 nM), F93Y (9.4 ± 2.3 nM) and N89D (19 ± 1 nM) all showed a slight decrease compared to wild type FnFld (34 ± 6 nM, Table 3.3). Y56W was the only variant to demonstrate an increase in Kd (690 ± 60 nM). The association rate constant (kon) was determined by stopped-flow spectrofluorimetry under pseudo first-order conditions where apo-FnFld was rapidly mixed with a >7.5-fold molar excess of FMN. As for the wild type protein, the monophasic change in fluorescence emission was fitted to a single exponential equation to give a pseudo-first order rate constant (kobs) (Figure 3.3B). The association rate constant (kon) was obtained from a plot of kobs against the FMN concentration. As presented in Table 3.3, there was no significant change in kon for Y56W (23 ± 2 mM-1 s-1) and N89D ( 23 ± 1 mM-1 s-1), while the kon for F93Y (38 ± 2 mM-1 s-1) and K13G (46 ± 1 mM-1 s-1) displayed a modest increase compared to wild type FnFld. There was a minor decrease in koff for K13G (4.0 ± 0.2 Ã 10-4 s-1), F93Y (3.6 ± 0.9 Ã 10-4 s-1) and N89D (4.4 ± 0.2 Ã 10-4 s-1) compared to wild type Fld (7.2 ± 1.3 Ã 10-4 s-1). The Y56W substitution resulted in a ~20-fold increase in koff (157 ± 14 Ã 10-4 s-1). 77 Table 3.3 Association (kon), dissociation rate constants (koff) and equilibrium dissociation constant (Kd) for FMN binding to variants of apo-Fld in 10 mM HEPES-NaOH, pH 7.0 at 25 oC. Variant kon (mM-1 s-1) Kd (nM) koff (10-4 s-1) FnFld 21 ± 1 34 ± 6 7.2 ± 1.3 K13G 46 ± 1 9.0 ± 0.8 4.0 ± 0.2 Y56W 23 ± 2 690 ± 60 157 ± 14 N89D 23 ± 1 19 ± 1 4.4 ± 0.2 F93Y 38 ± 2 9.4 ± 2.3 3.6 ± 0.9 Figure 3.3 A) Change in fluorescence emission of FMN following the addition of apo-forms of wild-type FnFld (squares), K13G (open circles), Y56F (closed triangle), F93Y (closed circle), and N89D (down triangle). The titration was performed in 10 mM HEPES-NaOH, pH 7.5 at 25 oC with an IS 2 mM. To determine the equilibrium dissociation constant, data points were fitted to a quadratic binding isotherm (Equation 2.1). B) Linear regression of recorded kobs plotted against FMN concentration for apo-forms of wild-type FnFld (squares), K13G (open circles), Y56F (closed triangle), F93Y (closed circle), and N89D (down triangle). The slope of this plot was used 78 to determine the association rate constant (kon). The experiment was performed 10 mM HEPES-NaOH, pH 7.5 at 25 oC. 3.3.3 Redox potentials of flavodoxins variants The redox potentials (Eox/sq and Esq/hq) of the Fld variants were determined from the titration of oxidised Fld with aliquots of sodium dithionite under an N2 atmosphere (Figure 3.4, Table 3.4). As with the wild type FnFld, the Eox/ sq and Esq/hq values were obtained by plotting the changes in absorbance at 580 nm against the reduction potential normalised to the standard hydrogen electrode. The greatest change in Eox/sq was observed for K13G (-11 ± 3 mV) and F93Y (-97 ± 2 mV), whereas the largest change in Esq/hq was observed for Y56W (-280 ± 3 mV) and N89D (-301 ± 2 mV). As for wild type FnFld, free energies associated with the binding of FMNsq (ÎGsq) and FMNhq (ÎGsq) were determined from Eox/sq and Esq/hq using equations 2.3 and 2.4 and constructed through the thermodynamic loop (Figure 2.1). The values of ÎG associated with the binding of the three redox states of flavin cofactor to the protein are presented in Table 3.4. The ÎGox recorded for FMNox binding to K13G (-46 kJ/mol) and F93Y (-46 kJ/mol) were more negative than wild type FnFld, signifying stronger binding of FMNox to the apo-protein. In contrast ÎGox associated with Y56W (-38 kJ/mol) was less negative than wild type FnFld due to weaker binding of FMNox to the apo-protein. The ÎGsq determined for FMNsq binding to K13G (-67 kJ/mol) and N89D (-63 kJ/mol) demonstrated tighter binding of FMNsq than observed for wild type FnFld. Whereas, Y56W (-56 kJ/mol) and F93Y (-59 kJ/mol) displayed weaker binding to FMNsq. Finally, there was stronger binding of FMNhq to Y56W (-45 kJ/mol), F93Y (-51 kJ/mol) and N89D (-51 kJ/mol) and weaker binding of FMNhq to K13G (-59 kJ/mol) compared to wild type FnFld. 79 K13G B A K13G C D F56W F56W E F Y93F Y93F 80 Figure 3.4 Overlaid absorption spectra of A) K13G, C) Y56W, E) F93Y and G) N89D acquired during chemical reduction of the flavin cofactor. Solid lines denote spectra associated with FMNox to FMNsq transition and dotted lines represent the transition from FMNsq to FMNhq. Plot of the absorbance at 580 nm of B) K13G, D) Y56W, F) F93Y and H) N89D against the reduction potential normalised to the standard hydrogen electrode. The data were fitted to equation 2.2 to determine for Eox/sq and Esq/hq. Table 3.4 Comparison of the redox potentials and free energies associated with FMN binding in different redox states to the different apo-flavodoxin variants. Variant Eox/sq (mV) Esq/hq (mV) ÎGox (kJ/mol) ÎGsq (kJ/mol) ÎGhq (kJ/mol) FnFld -43 ± 2 -256 ± 3 -43 -61 -53 K13G -11 ± 3 -262 ± 3 -46 -67 -59 Y56W -51 ± 3 -280 ± 3 -38 -56 -45 F93Y -97 ± 2 -259 ± 3 -46 -59 -51 N89D -39 ± 2 -301 ± 2 -44 -63 -51 FMN free56 -238 -172 - - - N89D N89D H G 81 3.4 Discussion 3.4.1 K13 accounts for slight decrease in FMN binding affinity The K13G phosphate loop in the FMN bound state does not display any significant structural difference compared to wild-type FnFld (Figure 3.2). There are minor changes to the Ψ and Ï angles, but these do not have an impact on the hydrogen bonds formed between residues on the P-loop and the FMN ribityl phosphate. The OH-N hydrogen bond formed between K13/G13 and the side chain on T10 is 2.8 Ã in both wild type FnFld and the K13G variant. Despite similar hydrogen bond contacts, replacement of K13 with a glycine results in 4-fold tighter binding of FMN to apo-K13G, due a 2-fold increase in kon and a ~2-fold decrease in koff. The structural origin of this modest increase in binding affinity is unclear. The crystal structure of K13G reveals a 5o shift in the position of the FMN cofactor (Figure 3.5) with respect to F93 in comparison to the wild type protein which may impact binding affinity. The more electropositive Eox/sq redox couple in K13G is due to improved stability of the FnFld-FMNsq complex, but again the structural rationale for this improved binding is unclear in the absence of a crystal structure of the reduced FnFld-FMNsq complex. 82 Figure 3.5 Overlay of the crystal structures of wild-type FnFld (pink) and K13G (blue) comparing the position of the FMN cofactor relative to residues Y56 and F93. The FMN cofactors are show in grey (wild-type FnFld) and yellow (K13G). 3.4.2 Y56 increases FMN binding affinity Changing Y56 to a tryptophan resulted in a 20-fold reduction in the binding affinity of FMN to apo-Y56W, Y56W was the only variant to reduce FMN binding affinity. The increase in Kd for Y56W is entirely attributed to a 20-fold increase in koff. The unchanged kon implies that this substitution does not impact the electrostatic and hydrophobic interactions that control initial binding between apo-FnFld and the FMN cofactor. In the structure of wild-type FnFld, the phenolic group of Y56 forms a hydrogen bond with the ribityl phosphate. Other long chain flavodoxins such as ones from Anabaena, S. elongates, E. coli and C. crispus harbour a conserved tryptophan residue in this position.48, 171, 172 This tryptophan forms a hydrogen bond between the NH of the indole ring and the phosphate group. Therefore, it is likely that this hydrogen bond is 83 maintained in Y56W. However, the distance of the OH-O hydrogen bond in wild-type FnFld is slightly shorter (2.5 Ã ) than the NH-O hydrogen bond seen in Anabaena (3.0 Ã ) (Figure 3.6). If this increased distance is also seen in the Y56W variant, then this might result in a weaker hydrogen bond and an increased koff. Figure 3.6 Overlay of the crystal structures of wild-type FnFld (pink) and Anabaena Fld (cyan, PDB entry code 1FLV), showing the hydrogen bond between Y56/W57 to the ribityl phosphate group of FMN. FMN cofactors are show in grey (wild-type FnFld) and yellow (Anabaena Fld). Another possible factor attributing to weaker binding in Y56W may be a conformational alteration of the 50âs loop caused by this substitution. This region is highly flexible and plays an important role in binding the oxidised form of the FMN cofactor.173 However, to confirm if there is a conformational change, the crystal structure of the Y56W variant will need to be determined. 84 The Y56W substitution did not appear to affect the stability of the apo-Y56W-FMNsq complex and the Eox/sq for this variant is similar to wild type FnFld. However, the Esq/hq was ~25 mV more electronegative. The higher electron density and larger ring size of the tryptophan indole ring compared to the phenolic ring of tyrosine may slightly destabilise the negatively charged Y56W-FMNhq complex. Weaker binding of FMNhq would result in a more electronegative Esq/hq value. 3.4.3 N89 results in a more electropositive Esq/hq N89 present in FnFld aligns with a conserved aspartic acid residue in Flds.57, 62 The carboxylate side chain of the aspartic acid residue (D90 in S. elongates and D95 in D. vulgaris) is < 6 Ã from FMN N1 (Figure 3.7). It is one of 5-6 acid residues that are > 13 Ã from the FMN N1. The negative change associated with these residues collectively produces an unfavourable environment for the negatively charged FMN hydroquinone. Consequently, they destabilise the formation of the apo-N89D-FMNhq complex, resulting in a more electronegative Esq/hq. Figure 3.7 A) Position of D90 relative to FMN N1 in Fld from S. elongates (PDB entry code 1CZL) B) Position of D95 relative to FMN N1 in Fld from D. vulgaris (PDB entry code 3KAP) C) Position of N89 relative to FMN N1 in FnFld. 85 The effects of this single carboxylate group on the FMNsq/hq potential was first observed for S. elongates in D. vulgaris. The D90N variant in S. elongates resulted in ~40 mV more electropositive Esq/hq.57 The D95N mutant in D. vulgaris caused Esq/hq to become ~45 mV more electropositive.62 Therefore, it would be expected that N89D should produce a more electronegative Esq/hq value. This is exactly what was observed experimentally; the N89D variant had an Esq/hq value that was ~45 mV more electronegative compared to the Esq/hq of wild-type FnFld. 3.4.4 Role of F93 The crystal structure of wild-type FnFld shows that F93 along with Y56 form a binding pocket for the FMN isoalloxazine ring. The F93Y variant demonstrated a 4-fold increase in FMN binding affinity compared to wild-type FnFld. This is attributed to a 2-fold increase in kon and a 2-fold decrease in koff. In addition, the Eox/sq was found to be ~50 mV more electronegative, but there was no change in Esq/hq. A reverse mutation in Anabaena Fld (Y94F) demonstrated the opposite effect, a decrease in FMN binding affinity and a ~30 mV more electropositive value for Eox/sq.50 Thus, it is not surprising that F93Y displayed matching characteristics. The only difference between phenylalanine and tyrosine is the presence of a hydroxide group on the C4 position. This makes tyrosine a more electron-rich amino acid relative to phenylalanine. A flanking tyrosine may result in in preferential stabilisation of the apo-FnFld-FMNox complex because of stronger dispersion forces between the electron-rich tyrosine and the oxidised isoalloxazine ring. 86 3.5 Concluding remarks In this study we were able to establish the significance of four naturally occurring substitutions found in the primary sequence of FnFld. Three of these substitutions (Y56, N89 and F93) account for the electropositive reductions observed with the FMN-FnFld complex. Y56 and N89 contribute towards the electropositive values of Esq/hq. This is likely because they do not provide a negative charge close to the FMN N1, therefore they do not destabilise the apo-FnFld-FMNhq complex. F93 contributed towards the electropositive value for Eox/sq. This may be accounted for by the electron-poor phenylalanine (relative to tyrosine) having weaker dispersion forces with the isoalloxazine ring on FMNox. It is likely that this affect is also responsible for the reduced FMN binding affinity caused by F93 compared with the conserved sequence. Y56 was the only substitution that led to a greater binding affinity. This may be due to the formation of a tighter hydrogen bond between Y56 and the FMN ribityl phosphate. The other three substitutions all show a decrease in binding affinity with FMN. The bulky K13 may increase the size of the conformational shift undertaken by the P-loop following FMN binding. This could account for the decreased kon and increase koff. N89 is a neutral residue that replaces a negatively charged aspartic acid. The loss of this charge may partially stabilise the apo-FnFld-FMNhq complex, resulting in a more electropositive Esq/hq. These data have provided us with much more insight into the roles of these amino acid substitutions. Combining these data with the data discussed in Chapter 2 enabled us to determine that FnFld has many unique properties that may impact its biological function as an electron transfer protein. The unique positive binding cleft on FnFld may prove beneficial for the design of novel drugs to block the electron transfer properties of this protein and the growth of F. nucleatum. In addition, this research provides greater insight into the mechanisms and redox properties of 87 flavodoxins and may also be used for making modifications to existing flavodoxins or the development of new flavodoxins. This could have uses in genetically modifying organisms that rely on ferredoxin as their primary electron transfer protein to express flavodoxin in order to improve their survival in low iron conditions. 88 4. Lanthionine Synthase 4.1 Chapter Summary F. nucleatum encodes for lanthionine synthase (LS), a PLP-dependent enzyme that generates H2S and lanthionine through the β-replacement of L-cysteine by a second molecule of L-cysteine. Lanthionine is an important amino acid for F. nucleatum as it is incorporated into the peptidoglycan layer as part of the peptide cross-link. The amino acid sequence for LS is similar to cystathionine β-synthase (CBS) and O-acetylserine sulfhydrylase (OASS), two enzymes that elicit catalytic promiscuity with respect to the type of β-replacement reaction they catalyse. Give the catalytic promiscuity of this class of enzymes, we sought to uncover what reactions LS can catalyse and to confirm whether the production of lanthionine is the physiologically role of the enzyme in F. nucleatum. In addition, we used stopped-flow spectroscopy to track the formation and decay of the PLP intermediates that form during the catalytic cycle. Finally, we sought to gain additional structural insight into the role of S234, which is part of a unique loop (233MSIG236) that interacts with the thioether product. Kinetic analysis reveals that LS does elicit some substrate promiscuity. It displays OASS activity which involves the conversion of O-acetyl-L-serine and HS- to cysteine and acetate. However, the turnover for this reaction in LS was >30-fold lower than other OASS enzymes. This was likely accounted for by slower conversion to the α-aminoacrylate intermediate, as determined by stopped-flow spectroscopy. In addition, this reaction was not considered physiologically relevant to F. nucleatum as the bacterial genome does not encode for the enzyme that converts L-serine to O-acetyl-L-serine substrate. Kinetic and HPLC confirmed that LS preferentially catalyses 89 the β-replacement of L-cysteine with a second molecule of L-cysteine to produce L-lanthionine and H2S. Replacement of S234 with alanine did not affect the OASS activity of the enzyme or the rate of H2S formation in the presence of L-cysteine. However, the substitution led to a 2000-fold reduction in kon for D,L-lanthionine binding to LS. It also caused the reaction to become reversible, in that S234A variant can also catalyse the β-replacement of L-lanthionine by sulfide to form L-cysteine. Finally, we tested the binding of nine product analogues of L-lanthionine and identified several compounds that bound to the LS active site. This may help us in the development of new inhibitors that may form the basis of new therapeutics to treat F. nucleatum infections. 4.2 Material and Methods 4.2.1 Materials Restriction enzymes were from New England Biolabs (Ipswich, MA, USA). Oligonucleotides were purchased from Integrated DNA Technologies (Coralville, IA, USA). F. nucleatum genomic DNA was from the American Tissue Culture Collection (Manassas, VA, USA). XL1-Blue competent cells and PfuTurbo DNA polymerase were from Agilent (Santa Clara, CA, USA). The DCâ¢ Protein Assay was from Bio-Rad (Hercules, CA, USA) and the E.Z.N.A plasmid mini kit was from VWR (Radnor, PA, USA). 4.2.2 Cloning petFld into pET15b plasmid The gene coding for LS (FN1220) was amplified from the genomic DNA of F. nucleatum strain ATCC 25586 with PfuTurbo DNA polymerase using primers 5â²â G GAA TTC CAT ATG 90 TTG AGA ATA AAG TTT GAT AGG AGG â3â² (forward primer) and 5â²â CGG GAT CCT TAT AAA TCA CAA ATA TTA GAT AAA TAT TTT TC â3â² (reverse primer) that included NdeI and BamHI restriction sites (shown in bold).111 The amplified PCR product was inserted into the NdeI and BamHI sites of the pET15b vector. The PCR reaction mixture contained PfuTurbo DNA polymerase (0.05 U/μL), PfuTurbo reaction buffer, 0.2 mM of each dNTP, 1.0 ng/mL F. nucleatum genomic DNA, 2.5 ng/mL of forward primer and 2.5 ng/mL of reverse primer in a total volume of 50 µL. The gene was amplified by cycling the reaction mixture 30 time through the following temperatures: 95 oC (30 s), 53 oC (30 s) and 72 oC (60 s) following incubation at 95 oC for 2 minutes. The PCR product, cleaved with NdeI and BamHI, was ligated into a pET15b plasmid digested with the same restriction enzymes. The ligated product was transformed into the XL1-Blue competent cells, and the cell mixture was plated onto LB-agar plates supplemented with ampicillin (100 µg/mL). Plasmid from a transformed cell was isolated using an E.Z.N.A plasmid mini kit.148 DNA sequencing by the Sequencing and Bioinformatics Consortium at UBC Vancouver confirmed the presence of the correct LS coding sequence in the pET15b plasmid. 4.2.3 Purification and expression of wild type lanthionine synthase Plasmid harbouring the cDNA for LS (named petLS) was transformed into E. coli strain Rosetta(DE3)pLysS for protein expression. A starter culture was prepared by inoculating 200 mL of LB media, supplemented with ampicillin (100 µg/mL) and chloramphenicol (34 µg/mL), with a single colony generated from a transformed cell. The starter culture was incubated for approximately 16 hours at 37 oC. Aliquots (10 mL) of the starter culture were added to 500 mL of Terrific Broth, supplemented with ampicillin (100 µg/mL) and chloramphenicol (34 µg/mL). The culture was incubated at 30 oC at 200 rpm until the optical density at 600 nm was ~0.6. Isopropyl 91 β-D-1-thiogalactopyranoside (ITPG) was added to cell cultures to a final concentration of 0.2 mM. The cultures continued to incubate for approximately 16 hours at 25 oC at 200 rpm. Cell cultures were centrifuged at 6371g for 15 min, and cell pellets were pooled together and stored at -80 oC until protein isolation. Cells (30 g) were resuspended in 200 mL of 50 mM Tris-HCl, pH 7.5 containing protease inhibitors phenylmethylsulfonyl fluoride (1.0 mM) and benzamidine (0.1 mM). The cell suspension was sonicated on ice at 32 % power for 8 s at 1 min intervals for 1 hour. The cell suspension was centrifuged at 39191g for 60 min at 4 oC. The pooled supernatants were applied to a Ni-nitrilotriacetate column (16 Ã 30 mm) (MilliporeSigma, MA, USA) equilibrated with 50 mM Tris-HCl, pH 7.5 and 0.02 M imidazole. Protein was eluted from the column using a step gradient: 5 CV of 100% Buffer A (50 mM Tris-HCl, pH 7.5 with 0.02 M imidazole) and 0% Buffer B (50 mM Tris-HCl, pH 7.5 and 0.5 M imidazole), 5 CV of 4 % Buffer B, 5 CV of 60 % Buffer B and 5 CV of 100 % Buffer B. Elution of the protein was monitored at 280 nm. Protein eluted at 60% Buffer B was pooled and dialysed against 4 L of 50 mM Tris-HCl, pH 7.5 for 16 hr. The dialysate was concentrated to ~10 mL using a centricon (Vivaspin 20, GE Healthcare, IL, USA) with a 30 kDa MW cut-off filter. The protein was then applied to a Q-sepharose HP column (26 Ã 140 mm) equilibrated with 50 mM Tris-HCl, pH 7.5. The protein was eluted using a linear gradient from 100 % Buffer C (50 mM Tris-HCl, pH 7.5) to 100 % Buffer D (50 mM Tris-HCl), pH 7.5, 0.5 M NaCl over 8 CV. Fractions containing protein were pooled and concentrated to 10 â 15 mL. SDS-PAGE was performed to assess the purity of the protein. Stocks of the protein containing 20% glycerol were prepared and stored at -80 oC. The concentration of the protein was determined through a Lowry assay and by measuring the absorbance at 277 nm using an extinction coefficient of 33.5 mM-1 cm-1. 92 4.2.4 Variants produced by site-directed mutagenesis The S234A variant of LS was created by site directed mutagenesis using the QuikChange method with 5â-CAT AAA ATT CAA GGT ATG GGA ATG GCT ATA GGT GGT ATT CCA GCA GTT TAT G-3â and 5â C ATA AAC TGC TGG AAT ACC ACC TAT AGC CAT TCC CAT ACC TTG AAT TTT ATG-3â as a forward and reverse primers, respectively .169 The presence of the mutation in LS and the absence of PCR-induced errors was confirmed by DNA sequencing. S234A was expressed and isolated following the same protocol for wild type LS described in Section 4.2.3. 4.2.5 Steady-state kinetic assays measuring H2S production The production of H2S by wild type LS and S234A in the presence of L-cysteine was monitored by a continuous spectrophotometric assay that followed the formation of lead sulfide from lead acetate and H2S. The reactions were performed at 10 oC, 25 oC and 37 oC for wild type LS and at 25 oC for S234A. The reaction mixture contained 4 mM lead acetate and varying concentrations of L-cysteine (0.1 mM to 100 mM). The mixture was made up to 1 mL with 100 mM HEPES-NaOH (pH 7.4). The reaction was initiated with the addition of 10 μL of 3.2 μM LS to give a final concentration of 32 nM. The initial velocity of the reaction was determined by measuring the change in absorbance at 390 nm (ÎÎµ = 5500 M-1 cm-1) using a UV/vis spectrophotometer (Lambda 265, Perkin Elmer, MA, USA) over the course of two minutes. The initial rate of H2S production was plotted against L-cysteine concentration and fitted to Michaelis-Menten equation (Equation 4.1). Equation 4.1: ð£ =ðððð¥[ð]ð¾ð+[ð] 93 4.2.6 Steady-state kinetic assays measuring L-cysteine production An acid-ninhydrin assay was used to follow the OASS activity of wild type LS and S234A.174 The reaction was performed in 100 mM HEPES-NaOH (pH 7.4) at 25 oC. A 1 mL reaction contained either 2.5 mM Na2S and varying concentrations of O-acetyl-L-serine (0.1 to 5 mM), 2.5 mM Na2S and varying concentrations of D,L-lanthionine (0.5 mM to 20 mM) or 5 mM O-acetyl-L-serine and varying concentrations of Na2S (0.1 to 2.5 mM). The reaction was initiated with the addition of 50 nM LS. At three time points (1 min, 2 min and 4 min), aliquots (200 μL) of the reaction mixture were quenched with the addition of 16 % (v/v) trichloroacetic acid (40 μL). An acid-ninhydrin solution (240 μL) and glacial acetic acid (240 μL) were then added to the quenched solution. The solution was then heated at 100 °C for 4 min. The reaction mixture was cooled on ice for 5 mins and then cold 100% ethanol (720 μL) was added. The absorbance at 560 nm of the reaction mixture was measured using a UV/visible spectrophotometer (Lambda 265, Perkin Elmer, MA, USA). The concentration of L-cysteine was determined from a standard curve using a 240 µL solution of L-cysteine (0.01 â 0.5 µmol) in place of the quenched reaction mixture. The initial rate of L-cysteine formation as determined from a plot of L-cysteine concentration against time. The initial rate of L-cysteine production was plotted against D,L-lanthionine, O-acetyl-L-serine or Na2S concentration and the data were fitted to Michaelis-Menten equation (Equation 4.1) to obtain kcat and Km. 4.2.7 High-performance liquid chromatography (HPLC) Products from reactions potentially facilitated by wild type LS and S234A were determined by reverse-phase HPLC. To confirm that lanthionine is the principle product formed by LS in the presence of L-cysteine, a 1 mL reaction mixture containing L-cysteine (1.0 mM) and LS (100 nM) 94 in 40 mM potassium phosphate pH 7.6 was incubated at 37 oC for 120 mins. The enzyme was then filtered from the reaction mixture by centrifugation using a Microcon YM-10 filter (10 kDa cut-off, Amicon). Following this step, 500 µL of a dansyl chloride solution (1.5 mg/mL) was added to the reaction mixture. The mixture was then incubated at room temperature for 30 mins while avoiding exposure to light. An aliquot (100 µL) of 2% ethylamine was then added to the reaction mixture and the solution was inverted until the yellow colour faded. Aliquots (10 µL) of each sample were injected into a C-18 column (Agilent Poroshell 210, 4.6Ã100 mm) and the dansylated products were separated at a flow rate of 0.5 ml/min at 40°C with a mobile phase of 33.25/66.75 (v/v) methanol/water containing 0.6â% (v/v) glacial acetic acid and 0.008â% (v/v) triethylamine. The appearance of dansylated products was monitored at 338 nm. The experiment was repeated using a reaction mixture containing 0.5 mM L-homocysteine and 0.5 mM L-serine or L-cysteine to assess the ability of LS to generate L-cystathionine from the β-replacement of either L-serine or L-cysteine with L-homocysteine. Finally, the experiment was repeated using 0.5 mM L-serine and L-cysteine to assess production of L-lanthionine from L-serine and L-cysteine. 4.2.8 Stopped-flow spectroscopy The pre-steady state kinetic properties describing the interaction between wild type LS or S234A and various substrates were investigated using a SF-61DX2 stopped-flow spectrophotometer (TgK Scientific). The stopped-flow reactions were performed in 100 mM HEPES-NaOH (pH 7.4) at 10 °C. A solution of enzyme (20-50 μM) was mixed with varying concentrations of either L-cysteine, O-acetyl-L-serine or D,L-lanthionine under the pseudo-first order conditions. Absorbance changes over time were recorded at a single wavelength using a photomultiplier and at multiple wavelengths (300 to 700 nm) using a photodiode array detector. The absorbance traces were fitted to either a single or double exponential equation using Kinetic 95 Studio (TgK Scientific). If the observed rate constant displayed a hyperbolic dependence on the substrate concentration, then the data were fitted to a hyperbolic function (Equation 4.2). Equation 4.2 ðððð =ðððð[ð¿]ð¾ð+[ð¿] where Kd is the dissociation constant, [L] is the substrate concentration and klim is the limiting first order rate constant for either external aldimine formation or formation of the α-aminoacrylate intermediate. 4.2.9 Fluorescence titrations The equilibrium dissociation constants (Kd) for the binding of D,L-lanthionine and nine product analogues to wild type LS were determined through fluorescence titrations. The change in fluorescence corresponds to formation of the external aldimine with the incoming ligand. The change in fluorescence of the PLP cofactor was followed by exciting the cofactor at its absorbance maxima (408 nm) and recording at the emission maxima of 545 nm. All experiments were performed in 100 mM HEPES-NaOH, pH 7.4 at 25 oC. A 3 mL solution of 0.1 µM of wild type LS was placed in a fluorescence cuvette and inserted in LS50âB fluorescence spectrophotometer (Perkin-Elmer, MA, USA). Aliquots (1 µL) of the ligand were added to increase the concentration by either 2, 5 or 10 µM. The Kd was determined by fitting the change in fluorescence to a quadratic binding isotherm (Equation 4.3). Equation 4.3 ð¥ð¹ = (ð¥ð¹ððð¥2ð¸0)(ð¿0 + ð¸0 + ð¾ð â [(ð¿0 + ð¸0 + ð¾ð)2 â 4ð¿0ð¸0]12) 96 where ÎF is the change in fluorescence emission, ÎFmax is the maximum change in fluorescence emission, Eo is the total enzyme concentration, Lo is the total ligand concentration, and Kd is the equilibrium dissociation constant for the external aldimine of LS. 4.3 Results 4.3.1 Properties of purified lanthionine synthase The cDNA corresponding to LS was successfully inserted into pET15b vector. Wild type and S234A were recombinantly expressed in E. coli and purified by Ni-NTA and anion exchange chromatography. The homogeneity and approximate size of the proteins was confirmed by SDS-PAGE. Wild type LS appeared as two bands with apparent molecular masses of ~38 and 33 kDa (Figure 4.1) which are both close to the expected molecular weight of 34.6 kDa. The two bands were excised and sent to the Proteomics Core Facility at UBC Vancouver. The bands were digested with trypsin and ran on a Q-ToF mass spectrometer. A database search against the Fusobacterium protein database was performed to identify the two bands. Both bands were found to contain LS and it was unclear why they separated into distinct bands. One possibility is separation due to a post-translational modification. 97 Figure 4.1 SDS 10% polyacrylamide gel showing wild type lanthionine synthase. Lane 1, BLUeye Pre-stained protein ladder (Sigma); Lane 2, crude extract; Lane 3, protein eluted from Ni-NTA column; Lane 4, protein eluted from Q-sepharose column. The UV-visible absorbance spectrum of LS shows a major peak at 406 nm and a minor peak at 320 nm (Figure 4.2). The predominant peak at 406 nm represents the internal aldimine with a Schiff base linkage. The addition of L-cysteine and D,L-lanthionine caused a blue-shift to an absorption maximum of 428 nm and small increase in absorbance at 320 nm. This shift likely represents the formation of an external aldimine with the product L-lanthionine. The addition of O-acetyl-L-serine to LS led to an increase in absorbance and a blue shift to 463 nm, that was accompanied by significant increase in absorption at 320 nm. This spectral intermediate likely represents the formation of an α-aminoacrylate intermediate that forms following β-elimination of the acetyl group from the O-acetyl-L-serine external aldimine. Lane 1 Lane 2 Lane 3 Lane 4 98 Figure 4.2 Absorption spectra of the PLP cofactor from wild type lanthionine synthase (black line) following a 5 min incubation with 1 mM L-cysteine (black dashes), 1 mM D,L-lanthionine (black dots) and 1 mM O-acetyl-L-serine (grey dashes). All spectra were recorded in 100 mM HEPES-NaOH (pH 7.4) at 25 oC. The UV-visible absorbance spectrum of S234A shows a major peak at 406 nm and a minor peak at 320 nm (Figure 4.3). The predominant peak at 406 nm represents the internal aldimine with a Schiff base linkage. The addition of L-cysteine and D,L-lanthionine caused a blue-shift to an absorption maximum of 428 nm and an increase in absorbance at 320 nm. This shift likely represents the formation of an external aldimine with the product L-lanthionine. The addition of O-acetyl-L-serine to LS led to an increase in absorbance and a blue shift to 463 nm, that was accompanied by significant increase in absorption at 320 nm. This spectral intermediate likely represents the formation of an α-aminoacrylate intermediate that follows β-elimination of the acetyl group from the external aldimine with O-acetyl-L-serine. 99 Figure 4.3 Absorption spectra of the PLP cofactor from S234A (black line) following 5 min incubation with 1 mM L-cysteine (black dashes), 1 mM D,L-lanthionine (black dots) and 1 mM O-acetyl-L-serine (grey dashes). All spectra were recorded in 100 mM HEPES-NaOH (pH 7.4) at 25 oC. 4.3.2 Steady-state kinetics analysis of H2S production The steady-state kinetic parameters associated with the production of H2S by lanthionine synthase were determined by the lead acetate assay described in section 4.2.5. The initial rate of H2S formation elicited a hyperbolic dependence on L-cysteine concentration and the data were fitted to the MichaelisâMenten equation (Equation 4.1, Figure 4.4). At 25 oC, the turnover rate of L-cysteine by wild type LS was determined to be 8.2 ± 0.2 s-1 with a KM(L-cys) of 1.8 ± 0.1 mM and a kcat/KM of 4600 ± 400 s-1 M-1 (Table 4.1). S234A had a 7-fold higher kcat than wild type LS (58.1 ± 2.9 s-1), however it also had a 9-fold higher KM for L-cysteine (18.1 mM). This resulted in the 100 catalytic efficiency of S234A being ~70% of wild type LS (3200 ± 600 s-1 M-1). At 37 oC the kcat for wild type LS was 16.6 ± 0.3 s-1, the KM(L-cys) was 1.7 ± 0.1 mM and kcat/KM was 9800 ± 600 M-1 s-1. At 10 oC the kcat for wild type LS was 3.9 ± 0.1 s-1, the KM was 0.9 ± 0.1 mM and kcat/KM was 4300 ± 500 M-1 s-1. The rate of formation of H2S by wild type LS in the presence of 20 mM L-cysteine did not change following the addition 20 mM L-homocysteine which indicates that LS is likely inefficient at catalysing the production of L-cystathionine through the β-replacement of L-cysteine with L-homocysteine, which is one of the H2S forming reactions of CBS. Table 4.1 Steady-state kinetic parameters for formation of H2S by lanthionine synthase and the S234A variant in the presence of L-cysteine. Reactions were performed at 25 oC in 100 mM HEPES-NaOH (pH 7.4). Enzyme kcat (s-1) KM(mM) kcat/KM (s-1 M-1) Wild type LS 8.2 ± 0.2 1.8 ± 0.1 4600 ± 400 S234A 58.1 ± 2.9 18.1 ± 2.5 3200 ± 600 Figure 4.4 Michaelis-Menten plot of the production of H2S at 25 oC in the reaction of L-cysteine catalysed by A) wild type LS and B) S234A. A B 101 4.3.3 Steady-state kinetic analysis of L-cysteine production The steady-state kinetic parameters associated with the production of L-cysteine by lanthionine synthase were determined by the acid-ninhydrin assay described in Section 4.2.6. The initial rate of L-cysteine formation elicited a hyperbolic dependence on O-acetyl-L-serine concentration and consequently the initial velocity data were fitted to the MichaelisâMenten equation (Equation 4.1, Figure 4.5). The turnover rate of O-acetyl-L-serine by wild type LS was determined to be 6.3 ± 0.2 s-1 with a KM(OAS) of 0.41 ± 0.04 mM and a kcat/KM of 15 ± 2 s-1 mM-1 (Table 4.2). With wild type LS the KM for HS- was 0.17 ± 0.03 mM and the catalytic efficiency was 40 ± 9 s-1 mM-1. S234A had a 3-fold higher kcat than wild type LS (16.6 ± 0.9 s-1); however, it also had a 5-fold higher KM for O-acetyl-L-serine (2.2 mM). This resulted in the catalytic efficiency of S234A being ~50% of wild type LS (7.5 ± 1.8 s-1 mM-1). There was no observable production of L-cysteine in the presence of D,L-lanthionine with wild type LS. However, S234A was able to catalyse the β-replacement of L-lanthionine by sulfide to produce L-cysteine with a turnover rate of 8.8 ± 0.2 s-1. The KM for D,L-lanthionine was 1.1 ± 0.1 mM and the catalytic efficiency was 8.0 ± 0.9 s-1 mM-1. Table 4.2 Steady-state kinetic parameters for formation of L-cysteine by lanthionine synthase and the S234A variant in the presence of O-acetyl-L-serine and Na2S. Reactions were performed at 25 oC in 100 mM HEPES-NaOH (pH 7.4). Enzyme kcat (s-1) KM(OAS) (mM) kcat/KM(OAS) (s-1 mM-1) KM(HS-) (mM) kcat/KM(HS-) (s-1 mM-1) Wild type LS 6.3 ± 0.2 0.41 ± 0.04 15 ± 2 0.17 ± 0.03 40 ± 9 S234A 16.6 ± 0.9 2.2 ± 0.4 7.5 ± 1.8 - - 102 Figure 4.5 Michaelis-Menten plots of the production of L-cysteine at 25 oC in the reaction of O-acetyl-L-serine and Na2S catalysed by A) wild type LS and B) S234A. 4.3.4 HPLC analysis of products The ability of wild type LS to catalyse four different reactions associated with CBS was analysed by reverse phase HPLC. These reactions were: 1) the production of L-lanthionine from two equivalents of L-cysteine, 2) the production of L-cystathionine from L-cysteine and L-homocysteine, 3) the production of L-cystathionine from L-serine and L-homocysteine and 4) the production of L-serine and H2S from the β-replacement L-cysteine by H2O (Figure 4.6). A B 103 Figure 4.6 Four reactions catalysed by human CBS. Reaction 1 is the formation of L-lanthionine from the β-replacement L-cysteine by a second molecule of L-cysteine. Reaction 2 is the β-replacement of L-cysteine with L-homocysteine to produce L-cystathionine and H2S, Reaction 3 is the β-replacement of L-serine with L-homocysteine to produce L-cystathionine and H2O. Reaction 4 is the formation of L-serine and H2S from the β-replacement L-cysteine by H2O. Reaction 3 is the canonical reaction of CBS, while reactions 1,2 and 4 are H2S forming reactions of CBS, with reaction 2 being the most efficient. Following a 2 hr incubation of wild type LS with L-cysteine, HPLC analysis revealed that lanthionine was the only product (Figure 4.7A). The HPLC chromatogram show that there was a negligible peak associated with L-cysteine starting material. These data demonstrate that LS can catalyse Reaction 1 with a >95% yield after 2 hr and does not catalyse reaction 4. 104 A 2 hr incubation of wild type LS with L-cysteine and an equimolar amount of L-homocysteine led to a major peak associated with lanthionine, a minor peak associated with cystathionine and a small peak associated with the L-homocysteine starting material (Figure 4.7B). These data show that LS can also catalyse Reaction 2 but is selective towards Reaction 1. Incubation of wild type LS with L-serine and either L-homocysteine or L-cysteine over 2 h did not show the production of any new products (Figure 4.8). Thus, LS is not capable of catalysing Reaction 3. Figure 4.7 Analysis of dansylated products using reverse phase HPLC. A) Products obtained following a 2 h incubation of LS (100 nM) with L-cysteine (0.5 mM) (top), pure D,L-lanthionine standard (middle), pure L-cysteine standard (bottom). B) Products obtained following a 2 h A B 105 incubation of LS (100 nM) with L-cysteine (0.5 mM) and L-homocysteine (0.5 mM) (top), pure L-cystathionine standard (middle), pure L-homocysteine standard (bottom). Figure 4.8 Analysis of dansylated products using reverse phase HPLC. A) Products obtained following a 2 h incubation of LS (100 nM) with L-serine (0.5 mM) and L-homocysteine (0.5 mM) (top), pure L-cystathionine standard (middle), pure L-serine standard (bottom). B) Products obtained following a 2 h incubation of LS (100 nM) with L-cysteine (0.5 mM) and L-serine (0.5 mM) (top), pure L-cysteine standard (middle), pure L-serine (bottom). The ability of S234A to catalyse Reactions 1, 2 and 4 (Figure 4.6) was also analysed by reverse phase HPLC. Following a 2 hr incubation of wild type S234A with L-cysteine, lanthionine was the only product detected, but there was significant peak associated with the L-cysteine starting material (Figure 4.9A). This would suggest that S234A can catalyse Reaction 1, but not A B 106 Reaction 4. However, the large peak associated with L-cysteine, supports the steady state assays that reveal that S234A variant can catalyse the β-replacement of L-lanthionine by sulfide to form L-cysteine. A 2 hr incubation of S234A with L-cysteine and an equimolar amount of L-homocysteine led to two similarly sized peaks that were associated with lanthionine and cystathionine (Figure 4.9B). There was also a negligible peak associated with the L-homocysteine starting material. These data suggest that LS catalyses both Reactions 1 and 2, with no preference towards either reaction. Figure 4.9 Analysis of dansylated products using reverse phase HPLC. A) Products obtained following a 2 h incubation of S234A (100 nM) with L-cysteine (0.5 mM) (top), pure L-cysteine standard (middle), pure D,L-lanthionine standard (bottom). B) Products obtained following a 2 h A B 107 incubation of S234A (100 nM) with L-cysteine (0.5 mM) and L-homocysteine (0.5 mM) (top), pure L-cystathionine standard (middle), pure L-homocysteine standard (bottom). 4.3.5 Pre-steady-state kinetic analysis with L-cysteine Rapid mixing of wild type LS (66 µM) with L-cysteine (2 mM) resulted in multiphasic absorbance changes in the PLP cofactor (Figure 4.10). From 0. 75 to 11 ms, there is a slight decrease in the absorbance maxima at 406 nm; this decrease coincided with a small increase in absorbance in λmax at 470 and 320 nm (Figure 4.10A). This transition had isosbestic points at 353 and 437 nm. This kinetic event likely reflects the formation of α-aminoacrylate intermediate in both tautomeric states from the internal aldimine.175 From 11 ms to 3.9 s there was a decrease in absorbance at both λmax at 406 nm and 470 nm and a further increase at 320 nm (Figure 4.10B). This second kinetic phase may reflect the continued conversion from the internal aldimine to the α-aminoacrylate alongside a shift in the equilibrium of the tautomeric states of the α-aminoacrylate in favour the enol-imine form over the keto-enamine form (Figure 4.11).94 Finally, from 17.6 s to 146 s there was a decrease in absorbance at 320 nm and an increase at 428 nm (Figure 4.10C). This final kinetic phase likely corresponds to the formation of the external aldimine with L-lanthionine following nucleophilic attack by a second molecule of L-cysteine.176 108 Figure 4.10 Time resolved spectra following rapid mixing of L-cysteine (2 mM) and wild type LS (66 µM) in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. The spectra were acquired on a log time base over 150 s and are split into three domains, a subset of the spectra are shown. A) 0.75 ms to 11.3 ms, B) 11 ms to 3.9 s and C) 17.6 s to 148 s. The arrows indicate the direction of the absorption change. A B C 109 Figure 4.11 The conversion of the PLP cofactor from the internal aldimine in LS to the α-aminoacrylate intermediate and then to the external aldimine with lanthionine. The α-aminoacrylate intermediate exists in one of two tautomeric states, the enol-imine form and the keto-enamine form. Single-wavelength stopped-flow spectroscopy was used to measure rate constants associated with these three distinct kinetic events. The absorbance traces at 320 and 406 nm were biphasic over 1 s. Consequently, they were fitted to a standard double exponential equation (Figure 4.12A/B, Table 4.3). The initial observed rate constant from the first kinetic phase (kobs1) acquired at 320 and 406 nm displayed a linear dependence on L-cysteine concentration (Figure 4.12C). The bimolecular rate constants, determined from the slope, were 35.3 ± 1.6 mM-1 s-1 (406 nm) and 36.1 ± 2.8 mM-1 s-1 (320 nm). 110 The observed rate constant for the second kinetic phase (kobs2) elicited a hyperbolic dependence on the concentration of L-cysteine (Figure 4.12D). At 406 nm the hyperbolic fit of kobs2 gave a binding constant (Kd) for L-cysteine of 0.57 ± 0.08 mM and a klim of 3.6 ± 0.13 s-1. At 320 nm the Kd for L-cysteine was 0.78 ± 0.05 mM and klim was estimated at 4.5 ± 0.13 s-1. Under the reaction conditions of this experiment, klim is associated with the limiting rate constant for the formation of the α-aminoacrylate intermediate. The absorbance change in λmax at 428 nm was monophasic from 3 to 350 s with L-cysteine (Figure 4.13). Fitting the trace to a single exponential equation gave an observed rate constant of 0.014 ± 0.008 s-1. The observed rate constant associated with this catalytic step is too slow to accurately reflect the true kinetic properties of LS. This attenuated observed rate constant may be due to the accumulation of a dead-end complex formed from LS and L-lanthionine during acquisition of data. Table 4.3 Pre-steady state kinetic parameters for LS and S234A with L-cysteine determined following absorption changes at 320 nm and 406 nm. Enzyme Wavelength (nm) kon (mM-1 s-1) Kd (mM) klim (s-1) Wild type 320 36.1 ± 2.8 0.78 ± 0.05 4.5 ± 0.13 Wild type 406 35.3 ± 1.6 0.57 ± 0.08 3.6 ± 0.13 S234A 406 14.5 ± 0.6 2.2 ± 0.5 2.7 ± 0.2 111 Figure 4.12 Single wavelength absorbance trances acquired at A) 320 nm and B) 406 nm following rapid mixing of wild type LS (66 µM) with varying concentration of L-cysteine (1 â 10 mM). Arrows show the direction of the absorbance traces as L-cysteine concentration increases. These traces were fitted to a double exponential equation to generate kobs1 and kobs2. C) Linear dependence of kobs1 with wild type LS measured at 320 nm (grey) and 406 nm (black) on L-cysteine concentration. D) Hyperbolic dependence of kobs2 measured at 320 nm (grey) and 406 nm (black) on L-cysteine concentration. All reactions were performed in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. 112 Figure 4.13 Averaged single wavelength absorbance trace at 428 nm obtained after mixing wild type LS (66 μM) with 2 mM L-cysteine. The absorbance trace was fitted to a single exponential equation. Absorbance changes in the PLP spectra upon rapid mixing of S234A (22 µM) with L-cysteine (Figure 4.14, Table 4.3) leads to two resolvable kinetic events. The photodiode array spectra show that from 0.8 ms to 300 ms there is a decrease in absorbance at λmax at 406 nm and increases at ~470 nm and 320 nm, which likely represents the formation of the α-aminoacrylate intermediate in both tautomeric states. The larger increase at 470 nm compared to the wild type enzyme suggests that the S234A substitution shifts the enzyme to the open conformation of the enzyme which favours the keto-enamine state over the enol-imine state.94 The second kinetic phase from 0.9 s to 40 s consists of an increase in the λmax at 406 nm and decreases at ~470 and 320 nm. The second kinetic phase represent the decay of the α-aminoacrylate intermediate. Single wavelength stopped flow absorbance traces at 406 nm were fitted to a double exponential equation (Figure 4.15A). The initial observed rate constant (kobs1) 113 measured at 406 nm showed a linear dependence on L-cysteine concentration (Figure 4.15B) with a kon of 14.5 ± 0.6 mM-1 s-1. The second rate constants (kobs2) showed a hyperbolic dependence on L-cysteine concentration (Figure 4.15C) with Kd of 2.2 ± 0.5 mM and klim of 2.7 ± 0.2 s-1. Under the reaction conditions of this experiment, klim is associated with the limiting rate constant for the formation of the α-aminoacrylate intermediate. Figure 4.14 Time resolved spectra following rapid mixing of L-cysteine (2 mM) and S234A (22 µM) in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. The spectra were acquired on a log time base over 40 s and are split into two time domains, a subset of the spectra is shown, A) 0.8 ms to 300 ms, B) 0.9 s to 40 s. A B 114 Figure 4.15 A) Single wavelength absorbance trances acquired at 406 nm following rapid mixing of S234A (22 µM) with varying concentration of L-cysteine (1 â 10 mM). Arrows show the direction of the absorbance traces as L-cysteine concentration increases. These traces were fitted to a double exponential equation to generate kobs1 and kobs2. B) Linear dependence on L-cysteine concentration of kobs1 measured at 406 nm with S234A. C) Hyperbolic dependence on L-cysteine concentration of kobs2 measured at 406 nm with S234A. All reactions were performed in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. 4.3.6 Pre-steady-state kinetic analysis with D,L-lanthionine The absorbance changes in the PLP spectra following rapid mixing of wild type LS (58 µM) and D,L-lanthionine are presented in Figure 4.16. The time-resolved photodiode array spectra 115 from 2 ms to 150 s revealed two resolvable kinetic events. From 2 ms to 6.4 s there is a decrease in λmax at 406 nm which coincides with an increase in absorbance at 320 nm. This initial kinetic event may represent the formation of a gem-diamine intermediate (Figure 4.17).176 From 6.4 s to 148 s there is shift in the λmax from 406 nm to 428 nm and a decrease in absorbance at 320 nm. This second kinetic phase is likely the result of the conversion of the gem-diamine intermediate to the external aldimine with lanthionine. Figure 4.16 Time-resolved UV-visible absorption spectra following rapid mixing of wild type LS (62 µM) with D,L-lanthionine (2 mM) over the course of 150 s. A subset of the acquired spectra are shown and arrows indicate the direction of the absorption change over time. The reaction was performed in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. 116 Figure 4.17 Formation of the gem-diamine intermediate and external aldimine following addition of lanthionine to LS. Single wavelength stopped-flow absorbances changes at 320 nm, 406 nm and 428 nm were monophasic (Figure 4.18A/C/E). The observed rate constants for 320 nm and 406 nm were linearly dependent of concentration of D,L-lanthionine (Figure 4.18B/D). The bimolecular rate constants at 406 nm and 320 nm were 107 ± 9 mM-1 s-1 and 87 ± 7 mM-1 s-1, respectively (Table 4.4). The observed rate constants associated with the absorbance change at 428 nm elicited a hyperbolic dependence of D,L-lanthionine concentration (Figure 4.18F), yielding a Kd for D,L-lanthionine of 0.24 ± 0.02 mM and a limiting rate constant (klim) for the formation of the external aldimine of 0.21 ± 0.01 s-1. 117 Table 4.4 Pre-steady state kinetic parameters for LS and S234A with D,L-lanthionine determined following absorption changes at 320 nm, 406 nm and 428 nm. Enzyme Wavelength (nm) kon (mM-1 s-1) Kd (mM) klim (s-1) Wild type 320 87 ± 7 ND ND Wild type 406 107 ± 9 ND ND Wild type 428 ND 0.24 ± 0.02 0.21 ± 0.01 S234A 406 0.046 ± 0.003 ND ND ND = not determined Figure 4.18 Single wavelength stopped-flow absorbance changes at A) 320 nm, C) 406 nm and E) 428 nm. Arrows denote the direction of absorbance changes following increase D,L-118 lanthionine concentration. The resulting observed rate constants were fitted to either linear or hyperbolic equations for B) 320 nm, D) 406 nm and F) 428 nm. All reactions were performed in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. Absorbance changes in the PLP spectra upon rapid mixing of S234A (21 µM) with D,L-lanthionine led to two resolvable kinetic events (Figure 4.19). The photodiode array spectra show that from 1 ms to 132 s there is a decrease in absorbance at λmax at 406 nm and increases at ~470 nm. The peak at 470 nm likely represents formation of the α-aminoacrylate intermediate; thus, unlike the wild type enzyme, S234A appears to catalyse the reversible the β-replacement of L-lanthionine by sulfide. The second slower kinetic phase from 251 s to 1424 follows decay of the λmax at ~470 nm and the α-aminoacrylate intermediate. The spectra after 1425 s was almost identical to the spectra at 1 ms. Single wavelength absorbance traces at 406 nm were fitted to a single exponential equation (Figure 4.20A). The initial observed rate constants (kobs) measured at 406 nm showed a linear dependence on D,L-lanthionine concentration (Figure 4.20B) with kon of 0.046 ± 0.003 mM-1 s-1, 2000-fold slower compared to wild type LS. This suggests that binding of D,L-lanthionine is significantly impeded by the absence of the serine residue. 119 Figure 4.19 Time resolved spectra following rapid mixing of D,L-lanthionine (2 mM) and S234A (21 µM) in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. The spectra were acquired on a log time base over 1424 s and are split into two domains, a subset of the spectra is shown, A) 1 ms to 132 s and B) 251 s to 1424 s. Figure 4.20 A) Single wavelength absorbance trances acquired at 406 nm following rapid mixing of S234A (21 µM) with varying concentration of D,L-lanthionine. The arrow shows the A B 120 direction of the absorbance traces as D,L-lanthionine concentration increases. B) Linear dependence on D,L-lanthionine concentration of kobs measured at 406 nm with S234A. Reactions were performed in 100 mM HEPES-NaOH (pH 7.4) at 10 oC. 4.3.7 Pre-steady-state kinetic analysis with O-acetyl-L-serine Multiwavelength stopped-flow data following rapid mixing of wild type LS (64 µM) and O-acetyl-L-serine (2 mM) were collected over 3 s on a log time base (Figure 4.21, Table 4.5). The spectra show a decrease in absorbance at 406 nm, concomitant with an absorbance increase at 320 and 460 nm. Two isosbestic points appear at 354 and 423 nm. Single wavelength absorbance traces at 406 and 460 nm were biphasic and consequently were fitted to a double exponential equation (Figure 4.22A/B). The initial observed rate constant (kobs1) was linearly dependent on O-acetyl-L-serine concentration and yielded bimolecular rate constants of 2.0 ± 0.1 mM-1 s-1 (406 nm) and 2.2 ± 0.1 mM-1 s-1 (460 nm) (Figure 4.22C). This biomolecular rate constant corresponds to the formation of a mixture of the enol-imine (absorbance maxima at 320 nm) and the keto-enamine (absorbance maxima at 460 nm) forms of the α-aminoacrylate (Figure 4.23).94 There was no detectable accumulation of the external aldimine with O-acetyl-L-serine. The second observed rate constant elicited a hyperbolic dependence on the concentration of O-acetyl-L-serine (Figure 4.22D). The Kd at 406 nm was 1.32 ± 0.06 mM and the klim was 1.49 ± 0.02 s-1. At 460 nm the Kd was determined as 0.89 ± 0.15 mM and the klim was 1.43 ± 0.08 s-1. The Kd reflects the binding constant for O-acetyl-L-serine while the klim corresponds to the limiting rate for formation of the α-aminoacrylate intermediate. 121 Table 4.5 Pre-steady state kinetic parameters for LS and S234A with O-acetyl-L-serine determined following absorption changes at 406 nm and 460 nm. Enzyme Wavelength (nm) kon (mM-1 s-1) Kd (mM) klim (s-1) Wild type 406 2.0 ± 0.1 1.32 ± 0.06 1.49 ± 0.02 Wild type 460 2.2 ± 0.1 0.89 ± 0.15 1.43 ± 0.08 S234A 406 1.0 ± 0.1 2.2 ± 0.8 1.5 ± 0.2 S234A 460 1.6 ± 0.1 1.2 ± 0.2 0.97 ± 0.06 Figure 4.21 A subset of the time resolved spectra following rapid mixing of O-acetyl-L-serine (2 mM) and wild type LS (64 µM) in 100 mM HEPES-NaOH (pH 7.4) at 10 oC measured on a log time base over the course of 3 s. 122 Figure 4.22 Single wavelength absorbance traces following rapid mixing of 64 µM LS with varying concentrations of O-acetyl-L-serine at A) 406 nm and B) 460 nm over a log time scale. Arrows indicate the direction of absorbance as O-acetyl-L-serine concentration increases. C) Linear dependence of kobs1 measured at 406 nm (black) and 460 nm (grey) on O-acetyl-L-serine concentration. D) Hyperbolic dependence of kobs2 measured at 406 nm (black) and 460 nm (grey) on O-acetyl-L-serine concentration. 123 Figure 4.23 The formation of the two tautomeric states (the enol-imine form and the keto-enamine) of the internal aldimine, external aldimine and α-aminoacrylate between LS and O-acetyl-L-serine. Multiwavelength stopped-flow data following rapid mixing of S234A (17 µM) and O-acetyl-L-serine (2 mM) were collected over 9 s on a log time base (Figure 4.24, Table 4.5). Like with wild type LS, the spectra show a decrease in absorbance at 406 nm, concomitant with an absorbance increase at 320 and 460 nm. Two isosbestic points appear at 354 and 423 nm. Single wavelength absorbance traces at 406 and 460 nm were biphasic and consequently were fitted to a double exponential equation (Figure 4.25A/B). The initial observed rate constant 124 (kobs1) was linearly dependent on O-acetyl-L-serine concentration and yielded bimolecular rate constants of 1.0 ± 0.1 mM-1 s-1 (406 nm) and 1.6 ± 0.1 mM-1 s-1 (460 nm) (Figure 4.25C). These bimolecular rate constants correspond to the formation of a mixture of the enol-imine and the keto-enamine forms of the α-aminoacrylate. Like the wild type enzyme, there was no detectable accumulation of the external aldimine with O-acetyl-L-serine. The second observed rate constant elicited a hyperbolic dependence on the concentration of O-acetyl-L-serine (Figure 4.25D, Table 4.5). The Kd at 406 nm was 2.2 ± 0.8 mM and the klim was 1.5 ± 0.2 s-1. At 460 nm the Kd was 1.2 ± 0.2 mM and the klim was 0.97 ± 0.06 s-1. The Kd reflects the binding constant for O-acetyl-L-serine while the klim corresponds to the limiting rate for formation of the α-aminoacrylate intermediate. These data are similar to the data for wild type LS. This suggests that S234 does not impact the mechanism involved in binding of O-acetyl-L-serine and subsequent formation of the α-aminoacrylate. Figure 4.24 A subset of the time resolved spectra following rapid mixing of O-acetyl-L-serine (2 mM) and S234A (17 µM) in 100 mM HEPES-NaOH (pH 7.4) at 10 oC measured on a log time base over the course of 9 s. 125 Figure 4.25 Single wavelength absorbance traces following rapid mixing of 17 µM S234A with varying concentrations of O-acetyl-L-serine at A) 406 nm and B) 460 nm over a log time scale. Arrows indicate the direction of absorbance change as O-acetyl-L-serine concentration increases. C) Linear dependence of kobs1 measured at 406 nm (black) and 460 nm (grey) on O-acetyl-L-serine concentration. D) Hyperbolic dependence of kobs2 measured at 406 nm (black) and 460 nm (grey) on O-acetyl-L-serine concentration. 4.3.8 LS binding to products and product analogues Fluorescence titration with wild type LS was performed to determine the effect of stereochemistry on ligand binding. The equilibrium dissociation constant (Kd) for the binding of multiple ligands (Figure 4.26) to wild type LS was determined through a series of fluorescence titrations as described in section 4.2.9. The change in fluorescence was fitted to a quadratic binding 126 isotherm (Equation 4.3, Figure 4.26). The ligands with tightest binding to LS were meso-2,6-diaminopimelic acid, D,L-lanthionine and D,L-2-aminopimelic acid where the Kd were 1.2 ± 0.1 µM, 2.3 ± 0.1 µM and 3.5 ± 0.5 µM respectively (Table 4.6). The weakest binding ligand was 2(S)-amino-6-boronohexanoic acid, with a Kd of 51.5 ± 3.9 µM and the only ligand not to bind was S-(2-boronoethyl)-L-cysteine. These data show that LS is stereoselective towards the meso-stereoisomer, as there is 15-fold tighter binding for meso-2,6-diaminopimelic acid compared to the L,L-stereoisomer. 127 Table 4.6 The equilibrium dissociation constants for the ten ligands with wild type LS Ligand Structure Kd (µM) 2-(S)-amino-6-boronohexanoic acid 51.5 ± 3.9 S-(2-boronoethyl)-L-cysteine No Binding Observed S-(2-aminoethyl)-L-cysteine 16.6 ± 1.6 D,L-2-aminopimelic acid 3.5 ± 0.5 S-(2-carboxyethyl)-L-cysteine 5.6 ± 0.4 D,L-propargylglycine 29.5 ± 2.7 meso-2,6-diaminopimelic acid 1.2 ± 0.1 L,L-2,6-diaminopimelic acid 16.3 ± 0.8 D,L-2,6-diaminopimelic acid (racemic mixture) 19.8 ± 1.2 D,L-lanthionine (racemic mixture) 2.3 ± 0.1 128 Figure 4.26 Change in fluorescence emission of wild type LS following the addition of A) D,L-lanthionine, B) D,L-2,6-diaminopimelic acid and C) meso-2,6-diaminopimelic acid. All experiments were performed in 100 mM HEPES-NaOH (pH 7.4) at 25 oC. 4.4 Discussion LS is the primary producer of H2S in F. nucleatum through the β-condensation of two molecules of L-cysteine to produce L,L-lanthionine.69 In bacteria, H2S has been shown to provide defence against reactive oxygen and is used as a major cytoprotectant and redox modulator.79, 81 However, H2S has been found to stimulate the proliferation and migration of colorectal cancer cells and is able to inhibit cell apoptosis.177 These show that H2S plays an important role in tumorigenesis and the progression of colorectal cancers. Lanthionine plays a vital role in F. nucleatum as it forms part of a peptide link in the peptidoglycan layer.178 Therefore, LS may make a good target for therapeutics for treating F. nucleatum infections. Characterisation of LS may also be useful in the design of novel synthetic routes for lanthionine and lanthionine analogues. 4.4.1 Preferred reactions of wild type LS Our studies of LS show that like CBS and OASS, LS does show catalytic promiscuity with regards to the type of β-replacement reaction that it catalyses; however, our kinetic studies indicate A B C 129 that it is catalytically tailored to produce lanthionine and H2S through the condensation of two equivalents of L-cysteine. CBS from both humans and yeast can catalyse the formation of lanthionine from two equivalents of L-cysteine. A comparison of the LS steady state parameter with human and yeast CBS indicate that LS is more efficient at this reaction. As presented in Table 4.7, the catalytic efficiency towards L-cysteine was 40-fold and 300-fold higher in LS, compared to yeast and human CBS, respectively. This can be attributed to LS possessing both a higher turnover rate and a lower KM for L-cysteine. However, unlike CBS, LS is less efficient at catalysing the β-replacement of L-cysteine with L-homocysteine. The addition of L-homocysteine to the reaction mixture did not affect the rate of H2S formation. Moreover, HPLC analysis revealed that in the presence of both L-cysteine and L-homocysteine LS was more proficient at catalysing the β-replacement with a second molecule of L-cysteine to produce lanthionine. Table 4.7 Steady-state kinetic parameters for formation of H2S in the presence of L-cysteine by lanthionine synthase, yeast CBS and human CBS. Reactions were performed at 37 oC in 100 mM HEPES-NaOH (pH 7.4). Enzyme kcat (s-1) KM(mM) kcat/KM (s-1 M-1) Wild type LS 16.6 ± 0.3 1.7 ± 0.1 9800 ± 600 Yeast CBS99 8.2 ± 1.2 33 ± 4 250 ± 50 Human CBS99 0.86 ± 0.01 27 ± 4 32 ± 4 LS cannot perform the canonical reaction of CBS, the β-substitution of L-serine with L-homocysteine to form L-cystathionine (Reaction 3 from Figure 4.6). This may be because LS cannot catalyse the α,β-elimination of H2O from the PLP-serine external aldimine. A comparison 130 of the X-ray crystal structures of LS (PDB entry code 5XEN) and yeast CBS (PDB entry code 6C2Q) both in the external aldimine forms with L-serine provide a plausible explanation. In the crystal structure of yeast CBS, the side chains of S82 and Y248 engage in a hydrogen bond with the β-hydroxyl group of serine (Figure 4.26A). These two residues are conserved in the CBS family. In the PLP-aminoacrylate structure of yeast CBS (PDB entry code 6C2Z), S82 and Y248 coordinate to a water molecule (Figure 4.27B) and it is hypothesised that these two residues promote α,β-elimination of H2O from the PLP-serine external aldimine. However, in the crystal structure of LS bound to serine, there is only one hydrogen bond interaction between β-hydroxyl group of serine and S80 (Figure 4.28). Consequently, LS cannot catalyse the α,β-elimination of H2O. Figure 4.27 Crystal structure of yeast CBS in the A) PLP-serine external aldimine state (PDB entry code 6C2Q) and B) α-aminoacrylate state (PDB entry code 6C2Z). 131 Figure 4.28 Crystal structure of LS bound to serine (PDB entry code 5XEN). We showed that LS possess OASS activity as it can catalyse the production of L-cysteine and acetate through the β-substitution of O-acetyl-L-serine with HS-. However, as presented in Table 4.8, the rate of O-acetyl-L-serine turnover was 20-fold and 260-fold lower than for OASS from S. typhimurium and A. thaliana respectively.94, 95 In addition, the kcat/KM for HS- in OASS from S. typhimurium approaches the limits of diffusion (22000 s-1 mM-1). This rate constant is 550-fold higher than kcat/KM for HS- in LS. Consequently, these data suggest HS- canât bind as efficiently to the PLP-aminoacrylate form of LS compared with other OASS. The kcat/KM for O-acetyl-L-serine is 3-fold lower than for HS-; in addition, the klim for the formation of the α-aminoacrylate with O-acetyl-L-serine is comparable to the turnover number measured for L-cysteine formation. These data would suggest that the β-elimination of the acetyl group is likely the rate-determining step. In plant and bacteria, OASS it is part of the main cysteine biosynthetic pathway alongside serine acetyltransferase (SAT) (Figure 4.29).90 SAT catalyses the acylation of L-serine in a reaction with acetyl-CoA forming CoA-SH and O-acetyl-L-serine.95 This acts as the source of O-132 acetyl-L-serine for OASS. However, the F. nucleatum genome lacks the gene coding for SAT.111 Consequently, it does not appear to be physiologically relevant for LS to catalyse the OASS reaction. Figure 4.29 The bacterial biosynthetic pathway for the production of L-cysteine from L-serine utilising SAT and OASS. Table 4.8 Steady-state kinetic parameters for formation of L-cysteine in the presence of O-acetyl-L-serine by lanthionine synthase, OASS from S. typhimurium and OASS from A. thalian. All reactions were performed at 25 oC. The pH for the reaction with wild type LS was 7.4, for OASS from S. typhimurium and A. thaliana it was 7.0. Enzyme kcat (s-1) KM(OAS) (mM) kcat/KM(OAS) (s-1 mM-1) KM(HS-) (mM) kcat/KM(HS-) (s-1 mM-1) Wild type LS 6.3 0.41 15 0.17 40 OASS from S. typhimurium94 130 1 130 0.0006 22000 OASS from A. thaliana95 1780 1.4 1270 0.22 9860 133 4.4.2 Binding of potential inhibitors We established that eight of the nine product analogues tested were able to bind to wild type LS. Amongst the product analogues, the highest binding affinity was seen for meso-2,6-diaminopimelic acid and D,L-2-aminopimelic acid. These are both compounds that lack a thioether group. The weakest binding affinity was seen in 2-(S)-amino-6-boronohexanoic acid and S-(2-boronoethyl)-L-cysteine did not bind. This would suggest that the bulky boronic acid group prevents binding to the LS active site. 2,6-diaminopimelic acid is an analogue of lanthionine, with the only difference being the absence of the thioether. The racemic mixture of D,L-lanthionine had 6-fold tighter binding than the racemic mixture of D,L-2,6-diaminopimelic acid, suggesting that the thioether group is important for product binding. However, meso-2,6-diaminopimelic acid had 15-fold tighter bind than L,L-2,6-diaminopimelic acid. Future inhibitor studies will build on these discoveries to optimise the ligands and find the most effective inhibitors. The effectiveness of these inhibitors at stopping or slowing growth of F. nucleatum in vivo will also need to be assessed. 4.4.3 Structural insight into the role of S234 S234 is part of a loop comprising residues 233MSIG236 that undergoes a substrate-induced conformational shift to coordinate with the lanthionine product as it is formed. The β-hydroxyl group of S234 forms a OH-O hydrogen bond to the carboxylate group of lanthionine (Figure 4.30). A sequence alignment between LS and several OASS enzymes, found that the residues 234SI235 was a unique insertion (Figure 4.31).109, 179-181 By converting S234 to an alanine there was a 30% decrease in catalytic efficiency in the production of lanthionine from β-replacement of L-cysteine 134 with a second molecule of L-cysteine. Moreover, the HPLC data showed that the S234A variant was less selective towards the production of lanthionine over cystathionine than wild type LS. These data suggest that the 234SI235 insertion plays a vital role in coordinating binding of the second L-cysteine molecule, stabilising the formation of lanthionine and preventing other potential substrates, such as L-homocysteine from binding. It also prevents the reverse reaction from occurring, the β-replacement of L-lanthionine by sulfide to produce L-cysteine. Figure 4.30 Crystal structure of the active site of wild type LS with A) no ligand (PDB entry code 5XEO) and B) L-lanthionine (PDB entry code 5XEM). This figure shows the hydrogen bond formed between S234 and the carboxylate group of L-lanthionine. A B 135 Figure 4.31 Sequence alignment performed using NCBI BLAST between F. nucleatum LS, OASS from A. thaliana (UniProtKB Q8RE94) and OASS from S. typhimurium (UniProtKB P47998). The unique 234SI235 insertion is highlighted in red. 4.5 Concluding remarks We have determined that the physiological reaction of LS from F. nucleatum is the production of lanthionine and H2S from two equivalents of L-cysteine. Kinetic and HPLC analysis showed a preference to produce lanthionine from L-cysteine, even in the presence of L-homocysteine. This would be physiologically significant as meso-lanthionine is a vital component of the F. nucleatum peptidoglycan layer. LS did show some OASS activity; however, the catalytic turnover was >30-fold lower than OASS from other sources. In addition, this reaction does not appear to be physiologically necessary for F. nucleatum as the bacteriumâs genome lacks the gene coding for SAT, the enzyme responsible for the production of O-acetyl-L-serine used by OASS. S234 likely plays an important role in the production of lanthionine by LS by forming a hydrogen bond with the carboxylate group. Replacement of S234 with alanine resulted in a 2000-fold lower value for kon for D,L-lanthionine. This substitution also resulted in greater catalytic promiscuity in LS, with there being approximately equal production of L-lanthionine and cystathionine when LS was incubated with both L-cysteine and L-homocysteine. In addition, the substitution enabled the β-replacement of L-lanthionine by sulfide to take place, a reaction that did F. nucleatum LS 217 LLSKGYIGPHKIQGMGMSIGGIPAVYDGSLADD 249 S. typhimurium OASS 216 AGEEIKPGPHKIQGIGA--GFIPGNLDLKLIDK 246 A. thaliana OASS 212 ILSGGKPGPHKIQGIGA--GFIPSVLNVDLIDE 242 136 not occur in wild type LS. These data show that the 233MSIG236 loop stabilises the formation of lanthionine. Consequently, wild type LS has a preference for the production of lanthionine through the condensation of two equivalents of L-cysteine. We have determined that LS is likely the main source of L-lanthionine produced by F. nucleatum, we anticipate that there is likely a racemase or epimerase that converts L-lanthionine to meso-lanthionine. This will make LS a good drug target due to the vital role of meso-lanthionine in the F. nucleatum peptidoglycan layer. Preventing the production of meso-lanthionine may weaken the bacterial membrane and result in death of the bacterium. In addition, this research may provide the basis for developing novel synthetic routes for lanthionine and lanthionine analogues. Finally, we established that there are multiple product analogues that can bind to the LS active site and may act as inhibitors. The next stage of this research will be a continuation of finding and testing potential inhibitors of LS. 137 5. Ornithine Decarboxylase/Arginase 5.1 Chapter Summary Ornithine decarboxylase/ arginase (ODA) is a bifunctional enzyme unique to F. nucleatum. The N-terminal half (residues 1 â 500) comprises the ornithine decarboxylase component and the C-terminal half (residues 500 -700) comprises the arginase component. Ornithine decarboxylase (ODC) is a PLP-dependent enzyme that converts L-ornithine to putrescine. The ODC component of ODA is a group I PLP-dependent decarboxylase and sequence alignment of ODA with other members of this group identified three putative active site residues (H119, D199 and K230) in the PLP binding pocket. The catalytic importance of these residues was confirmed through site-directed mutagenesis. H119 was replaced with a glutamine (H119Q) as the side chain is a similar size to the imidazole group, but it is unable to donate a proton during catalysis. D199 was changed to alanine to remove the negative charge associated with the carboxylate group. Finally, K230 was converted to methionine (K230M) as the side chain is of similar size; but the absence of a primary amine will prevent formation of an imine linkage with the PLP. These three active site variants (H119Q, D199A and K230M) did not show detectable activity even when supplemented with PLP. Arginases catalyse the conversion of L-arginine to L-ornithine and urea. Arginases incorporate two Mn2+ ions though a conserved network of active site residues. The divalent metals coordinate to a hydroxide ion that initiates nucleophilic attack on the guanidium group of L-arginine. Sequence alignment of the arginase component of ODA with arginases from a variety of organisms revealed two unusual amino acid substitutions in the conserved metal coordinating shell of ODA. Metal analysis by ICP-MS showed that the wild type enzyme does not bind Mn2+. 138 However, restoration of ODA metal coordinating shell through site directed mutagenesis resulted in an increase in Mn2+content, but a modest decrease in catalytic activity. The kcat and KM(L-arg) for the combined arginase and ornithine decarboxylase activity of wild type ODA was determined to be 3.0 s-1 and 10 mM respectively. When the individual components of ODA (arginase and ODC) were sub-cloned and expressed, the arginase component was found to be inactive on its own, while the kcat and KM(L-orn) for the ODC component were found to be 6.7 s-1 and 1.8 mM respectively. Gel filtration revealed that wild-type ODA formed an octamer in solution, while the arginase component forms a tetramer and a dimer, and the ODC component is dimeric. The activity of ODA was not affected in the presence of known inhibitors of human arginase or an inhibitor of a structurally related bacterial PLP-dependent decarboxylases. These data further suggest that ODA possesses a distinct active site that warrants further investigation through structural analysis by X-ray crystallography. A novel active site may also prove promising for the development of novel therapeutics for the treatment of F. nucleatum infections. 5.2 Materials and Methods 5.2.1 Materials Q5® High Fidelity DNA polymerase and restriction enzymes were from New England Biolabs (Ipswich, MA, USA). Oligonucleotides were purchased from Integrated DNA Technologies (Coralville, IA, USA). F. nucleatum genomic DNA was from the American Tissue Culture Collection (Manassas, VA, USA). E. coli XL1-Blue competent cells were from Agilent, (Santa Clara, CA, USA). The DCâ¢ Protein Assay was from Bio-Rad (Hercules, CA, USA) The E.Z.N.A plasmid mini kit was from VWR (Radnor, PA, USA). 139 5.2.2 Cloning gene for ODA into pET15b plasmid The gene coding for ODA (Fn0501) was amplified from the genomic DNA of F. nucleatum strain ATCC 25586 with Q5® High Fidelity DNA polymerase using primers 5â-AGA ATT CCA TAT GTC TAA ATT AGA CCA AAA TAA G -3â (forward primer) and 5â- GTG GAT CCT TAA TAA TCT GGG TTC ATC ATA TAT TG- 3â (reverse primer). The primers incorporated NdeI (forward primer) and BamHI (reverse primer) restriction sites (shown in bold).The 50 µL PCR reaction mixture contained Q5® High Fidelity DNA polymerase (0.02 U/µL), Q5® Reaction Buffer, dNTP (0.2 mM), F. nucleatum genomic DNA (1 ng/mL), forward primer (2.5 ng/mL) and reverse primer (2.5 ng/mL). The gene was amplified by cycling the reaction mixture 30 times through the following temperatures: 95 oC (45 s), 55 oC (30 s) and 72 oC (360 s) following an incubation at 95 oC for 2 minutes. The PCR product, cleaved with NdeI and BamHI, was then ligated into pET15b plasmid digested with the same restriction enzymes. The ligated product was transformed into E. coli XL1-Blue and then the cell mixture was plated onto LB-agar plates supplemented with ampicillin (100 µg/mL). The cloning strategy incorporated a hexahistidine tag onto the N-terminus of the protein to aid in protein purification. Plasmid from a transformed cell was isolated using an E.Z.N.A plasmid mini kit.148 The successful cloning of the gene was confirmed by DNA sequencing by the Sequencing and Bioinformatics Consortium at UBC Vancouver. 5.2.3 Purification and expression of wild-type ODA The plasmid harbouring the cDNA for ODA (named petODA) was transformed into E. coli strain Rosetta(DE3)pLysS for protein expression. A starter culture was prepared by inoculating 140 200 mL of LB media, supplemented with ampicillin (100 µg/mL) and chloramphenicol (34 µg/mL), with a single colony generated from a transformed cell. The starter culture was incubated for approximately 16 hours at 37 oC. Aliquots (10 mL) of the starter culture were added to 500 mL of Terrific Broth, supplemented with ampicillin (100 µg/mL) and chloramphenicol (34 µg/mL). The culture was incubated at 30 oC at 200 rpm until the optical density at 600 nm was ~0.6. Isopropyl β-D-1-thiogalactopyranoside (ITPG) was added to cell cultures to a final concentration of 0.2 mM. The cultures continued to incubate for approximately 16 hours at 25 oC at 200 rpm. Cell cultures were centrifuged at 6371g for 15 min, and cell pellets were pooled together and stored at -80 oC until protein isolation. Cells (30 g) were resuspended in 200 mL of 50 mM Tris-HCl, pH 7.5 containing protease inhibitors phenylmethylsulfonyl fluoride (1.0 mM) and benzamidine (0.1 mM) and MnCl2 (1 mM). The cell suspension was sonicated on ice at 32 % power for 8 s at 1 min intervals for 1 hour. The suspension was centrifuged at 39191g for 60 min at 4 oC. The pooled supernatants were applied to a Ni-nitrilotriacetate (NTA) column (16 Ã 30 mm) (MilliporeSigma, MA, USA) equilibrated with 50 mM Tris-HCl, pH 7.5 and 0.02 M imidazole. Protein was eluted from the column using a step gradient: 5 CV of 100% Buffer A (50 mM Tris-HCl, pH 7.5 with 0.02 M imidazole) and 0% Buffer B (50 mM Tris-HCl, pH 7.5 and 0.5 M imidazole), 5 CV of 4 % Buffer B, 5 CV of 60 % Buffer B and 5 CV of 100 % Buffer B. Elution of protein was monitored at 280 nm. Protein eluted at 60% buffer B was pooled and dialysed against 4 L of 50 mM Tris-HCl, pH 7.5 for 16 hr. The dialysate was concentrated to ~10 mL using a centricon (Vivaspin 20, GE Healthcare, IL, USA) with a 30 kDa MW cut-off filter. The protein was then applied to a Q-sepharose HP column (26 Ã 140 mm) equilibrated with 50 mM Tris-HCl, pH 7.5. The protein was eluted using a linear gradient from 100 % Buffer C (50 mM Tris/HCl, pH 7.5) to 100 % Buffer D (50 mM Tris/HCl), pH 7.5, 0.5 M 141 NaCl over 8 CV. Fractions containing protein were pooled and concentrated to 10 â 15 mL. SDS-PAGE gel was performed to assess the purity of the protein. Stocks of the protein containing 20% glycerol were prepared and stored at -80 oC. The concentration of the protein was determined through a Lowry assay and by measuring the absorbance at 277 nm using an extinction coefficient of 33.5 mM-1 cm-1. ODA isolated for ICP-MS analysis, followed the same purification strategy except that the recombinant protein was expressed in minimal media following a published protocol for metal analysis of formiminoglutamase from Trypanosoma cruzi.182 The minimal media contained 1Ã M9 salts, 18 mM D-(+)glucose, 1% casamino acids, 2 mM MgSO4 and 100 µM CaCl2 in a total volume of 500 mL. The 500 mL cultures were also supplemented with 100 μg/mL of ampicillin and 35 μg/mL of chloramphenicol. Following the Ni-NTA chromatographic step, ODA was dialysed for 16 hr against 4 L of 50 mM Tris-HCl, pH 7.5 in the presence of 500 units of thrombin (GE Biosciences) to cleave the N-terminal hexahistidine tag. The dialysate was applied to Ni-NTA column (16 x 30 mm) equilibrated with 50 mM Tris/HCl, 0.02 M imidazole, pH 7.5. The column was washed with 3 CV of the same buffer and the flow-through containing the cleaved ODA was collected. These protein stocks were stored at -80 oC but did not have glycerol added. 5.2.4 Variants produced by site-directed mutagenesis The following variants of ODA (H119Q, D199A K230M, S609D, S609A, N713D and the double mutant S609D/N713D) were created by site directed mutagenesis using PfuTurbo DNA Polymerase (Agilent, CA, USA) using the QuikChange method.169 The primers used for each variant are shown in Table 5.1. 142 Table 5.1 Sequence of oligonucleotides used in site directed mutagenesis to produce the ODA variants. Bold type highlights the sequence of the mutated codon. Primer Sequence Forward K230M 5â GT ACT CAA AGT ACA CAT ATG ATT TTA GGT GCT ATG Reverse K230M 5â CAT AGC ACC TAA AAT CAT ATG TGT ACT TTG AGT AC Forward H119Q 5â GTA CCA AGA AAT GTT CAA AAA TCA GTT TCA GCA GG Reverse H119Q 5â CC TGC TGA AAC TGA TTT TTG AAC ATT TCT TGG TAC Forward D199A 5â GAT ATA CCT CTT ATT GTA GCT GAA GCT CAT GGA CCA C Reverse D199A 5â G TGG TCC ATG AGC TTC AGC TAC AAT AAG AGG TAT ATC Forward S609A 5â GGA GTT TTA TGG ATA GCT GCT CAT GGA GAT ATG Reverse S609A 5â CAT ATC TCC ATG AGC AGC TAT CCA TAA AAC TCC Forward S609D 5â GGA GTT TTA TGG ATA GAT GCT CAT GGA GAT ATG Reverse S609D 5â CAT ATC TCC ATG AGC ATC TAT CCA TAA AAC TCC Forward N713D 5â CAT ATC AGT ATA GAT ATG GAT GTA TTT GAC CCA GAA ATT GC Reverse N713D 5â GC AAT TTC TGG GTC AAA TAC ATC CAT ATC TAT ACT GAT ATG The presence of the mutation in ODA and absence of PCR-induced errors was confirmed by DNA sequencing. These variants were expressed and isolated following the same protocol for wild type ODA. The S609D/N713D variant was also expressed using the minimal media protocol for ICP-MS analysis. 5.2.5 Sub-cloning ODC and arginase component parts Plasmid constructs for the expression of the ODC component (residues 1 to 329) (ODC/ODA) and the arginase component (residues 462 to 760; Arg/ODA) of ODA were generated by PCR amplifying the appropriate coding sequences from pETODA vector using PfuTurbo DNA polymerase. The oligonucleotides for the ODC component, 5â-AGA ATT CCA TAT GTC TAA ATT AGA CAA AAT AAG-3â (forward primer) and 5â- ACG GAT CCT TCA GTG TAT AAG 143 TAA ATA GC -3â (reverse primer), and for the arginase component 5â- AGA ATT CCA TAT GAA AAA TGT TCT TAT TGG AGG-3â (forward primer) and 5â GCG GAT CCT TAA TAA TCT GGG TTC ATC-3â (reverse primer), were designed to include NdeI and BamHI restriction sites to facilitate cloning into pET15b. The conditions used for PCR were denaturation at 95 oC for 2 min, followed by 30 cycles at 95 oC for 30 s, 55 oC for 30 s and 72 oC for 3 min. The PCR products were digested with NdeI and BamHI and sub-cloned into pET15b. The constructs were sequenced to ensure no PCR-induced error had been introduced. The expression plasmids were subsequently transformed into E. coli strain Rosetta(DE3)pLysS for the production of the recombinant protein. The genes were then expressed and purified via the same method as wild type ODA. 5.2.6 Measuring the rate of CO2 formation with L-arginine and L-ornithine ODC activity of full length ODA, the isolated ODC component and the H119Q, D119A and K230M ODA variants was determined using a coupled spectrophotometric assay developed by Smithson et al (2010).183 The assay monitors the production of CO2 by ODC in the presence of L-ornithine. Under basic buffering conditions CO2 converts to bicarbonate, which in the presence of phosphoenolpyruvate carboxylase is added to phosphoenolpyruvate to form oxaloacetate. Malate dehydrogenase then reduces oxaloacetate to from malate in the presence of NADH. The amount of NADH converted to NAD+ is determined by following the absorbance change at 340 nm using an extinction coefficient of 6.22 mM-1 cm-1. This assay was performed at 25 °C in a glove box (Belle Technology, Weymouth, UK) maintained under 99.9% nitrogen gas to drastically reduce atmospheric CO2 interfering with the reaction. The reaction buffer, 75 mM Tris-HCl containing 25 mM NaCl and 8.0 mM MgSO4 was 144 purged for two hours with N2 and then placed in the glove box for 16 hours to equilibrate. Powdered forms of phosphoenolpyruvate, NADH, L-ornithine and phosphoenolpyruvate carboxylase were introduced to the glove box and dissolved in the N2 purged reaction buffer. Small volumes (300 µL) of malate dehydrogenase and ODA were brought into the anaerobic environment and allowed to equilibrate for 30 mins. The 1 mL reaction mixture contained 100 µM phosphoenolpyruvate, 1.0 units phosphoenolpyruvate carboxylase, 4.0 units of malate dehydrogenase, 0.2 mM NADH and variable concentrations of L-ornithine (0.25 to 10 mM). The reaction was initiated with the addition of 20 nM ODA. The oxidation of NADH was observed by monitoring the decrease in absorbance at 340 nm (ÎÎµ = 6.22 mM-1 cm-1) using a UV/vis spectrophotometer (Lambda 265, Perkin Elmer, MA, USA) over the course of 8 mins at 25 oC. The initial rate of NADH oxidation was plotted against L-ornithine concentration and fitted to the Michaelis-Menten equation (equation 5.1). Equation 5.1: ð£ =ðððð¥[ð]ð¾ð+[ð] The combined activity of ODA (i.e. the activity of the arginase and ornithine decarboxylase) was determined with the same coupled spectrophotometric assay except that variable concentrations of L-arginine (0.5 to 100 mM) were used in place of L-ornithine. The initial rate of NADH oxidation was plotted against L-arginine concentration and fitted to the Michaelis-Menten equation (equation 5.1). This kinetic assay was performed for wild type ODA, all ODA variants (except D199A) and the isolated arginase domain. Kinetic assays of the ODA H119Q and K230M variants and the isolated arginase component also contained an excess of the isolated ODC (0.1 µM) to compensate for the fact that the ODC domain was either removed or 145 inactivated through mutagenesis. The coupled assay was also used to measure the activity of wild type ODA in the presence of putative inhibitors of the ODC active site (2-(difluoromethyl)ornithine), (2-(difluoromethyl)arginine) and the arginase active site (S-(2-boronoethyl)-L-cysteine and 2(S)-amino-6-boronohexanoic acid).125, 144, 184 5.2.7 Rate of urea formation by ODA and the isolated arginase component The arginase activity of ODA was initially measured through a coupled spectroscopic assay developed by Ozer et al (1985).185 The assay measures the rate of urea formation in the presence of L-arginine using urease and glutamate dehydrogenase as coupling enzymes. Urea produced by arginase undergoes hydrolysis to ammonia and CO2 by urease. Ammonia is consumed by glutamate dehydrogenase. This reaction uses NADPH as a source of electrons and hence the rate of the reaction can be followed through the oxidation of the pyridine nucleotide. The 1 mL reaction mixture contained 40.0 units of urease, 3 units of glutamate dehydrogenase, 1.0 mM α-ketoglutarate, 0.1 mM NADPH and variable concentrations of L-arginine (0.5 to 20 mM). The reaction was initialised by the addition of 400 nM ODA. Oxidation of NADPH was observed by monitoring the decrease in absorbance at 340 nm (ÎÎµ = 6.22 mM-1 cm-1) using a UV/vis spectrophotometer (Lambda 365, Perkin Elmer, MA, USA) over the course of 8 mins at 25 oC. The initial rate of NADPH oxidation was plotted against L-arginine concentration and fitted to the Michaelis-Menten equation (equation 5.1). 5.2.8 Determining quaternary structure via gel filtration chromatography The quaternary structure of ODA, ODC/ODA and Arg/ODA were determined through size exclusion chromatography using a HiLoad Superdex 200 PG (1.6 Ã 60 cm) column (GE Healthcare) equilibrated with 50 mM potassium phosphate pH 7.2, 150 mM NaCl. Aliquots (500 146 µL) of protein standards: thyroglobulin, ferritin, aldolase, conalbumin, ovalbumin and blue dextran 2000 (GE Healthcare) at 5 mg/mL, were loaded onto the column and eluted at a flow rate of 1.0 mL/min using the equilibration buffer. The volume the protein was eluted (Ve) was measured and the proportion of pores available to the molecule (Kav) was determined using equation 5.2. Equation 5.2: ðð = ðð + ð¾ðð£(ðð¡ â ðð) where Vo is the void volume (elution volume of blue dextran) and Vt is the total volume. Kav was fitted against molecular weight to produce a calibration curve. A 500 uL aliquot of ODA was then loaded onto the column and eluted with an isocratic gradient to find Ve for ODA and equation 5.2 was used to find Kav for ODA. The molecular weight of ODA was determined from the calibration curve. 5.2.9 Divalent metal content of ODA by ICP-MS The Mn2+ content of wild type ODA and the S609D/N713D mutant was determined via ICP-MS with the assistance of Dr. Mark Button at the FiLTER Laboratory, UBC Okanagan. For metal analysis, recombinant ODA was expressed in minimal media and the hexahistidine tag was removed (as described in section 5.2.3). The protein was concentrated to 5 mL using a centricon (Vivaspin 20, GE Healthcare, IL, USA) with a 30 kDa MW cut-off filter. Five mL of the flow through was collected from the centricon and was used as a blank for ICP-MS. The protein samples were run on an 8900 ICP-QQQ-MS (Agilent, CA, USA) with an ASX520 autosampler (Cetac, AB, Canada). The instrument was optimised for maximum sensitivity by maintaining the oxide and divalent ion ratios below 147 2%. The instrument was calibrated with a four-point curve ranging from 0 â 100 ppb of multi element standard. The four elemental internal standards were Sc, Y, In and Tb. 5.3 Results 5.3.1 Properties of purified ODA and the isolated ODC and arginase components The cDNA encoding ODA, ODC/ODA and Arg/ODA were successfully inserted into the pET15b vector. All three proteins and the active site variants were recombinantly expressed in E. coli and purified by affinity and ion exchange chromatography. Approximately 50 mg of ODA, 40 mg of ODC/ODA and 20 mg of Arg/ODA was obtained from 3 L of bacterial culture. The homogeneity and approximate size of the proteins was confirmed by SDS-PAGE (Figure 5.1). The apparent molecular masses of ODA (~90 kDa), ODC/ODA (~53 kDa) and Arg/ODA (~30 kDa) are similar to their expected molecular weights of 87.4, 54.2 and 30.3 kDa. 148 Figure 5.1 A) SDS 8% polyacrylamide gel showing the purification of wild type ODA. Lane 1, BLUeye pre-stained protein ladder (Sigma); Lane 2, crude extract; Lane 3, flow through from the Ni-NTA column; Lane 4, protein eluted from Ni-NTA column; Lane 5, protein eluted from Q-sepharose column B) SDS 10% polyacrylamide gel showing wild type ODA, the isolated ODC component and the isolated arginase component following purification by affinity chromatography and anionic exchange chromatography. Lane 1, protein molecular weight marker; Lane 2, purified wild type ODA; Lane 3, purified ODC domain; Lane 4 purified arginase domain. Lane 1 Lane 2 Lane 3 Lane 4 Lane 5 A Lane 1 Lane 2 Lane 3 Lane 4 B 149 Wild-type ODA eluted from the size exclusion chromatograph with a total molecular weight of 703 kDa. These data suggest that ODA, with a monomeric mass of ~87 kDa forms an octamer in solution (Figure 5.2, Table 5.2). The isolated ODC domain eluted as a dimer, while the isolated arginase component eluted as a mixture of two different quaternary conformations: a tetramer (57%) and a dimer (43%). Table 5.2 Number of subunits formed by ODA and the isolated arginase and ornithine decarboxylase components determined by size-exclusion chromatography. Enzyme Total Mr (kDa) Mw of subunit (kDa) Number of subunits Quaternary structure Wild type ODA 703 87.4 8.04 Octamer Arg/ODA 126 (57%), 59 (43%) 30.3 4.16 (57%), 1.95 (43%) Tetramer (57%), Dimer (43%) ODC/ODA 107 54.2 1.97 Dimer Figure 5.2 Calibration curve determined from gel filtration showing how the quaternary structure of ODA and the separated component parts were determined. Grey points represent standards and black represents ODA and the separated components. 150 Purified wild type ODA and the isolated ODC component were a bright yellow colour due to the presence of the PLP cofactor. The isolated arginase component was colourless. The UV-visible absorbance spectra of ODA and the isolated ODC component show a major peak at 420 nm and a minor peak at 320 nm (Figure 5.3). The prominent peak at 420 nm indicates that the primary tautomeric form of the PLP cofactor in the internal aldimine state is the keto-enamine with the imine nitrogen protonated and hydrogen-bonded to the 3â²-oxygen of the PLP cofactor. The addition of L-ornithine to full length ODA caused a decrease in the absorption peak at 420 nm and a slight red-shift to an absorption maximum of 418 nm. There was also a slight absorbance increase at 320 nm. These absorbance shifts represent conversion to the external aldimine with putrescine and the increase in absorbance at 320 nm indicates that the tautomeric equilibrium of the PLP cofactor is shifted to the enolimine state. The addition of L-arginine or L-lysine to purified ODA did not cause a shift in the PLP absorption profile, signifying that the enzyme is specific for L-ornithine (Figure 5.4). The K230M and H119Q variants were in their apo-form as they did not show a PLP absorbance peak (Figure 5.5). The 420 nm absorbance peak of D119A was half of that of wild type ODA revealing that the mutation results in weaker cofactor binding. 151 Figure 5.3 UV/visible spectra of wild type ODA (black line) the isolated ODC component (dark grey dashes) and the isolated arginase component (light grey dots). Figure 5.4 UV/visible spectra of wild type ODA in the presence of no substrate (black line), L-ornithine (black dots), L-arginine (grey dots) and L-lysine (grey dashes). 152 Figure 5.5 UV/visible spectra of wild type ODA (black line), D119A (grey dots), H119Q (grey dashes) and K230M (black dots). 5.3.2 Steady state kinetic analysis of wild type ODA and the isolated ODC component with L-ornithine as the substrate Initial velocity data gathered from the kinetic assays were fit to the Michaelis-Menten equation (Figure 5.6) to determine the turnover rate (kcat), the Michaelis constant (KM(L-orn)) and the catalytic efficiency (kcat/KM(L-orn)). Wild type ODA gave a turnover rate of 12.2 ± 0.3 s-1 (Table 5.3) and a KM(L-orn) of 1.8 ± 0.1 mM. All three ODC active site variants (H119Q, K230M, D199A) did not show detectable activity even with the addition of exogenous PLP (100 nM) to the reaction mixture. The isolated ODC component elicited a similar KM(L-orn) (1.7 ± 0.2 mM) as full length wild type ODA, but the kcat (6.7 ± 0.3 s-1) was ~2-fold lower. 153 Figure 5.6 Michaelis-Menten plot of the production of CO2 in the presence of L-ornithine and wild type ODA (black line) and ODC/ODA (dashed grey). Table 5.3 Kinetic parameters for the ODA and ODC/ODA in the presence of L-ornithine. Enzyme kcat (s-1) KM(L-orn) (mM) kcat/KM (mM-1 s-1) Wild type ODA 12.2 ± 0.3 1.8 ± 0.1 6.8 ± 0.5 ODC/ODA 6.7 ± 0.3 1.7 ± 0.2 3.9 ± 0.6 5.3.3 Kinetic analysis of wild type ODA and the isolated arginase component with L-arginine as the substrate Initially, we attempted to measure the arginase activity (the hydrolysis of L-arginine to L-ornithine and urea) of full-length ODA and the isolated arginase component through a spectroscopic assay that couples the rate of urea formation with the oxidation of NADPH 154 (described in section 5.2.6). This assay did not work as intended as the arginase turnover rate was ~10-fold lower than the turnover rate determined for the full-length enzyme using the CO2 coupled assay described below. The combined activity of full-length wild type ODA (the arginase and ornithine decarboxylase activities) was determined by measuring the amount of CO2 produced using a couple spectrophotometric assay in the presence of L-arginine. The initial velocity data generated at increasing concentrations of L-arginine were fitted to the Michaelis Menten equation giving a kcat of 3.0 ± 0.1 s-1 and a KM(L-arg) of 10 ± 1 mM (Figure 5.7, Table 5.4). The S609D and N713D arginase active site variants present in full length ODA, elicited a modest decrease in catalytic activity with kcat values of 2.5 ± 0.1 s-1 and 2.6 ± 0.1 s-1 respectively. S609A and the double variant, elicited a ~2-fold reduction in activity with kcat values of 1.4 ± 0.1 s-1 and 1.7 ± 0.1 s-1 respectively. The KM(L-arg) for all the arginase active site variants showed a slight decrease from that of wild type ODA. Notably, the isolated arginase component and the K230M and H119Q variants present in full length ODA did not show CO2 production even in the presence of excess amount of the individual ODC component. The lack of CO2 production in the K230M and H119Q variants suggest that the L-ornithine generated in the functional arginase component of the variant does not diffuse to the functional ODC component present in the reaction mixture. 155 Figure 5.7: Michaelis-Menten plot of the production of CO2 in the presence of L-arginine and wild type ODA (black line), S609A (light grey line), S609D (dark grey dotted), N713D (light grey dashes) or S609D/N713D (black dashes) Table 5.4: Steady-state kinetic parameters for the ODA variants in the presence of L-arginine. Enzyme kcat (s-1) KM(L-arg) (mM) kcat/KM (M-1 s-1) Wild type ODA 3.0 ± 0.1 10 ± 1 310 ± 30 S609D 2.5 ± 0.1 12 ± 1 210 ± 20 S609A 1.7 ± 0.1 18 ± 3 90 ± 20 N713D 2.6 ± 0.1 12 ± 1 220 ± 20 S609D/N713D 1.4 ± 0.1 14 ± 2 100 ± 20 K230Ma - - - H119Qa - - - Arg/ODAa - - - aNo activity was detected even in the presence of ODC/ODA 156 5.3.4 No inhibition observed of wild type ODA The effects of two known inhibitors of human arginase and one inhibitor of a group III PLP-dependent decarboxylase on the activity of wild type ODA was examined. The CO2 coupled spectroscopic assay was used to measure the effects of the inhibitor. S-(2-boronoethyl)-L-cysteine and 2(S)-amino-6-boronohexanoic acid, two potent inhibitors of human arginase, did not show an effect on ODA activity when L-arginine was present at 10 mM and the inhibitors were at a concentration of 10 mM.144 Likewise, DL-α-(difluoromethyl)arginine, an irreversible inhibitor of bacterial arginine decarboxylases, did not inhibit ODA activity at a L-ornithine concentration of 1.5 mM and inhibitor concentrations of 10 mM. 5.3.5 Metal Analysis of ODA and the S609D/N713D variant ICP-MS analysis revealed that no elemental Mn2+ was bound to wild type ODA. However, the S609D/N713D variant, which was created to resemble the conserved divalent metal chelating active site of arginase was found to have 1.3 μM Mn2+ in a 15 μM solution of protein, which is ~20-fold lower than what would be anticipated for a protein binding two Mn2+ cations. It appears from these data that the restoration of the ODA active site to conserved inner sphere coordination shell does partially improve Mn2+ binding. 5.4 Discussion 5.4.1 Advantages of bifunctional enzyme to F. nucleatum ODA is a bifunctional enzyme that appears to be unique to Fusobacterium. A protein-protein BLAST analysis of the ODA primary sequence against non-redundant protein sequence database only retrieves homologous protein sequences from other members of the Fusbacteria genera (i.e. Fusobacterium hwasookii and Fusobacterium canifelium). 157 There are three common types of bifunctional enzymes: 1) those that catalyse consecutive reactions and exhibit substrate channelling, 2) those that catalyse consecutive reactions but donât exhibit substrate channelling and 3) those catalysing opposing reactions.186 There are numerous benefits to substrate channelling, these include: isolating intermediates from competing reactions, protecting unstable intermediates, removing unfavourable energetics associated with desolating substrates and enabling more rapid transfer of substrates between active sites.187 Consequently, it is often evolutionary advantageous for an organism to incorporate bifunctional enzymes. Since ODA catalyses two reactions that are consecutive within the metabolic pathway, it must belong to one of the first of these two categories. It may be beneficial for F. nucleatum to fuse together arginase and ornithine decarboxylase as it will enable more efficient conversion of L-arginine to putrescine and urea. Putrescine is a vital compound in the synthesis of the polyamines spermidine and spermine in many organisms.119 Analysis of the distribution of polyamines in F. nucleatum subsp nucleatum (strain JCM 8532) revealed that the most abundant polyamine was the tetraamine spermine, with a small fraction of the triamine spermidine; putrescene was not detected.188 However, analysis of the F. nucleatum genome reveals that it does not encode a gene product capable of spermidine or spermine synthesis.111 This suggests that F. nucleatum incorporates polyamines from its surrounding environment and doesn`t produce them itself. Indeed, F. nucleatum does possess the membrane bound components (pot-A, -B and -C) of the spermidine/putrescine ATP-driven transport system seen in bacteria such as E. coli.111, 189 There is a gene (FN0129) found in the genome for F. nucleatum that may encode for urease. Hence, the urea produced from ODA is likely converted to ammonium and CO2 though the action of this urease. The production of NH4+ can protect the bacterium from acidic environments present in the 158 mouth.190 Thus, urea formation may be a very important biological role of ODA for F. nucleatum survival. 5.4.2 Comparison of kinetic activity The kcat of the combined activity of ODA (3.0 s-1) is 4-fold lower than the kcat measured just for the ornithine decarboxylase activity of the full-length enzyme (12.2 s-1). These data suggest that either substrate transfer between the active sites or the arginase catalytic activity is the rate determining step of catalysis. Interestingly, the arginases typically exhibit higher turnover number compared to ornithine decarboxylases. For example, the kcat of rat liver arginase I and human arginase II are 250 s-1 and 203 s-1, respectively.138, 191 In contrast, the kcat of ornithine decarboxylases from Trypanosoma brucei ODC (7.0 s-1) and human (5.3 s-1), is significantly slower.183 This might suggest that transfer of the substrate is slower, but direct measure of the arginase activity of ODA is required before any conclusions can be drawn. The value of KM(L-arg) (10 mM) for ODA was comparable to that of rat liver arginase I (1.4 mM) and human type II arginase (4.8 mM). The kcat for the isolated ODC component was ~2-fold lower compared to full length ODA indicating that the presence of the arginase component increases the ornithine decarboxylase activity of the enzyme. There was no change to the KM between ODA and ODC/ODA suggesting that the arginase component did not affect L-ornithine binding. 5.4.3 Comparison of quaternary structures The isolated arginase component forms dimer and tetramer complexes in solution. While most mammalian arginase are trimeric with C3 symmetry, bacterial and some protozoa arginases are monomeric, dimeric or hexametric with D3 symmetry.133, 192, 193 The isolated ODC component 159 forms a dimer. This contrasts with another type I PLP dependent decarboxylase, ODC from Lactobacillus sp. 30a, which forms a dodecamer (oligomer of six dimers).117The oligomerisation of the dimers into a dodecamer is due to the presence of a 107 residue N-terminal domain called the wing domain (Figure 5.8). The wing domain comprises one β-sheet and four flanking α-helices. This region is isolated from the rest of the subunit and points into the central cavity region, where it can interact with neighbouring subunits to stabilise the formation of the dodecamer. The amino acid of sequence of ODA does not include this wing domain; thus, it is not surprising that ODA does not oligomerise to a dodecamer. Figure 5.8 The crystal structure of the dimerised from of ODC from Lactobacillus sp. 30a (PDB entry code 1ORD) showing the wing domain (pink) projecting from the rest of the protein. 160 5.4.4 Ornithine decarboxylase active site residues successfully identified All three ODC active site variants (H119Q, D199A and K230M) resulted in undetectable enzymatic activity. The lack of yellow pigmentation in the isolated H119Q and K230M variants indicates a disruption in PLP binding. The former variant may sterically restrict PLP binding, while the latter prevents formation of the internal aldimine. Improper PLP binding likely accounts for the undetectable L-ornithine turnover observed for H119Q and K230M. The D119A variant removes the salt bridge formed between the aspartic acid carboxylate with the pyridinium N on PLP.85, 117 This salt bridge has been shown in related enzymes to stabilise the quinonoid intermediate formed after the Cα decarboxylation step. The absence of activity observed for D119A is likely attributed to this variant destabilising the quinonoid intermediate. 5.4.5 ODA may function via a unique arginase mechanism In the arginase component, a series of mutations were made in order to revert the active site to the conserved first sphere coordination shell observed for arginase. S609 was replaced by aspartic acid (S609D), asparagine was converted to aspartic acid (N713D) and a double mutant containing both these mutations was produced (S609D/N713D). Finally, a fourth mutant was made that replaced S609 with an alanine reside (S609A). Each of the active site variants elicited a modest decrease in kcat. S609D and N713D showed a 14% reduction in kcat, while S609D/N713D and S609A elicited 2-fold decrease in kcat compared to wild type ODA. Variants of human arginase where D234 (the residue corresponding to N713 in ODA) was altered to either glutamic acid, alanine or histidine, demonstrated substantial drops in activity. This decrease in catalytic activity indicates that D234 is vital for catalysis by arginase. 161 The modest decrease in activity for S609D, N713D and S609D/N713D combined with the fact that Mn2+ does not appear to bind the active site indicates that the network of residues in ODA does not serve to coordinate to Mn2+. This suggests that another divalent metal made be required L-arginine hydrolysis by ODA. A structurally distinct arginase active site in ODA is also supported through inhibition experiments with S-(2-boronoethyl)-L-cysteine and 2(S)-amino-6-boronohexanoic acid. These inhibitors work by forming a boronate ion which mimics the tetrahedral intermediate formed during nucleophilic attack by the hydroxide ion. In human type II arginase, S-(2-boronoethyl)-L-cysteine and 2(S)-amino-6-boronohexanoic acid have Ki values of 0.31 and 0.25 µM respectively.144 However, both inhibitors failed to inhibit ODA activity, suggesting a structurally distinct binding pocket and perhaps a different mode of catalysis for the hydrolysis of L-arginine. 5.4.6 Substrate channelling from the arginase domain to the ODC domain When the activity of H119Q and K230M were tested in the presence of excess ODC/ODA there was no detectable CO2 formation. This suggest that the L-ornithine produced for the function arginase component of H119Q and K230M cannot be released from the enzyme and must interact with the circulated ODC/ODA. Thus, L-ornithine produced for the arginase active site is channelled directly to the ODC active site. 5.5 Concluding remarks From the experiments performed on the ODC component of ODA, we were able to confirm that H119, D199 and K230 are catalytically important residues in the active site. We established that K230 was the lysine residue that covalently bound to PLP during the formation of the internal aldimine. 162 The arginase component appears to demonstrate a unique mechanism that does not utilise Mn2+ as part of its catalytic mechanism. Known arginase inhibitors BEC and ABH did not have any effect on the activity of ODA. There also seems to be substrate channelling between the ODC and arginase components, consequently, inhibiting the ODC component may also inhibit the arginase component. ODA shows unique properties that may make it a good drug target for treatment of conditions associated with F. nucleatum. Thus far, we have been unable to successfully inhibit either domain of ODA. Further research on this enzyme could include a more comprehensive screening for inhibitors and determining the full crystal structure of the enzyme. 163 6. Conclusion Fusobacterium nucleatum is a gram-negative bacterium found in the human oral cavity. Over the past few decades F. nucleatum has been shown to be linked with several diseases. These include periodontal diseases, abnormal pregnancy outcomes, colorectal cancers. Therefore, finding therapeutics that selectivity target F. nucleatum is an important area of research. Some therapeutics work by inhibiting specific proteins that perform vital functions for the survival of the organism. By disabling these proteins, growth of the bacteria can be slowed or stopped, and which will improve the health outcomes for patients with F. nucleatum infections. In this thesis I have characterised three proteins encoded for by the F. nucleatum genome that are potential drug targets. These three proteins are: an electron transfer flavodoxin (Fld), lanthionine synthase (LS) and a bifunctional ornithine decarboxylase/ arginase (ODA). We have found that all three of these proteins show unique properties that may enable them to be selectively targeted by inhibitors. To confirm the importance of these proteins, in future studies we will produce knockout strains of F. nucleatum that are deficient in each of the genes encoding for these three proteins. This will be done in collaboration with Dr. Daniel Slade at Virginia Tech. Dr. Sladeâs lab have grown F. nucleatum under anaerobic conditions and created gene knock-outs for phospholipase A1 autotransporter FplA.194 This was performed by creating a FplA deletion strain in F. nucleatum using a single homologous crossover plasmid developed from a Clostridium shuttle vector and a recombinant method involving restriction endonuclease digestion to delete a specific internal fragment of the plasmid.195 A similar method can be used to create Fld, LS and ODA knockout strains. Comparison between growth of F. nucleatum with and without these genes should enable us to confirm if they are essential for the survival of the organism. We would anticipate that both ODA and LS are essential, as they both have products that we believe to be vital for the survival 164 of the organism which are not readily available for scavenging in the local environment of the organism. However, since there are seven Flds encoded for by the F. nucleatum genome, it is possible we may need to knockout all seven in order to impede growth of the bacterium. F. nucleatum is a flavodoxin specialist, in that it encodes for seven flavodoxin genes. Unlike many bacteria, it does not possess the iso-functional Fe-S dependent ferredoxin which may be a survival strategy for avoiding the toxic effects of H2O2 produced by neighbouring bacteria. Expansion of the flavodoxin pool may enable F. nucleatum to differentially control the flux of electron through different metabolic pathways. Chapters 2 and 3 of this thesis focuses on the characterisation of one of the seven flavodoxins produced by F. nucleatum. There are a number of conserved residues in the flavodoxin family that play an important role in tuning the FMN reduction potentials by modulating the binding affinity for different redox forms of the flavin cofactor. In Chapter 2, we found that the binding affinity of FMN to apo-FnFld was reduced compared with other related flavodoxins and that there was an increase in FMN binding affinity following an increase in ionic strength of the solution. This is attributed to a significant conformational shift of F93 observed in the crystal structure between apo- and hold-FnFld. These data suggest that FMN binding is initially driven via binding of the ribityl phosphate to the P-loop. FnFld was revealed to have a more electropositive Eox/sq and Esq/hq in comparison with other long-chain flavodoxins. The shift in Eox/sq is likely the result of the absence of a hydrogen bond between FMN N5 and the 50âs loop. This was confirmed by the crystal structure of holo-FnFld, which found the distance between FMN N5 and the closest hydrogen bond donor (NH on V58) to be 4.9 Ã , too far apart to form a hydrogen bond. The electropositive shift in Esq/hq may be attributed to 165 tighter binding of the negatively charged FMNhq. This tighter binding most likely the result of the near absence of destabilising negatively charged residues near the FMN N1. In Chapter 3, we established the significance of the four naturally occurring substitutions found in the primary sequence of FnFld. We found that Y56 and N89 contribute towards the electropositive values of Esq/hq. These do not provide a negative charge close to the FMN N1, therefore there is no destabilisation of the apo-FnFld-FMNhq complex. F93 contributed towards the electropositive value for Eox/sq and decrease in binding affinity to FMNox. This may be accounted for by weaker dispersion forces with the isoalloxazine ring on FMNox due to the electron-poor phenylalanine (relative to the conserved tyrosine). K13 and N89 also led to lower FMN binding affinity. With K13, this may be accounted for by the bulky lysine residue, which may increase the size of the conformational shift undertaken by the P-loop following FMN binding. N89 is a neutral residue that replaces a negatively charged aspartic acid, this results in a more electropositive Esq/hq due to partial stabilisation of the apo-FnFld-FMNhq complex. Y56 led to a greater FMN binding affinity, likely due to the formation of a tighter hydrogen bond between Y56 and the FMN ribityl phosphate. We have determined that FnFld has many unique properties impacting its biological function as an electron transfer protein. These may prove beneficial for the design of novel drugs to block the electron transfer properties of this protein and the growth of F. nucleatum. These insights into flavodoxins could also be used to design novel flavodoxins which may have other biotechnological applications in techniques such as genetic modification. For example, it may be possible to express flavodoxin in organisms that usually utilise ferredoxin to improve their survival in low iron conditions. It is conceivable that each flavodoxin produced by F. nucleatum is specific for a redox partner protein, although this will require more in-depth biochemical analysis. This will involve 166 identifying and expressing all the other flavodoxins and the Fld redox partners, such as pyruvate-flavodoxin oxidoreductase (PFOR) and flavodoxin-quinone reductase (FqrB), encoded by the F. nucleatum genome. We will then be able to characterise these proteins and perform biochemical assays to determine any selectivity between flavodoxins and the redox partners. This will help us establish a clearer understanding for energy transfer in F. nucleatum. In Chapter 4, we characterised a unique lanthionine synthase (LS), a unique PLP-dependent enzyme that generates H2S and L-lanthionine through the β-replacement of L-cysteine by a second molecule of L-cysteine. Lanthionine is a vital amino acid for F. nucleatum as meso-lanthionine (which L-lanthionine is a precursor) is incorporated into the peptidoglycan layer as part of the peptide cross-link. Therefore, targeting LS with a selective inhibitor may prevent production of lanthionine, thus weakening the bacterial membrane and resulting in the death of the bacterium. We confirmed that the likely physiological reaction of LS from F. nucleatum is the production of L-lanthionine and H2S from two equivalents of L-cysteine. Despite showing OASS activity, we concluded that this is unlikely to be physiologically necessary for F. nucleatum, as it lacks the gene coding for SAT, the precursor enzyme in the cysteine biosynthetic pathway. We identified a unique insertion of the residues 234SI235 compared with OASS enzymes. Our results suggest that the β-hydroxyl group of S234 forms a OH-O hydrogen bond to the carboxylate group of lanthionine. This shows that the 234SI235 insertion plays a vital role in coordinating binding of the second L-cysteine molecule. In addition, it stabilises the formation of lanthionine and prevents the reverse reaction, the β-replacement of L-lanthionine by sulfide to form L-cysteine, from occurring. This research may also be used to help develop new synthetic routes to form lanthionine or lanthionine analogues. 167 The next stage of this research will be to build on the inhibition studies discussed in this thesis. We will identify what type of compound make the most efficient inhibitors and will determine the crystal structure of LS bound to these compounds in order to determine how these compounds fit into the binding pocket and how they may be optimised. We will then need to test the inhibitors in vivo by growing F. nucleatum and treating the bacteria with the inhibitors to observe if they affect its growth. Finally, in Chapter 5 we characterised a unique bifunctional Ornithine decarboxylase/ arginase (ODA). The N-terminal half of the protein (residues 1 â 329) contains the ornithine decarboxylase (ODC) component, which catalyses the decarboxylation of L-ornithine to putrescine and CO2. The C-terminal half (residues 462 â 760), contains the arginase component, which catalyses hydrolysis of L-arginine to L-ornithine and urea. Since the F. nucleatum genome lacks genes that encode enzymes capable of spermidine or spermine synthesis from putrescine. It is likely that the main purpose of ODA in F. nucleatum is the production of urea. The urea produced from ODA is likely converted to ammonium and CO2 though the action of a urease encoded for by the F. nucleatum genome. NH4+ can protect the bacterium from acidic environments present in the mouth, making urea formation a very important biological role of ODA for F. nucleatum survival. We established that H119, D199 and K230 were amongst the amino acid residues that made up the active site of the ODC component. We found that the arginase component demonstrates a unique mechanism that does not utilise Mn2+ as part of its catalytic mechanism. There also seems to be substrate channelling between the ODC and arginase components preventing release of L-arginine in the absence of the ODC component. Consequently, inhibiting the ODC component may also inhibit the arginase component. ODA shows unique properties and structures, this may make it a good drug target for treatment of conditions associated 168 with F. nucleatum. This research may also be used to help the development of new synthetic routes for ornithine, putrescine and other product analogues. 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Thesis/Dissertation
eng
Biochemistry and Molecular Biology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Characterisation of three cofactor-containing proteins from the oral bacterium Fusobacterium nucleatum
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