Science, Faculty of
Chemistry, Department of
DSpace
UBCV
Cleveland, Maria
Doctor of Philosophy - PhD
University of British Columbia
The utilization and valorization of biomass has been proposed as a sustainable alternative to petroleum-based energy, chemicals and materials. Microbial enzymes can be used as selective catalysts for the depolymerization and functionalization of plant material. In this regard, the copper radical oxidases (CROs) from Auxiliary Activity Family 5 subfamily 2 (AA5_2) are attractive targets since they oxidize primary alcohols in a chemo-selective manner to the corresponding aldehydes without utilizing complex and expensive organic cofactors. AA5_2 was first defined by the archetypal galactose oxidase from Fusarium graminearum, however, other fungal AA5_2 members have recently been shown to comprise a wide range of specificities for aromatic, aliphatic and furan-based alcohols. This suggests a broader substrate scope of CROs for biocatalytic applications. However, only 10% of the annotated AA5_2 members have been characterized to date. The work presented in this thesis describes the biochemical characterization of fourteen fungal copper radical oxidases, recombinantly produced in Pichia pastoris, encompassing galactose oxidases, raffinose oxidases, broad spectrum alcohol oxidases, non-carbohydrate alcohol oxidases and aryl alcohol oxidases. Detailed product analysis was conducted of carbohydrates, furans and glycerol oxidized by these novel enzymes to further explore their potential applications. Of note, we observed direct oxidation of 5-hydroxymethylfurfural (HMF), an important platform chemical derived from biomass, to 5-formyl-2-furoic acid (FFCA) by six enzymes. One enzyme facilitated the direct one-pot conversion of HMF to 2,5-furandicarboxylic acid (FDCA), a building block for bio-plastic production. Through docking of galactose into homology models of these newly characterized enzymes, and comparing it to the archetypal galactose oxidase, key amino acid residues important for substrate specificity have been proposed. Finally, the potential of enzyme immobilization using the cross-linked enzyme aggregate (CLEA) strategy has been investigated for a specific alcohol oxidase to increase its utilization in biocatalysis. This work expands our understanding of the catalytic diversity of copper radical oxidases from AA5_2 and doubles the number of characterized AA5_2 members. Detailed sequence and enzymatic analysis highlights their structure-function relationships while product analysis informs their possible biotechnological applications in bio-products manufacturing. Furthermore, immobilization was shown to be a promising avenue for increasing the biocatalytic potential of CROs.
INVESTIGATING CATALYTIC DIVERSITY IN COPPER RADICAL OXIDASES FOR POTENTIAL BIO-PRODUCT APPLICATIONS by Maria Cleveland B.Sc., Queens University, 2014 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Chemistry) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) August 2021 © Maria Cleveland, 2021 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: INVESTIGATING CATALYTIC DIVERSITY IN COPPER RADICAL OXIDASES FOR POTENTIAL BIO-PRODUCT APPLICATIONS submitted by Maria Cleveland in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Chemistry Examining Committee: Dr. Harry Brumer, Professor, Chemistry, UBC Supervisor Dr. Stephen Withers, Professor, Chemistry, UBC Supervisory Committee Member Dr. David Perrin, Professor, Chemistry, UBC University Examiner Dr. Kirsten Wolthers, Associate Professor, Chemistry, UBC University Examiner Additional Supervisory Committee Members: Dr. Pierre Kennepohl, Associate Professor, Chemistry, UBC Supervisory Committee Member Dr. Mark MacLachlan, Professor, Chemistry, UBC Supervisory Committee Member iii Abstract The utilization and valorization of biomass has been proposed as a sustainable alternative to petroleum-based energy, chemicals and materials. Microbial enzymes can be used as selective catalysts for the depolymerization and functionalization of plant material. In this regard, the copper radical oxidases (CROs) from Auxiliary Activity Family 5 subfamily 2 (AA5_2) are attractive targets since they oxidize primary alcohols in a chemo-selective manner to the corresponding aldehydes without utilizing complex and expensive organic cofactors. AA5_2 was first defined by the archetypal galactose oxidase from Fusarium graminearum, however, other fungal AA5_2 members have recently been shown to comprise a wide range of specificities for aromatic, aliphatic and furan-based alcohols. This suggests a broader substrate scope of CROs for biocatalytic applications. However, only 10% of the annotated AA5_2 members have been characterized to date. The work presented in this thesis describes the biochemical characterization of fourteen fungal copper radical oxidases, recombinantly produced in Pichia pastoris, encompassing galactose oxidases, raffinose oxidases, broad spectrum alcohol oxidases, non-carbohydrate alcohol oxidases and aryl alcohol oxidases. Detailed product analysis was conducted of carbohydrates, furans and glycerol oxidized by these novel enzymes to further explore their potential applications. Of note, we observed direct oxidation of 5-hydroxymethylfurfural (HMF), an important platform chemical derived from biomass, to 5-formyl-2-furoic acid (FFCA) by six enzymes. One enzyme facilitated the direct one-pot conversion of HMF to 2,5-furandicarboxylic acid (FDCA), a building block for bio-plastic production. Through docking of galactose into homology models of these newly characterized enzymes, and comparing it to the archetypal galactose oxidase, key amino acid residues important for substrate specificity have been iv proposed. Finally, the potential of enzyme immobilization using the cross-linked enzyme aggregate (CLEA) strategy has been investigated for a specific alcohol oxidase to increase its utilization in biocatalysis. This work expands our understanding of the catalytic diversity of copper radical oxidases from AA5_2 and doubles the number of characterized AA5_2 members. Detailed sequence and enzymatic analysis highlights their structure-function relationships while product analysis informs their possible biotechnological applications in bio-products manufacturing. Furthermore, immobilization was shown to be a promising avenue for increasing the biocatalytic potential of CROs. v Lay Summary Climate change is a pressing worldwide concern and changing the economy from petroleum-based to bio-based is a strategy to mitigate this issue. In a bio-based economy, plant resources are utilized to produce energy, chemicals, and materials. One avenue to pursue this goal is to use enzyme as catalysts for the breakdown and modification of biomass. Processes for the degradation and subsequent fermentation of cellulose rich plant materials (eg. switchgrass and corn) have been developed to produce bio-ethanol as fuel. Furthermore, the use of enzymes to manufacture chemicals and pharmaceuticals is a growing area of research. In this thesis, I produced and characterized fourteen new fungal copper-radical oxidase (CRO) enzymes that have a diverse range of activities on a wide variety of substrates found within biomass or industrially relevant. A large focus of the work was to investigate the potential applications of these enzymes for bio-product manufacturing and to increase their potential utilization in biocatalysis. vi Preface Chapter 1: This chapter was written by the author (Maria Cleveland) with feedback from Prof. Harry Brumer (Ph.D. supervisor). Chapter 2: This chapter was written by the author (Maria Cleveland), who also produced the recombinant enzymes, performed all enzyme kinetic and product analysis. Dr. Michael Lafond cloned and generated expression strains. Roderick (Fan) Xia and the author (Maria Cleveland) optimized and performed glyceraldehyde stereochemistry analysis with support from Dr. Ryan Chung and Dr. Paul Mulyk, under the supervision of Prof. Jason Hein. Prof. Harry Brumer (Ph.D. supervisor) reviewed and revised the manuscript. A version of this chapter has been submitted and accepted for publication: Cleveland, M., Lafond, M., Xia, F. R., Chung, R., Mulyk, P., Hein, J., & Brumer, H. Two Fusarium copper radical oxidases with high activity on aryl alcohols. Biotechnol. Biofuels 2021, 14 (1), 138. Chapter 3: This chapter was written by the author (Maria Cleveland), who also performed the enzyme cloning, biochemistry, enzyme kinetic and product analysis. Dr. Yann Mathieu performed sequence analysis, bioinformatics, and assisted with the enzyme cloning. David Ribeaucourt and Mireille Haon performed recombinant protein production and initial kinetic measurements, under the supervision of Prof. Michael Lafond and Prof. Jean-Guy Berrin. The author (Maria Cleveland) performed the glycerol oxidation experiments and the analysis was performed with assistance from Dr. Paul Mulyk, under the supervision of Prof. Jason Hein. Prof. Harry Brumer (Ph.D. supervisor) reviewed and revised the chapter. A version of this chapter will be submitted for publication. vii Chapter 4: This chapter was written by the author (Maria Cleveland), who also performed the experiment. The HPLC analysis was performed through collaboration with Dr. Jason Hein’s lab. Prof. Harry Brumer (Ph.D. supervisor) reviewed and revised the chapter. Chapter 5: This chapter was written by the author (Maria Cleveland) with feedback from Prof. Harry Brumer (Ph.D. supervisor). viii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ....................................................................................................................... viii List of Tables .............................................................................................................................. xiv List of Figures ...............................................................................................................................xv List of Symbols .......................................................................................................................... xvii List of Abbreviations ............................................................................................................... xviii Acknowledgements .................................................................................................................. xxiv Dedication ................................................................................................................................. xxvi Chapter 1: Introduction ................................................................................................................1 1.1 Biorefineries and Biocatalysis ........................................................................................ 1 1.2 Bio-redox Catalysis ......................................................................................................... 4 1.3 Copper Radical Oxidases ................................................................................................ 7 1.3.1 Tertiary Structure ........................................................................................................ 8 1.3.2 Reaction Mechanism ................................................................................................. 10 1.3.3 Redox Potential ......................................................................................................... 12 1.3.4 Substrate Specificity ................................................................................................. 13 1.3.5 Biological Role ......................................................................................................... 18 1.3.6 Applications .............................................................................................................. 20 1.4 Enzyme Immobilization ................................................................................................ 22 ix 1.4.1 Immobilization Methods ........................................................................................... 24 1.4.2 Cross-Linked Enzyme Aggregates (CLEA) ............................................................. 27 1.5 Thesis Aims .................................................................................................................. 30 Chapter 2: Two Fusarium Copper Radical Oxidases with High Activity on Aryl Alcohols 32 2.1 Introduction ................................................................................................................... 32 2.2 Results and Discussion ................................................................................................. 35 2.2.1 Bioinformatics........................................................................................................... 35 2.2.2 Protein Production and Substrate Screening ............................................................. 39 2.2.3 Michaelis-Menten Kinetics and Product Analysis .................................................... 42 2.2.3.1 Aryl Alcohol Oxidation .................................................................................... 44 2.2.3.2 Saccharide Oxidation ........................................................................................ 48 2.2.3.3 Glycerol Oxidation............................................................................................ 57 2.3 Conclusion .................................................................................................................... 58 2.4 Materials and Methods .................................................................................................. 60 2.4.1 Chemicals and Enzymes ........................................................................................... 60 2.4.2 Sequence and Bioinformatic Analysis ...................................................................... 60 2.4.3 DNA Cloning ............................................................................................................ 61 2.4.4 Protein Production and Purification .......................................................................... 62 2.4.5 Analytical Protein Deglycosylation .......................................................................... 64 2.4.6 pH and Temperature Profile ...................................................................................... 64 2.4.7 Temperature Stability................................................................................................ 65 2.4.8 Enzyme Kinetics ....................................................................................................... 65 2.4.9 Computational Docking Studies ............................................................................... 66 x 2.4.10 Synthesis of Methyl α/β-D-fructofuranosides and Methyl β-D-fructopyranoside 67 2.4.11 Enzyme Product Analysis ..................................................................................... 68 Chapter 3: Biochemical survey of Copper Radical Oxidase specificity in Auxiliary Activity Family 5.........................................................................................................................................72 3.1 Introduction ................................................................................................................... 72 3.2 Results ........................................................................................................................... 74 3.2.1 Molecular Diversity of AA5_2 Members ................................................................. 74 3.2.2 Protein Production and Initial Biochemical Characterization .................................. 78 3.2.3 Extended Substrate Screening and Michaelis-Menten Kinetics ............................... 80 3.2.3.1 Galactose 6-oxidases ......................................................................................... 83 3.2.3.1.1 ExeGalOx .................................................................................................... 83 3.2.3.1.2 MreGalOx ................................................................................................... 83 3.2.3.1.3 FoxGalOxB ................................................................................................. 84 3.2.3.1.4 PfeGalOx ..................................................................................................... 85 3.2.3.2 Raffinose Oxidases ........................................................................................... 85 3.2.3.2.1 UmaRafOx .................................................................................................. 86 3.2.3.2.2 PhuRafOx .................................................................................................... 86 3.2.3.3 Broad-spectrum Alcohol Oxidases ................................................................... 87 3.2.3.3.1 AflAlcOx...................................................................................................... 87 3.2.3.3.2 PruAA5_2 (PruAlcOx) ............................................................................... 87 3.2.3.3.3 FoxAlcOx .................................................................................................... 89 3.2.3.3.4 PorAlcOx .................................................................................................... 89 3.2.3.4 Non-carbohydrate Alcohol Oxidases ................................................................ 90 xi 3.2.3.4.1 GciAlcOx .................................................................................................... 90 3.2.3.4.2 AsyAlcOx .................................................................................................... 91 3.2.4 Application to Bioproduct Conversion ..................................................................... 91 3.2.4.1 HMF .................................................................................................................. 92 3.2.4.2 Glycerol............................................................................................................. 94 3.3 Discussion ..................................................................................................................... 96 3.4 Conclusion .................................................................................................................. 101 3.5 Materials and Methods ................................................................................................ 102 3.5.1 Chemicals and enzymes .......................................................................................... 102 3.5.2 Sequence Analysis and Bioinformatics .................................................................. 102 3.5.3 DNA Cloning and Recombinant Strain Production ................................................ 104 3.5.4 Protein Production and Purification ........................................................................ 105 3.5.5 Initial Activity Test ................................................................................................. 108 3.5.6 Plate Assays ............................................................................................................ 109 3.5.7 Michaelis-Menten Kinetics ..................................................................................... 110 3.5.8 Computational Docking Studies ............................................................................. 111 3.5.9 Enzyme Product Analysis ....................................................................................... 111 Chapter 4: Cross-linked Enzyme Aggregates of a Copper Radical Alcohol Oxidase for Increased Potential in Biocatalysis ...........................................................................................114 4.1 Introduction ................................................................................................................. 114 4.2 Results ......................................................................................................................... 117 4.2.1 CgrAlcOx, Catalase and HRP CLEA ..................................................................... 117 4.2.1.1 Screening CLEA Preparation Conditions ....................................................... 117 xii 4.2.1.2 Comparison of CLEA and combi-CLEA Activity.......................................... 119 4.2.2 Replacement of HRP with Potassium Ferricyanide ................................................ 123 4.3 Discussion ................................................................................................................... 128 4.4 Conclusions ................................................................................................................. 132 4.5 Materials and Methods ................................................................................................ 133 4.5.1 Chemicals and Enzymes ......................................................................................... 133 4.5.2 HPLC Method ......................................................................................................... 133 4.5.3 CRO Activity Assay ............................................................................................... 134 4.5.4 Precipitant and Glutaraldehyde Inhibition Assay ................................................... 134 4.5.5 CLEA and combi-CLEA Preparation ..................................................................... 134 4.5.6 CLEA Activity Assay ............................................................................................. 135 4.5.7 Reusability .............................................................................................................. 137 4.5.8 Non-CLEA Reaction Monitoring ........................................................................... 137 Chapter 5: General Conclusions and Future Directions ........................................................138 5.1 General Conclusions ................................................................................................... 138 5.2 Future Directions ........................................................................................................ 140 References ...................................................................................................................................144 Appendices ..................................................................................................................................162 Appendix A : Supporting Information for Chapter 2: Two Fusarium Copper Radical Oxidases with High Activity on Aryl Alcohols...................................................................................... 162 A.1 Supporting Tables ................................................................................................... 162 A.2 Supporting Figures .................................................................................................. 171 xiii Appendix B : Supporting Information for Chapter 3: Biochemical survey of Copper Radical Oxidase specificity in Auxiliary Activity Family 5 ................................................................ 207 B.1 Supporting Tables ................................................................................................... 207 B.2 Supporting Figures .................................................................................................. 226 Appendix C : Supporting Information for Chapter 4: Cross-linked Enzyme Aggregates of a Copper Radical Alcohol Oxidase for Increased Potential in Biocatalysis .............................. 257 C.1 Supporting Tables ................................................................................................... 257 C.2 Supporting Figures .................................................................................................. 257 xiv List of Tables Table 2.1. Substrate specificity of FgrAAO and FoxAAO .......................................................... 43 Table 2.2. Percent conversion of HMF, DFF and HMFCA by FgrAAO and FoxAAO .............. 47 Table 3.1. Key amino acid residues in AA5_2 proteins compared to other characterized members....................................................................................................................................................... 77 Table 3.2. Initial rate kinetics of active AA5_2 enzymes ............................................................. 82 Table 3.3. Percent conversion of HMF, HMFCA, DFF and FFCA by PruAA5_2A (PruAlcOx), AflAlcOx, GciAlcOx, PorAlcOx and FoxGalOxB ....................................................................... 93 Table 3.4. Stereochemical outcome of glycerol oxidation by AA5 CROs ................................... 95 Table 4.1. Experimental conditions for CLEA preparation ........................................................ 135 xv List of Figures Figure 1.1. Diagram of biorefinery applications ............................................................................. 3 Figure 1.2. Distribution of reaction types for industrial bio-oxidation reactions ........................... 4 Figure 1.3. Reactions catalyzed by oxidative enzymes .................................................................. 5 Figure 1.4. Crystal structure of copper radical oxidases ................................................................. 9 Figure 1.5. Active site residues of copper radical oxidases .......................................................... 10 Figure 1.6. Reaction mechanism of copper radical oxidases ........................................................ 12 Figure 1.7. Substrates for archetypal galactose oxidase (FgrGalOx) ........................................... 14 Figure 1.8 Substrates for CgrAlcOx (A.) and CgrAAO (B.)........................................................ 18 Figure 1.9. Enzyme immobilization methods ............................................................................... 24 Figure 1.10. Cross-linking enzyme aggregates procedure ............................................................ 27 Figure 2.1. Molecular phylogeny of 45 AA5_2 members ............................................................ 36 Figure 2.2. Active site amino acids in AA5_2 members .............................................................. 38 Figure 2.3. Initial activity screens of FgrAAO and FoxAAO ...................................................... 41 Figure 2.4. HMF oxidation ........................................................................................................... 46 Figure 2.5. Molecular docking of galactose (cyan) in FgrGalOx (PDB ID 1GOF)56 (A) and a Phyre2 homology model of FgrAAO (B) using AutoDock Vina as implemented in Chimera ... 49 Figure 2.6. Summary of observed hydrated aldehyde (gem-diols) products from the oxidation of methyl fructosides, anhydro sugars and raffinose by FgrAAO .................................................... 51 Figure 2.7. Complete analysis of oxidation reaction components of FgrAAO with (A) methyl β-D-fructopyranose (1) and (B) methyl β-D-fructofuranose (2) ...................................................... 53 xvi Figure 2.8. Molecular docking (AutoDock Vina as implemented in Chimera) of (A) methyl β-D-fructopyranoside (3), (B) methyl β-D-fructofuranoside (2) and (C) methyl α-D-fructofuranoside (1) in FgrAAO .............................................................................................................................. 56 Figure 3.1. Sequence relationships of 623 AA5_2 catalytic modules .......................................... 76 Figure 3.2. Initial activity screens of expressed AA5_2 ............................................................... 79 Figure 3.3. Extended activity screening of AA5_2 enzymes........................................................ 81 Figure 3.4. Oxidation pathways of HMF and Glycerol ................................................................ 92 Figure 3.5. Molecular docking of galactose (cyan) into the experimental tertiary structure of FgrGalOx (PDB ID 1GOF) .......................................................................................................... 97 Figure 4.1. Effect of cross-linker concentration on CLEA/combi-CLEA activity ..................... 119 Figure 4.2. Comparison of percent conversion of CgrAlcOx CLEA with soluble accessory enzymes, combi-CLEA and soluble enzymes for benzyl alcohol oxidation .............................. 121 Figure 4.3. Influence of ferricyanide concentration on CLEA and combi-CLEA percent conversions ................................................................................................................................. 125 Figure 4.4. Time dependent conversion of 10 mM benzyl alcohol by CgrAlcOx CLEA and combi-CLEA ............................................................................................................................... 126 Figure 4.5. Effect of the addition of a new component on the percent conversion of 10 mM benzyl alcohol by CgrAlcOx CLEA ........................................................................................... 128 xvii List of Symbols Å Angstrom = 10-10 meters α Alpha β Beta ˚C Degrees Celsius ε Extinction coefficient δ Chemical shift in NMR spectroscopy % Percent g Gravitational field strength λmax Lambda max (k)Da (Kilo)dalton [S] Substrate concentration xviii List of Abbreviations AA Auxiliary Activity Family AA5 Auxiliary Activity Family 5 AA5_1 Auxiliary Activity Family 5 Subfamily 1 AA5_2 Auxiliary Activity Family 5 Subfamily 2 AAO Aryl alcohol oxidase ABTS 2.2’-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid) Afl Aspergillus flavus AlcOx Alcohol oxidase Arg (R) Arginine Asy Aspergillus sydowii BMGY Buffered complex glycerol medium BMMY Buffered complex methanol medium BSA Bovine serum albumin CAT Catalytic CAZy Carbohydrate-Active enZYmes CBM Carbohydrate binding module Cca Corynespora cassiicola cDNA complementary Deoxyribonucleic acid Cgl Colletotrichum gloeosporioides Cgr Colletotrichum graminicola Chi Colletotrichum higginsianum CLEA Cross-linked enzyme aggregates xix CRO Copper radical oxidase Cu Copper CuSO4 Copper Sulfate Cys (C) Cysteine DFF 2,5-Diformylfuran EC Enzyme Commission ESI Electrospray ionization ET Electron transfer EtOAc Ethyl acetate Exe Exophiala xenobiotica FAD Flavin adenine dinucleotide FDCA 2,5-Furandicarboxylic acid FFCA 2,5-Formylfurancarboxylic acid Fgr Fusarium Graminearum FMN Flavin mononucleotide Fox Fusarium Oxysporum GalOx Galactose Oxidase Gci Glomerella cingulate GDP Gross domestic product h Hour H Proton H2O2 Hydrogen peroxide ha Hectare xx HAT Hydrogen atom transfer HCl Hydrochloric Acid His (H) Histidine HisTrap Nickel affinity column HMF 5-Hydroxymethylfurfural HMFCA 5-Hydroxymethyl-2-furancarboxylic acid HPLC High-performance liquid chromatography HRP Horseradish peroxidase IMAC Immobilized Metal Affinity Chromatography kcat Unimolecular rate constant kcat/KM Catalytic efficiency KM Michaelis constant L Litre LBLS Luria Bertani Low Salt LC-MS Liquid chromatography–mass spectrometry Lys (K) Lysine M Molar m/z Mass-to-charge ratio MeOH Methanol mg Milligram MHz Megahertz min Minute mL Millilitre xxi mM Millimolar mmol Millimole mol Mole Mre Mytilinidion resinicola Mw Molecular weight MWCO Molecular weight cut-off NaCl Sodium chloride NAD(P)H Reduced nicotinamide adenine dinucleotide (phosphate) Ni-NTA Ni nitrilotriacetic acid nm Nanometre nmol Nanomole NMR Nuclear magnetic resonance OH Hydroxyl group P. pastoris Pichia pastoris PDB Protein Data Bank Pfe Penicillium fellutanum Phe (F) Phenylalanine Phu Pseudozyma hubeiensis Por Pyricularia oryzae ppm Parts per million Pru Penicillium rubens psi Pounds per square inch PT Proton transfer xxii RafOx Raffinose oxidase Rf Retention factor rpm Revolutions per minute RT Room temperature Rz Ratio of absorbance due to hemin to absorbance due to protein SDS-PAGE Sodium dodecyl sulfate-polyacrylamide SEC Size exclusion chromatography s seconds Ser (S) Serine Sto Stogonospora sp Trp (W) Tryptophan Tyr (Y) Tyrosine U Enzyme unit Uma Ustilago maydis UNK Unknown module UV Ultraviolet V Voltage v/v Volume by volume VIS Visible w/v Weight by volume YPD Yeast extract peptone dextrose μg Microgram μL Microlitre xxiii μM Micromolar µm Micrometre xxiv Acknowledgements I would like to express my sincere gratitude to all of the family, friends, mentors and colleagues that have supported me throughout my graduate degree. Foremost, I would like to thank my supervisor, Dr. Harry Brumer, for providing me the opportunity to pursue a PhD is his lab, for offering scientific guidance through my degree, for supplying the resources to pursue a degree in between the field of chemistry and biology, and for developing my writing skills. I would also like to thank my colleagues in the Brumer lab, especially the CRO team: Dr. Yann Mathieu, Dr. Stephanie Forget, and Roderick Xia. The three of you provided me with much needed scientific steering and our discussions were always insightful and motivational. A special thank you to Dr. Julie Grondin, Dr. Laleh Solhi and Dr. Shaheen Shojania for the friendly conversations and lending a sympathetic ear during lunchtime. And of course, thank you to all the other members of the Brumer Lab, past and present, for your mentorship and support. Each and every one of you holds a special place in my heart and I will not forget the kind and fun lab environment that all of you created. I would like to acknowledge all the current and former members of the many UBC chemistry organizations I have had the pleasure to be a part of. Whether organizing social events with CGSS, developing an action plan on sustainability efforts with GCU or discussing important societal issues with CDI, all of these activities were critical for my success in my graduate degree and have motivated to lead a life of service to my communities. I am grateful to the members of my supervisory committee Dr. Stephen Withers, Dr. Pierre Kennepohl, and Dr. Mark MacLachlan, who have provided feedback and guidance over the years. xxv Last but foremost, I want to thank my family for always being there for me in times of need. My parents, Dr. Tatiana Knodel and Dr. Bruce Cleveland, have fostered my love of science in me from a very young age. They were both amazing role models demonstrating on a daily basis the qualities of great scientist. They have always been supportive and encouraging of my dreams and I will appreciate their love and kindness forever. And my deepest gratitude goes to my partner, Dr. Nick Hein, who has been my rock for the past four years. It has been a unique experience going through graduate school with someone who can relate to almost all my experiences and has been there with me to celebrate my achievements and to pick me up when times our rough. You are one of the most patient, kind, loving and affectionate people I know and thank you for believing in me when I doubted myself. xxvi Dedication I would like to dedicate this thesis to my parents, Dr. Tatiana Knodel and Dr. Bruce Cleveland, and to my partner, Dr. Nicholas Hein. Their enduring love and compassion gave me the strength to continue during difficult times. 1 Chapter 1: Introduction 1.1 Biorefineries and Biocatalysis With over 347 million hectares (ha) of forest, Canada has 9% of the world’s forest and the forestry industry contributed 23.7 billion (1.1%) to Canada’s nominal GDP in 2019.1 The wood fibers associated with the foresting industry are a common feedstock for traditional products such as pulp, paper and lumber. However, in Canada, newsprint production declined 70% from 1990 compared to 2019 due to increased global competition and the prevalence of digital media.1 Hence, the need to modernize the output of the forestry industry is paramount for the continued economic growth of such a fundamental Canadian industry. Consequently, the functionalization and valorization of traditional biomass has been a growing field for many years. In the annual report on the State of the Canadian Forests it was noted that advances in bio-based products are expected to contribute up to 50% of consumer products by the year 2050.1 Furthermore, the diversification of the forest sector through the growth of the circular bioeconomy, which encompasses bio-chemicals, bio-materials and bioenergy, would create new opportunities and jobs in the country. In addition, the devastating effects of climate change such as frequent severe storms and hurricanes, devastating forest fires, rising seas levels, warming oceans, species extinctions and more, have increased world-wide concern of greenhouse gas emissions from our society’s usage of fossil fuels.2 The push towards a low-carbon circular bio-based economy is essential to mitigate the current high-carbon lifestyle western society has exploited for decades.3 In this regard, an integrated biorefinery is a major step towards transitioning to a low-carbon circular bio-based economy. The term biorefinery describes a facility analogous to a petroleum refinery, which utilizes biomass as the carbon source. Biorefineries can produce a 2 multitude of varied outputs such as traditional forestry products (pulp, paper and lumber), as well as first/second generation biofuels and more complex bio-products, bio-materials and bio-chemicals (Figure 1.1).4-6 Recently, there has been strong interest in converting cellulose-rich biomass to ethanol through hydrolysis of plant material to glucose and subsequent fermentation. This process can be energy intensive and its viability is highly dependent on the feedstock cost relative to the market price of ethanol.7 Due to the potential negative impact of feedstock sourcing on biodiversity, land use and competition with food crops, a more recent line of inquiry has been in second generation bio-fuels.8 This process involves degrading complex woody biomass containing lignin, hemicellulose and cellulose to simple sugars though depolymerization and fermentation. Due to the higher degree of complexity of the feedstock, second generation biofuels are a difficult process to optimize to be competitive in the current fuel market.9 The utilization of forest biomass for bio-fuels production is a great opportunity to investigate and invest in for the Canadian forestry industry in order to reduce the consumption of fossil fuels in the transportation sector. However, all bio-fuel derived processes seek to take an intricate natural substrate full of inherent chemical complexity and to deconstruct it to simple component parts. A different approach would involve exploiting the underlying functionality of biomass to produce high-value bio-chemicals and bio-materials through partial degradation and selective derivatization, as this would be a more efficient and economical approach to biomass utilization than full catabolism to low-value fuels.10, 11 3 Figure 1.1. Diagram of biorefinery applications (adapted with permission from FPAC and FPInnovations, 20116) Furthermore, biomass is complex and contains many component parts such as fibers, carbohydrates and phenolics which would all be utilized individually to produce an assortment of products in an integrated biorefinery.12 In nature, biomass is efficiently degraded by a variety of fungal species though secreted enzymes. These biocatalysts have undergone selective pressure to become efficient and selective and can be utilized for industrial purposes to depolymerize, hydrolyze and oxidize a multitude of different biomass feedstocks.13-21 A wide diversity of microorganisms have been mined for new and improved natural biocatalysts for a wide range of applications in the detergent, food, feed and biofuels industries.22, 23 Biocatalysis has recently garnered a lot of industrial interest, particularly in the pharmaceutical industry, as a green alternative to traditional synthetic routes.24 There are many key advantages to biocatalysis such as 1) enzymes being renewable and biodegradable catalysts; 2) enzymes are selective chiral catalysts, often with inherent regio- and stereo-selectivities, that can be essential in synthetic 4 schemes; 3) enzymes can be easily altered through rational protein engineering or directed evolution to obtain increased activity, increased stability or different selectivity using well-stablished genetic tools; 4) enzymes can help current synthetic processes by shortening synthetic routes and discontinuing the usage of protecting groups; 5) lowering industrial costs by performing reactions under milder conditions such as lower pressures and temperatures and with low catalyst loading, and 6) enzyme can be immobilized to increase their stability and reusability.14, 25, 26 Biocatalysis is a sustainable technology based upon the development of new enzymes with favourable industrial properties and improved process engineering and the potential for enzyme immobilization to increase performance and commercial success. 1.2 Bio-redox Catalysis Bio-redox catalysis accounts for one third of all enzymatic processes that are performed commercially, approximately half of which are oxidation reactions.14 Most of those are enantioselective oxyfunctionalizations such as hydroxylation, dihydroxylation, epoxidation and Baeyer-Villiger oxidations (Figure 1.2).27 Figure 1.2. Distribution of reaction types for industrial bio-oxidation reactions (adapted with permission from Hollmman et al. 201114) 5 The oxidation of diverse aliphatic and aromatic alcohols is common in nature and has significant utility in a variety of biotechnological applications.13-15 Four main enzymatic activities have been employed for industrial oxidations: dehydrogenases, oxidases, oxygenases and peroxidases (Figure 1.3). These enzymes function on a vast array of substrates including primary and secondary alcohols, benzylic alcohols, and certain simple sugars. However, despite these enzymes being effective catalysts, a few drawbacks exist limiting their broad industrial utilization such as narrow substrate scope, cofactor dependence, stability and cost. Figure 1.3. Reactions catalyzed by oxidative enzymes A. Dehydrogenases, B. Oxidases, C. Oxygenases, D. Peroxidases (adapted with permission from Dong et al. 201828) Dehydrogenases reversibly oxidize alcohols and amines, and the most studied class being alcohol dehydrogenases (ADHs)27, 29 with many thermostable enzymes from a variety of different organisms.30-33 ADHs can possess catalytic activity in media with high concentrations of organic solvents such as the enantioselective oxidation of sec-alcohol and the asymmetric reduction of ketones, using acetone and 2-propanol.34-36 6 Oxidases catalyze the electron transfer from their substrate to molecular oxygen with concomitant formation of water or peroxide as the byproduct. Many oxidases acting on specific substrates such as aliphatic alcohol oxidases, polyol oxidases (including glucose oxidases, pyranose-2-oxidase and alditol oxidase), glycolate oxidase and cholesterol oxidase have been found in a variety of organisms.14 With regards to amine oxidation, the flavin-dependent monoamine oxidase from Aspergillus niger (MAO-N) has been used as a starting point to create variants capable of oxidizing primary, secondary and tertiary amines.37 Of significant industrial interest was the discovery of a flavin-dependent 5-hydroxymethyl furfural oxidase (HMF oxidase) capable of converting HMF into FDCA through a three-step pathway.38 HMF can be made sustainably from fructose39 and FDCA can be used in the production of bio-based plastics.40 Oxygenases incorporate molecular oxygen directly into organic molecules through two different mechanisms: monooxygenases add a single oxygen atom to a molecule while the other oxygen atom is converted to water though the use of a donor, while dioxygenases incorporate molecular oxygen into the substrate through a peroxy-species.29 Oxygenases produce potent oxygenation agents capable of epoxidation, aromatic hydroxylation and Bayer-Villiger chemistry. Cytochrome P450 monooxygenases containing a heme iron complex and mono-/multi-nuclear Fe-monooxygenases are examples of oxygenases with industrial potential due to the potential of harnessing their tremendous substrate scope for chemical synthesis.41, 42 Lastly, peroxidases perform a two-electron oxidation using hydrogen peroxide and producing water. These catalysts can contain protoporphyrin IX, selenium, vanadium or manganese metal ions and perform chemical transformations similar to P450 monooxygenases.29 7 Most of the aforementioned enzymes require an organic cofactor in order to perform their oxidative chemistry. These include, but are not limited to, nicotinamide cofactors (NADH or NADPH), which are required for most dehydrogenases and oxygenases, whereas oxidases can require a flavin cofactor (FAD or FMN).13 These cofactors are often expensive and could require whole cell regeneration systems for industrial utilization, which can be costly and problematic. Furthermore, many enzymes with oxidative chemistry have the potential to produce the carboxylic acid from their alcohol substrates. Depending on the desired final product, the reactive aldehyde may be a necessary intermediate and the problem of enzymatic over-oxidation can limit the utilization of these biocatalysts. In this regard, a class of oxidative enzymes that has been overlooked, until recently, for their potential industrial applicability are copper radical oxidases (CROs). CROs are stereo- and regio-selective organic cofactor-independent catalysts active on a variety of carbohydrates, aliphatic and aromatic alcohols. These enzymes only require an inorganic copper metal ion for their mechanism and molecular oxygen as a co-substrate. 1.3 Copper Radical Oxidases Copper radical oxidases comprise the Auxiliary Activity Family 5 (AA5) in the Carbohydrate-Active Enzymes (CAZy) classification.43 CROs are classified in two subfamilies, where subfamily AA5_1 contains characterized glyoxal oxidases (EC 1.2.3.15)44 and subfamily AA5_2 contains galactose oxidases (EC 1.1.3.9)45 in addition to the more recently discovered aliphatic alcohol oxidases (EC 1.1.3.13)46, 47 and aryl alcohol oxidase (EC 1.1.3.7).48 The most studied enzyme in subfamily AA5_1 is the glyoxal oxidase from Phanerochaete chrysosporium discovered in 1987.49 For subfamily AA5_2, the archetypal galactose-6 oxidase from Fusarium graminearum (FgrGalOx) was first reported in 1959 from cultures of Polyporus circinatus (later 8 named Fusarium graminearum50, 51). This first report already established FgrGalOx as a metalloenzyme; its copper requirement was then confirmed by Amaral et al. in 1963.52 Although known for decades, both subfamilies have undergone limited characterization. With over 650 publicly available putative AA5 sequences currently in the CAZy database, only 5 members from the AA5_1 and 11 from the AA5_2 have been characterized with 3 crystal structures determined only for AA5_2 members. Moreover, until 2015 the characterized enzymes from the AA5_2 subfamily were found to exhibit mainly galactose oxidase activity, but in the past five years five novel non-carbohydrate oxidase enzymes were found, in addition to a raffinose oxidase, therefore expanding the substrate diversity of the AA5_2 subfamily.46-48, 53 Furthermore, an enzyme from Penicillium rubens Wisconsin 54–1255 (PruAA5_2A) has been identified as a copper radical oxidase with functional overlap across the AA5 family substrates.54 1.3.1 Tertiary Structure As CROs, AA5_1 and AA5_2 share the same tertiary structure, despite having low sequence identity and differing substrate specificities.43, 55 Furthermore, all characterized enzymes possess a seven-bladed β-propeller fold (Figure 1.4).46, 48, 56 The archetypal GalOx is made up of three domains: domain 1 has a “β sandwich” structure identified as a carbohydrate binding module (CBM32) with affinity for galactose,56, 57 domain 2 is the catalytic domain which has the pseudo symmetric seven-bladed β-propeller fold, and domain 3, which is the smallest, forms a hydrogen bonding network to stabilize domain 2. The seven-bladed β-propeller domain58 has been found in other non-related proteins: methylamine dehydrogenase59 and UV-B sensing protein in plants.60 In contrast to FgrGalOx, the Colletorichum graminicola and Colletotrichum gloeosporioides aliphatic alcohol oxidases do not possess any CBM,46 while the Colletorichum 9 graminicola aryl alcohol and raffinose oxidases have a PAN domain present instead.48, 53 In addition, the fusion of a galactose oxidase with a CBM29 has shown an increase in catalytic efficiency of the construct on galactose-containing hemicelluloses compared to WT.61 Figure 1.4. Crystal structure of copper radical oxidases FgrGalOx (PDB ID 1GOF), Copper ion in orange and B. CgrAlcOx (PDB ID 5C86), Copper ion in grey(adapted with permission from Yin et al. 201546) The redox-active center of AA5_2 oxidases is composed of a copper ion stabilized by two tyrosines and two histidines (in FgrGalOx, these are Tyr272, Tyr495, His496, His581; Figure 1.5), resulting in a distorted square pyramidal geometry.45, 62 AA5_2 proteins fall into the type 2 “non-blue” copper category of copper containing proteins due to their nitrogen and oxygen ligands,56 as opposed to type 1 copper “blue” proteins with nitrogen and sulfur ligands and type 3 copper proteins containing metal dimers.63 The very unique feature of this enzyme family is the covalently linked equatorial tyrosine, with an adjacent cysteine, by a thioether bond (Tyr 272 and Cys 228 in the archetypal 10 FgrGalOx) (Figure 1.5).56, 64 In the presence of cupric ions, Tyr272 is spontaneously cross-linked to Cys228, creating a tyrosyl-cysteine (Tyr-Cys) thioether linkage and a ring system that stabilizes the active [Cu2+ Tyr•] metallo-radical complex that forms in the presence of dioxygen.65 This cross-linking has been proposed to stabilize the radical though delocalization on the equatorial tyrosine during catalysis.45 Another interesting feature of AA5_2 members is the presence of a “stacking” tryptophan (W290 in FgrGalOx) whose indole ring lies above the covalently linked Tyr-Cys. This feature is thought to be crucial for reactivity by stabilizing the radical species either by delocalization and/or by protecting the thioether bond from solvent.66 Figure 1.5. Active site residues of copper radical oxidases FgrGalOx, Cu ion in green; B. Active site of CgrAlcOx overlayed with FgrGalOx, Cu ion in gray (adapted with permission from Yin et al. 201546) 1.3.2 Reaction Mechanism The oxidation of a primary alcohol to the corresponding aldehyde with concomitant reduction of oxygen to hydrogen peroxide is a two-electron process in AA5_2 oxidases that is mediated through the free-radical-coupled copper complex (Figure 1.6). The catalytic mechanism 11 of subfamily 2 progresses through a two-step process: the first half-reaction performs the oxidation of the substrate, while the second half-reaction regenerates the oxidation state of the active-site copper with concurrent reduction of molecular oxygen to hydrogen peroxide. The first half-reaction begins with a proton transfer (PT) from the substrate hydroxyl group to the oxygen of the axial tyrosine, followed by a hydrogen atom transfer (HAT) from the alkoxy carbon to the tyrosine-cysteine radical cofactor. Lastly, an electron is transferred (ET) from the substrate to generate the aldehyde while reducing the copper center to Cu(I). The second half-reaction involves the same steps as the first reaction but in the opposite order: an electron transfer from the now Cu(I) metal to a bound molecular oxygen molecule producing superoxide, a HAT from the cross-linked cysteine-tyrosine cofactor to the superoxide and finally a proton transfer from the axial tyrosine to the now hydroperoxide to yield hydrogen peroxide and the re-oxidized Cu(II) metal center that is ready to perform further turnover. Much spectroscopic evidence has been produced to support this proposed catalytic cycle.45, 62, 67, 68 Moreover, theoretical studies have provided evidence for this mechanism and related biomimetic models possess mechanistic similarities.69, 70 In the case of FgrGalOx, the enzyme exhibited no pH-dependent spectral change and the proton transfer was instead sensitive to temperature.71 Other work has focused on identifying parameters to achieve optimal activity of CROs to inform further process optimization for industrial utilization. FgrGalOx has been shown to have increased activity with the addition of peroxidases such as horse-radish peroxidase (HRP) and catalase72 and potassium ferricyanide has also been identified as a small molecule activator.73 A comparative work summarizing the effect of pH, oxygen saturation, and the source of a co-oxidant, either HRP, potassium ferricyanide or magnesium (III) acetate has been explored on galactose oxidase variants74, 75 and on CgrAlcOx.76 12 Figure 1.6. Reaction mechanism of copper radical oxidases A. First half-reaction – oxidation of substrate. B. Second-half reaction – regeneration of active site radical. PT – proton transfer, HAT – hydrogen atom transfer, ET – electron transfer (based on Humphreys et al. 200968) 1.3.3 Redox Potential A particularly interesting feature of AA5_2 enzymes is the presence of a redox active Tyr-Cys cofactor which exhibits an unusually low reduction potential (+275 mV)77-79 compared to unmodified tyrosine in solution (> +800 mV) or in other enzymatic systems.80 This is thought to be accomplished through several factors including π-stacking with aromatic residues and the electron donating effect of the thioether linkage, increasing the stability of the protein free radical.45, 66, 81 The presence of the Cys-Tyr cofactor showed only a minimal effect on the inner sphere of the copper atom coordination geometry in FgrGalOx, but facilitates delocalization of the redox active orbital onto the thioether sulfur from Cys228, thereby reducing the spin coupling between the Cys-Tyr radical and the Cu(II) site.82 Of note, in glyoxal oxidases (family AA5_1), the Tyr is natively a His instead,83, 84 and has a reduction potential of around +640 mV.44 This difference in oxidizing power could explain why FgrGalOx typically acts on hydroxyl groups 13 yielding aldehyde products, whereas the main activity of glyoxal oxidases is to convert aldehydes to carboxylic acids.79 In FgrGalOx, a Trp290His substitution increased the redox potential of the resulting enzyme from +400 mV to +730 mV;85 however, it also decreased the catalytic efficiency by 1000-fold67 and perturbed the stability of the [Cu2+ Tyr•] metallo-radical complex at neutral pH.86 The novel CgrAlcOx and CgrAAO are speculated to have a higher reduction potential than FgrGalOx due to their “stacking” amino acid substitution.46, 48 1.3.4 Substrate Specificity Currently, AA5_2 enzymes are classified as galactose oxidases based on the archetypal galactose oxidase from Fusarium graminearum, which catalyzes the regioselective oxidation of the C6-hydroxyl group on the monosaccharide galactose.87 Besides its ability to oxidize the 6-hydroxyl of D-galactose and galactosamine,88 the range of substrates targeted by FgrGalOx includes galactose derivatives such as 1-methyl--D-galactopyranoside,89 and galactose-containing oligo-and polysaccharides including: lactose, melibiose, raffinose, xyloglucan, galactomannan and galactoglucomannan (Figure 1.7).90, 91 The specific pattern of galactose branching and its relative proportion to other sugars within the same carbohydrate influence their enzymatic oxidation efficiency. Guar gum, locust bean gum, spruce galactomannan, tamarind xyloglucan, corn arabinoxylan and larch arabinogalactan have all been tested and are substrates of FgrGalOx. Although turnover (kcat) values of FgrGalOx for polysaccharides are 100 times lower for D-galactose, the enzyme shows substantially higher catalytic efficiencies (kcat/KM) on polysaccharides, due to comparatively low affinity (KM) values.61 Some sugar alcohols have also been tested such as D-threitol and xylitol with 10-15% yields,92 as well as 3,3-difluoro-4,4,-difluoro-galactose (3,3,4,4-tetrafluoro-6-(hydroxymethyl)tetrahydro-pyran-2,5-diol).93 14 Figure 1.7. Substrates for archetypal galactose oxidase (FgrGalOx) The broad range of galactose-containing oligo- and polysaccharides accepted by FgrGalOx is partly explained by the relatively open substrate-binding domain and shallow active site of this enzyme.46, 64, 94 At the same time, the strict regioselectivity of FgrGalOx towards the C6 primary hydroxyl group of galactose and terminal galactosyl residues in polysaccharides has been attributed to the hydrogen bonding between Arg330 and the axial orientation of the galactosyl C4 hydroxyl in accepted substrates.64, 95 Studies have been performed on the archetypal FgrGalOx to understand its remarkable selectively for galactose. Rogers et al. 2007 have shown that when the “stacking” tryptophan in position 290 is mutated to a phenylalanine, the KM of the enzyme for its natural substrate galactose increased 36-fold, demonstrating a significant loss of catalytic efficiency due to less efficient binding of the substrate.66 This has been hypothesized to be due to the loss of a key hydrogen bonding interaction between the indole nitrogen of the tryptophan and 15 the oxygen in the galactose ring. Furthermore, this specific residue participates in π-π stacking with the cysteine-tyrosine cofactor that is crucial for the oxidative mechanism of the enzyme to shield the radical from exposure to solvent.45 A variety of different mutagenesis and directed evolution experiments have been performed to increase the catalytic efficiency of the archetypal FgrGalOx96, 97, ultimately culminating in increased thermostability and activity with the variant M1 (harboring the P136, S10P, M70V, P136, G195E, V494A and N535D mutations)98 or multiple variants including mutations C383S, V494A, Y436H in single, double and triple mutant combinations.99 Various engineering efforts have been performed on FgrGalOx, as most recently reviewed.90 Many studies have been conducted in attempts to change substrate specificity through key point mutations. For instance, by combining structure-guided approaches and combinatorial library construction with random and saturation mutagenesis, Sun et al. (2002) created a variant that oxidized D-glucose and various alcohols, including aliphatic alcohols, while the wild-type enzyme was essentially inactive on these substrates.100 This variant, named M3, (aka M-RQW for the mutations R330K, Q406T, and W290F) has an activity on D-glucose 1000-fold higher than the wild type; however, its activity on D-galactose was reduced by more than 1000 times.100 Another variant produced by combining structure-based computational protein design with library-based enzyme screening resulted in Des3-2 (for the mutations Q326E, Y329K and R330K) exhibiting a 4-fold increase in specific activity towards glucose, and a 224-fold increase for galactose compared to M3.95 Furthermore, the mutations R330A or R330K produced variants with increased catalytic efficiency on fructose, at the expense of galactose.101 Therefore, R330 is implicated in substrate recognition, which accounts for its crucial role in modulating the substrate specificity of these mutants. 16 Conversely, a C383S mutation increases the enzyme catalytic efficiency towards guar gum and methyl galactose by possibly causing an increase in the hydrogen bonding network at the substrate binding site, which could reduce the KM.97, 99 Other studies have identified FgrGalOx mutants with increased activity towards mannose (variant H1 with R330K mutation in M1 background)102 and N-acetylglucosamine (variant F2 with R330K, W290F, Q406E, Y405F mutations in M1 background).102 Besides carbohydrates, variants of FgrGalOx have been engineered to oxidize secondary alcohols (variant M3-5 similar to M3 with a methionine at the 330 position)103 and amino alcohols (variants F2 and M3-5).104 Variant M3-5 has also been shown to oxidize some benzylic alcohols to the corresponding carboxylic acid.105 As these data indicate, a plethora of mutagenesis studies have been conducted on FgrGalOx, but efforts to expand its activity beyond galactose have generally had limited success. Several other AA5_2 members from the Fusarium genera, such as F. oxysporum106 and F. acuminatum107 showed substrate preferences comparable to FgrGalOx. However, in recent years, AA5_2 oxidases with distinct substrate profiles have been discovered. Most notably, two AA5_2 enzymes from Colletotrichum graminicola and Colletotrichum gloeosporioides (CgrAlcOx and CglAlcOx) were essentially inactive on galactose and galactosides, but efficiently oxidized the primary hydroxyl of diverse aliphatic alcohols (Figure 1.8).46 Specifically, CgrAlcOx has been found to exhibit high specificity towards aliphatic alcohols such as 1-butanol and was also highly active on benzyl alcohol, demonstrating almost complete conversion to the aldehyde product.46 Moreover, CgrAlcOx also catalyses the oxidation of cinnamyl alcohol, which has a pungent fragrance. The oxidation of aromatic alcohols by CgrAlcOx has biocatalytic potential, since the utilization of enzymes within the food and fragrance industry has been a growing trend.24 Overall, these two alcohol oxidases (AlcOx) have similar tertiary structures and active site architectures as 17 the classical FgrGalOx, but the most notable differences are the absence of the family 32 carbohydrate-binding module (CBM 32) and a tryptophan-to-phenylalanine substitution in the “stacking” residue. In addition, if this phenylalanine in CgrAlcOx is substituted for a tryptophan almost all activity on alkanols is lost but the ability to more effectively oxidize conjugated alcohols is gained.46 This mutation indicates that this aromatic residue likely plays an important role in the substrate specificity of these enzymes, but it alone does not determine the selectivity of their reactivity. Similarly, CgrRafOx from Colletotrichum graminicola showed only negligible activity on galactose, and in this enzyme the N-terminal CBM32 of FgrGalOx is replaced by a PAN_1 domain (PF00024). CgrRafOx was not active towards aliphatic alcohols, however, it showed the highest activity for tri- and di-saccharides such as raffinose and melibiose, along with the glycolaldehyde dimer.53 More recently, the last of three AA5_2 found within C. graminicola genome has been characterized and classified as an aryl-alcohol oxidase (Figure 1.8), an activity restricted only to AA3_2 members so far. Moreover, this enzyme has a high specific activity towards 5-hydroxymethylfurfural, a compound generated during biomass treatment for biofuel production known to be toxic for microbes used in bioprocesses.48 In these newly discovered CROs (CgrRafOx and CgrAAO), the “stacking” active residue is a tyrosine instead of a tryptophan (as in FgrGalOx) or a phenylalanine (as in CgrAlcOx). While the role of this tyrosine has not been investigated in CgrRafOx, in CgrAAO its substitution by a phenylalanine directed its oxidative activity towards aryl alcohols and furans rather than carbohydrates and its substitution by a tryptophan modulated its preference towards carbohydrates.48 Two alcohol oxidases were characterized from the rice blast pathogen Pyricularia oryzae (PoAlcOx) and from Colletotrichum higginsianum (ChAlcOx) with both enzymes possessing 18 prominent activity on n-butanol, ethanol, 1,3-butanediol and glycerol.47 PoAlcOx contained a WSC domain that was able to bind xylans and fungal chitin/β-1,3-glucan, implicating the domains involvement in enzyme anchoring on the plant surface.47 Figure 1.8 Substrates for CgrAlcOx (A.) and CgrAAO (B.) Finally, a recently characterized CRO from Penicillium rubens Wisconsin 54–1255 (PruAA5_2A) has been shown to have the highest carbohydrate oxidase activity on the galactose-containing oligosaccharide raffinose, but also had the highest catalytic efficiency on glycolaldehyde dimer, typically a substrate for AA5 subfamily 1 enzymes.54 Due to this broad substrate profile the authors concluded that this enzyme may be a functional intermediate with an overlap of activities across the AA5 substrates.54 PruAA5_2A also has a serine at position 383 and an alanine at position 436 which have been shown to be important for enhanced catalysis.97 1.3.5 Biological Role The biological role of the AA5 subfamily 2 enzymes has remained elusive even though the mechanism of their activity has been extensively studied. These copper radical oxidases produce hydrogen peroxide during their catalytic cycle and this has been speculated to be coupled to 19 oxidative lignocellulosic degradation.84, 108 This process occurs using secreted peroxidases in white rot fungi or non-enzymatic Fenton chemistry in brown rot fungi.84 According to the CAZy database43l, AA5_2 sequences have been identified in one archaea species, 4 pathogenic species in the genus Burkholderia and other bacterial strains, and most sequences are found in many fungal plant pathogens, some of which attack food crops such as wheat and barley (Fusarium graminearum),109 potatoes (Fusarium sambucimum),110 rice (Fusarium fujikuroi111 and Magnaporthe oryzae112), maize (Fusarium verticilloides113 and Colletotrichum graminicola114), banana (Fusarium oxysporum f. sp. Cubense)115 and others. Characterized AA5 subfamily 2 enzymes have all been found in fungal saprotrophs and phytopathogens. The characterized galactose oxidases from this subfamily all come from a variety of species in the genus Fusarium, more specifically: the archetypal GalOx from Fusarium graminearum,51 Fusarium oxysporum,106 and Fusarium sambucinum.89 Characterized enzymes with novel activities all come from the genus Colletotrichum and Pyricularia. Specifically, aliphatic alcohol oxidases were from Colletotrichum graminicola, Colletotrichum gloisporioides,46 Colletotrichum higginsianum and Pyricularia oryzae,47 while the raffinose and aryl-alcohol oxidases were both from Colletotrichum graminicola.48, 53 AA5 enzymes have been speculated also to be important for fungal virulence. However, to our knowledge, no deletion study has been performed for any AA5_2 member. In 2005, Leuthner et al. showed that the deletion of the AA5_1 member glyoxal oxidase abolished the pathogenicity of Ustilago maydis.108 In addition, the deletion of a glyoxal oxidase from Streptomyces lividans showed loss of glycan accumulation at hyphal tips.116 This further indicates the importance of AA5 enzymes in the infectious behaviour of fungi. A putative galactose oxidase has been identified in 20 an allergen from Artemisia pollen.117 Furthermore, a putative galactose oxidase, RUBY, has been demonstrated to promote the cell-to-cell adhesion in the seed coat epidermis of Arabidopsis.118 1.3.6 Applications Glycosylation is a common posttranslational modification and glycoconjugates are an integral component of the immune system, hence the profiling of glycosylation is a critical area of research.119, 120 However, the diversity of glycans on proteins and the non-templated nature of glycan biosynthesis creates the need for multitude of labeling techniques. The first use of the archetypal FgrGalOx was in 1965 for the histochemical examination of mucus-secreting cells in bovine specimens.121 Since then, the use of FgrGalOx oxidative properties in glycoproteins labelling has been an extensive area of research for multiple decades. FgrGalOx mediated oxidation and subsequent conjugation with biotin hydrazide,122, 123 rhodamine and fluorescein containing hydrazides124 or by radio-labelling tritiated sodium borohydride (NaB3H4),125 has been used to visualize the terminal galactosyl and N-acetylgalactosamine residues on cells. A strategy that uses galactose oxidase treatment followed by aniline-catalyzed oxime ligation (GAL) has also been employed for glycoprotein labelling.126 Furthermore, the use of FgrGalOx variants has been explored for the labelling of galactose-containing carbohydrates with M1, mannose-containing carbohydrates with H1 and N-acetylglucosamine with F2.102 H1 has also been used in labeling cells that express mannosylated glycoproteins on their surface.102 As well, the human xeno-autoantigen N-glycolylneuraminic acid (Neu5Gc) has been selectively labeled using the mutant F2 and attached to a complex glycoprotein with similar glycans present.127 Functionalization of polysaccharides using carbohydrate oxidases was first presented over three decades ago, when various derivatives of hemicellulosic polysaccharides were prepared 21 using FgrGalOx.128, 129 In addition, primary alcohol oxidation mediated by AA5_2 enzymes generates a reactive aldehyde that allows further modifications to produce a variety of compounds such as uronic acids,130, 131 indium-mediated allylation/propargylation,132, 133 and deoxy sugars131, 133 that have serve as platform for further modifications. A beneficial characteristic of these manipulations is that they can be performed right after the enzymatic oxidation in a one-pot reaction.90 The use of FgrGalOx to functionalize polysaccharides has been successful for the development of clickable cellulose surfaces,134 carboxylated polysaccharides as highly reactive galactoaldehydes,135 and selectively allylated polysaccharides.132 Due to the intrinsic affinity of hemicellulose to cellulose, the polysaccharide derivatives were adsorbed to functionalize the cellulosic surfaces, providing the basis for advanced biomaterials with predefined properties. Enzymatically oxidized polysaccharides have also been used for the creation of aerogels relevant to food and medical applications.136 Furthermore, a biosensor for esterase activity has been developed through oxidation of xyloglucan by FgrGalOx followed by propargylation.137 The modified carbohydrate is then adsorbed to a cellulosic surface and the biosensor was attached via click-chemistry.137 FgrGalOx has also been used as a sensing tool in multiple applications. Galactose oxidase, in concert with glucose oxidase and pyranose 2-oxidase, has been used for the removal of molecular oxygen from water and soil samples used in the monitoring of nitrate ions.138 FgrGalOx has been used in biosensing for the detection of lactose in milk and other dairy products.92 Many biosensors have been produced by immobilizing galactose oxidase on a variety of surfaces.139-141 The catalytic promiscuity of galactose oxidase has also been used as a green method for the enzymatic one‐pot conversion of alcohols into the corresponding nitriles.142 Furthermore, variant M3-5 of FgrGalOx has been used in an enzyme cascade reaction for the synthesis of 22 furandicarboxylic acid (FDCA) from the biomass derived hydroxy-methyl furfural (HMF) with the addition of an aldehyde oxidase in a one-pot tandem reaction.143 A highly evolved version of FgrGalOx to obtain the reverse stereo-selectivity has been used in the pharmaceutical industry, showcased by the industrial biocatalytic cascade for the synthesis of the HIV drug treatment islatravir.144 Process requirements of the galactose oxidase mutant M3-5, specifically the presence of accessory enzymes, choice of buffer, and oxygen supply, have been identified and optimized for the oxidation of alcohols on an industrial scale.74 1.4 Enzyme Immobilization Enzymes can be costly to produce and difficult to recover from an aqueous medium after performing catalysis, leading to their slow adoption by industry. One strategy to improve the industrial applicability and cost-effectiveness of enzymatic catalysts is to convert them from homogenous to heterogenous catalysts through immobilization.26 This technique can increase enzyme thermostability and stability in non-aqueous solvents, while allowing for its recovery and re-use while maintaining its catalytic activity. Furthermore, enzyme immobilization can enable new applications that were previously not chemically or economically viable. The first industrial application of immobilized enzymes was in the production of amino acids (L-methionine, L-phenylalanine, and L-valine) in Japan in 1969.145 Since then, two important industrial milestones were the use of immobilized glucose isomerase and penicillin amidase for large-scale applications.146, 147 Glucose isomerase converts glucose to fructose and has been used in one of the most commercially significant advances for the production of high-fructose corn syrup (HFCS). Prior to this breakthrough in 1976, most sucrose was sourced from sugar beets and sugar cane. To this day, glucose isomerase still controls the biggest share of the food industry market. Two main 23 methods are used in the industrial immobilization of glucose isomerase: cell-free enzyme immobilization and whole-cell immobilization. Since glucose isomerase is an intracellular protein, most companies (Nagase, Solvay, Novo-Nordisk and others) use different whole-cell immobilization strategies including heat-treated cells bound to anion exchange resin, cells occluded in gelatin followed by glutaraldehyde, polyamine glutaraldehyde cross-linked cells that are extruded and granulated, chitosan-treated glutaraldehyde cross-linked cells and glutaraldehyde cross-linked cells that have been extruded.146 Conversely, some cell free methods exist for HFCS production utilizing a variety of carrier-resins and chemical cross-linking techniques. Commercial examples include CPC (Enzyme Biosystems) usage of an anion-exchange resin, Clinton Corn Processing Company utilization of glucose isomerase immobilized on DEAE-cellulose, Genencor International usage of DEAE-cellulose agglomerated with polystyrene and TiO2, Solvay adsorption of glucose isomerase onto SiO2 particles followed by cross-linking with glutaraldehyde and Honeywell Universal Oil Products utilization of polyethyleneimine-treated alumina with glutaraldehyde cross-linked enzyme.146 The other enzyme that has attracted significant attention is the penicillin amidase. During the 1960s, the medical community encountered infections that were resistant to the antibiotic penicillin, and pharmaceutical companies explored penicillin derivatives to combat this problem. At that time, the chemical processes used for the hydrolysis of penicillin to acquire its backbone structure were tedious, complex and required large amount of toxic and expensive chemicals. An enzymatic route for hydrolysis was more elegant had not been performed at a large scale up to this point. The original industrial patent for penicillin amidase immobilization by Bayer used a copolymer composed of acrylamide, NN′-methylene bisacrylamide and maleic acid in the presence of propionitrile and ammonium peroxydisulfate at pH 7.6 under nitrogen.147 This 24 polymerization strategy had excellent properties including, the high yield of active enzyme bound (92%), increased stability of the biocatalyst (>> 30 batches), and a high yield of the target product 6-aminopenicillanic acid (91%). 1.4.1 Immobilization Methods Many different methods for enzyme immobilization have been studied and different perspectives for their classification exist. Some methods overlap in properties and often they could be combined to enhance maximum catalytic productivity. Furthermore, the choice of the immobilization method depends on the overall process and a range of methodologies should be explored depending on the enzyme and its application. Some important immobilization methods include: adsorption, entrapment/encapsulation, cross-linking to a surface and cross-linked enzyme aggregates (Figure 1.9). Figure 1.9. Enzyme immobilization methods (re-used with permission from Rehm et al. 2016148) 25 Adsorption of an enzyme onto a carrier surface is one the oldest methods for enzyme immobilization.149 Its simplicity allows it to be performed under mild conditions with the potential for reversibility. A variety of different non-covalent interactions such as physical adsorption, ionic bonding, coordination chemistry though metal binding and affinity adsorption can be exploited through this method.150 However, the inherently weak interactions involved in this method often causes enzyme leakage from the carrier due to the use of water, high ionic strength buffers, high or low pH and other harsh industrial conditions.26, 150 Entrapment/encapsulation of an enzyme involves inclusion of the catalyst in an organic or inorganic polymer matrix.150 Some of the earliest examples of enzyme entrapment used gel-like matrixes.151, 152 Generally, whole cells or purified enzymes are dispersed into the polymer as an immiscible medium followed by gelation or polymerization. The correct pore size of the polymer matrix is of great importance in order to diminish enzyme leakage from the entrapped particles, and to allow for the efficient diffusion of the substrates/products in and out of the matrix.150 Most entrapment immobilization strategies occur in water-based environments and can cause high swelling and some matrixes are not suitable for industrial conditions. Furthermore, physical constraints are often too weak to prevent enzyme leakage and covalent attachments are necessary. Cross-linking an enzyme to a surface is one of the most used methodologies for enzyme immobilization. Covalent bonds provide a strong linkage between the enzyme and the carrier, diminishing the potential for enzyme leakage. This is an important consideration owing to the cost of enzyme production in an industrial context. Generally in this method, the covalent bond is between an active amino group on the enzyme to an active functional group on the carrier.150 This allows for a vast variety of combinations and design opportunities between an assortment of 26 carrier options with wide chemical and physical diversity that needs to be tailored to each enzyme of interest. Due to the complexity of this method, many considerations must be addressed for each enzyme immobilization project such as the physical and chemical properties of the carrier, the enzyme orientation and conformation after immobilization, the chemistry and number of linkages between the enzyme and the carrier, and the enzyme distribution on/within the carriers.150 Covalent cross-linking for the purpose of enzyme immobilization must always balance the loss of activity due to diffusion constraints of a heterogenous catalyst versus the increased stability and potential for catalyst recyclability. Common supports to this methodology include synthetic polymers, biopolymers and inorganic solids such as silicas.153, 154 A variety of different carriers exploiting different chemical functionalities including polyanhydride, polycarbonate, polyaldehyde, polyepoxide, polyisocyanate and others have been synthesized. A specific example of the first commercial synthetic oxirane polymer used for enzyme immobilization was Eupergit C. Eupergit C is a solid support made up of 100-250 μm beads synthesized by the copolymerization of N,N-methylene-bis-(methacrylamide), glycidyl methacrylate, allyl glycidyl ether and methacrylamide.155 Eupergit C is a chemically and mechanically stable material of which several tons are employed per year in various industrial biocatalytic processes.155, 156 However, decreased activity caused by introduction of a non-active support can contribute to lower space-time yields of the catalyst.157 A carrier-free method with stable covalent cross-linking can overcome some of these challenges. Cross-linking enzyme aggregates (CLEA) is a simple method which involves the precipitation of enzymes and subsequent cross-linking, often with glutaraldehyde, to form enzyme aggregates. This is the primary methodology used in Chapter 4 and will be discussed in detail in the next section. 27 1.4.2 Cross-Linked Enzyme Aggregates (CLEA) Cross-linking of soluble enzymes was first reported in the 1960s, but the resulting particles were often unstable, had low activity and were difficult to reproduce.26 In the early 1990s, a technique to make cross-linked enzyme crystals (CLEC) was introduced for the potential production of industrial biocatalysts.158 CLECs had good stability, high activity and were easy to re-use. However, the requirement for enzyme crystallization limited its broader application. An alternative method called CLEA was developed to increased enzyme stabilization, retention of activity and the reproducibility of enzyme preparations via their precipitation from solution without affecting the tertiary structure of the catalysts, followed by cross-linking (Figure 1.10). This methodology was first used on penicillin G acylase for the synthesis of ampicillin in a broad range of organic solvents.159 Figure 1.10. Cross-linking enzyme aggregates procedure (adapted with permission from Yamaguchi et al. 2018160) Optimization of CLEAs preparation for a novel enzyme includes adjusting the choice of precipitant and cross-linker, length of cross-linking, and the ratio of cross-linker to enzyme. Different precipitants can be used such as salts, organic solvents and non-ionic polymers and can induce distinct enzyme conformations and have an effect on the catalytic activity of the CLEA 28 particles.161 Different cross-linking reagents can be used for the preparation of CLEAs but the most common is glutaraldehyde.26 This small molecule reacts with the amino groups on the enzyme surface and forms a covalent bond between enzymes. Other cross-linking reagents have been proposed for CLEAs with low activity, such as dextran polyaldehyde162 or polyamines for enzymes possessing a few surface lysines.163 The degree of cross-linking in the CLEA particle can affect its catalytic activity and can be modulated by tuning the ratio of cross-linker to enzyme.164 Too little cross-linker can cause enzyme leaching from the CLEA particle, while too much can cause the enzymes to be over cross-linked and diminish their activity due to a decreased flexibility and hindered access to the active site.165 Immobilization of enzymes using CLEAs has been shown to retain enzyme activity in many cases, however, loss of activity can also be observed. Some preparations have utilized additional additives such as Triton X166 or bovine serum albumin (BSA)167 to help retain the enzymatic activity of the CLEAs. Nevertheless, the benefit of increased stability, tolerance to organic solvent and reusability can outweigh the loss of enzyme activity when forming CLEA particles. Enzymes immobilized using CLEA have shown to possess increased stability, shelf life,26, 165, 168, 169 and thermostability.170-172 For example, penicillin G acylase CLEAs prepared with tert-butanol displayed higher thermostability compared to when DME or PEG were used as precipitants.173 Furthermore, CLEAs preparation can broaden the pH stability range of enzymes, which can be important for certain applications. CLEAs of a peroxidase from Brassica rapa were shown to be active at more acidic pH values than the free enzyme,174 while glutamate decarboxylase CLEAs were able to remain active at more alkaline pH, allowing for higher production of γ-aminobutyric acid.175 Solvent stability is also a benefit from the creation of 29 CLEA particles.176 Lastly, storage stability has been increased with CLEA technology.177-179 For example, CLEAs made with amyloglucosidase retained 98% of their initial activity after 2 months of storage at 4 oC compared to the free enzyme that lost more than 50% of its initial activity after only 5 days using the same storage conditions.180 Lastly, there are many examples in the literature of CLEAs maintaining activity over multiple re-use cycles.26, 164, 165, 168, 169 A variety of different enzyme classes have been immobilized with CLEAs such as hydrolases, proteases, amidases, esterase, nitrilases, oxidoreductases, lyases and others.168, 169 Of specific note are the many CLEA particles formed with serine proteases for organic synthesis applications181-183 and the various preparations of lipase CLEAs.184-186 Some oxidoreductases have also been incorporated into CLEA particles such as: glucose oxidase and galactose oxidase.131, 187 In addition, enzyme immobilization using CLEAs can be done directly from a cell lysate without further purification since the first step in the preparation is precipitation, which is an established technique for enzyme purification. Immobilization of enzymes using CLEAs can be more cost effective for an industrial application since this methodology is carrier-free, eliminating the cost of a catalytically inactive support.168 Furthermore, diffusion limitations are not a significant issue for CLEAs due to the porosity of the particles and the tunability of particle size by variation in the enzyme to cross-linker ratio.165 One drawback of the CLEA methodology is that it must be optimized for each enzyme of interest; however, this process could be automated. In addition, resulting CLEA particles are often small (<50 μm), which can be an issue in some industrial applications. Overall, CLEAs allow for a more cost-effective process due to being carrier-free, the particles generally have 30 increased stability to heat and organic solvents, do not leach catalysts in aqueous media and are easily recyclable. Due to the simplicity of this cross-linking method, multiple enzymes can be co-immobilized together to form combi-CLEAs. Combi-CLEAs are useful for applications involving enzyme cascade reactions that can be performed in one-pot. Combi-CLEAs often display better results when compared to the soluble enzyme counterparts due to the increased proximity of intermediates to the second enzyme. Combi-CLEAs have been utilized in different synthetic cascade reactions such as the resolution of S-mandelic acid,188, 189 production of α-hydroxycarboxylic amides190 and the production of alkyl ferulates.191 Additional examples include a mixture of pectinase, xylanase and cellulase that have been co-immobilized for a potential use in cellulosic bio-fuel production and the one-pot biomass conversion of sucrose to trehalose using tri-enzyme combi-CLEAs.192, 193 Other applications of combi-CLEAs, besides organic synthesis, are the biodegradation of acetaminophen in waste water employing a laccase and tyrosinase enzyme mixture194 and the co-immobilization of a peroxidase with a glucose oxidase for removal of endocrine disruptors.195 Further developments in the field of CLEAs include the introduction of magnetic CLEAs, incorporation of nano-particles and the use of supports for CLEAs.26, 164, 165, 169 1.5 Thesis Aims The central aims of this thesis are to characterize homologs belonging to the Auxiliary activity family 5 subfamily 2 enzymes, to investigate their biotechnological applications, and to improve their potential in biocatalysis using immobilization strategies. At the start of this project, only 5% of AA5_2 enzymes had been characterized from the publicly available sequences found in the CAZy database. Within the characterized members, two distinct enzymatic functions were 31 observed: galactose oxidase (FgrGalOx) and aliphatic/aromatic alcohol oxidase (CgrAlcOx). In order to develop a framework for the structural determination of their substrate specificity, more members of the AA5_2 subfamily needed to be produced and biochemically characterized. Bioinformatic analysis aided in selection of sequences targeted for production. The conventional ABTS-HRP assay for copper radical oxidase was utilized for the detailed biochemical characterization of AA5_2 homologs. Sequence information was analyzed in conjunction with kinetic data and docking studies to inform structure-function relationships. Characterization techniques were employed to assess the oxidative product of the novel CROs for their biotechnological potential. Lastly, enzyme immobilization using the cross-linking enzyme aggregate (CLEA) method was employed to improve the biocatalytic potential of CROs, specifically the Colletotrichum graminicola alcohol oxidase (CgrAlcOx). The work presented in this thesis demonstrates the diversity of activity found within the AA5_2 subfamily and offers potential avenues for further biocatalytic applications of CROs. 32 Chapter 2: Two Fusarium Copper Radical Oxidases with High Activity on Aryl Alcohols 2.1 Introduction Our dependency on non-renewable resources and the associated environmental consequences pose as an existential crisis to human society.3, 2 One avenue to reduce consumption of fossil petroleum is to utilize biomass as a renewable, alternative carbon source. In this regard, the development of integrated biorefineries that produce multiple products, including fuels, chemicals, and materials (collectively “bio-products”) would advance the goal to lower net carbon emissions.4, 5 In particular, harnessing the inherent chemical functionality of biomass constituents to produce high-value bio-products through selective derivatization is currently envisioned as a better value proposition than complete saccharification and fermentation to commodity fuels.10 In nature, terrestrial biomass is efficiently transformed and degraded by fungi via a plethora of specialized enzymes that could be harnessed in biocatalytic processes for the industrial production of bio-products.13-15 In particular, fungi have a rich history as sources of hydrolytic enzymes for biomass saccharification.19, 196-198 Fungi also produce a diversity of redox enzymes, which act on a wide range of substrates.43 Notably, there is increasing interest in the use of redox enzymes to replace chemical oxidants, many of which generate hazardous waste co-products. Indeed, bio-oxidation reactions account for one-sixth of the biocatalysis performed industrially, including hydroxylation, Bayer-Villiger catalysis, alcohol and amine oxidation, and other transformation.14, 27 Presently, the majority of these transformations are performed by 33 FAD- and NAD(P)-dependent enzymes, which require complex organic cofactors, and in the latter case, cofactor recycling strategies.13 In contrast, copper-radical oxidases (CROs),84 have received considerably less attention regarding their potential as industrial biocatalysts.14, 199 CROs comprise Auxiliary Activity Family 5 in the Carbohydrate-Active Enzymes (CAZy) and thus share similar tertiary structures and active sites containing a mononuclear copper center.46, 48, 56, 84 The AA5 family has been further divided into two subfamilies based on molecular phylogeny. Subfamily 1 (AA5_1) is comprised of (methyl)glyoxal oxidases (E.C. 1.2.3.15) that oxidize select aldehydes, likely via the hydrated gem-diol, to the corresponding carboxylic acid with concomitant reduction of O2 to H2O2 44. Subfamily 2 (AA5_2) contains the founding galactose oxidases (GalOx, E.C.1.1.3.9)45, 50, 51, 87, 200 as well as the more recently discovered general alcohol oxidases (AlcOx, E.C. 1.1.3.13),46, 47 and aryl alcohol oxidases (AAO, E.C. 1.1.3.7),48 all of which convert the primary alcohol of the substrate to the corresponding aldehyde in an analogous two electron oxidation. The specific biological roles of CROs are currently unknown, although they have been speculated to play a role in oxidative lignocellulose degradation and have been linked to fungal pathogenesis.84, 108 The archetypal galactose oxidase from the phytopathogen Fusarium graminearum (FgrGalOx) has been used in many biotechnological applications such as glycoprotein labelling, 121, 122, 124, 126 construction of lactose biosensors,92,chemo-enzymatic modification of galactose and galactosides,131 and complex polysaccharide modification for the development of functional materials.128, 130, 132, 133, 136 As the only characterized member of AA5 for many years, FgrGalOx has been the subject of many protein engineering and directed evolution experiments to broaden substrate scope to include activity on glucose,100, 95 fructose,101 mannose,102 N-34 acetylglucosamine,102 secondary alcohols,103 amino alcohols,201 and benzyl alcohols.105 Other studies have been performed to increase the catalytic efficiency of FgrGalOx.96-99 Especially notable, a highly evolved variant of FgrGalOx was central to the recent industrial development of a biocatalytic synthesis of the drug islatravir.144 Thus, there is significant scope to expand the application space of CROs to meet the need for new oxidation catalysts dependent on substrate range.14, 199, 202, 203 The diversity of CROs from AA5 remains largely unexplored, but features a breadth of sequences from fungal sources that can be mined for potentially new industrial applications. Of the hundreds of publicly available putative AA5 sequences currently in the CAZy database,43 only ten AA5_2 members from the subfamily have been biochemically characterized, with the majority (five) characterized as galactose or galactoside oxidases (four from Fusarium species).53, 89, 106, 204 In this context, we have recently revealed novel non-carbohydrate-active CROs in AA5_2 from the phytopathogenic fungus Colletotrichum graminicola: CgrAlcOx displays high activity on primary aliphatic, benzyl, and other unsaturated alcohols,46 while CgrAAO is most active on a range of benzyl alcohols and is also active on hydroxymethylfurfural (HMF, EC 1.1.3.47).48 Intrigued by this broader catalytic potential of AA5_2, we have used molecular phylogeny to guide the selection of two distinct homologs from F. graminearum and F. oxysporum for recombinant expression and biochemical characterization. Notably, both enzymes had low activity on carbohydrates, but displayed high activity on aryl alcohols, such as veratryl alcohol, and have considerable activity on HMF and oxidized congeners. Detailed enzyme kinetic and product analysis, together with molecular modelling of substrate recognition, allowed us to rationalize this specificity profile with regard to the composition of the active site. 35 2.2 Results and Discussion 2.2.1 Bioinformatics Building upon our previous discovery of unique substrate specificities among Colletotrichum AA5_2 homologs,46, 48 we were keen to explore alternative homologs of AA5_2 members from Fusarium species, from which a number of galactose oxidases have been described,89, 106, 204 including the archetypal FgrGalOx.45, 56, 62, 66 A molecular phylogeny was generated to guide target selection, using 45 AA5_2 catalytic modules selected from the CAZy database,43 including previously characterized homologs from Fusarium and Colletotrichum, (Figure 2.1). We observed that a homolog from Fusarium graminearum (GenBank XP_011322138) grouped into a strongly supported clade that was notably distant from that containing FgrGalOx. This clade is also distinct from the early diverging clade comprising the Colletotrichum AlcOxs and AAO. To our knowledge, there are no biochemically characterized members of this clade,205 thus warranting further study. We also selected the homolog from Fusarium oxysporum (GenBank XP_018246910) for characterization, as a second example from this clade. Hereafter, these homologs will be referred to as FgrAAO and FoxAAO, respectively, based on subsequent substrate specificity analysis (vide infra). 36 Figure 2.1. Molecular phylogeny of 45 AA5_2 members The sequences of two AA5_1 (methyl)glyoxal oxidase catalytic modules108, 206 were included as an outgroup to root the tree (highlighted in pink). Clades containing characterized Colletotrichum enzymes are highlighted in blue, FgrAAO and FoxAAO in purple and FgrGalOx in green. GenBank Accessions are given for all sequences from the public CAZy database (February 2021) with the genus and species names of source organisms with E.C. numbers also listed. Data from previous studies comprise AAO95371 (FgrGalOx)56, 62, 66, AHA90705.1 (FoxGalOx)106, AIR07394.1 (FsaGalOx),89 AJE27923 (FsuGalOx),204 ADG08188.1 and ADG08187.1 (FveAA5 and FsuAA5),205 XP_011322138 and XP_018246910 (FgrAAO and FoxAAO, this study), EFQ27661.1 (CgrAAO)48, EFQ30446 and ELA25906.1 (CgrAlcOx and CglAlcOx),46 EFQ36699.1 (CgrRafOx),53 OBR05259.1 (ChiAlcOx) and XP_003719369.1 (PoAlcOx).47 37 Concordant with this phylogeny, a protein sequence alignment of FgrAAO and FoxAAO revealed that they are highly similar (91% pairwise identity, Table A1). FgrAAO and FoxAAO have ca. 60% pairwise identity versus the AA5 archetype FgrGalOx56, 64 and with characterized GalOx from F. oxysporum, F. sambucinum and F. subglutinans89, 106, 204 (Figure 2.1 and Table A1). On the other hand, FgrAAO and FoxAAO have low sequence similarity with the characterized Colletotrichum AA5 members, CgrAlcOx and CgrAAO (ca. 50% identity, Table A1). Particularly notable, FgrAAO and FoxAAO share high sequence identity (80%) with orthologs from F. verticillioides and F. subglutinans, which have been cloned but not characterized in recombinant form (Figure 2.1 and Table A1).205 It is worth underscoring that the phylogeny shown in Figure 2.1 was calculated based on AA5 catalytic modules only, and that FgrAAO and FoxAAO have the same modular organization as FgrGalOx, viz. an N-terminal carbohydrate-binding module (CBM32)207 in tandem with the AA5_2 module. In contrast, CgrAlcOx and CgrAAO lack an N-terminal carbohydrate-binding module (Figure 2.2). Detailed sequence analysis indicated that key active-site and secondary shell radical-stabilizing residues are largely conserved between FgrGalOx, FgrAAO and FoxAAO (Figure 2.2). A notable exception is the Arg-to-Lys substitution in FgrAAO and FoxAAO corresponding to K330 in FgrGalOx. Site-directed mutation of this residue in FgrGalOx has been shown to increase relative activities on fructose (in wild-type FgrGalOx)101 and mannose (in M1 variant),102 thus highlighting a role in substrate specificity. Mutation of the active-site residues G195 and Q326 in FgrGalOx has been shown to improve protein stability131 and alter substrate specificity102, respectively, while the distal residues C383, Y436, and V494 have been implicated in affecting catalytic efficiency.96-99 FgrAAO and 38 FoxAAO are distinguished from the wild-type FgrGalOx in these positions and differ from each other at position Y436 (Figure 2.2). Figure 2.2. Active site amino acids in AA5_2 members A. Key amino acid residues in the two candidate enzymes compared to other characterized AA5_2 copper radical oxidases. Non-conserved amino acids in relation to FgrGalOx are bolded. Fungal species: Fgr, Fusarium graminearum; Fox, Fusarium oxysporum; Cgr, Colletotrichum graminicola. B. Active site of FgrGalOx. Left panel - full protein, right panel - zoom of active site. Colours are coordinated with amino acid residues in Panel A. 39 2.2.2 Protein Production and Substrate Screening cDNA corresponding to the mature, full-length FgrAAO and FoxAAO proteins comprising the N-terminal CBM32 and C-terminal AA5_2 module (lacking the native signal peptide) were cloned into pPICZα-A vector in-frame with a Saccharomyces cerevisiae α-factor secretion signal peptide and a C-terminal hexa-histidine tag. Both proteins were successfully produced in Pichia pastoris X33 and purified by a two-step IMAC-SEC protocol, with typical yields of 1 mg of protein per 400 mL of expression medium. Notably, FoxAAO eluted as a single pure peak from the size exclusion column, whereas FgrAAO eluted as two peaks, the second of which contained pure enzyme, as determined by SDS-PAGE (Figure A2). Both protein production levels are significantly lower (200-fold) than recombinant wild-type FgrGalOx208 produced in P. pastoris. FgrAAO and FoxAAO (2.5 mg.L-1) produce similarly to FoxGalOx (10.6 mg.L-1)106 and had production values lower than wild-type FgrGalOx and FgrGalOx M1 constructs with histidine tags (120 mg.L-1 and 110 mg.L-1, respectively),209 CgrAlcOx (30-40 mg.L-1),46 and CgrAAO (42.5 mg.L-1)48. In light of the sequence similarity of FgrAAO and FoxAAO to FgrGalOx (Table A1), galactose was selected as an initial substrate for optimum pH and temperature determination. Both recombinant enzymes were active in the pH range from 5 – 10, with bell-shaped pH-rate profiles and optimum pH values of ca. 7.5 in sodium phosphate buffer (Figure A3). This pH optimum is similar to other characterized AA5_2 members.46, 48, 89, 106, 210 FgrAAO and FoxAAO were rapidly inactivated above 65 °C (Figure A4). Additional stability assays indicated that both enzymes were stable at 30 °C over ca. 6 h, but lost a significant amount of activity with extended incubation at this and higher temperatures (Figure A5). Overall, FgrAAO and FoxAAO appear to have lower thermostability than other AA5_2 members.46, 106 40 FgrAAO and FoxAAO were then screened against a broad panel of primary alcohols, including carbohydrates, alkanols, and aryl alcohols of biological and synthetic relevance (Figure 2.3 and Table A2). Concordant with our initial activity assays, galactose and the galactosylated di- and trisaccharides melibiose, lactose, and raffinose were competent substrates of FgrAAO and FoxAAO, with specific activities up to 3 mol min-1 mg-1 (Figure 2.3 and Table A2). In contrast, activity on xyloglucan or galactomannan, which bear terminal galactosyl (t-Gal) branch residues, was extremely limited. Strikingly, the highest monosaccharide activity was observed with mannose, while activity on the ketose, fructose, was comparable to that on galactose (Figure 2.3 and Table A2). Yet, very limited activity was detected on sucrose (α-D-glucopyranosyl-(1→2)-β-D-fructofuranoside) and β-(2→1) fructo-oligosaccharides. 41 Figure 2.3. Initial activity screens of FgrAAO and FoxAAO Measurements were performed in triplicate at 25 °C in 50 mM sodium phosphate buffer, pH 7.5, using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 2.5 mg mL-1 for polysaccharides, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal, aryl alcohols and furans and 10 mM for secondary alcohols. Reactions were started with the addition of 6 pmol – 1 nmole of purified enzyme (Table A2). Primary alcohols, including a range of alkanols, diols, and alditols were generally not substrates for FgrAAO and FoxAAO, although glycerol was a competent substrate with a specific activity of 1.5 μmol min-1 mg-1 (one-half that of galactose). Although benzyl alcohol and cinnamyl alcohol were substrates for both enzymes, with cinnamyl alcohol preferred, the highest activities for FgrAAO and FoxAAO were observed with the substituted aryl alcohols, 3-42 methoxybenzyl alcohol (m-anisyl alcohol) and veratryl alcohol (3,4-dimethoxybenzyl alcohol). At the same time, 4-hydroxybenzyl alcohols (i.e., 4-hydroxybenzyl, vanillyl, and coniferyl alcohols) were not tolerated by either enzyme, and low activity on 4-methoxybenzyl alcohol (p-anisyl alcohol) was also observed. FgrAAO and FoxAAO also exhibited activity on the furan substrates hydroxymethylfurfural (HMF), 5-(hydroxymethyl)furan-2-carboxylic acid (HMFCA) and 2,5-furandicarboxaldehyde (DFF) at levels similar to that of galactosyl substrates. This activity profile is notably distinct from that of the recently characterized CgrAAO,48 as well as CgrAlcOx.46 In particular, CgrAAO does not show a significant preference for the substitution pattern on the aromatic ring of benzyl alcohols.48 The regioselectivity of FgrAAO and FoxAAO is similar to the engineered FgrGalOx variant M3-5, which has 1.4-fold higher activity on 3-methoxybenzyl alcohol than on 4-methoxybenzyl alcohol.105 2.2.3 Michaelis-Menten Kinetics and Product Analysis The results of our initial activity screen, in particular the predominant activity on 3-methoxybenzyl alcohol and veratryl alcohol, suggests that both enzymes may be best named as aryl alcohol oxidases (EC 1.1.3.7). However, this is not unequivocal as select carbohydrates and furans exhibited significant specific activity values at the substrate concentrations tested. To further define the substrate specificity of FgrAAO and FoxAAO, Michaelis-Menten kinetic analysis was performed to determine kcat and KM values for the lead substrates (Figure A6 and Table 2.1). In these cases, assays of FgrAAO and FoxAAO were performed at 35 °C to balance optimal enzyme activity with short-term stability (vide supra). 43 Table 2.1. Substrate specificity of FgrAAO and FoxAAO * Substrate FgrAAO FoxAAO KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) Carbohydrates Galactose 1700 ± 150 21 ± 1.0 12 1600 ± 150 23 ± 1.2 14 Mannose 450 ± 74 36 ± 1.2 79 620 ± 57 19 ± 0.35 31 Fructose 460 ± 91 9.7 ± 0.55 21 510 ± 82 16 ± 0.72 31 Polyols Glycerol 2200 ± 360 17 ± 1.2 7.7 2700 ± 403 24 ± 1.6 8.8 Aryl alcohols Benzyl alcohol 86 ± 12 37 ± 2.8 430 30 ± 19 31 ± 4.2 1000 (350 ± 20)† Cinnamyl alcohol n.d.§ n.d.§ 3000 n.d.§ n.d.§ 3200 4-methoxybenzyl (p-anisyl) alcohol 59 ± 7.2 32 ± 1.8 540 51 ± 12 34 ± 2.9 290 3-methoxybenzyl (m-anisyl) alcohol 6.5 ± 1.5 52 ± 1.5 8000 6.7 ± 0.59 34 ± 1.7 5100 3,4-dimethoxybenzyl (veratryl) alcohol 3.7 ± 0.75 41 ± 4.0 11000 3.0 ± 0.24 25 ± 1.2 8300 Furans HMF 14 ± 2.6 29 ± 1.7 2100 17 ± 4.9 26 ± 2.6 1500 DFF 10 ± 2.9 3.6 ± 0.27 360 3.7 ± 1.0 1.4 ± 0.13 380 HMFCA 4.4 ± 0.90 3.8 ± 0.33 860 3.1 ± 0.31 5.2 ± 0.14 1700 * Errors represent the standard deviation from the mean values. † KM, kcat, and kcat/KM values calculated from fitting of the standard Michaelis-Menten equation to the data. Value in parentheses is the kcat/KM value obtained from a linear fit to the data in the substrate range 1 – 50 mM (v0 = (kcat/KM)[S]). Fitting of the data with a modified Michaelis-Menten equation accounting for substrate inhibition was unsuccessful due to very high errors on individual parameters (see Figure A6). § Individual KM and kcat values not determinable due to substrate solubility limitations; kcat/KM values obtained from slope of linear v0 versus [S] plots (Figure A6). 44 This analysis revealed that the monosaccharides galactose, mannose, and fructose, and the triol glycerol, are in fact poor substrates for both FgrAAO and FoxAAO. Although kcat values for these substrates are comparable to those of benzyl alcohols and furans, KM values are in the 0.5 – 2.0 M range. This indicates poor active-site affinity and selectivity for these polyhydroxylated compounds, with kcat/KM values only in the range 10-80 M-1s-1. Specifically, galactose was indeed a poor substrate for both FgrAAO and FoxAAO, displaying ~1000-fold lower kcat/Km values than the archetypal FgrGalOx,101 due to ca. 25-fold higher KM values and ca. 50-fold lower kcat values. In contrast, kcat/KM values for competent benzyl alcohol substrates are in the range 103-104 M-1s-1, while that of HMF is ca. 2000 M-1s-1 for both enzymes. In these cases, this is largely due to KM values in the millimolar range (Table 2.1). 2.2.3.1 Aryl Alcohol Oxidation Benzyl alcohols. The much higher specificity constants of FgrAAO and FoxAAO on benzyl and furan alcohols validates the designation of these enzymes as aryl alcohol oxidases (EC 1.1.3.7). Concordant with their grouping in the same phylogenetic clade (Figure 2.1), both enzymes displayed similar catalytic parameters on those substrates for which detailed kinetics were performed, with the highest activities toward veratryl alcohol, 3-methoxybenzyl (m-anisyl) alcohol, and cinnamyl alcohol (Table 2.1). kcat/KM values for these substrates were also similar to those observed for CgrAAO 48 and FgrGalOx 211, for which these are secondary substrates (Table A4). Distinctly, the broad-spectrum aliphatic/aryl alcohol oxidase CgrAlcOx46 is ca. 2 orders of magnitude more efficient than FgrAAO and FoxAAO on benzyl and cinnamyl alcohols, but does not oxidize other substrates of these enzymes, i.e., 4-methoxybenzyl alcohol (p-anisyl alcohol) and 4-hydroxybenzyl alcohol (Figure 2.3 and Table 2.1). Together, specific activity and Michaelis-Menten kinetic data for FgrAAO and FoxAAO indicate that these 45 enzymes favor 3-methoxy substitutions on the benzyl ring, and strongly disfavor substrates with 4-hydroxy substitutions (Figure 2.3 and Table 2.1). In the absence of an enzyme-substrate complex structure, it is presently unclear how a balance of electronic (e.g., the Hammett substituent constants for 3-methoxybenzyl, 4-methoxybenzyl, and veratryl alcohols are +0.12, -0.27, and -0.15, respectively212) and steric effects (e.g., specific recognition of the 3-methoxy group) dictate the observed specificities. Interestingly, the R330K mutant of FgrGalOx showed a 63-fold decrease in activity on 3-methoxybenzyl alcohol compared to WT FgrGalOx.101 This result juxtaposes our current observations that FgrAAO and FoxAAO natively possess an R330K mutation, yet exhibit a 2-fold increase in catalytic efficiency on 3-methoxybenzyl alcohol compared to WT FgrGalOx. These differences might be explained by the additional Q326E substitution that may make the FgrAAO and FoxAAO active sites more accessible to the methoxy-substituted substrate. Furans. The observation of demonstrable 5-(hydroxymethyl)furfural oxidation by FgrAAO and FoxAAO is also consistent with the assignment of these enzymes as aryl alcohol oxidases (see EC 1.1.3.47 vs. EC 1.1.3.7). The conversion of HMF from biomass sources into the bis-functional polymer precursors DFF and FDCA is of significant contemporary interest.40, 213 Select oxidases are capable of sequential oxidation of HMF to the various intermediates shown in Figure 2.4.27, 38, 143, 214-216 Hence, we analyzed the reaction kinetics and products of FgrAAO and FoxAAO in each of these steps. 46 Figure 2.4. HMF oxidation [O] represents a generic oxidant. The symbols next to each compound are used to denote the corresponding peaks in the NMR spectra comprising Figures A7-A9. FgrAAO and FoxAAO display similar specificity constants on HMF to the eponymous bacterial HMF oxidase (a GMC superfamily oxidoreductase),38 and exhibit higher specificity constants than other enzymes with HMF oxidase activity, including some glyoxal oxidases from AA5_1 (Table A5).214-216 However, the recently reported CgrAAO48 has 10-fold higher activity on HMF than both FgrAAO and FoxAAO (Table A5). Product analysis by 1H-NMR spectroscopy following extended incubation with HMF revealed a product mixture of DFF and FFCA (Figure A7), consistent with the initial formation of DFF and subsequent oxidation to FFCA, presumably via the aldehyde hydrate.44, 216 Indeed, the conversion of DFF to FFCA was directly demonstrated by initial-rate kinetics and product analysis, although this oxidation step was considerably slower (Table 2.2 and Figure A8). Furthermore, 1H NMR analysis following the incubation of FgrAAO or FoxAAO with HMF at 1.5 h and 3 h did not indicate the formation of HMFCA, which argues against the lower pathway 47 shown in Figure 2.4. Neither FgrAAO nor FoxAAO was able to oxidize FFCA to FDCA under the conditions tested, perhaps due to the low degree of hydration of FFCA compared with DFF 216. Furfural was also not oxidized by either enzyme in overnight reactions. Interestingly, HMFCA was also a good substrate for FgrAAO and FoxAAO, albeit with lower kcat and KM values than HMF, but with significant conversion to FFCA in extended incubations (Table 2.2 and Figure A9). In total, the specificity profiles of FgrAAO and FoxAAO are significantly different than those of other oxidases that are active on HMF and its derivatives,38, 143, 214-216 as summarized in Table A5. Table 2.2. Percent conversion of HMF, DFF and HMFCA by FgrAAO and FoxAAO Substrate FgrAAO FoxAAO Products (% conversion) Products (% conversion) HMF DFF (69%), FFCA (31%) DFF (76%), FFCA (24%) DFF FFCA (49%) FFCA (51%) HMFCA FFCA (84%) FFCA (96%) FFCA 0% 0% It is important to note that FgrAAO and FoxAAO are both able to convert HMF to FFCA, which is a step further than CgrAAO.48 The bacterial HMF oxidase is able to convert HMF fully to FFCA as well, but its use in industry may be hindered by its cofactor dependent mechanism.28, 38, 217, 218 Full enzymatic conversion of HMF to FDCA is ideal, but FgrAAO and FoxAAO show the possibility of an extension of current biocatalytic processes to FFCA with a singular enzyme, instead of a complex co-factor dependent enzymatic cascade. Furthermore, FFCA itself has been used as precursor for biofuels, surfactants and resins.219 48 2.2.3.2 Saccharide Oxidation As indicated above, monosaccharides, and galactose in particular, are comparatively poor substrates for FgrAAO and FoxAAO. Nonetheless, the observation of activity on galactose, mannose, and fructose deserves further comment. As listed in Table 2.1, the hexopyranoses galactose and mannose exhibit similar kcat values, with the apparently greater selectivity for mannose originating from a < 4-fold lower KM value. The activity on fructose is likewise weak, but of particular interest as this 2-ketose presents two potentially oxidizable primary hydroxyl groups in the predominant furanose form. Galactose. Although the molecular basis for the specificity difference between FgrAAO and FoxAAO and the archetypal FgrGalOx is unclear in the absence of any three-dimensional substrate or product complexes of AA5 members,46, 48, 56, 65, 66, 94, 99, 101, 102 sequence analysis and molecular modelling provides some insight. In particular, there is literature precedent for docking ligand structures into models of AA5_2 variants for activity analysis.95, 127 Foremost, substitution of R330 with lysine in FgrGalOx has been shown to dramatically decrease catalytic efficiency towards galactose through a 9-fold increase in KM and a 5-fold decrease in kcat (Table A3).101 This arginine, which is likewise replaced by lysine in FgrAAO and FoxAAO (Figure 2.2), has also been speculated to be involved in hydrogen bonding to the -OH groups at C3 and C4 of galactose by FgrGalOx.100 Molecular modelling (Figure 2.5) supported this proposal and indicated that R330 is positioned in FgrGalOx (PDB ID 1GOF)56 to make two H-bonds with C3-OH and C4-OH (1.8 and 2.4 Å, respectively). In contrast, when galactose is docked into a Phyre2 homology model of FgrAAO, (based on FgrGalOx, PDB ID 2VZ1, as the template; 63% i.d., confidence value 100.0), only one potential H-bond is predicted between K330 and the C3-OH of galactose (2.0 Å). Although 49 hypothetical, these structures implicate variation of this residue as a carbohydrate specificity determinant among AA5_2 members. Notably, CgrAlcOx and CgrAAO, which have similarly low activity on galactose as FgrAAO and FoxAAO, also have a residue other than lysine at this position (Figure 2.2). Furthermore, FgrAAO and FoxAAO have catalytic efficiencies on galactose that are more similar to the FgrGalOx mutant Des 3-2 (Q326E, Y329K and R330K)95 (Table A3), highlighting the importance of E326 and K330, which are both found natively in the homologs. The combination of mutations at distal positions (C383, Y436 and V494) might also contribute to the lower activity of FgrAAO and FoxAAO, as these residues have been implicated in changing the catalytic efficiency towards galactose (Table A3).96-99 Figure 2.5. Molecular docking of galactose (cyan) in FgrGalOx (PDB ID 1GOF)56 (A) and a Phyre2 homology model of FgrAAO (B) using AutoDock Vina as implemented in Chimera Copper (dark orange), W290 (yellow), Y329 (green), R330/K330 (magenta). Ligand coordination is indicated, specifying the distances (black). Mannose. The differences in catalytic constants between galactose and mannose are slight, and are thus difficult to rationalize. However, it is worth noting that this is the first report, to our knowledge, of an AA5_2 member that demonstrates a higher native catalytic efficiency on a non-galactosyl carbohydrate. The wild-type FgrGalOx has no reported activity on mannose 50 while the engineered variants M1 and M3 have relatively low specific activity compared to FgrAAO and FoxAAO on this monosaccharide.98, 102 Furthermore, the enzymes presented in this study have a 2-fold higher catalytic efficiently that the FgrGalOx mutant H1 (Table A3).102 These similarities could be caused by the analogous R330K substitution that is also present in the H1102 and M3 variants.98, 102 Our attempts to model the binding of mannose to FgrGalOx and FgrAAO were inconclusive, precluding further discussion. Fructose. The only other enzyme from AA5_2 with the ability to oxidize fructose is the engineered variant FgrGalOx R330K,101 versus which FgrAAO and FoxAAO have comparably low activities (Table A3). Nonetheless, this raises the question of which of the primary hydroxyl groups of fructose might be oxidized by FgrAAO and FoxAAO. Hence, we synthesized methyl β-D-fructopyranoside (1), methyl β-D-fructofuranoside (2) and methyl α-D-fructofuranoside (3) (Figures A10-11) by Fisher glycosylation220 to eliminate complications due to mutarotation of the free sugar (Figure 2.6). 51 Figure 2.6. Summary of observed hydrated aldehyde (gem-diols) products from the oxidation of methyl fructosides, anhydro sugars and raffinose by FgrAAO 52 1D (1H, 13C{1H}) and 2D NMR (HSQC and HMBC) spectroscopy revealed that the two enzymes produced identical oxidation products (Figure 2.7, Tables A6-A7 and Figures A12-17). The reaction of methyl β-D-fructopyranoside (1) with FgrAAO and FoxAAO resulted in 19% and 17% conversion, respectively, as estimated by integration of 1H NMR spectra. The only product observed in both cases was the hydrated aldehyde (7), as confirmed by detailed 1D and 2D NMR analysis (Figures 2.7, A12 and A15A-17A). Specifically, no aldehyde signal was observed in the range 9-10 ppm in 1H NMR spectra, while a peak at 5.17 ppm corresponding to the C-H of a gem-diol221 correlated to the quaternary carbon at C2, thus confirming the site of oxidation as the primary hydroxyl group of C1 (Figures A15A and A17A). Turning our attention to the furanosides, methyl β-D-fructofuranoside (2) was converted to a single product (8) (Figures 2.7, A13 and A15B-17B) by FgrAAO or FoxAAO with 16% and 11% conversion, respectively. In these cases, a distinct doublet was observed at 4.85 ppm, which was attributed to the C-H of a hydrated aldehyde. This doublet correlated to C5 in the HMBC spectrum, signifying that the primary hydroxyl of C6 was the site of oxidation of methyl β-D-fructofuranoside (Figures A15B-17B). 53 Figure 2.7. Complete analysis of oxidation reaction components of FgrAAO with (A) methyl β-D-fructopyranose (1) and (B) methyl β-D-fructofuranose (2) The 1H-13C HSQC used for the identification of the FgrAAO oxidation products. Each black crosspeak corresponds to a 1JC,H-coupling of the substrate and the red crosspeak corresponds to a 1JC,H-coupling of the product interaction from the HSQC experiment. Two carbon peaks at 63.1 ppm and 72.7 ppm have been attributed to an impurity that is present in either the HRP, catalase or buffer solutions. The enzymes were removed by ultrafiltration, and the filtrate was frozen with liquid nitrogen, lyophilized and resuspended in D2O. 54 On the other hand, the incubation of methyl α-D-fructofuranoside (3) with FgrAAO or FoxAAO showed no indication of product formation (Figures A14 and A15C-A17C). Thus, to further explore potential steric and stereochemical factors effecting enzymatic oxidation, 2,5-anhydro-D-glucitol (4) and 2,5-anhydro-D-mannitol (5) were tested as analogs of methyl α-D-fructofuranoside and methyl β-D-fructofuranoside, respectively. Notably, no oxidation was observed with 2,5-anhydro-glucitol, however 6% oxidation was observed for 2,5-anhydro-mannitol, which is concordant with our observations with the methyl fructofuranosides. The low degree of conversion of 2,5-anhydro-mannitol made full spectral assignment challenging, however, we were able to observe correlation peaks in the HSQC and HMBC (Figure A18), which were consistent with oxidation of a single primary hydroxyl group to an aldehyde that was subsequently hydrated to the gem-diol (9). In this context, it is important to note both primary hydroxyl groups of 2,5-anhydro-mannitol are equivalent, due to C2 symmetry. To assist in rationalizing these results, we modelled the binding of the three methyl fructosides (1-3) to a structural homology model of FgrAAO. A structural model was obtained for methyl β-D-fructopyranoside (1) with C1-OH oriented towards the copper and the methoxy group positioned toward an active site pocket containing P463, Y405 and K330 (Figure 2.8). However, in this model, Tyr272 was 3.7 Å away from the protons on the primary alcohol, suggesting a catalytically inactive pose. Using the same grid search area, a productive complex with methyl β-D-fructofuranoside (2) was obtained, in which the C6-OH was pointing towards the copper center and the anomeric methoxy group was pointing away from the substrate recognition residues and towards F464 and F194 (Figure 2.8). In this model, the catalytic Tyr272 was located at a reasonable distance away from the abstractable hydrogen (2.0 Å). Most notably, the alternate structure, with the C1-OH oriented towards the copper, was not observed 55 by modeling. Although hypothetical, these results are consistent with our experimental product analysis (Figure 2.6). When docking was attempted with methyl α-D-fructofuranoside (3), only one model was produced with the methoxy pointed towards the copper center, which represents a non-productive binding mode and is consistent with experimental results (Figure 2.8). Notably, this pose constitutes a flip of the furanose ring relative to that observed for methyl β-D-fructofuranoside, which places C-6 distal from the copper center. Together, these models suggest that binding modes that direct the anomeric methyl group into the protein are disfavored, thus rationalizing the observed specificity. 56 Figure 2.8. Molecular docking (AutoDock Vina as implemented in Chimera) of (A) methyl β-D-fructopyranoside (3), (B) methyl β-D-fructofuranoside (2) and (C) methyl α-D-fructofuranoside (1) in FgrAAO Copper (dark orange), methoxy group on sugar (dark blue), W290 (yellow), Y329 (green), K330 (magenta), E326 (blue), Y405 (orange), Q406 (cyan), P463 (pink), F194 (red), Q195 (black). C1-OH and C6-OH of each sugar are labelled. In light of the observed 6-O-oxidase activity on methyl β-D-fructofuranoside, as well as the oxidase activity observed on the classical GalOx substrate raffinose [β-D-fructofuranosyl-O-57 α-D-glucopyranosyl-(1→6)-α-D-galactopyranoside] (6) (Figure 2.3), the potential for multiple oxidations of the trisaccharide was investigated by NMR spectroscopy. Thus 1H 1D-TOCSY, via selective irradiation of peaks at 6.19 ppm and 5.12 ppm that appeared following incubation with FgrAAO, indicated oxidation and hydration of the C-6 hydroxyl groups of both galactose and fructose C6, respectively (full assignments of the proton and carbon spectra, for raffinose and the oxidation product using 1D (1H, 13C{1H}, TOCSY) and 2D (HSQC, HMBC and TOCSY) NMR, Figures A19-A22, are provided in Table A8). Specifically, no peaks corresponding to oxidation of C1 on the fructosyl residue were observed. Based on the NMR analysis, 87% of unoxidized raffinose remained after incubation with FgrAAO. Multiple products, including two mono-oxidized species and di-oxidized species (10), could be present, but due to low conversion no quantitative data could be obtained. The observation of the oxidation of both terminal primary hydroxyl groups in raffinose (Figure 2.6) by FgrAAO is consistent with the similar kcat/KM values for the component monosaccharides, galactose and fructose. It would also appear that oxidation of C1 of the fructofuranosyl residue of raffinose is precluded sterically. The bis-oxidation of raffinose by FgrAAO suggests a potential route to valorize raffinose as a chemical building-block, as has previously been explored with lactose and FgrGalOx.131 2.2.3.3 Glycerol Oxidation Glycerol is a prochiral compound that is produced in significant quantities annually, including as a byproduct of the biodiesel industry.222 Hence, there is significant interest in the valorization of glycerol by desymmetrization. It has been previously reported that the wild-type FgrGalOx oxidizes glycerol stereoselectively to L-glyceraldehyde.223 Moreover, FgrGalOx variants have been integral to the desymmetrization of a key 2-alkynyl-glycerol derivative in the 58 synthesis of a key antiviral drug.144 On the other hand, we have also recently shown that CgrAlcOx favours oxidation of glycerol to D-glyceraldehyde.224 Here, comparative analysis indicates that FgrAAO and FoxAAO both oxidize glycerol predominantly to produce L-glyceraldehyde, with enantiomeric ratios (er) of 93:7 and 91:9 (L-:D-), respectively (Figures A23-A24). Under similar conditions, FgrGalOx likewise favored the L-isomer, as expected, with an er of 96:4 (L-:D-), whereas the er for CgrAlcOx was 10:90 (L-:D-). The similar stereoselectivities of the Fusarium AA5 CROs may be understood in terms of their conserved active-site motifs (Figure 2.2). Nonetheless, the ability of these enzymes to selectively produce the unnatural and more valuable L-isomer may find biotechnological application. In this context, we note that FgrGalOx is ca. 2-fold more active than FgrAAO and FoxAAO. 2.3 Conclusion In summary, a detailed biochemical characterization of two orthologous CROs from Fusarium graminearum and Fusarium oxysporum, which segregate into a previously uncharacterized clade with AA5_2, revealed a distinct substrate specificity profile versus known GalOx,45, 53, 56, 89, 106, 204 AlcOx,46, 47 and AAO48 within the family. Despite strong conservation of active site architecture compared to the archetypal FgrGalOx, both FgrAAO and FoxAAO have a novel native preference for mannose and fructose over galactose or galactosides, and a clear preference for aryl alcohols, specifically 3-methoxybenzyl alcohol and veratryl alcohol. Unlike CgrAlcOx,46 FgrAAO and FoxAAO are poorly active on unactivated alkanols, with the exception of the aforementioned saccharides and glycerol. While the nomenclature of oxidases is fraught with difficulty due to overlapping definitions of EC 1.1.3.7 (aryl-alcohol oxidase), EC 1.1.3.13 (alcohol oxidase), and EC 1.1.3.47 (5-(hydroxymethyl)furfural oxidase), detailed kinetic 59 data (Table 2.1 and Figure 2.3) support denoting these F. graminearum and F. oxysporum CROs as aryl-alcohol oxidases. High KM values and low specificity constants (kcat/KM values) do not suggest that galactose 6-oxidation (EC 1.1.3.9) is the natural activity of these enzymes. Finally, we note that the physiological substrate of these enzymes is currently unknown and may not be represented in our panel of compounds. In this context, it is notable that FgrAAO and FoxAAO group together with orthologs from F. verticillioides and F. subglutinans in a clade previously designated as “gaoB” (galactose oxidase B)56 (Figure 2.1). However, the F. verticillioides and F. subglutinans gene products have not been biochemically characterized.205 Our kinetic data on FgrAAO and FoxAAO, like data on the Colletotrichum homologs,46-48 indicate that caution is warranted in extending functional predictions in AA5_2 on the basis of bioinformatics alone. Indeed, our analysis would suggest that the F. verticillioides and F. subglutinans orthologs are likely to also be predominant aryl alcohol oxidases. Within this clade, an arginine to lysine substitution in the active site may be a key specificity determinant. Finally, the ability of FgrAAO and FoxAAO to oxidize HMF and derivatives, as well as glycerol, suggests possible applications of these CROs in biocatalysis. Like FAD-dependent oxidases, CROs use molecular oxygen as a co-substrate, thus avoiding the requirement for co-factor regeneration of NAD-dependent oxidases. At the same time, the facile production of AA5 CROs in P. pastoris and E. coli enables further tuning of specificity for biocatalytic applications, which will be further informed by enzyme structure-function relationships, including those described here. Indeed, the replacement of chemical oxidants with potentially greener biocatalysts is a topic of considerable contemporary interest. 60 2.4 Materials and Methods 2.4.1 Chemicals and Enzymes Wild-type galactose oxidase from Fusarium graminearum and alcohol oxidase from Colletotrichum graminicola were produced in Pichia pastoris and purified as previously described.46, 209 Distilled water was used for the preparation of all buffers and stock solutions unless stated otherwise. Catalase from bovine liver (2000–5000 units per mg protein, Sigma) and horseradish per-oxidase (Rz > 3, 300 units per mg, Bio Basic Canada Inc.), obtained as lyophilized powders, were used as received. Other substrates and reagents were purchased from commercial sources (Sigma-Aldrich, VWR or Fisher) and used without further purification. 2.4.2 Sequence and Bioinformatic Analysis Sequences of FgrAAO (GenBank XP_011322138) and FoxAAO (GenBank XP_018246910) were taken from the National Center for Biotechnology Information (NCBI). Percent identity analysis was performed using the full-length sequences of FgrAAO, FoxAAO and other AA5_2 members from the Fusarium and Colletotrichum genus using MatGat.225 An alignment using the online Genome Net CLUSTALW tool was performed with the two new enzymes presented in this study and compared to the characterized galactose oxidase from Fusarium graminearum (FgrGalOx) (GenBank: P0CS93.1), alcohol oxidase (CgrAlcOx) (Gen Bank: EFQ30446.1) and aryl oxidase (CgrAAO) (Gen Bank: EFQ 27661.1) from Colletotrichum graminicola. In addition, Phyre2226 was used to make three-dimensional homology models for FgrAAO and FoxAAO. The structural models were aligned to the crystal structures of FgrGalOx (PDBID 1GOF)56 and CgrAlcOx (PDBIID 5C86)46 for spatial amino acid comparison to give further support to the alignment. 61 In addition, two AA5_1 sequences and 62 sequences of AA5_2 members from a variety of fungal species were extracted from the public CAZY database43 (February 2021) and aligned using MUSCLE.227 Where present, signal sequences and additional domains, such as carbohydrate-binding modules, were removed. Any redundant sequences were excluded and the resulting catalytic domain sequences were realigned. A maximum-likelihood phylogenetic tree was estimated by RAxML version 8.2.12228 using 47 sequences as inputs on the CIPRES gateway229 with automatic bootstrapping terminating at 650 bootstrap replicates. The resulting phylogeny was visualized with FigTree (http://tree.bio.ed.ac.uk/software/figtree/). 2.4.3 DNA Cloning cDNA encoding FgrAAO (Genbank ID XP_011322138) and FoxAAO (Genbank ID XP_018246910) without the predicted native signal peptide and including a C-terminal His6 tag-encoding sequence were commercially synthesized in a codon-optimized form. The sequences were cloned into the P. pastoris expression vector pPICZα-A using the EcoRI and XbaI restriction sites flush with the sequence encoding the S. cerevisiae α-factor signal peptide by Genescipt. The resultant constructs were transformed into chemically competent E. coli DH5α by heat shock. In both cases, transformants were grown overnight at 37 °C on Luria-Bertani-Low-Salt (LBLS) agar plate and selected against 25 μg mL−1 Zeocin (Invitrogen). Surviving colonies were picked and grown overnight at 37 °C in 5 mL LBLS medium with 25 μg mL−1 Zeocin. Plasmids from the overnight cultures were extracted using a commercial miniprep-kit (Geneaid, New Taipei City, Taiwan). The presence of the pPICZα-A-FgrAAO and pPICZα-A-FoxAAO constructs in positive clones was checked by agarose (1%; w/v) gel electrophoresis and the correct insertion of both genes into the corresponding vectors was verified by DNA sequencing. 62 Transformation of constructs into P. pastoris X33 was performed by digesting 6.5-14 μg of plasmid DNA for 4 h at 37 °C with PmeI (New England Biolabs). The linearized plasmid was purified using an ion exchange column (Omega). The resulting digested plasmids were transformed into electrocompetent P. pastoris X33 cells prepared on the same day.230 Transformants were grown on yeast extract peptone dextrose (YPDS) agar plates for 2 days at 30 °C and selected against 100 or 500 μg mL−1 Zeocin. Four colonies selected from 500 μg mL−1 Zeocin plates were inoculated into 15 mL sterile conical tubes containing 2 mL of YPD and grown for 6 hours at 30 °C shaking at 200 rpm. At OD ~0.15, 200 μL of the growing culture was transferred to 10 mL of buffered complex glycerol medium (BMGY) and grown in a shaking incubator overnight at 30 °C and 200 rpm. Cells were then pelleted by centrifugation at 3000 g for 10 min at 25 °C, the BMGY medium was discarded and replaced with 4 mL of buffered complex methanol medium (BMMY) containing 0.5% (v/v) methanol. The cultures were shaken at 200 rpm over 3 days at either 25 or 16 °C with regular feeding of 0.5% methanol every 24 h to ensure continued protein expression. Secreted proteins were separated from the cells by centrifugation at 4000 rpm for 10 min and protein production was monitored using SDS-PAGE. The clone yielding the highest amount of protein was retained for large-scale production. 2.4.4 Protein Production and Purification Large Scale Protein Production Single colonies of Pichia pastoris X33 expressing clones were individually streaked onto agar plates containing 500 μg mL-1 of the antibiotic Zeocin and grown for 2 days in the dark in a 30 oC incubator. Precultures containing 4 mL of YPD and 500 μg mL−1 Zeocin were inoculated using a single colony and shaken at 30 oC at 250 rpm for 10 h. Biomass production was initiated by the addition of the preculture into 1 L of BMGY media shaken in 4 L beveled flasks at 30 oC 63 at 250 rpm overnight. Once the BMGY cultures reached an OD600 of 6-12, the cells were harvested by centrifugation at 3000 g for 15 mins at room temperature. The cells were then resuspended using 400 mL of BMMY media supplemented with 0.5 mM copper sulfate containing 3% methanol and transferred to 1L flasks with a foam cap. The flasks were shaken at 250 rpm at 16 °C for three days. The cultures were fed 1% (v/v) methanol every 24 h to maintain continued expression of the recombinant protein. On the last day of methanol induction, the desired proteins were separated from the cells by centrifugation at 3000 g for 30 mins at 4 °C. The supernatant was quickly decanted, filtered through 0.45 μm membrane and stored at 4 oC until purification. Protein Purification The pH of the liquid medium which contained FgrAAO or FoxAAO was raised before purification to 7.5-8.0 by the dropwise addition of 1 M NaOH to the stirred solution. The medium was then filtered through a 0.45 μm membrane and allowed to equilibrate for at least 12 h at 4 oC. The supernatant was allowed to pass through a 5 mL pre-packed Ni-NTA column, pre-equilibrated with 50 mM sodium phosphate buffer at pH 7.5 with 300 mM NaCl with 10 mM imidazole at 5 mL/min. The column was washed with 5 column volume of equilibration buffer containing 50 mM sodium phosphate buffer at pH 7.5 with 300 mM NaCl and 10 mM imidazole at 4.5 mL/min. Proteins were eluted with a linear gradient of 2% to 100% of 500 mM imidazole in a 50 mM sodium phosphate buffer with 300 mM NaCl at 5 mL/min. The total elution volume was 125 mL collected in 1 mL fractions. Both FgrAAO and FoxAAO eluted at ~140 mM imidazole. The proteins were then concentrated using a 30,000 MWCO Vivaspin centricone and 0.5 – 1 mL was loaded onto a Superdex 75 size-exclusion column pre-equilibrated with 50 mM sodium phosphate buffer at pH 7.5 at 1 mL/min. A total volume of 200 mL of equilibration 64 buffer was used through the column at 1 mL/min. FoxAAO eluted at 102 mL as one singular peak, while FgrAAO eluted as two peaks at 72 and 104 mL. SDS-PAGE was performed using pre-cast 4-20% (w/v) polyacrylamide gel in the presence of 2% (w/v) SDS under reducing conditions. Proteins were visualized using Coomassie blue R-250. Protein concentrations were determined by measuring A280. The extinction coefficients were calculated using the ProtParam tool on the ExPASy server. 2.4.5 Analytical Protein Deglycosylation The presence of protein glycosylation on FgrAAO and FoxAAO was assessed by treatment with N-glycosidase F from Flavobacterium meningosepticum (PNGaseF, New England Biolabs). Deglycosilation experiments were performed under denaturing condition by adding 3 μg of protein to 10X Glycoprotein Denaturing Buffer and heated for 10 min at 100 °C. The samples were subsequently diluted to 20 μL with GlycoBuffer 2 and tertigol-type NP-400 detergent. Finally, 1 μL of PNGaseF was added to the sample and incubated for 1 hour at 37 °C. Changes in protein mobility was assessed by SDS-PAGE stained with Coomassie blue R-250. 2.4.6 pH and Temperature Profile The enzyme activity across a wide range of pH values was determined using phosphate citrate (pH 4.0-7.0), sodium phosphate (pH 5.5-8.5) and glycine-NaOH (pH 8.5-11.0) buffers. Enzyme activity was measured using the HRP – 2,2’-azinobis(3-ethylbenzthiazoline-6-sulfonic acid) (ABTS)-coupled assay with 300 mM of galactose at 25 oC. The optimum temperature was determined using the same coupled assay using 50 mM sodium phosphate buffer at pH 7.5 with 1 μg mL-1 as the final enzyme concentration and 300 mM galactose as the substrate at temperatures ranging from 15 °C to 80 °C. 65 2.4.7 Temperature Stability Temperature stability of both FgrAAO and FoxAAO was determined by diluting the stock protein in 50 mM sodium phosphate buffer at pH 7.5 to obtain a final protein concentration of 0.2 mg mL-1 for both proteins studied. The diluted protein was then pre-incubated in a thermocycler at 30 °C, 39 °C, 49 °C and 60 °C. Samples were taken out at different time interval and the activity of the proteins was measured using the HRP-ABTS coupled assay with 300 mM galactose as the substrate at 30 °C. 1 μg mL-1 was the final enzyme concentration in the reactions. 2.4.8 Enzyme Kinetics The colorimetric HRP-ABTS coupled assay was used to determine the kinetics of the enzymatic oxidation of substrates. The oxidation of the alcohol group on the substrates by AA5 enzymes consumes 1 equivalent of O2 and produces 1 equivalent of H2O2. The oxidation of ABTS (λmax = 420 nm, ε = 36,000 M-1cm-1) is catalyzed by the enzymes HRP using 2 equivalents of H2O2. This is a standard assay used to monitor initial rate kinetics for AA5 enzymes.209 This assay was optimized for sensitivity and linearity. The activity of FgrAAO and FoxAAO was surveyed on a variety of substrates using this coupled assay comprised of 50 mM sodium phosphate buffer at pH 7.5, 0.46 mM ABTS and 30 U/mL of HRP at 25 oC. The initial substrate screen included carbohydrates, polyols, diols and primary alcohol substrates at 300 mM, polysaccharides at 2.5 mg mL-1, benzyl alcohol and galactitol at 30 mM, methyl glyoxal, aryl alcohols and furans at 5 mM and 10 mM for secondary alcohols. One unit of AA5 enzyme activity was defined as the amount of enzyme required to oxidize 2 μmol of ABTS per minute, which is equivalent to the consumption of 1 μmol of oxygen per minute. 66 To determine Michaelis-Menten parameters of FgrAAO and FoxAAO, different concentrations of substrates were used over the range of 10-2500 mM for galactose, 10-4000 mM for fructose, 5-3000 mM for mannose, 25-7000 mM for glycerol, 10-250 mM for benzyl alcohol, 0.5-13 mM for cinnamyl alcohol, 1-200 mM for p-anisyl alcohol for FgrAAO and 1-100 mM for FoxAAO, 0.25-100 mM for m-anisyl alcohol for FgrAAO and 0.25-50 mM for FoxAAO, 0.25-10 mM for veratryl alcohol, 1-80 mM for HMF, 0.25-20 mM for HMFCA and 0.1-40 mM for DFF for FgrAAO and 0.5-20 mM for FoxAAO. The reactions were performed at 35 oC in 50 mM sodium phosphate buffer at pH 7.5 with 0.46 mM ABTS, 30 U/mL of HRP and using 2.1 to 28 pmol of purified FgrAAO and FoxAAO. Data were fit with the Michaelis−Menten equation using OriginPro software (OriginLab 9.55). 2.4.9 Computational Docking Studies Molecular docking simulations were performed using the CHIMERA software from UCSF Resource for Biocomputing, Visualization, and Informatics.231 The FgrGalOx crystal structure (PDB 1GOF) and a Phyre model of FgrAAO were used to generate the receptors for simulations. Ligands were either extracted from other PDB files or build from components extracted from other PDB files. Ligands and receptors were first prepared for docking in chimera by adding hydrogens and assigning proper protonation states. The docking simulation itself was performed using Autodock VINA, run within CHIMERA, with the AMBER03 force field.232 Appropriate simulation cells were defined for the respective docking simulations. For docking of galactose and mannose, a 7 Å cell with the copper atom bordering the z-coordinate edge was chosen. Galactose gave a reasonable binding pose with both FgrGalOx and FgrAAO, while mannose lowest energy structures produced the sugar in unproductive binding positions. For the docking of methyl α/β-fructofuranosides and methyl β-fructopyranoside, a 6x7x7 Å cell was selected 67 with the copper atom bordering the z-coordinate edge. Reasonable functional binding poses were calculated for methyl β-fructofuranosides and methyl β-fructopyranoside, and an unproductive for catalytic turnover binding structure was calculated for methyl α-fructofuranosides. The created receptor-ligand complex structures were further processed using the PyMOL software from Schrodinger LLC involving the identification of ligand-receptor interaction and for the determination of distances. 2.4.10 Synthesis of Methyl α/β-D-fructofuranosides and Methyl β-D-fructopyranoside The 1-O-methyl D-fructose derivatives were prepared according to a modified procedure.220 To a solution of D-fructose (1.00 g, 0.111 mol) in MeOH (50 mL) 50 μl of acetyl chloride (AcCl) was added at RT and stirred for 25h. The reaction was neutralized with Dowex 66 Free Base resin, filtered, washed with MeOH and concentrated. Half the crude mixture was purified by anion exchange chromatography (Dowex 1 x 8 200-400, -OH form; distilled water as the mobile phase), which yielded three products: methyl α-D-fructofuranoside (0.193 g, 39%, Rf 0.58 [6:4:1, EtOAc-i-PrOH-H2O]), methyl β-D-fructofuranoside (0.064 g, 13%, Rf 0.46), methyl β-D-fructopyranoside (0.084 g, 17%, Rf 0.34) and a mixture of methyl α/β-D-fructofuranoside (0.004 g, 0.8%) and methyl β-D-fructofuranoside/ β-D-fructopyranoside (0.141 g, 29%) all as clear gels, giving a total purification yield of 99%. NMR experiments were run on a Bruker AV III HD 400 MHz spectrometer equipped with a BBFO smart probe. 1H, 1H,13C HSQC and 1H,13C HMBC NMR spectra were used to characterize the compounds. Compound methyl α-D-fructofuranoside: 1H NMR (400 MHz, D2O): δ= 3.22 (s, 3H, H7), 3.68 (dd, J = 32.6, 12.3 Hz, 4H overlapped with H6, H1), 3.72 (m, 4H overlapped with H1, H6), 3.85 (m, 1H, H4), 4.05 (t, J = 7.8 Hz, 1H, H5), 4.17 (d, J = 8.2 Hz, 1H, H3). 13C{1H} NMR (100.6 MHz, D2O): δ 49.5 (C7), 60.5 (C1), 63.3 (C6), 75.7 (C5), 77.6 (C3), 68 82.0 (C4), 104.4 (C2). Compound methyl β-D-fructofuranoside: 1HNMR (400 MHz, D2O): δ 3.32 (s, 3H, H7), 3.73 (dd, J = 49.7, 12.8 Hz, 4H overlapped with H6, H1), 3.75 (dd, J = 52.6, 12.9 Hz, 4H overlapped with H1, H6), 3.96 (m, 2H, H4 and H5), 4.10 (m, 1H, H3). 13C{1H} NMR (100.6 MHz, D2O): δ= 48.8 (C7), 58.4 (C1), 61.9 (C6), 77.9 (C5), 80.7 (C3), 83.8 (C4), 108.9 (C2). Compound methyl β-D-fructopyranoside: 1HNMR (400 MHz, D2O): δ= 3.29 (s, 3H, H7), 3.76 (dd, J = 31.4, 13.0 Hz, 4H overlapped with H1, H6), 3.77 (s, 4H overlapped with H6, H1), 3.85 (m, 1H, H4), 3.92 (d, J = 10.0 Hz, 1H, H3), 3.97 (m, 1H, H5). 13C{1H} NMR (100.6 MHz, D2O): δ 49.1 (C7), 61.6 (C1), 65.2 (C6), 68.6 (C3), 69.3 (C5), 70.0 (C4), 101.2 (C2). 2.4.11 Enzyme Product Analysis Oxidation of Methyl α/β-D-fructofuranosides and Methyl β-D-fructopyranoside Reactions containing 20 mg of substrate (methyl α/β-D-fructofuranosides and methyl β-D-fructopyranoside) and 1 mg mL-1 of both catalase and HRP were initiated by the addition of 300 μg of purified FgrAAO and FoxAAO in a final volume of 1 mL (50 mM sodium phosphate buffer, pH 7.5). Reactions were stirred at 400 rpm at room temperature for 16 hours, at which time the enzymes were removed by ultrafiltration (10 kDa cut-off Centricon-Millipore, Billerica, MA, USA). The products were collected, frozen in liquid nitrogen and lyophilized for 4 days. The resulting powders were resuspended in D2O and a preliminary NMR analysis was conducted. The samples were subsequently lyophilized for 24 hours and a second reaction was performed under the previously mentioned conditions with 500 μg of purified enzyme added. The reaction was stirred for 25 hours and the same work-up procedure was followed. NMR experiments were run on a Bruker AV III HD 400 MHz spectrometer equipped with a BBFO smart probe. 1H, 13C, HSQC and HMBC NMR spectra were calibrated to the 69 internal acetone peak (0.34M) (2.22 ppm and 30.89 ppm, respectively). The integration value for relevant peak areas were used to determine percent conversions. Oxidation of 2,5-anhydromannitol and 2,5-anhydroglucitol Reactions containing 20 mg of substrate (2,5-anhydro-D-mannitol and 2,5-anhydro-D-glucitol) and 1 mg mL-1 of both catalase and HRP (dissolved in D2O) were initiated by the addition of 700 μg of purified FgrAAO in a final volume of 1 mL (50 mM sodium phosphate buffer, pH 7.5 previously lyophilized and resuspended in D2O). Reactions were stirred at 400 rpm at room temperature for 24 hours, at which time the enzymes were removed by ultrafiltration (5 kDa cut-off Centricon-Millipore, Billerica, MA, USA). The products were collected and NMR analysis was conducted. NMR experiments were run on a Bruker AV III HD 400 MHz spectrometer. 1H, 13C, HSQC and HMBC NMR spectra were calibrated to the internal acetone peak (0.34M) (2.22 ppm and 30.89 ppm, respectively). The integration value for relevant peak areas were used to determine percent conversions. Oxidation of Raffinose Reactions containing 20 mg of raffinose and 1 mg mL-1 of both catalase and HRP were initiated by the addition of 800 μg of purified FgrAAO in a final volume of 1 mL (50 mM sodium phosphate buffer, pH 7.5). A control reaction without FgrAAO was performed using the same protocol. Reactions were stirred at 400 rpm at room temperature for 25 hours, at which time the enzymes were removed by ultrafiltration (5 kDa cut-off Centricon-Millipore, Billerica, MA, USA). The products were collected, frozen in liquid nitrogen and lyophilized for 3 days. The resulting powders were resuspended in D2O and NMR analysis was conducted. NMR experiments were run on a Bruker AV III HD 400 MHz spectrometer. 1H, 13C, HSQC, HMBC and TOSCY spectra were calibrated to the internal acetone peak (0.34M) (2.22 ppm and 30.89 70 ppm, respectively). The integration value for relevant peak areas were used to determine percent conversions. Oxidation of HMF, DFF, HMFCA, FDCA and Furfural Reactions containing 10 mM of substrate (HMF, DFF, HMFCA, FDCA and furfural) and 1 mg mL-1 of both catalase and HRP were initiated by the addition of 60 μg of purified FgrAAO and FoxAAO in a final volume of 1 mL (50 mM sodium phosphate buffer, pH 7.5). For each reaction a negative control was performed with identical conditions omitting the purified enzyme. Reactions were stirred at 400 rpm at room temperature for 16.5 hours, at which time the enzymes were removed by ultrafiltration (5 kDa cut-off Centricon-Millipore, Billerica, MA, USA). D2O was added to the filtrate to a final composition of 10% (v/v). NMR experiments were run on a Bruker AV III HD 400 MHz spectrometer equipped with a BBFO smart probe. 1H NMR spectra were collected with water suppression (4.7 ppm) using a standard pre-saturation pulse sequence. Chemical shifts were calibrated to the internal HOD peak (4.7 ppm). Standards of all substrates were used to identify distinct chemical shifts for each molecule. The integration value for relevant peak areas were used to determine percent conversions. Oxidation of Glycerol Reaction containing 0.54 M glycerol and 1 mg mL-1 of both catalase and HRP were initiated by the addition of 800 μg of purified FgrGalOx, CgrAlcOx, FgrAAO and FoxAAO in a final volume of 1 mL (50 mM sodium phosphate buffer, pH 7.0-8.0). Reactions were left at room temperature for 24 hours, at which time the enzymes were removed by ultrafiltration (5 kDa cut-off Centricon-Millipore, Billerica, MA, USA). 20 mg of 2,4-dinitrophenyl hydrazine was added to the Eppendorf and the reaction mixture was incubated in a heat block at 50 ℃ for 6 hours. A TLC plate was used to check formation of desired product (Rf = 0.45, 100% EtOAc). 71 Subsequently, the solution was purified via preparatory TLC and the desired hydrazone was mechanically isolated and dissolved in MeOH. The solution was then filtered and concentrated. The composition of the purified glyceraldehyde-hydrazones was analyzed on HPLC (3 μL injection, Chiracel® IA-3). Eluents used for HPLC methods, water with 0.1% formic acid (A) and methanol (B) were LC-MS grade (Optima, Fisher). For separation between L-and D-glyceraldehyde-hydrazones an isocratic method using 60% A, 40% B with a flow rate of 0.65 mL/min was used with a 12 minute stop time with UV detection at 360 nm. L-glyceraldehyde-hydrazone eluted at 2.30 min and D-glyceraldehyde-hydrazone eluted at 2.76 min. ESI mass spectra were also collected in positive mode scan for m/Z 95 – 500 running at 0.8 s/cycle. drying gas = 5.0 L/min, nebulizer pressure = 50 psi, gas temperature = 300 °C, capillary voltage = 4000 V. 72 Chapter 3: Biochemical survey of Copper Radical Oxidase specificity in Auxiliary Activity Family 5 3.1 Introduction Global climate change due to human activities is a pressing issue confronting our society (https://climate.nasa.gov/scientific-consensus/).233 One avenue to decrease reliance on petroleum is to increase the utilization of plant biomass as a renewable carbon feedstock.2, 3 Plant biomass is an abundant source of complex polysaccharides, polyaromatics (i.e., lignin), and low molecular weight extractives (e.g. lipids, terpenes, etc.), each of which can potentially be valorized via (bio)catalytic routes. Foremost, the development of so-called “biofuels”, e.g. ethanol fermented from sugar feedstocks, has received the widest attention, but faces certain economic and scalability challenges.7, 9, 234 In parallel, there is a growing appreciation that upgrading plant biomass components to produce high-value biochemicals and biomaterials may provide solutions to lower reliance on fossil petroleum beyond the energy sector.4, 10, 11, 235, 236 The utilization of plant biomass poses many challenges due to inherent chemical and structural complexity, which make fractionation and chemical transformation difficult. Yet, a wide diversity of microorganisms have developed specialized biocatalytic systems of hydrolytic and oxidative enzymes to unlock this natural resource.13-21, 43, 237 Informed by decades of fundamental biochemical research,238-241 industrial enzyme “cocktails” have been developed for complete saccharification of plant biomass, especially starch and cellulose fractions, for fermentation to biofuels.197, 198, 242 Spurred by a growing track record of successful applications and significant industrial infrastructure for scalable production, enzyme-based biocatalysis is enjoying a period of sustained and growing interest.24, 243, 244 In particular, there is strong demand to replace chemical 73 oxidants, many of which produce hazardous waste, with “greener” alternatives, making the study of oxidative enzymes attractive in this context.14, 24 One class of oxidases that has shown promise for the valorization of plant biomass is the copper radical oxidases (CROs) that comprise Auxiliary Activity Family 5 (AA5) in the Carbohydrate-Active Enzymes (CAZy) classification.43 These enzymes contain a mono-nuclear copper radical center coordinated by a distinct crosslinked tyrosine-cysteine active-site residue, which catalyzes two-electron oxidations with concomitant reduction of oxygen to hydrogen peroxide.45, 84 Within AA5, two subfamilies have been identified: Characterized subfamily 1 (AA5_1) enzymes have predominant activity on glyoxal and methylglyoxal (EC 1.2.3.15).44 Subfamily 2 (AA5_2) contains the archetypal CRO, galactose 6-oxidase (GalOx, EC1.1.3.9),45, 64 as well as more recently discovered general alcohol oxidases (AlcOx, EC 1.1.3.13)46, 47 and aryl alcohol oxidases (AAO, EC 1.1.3.7).48 The archetypal galactose oxidase from the phytopathogenic fungus Fusarium graminearum (FgrGalOx) and its engineered variants have found widespread biotechnological applications, including glycoprotein labelling, biosensor development, polysaccharide functionalization, and other applicants involving the oxidation of carbohydrates and other alcohols.92, 121, 122, 124, 126, 128, 130-136 Of particular note, a highly engineered version of FgrGalOx was employed recently in the biocatalytic synthesis of the HIV drug islatravir144. Thus, there is sustained interest to expand the utilization of CROs industrially, including through the discovery of homologs with inherently diverse catalytic specificities.14, 203, 245 Currently, however, very few (currently nineteen) CROs from AA5 have been biochemically characterized,46-48, 53, 54, 62, 89, 106, 108, 118, 204, 206, 246, 247 the majority of which originate from two fungal genera: Fusarium45, 89, 106, 204, 247 and Colletotrichum46-48, 53 (for an updated list see 74 http://www.cazy.org/AA5_characterized.html, ref.43). Nonetheless, recent studies have revealed homologs in these fungi with distinct specificities for alkyl and aryl alcohols, indicating that the large sequence space of uncharacterized AA5 members presents an opportunity to find new biocatalysts.46, 48, 247 Inspired by these results, we were keen to continue mapping activities across the AA5_2 subfamily, drawing upon a growing array of new fungal genomes. Hence, in the present study we have combined molecular phylogeny and sequence similarity network analysis to guide the selection of 40 AA5_2 members from diverse fungi for recombinant expression in the yeast Pichia pastoris. Twelve of these were successfully produced and biochemically characterized, revealing new galactose 6-oxidases, broad-specificity primary alcohol oxidases, and non-carbohydrate alcohol oxidases. 3.2 Results 3.2.1 Molecular Diversity of AA5_2 Members A manually curated multiple protein sequence alignment of over 623 AA5_2 catalytic modules, with signal peptides and additional domains removed, was used to generate a Maximum Likelihood (ML) phylogeny and a sequence similarity network (SSN) (Figure 3.1). Using AA5_1 glyoxal oxidases108, 248 as an outgroup, 39 subgroups were identified based on tree topology and generally high supporting bootstrap values (Figure 3.1A). As indicated, the previously characterized AA5_2 members are broadly dispersed in the tree, with some enzymes clustered into more well-represented clades. The corresponding SSN at an alignment score (bitscore) cut-off of 550 was consistent with the grouping observed in the phylogenetic tree and subsequently used for sequence selection and mapping of biochemical data (Figure 3.1B). In Table 3.1 and subsequently in the text, the targets are named according to their predominant activities as determined by subsequent kinetic assays (vide infra). 75 76 Figure 3.1. Sequence relationships of 623 AA5_2 catalytic modules (A) Phylogenetic tree with bootstrap values supporting the 39 subgroups are indicated at each node/branch. (B) Sequence similarity network (SSN). Each node corresponds to one of the 623 catalytic modules used as an input to build the SSN created in Cytoscape with yFiles Organic layout.249 Edges represent an alignment bitscore threshold of 550, which clusters the sequences into groups as analogous to those observed in (A). In both panels, AA5 members with previously available biochemical data are coloured in black and indicated as (methyl)glyoxal oxidases (UmaGlyOx,108 PchGlyOx1 and PchGlyOx2,248 TciGlyOx1 and TciGlyOx2246) (comprising the outgroup), galactose oxidases (FgrGalOx,56, 62, 66 FoxGalOx,106 FsaGalOx,89 FsuGalOx,204 FveGalOx, and FsuGalOx205), alcohol oxidases (CgrAlcOx and CglAlcOx,46 ChiAlcOx and PoAlcOx,47 raffinose oxidase (CgrRafOx),53 AA5_2 oxidase (PruAA5_2A (PruAlcOx))54 and the aryl alcohol oxidase (CgrAAO).48 Enzymes first characterized in the present study are coloured in neon green. Two selected sequences for which characterization was published contemporaneously, PruAA5_2A (PruAlcOx)54 and PorAlcOx47 are coloured in grey. To further inform target selection, we mapped the variability of the amino acids involved in catalysis and substrate recognition, as determined by mutagenesis and/or structural analysis of AA5_2 members,66, 95, 96, 98-103 onto the SSN (Tables 3.1, B1, and Figure B1). Thus, 40 targets were selected to represent the maximum diversity of fungal ecology and sequence identity possible in this sample size (Table 3.1 and Figure B1). 77 Table 3.1. Key amino acid residues in AA5_2 proteins compared to other characterized members * Enzyme Organism GenBank/JGI mycocosm Accession Active Site Amino Acids Distal Amino Acids Modularity Radical Stabilization Substrate Recognition/Active Site Cavity Shape Catalytic Efficiency FgrGalOx Fusarium graminearum AAO95371 W290 F194 Q326 Y329 R330 Q406 P463 C383 CBM 32-CAT CgrAlcOx Colletorichum graminicola EFQ30446 F W G F M T L C CAT CgrAAO Colletorichum graminicola EFQ27661 Y F E W R T V C PAN 1-CAT FgrAAO Fusarium graminearum XP_011322138 W F E Y K Q P N CBM 32-CAT ExeGalOx Exophiala xenobiotica KIW55415 W F Q F R Q P C CBM-CBM 32-CAT MreGalOx Mytilinidion resinicola XP_033570565 W F Q F R Q P N CBM-CBM 32-CAT FoxGalOxB Fusarium oxysporum FOTG_04629 W M Q F R Q P C UNK-CBM 32-CAT PfeGalOx Penicillium fellutanum 382062 W Y Q Y R Q P S CBM 32-CAT UmaRafOx Ustilago maydis XP_011389156 Y F A W R S G C UNK-CAT PhuRafOx Pseudozyma hubeiensis XP_012186969.1 Y F A W R S G C UNK-CAT AflAlcOx Aspergillus flavus KAF7627372 W Y L Y H E P S UNK-CBM 32-CAT PruAA5_2A (PruAlcOx) Penicillium rubens CAP96757 W Y D Y R E P S CBM 32-CAT FoxAlcOx Fusarium oxysporum FOPG_18201 W F D S K A Q C CBM 32-CAT PorAlcOx Pyricularia oryzae XP_003719369 F F G L Y T L C WSC-CAT GciAlcOx Glomerella cingulata 1901294 F W L F M T L C CAT AsyAlcOx Aspergillus sydowii XP_040706357 F Y H D R V P C UNK-CBM 32-CAT StoAA5 Stogonospora sp OAK97814 W Y E F R D G C CAT CcaAA5 Corynespora cassiicola PSN67470 W A A N E H G C CAT * Non-conserved amino acids in relation to FgrGalOx are bolded. Fungal species: Fgr, Fusarium graminearum; Cgr, Colletotrichum graminicola; Exe, Exophiala xenobiotica; Mre Mytilinidion resinicola; Fox, Fusarium oxysporum; Afl, Aspergillus flavus; Pru, Penicillium rubens; Pfe, Penicillium fellutanum; Por, Pyricularia oryzae; Gci, Glomerella cingulate; Uma, Ustilago maydis; Phu, Pseudozyma hubeiensis; Asy, Aspergillus sydowii; Sto, Stogonospora sp; Cca, Corynespora cassiicola. Modularity: Carbohydrate binding domain (CBM), catalytic domain (CAT), PAN_1 (PF00024), WSC (PF01822), unknown domain (UNK). 78 3.2.2 Protein Production and Initial Biochemical Characterization cDNAs corresponding to the full-length sequence of 40 candidate enzymes, i.e., including any carbohydrate binding modules (CBM250), PAN_1 domains (PFAM, PF00024), WSC domains (PFAM , PF01822), or unknown domains (UNK) in tandem with the catalytic AA5 module, were cloned into the pPICZα-A or pPICZα-C vector in-frame with the Saccharomyces cerevisiae α-factor secretion signal and a C-terminal hexa-histidine tag. Fifteen proteins were thus produced successfully in Pichia pastoris X33 and purified via Immobilized Metal Affinity Chromatography (IMAC), followed by buffer-exchange using size-exclusion chromatography (Figure B2). All proteins were produced full-length except FoxGalOxB, for which a truncated construct was purified (Figure B2). Typical protein yields varied between 2.2 up to 65 mg L-1 based on the original volume of buffered complex methanol medium (Table B3). Overall, protein production levels were comparable to previous AA5_2 preparations from P. pastoris, e.g. : FgrAAO and FoxAAO (2.5 mg L-1),247 FoxGalOx (10.6 mg L-1),106 CgrAlcOx (30 – 40 mg L-1),46 CgrAAO (43 mg L-1),48 and the wild-type and M1 variants of FgrGalOx (120 mg L-1 and 110 mg L-1, respectively).209 The fourteen successfully produced proteins were initially screened for activity using three compounds representing efficient substrates of previously characterized AA5_2 members, viz. galactose and raffinose for galactose 6-oxidases and benzyl alcohol for general alcohol oxidases (Figure 3.2). As expected, a range of specificities was observed, with some enzymes clearly displaying higher specific activity on benzyl alcohol (AflAlcOx and GciAlcOx) or galactosyl moieties (FoxGalOxB, MreGalOx and ExeGalOx), while for others this preference was equivocal (PfeGalOx and PruAlcOx). Notably, several targets had relatively low specific activity on all of the three substrates tested: UmaRafOx, PhuRafOx, CcaAA5, AsyAlcOx, FoxAlcOx, PorAlcOx, 79 and StoAA5. P. pastoris constructs of StoAA5 and CcaAA5 produced soluble protein but the specific activity of the enzyme was not consistent between productions, hence no further work was conducted on these two homologs. Figure 3.2. Initial activity screens of expressed AA5_2 Measurements were performed in triplicate at 30 °C in 100 mM sodium phosphate buffer pH 8.0 using the HRP/ABTS assay. Activities were determined using 300 mM galactose or raffinose, or 30 mM benzyl alcohol. Reactions were started with the addition of 0.5 mM – 1.95 μM of purified enzyme (Table B4). 80 Based on the results of the initial activity screen, either galactose or benzyl alcohol was chosen as the substrate for the determination of the pH-rate profile and temperature stability of each target. Exceptionally, glycerol was used as the substrate for UmaRafOx and PhuRafOx following further substrate screening (specific activities > 3.0 μmol min-1 mg-1) because these enzymes displayed very low specific activities on the three initial substrates (< 0.25 μmol min-1 mg-1). All recombinant enzymes were active between pH 6.0 – 8.5 (Table 3.2) and most exhibited bell-shaped pH-rate profiles with optima between 7.0 – 8.5 in sodium phosphate buffer (Figure B5), akin to other characterized AA5_2 enzymes.46, 48, 89, 106, 251 Distinctly, MreGalOx and ExeGalOx had optimum values at pH 6.0 and 6.5, respectively, while UmaRafOx and PhuRafOx were most active in glycine-NaOH buffer at pH 8.5. Most AA5_2 enzymes were stable at 30 °C, similar to some previously characterized members.46, 48 Three enzymes (FoxGalOxB, AflAlcOx and PruAlcOx) were stable between 35 °C and 40 °C, similar to FoxGalOx (40 °C ),106 and three enzymes (ExeGalOx, MreGalOx and FoxAlcOx) exhibited higher thermostabilities than previously characterized CROs (45 °C - 55 °C) (Table 3.2 and Figure B6). Cumulatively, these results are consistent with mesophilic nature of their natural hosts. Due to instrument limitations, the substrate screening were performed at ambient temperature while subsequent Michalis-Menten kinetic assays were performed at temperatures that balanced optimal enzyme activity with short-term stability. 3.2.3 Extended Substrate Screening and Michaelis-Menten Kinetics After establishing preferred assay parameters for the twelve demonstrably active AA5_2 targets, these enzymes were screened in a parallel, multi-well plate assay against a broader panel of primary alcohol-containing substrates comprising carbohydrates, alkanols, and aryl alcohols with relevance to biomass utilization, pharmaceutical preparation or bio-material synthesis 81 (Figure 3.3). Based upon the outcome of this extended screen, key substrates were selected based on specific activity values for detailed Michaelis-Menten kinetic analysis (Table 3.2 and Figures B7 – B18). The kinetic data for the top substrates of each enzyme are abstracted into Figure 3.3 and Table 3.2. Taken together, these assays enabled us to group the target enzymes by their predominant activities, which are discussed below in the context of the molecular phylogeny and SSN clustering. Figure 3.3. Extended activity screening of AA5_2 enzymes Measurements were performed in triplicate at room temperature in 100 mM buffer, using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 2.5 mM for aldehydes, aryl alcohols and furans. Reactions were started with the addition of 30 pmol – 0.26 nmol of purified enzyme 82 Table 3.2. Initial rate kinetics of active AA5_2 enzymes Enzyme Substrate KM (mM) kcat (s-1) kcat/KM (M-1.s-1) Adjusted R-square value Condition ExeGalOx Galactose 25 ±4.0 130 ±3.7 (5.2 ±0.8) x 103 0.96 pH 6.0, 49 °C Melibiose 25 ±0.6 37 ±0.2 (1.5 ±0.03) x 103 1.0 MreGalOx Galactose 64 ±4.8 32 ±0.7 (5.0 ±0.4) x 102 0.99 pH 6.0, 55 °C Melibiose n.d.§ n.d.§ (1.8 ±0.8) x 102 0.99 FoxGalOxB Galactose 24 ±1.3 81 ±1.0 (3.4 ±0.2) x 103 1.0 pH 8.0, 39 °C Melibiose n.d.§ n.d.§ (5.9 ±1.7) x 103 0.95 PfeGalOx Glycerol 69 ±1.6 140 ±1.1 (2.0 ±0.1) x 103 0.95 pH 8.5, 30 °C Galactose 28 ±6.7 140 ±4.5 (5.0 ±1.2) x 103 1.0 UmaRafOx Glycerol 2900 ±720 130 ±24 45 ±14 1.0 pH 8.5, 30 °C Raffinose 450 ±94 130 ±15 290 ±69 0.99 PhuRafOx Glycerol 2900 ±390 56 ±3.8 19 ±2.9 0.99 pH 8.5, 30 °C Raffinose 410 ±33 8.8 ±0.5 21 ±2.1 1.0 AflAlcOx Benzyl Alcohol 19 ± 1.9 410 ±11 (2.1 ±0.2) x 104 1.0 pH 8.5, 39 °C HMF 5.0 ± 1.3 190 ± 20 (3.8 ±1.1) x 104 0.94 Galactose 980 ± 61 400 ±10 (4.1 ±0.3) x 102 1.0 PruAA5_2A (PruAlcOx) Glycerol 34 ±1.7 170 ±3.0 (4.9 ±0.3) x 103 0.99 pH 7.5, 35 °C Galactose 110 ±18 220 ±8.6 (2.0 ±0.3) x 103 0.98 Benzyl Alcohol 9.2 ±1.8 110 ±5.0 (1.2 ±0.2) x 104 0.97 FoxAlcOx Benzyl Alcohol 230 ±70 24 ±4.5 100 ±37 0.98 pH 8.5, 45 °C Galactose 2000 ±650 8.3 ±1.7 4.1 ±1.6 0.98 PorAlcOx Benzyl Alcohol 1.3 ± 0.10 8.6 ±0.15 (6.6 ±0.5) x 103 0.99 pH 7.5, 30 °C Butanol 4.6 ± 0.30 7.9 ±0.09 (1.7 ±0.1) x 103 0.99 Glycerol 57 ± 8.1 8.5 ±0.32 (1.5 ±0.2) x 102 0.98 1,4-Butanediol 4.3 ± 0.56 9.5 ±0.51 (2.2 ±0.3) x 103 0.97 GciAlcOx Benzyl Alcohol 3.6 ±0.8 350 ±20 (9.7 ±2.2) x 104 0.95 pH 8.0, 30 °C HMF 0.09 ±0.01 240 ±16 (2.7 ±0.3) x 106 0.99 1,4-Butanediol 3.9 ±0.4 330 ±13 (8.5 ±0.9) x 104 0.98 AsyAlcOx Benzyl Alcohol 79 ±22 17 ±1.8 (2.2 ±0.6) x 102 0.96 pH 8.0, 30 °C Glycerol 700 ±80 38 ±2.0 54 ±6.8 1.0 * Error bars represent the standard deviation from the mean values. § Individual KM and kcat values not determinable; kcat/KM values obtained from slope of linear v0 versus [S] plots (see panels B in Figures B7-B18). 83 3.2.3.1 Galactose 6-oxidases Enzymes in this group predominately possess the canonical galactose-6 oxidase activity (EC 1.1.3.9) that first defined the family.51, 87, 210 As such, they have minimal to no activity on other polyols, diols, and other non-carbohydrate primary alcohols. The activity profile of these enzymes is comparable to the archetype, FgrGalOx.203, 252 3.2.3.1.1 ExeGalOx ExeGalOx is located in a separate clade in the phylogenetic tree and it possesses low sequence similarity to its closest homologs FoxGalOxB and the archetypal FgrGalOx (40% and 47%, respectively) (Figure 3.1 and Table B2). ExeGalOx displayed moderate specific activity on galactose and showed lower but equivalent specific activities on raffinose and melibiose, hence we characterized this enzyme as a galactose oxidase (EC 1.1.3.9) (Table B5 and Figure B7). Compared to the FgrGalOx, ExeGalOx had a 3-fold lower catalytic efficiency on galactose due to a lower kcat even though the KM of ExeGalOx was 4-fold lower (Tables 3.2, B11 and Figure B7). In addition, ExeGalOx had a 3-fold lower catalytic efficiency on melibiose compared to galactose, which reflects a lower turnover rate since the KM for both substrates is the same (Table 3.2 and Figure B7). 3.2.3.1.2 MreGalOx MreGalOx also is located in a separate clade sharing low sequence identity with the closest neighbouring homologs: 35% vs. AsyAlcOx and 39% vs. AflAlcOx (Figure 3.1 and Table B2). MreGalOx exhibited low production yields and lower activity compared to other enzymes presented in this study. No activity was observed with many substrates above the limit of detection of our assay (9 x 10-4 μmol min-1 mg-1 using 0.85 μM of purified enzyme). Nonetheless, MreGalOx demonstrated low specific activity on galactose and comparable specific 84 activities on raffinose and melibiose, thus we defined this enzyme as a galactose oxidase (EC 1.1.3.9) (Figure B8 and Table B5). Compared to the FgrGalOx, MreGalOx had a 20-fold lower catalytic efficiency on galactose (Tables 3.2, B11 and Figure B8). Comparing the activity on melibiose versus galactose, MreGalOx had a 3-fold higher catalytic efficiency on melibiose, however KM and kcat parameters could not be reliably determined due to apparent substrate inhibition (Table 3.2 and Figure B8). 3.2.3.1.3 FoxGalOxB FoxGalOxB is located in a separate clade next to that containing the archetypal FgrGalOx and other characterized galactose 6-oxidases (Figure 3.1). Only 40% identity is shared between FoxGalOxB and FgrGalOx, despite the close phylogenetic association (Table B2). The full substrate profile of FoxGalOxB lead us to define it as a galactose oxidase (EC 1.1.3.9). Since a galactose oxidase has already been characterized in Fusarium oxysporum,106 we have named this homolog FoxGalOxB.253 FoxGalOxB showed high specific activity on galactose and melibiose, followed by raffinose and then lactose, whereas low specific activity was observed on polyols, diols, primary and aryl alcohols (Figure B9 and Table B6). In addition, FoxGalOxB showed low specific activity on both HMF and HMFCA (3.3 mol min-1 mg-1) with low specific activity on DFF and no activity on FFCA (Figure B9 and Table B6). Compared to the archetypal FgrGalOx, FoxGalOxB had a 3-fold lower catalytic efficiency on galactose (Tables 3.2, B11 and Figure B9). Substrate inhibition was also observed on melibiose with FoxGalOxB and an overall 2-fold decrease in catalytic efficiency compared to galactose was found (Table 3.2 and Figure B9). 85 3.2.3.1.4 PfeGalOx PfeGalOx is located in the same clade as PruAA5_2 (PruAlcOx) (56% identity) and is located next to the clade containing AflAlcOx (51% identity) (Figure 3.1 and Table B2). Analogous to FoxGalOxB, PfeGalOx displayed high specific activity on galactose, however other galactosylated carbohydrates showed moderate activity, with highest activity observed on raffinose, followed by lactose, and lastly melibiose (Figure B10 and Table B6). Interestingly, PfeGalOx also showed prominent specific activity on xylose (Figure B11 and Table B6), which is a novel activity among CROs. Furthermore, PfeGalOx displayed high specific activity on glycerol, and moderate activities on benzyl alcohol, butanol and 1,2-propanediol/1,3-propanediol, yet low activity on other primary alcohols (Figure B10 and Table B6). Considering the full substrate profile of PfeGalOx, we have defined this enzyme to be a galactose oxidase (EC 1.1.3.9). PfeGalOx had a 2-fold lower catalytic efficiency on galactose than FgrGalOx101 attributable to both an 8-fold decrease in kcat and a 4-fold decrease in KM (Tables 3.2, B11, and Figure B10). Furthermore, PfeGalOx showed a 2-fold higher catalytic efficiency on glycerol than CgrAlcOx,46 however it had a 2.5-fold higher catalytic efficiency on galactose compared to glycerol (Tables 3.2, B12, and Figure B10). 3.2.3.2 Raffinose Oxidases Raffinose oxidases, from the perspective of specificity, can be considered as a subclass of the galactose 6-oxidizes, because these enzymes oxidase 6-OH of the galactosyl residue of this trisaccharide. Nonetheless, these enzymes exhibit a distinct specificity for raffinose over galactose and other galactosides, as has been demonstrated for CgrRafOx.53 These enzymes are also comparably proficient at oxidizing glycerol, some diols, and glycolaldehyde dimer. 86 3.2.3.2.1 UmaRafOx UmaRafOx is located in a distinct clade, which also contains PhuRafOx (81% identity) and the previously characterized CgrRafOx (43% identity) (Figure 3.1 and Table B2). UmaRafOx showed low specific activity on glycerol, compared to other enzymes in the present study. Nonetheless, the highest specific activity was observed on this polyol for UmaRafOx (Figure B11 and Table B7). UmaRafOx showed an overall low specific activity on carbohydrates, however, the substrate with the second-highest specific activity was raffinose (Figure B12 and Table B7). UmaRafOx also displayed a similar specific activity to raffinose on glycolaldehyde dimer, however no activity on methyl glyoxal was observed (Figure B12 and Table B7). UmaRafOx had 20-fold lower catalytic efficiencies on glycerol compared to CgrAlcOx,46 and 15-fold higher catalytic efficiencies on glycerol compared to CgrRafOx (Tables 3.2 and B12). The catalytic efficiencies of UmaRafOx on raffinose is 6-fold higher than CgrRafOx53 (Table 3.2). Based on these kinetic results, UmaRafOx we defined UmaRafOx as a raffinose oxidase (EC 1.1.3.-). 3.2.3.2.2 PhuRafOx PhuRafOx has a specific activity profile similar to UmaRafOx displaying high activity on glycerol, followed by raffinose (Figure B12 and Table B7). PhuRafOx also showed low specific activity on glycolaldehyde dimer with equivalent specific activity observed on 1,3-propanediol (Figure B12 and Table B7). PhuRafOx had 50-fold lower catalytic efficiencies on glycerol compared to CgrAlcOx,46 and 6-fold higher catalytic efficiencies on glycerol compared to CgrRafOx (Tables 3.2 and B12). The catalytic efficiency of PhuRafOx on raffinose is 2-fold lower than CgrRafOx, however, taking into account the fully distinct substrate profile we characterize PhuRafOx as a raffinose oxidase (EC 1.1.3.-). 87 3.2.3.3 Broad-spectrum Alcohol Oxidases Enzymes in this group exhibit a broad catalytic range, i.e. a lack of selectively, for carbohydrates including galactose and galactosides, linear alcohols including diols and polyols, as well as diverse aryl alcohols including benzyl alcohols and furan derivatives. At the same time, these enzymes demonstrate distinct substrate preferences within these compound classes. The activity profiles of these enzymes are reminiscent of those of FgrAAO247 and CgrAAO.48 3.2.3.3.1 AflAlcOx AflAlcOx is located in a separate clade in the phylogenetic tree and shares moderate sequence identity with its closest homologs PfeGalOx and PruAA5_2 (PruAlcOx) (51% and 48%, respectively) (Figure 3.1 and Table B2). AflAlcOx showed high specific activity on benzyl alcohol and displayed moderate specific activity on galactose, glycerol, diols, butanol, HMF and other aryl alcohols (Figure B13 and Table B8). Due to this broad range of activities, we have defined AflAlcOx as a general alcohol oxidase (EC 1.1.3.13 and EC 1.1.3.7). AflAlcOx showed a 7-fold lower catalytic efficiency compared to CgrAlcOx on benzyl alcohol (Table B12) and showed a higher catalytic efficiency on HMF than CgrAAO, bacterial HMFO (a GMC superfamily oxidoreductase) and FgrAAO (2-fold, 5-fold and 20-fold, respectively) (Table B13). Even though AflAlcOx displayed prominent specific activity on galactose, the catalytic efficiency of AflAlcOx on benzyl alcohol was 50-fold higher than on galactose (Table 3.2 and Figure B13). Furthermore, AflAlcOx had 25-fold lower catalytic efficiency on galactose than the archetypal FgrGalOx, due to by a 10-fold increase in KM and 3-fold decrease in kcat compared to FgrGalOx (Table B11). 3.2.3.3.2 PruAA5_2 (PruAlcOx) PruAA5_2 (PruAlcOx) is located in the same clade as PfeGalOx (56% identity) and shares 48% identity with AflAlcOx located in the neighboring clade (Figure 3.1 and Table B2). 88 PruAA5_2 (PruAlcOx) displayed high specific activity on glycerol, moderate activities on 1,2-propanediol and 1,3-propanediol and showed decreased specific activities with increasing chain length on other diols (Figure B14 and Table B8). PruAA5_2 (PruAlcOx) also showed high specific activity on galactose, but low activity was observed for other carbohydrates. Furthermore, high specific activity was also observed on benzyl alcohol and PruAA5_2 (PruAlcOx) showed moderate specific activity on HMF with lower specific activity observed on HMFCA, very low activity on DFF and no activity on FFCA (Figure B14 and Table B8). Based on this wide substrate scope, we have defined PruAA5_2 (PruAlcOx) as a general alcohol oxidase (EC 1.1.3.13 and EC 1.1.3.7). PruAA5_2 (PruAlcOx) displayed catalytic efficiencies 5-fold higher on glycerol and 10-fold lower on benzyl alcohol than CgrAlcOx46 (Table B12). Despite PruAA5_2 (PruAlcOx) possessing a high specific activity on galactose, similar to AflAlcOx, the catalytic efficiency of PruAA5_2 (PruAlcOx) on benzyl alcohol was higher than on galactose (25-fold) (Table B11). PruAA5_2 (PruAlcOx) has a similar KM value compared to FgrGalOx on galactose and shows a 5-fold lower turnover rate (Table B11). During the course of the present study PruAA5_2 (PruAlcOx) was characterized as exhibiting an activity profile combining those of AA5_1 and AA5_2 members.54 Here, we observed generally higher activities on all tested substrates and a different specificity profile. In our hands, PruAA5_2 (PruAlcOx) possessed a higher specific activity on galactose rather than raffinose54 (9-fold) and the highest specific activity was observed on glycerol rather than the glycolaldehyde dimer54 (2-fold) at the equivalent concentrations as the previous study. Furthermore, our enzyme exhibited substantial specific activity on benzyl alcohol (30 mM – 84 mol min-1 mg-1) while the previous study did not observe any activity on this substrate at 300 89 mM.54 The reasons for these discrepancies are unclear, but we note that P. pastoris SMD1168H and fed-batch production was used previously vs. P. pastoris X33 shake-flask production here. 3.2.3.3.3 FoxAlcOx FoxAlcOx is located in the same clade as the recently characterized FgrAAO and FoxAAO (67% identity). This clade is located beside the classical galactose oxidases and FoxAlcOx shares 58% identity with FgrGalOx. FoxAlcOx has generally low specific activity on all substrates tested and displayed the lowest specific activity out of all the enzymes presented in this study (Figure B15 and Table B9). FoxAlcOx showed the highest specific activity on benzyl alcohol with the second highest specific activity observed on galactose and the aldehyde compounds, i.e. methyl glyoxal and glycolaldehyde dimer (Figure B15 and Table B9). The catalytic efficiency of FoxAlcOx on benzyl alcohol is 1400-fold lower than CgrAlcOx46 and 4-fold lower than FgrGalOx211 (Table B12). Due to this broad substrate range, which includes carbohydrates, aryl alcohols and aldehydes (which likely react with the enzyme in their hydrated forms,44, 84 we have classified this FoxAlcOx as a general alcohol oxidase (EC 1.1.3.13 and EC 1.3.3.7). Since the second highest specific activity for FoxAlcOx was observed on galactose, Michaelis-Menten kinetics was performed for this substrate. FoxAlcOx displayed a 2500-fold lower catalytic efficiency on galactose than FgrGalOx (Table B11). 3.2.3.3.4 PorAlcOx PorAlcOx is located in the same distinct cluster as GciAlcOx (57% identity) and CgrAlcOx (55% identity) (Figure 3.1 and Table B2). Similar to MreGalOx, PorAlcOx was produced with low yields (Table B3). Nevertheless, comparable moderate specific activities were measured on polyols, diols, primary alcohols and aryl alcohols with notable activity on benzyl alcohol and HMFCA (Figure B16 and Table B10). In contrast to AflAlcOx and 90 PruAA5_2 (PruAlcOx), PorAlcOx showed higher activity on HMFCA versus HMF while also displaying low activities on DFF and FFCA (Figure B16 and Table B10). The broad substrate profile of PorAlcOx is indicative of a general alcohol oxidase (EC 1.1.3.13) (Figure B16 and Table B10). PorAlcOx had lower catalytic efficiencies compared to CgrAlcOx,46 i.e. kcat/KM values were 20-fold less on benzyl alcohol, 40-fold less on 1,4-butanediol, 80-fold less on butanol and 6-fold less on glycerol (Table 3.2 and B12). During the course of our study, PorAlcOx was characterized using ethanol, butanol, 1,3-butanediol, and glycerol as substrates. Although we did not obtain kinetic parameters on all of the aforementioned substrates, our catalytic efficiencies on butanol and glycerol are comparable to the previously reported values47. 3.2.3.4 Non-carbohydrate Alcohol Oxidases Enzymes in this group clearly appear to reject mono-, di-, and trisaccharides, mostly notably including galactose and raffinose, in favor of alkanols and aryl alcohols. At the same time, these enzymes are generally proficient on linear diols and polyols. The activity profiles of these enzymes are reminiscent of those of CgrAlcOx and CglAlcOx.46 3.2.3.4.1 GciAlcOx GciAlcOx is located in the same clade as PorAlcOx, CgrAlcOx and ChiAlcOx (Figure 3.1). Furthermore, it shares 79% identity with CgrAlcOx (Table B2). GciAlcOx showed equally high specific activities on all diols, glycerol, primary alcohols, benzyl alcohols and substituted aryl alcohols, hence we characterized this enzyme as a general alcohol oxidase (EC 1.1.3.7 and EC 1.1.3.13) (Figure B17 and Table B10). GciAlcOx had catalytic efficiencies comparable to CgrAlcOx46 on benzyl alcohol and 1,4-butanediol and are 50-fold and 40-fold higher, respectively, compared to CgrAAO48 (Tables 3.2 and B12). Furthermore, GciAlcOx displayed high specific activity on HMF and while showing almost no activity on DFF and FFCA (Figure 91 B17 and Table B10). GciAlcOx had the highest reported catalytic efficiency to-date on HMF with a 140-fold higher kcat/KM value than CgrAAO48 and a 380-fold higher kcat/KM value than the bacterial HMFO216 (a GMC superfamily oxidoreductase) (Table B13). As such, GciAlcOx may also be classified as an HMF oxidase (EC 1.1.3.47). 3.2.3.4.2 AsyAlcOx AsyAlcOx is located in separate clade and shares only 45% identity with AflAlcOx in the neighbouring clade (Figure 3.1 and Table B2). AsyAlcOx possessed moderate specific activity on aryl alcohols and diols, with notable specific activity on benzyl alcohol, thus we defined this enzyme as a general alcohol oxidase (EC 1.1.3.13 and EC 1.3.3.7). AsyAlcOx showed low specific activity on the propanediols and displayed enhanced specific activity with increased carbon chain length culminating with 1,6-hexanediol (Figure B18 and Table B9). Michaelis-Menten kinetics indicated that AsyAlcOx had over 600-fold and 17-fold lower catalytic efficiencies on benzyl alcohol and glycerol, respectively, than CgrAlcOx46 (Tables 3.2, B11, and Figure B18). 3.2.4 Application to Bioproduct Conversion CROs from AA5 are finding increasing applications in the conversion of small molecules to valuable chemical intermediates.74, 76, 92, 131, 133, 144, 203, 245 Among these, HMF from the thermochemical breakdown of plant biomass252, 253 and glycerol, a by-product of the biodiesel industry,222 are of particular interest. On one hand, HMF can be converted into the bi-functional polymer precursors DFF and FDCA by controlled oxidation (Figure 3.4A).40, 213 On the other hand, glycerol is an exemplar prochiral compound and its valorization through desymmetrization has garnered significant interest (Figure 3.4B). Relatedly, an evolved variant of FgrGalOx was 92 used for an essential desymmetrization of a key 2-ethynyl-glycerol intermediate in the biocatalytic cascade synthesis of the antiviral drug islatravir144. Figure 3.4. Oxidation pathways of HMF and Glycerol A. HMF oxidation pathway. The symbols next to each compound are used to denote the corresponding peaks in the NMR spectra comprising Figures B19-B23. B. Glycerol oxidation pathway showing L- and D-glyceraldehyde. [O] represents a generic oxidant. 3.2.4.1 HMF Enzyme with specific activities above 3 mol min-1 mg-1 on any furan substrate were incubated with either 10 mM HMF, HMFCA, DFF and FFCA, followed by direct quantitation of the product(s) 1H NMR (Table 3.3). Previous work has shown that CgrAAO oxidizes HMF to DFF while FgrAAO produced a mixture of DFF and FFCA as the terminal product of HMF oxidation.48, 247 93 Table 3.3. Percent conversion of HMF, HMFCA, DFF and FFCA by PruAA5_2A (PruAlcOx), AflAlcOx, GciAlcOx, PorAlcOx and FoxGalOxB AA5 CRO Substrate HMF HMFCA DFF FFCA FoxGalOxB DFF (29%), FFCA (4%) FFCA (17%) FFCA (12%) FDCA (7%) AflAlcOx DFF (52%), FFCA (48%) FFCA (100%) FFCA (40%) 0% PruAA5_2 (PruAlcOx) DFF (52%), FFCA (48%) FFCA (100%) FFCA (38%) FDCA (5%) GciAlcOx FFCA (91%), FDCA (9%) FFCA (92%), FDCA (8%) FFCA (87%), FDCA (13%) FDCA (24%) PorAlcOx DFF (49%), FFCA (2%) FFCA (83%), FDCA (17%) FFCA (72%), FDCA (28%) FDCA (49%) GciAlcOx showed 100% conversion of HMF to 91:9 mixture of FFCA and FDCA, respectively (Table 3.3 and Figure B19). Based on experiments with the pathway intermediates HMFCA and DFF, GciAlcOx is able to oxidize both furans efficiently with the major product consisting of FFCA with some FDCA observed (Table 3.3 and Figure B19). PorAlcOx, which belongs to the same cluster as GciAlcOx, showed incomplete conversion of 10 mM HMF producing predominantly DFF with 2% FFCA observed (Table 3.3 and Figure B20). Interestingly, PorAlcOx showed full conversion of HMFCA and DFF to mostly FFCA and partial conversion to FDCA was observed (Table 3.3 and Figure B20). PruAA5_2A (PruAlcOx) oxidized 100% of 10 mM HMF and peaks corresponding to DFF, including its hydrate form, and FFCA were observed (Table 3.3 and Figure B21). AflAlcOx showed similar percent conversions to PruAA5_2A (PruAlcOx) for HMF, HMFCA, and DFF (Table 3.3 and Figure B22). The main difference, no conversion was observed with FFCA for AflAlcOx. FoxGalOxB displayed low percent conversion of HMF and had a similar product profile to PruAA5_2A (PruAlcOx) (Table 3.3 and Figure B23). 94 From previous work, FgrAAO, an aryl alcohol oxidase from Fusarium graminearum, was shown to produce similar ratios of products in overnight incubations with furan-containing substrates compared to PruAA5_2A (PruAlcOx), AflAlcOx and FoxGalOxB.247 It was noted for FgrAAO, and other CROs, that perhaps the low degree of hydration of FFCA compared with DFF causes the reactions with furans to end with FFCA.216 In contrast, the increased production of FDCA observed for both GciAlcOx and PorAlcOx may be caused by the active site architecture of these enzymes being able to accommodate the aldehyde form of FFCA which results in higher percent conversion to FDCA compared to other CROs 3.2.4.2 Glycerol It has been previously reported that the wild-type FgrGalOx and FgrAAO oxidize glycerol stereoselectively to L-glyceraldehyde, while CgrAlcOx produces the natural D-glyceraldehyde isomer.223, 247 To explore the stereoselectivity of the CROs in the present study, those with specific activities toward glycerol of greater than 3 mol min-1 mg-1 were incubated with 0.54 M glycerol for 24 hours, followed by chiral HPLC analysis of the resulting 2,4-dinitrophenyl glyceraldehyde-hydrazones (Table 3.4 and Figure B24). 95 Table 3.4. Stereochemical outcome of glycerol oxidation by AA5 CROs AA5 CRO Product L-glyceraldehyde hydrazone (%) D-glyceraldehyde hydrazone (%) FgrGalOx 96 4 CgrAlcOx 10 90 FoxGalOxB 79 21 PfeGalOx 71 29 UmaRafOx 84 16 PhuRafOx 82 18 AflAlcOx 88 12 PruAA5_2A (PruAlcOx) 96 4 GciAlcOx 3 97 PorAlcOx 11 89 AsvAAO 77 23 GciAlcOx and PorAlcOx showed a preference for producing D-glyceraldehyde with an er of (3:97 and 11:89, respectively) (L-:D-) which is expected, since both share higher identity with CgrAlcOx (79% and 55%, respectively) than with FgrGalOx (38% and 34%). On the other hand, all the other enzymes tested for the stereo-selectivity of glycerol oxidation showed strong preference for the formation of L-glyceraldehyde (Table 3.4 and Figure B24). These enzymes all have a higher identity to FgrGalOx (52 – 36%) than CgrAlcOx (33 – 23%) (Table B2), providing further evidence that the different amino acid scaffolds of the active site of FgrGalOx and CgrAlcOx direct the stereo-selective oxidation of achiral glycerol. 96 3.3 Discussion Biocatalysis will play an increasingly important role in the development of a sustainable economy, and the discovery of novel enzymes, particularly oxidases, is essential to support this transition. Only recently has it been appreciated that the range of native catalytic activities found among the copper radical oxidases from Auxiliary Activity Family 5 Subfamily 2 members extends beyond the founding, classical galactose 6-oxidases.46-48 In the present study, we have nearly doubled the number of biochemically characterized AA5_2 members to reveal a diverse spectrum of CRO specificities. These comprise four canonical galactose-6-oxidases (ExeGalOx, MreGalOx, FoxGalOxB and PfeGalOx), two raffinose-specific galactosyl oxidases (UmaRafOx and PhuRafOx), four broad spectrum alcohol oxidases (AflAlcOx, PruAA5_2A (PruAlcOx), FoxAlcOx and PorAlcOx), and two alcohol oxidases that notably do not address carbohydrates (GciAlcOx and AsyAlcOx). Mapping these activities onto the current phylogenetic landscape of AA5_2 indicated that although a number of clades demonstrate conserved substrate specificities, galactosyl versus general alcohol oxidase activity was not monophyletic. However, detailed analysis of active-site residues, supported by structural homology modeling based on FgrGalOx (PDB ID 1GOF)56 and computational docking of galactose (Figures 3.5 and B25-B27), does allow us to rationalize some structure-function relationships among the CROs in the absence of experimental tertiary structures of these homologs. 97 Figure 3.5. Molecular docking of galactose (cyan) into the experimental tertiary structure of FgrGalOx (PDB ID 1GOF)56. Copper (dark orange), W290 (yellow), Y329 (green), R330 (magenta), F494 (brown) and F194 (red). Ligand coordination is indicated, specifying the distances (black). Four CROs displayed similar substrate profiles to the archetypal galactose oxidase, however, ExeGalOx, MreGalOx, FoxGalOxB have a distinct difference with a phenylalanine at the position corresponding to Y329 in FgrGalOx (Table 3.1). Although this residue has been suggested to form a hydrogen bond with the 1-OH of galactose95 (Figure 3.5), ExeGalOx, MreGalOx, and FoxGalOxB all have lower KM values than the archetype (Table B11), thus indicating that tyrosine is dispensable to maintain efficient galactose binding. Computational docking suggests that other predicted hydrogen bonds to active-site arginine and tryptophan residues may be sufficient for galactose binding in these enzymes, such as seen in ExeGalOx (Figure B25B). Consistent with this hypothesis, three broad spectrum alcohol oxidases (FoxAlcOx, AflAlcOx and PruAlcOx) also showed prominent activity on galactose. Of these, the enzymes with 98 KM values equal to, or lower, than FgrGalOx on galactose retain an arginine at the corresponding position (Arg330, Table 3.1 and B10). This amino acid has been speculated to make two hydrogen bonds with the 3-OH and 4-OH of galactose,100 which is supported by docking analysis of FgrGalOx and ExeGalOx (Figures 3.5 and B26B). However, enzymes with a lysine (FoxAlcOx) or a histidine (AflAlcOx) at this position have higher KM values, possibly caused by the loss of hydrogen bonding interactions. FoxAlcOx possess a lysine that only forms one predicted hydrogen bond to galactose (2.7 Å) (Figure B26C), and the limited activity of this homolog is similar to previously studied AA5_2 members with this substitution101, 247 (Table B11). Likewise, the longer effective distance between the histidine and galactose modelled in AflAlcOx likely diminishes the potential of hydrogen bonding (Figure B26D). With the exception of CgrAAO48 and AsyAlcOx, other enzymes with no activity on galactose feature a methionine or tyrosine residue at the position corresponding to Arg330 in FgrGalOx (Table 3.1). Another residue that appears to be important in determining galactose binding is F194, which together with F464, is speculated to form a hydrophobic wall in FgrGalOx.100 Our models indeed support that this wall may interact with the comparatively hydrophobic bottom face (defined by the axial H-1, H-3, and H-5) of -galactopyranoside, as well as the exocyclic CH2 group (Figure 3.5). Enzymes with a substitution at the position corresponding to F194 in FgrGalOx and demonstrable galactose oxidase activity (> 60 mol min-1 mg-1, viz. PfeGalOx, AflAlcOx, PruAA5_2A (PruAlcOx) with tyrosine and FoxGalOxB with methionine), all have lower kcat values and overall lower catalytic efficiencies, despite some having lower KM values (PfeGalOx and PruAA5_2A (PruAlcOx) (Table B11). Modeling of PfeGalOx, AflAlcOx, and PruAA5_2A (PruAlcOx) with galactose also suggests that the corresponding tyrosine can also form a hydrophobic wall with the bridging phenylalanine. However, this tyrosine is computed to 99 be within hydrogen bonding distance to the ring oxygen of galactose and, consequently, may orient the 6-OH further away from the copper center compared to FgrGalOx, thereby attenuating activity (Figure B27B-D). Confoundingly, FoxGalOxB, has a methionine in the position corresponding to position 194 in FgrGalOx (Table 3.1). Despite the obvious disruption of the hydrophobic wall (Figure B27E), FoxGalOxB possesses predominant GalOx activity, albeit with a low turnover rate. The binding mode of the galactose in our model appears to be non-productive, since neither of the hydrogen atoms on the hydroxymethyl group are positioned to interact with the catalytic tyrosine (Y272 in FgrGalOx) for abstraction. Although this modelled complex precludes definitive analysis in the absence of site-directed mutagenesis and kinetic analysis of variants, it is notable that the low turnover rate observed for FoxGalOxB is matched by a low KM value. The same hydrogen bonds between galactose and arginine and tryptophan indicated in other enzymes are maintained FoxGalOxB, consistent with this observation (Table B11). In addition to the obvious residues lining the active site pocket, W290 in FgrGalOx is a second-shell amino acid which strongly affects catalysis in AA5_2 CROs. FgrGalOx W290 stacks with covalent tyrosine-cysteine radical that is central to the mechanism of CROs, and potentially makes a hydrogen bond to the ring oxygen of galactose (Figure 3.5). Mutagenesis of W290 significantly diminishes catalytic efficiency.66 The homologs CgrAAO and CgrAlcOx possess a tyrosine and a phenylalanine at this position, respectively, site-directed mutation of which to Trp modulates the relative activities toward carbohydrates versus other primary alcohols.46, 48 On the other hand, the recently characterized FgrAAO and FoxAAO possess a tryptophan at the same position, yet have much higher catalytic efficiencies on aryl alcohols than galactose.247 As such, the presence of this Trp as a galactose specificity determinant is somewhat ambiguous. However, 100 we note that three general alcohol oxidases (PorAlcOx, GciAlcOx, and AsyAlcOx) and the two raffinose oxidases (UmaRafOx, PhuRafOx), all of which showed low or no activity on galactose (< 3 mol min-1 mg-1) do not carry a Trp at this position. PorAlcOx, GciAlcOx, and AsyAlcOx have a phenylalanine at this position, while the raffinose oxidases have a tyrosine (Table 3.1). The other enzymes in our study that oxidize galactose all have the W290 residue conserved, like the canonical FgrGalOx. The last amino acid that warrants comment is C383, which lies at the back of the active site pocket but does not directly interact with galactose in FgrGalOx. This residue has been shown previously to affect FgrGalOx catalytic efficiency on galactose,96 as its replacement with a serine increased the catalytic efficiency of the mutant while an asparagine substitution decreases the catalytic efficiency 10-fold.96, 247 X-ray crystallography demonstrated that the C383S mutant of FgrGalOx participates in the H-bonded water network, stabilizing the backbone nitrogen between residues Tyr495 and His496 possibly increasing activity.99 In our study, AflAlcOx, PruAlcOx and PfeGalOx have some of the highest catalytic efficiencies from all enzymes produced in this study and they all natively have the C383S substitution (Table 3.1). Furthermore, MreGalOx has the same active site structure as ExeGalOx but has a 10-fold decreased catalytic efficiency on galactose (Table 3.2). MreGalOx has an asparagine at the position corresponding to C383 in FgrGalOx which, due to a bulkier side chain, could disrupt the hydrogen bonding network, thereby lowering the activity of the enzyme. Although the contributions of individual active-site amino acids to the specificity of AA5_2 CROs is not completely unambiguous, we can nonetheless discern some general trends based on kinetic observations for the complete set of enzymes characterized to-date (based on FgrGalOx residue numbering): the presence of R330 is crucial for low KM values on galactose and 101 galactosides while its replacement generally favours non-carbohydrate oxidation. The neighbouring amino acid, Y329, is not a key determinant of galactose oxidase binding/activity and a variety of amino acid residues are found at this position in CROs (Figure B1F). In contrast, F194 is important for efficient turnover rates for galactose and although this amino acid is mostly conserved among AA5_2 sequences (Figure B1D), the majority of other sequences regardless of carbohydrate or non-carbohydrate activity contain an aromatic residue at this position likely to maintain the hydrophobic wall with F464. Lastly, the presence of W290 allows for prominent galactose oxidase activity but alone does not dictate the enzymes specificity. 3.4 Conclusion A growing body of functional information on AA5 members, including the detailed biochemical characterization of twelve fungal orthologs in our present study, is continuing to reveal a surprisingly broad catalytic diversity within this family of CROs. In this context, we note that automated genome annotations often label AA5_2 members as “galactose oxidases” based on sequence similarity to the archetype FgrGalOx, which misrepresents the wider potential of these enzymes. Thus, such annotations should be viewed with caution, and we would generally advocate that these proteins should be more conservatively annotated as “copper radical oxidases”84 until specificity can be reasonably established. We also note that the true physiological substrates of these enzymes are generally unknown, so specificities determined in vitro – including those here – may only be indicative. Nonetheless, it is revealing that AA5_2 comprises apparent galactose oxidases, raffinose oxidases, broad spectrum alkyl- and aryl-alcohol oxidases, and non-carbohydrate alcohol oxidases of potential use in the specific biocatalytic transformation of both carbohydrate and non-carbohydrate substrates, including bioproducts such as HMF and glycerol. 102 The replacement of chemical oxidants with redox enzymes is a topic of considerable contemporary interest, for which the palette of CROs constitutes an alternative to co-factor dependant oxidases. For example, GciAlcOx had the highest catalytic efficiency on HMF currently reported, compared to CgrAAO48 and FgrAAO,247 the bacterial HMF oxidase (a GMC superfamily oxidoreductase),38 and some glyoxal oxidases from AA5_1 (Table B12).214-216 On the other hand, AflAlcOx may find use in multiple possible biocatalytic applications because it has high activity on both carbohydrate and non-carbohydrate substrates. The optimal choice of enzyme for a specific biocatalytic process will of course be substrate-dependent, for which the current palette of AA5_2 CROs serves as an immediate resource and a starting point for future enzyme engineering efforts. 3.5 Materials and Methods 3.5.1 Chemicals and enzymes Distilled water was used for the preparation of all buffers and stock solutions unless stated otherwise. Catalase from bovine liver (2000–5000 units per mg protein, Sigma) and horseradish per-oxidase (Rz > 3, 300 units per mg, Bio Basic Canada Inc.), obtained as lyophilized powders, were used as received. Other substrates and reagents were purchased from commercial sources (Sigma-Aldrich, VWR or Fisher) and used without further purification. 3.5.2 Sequence Analysis and Bioinformatics A total of 247 bacterial and 103 fungal AA5_2 sequences were collected from the CAZy database in January 2019.43 Sequences of characterized AA5_1 members were also retrieved from the CAZy database. In addition, full length sequences of FgrGalOx (Uniprot P0CS93), CgrAlcOx (Genbank EFQ30446) and CgrAAO (Genbank EFQ27661), as well as their corresponding domains, were used as templates for BLAST analysis against all Fungi in the Mycocosm portal for 103 fungal genomics (https://genome.jgi.doe.gov/programs/fungi/index.jsf).254, 255 Where present, signal peptides and additional modules, such as carbohydrate-binding modules, were removed to isolate the catalytic modules for subsequent analysis. Catalytic modules sharing 100% identity were down-sampled to one sequence to eliminate redundancy. A multiple sequence alignment was created with MAFFT v.7.402 using the L-INS-I algorithm256, on the CIPRES Science Gateway (www.phylo.org).229 Using this alignment, catalytic modules having deletions/substitutions at key active site residues, namely Cys272 and Tyr316 (which form the cross-linked thioether-tyrosyl cofactor) and other copper coordinating residues of FgrGalOx64 were removed. Moreover, obviously erroneous sequences generated by incorrect splicing predictions were removed through identification of unusually long deletions/insertions. With this curated multiple sequence alignment, comprising 623 AA5_2 catalytic modules, a maximum likelihood phylogenetic tree was produced using RAxML v.8, with 100 bootstrap replications,228 on the CIPRES Science Gateway portal. The resulting tree was visualized using FigTree and monophyletic groups were supported by bootstrap values >75, and tree topology. SSNs were generated by computing BLASTP257 all-versus-all local alignments of the curated 623 AA5_2 catalytic domains using SSNpipe (https://github.com/ahvdk/SSNpipe), which generated the E-value, bit score, alignment length, sequence identity, and sequence similarity for all sequence pairs. The data were filtered using a bit score threshold between 500 to 600, with increments of 25, to generate the final SSNs. A bit score threshold of 550, clustering the sequences into groups, which resolves the same monophyletic groups as those observed in the phylogenetic tree, was retained. The SSNs were visualized with Cytoscape using yFiles Organic Layout,249 and colouring of each node was based on monophyletic groups inferred from the phylogenetic tree. 104 In addition, Phyre 2226 was used to make three-dimensional homology models for AflAlcOx, PruAA5_2A (PruAlcOx), PfeGalOx, ExeGalOx, FoxGalOxB, FoxAlcOx and MreGalOx. The structural models were aligned to the crystal structures of FgrGalOx (PDB ID 1GOF)56 and CgrAlcOx (PDB ID 5C86)46 for spatial amino acid comparison to give further support to the alignment. 3.5.3 DNA Cloning and Recombinant Strain Production cDNA encoding genes of interest without the predicted native signal peptide, and including a C-terminal His6 tag-encoding sequence, were commercially synthesized in a codon-optimized form (BioBasic, Markham, Canada) and cloned directly into pPICZα-A or pPICZα-C using the EcoRI, SfiI, ClaI, XbaI and NotI restriction sites flush with the sequence encoding the S. cerevisiae a-factor signal peptide, and transformed into chemically competent E. coli DH5α by heat shock. All recombinant proteins were produced using the in-house 3PE Platform (Pichia Pastoris Protein Express www.platform3pe.com) as described in Haon et al., 2015.258 The transformation of competent P. pastoris X33 cells with 5 µg of PmeI-linearized pPICZα-A or or pPICZα-C plasmids (Invitrogen, Cergy-Pontoise, France) containing the targeted sequences were done by electroporation. Each construct was spread onto plates containing either 100 or 500 µg mL-1 of Zeocin, and three P. pastoris transformants were isolated from each plate. Zeocin-resistant P. pastoris transformants were then screened for protein expression in 24-deepwell plates with the following conditions: induction of expression was done for 3 days at 25 °C in BMMY supplemented with CuSO4 (500 µM) and methanol (3%, v/v), with daily addition of 1% (v/v) methanol. The recombinant proteins were then purified from supernatant by affinity chromatography on Ni-NTA resin using automated procedure as described previously258. Finally, 105 the analysis of purified recombinant protein expression by SDS-PAGE (8% polyacrylamide precast E-PAGE gels, Invitrogen) using Coomassie blue staining, was used to select the best-producing transformants. 3.5.4 Protein Production and Purification Small Scale Protein Production and Purification The best-producing transformants were streaked on yeast extract peptone dextrose (YPD) agar plates containing Zeocin (100 µg mL-1) and allowed to grow three days (30 °C). One colony of the selected transformants served to inoculate 5 mL of YPD broth in 50 mL sterile conical tubes, and were incubated for five hours at 30 °C and 160 rpm in an orbital shaker. The preculture was used to inoculate (0.2%, v/v) 500 mL of BMGY, before incubation 16 hours (30 °C) until the OD600 nm reached 4-6. Then, the biomass was harvested by centrifugation (10 minutes, 16 °C, 5,000 g) and resuspended in 100 mL of BMMY medium supplemented with 500 µM of CuSO4 and 3% methanol (v/v) and incubated for 3 days in an orbital shaker (200 rpm, 25 °C), with daily feeding of 1% methanol (v/v). For PorAlcOx, StoAA5 and CcaAA5 clones, proteins expression was further improved by lowering induction temperature to 20 °C and increasing daily methanol feed to 3% (v/v). The cells were then harvested by centrifugation (10 minutes, 4 °C, 5,000 g), the supernatants were filtrated on 0.45 µm membrane (Millipore, Burlington, Massachusetts, USA) and stored at 4 °C prior to purification. The filtered supernatants were adjusted to pH 7.8 just before purification and filtered on 0.22 µm filters (Millipore, Molsheim, France) and were loaded onto a HisTrap HP-5 mL column (GE Healthcare, USA), equilibrated with buffer A (Tris-HCl 50 mM, pH 7.8, NaCl 150 mM, imidazole 10 mM) and connected to an Äkta Xpress system (GE Healthcare). (His)6-tagged recombinant proteins were eluted with 50% buffer B (Tris-HCl 50 mM pH 7.8, NaCl 150 mM, 106 imidazole 500 mM). After SDS-PAGE (10% polyacrylamide precast gel, Bio-Rad) analysis, fractions containing the recombinant enzymes were pooled, concentrated and exchanged to 50 mM Tris-HCl pH 7.0. The concentrated proteins were incubated overnight at 4 °C with 1 mM CuSO4 to ensure full copper loading and then buffer exchange in 50 mM Tris-HCl buffer pH 7.0 to remove CuSO4 excess. The protein concentration was determined by UV absorption at 280 nm using a Nanodrop ND-200 spectrophotometer (Thermo Fisher Scientific, USA) and calculated using theoretical molecular weight and the molar extinction coefficients were determined with ProtParam web tool (https://web.expasy.org/protparam/) (ExeGalOx ε = 142,522 M-1 cm -1; MreGalOx ε = 153,842 M-1 cm -1; FoxGalOxB ε = 140,907 M-1 cm -1; PfeGalOx ε = 131,967 M-1 cm -1; UmaRafOx ε = 109,062 M-1 cm -1; PhuRafOx ε = 105,052 M-1 cm -1; AflAlcOx ε = 134,885 M-1 cm -1; PruAA5_2 (PruAlcOx) ε = 126,802 M-1 cm -1; FoxAlcOx ε = 131,967 M-1 cm -1; PorAlcOx ε = 89,645 M-1 cm -1; GciAlcOx ε = 95,527 M-1 cm -1; AsyAlcOx ε = 123,947 M-1 cm -1; CcaAA5 ε = 77,475 M-1 cm -1; StoAA5 ε = 69,455 M-1 cm -1). Large Scale Protein Production and Purification Single colonies of Pichia pastoris X33 expressing clones were individually streaked onto agar plates containing 500 μg mL-1 of the antibiotic Zeocin and grown for 2 days in the dark in a 30 oC incubator. Precultures containing 5 mL of YPD and 0.4 mg mL-1 Zeocin were inoculated using a single colony and shaken at 30 oC at 250 rpm for 9 h. Biomass production was initiated by the addition of the preculture into 1 L of BMGY media shaken in 4 L beveled flasks at 30 oC at 250 rpm overnight. Once the BMGY cultures reached an OD600 of 6-12, the cells were harvested by centrifugation at 3000 g for 15 mins at room temperature. The cells were then resuspended using 400 mL of BMMY media supplemented with 0.5 mM copper sulfate containing 3% methanol and transferred to 1 L flasks with a foam cap. The flasks were shaken at 107 250 rpm at 16 °C for three days. The cultures were fed 1% (v/v) methanol every 24h to maintain continued expression of the recombinant protein. On the last day of methanol induction, the desired proteins were separated from the cells by centrifugation at 8000 g for 15 mins at 4 °C. The supernatant was quickly decanted, filtered through 0.45 μm membrane and stored at 4 oC till purification. The pH of the liquid medium which contained the proteins of interest was raised before purification to 7.5-8.0 by the dropwise addition of 1 M NaOH to the stirred solution. The medium was then filtered through a 0.45 μm membrane and allowed to equilibrate for at least 12 h at 4 ˚C. The supernatant was allowed to pass through a 5 mL pre-packed Ni-NTA column, pre-equilibrated with 50 mM sodium phosphate buffer at pH 7.5 with 300 mM NaCl and 10 mM imidazole at 5 mL/min. The column was washed with 10 column volume of equilibration buffer containing 50 mM sodium phosphate buffer at pH 7.5 with 300 mM NaCl and 10 mM imidazole at 5 mL/min. Proteins were eluted with a linear gradient of 2% to 100% of 500 mM imidazole in a 50 mM sodium phosphate buffer with 300 mM NaCl at 5 mL/min. The total elution volume was 125 mL collected in 1mL fractions. The fractions of interest were pooled and concentrated using a 30,000 MWCO Vivaspin centricone and 0.25 – 1 mL was loaded onto a Sephadex G-25 column pre-equilibrated with 50 mM sodium phosphate buffer at pH 7.5 at 5 mL/min. A total volume of 200 mL of equilibration buffer was passed through the column at 5 mL/min. The fractions of interest were pooled and concentrated using another 30,000 MWCO Vivaspin centricone. The proteins were aliquoted, flash frozen in liquid nitrogen and stored at -70 °C. SDS-PAGE was performed using pre-cast 4-20% (w/v) polyacrylamide gel in the presence of 2% (w/v) SDS under reducing conditions. Proteins were visualized using Coomassie blue R-250. 108 Protein concentrations were determined by measuring A280. The extinction coefficients were derived from the sequence using the ProtParam tool on the ExPASy server. 3.5.5 Analytical Protein Deglycosylation The presence of protein glycosylation on UmaRafOx, PhuRafOx, AflAlcOx, PruAA5_2A (PruAlcOx), ExeGalOx, MreGalOx, PfeGalOx, FoxGalOxB, FoxAlcOx, GciAlcOx, AsyAlcOx, PorAlcOx, StoAA5 and CcaAA5 was assessed by treatment with N-glycosidase F from Flavobacterium meningosepticum (PNGaseF, New England Biolabs). Deglycosilation experiments were performed under denaturing condition by adding 5 μg of protein to 10X Glycoprotein Denaturing Buffer and heated for 10 min at 100 °C. The samples were subsequently diluted to 20 μL with GlycoBuffer 2 and tertigol-type NP-400 detergent. Finally, 1 μL of PNGaseF was added to the sample and incubated for 1 hour at 37 °C. Changes in protein mobility was assessed by SDS-PAGE stained with Coomassie blue R-250. Molecular wights of proteins were estimated using a standard curve of the log (MW) versus Rf of the protein ladder (BLUelf). 3.5.6 Initial Activity Test The colorimetric HRP – 2,2’-azinobis(3-ethylbenzthiazoline-6-sulfonic acid) (ABTS)-coupled assay was used to determine the kinetics of the enzymatic oxidation of substrate. The oxidation of the alcohol group on the substrates by AA5 enzymes consumes 1 equivalent of O2 and produces 1 equivalent of H2O2. The oxidation of ABTS (λmax = 514 nm, ε – 36000 M- 1cm- 1)259 is catalyzed by the enzymes HRP using 2 equivalents of H2O2. This is a standard assay used to monitor initial rate kinetics for AA5 enzymes.209 This assay was optimized for sensitivity and linearity. The enzymes that expressed well from the production screen were tested for activity on 300 mM galactose, 300 mM raffinose and 30 mM benzyl alcohol. Enzyme 109 activity was measured using the HRP-ABTS coupled assay comprised of 100 mM sodium phosphate buffer at pH 8.0, 0.46 mM ABTS and 21 U/mL of HRP at 30 oC using a Cary 60 UV-VIS spectrometer. 3.5.7 Plate Assays Proof of Concept The enzyme AflAlcOx was used for the proof-of-concept experiments for the development of a medium through-put plate-based HRP-ABTS-coupled assay. The pH profile, temperature stability profile and partial substrate screen for AflAlcOx were performed initially using the standard HRP-ABTS coupled assay using a Cary 60 UV-VIS spectrometer with plastic 1 mL cuvettes (Figure B4). The same experiments using the same conditions were subsequently performed using a 96-well plate assay with BioTek Epoch microplate spectrophotometer. All experiments with AflAlcOx performed on the Cary 60 UV-VIS spectrometer were equivalent to experiments performed using the BioTek Epoch microplate spectrophotometer (Figure B4), validating the plate assay. This assay was optimized for sensitivity and linearity. A standard curve for the production of ABST radical was performed by incubating 7 mM ABST with 12.25 – 22.05 nmole potassium persulfate for 16 hours in the dark to correctly assess the pathlength of the 96-well plate259 (Figure B3). pH Profile Enzyme activity across a wide range of pH values was determined using phosphate citrate (pH 4.0-7.0), sodium phosphate (pH 5.5-8.5), glycine-NaOH (pH 8.5-11.0) and CHES (pH 8.5-10.50) buffers. Enzyme activity was measured using the HRP-ABTS coupled assay in a 96-well plate assay at RT in 100 mM buffer with either 300 mM of galactose, 30 mM benzyl alcohol or 300 mM of glycerol depending on the initial activity screen test. 110 Temperature Stability Temperature stability of candidate enzymes was determined by first diluting the stock protein in 50 mM sodium phosphate buffer at the optimum pH determined for each enzyme previously. The diluted protein was then pre-incubated in a thermocycler at 30 °C, 39 °C, 49 °C, 60 °C, 69 °C and 79 °C. Samples were taken out at different time interval and the activity of the proteins was measured using the HRP-ABTS coupled assay in a 96-well plate assay at RT with 300 mM galactose, 30 mM benzyl alcohol or 300 mM glycerol as the substrate based on previous experiments. Substrate Screen The activity of candidate enzymes was surveyed on a variety of substrates using the HRP-ABTS coupled assay comprised of 100 mM sodium phosphate buffer at pH value determined by the pH profile, 0.46 mM ABTS and 21 U/mL of HRP at RT using the 96-well plate assay method. The initial substrate screen included carbohydrates, polyols, diols and primary alcohol substrates at 300 mM, benzyl alcohol and galactitol at 30 mM, methyl glyoxal and glycolaldehyde dimer at 5 mM, aryl alcohols and furans at 2.5 mM. One unit of AA5 enzyme activity was defined as the amount of enzyme required to oxidize 2 μmol of ABTS per minute, which is equivalent to the consumption of 1 μmol of oxygen per minute. 3.5.8 Michaelis-Menten Kinetics To determine Michaels-Menten parameters of candidate enzymes, different concentrations of substrate solutions were used over the range of 0.1 - 1500 mM for galactose, 0.5 – 750 mM for melibiose, 0.1 – 285 mM for benzyl alcohol, 0.5 - 200 mM for butanol, 5 – 6000 mM glycerol, 0.1 – 200 mM 1,4-butanediol, and 0.01 – 25 mM HMF. The reactions were performed using the HRP-ABST coupled assay with 0.46 mM ABTS, 21 U/mL of HRP at 111 different temperatures in 100 mM buffer at different pH informed by the temperature stability assays and pH profile experiments using the Cary 60 UV-VIS spectrometer with plastic 1 mL cuvettes. More detailed information for the experimental conditions of each Michaelis-Menten experiment can be found in Table 3.2. Data were fit with the Michaelis−Menten equation using OriginPro software (OriginLab 9.55). 3.5.9 Computational Docking Studies Molecular docking simulations were performed using the CHIMERA software from UCSF Resource for Biocomputing, Visualization, and Informatics.231 The FgrGalOx crystal structure (PDB ID 1GOF) and a Phyre model of AflAlcOx, PruAlcOx, PfeGalOx, ExeGalOx, FoxGalOxB and FoxAlcOx were used to generate the receptors for simulations. Galactose was extracted from a different PDB file. Ligands and receptors were first prepared for docking in chimera by adding hydrogens and assigning proper protonation states. The docking simulation itself was performed using Autodock VINA, run within CHIMERA, with the AMBER03 force field.232 Appropriate simulation cells were defined for the respective docking simulations. For docking of galactose, a 7 Å, 6x6x7 Å or a 6 Å with the copper atom bordering the z-coordinate edge was chosen. Galactose gave a reasonable binding pose with both all enzymes used. The created receptor-ligand complex structures were further processed using the PyMOL software from Schrodinger LLC involving the identification of ligand-receptor interaction and for the determination of distances. 3.5.10 Enzyme Product Analysis Oxidation of HMF, DFF, HMFCA, FDCA and furfural 10 mM of substrate (HMF, DFF, HMFCA, and FDCA) and 1 mg mL-1 of both catalase and HRP were combined and the reaction was initiated by adding 60 μg of purified AA5_2 112 enzyme to a vial with 50 mM buffer in a final volume of 1 mL. Reactions were stirred at 400 rpm at room temperature for 17 hours followed by enzyme removal by ultrafiltration (5 kDa cut-off Vivaspin Sartorius, Stonehouse, UK). D2O was added to the filtrate to a final composition of 10% (v/v). NMR experiments were performed on a Bruker AVANCE 600 MHz spectrometer. 1H NMR spectra were collected with water suppression (4.7 ppm) using a standard pre-saturation pulse sequence. Chemical shifts were calibrated to the internal HOD peak (4.7 ppm). Standards of all substrates were used to identify distinct chemical shifts for each molecule. The integration value for relevant peak areas were used to determine percent conversions. Oxidation of Glycerol 0.54 M glycerol and 1 mg mL-1 of both catalase and HRP were combined and 700 μg of purified AA5_2 enzyme in a final volume of 1 mL (50 mM buffer) was added to start the reaction. Reactions were stirred at 400 rpm at room temperature for 24 hours, at which time the enzymes were removed by ultrafiltration (5 kDa cut-off Vivaspin Sartorius, Stonehouse, UK). 20 mg of 2,4-dinitrophenyl hydrazine was added to the Eppendorf and the reaction mixtures were incubated in a heat block at 50 ℃ for 6 hours with intermittent shaking. A TLC plate was used to check formation of desired product (Rf= 0.45 100% EtOAc). The solution was purified via preparatory TLC and the desired hydrazone was mechanically isolated and dissolved in MeOH. Subsequently, the solution was filtered, concentrated and the composition of the purified glyceraldehyde-hydrazones was analyzed on HPLC (3 μL injection, Chiracel® IA-3). Eluents used for HPLC methods, water with 0.1% formic acid (A) and methanol (B) were LC-MS grade (Optima, Fisher). For separation between L-and D-glyceraldehyde-hydrazones an isocratic method using 60% A, 40% B with a flow rate of 0.65 mL/min was used with a 12 minute stop time with UV detection at 360 nm. L-glyceraldehyde-hydrazone eluted at 2.30 min and D-113 glyceraldehyde-hydrazone eluted at 2.76 min. ESI mass spectra were also collected in positive mode scan for m/Z 95 – 500 running at 0.8 s/cycle. drying gas = 5.0 L/min, nebulizer pressure = 50 psi, gas temperature = 300 °C, capillary voltage = 4000 V. 114 Chapter 4: Cross-linked Enzyme Aggregates of a Copper Radical Alcohol Oxidase for Increased Potential in Biocatalysis 4.1 Introduction For centuries, humans have utilized microorganisms for a variety of purposes, such as the production of bread, beer and wine. The enzymatic nature of these processes was discovered and has been harnessed for the benefit of human society. Currently, enzymes are employed in a wide range of industrial processes and the number of applications has grown exponentially over the past decades, to include the pharmaceutical, chemical, textile, detergent, cosmetics, food and waste management fields.260 The use of enzymes in catalysis offers many key advantages, such as inherent regio- and stereo-selectivity, tunability of stability and activity through protein engineering, and allowing for less energy intensive reaction conditions.14, 25, 244 In particular, there is increasing interest in replacing chemical oxidants, many of which produce toxic waste products, with oxidoreductive enzymes.14, 24, 261 Out of all biocatalysis executed industrially, one-sixth are bio-oxidation reactions which include hydroxylation, Bayer-Villiger catalysis, alcohol and amine oxidation, and other transformations.14, 27 Currently, a large portion of these reactions use enzymes requiring complex organic cofactors, such as FAD- and NAD(P), the latter of which necessitates recycling strategies.13 In contrast, copper-radical oxidases (CROs),46, 48, 56, 84 found in the Auxiliary Activity Family 5 in the Carbohydrate-Active Enzymes (CAZy),43 perform two electron oxidations with concomitant reduction of O2 to H2O2 without organic cofactors and have received some attention regarding their potential as industrial biocatalysts.14, 199 Specifically, subfamily 2 (AA5_2) contains the archetypal galactose oxidase from Fusarium graminearum (FgrGalOx, 115 E.C.1.1.3.9),45, 50, 51, 87, 200 as well as the more recently discovered general alcohol oxidases (CgrAlcOx, E.C. 1.1.3.13),46, 47 and aryl alcohol oxidases (CgrAAO, E.C. 1.1.3.7)48 from Colletotrichum graminicola. AA5_2 enzymes oxidize a variety of primary alcohol-containing substrates to the corresponding aldehydes.45, 46, 48, 91 FgrGalOx, the founding member of the AA5_2, and its variants have been investigated for different biotechnological applications, which include glycoprotein labelling,121, 122, 124, 126 development of biosensors,92 chemo-enzymatic modification of polysaccharides for novel materials,128, 130-133, 136 and the industrial manufacturing of pharmaceuticals.144 A significant hurdle to the industrial utilization of CROs is their reliance on accessory enzymes, catalase (EC 1.11.1.6) and horseradish peroxidase (HRP, EC 1.11.1.7), to achieve optimum activity.134 In some cases, potassium ferricyanide and manganese (III) acetate have been shown to function as alternative oxidants to HRP.75, 76 Significant challenges remain that limit the widespread use of biocatalysis in industrial applications, including low substrate loading due to issues with solubility in aqueous mediums, relatively short lifespans of enzymatic catalysts and high costs of enzyme production.26 Enzyme immobilization can increase enzyme stability, catalyst recovery, and enable new applications that were previously not economically viable. For example, two prominent industrial processes using immobilized enzymes that comprise a large portion of the industrial enzyme market are the production of high fructose corn syrup with glucose isomerase and the synthesis of penicillin-based pharmaceuticals with penicillin amidase.146, 147 A variety of immobilization methods exist, such as adsorption, entrapment/encapsulation, cross-linking to a surface, and the preparation of cross-linked enzyme aggregates (CLEA). CLEA are produced by enzyme precipitation followed by cross-linking and have been shown to be effective for a variety of enzyme classes.26, 168, 169 The most commonly used cross-116 linker is glutaraldehyde, which reacts with free amino groups on enzyme surfaces.161, 168, 187 Cross-linking of soluble enzymes was first used in the 1960s but resulted in unstable particles that were difficult to reproduce.26 Cross-linking was then applied to enzyme crystals (CLEC) and showed good stability, high activity, and reusability, however the requirement of crystallization limited broader industrial applicability.158 CLEA showed the same advantages as CLEC and was first used on penicillin acylase in a broad range of organic solvents.159 Immobilization of enzymes using CLEA has been shown to increase enzyme stability (including temperature, pH, or solvent),170-172, 174-176 shelf life,26, 165, 168, 169, 177-179 and reusability.26, 164, 165, 168, 169 Due to the simplicity of the CLEA technique, it can be applied to processes involving enzyme cascades by co-cross-linking multiple enzymes forming “combi-CLEA”, such as the resolution of S-mandelic acid,188, 189 production of α-hydroxycarboxylic amides,190 the production of alkyl ferulates,191 and other processes.192-195 Furthermore, CLEA has been successfully used to independently immobilize a CRO (FgrGalOx),187 catalase,262 and HRP,263 which serves as a starting point for the potential immobilization of other CROs, in combination with accessory enzymes. We wanted to improve the application potential of CgrAlcOx for the synthesis of aryl aldehydes by investigating its immobilization for the oxidation of benzyl alcohol through CLEA. Benzyl alcohol was selected as the model substrate due to the high enzymatic conversion and its ease of product detection by UV. First, we optimized the preparation of CgrAlcOx CLEA and combi-CLEA with accessory enzymes (HRP and catalase) and investigated the conversion yield of benzyl alcohol with these immobilized catalysts. We also explored the utilization of potassium ferricyanide as a replacement for HRP to lower the potential cost of the process. 117 4.2 Results 4.2.1 CgrAlcOx, Catalase and HRP CLEA The CLEA method was used for the immobilization of CgrAlcOx, with glutaraldehyde selected as the cross-linker based on a long historical precedent.26 Experiments varying precipitant type and cross-linker concentration were performed to optimize CLEA activity. We investigated the difference between using CgrAlcOx CLEA alone or with soluble accessory enzymes and combi-CLEA prepared with all three enzymes co-immobilized (CgrAlcOx, catalase, and HRP). 4.2.1.1 Screening CLEA Preparation Conditions A variety of precipitants used in CLEA formation can have a detrimental impact on enzyme activity, thus a precipitant screen was conducted. A panel of organic solvents or saturated aqueous ammonium sulfate were incubated in volumetric excess with an aliquot of purified CgrAlcOx. Subsequently, the enzyme was resuspended with buffer and the standard ABTS-HRP coupled assay was used to determine the residual activity of the solubilized enzymes. CgrAlcOx showed no loss in enzyme activity after the initial aggregation step with 1-propanol, 2-propanol, tert-butanol and saturated ammonium sulfate (Table C1). Saturated ammonium sulfate, a well-known precipitant used in enzyme purification,264 and 1-propanol/2-propanol were not investigated further since a novel kinetic lag-phase was observed with the former, while the later alcohols are also substrates for CgrAlcOx.46 tert-butanol did not diminish the activity of CgrAlcOx after precipitation and is not a substrate of the alcohol oxidase, therefore it was chosen as the precipitant for subsequent CLEA optimization. Notably, this precipitant was also selected previously for the production of CLEA of the related copper radical oxidase, FgrGalOx.187 118 CgrAlcOx has 23 surface lysines potentially available for glutaraldehyde cross-linking.46 A glutaraldehyde inhibition study was first conducted with CgrAlcOx in solution and no loss of specific activity in the presence of 1-100 mM glutaraldehyde was observed (Figure C1). 4 mg/mL of CgrAlcOx was chosen for all CLEA experiments since at this enzyme concentration the presence of CLEA particles was visually confirmed. Based on previous research establishing the requirement of accessory enzymes (HRP and catalase) for optimal CRO activity,74, 76, 221 we surveyed two types of preparation: one composed of CgrAlcOx CLEA with HRP and catalase added in solution and the other with all three enzymes co-immobilized together to form combi-CLEA. Experiments to optimize cross-linker concentration were conducted and the best percent conversion (100%) of 1 mM benzyl alcohol was observed with CLEA prepared with 1 mM glutaraldehyde (Figure 4.1). Therefore, all further preparations of CgrAlcOx CLEA and combi-CLEA were performed with 1 mM glutaraldehyde as the optimum cross-linker concentration. As expected, shaking was required to obtain maximum conversion (Figure C2), thus all subsequent samples underwent shaking at 250 rpm. Previous research on CLEC has demonstrated lyophilization can be used as a method for stabilization without impacting reactivity.265 In our case, lyophilized CLEA showed a lower benzaldehyde production than non-lyophilized (Figure C3). 119 Figure 4.1. Effect of cross-linker concentration on CLEA/combi-CLEA activity CgrAlcOx CLEAs made of 0.8 nmol CgrAlcOx plus 3.3 nmol HRP and 25 nmol of soluble catalase (black square) or combi-CLEAs made of 0.8 nmol CgrAlcOx cross-linked with 3.3 nmol HRP and 25 nmol catalase (red circle) prepared with varying concentrations of glutaraldehyde cross-linker were incubated with 1 mM benzyl alcohol for 30 mins. 4.2.1.2 Comparison of CLEA and combi-CLEA Activity With optimized CLEA preparation conditions in-hand, we then assayed the percent conversion of CgrAlcOx CLEA alone, CgrAlcOx CLEA with soluble accessory enzymes and combi-CLEA prepared with all three-enzymes co-immobilized. In our initial screens, we noted that CgrAlcOx CLEA alone in the reaction, i.e. in the absence of catalase and HRP, only reached 45% conversion of 1 mM benzyl alcohol to benzaldehyde (Figure C3). On the other hand, CgrAlcOx CLEA to which these accessory enzymes were added in solution, as well as the combi-CLEA comprised of CgrAlcOx, HRP, and catalase, achieved quantitative conversion of 1 mM benzyl alcohol at three hours (100% and 95 ± 5% conversion, respectively) (Figure C2). These 120 data confirm that the addition of HRP and catalase is essential to achieve and maintain optimum activity of immobilized CgrAlcOx, as has been observed for this and other CROs in solution.74, 76, 221 With a goal of simplifying reaction conditions, we explored the effect of omitting buffer components and carrying-out substrate conversions in ultrapure water. To achieve this, 50 mM sodium phosphate buffer, pH 7.0, was selected as a compromise of the pH optimums of the individual enzymes.46, 266, 267 All CLEA reactions performed without buffer had lower conversion yields (~ 65% at 1 h) than the corresponding reactions performed in buffer (~ 100% at 1 h). Furthermore, all reactions with CgrAlcOx CLEA and soluble accessory enzymes (HRP and catalase) showed lower percent conversions at all time points than the corresponding reactions with combi-CLEA (Figure 4.2). However, the control reaction performed in water, where all enzymes are soluble, achieved 97% conversion in 10 min (Figure 4.2), indicating that immobilization of CgrAlcOx caused a decrease in the biotransformation velocity. 121 Figure 4.2. Comparison of percent conversion of CgrAlcOx CLEA with soluble accessory enzymes, combi-CLEA and soluble enzymes for benzyl alcohol oxidation CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase and 3.3 nmol HRP (black and grey) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 3.3 nmol HRP and 25 nmol catalase (dark and light red) were incubated 1 mM benzyl alcohol in either water (black square, dark red triangle) or 50 mM sodium phosphate buffer, pH 7.0 (grey circle and light red rhombus) for 10 min, 30 min and 60 min. A control reaction performed in water with only soluble enzymes was also assessed (blue pentagon). Benzaldehyde production was quantified by HPLC. To further simplify the reaction set-up, we wanted to explore the effect of immobilizing two-enzymes into combi-CLEA with the third enzyme in solution, while also assessing buffer dependence (Figure C4). Our control reaction containing only HRP-catalase combi-CLEA without soluble CgrAlcOx showed no conversion (Figure C4K-L). Combi-CLEA consisting of CgrAlcOx-catalase with soluble HRP showed higher percent conversion in water than in buffered 122 conditions (Figure C4A-B), while combi-CLEA made of CgrAlcOx-HRP with soluble catalase showed the opposite effect, i.e. higher percent conversion observed in buffered conditions (Figure C4E-F). Over 95% conversion was observed with combi-CLEA made of the accessory enzymes with soluble CgrAlcOx (Figure C4I-J). The three-enzyme combi-CLEA reacted in buffered conditions (Figure C4M-P) showed comparable conversions to other CLEA conditions in which CgrAlcOx was immobilized. However, the highest percent conversion in which CgrAlcOx was immobilized was observed for CgrAlcOx-catalase combi-CLEA with soluble HRP in benzyl alcohol without buffer (Figure C4A). This indicates that CLEA composed of two enzymes is potentially beneficial to achieve high conversion. One advantage of enzyme immobilization is the potential for increased reusability, as CLEA of a variety of enzymes have shown comparable activity over 3 – 30 cycles.268-270 The stability and reusability of CgrAlcOx CLEA and combi-CLEA were assessed with two consecutive one-hour cycles of 1 mM benzyl alcohol oxidation. Compared to the first cycle, the percent conversions observed in the second cycle were lower, however, during the second cycle, CLEA samples (dashed columns) showed on average higher percent conversions than the controls (Figure C5). This indicates that some mechanical stability has been imparted to the immobilized aggregates. Unfortunately, the highest observed conversion after the second cycle was only at 55%, which is a decrease in reusability compared to other systems in the literature.26, 164, 165 Biocatalytic processes seek to minimize the cost of downstream product recovery, and consequently, high product concentrations are beneficial.271 Therefore, higher substrate loadings were tested to determine the maximum concentration converted after 10 mins using immobilized CgrAlcOx. The percent conversion of CgrAlcOx CLEA with soluble catalase and HRP was lower than for the uncross-linked controls for reactions with both 10 mM and 30 mM benzyl alcohol 123 (Figure C6), which is in accordance with our results with 1 mM benzyl alcohol (Figure 4.2). However, combi-CLEA displayed comparable percent conversion to the controls with 10 mM benzyl alcohol, whereas with 30 mM benzyl alcohol the combi-CLEA have slightly lower conversions than the controls (Figure C6). This shows that the three-enzyme system CLEA (CgrAlcOx, HRP, and catalase) may not be the most effective immobilization technique for the oxidation of higher than 1 mM benzyl alcohol. However, our previous results with CgrAlcOx-catalase combi-CLEA with soluble HRP (Figure C4A) indicated that higher percent conversion may potentially be observed with just a two-enzyme combi-CLEA composed of CgrAlcOx and catalase with a soluble single electron oxidant. 4.2.2 Replacement of HRP with Potassium Ferricyanide HRP rescues CgrAlcOx from a catalytically inactive state by re-establishing the lost radical on the cysteine-tyrosine residue of CROs.45, 62, 68, 76 However, HRP is an expensive enzyme that is difficult to produce recombinantly, hence finding a small molecule to replace it would be desirable. Potassium ferricyanide (K3Fe(CN)6) has been previously identified to increase the activity of CROs74, 210 and it has been shown to perform the same role as HRP.76, 272 We assessed the pH dependence of immobilized CgrAlcOx with both soluble and co-immobilized catalase and varying concentration of K3Fe(CN)6 and found the best percent conversions were seen at pH 7.0 (Figures C7 – C8), which is concordant to the results previously obtained for homogenous catalysis with CgrAlcOx.76 In addition, reactions with higher concentrations of K3Fe(CN)6 showed slightly higher conversions (Figures C7 – C8). No difference was observed between reactions performed with CgrAlcOx-catalase CLEA versus CgrAlcOx CLEA with soluble catalase for 1 mM benzyl alcohol (Figure C7). However, CgrAlcOx-catalase CLEA yielded higher percent conversion than CgrAlcOx CLEA with soluble catalase when reacted with 10 mM benzyl alcohol (Figure C8). 124 The best conditions showed 70 – 80% conversion of 1 mM benzyl alcohol and 27% conversion of 10 mM benzyl alcohol in 30 min (Figures C7-C8), which is higher than what was observed for the three-enzyme system (vide supra). Subsequently, we monitored the reaction of 1 mM benzyl alcohol with 0.1 mM, 1 mM or 10 mM K3Fe(CN)6 for 1 and 2 hours in either water or buffered conditions (pH 7.0) and observed almost full conversion with 1 mM and 10 mM K3Fe(CN)6 with buffered combi-CLEA and 10 mM K3Fe(CN)6 with CgrAlcOx CLEA with soluble catalase without buffer (Figure 4.3). Incomplete conversion using 0.1 mM K3Fe(CN)6 was observed after 1 and 2 hours while all other conditions reached 100% conversion at 2 hours (Figure 4.3). Full oxidation of 1 mM benzyl alcohol can be achieved utilizing CLEA particles composed of CgrAlcOx and catalase with soluble potassium ferricyanide successfully replacing HRP. 125 Figure 4.3. Influence of ferricyanide concentration on CLEA and combi-CLEA percent conversions CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase (solid bars) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 25 nmol catalase (dashed bars), were incubated with 1 mM benzyl alcohol in water (red) or 50 mM sodium phosphate buffer, pH 7.0 (blue) and either 0.1 mM (pink and light blue), 1 mM (red and blue) or 10 mM (dark red and dark blue) K3Fe(CN)6 was present. Benzaldehyde production was quantified by HPLC after 1 or 2 hours. To overcome the limitation seen in the HRP system with higher substrate loadings, we investigated the potential of replacing HRP with ferricyanide by performing the reaction with 10 mM benzyl alcohols. After 1 hour, all trials reached between 40 – 50% maximum conversion before reactivity stalled (Figure 4.4). This plateau remained over extended reaction time and increasing the concentration of K3Fe(CN)6 did not increase the maximum conversion (Figure 4.4). 126 Doubling the initial concentration of CgrAlcOx in the CLEA particles did not affect the percent conversion after 4 hours, indicating that the stalling of the reaction is not dependent on the initial amount of CgrAlcOx (Figure C9). Furthermore, similar reactions stalling was observed when incubating either CgrAlcOx CLEA or combi-CLEA with 30 mM benzyl alcohol and varying K3Fe(CN)6 concentrations, which showed a comparable percent conversion at 30 min (9 ±3%) or 17 hours (15 ±3%) (Figure C10). Together, these results indicate that incubation time or initial CgrAlcOx concentration are not the reason for incomplete conversion of 10 mM and 30 mM benzyl alcohol. Figure 4.4. Time dependent conversion of 10 mM benzyl alcohol by CgrAlcOx CLEA and combi-CLEA A. Reaction performed in water. B. Reaction performed 50 mM sodium phosphate buffer, pH 7.0. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase (solid lines) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 25 nmol catalase (dashed lines), were incubated with 10 mM benzyl alcohol and either 10 mM (squares) or 100 mM (triangles) K3Fe(CN)6. For each time point benzaldehyde production was quantified by HPLC To investigate the possible reasons for incomplete conversion, we monitored by HPLC reactions containing equimolar concentration of benzyl alcohol and benzaldehyde with soluble CgrAlcOx, catalase, and K3Fe(CN)6. No product inhibition was observed (Figure C11), 127 concordant to previous work with CgrAlcOx in the presence of HRP and catalase76. A steady increase in conversion was observed over time, reaching 82 – 86% conversion of 10 mM benzyl alcohol after 15 hours (Figure C12). Furthermore, CLEA reactions (Figure 4.4) reached higher conversions with 10 mM benzyl alcohol at equivalent time points before 2 hours compared to non-CLEA reactions (Figure C12). These results indicated that the reaction rate may be increased with our immobilized particles and the conversion plateau is specific to CLEA. We subsequently investigated other factors contributing to reaction stalling such as enzyme deactivation (CgrAlcOx and/or catalase), depletion of K3Fe(CN)6 or oxygen limitation. First, conversion yields were measured using CLEA or combi-CLEA particles with 10 mM benzyl alcohol and 10 mM K3Fe(CN)6 in buffered conditions for 3 hours. Subsequently, either soluble CgrAlcOx, soluble catalase, K3Fe(CN)6, hydrogen peroxide or one CgrAlcOx CLEA particle were added into the reaction and percent conversions were determined 30 minutes after addition (Figure 4.5). The conversion yields increased for each additive, and the greatest changes occurred after the addition of hydrogen peroxide or soluble CgrAlcOx (Figure 4.5). H2O2 was used to deliver oxygen into the reaction and the observed increased conversion indicated that both enzymes (CgrAlcOx and catalase) were still active and that enough potassium ferricyanide remained for the reaction to continue. The increased conversion after addition of CgrAlcOx and CgrAlcOx CLEA is expected since this enzyme is the main catalyst for oxidation. Furthermore, the increased conversion observed after the introduction of other additives could be due to air exposure of the reaction mixture. 128 Figure 4.5. Effect of the addition of a new component on the percent conversion of 10 mM benzyl alcohol by CgrAlcOx CLEA CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase (solid bars) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 25 nmol catalase (dashed bars) were incubated with 10 mM benzyl alcohol and 10 mM K3Fe(CN)6 in 50 mM sodium phosphate buffer pH 7.0 for 3 hours, benzaldehyde production was quantified by HPLC (black bars). Next, either 0.8 nmol CgrAlcOx, 25 nmol catalase, 1 nmol K3Fe(CN)6, 16 nmol hydrogen peroxide or one particle of CgrAlcOx CLEA (0.8 nmol CgrAlcOx) were added to the reaction and 30 min after addition the benzaldehyde production was quantified again by HPLC (blue bars). 4.3 Discussion Capitalizing on the interest to replace chemical oxidants with oxidoreductases,244, 261 we wanted to investigate the biocatalytic potential of CgrAlcOx,46 a competent general alcohol oxidase, in the context of the pharmaceutical and fine chemicals industries. Biocatalysis still faces challenges for industrial applicability and enzyme immobilization can increase the stability and reusability of biocatalysts, thereby potentially lowering costs.26 We chose the cross-linked 129 enzyme aggregate (CLEA) strategy as a facile method to test if CgrAlcOx was amenable to enzyme immobilization. To achieve the optimum activity of CROs, accessory enzymes are required: catalase for the conversion of hydrogen peroxide, generated by the CRO during its catalytic cycle, into the co-substrate oxygen and water; and horseradish peroxidase (HRP) for rescuing the off-cycle, fully reduced form of CgrAlcOx by a single electron oxidation.72, 74, 76 In addition, potassium ferricyanide has been shown to function as a replacement of HRP in CRO mediated catalysis.74, 76 Low conversions were observed with only CgrAlcOx CLEA, confirming accessory enzymes are required even with an immobilized CRO (Figure C3). Hence all other experiments focused on either the immobilization of CgrAlcOx with soluble catalase and either HRP or K3Fe(CN)6 versus combi-CLEA containing multiple cross-linked enzymes. It is well established that ions and ionic strength influence the activity and stability of soluble enzymes,273 and thus we explored if the incorporation of CgrAlcOx into CLEA could alleviate the requirement of buffer. For reactions with 1 mM benzyl alcohol, higher conversions were observed in buffered samples for all types of CLEA with either HRP or K3Fe(CN)6 (Figures 4.2, 4.3, and C7). However, with increased substrate concentration, the importance of buffering the solution diminished (Figures 4.4 and C8-C10). The CLEA strategy may be a solution to minimize the cost of buffering, which would be significant on an industrial scale, since our results show that with increased substrate concentration there is less of a requirement to use buffer. For trials performed with HRP as the co-oxidant, combi-CLEA reactions showed higher conversions than CgrAlcOx CLEA with soluble enzymes, indicating a benefit of cross-linking all three enzymes (Figure 4.2). Since it has been proposed that HRP directly interacts with CgrAlcOx 130 in order to restore the active site radical,76 the requirement of physical proximity is being accommodated in the combi-CLEA since an enhancement in conversion is observed. However, when we individually immobilized CgrAlcOx with either catalase or HRP and supplemented the third enzyme in solution, we observed that the CgrAlcOx-catalase CLEA in water (Figure C4A) and CgrAlcOx-HRP CLEA in buffer (Figure C4F) showed higher percent conversion than other trials (Figure C4M-P). This indicates that while physical proximity of the enzyme in combi-CLEA is beneficial, the three-enzyme CLEA is ultimately not the best CgrAlcOx immobilization option for benzyl alcohol, which was further confirmed with 10 mM and 30 mM benzyl alcohol (Figure C6). The three-enzyme combi-CLEA formed visibly bigger aggregates than the CLEA made with only CgrAlcOx, possibly contributing to increased mass-transfer limitations. Furthermore, since HRP only possesses six lysine residues,274 its capacity to form stable CLEA could be limited. Substituting HRP, an expensive enzyme, for a relatively inexpensive small molecule would be beneficial for increasing the viability of this system by reducing the complexity of the cross-linked system. Since our previous results with CgrAlcOx-catalase CLEA with soluble HRP (Figure C4A) showed high percent conversion, we moved our attention to the two-enzyme CgrAlcOx-catalase CLEA with a soluble single electron oxidant. In addition, two small molecules (potassium ferricyanide and manganese (III) acetate) have been used successfully as HRP replacements.75, 76 Potassium ferricyanide was able to replace HRP within our CLEA system and displayed higher conversion for trials with K3Fe(CN)6 present than compared to the three-enzyme system for 1 mM benzyl alcohol (Figures 4.2 and C7). A similar observation can be made for experiments with 10 mM benzyl alcohol (Figures C6 versus C8), indicating the two-enzyme CLEA with potassium ferricyanide is a superior option for benzyl alcohol oxidation. Furthermore, 131 in all trials with K3Fe(CN)6, combi-CLEA yielded higher percent conversion than CgrAlcOx CLEA with soluble catalase, which is also a trend observed with the three-enzyme combi-CLEA However, with increased substrate concentration, the percent conversion with CgrAlcOx, catalase, and K3Fe(CN)6 plateaued at 50% after 2 hours (Figure 4.4). We added all reaction components individually after three hours and observed an increase in conversion for all conditions (Figure 4.5). The greatest observed increase in conversion was when hydrogen peroxide was added to deliver oxygen into the solution through the activity of catalase. Oxygen is required for the second half-reaction of the ping pong mechanism of CROs45, 62, 67, 68 and since oxygen has poor solubility in aqueous solutions (0.27 mM),275 it is conceivable that in the closed CLEA reactions all of the oxygen present at the start of the reaction was depleted. A simple solution to increase the concentration of oxygen would be to bubble air into the reaction mixture. While CgrAlcOx has been previously shown to be deactivated when exposed to aeration,76 the utilization of CgrAlcOx CLEA may provide extra stability. Other solutions such as controlled addition of hydrogen peroxide into the vessel or attempting other methods of immobilization adapted to flow chemistry could also be explored to overcome the reaction stalling. The second highest increase was observed with the addition of soluble CgrAlcOx, which was expected since we observed high percent conversion with soluble CgrAlcOx and combi-CLEA made of accessory enzymes (Figure C4I-J). The increase in percent conversion following the addition of extra catalase or K3Fe(CN)6 could be attributed to the additives themselves or could be due to oxygen entering the reaction for the short time the vessels were open. It has been previously observed that even small changes in the oxygen content can have large effects on the reaction rate of CROs.74 Overall, oxygen diffusion may be the limiting factor responsible for the reaction stalling observed with CgrAlcOx CLEA particles. 132 Lastly, one of the main advantages of enzyme immobilization is the ability to reuse the biocatalysts, thereby lowering the initial production cost of the application.26 We observed 55% conversion after one re-use cycle with different types of CgrAlcOx CLEA (Figure C5). Although previous work with laccase CLEA did not show appreciable decrease in activity after the first round of enzyme recycling, subsequent rounds did have decreased conversion and it was speculated the cause was mechanical loss of the CLEA particles.276 A similar effect could be occurring in our system and could potentially be overcome with higher cross-linker concentrations to make more mechanically robust CLEA particles. Furthermore, more optimization needs to be conducted to improve the currently observed limited reusability of CgrAlcOx CLEA including changing the proportion of enzymes and substituting HRP with K3Fe(CN)6. 4.4 Conclusions Few oxidoreductases have been immobilized into CLEA particles, with most work focused on laccases and glucose oxidase.195, 277, 278 A method for the preparation of cross-linked enzyme aggregates (CLEA) has been established for CgrAlcOx and optimized for precipitant type and cross-linker concentration. Full conversion of benzyl alcohol was achieved with combi-CLEA containing CgrAlcOx and accessory enzymes (HRP and catalase). Furthermore, potassium ferricyanide was successfully used as a replacement for HRP resulting in enhanced conversion with the CgrAlcOx-catalase CLEA. The dependence on buffer diminished as substrate loadings increased, however, the conversion plateaued at 50% possibly due to the lack of oxygen in the sealed reaction. Our work serves as a starting platform for the CLEA immobilization of CgrAlcOx and other CROs for biocatalytic applications. 133 4.5 Materials and Methods 4.5.1 Chemicals and Enzymes Distilled water was used for the preparation of all buffers and stock solutions unless stated otherwise. CgrAlcOx was expressed in Pichia pastoris and purified using Ni-NTA chromatography as previously reported.46 Catalase from bovine liver (2000–5000 units per mg protein, Sigma) and horseradish peroxidase (Rz > 3, 300 units per mg, Bio Basic Canada Inc.), obtained as lyophilized powders, were used as received. Benzyl alcohol (98%) and a 25% solution of glutaraldehyde in water was purchased from Sigma-Aldrich. Potassium ferricyanide was purchased from Amresco (ASC grade). Other substrates and reagents were purchased from commercial sources (Sigma-Aldrich, VWR or Fisher) and used without further purification. 4.5.2 HPLC Method The eluents used for the HPLC method were water with 0.1% formic acid (A) and acetonitrile (B) from Optima, Fisher (LS-MS grade). An Agilent 1200 HPLC system equipped with Poroshell 120 EC-C18 (2.2 × 50 mm, 2.7 μm) column and a Poroshell HPH-C18 (2.1 × 5 mm, 2.7 μm) guard column was used for the percent conversion analysis. For benzyl alcohol analysis, the HPLC method was initial 12.5% B with ramping to 62.5% B over 2.00 minutes, followed by an isocratic step of 100% B for 1.25 min. The flow rate was 0.65 mL/min. Benzyl alcohol was monitored at 210 nm and eluted at 0.6 min while benzaldehyde was monitored at 250 nm and eluted at 1.1 min. Substrate and product peak areas were converted to concentration using a standard curve comprised of triplicate measurements of benzyl alcohol and benzaldehyde at concentration of 0.1, 0.5, 1, 3, 5, and 7 mM. The calibration curves were constructed using the wavelength close to the absorption maximum of the analytes for maximum sensitivity (λ 210 for 134 benzyl alcohol and λ 250 for benzaldehyde). Either 0.3 μL, 0.5 μL, 0.7 μL, 1 μL, 3 μL or 5 μL were injected depending on concentration of the initial solution in order to be within the values of the calibration curve. 4.5.3 CRO Activity Assay The standard HRP – 2,2’-azinobis(3-ethylbenzthiazoline-6-sulfonic acid) (ABTS)-coupled assay209 with 3 mM of benzyl alcohol, 0.46 mM ABTS and 30 U/mL of HRP at 25 oC was performed to assess the activity of CgrAlcOx. The oxidation of the alcohol group on the substrates by AA5 enzymes consumes 1 equivalent of O2 and produces 1 equivalent of H2O2. The oxidation of ABTS: (λmax = 420 nm, ε = 36000 M-1cm-1) is catalyzed by the enzyme HRP using 2 equivalents of H2O2. 4.5.4 Precipitant and Glutaraldehyde Inhibition Assay 90 μL of precipitant was added to 10 μL of CgrAlcOx at 0.5 mg/mL and after 15 minutes 900 μL of buffer (50 mM sodium phosphate, pH 8.0) was added. The standard CRO activity assay was performed with 10 μL aliquot of the resuspended CgrAlcOx. The standard HRP-ABTS coupled assay was performed using 70 μg of CgrAlcOx preincubated with 0 mM, 1 mM, 25 mM, 75 mM and 100 mM glutaraldehyde in 50 mM sodium phosphate buffer, pH 8.0. 4.5.5 CLEA and combi-CLEA Preparation A general protocol was used to prepare different types of CLEA particles that included different combinations of 4 mg mL-1 CgrAlcOx, 332 μM HRP and/or 2480 μM catalase and different volumes of tert-butanol precipitant and glutaraldehyde cross-linker (Table 4.1). The enzyme(s) were distributed into a 1 mL or 2 mL Eppendorf tube. Subsequently, tert-butanol was added followed by varying concentrations of glutaraldehyde solution to obtain the desired final concentration. The suspension was mixed twice using a pipette and then samples were incubated 135 at RT for 30 min without shaking. Afterwards, samples were centrifuged at 10 000 g for 10 min at 4 ̊C, the liquid decanted and the pellet was washed with water. A second centrifugation was performed, solution decanted again and any excess visible liquid was removed. Some samples were frozen in liquid nitrogen and lyophilized overnight while others were immediately assayed using the CLEA activity assay described below. The two-enzyme CLEA had 10 μL of either soluble 1) 2480 μM catalase, 2) 332 μM HRP or 3) 4 mg mL-1 CgrAlcOx added so all three enzymes were present at the start of the incubation. A control reaction with only the two-enzyme combi-CLEA particles and no soluble enzyme were performed for all conditions. Table 4.1. Experimental conditions for CLEA preparation CLEA Type CgrAlcOx Volume (μL) catalase Volume (μL) HRP Volume (μL) Precipitant Volume (μL) Cross-linker Volume (μL) CgrAlcOx Only 10 / / 90 10 CgrAlcOx, catalase and HRP 10 10 10 270 30 CgrAlcOx and catalase 10 10 / 180 20 CgrAlcOx and HRP 10 / 10 180 20 Catalase and HRP / 10 10 180 20 4.5.6 CLEA Activity Assay To assay CLEA activity 1 mM, 10 mM or 30 mM benzyl alcohol diluted in water or 50 mM sodium phosphate buffer (pH 6.0, 7.0 or 8.0) was incubated with the CLEA particle for specific time intervals. To test the influence of agitation, some samples were left standing upright at RT while the remaining samples were placed horizontally on an orbital shaker (LAB-LINE instruments, Melrose Park, IL) set to 250 rpm. For trials with the three-enzyme system: either CgrAlcOx CLEA was mixed with 10 μL of 2480 μM catalase and 10 μL of 333 μM HRP in solution at the start of the incubation or combi-CLEA consisting of CgrAlcOx, catalase and HRP 136 were used. For trials substituting HRP with potassium ferricyanide: either CgrAlcOx CLEA was mixed with 10 μL of 2480 μM catalase and K3Fe(CN)6 or combi-CLEA made of CgrAlcOx-catalase was prepared and mixed with K3Fe(CN)6. Different concentrations of K3Fe(CN)6 were used (0.1 mM, 1 mM, 10 mM, 30 mM, 100 mM or 300 mM) and all samples were vortexed to obtain a homogenous solution with benzyl alcohol prior to incubation. For combi-CLEA particles consisting of CgrAlcOx and HRP, 10 μL of 2480 μM catalase was added at the start of the reaction and for combi-CLEA particles consisting of catalase and HRP, 10 μL of 4 mg mL-1 CgrAlcOx was added at the beginning of the incubation. At specified time points, samples for experiments with CgrAlcOx CLEA alone (Figure C3), determining optimum cross-linker concentrations (Figure 4.1) and to confirm the benefit of reaction shaking (Figure C2) were centrifuged at 10 000 g for 10 min at RT to stop the reaction. All other experiments were quenched with MeOH (99.8% purity). The proportion of reaction to MeOH was either 1:1, 1:2, 1:4 or 1:6 depending on the initial concentration of benzyl alcohol to allow the measurement to be within the linear range of the detector. Subsequently, the solutions were loaded onto 0.2 μm cellulose acetate spin filters (VWR, P/N: 2994-752) and centrifuged again. The filtrates were analyzed by HPLC. For most trials, control reactions with no cross-linking reagent were performed. One reaction with combi-CLEA in the presence of 1 mL of 1 mM benzyl alcohol mixed with 50% methanol was performed. To assess the stalled reactions, CLEA was incubated for 3 hours and then a 50 μL aliquot was quenched with MeOH and either 10 μL of 4 mg mL-1 CgrAlcOx, 10 μL of 2480 μM catalase, 10 μL of 100 mM K3Fe(CN)6, 10 μL of 1.6 M hydrogen peroxide or a new CgrAlcOx CLEA particle was added to the tubes. The reaction continued with shaking at 250 rpm and a 100 μL aliquot was quenched and analyzed by HPLC after 30 mins. 137 4.5.7 Reusability CLEA was incubated with 1 mL of 1 mM benzyl alcohol prepared in either water or 50 mM sodium phosphate buffer, pH 7.0 in 2 mL Eppendorf tubes and shaken at 250 rpm. Reactions were quenched after 1 hour by adding 300 μL of the reaction to 300 μL of methanol (99.8% purity), loaded onto a 0.2 μm cellulose acetate spin filters (VWR, P/N: 2994-752), and centrifuged at 10 000 g for 10 min at 4 C̊. The remaining reaction liquid was then decanted and a fresh solution of 1 mM benzyl alcohol prepared in either water or 50 mM sodium phosphate buffer, pH 7.0 was added and incubated at 250 rpm for another 60 min. The reaction was then quenched as previous described and the filtrate was analyzed by HPLC. The CLEA reactions were performed in duplicate. Control reactions with no cross-linking reagent were performed in parallel with the experimental reactions with either the precipitation of only CgrAlcOx or with the precipitation of all three enzymes. 4.5.8 Non-CLEA Reaction Monitoring The assay consisted of 5 mM or 10 mM benzyl alcohol, 0 mM or 5 mM benzaldehyde, 0.8 nmol CgrAlcOx, 25 nmol catalase, and K3Fe(CN)6 (5 mM, 10 mM, 50 mM or 100 mM) in either water or 50 mM sodium phosphate buffer, pH 7.0 (final volume 1 mL). All reaction components were mixed prior to addition of CgrAlcOx. Initiation time (t=0) was when CgrAlcOx was added. Reactions were analyzed by HPLC. 138 Chapter 5: General Conclusions and Future Directions 5.1 General Conclusions With the existential threat of climate change looming over society, a shift away from fossil fuel derived energy, chemicals and materials is necessary to disrupt the current trajectory towards catastrophe. The utilization of biomass and the establishment of biorefineries are key strategies to mitigate our greenhouse gas emissions by using renewable carbon for energy and fine chemicals production. Canada is advantageously placed to become a world leader in the circular bio-economy by possessing 9% of the world’s forest and having a well-established forestry industry.1 Biocatalysis can aide the valorization of the complex plant biomass by employing enzymes with inherent selectivity, lower toxicity, and possible cost-effectiveness through less energy intensive processes and catalyst recycling systems.26 Copper radical oxidases, specifically, can serve as catalysts for primary alcohol oxidation of carbohydrates and non-carbohydrates without the requirement of organic cofactors. Within copper radical oxidases characterized members of subfamily two (AA5_2), distinct and diverse enzymatic activities have been reported: galactose oxidase (FgrGalOx),45 aliphatic/aromatic alcohol oxidase (CgrAlcOx)46 and hydroxymethylfurfural oxidase (CgrAAO).48 The work described in this thesis focuses on the discovery and characterization of new catalysts within AA5_2 CROs, and subsequent exploration of enzyme immobilization to further their utilization in biocatalysis. In chapter 2 we described the biochemical characterization and detailed product analysis of two Fusarium AA5_2 enzymes we subsequently classified as aryl alcohol oxidases. FgrAAO and FoxAAO are the first reported aryl alcohol oxidases in this genus and further solidify the diverse nature of activities found in the AA5_2 subfamily. We determined that both enzymes can 139 oxidize hydroxymethyl furfural (HMF) directly to 5-formyl-2-furoic acid (FFCA) and also identified the oxidation sites of methyl fructosides, anhydro sugars and raffinose by FgrAAO to fully explore their biocatalytic potential for biomass functionalization. In chapter 3 we identified 40 target sequences across the AA5_2 subfamily through sequence similarity networks and were able to recombinantly produce and characterize twelve novel enzymes: four galactose oxidases (ExeGalOx, MreGalOx, FoxGalOxB and PfeGalOx), two raffinose oxidases (UmaRafOx and PhuRafOx), four broad spectrum alcohol oxidases (AflAlcOx, PruAA5_2A (PruAlcOx), FoxAlcOx and PorAlcOx), and two non-carbohydrate alcohol oxidases (GciAlcOx and AsyAlcOx). Of note, GciAlcOx had the highest catalytic efficiency on HMF currently reported and was able to partially oxidize it directly to FDCA, an important monomer for the synthesis of bio-plastics. Structure-function trends were determined through detailed analysis of the active-site residues combined with computational docking of galactose into structural homology modeling based on FgrGalOx. Based on FgrGalOx residue numbering, we found the presence of R330 being critical for low KM values while Y329 is not. Moreover, the presence of F194 allows efficient turnover while W290 allows prominent galactose oxidase activity but is not its sole determinant. Chapter 4 built on previous work to increase the industrial applicability of CROs74, 76 through utilizing the CLEA immobilization method on a general alcohol oxidase (CgrAlcOx). We were able to show that CgrAlcOx could be successfully incorporated into CLEA particles, however poor percent conversion of benzyl alcohol to benzaldehyde was observed using this enzyme alone. Thanks to the addition of accessory enzymes, (HRP and catalase) in either soluble form or combi-CLEAs, 100% conversion of 1 mM benzyl alcohol was achieved in 1 hour. Unfortunately, higher substrate loadings resulted in lower conversion yields with the three-enzyme 140 system. As a result, we investigated potassium ferricyanide (K3Fe(CN)6) as a potential HRP replacement and observed enhanced conversion with the CgrAlcOx-catalase combi-CLEA using 1 mM benzyl alcohol. However, the reaction stalled at 50% conversion for 10 mM benzyl alcohol even after 8 hours of incubation. Different causes for the plateau in conversion were explored and the hypothesis of oxygen limitation was proposed. Collectively, the work described in this thesis further enhances the knowledge of CROs, including their diverse activities, bio-product potential and applicability. Furthermore, this work doubled the number of characterized AA5_2 members while showcasing that their original classification as galactose oxidases misrepresents the broader potential of this enzyme subfamily. Indeed, we have discovered enzymes with broad substrate scopes consisting of both carbohydrates and non-carbohydrates, which could be useful in multiple possible biocatalytic applications such the functionalization of biomass for biosensors or the production of bio-plastics from HMF. We have also demonstrated the increased biocatalytic potential of CROs by immobilizing CgrAlcOx through cross-linking and successfully replaced the accessory enzyme HRP with K3Fe(CN)6 without a loss in product conversion. 5.2 Future Directions Despite my work, there are still unanswered question regarding CROs. The work presented in this thesis solely focused on fungal CROs. However, within the CAZy database,43, 237 sequences putatively coding for CROs are also found in bacteria and plant species. Bacterial sequences belong mostly to one genus (Burkholderia) and are almost exclusively annotated as AA5_2. On the other hand, sequences found within plant genomes have not yet been placed in either subfamily (AA5_1 or AA5_2), and come from a variety of organisms such as pineapple (Ananas comosus var. bracteatus), peanut (Arachis hypogaea), wild cabbage (Brassica oleracea), wild lettuce 141 (Lactuca saligna), Japanese rice (Oryza sativa subsp. japonica), almond (Prunus dulcis), and others. Although no recombinantly produced CRO from plants has been characterized, a putative galactose oxidase from Arabidopsis thaliana (RUBY) was recently demonstrated to play a role in forming cross-links between seed-coat mucilage components, thereby promoting the cell-to-cell adhesion of the seed coat epidermis.118 Since CROs are oxidases, it would be of interest to investigate the in vitro and in vivo activity of AA5 enzymes in plants and to identify how these enzymes impact their growth and development. Interestingly, based on a preliminary sequence comparison of RUBY with the canonical FgrGalOx, some key amino acid residues appear to be missing in the plant CRO, illustrating a potentially different active site architecture. Chapter 4 of this thesis focused on improving the applicability of CROs, specifically CgrAlcOx, in biocatalytic processes through immobilization into CLEA. However, more work should be done to assess the stability of these oxidative CLEA over time when exposed to vigorous bubbling of oxygen, and to explore the potential of this immobilization strategy for the oxidation of alkyl alcohols for which incomplete conversion was observed.76 Furthermore, limited recyclability was observed with the different CgrAlcOx CLEA systems, likely due to mechanical instability. Additional work could study the role increased cross-linker concentration has on the recyclability of CLEA. Moreover, a comparative study focusing on the reusability of CgrAlcOx-catalase CLEA with potassium ferricyanide would shed more light on the applicability of these CLEA in biocatalysis. Building off the last point, we explored the replacement of HRP with potassium ferricyanide in the context of CgrAlcOx CLEA immobilization. Potassium ferricyanide has been known to affect CRO activity,74, 210 however recently a different inorganic oxidant (manganese (III) acetate) was shown to function as an HRP replacement as well75. The potential for other 142 water soluble, stable and affordable oxidizing inorganic metal salts could be explored as CRO activators, furthering the biocatalytic applicability of AA5_2 enzymes. The work in this thesis also presented a proof of concept for CgrAlcOx immobilization through cross-linking, which opens the door to explore other immobilization techniques. Chemical cross-linking to a surface is one of the most employed methodologies and it often utilizes active amino groups for the formation of covalent linkages. Compared to CLEA, the method of cross-linking enzymes to a surface has the added variable of choosing the physical and chemical characteristics of the carrier.150 Of course, by changing the immobilization method, more experiments are necessary to determine the optimum protocol for carrier surface modification. A possible disadvantage with this method is the inability of enzymes in a multi-enzyme system to be physically close together, an apparent requirement for CRO biocatalysis. Nonetheless, cross-linking CROs to a surface offers numerous advantages for biocatalytic applicability that the CLEA method does not. For example, oxygen concentration being crucial for high CROs biotransformation efficiency, the surface cross-linked carrier could be incorporated into columns, which would allow for flow chemistry processes with continuous oxygen supplementation. Furthermore, CgrAlcOx CLEA showed some reusability, which could be improved with more mechanical stability that is often associated with cross-linking enzymes to a surface. Lastly, the work presented in this thesis also serves as a basis for the immobilization of other CROs due to the similarity of their tertiary structures that is shared between many AA5_2 members. While the CROs presented in this thesis were assessed for their activity, selectivity and biocatalytic potential, there is interest in studying these enzymes in their hosts organisms. The in vivo activity of these enzymes is not understood and their physiological substrates remain unknown. Since most AA5_2 sequences are found in pathogenic fungi, there exists a great 143 opportunity to explore a possible link between copper radical oxidases and the pathogenicity of their hosts. The mechanism of CROs has been extensively studied and during their catalytic cycle they produce hydrogen peroxide as a by-product, which has been speculated to be coupled with the oxidative degradation of lignocellulosic biomass.84, 108 Gene deletion has been used in experiments performed with AA5_1 glyoxal oxidases in two pathogenic fungi. The deletion mutant of Streptomyces lividans showed an accumulation of glycans in the hyphal tips116 while the deletion of the AA5_1 in Ustilago maydis abolished its pathogenicity.108 Whether fungal CROs aid in the degradation of lignocellulose or are directly involved in pathogenicity of their host fungi, work exploring their biological role could potentially address these unanswered questions that still persists. As a final remark, the potential for biocatalysis to transform our industrial landscape of energy production, chemicals manufacturing, and pharmaceuticals synthesis is vast. 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E.; Haroune, L.; Ba, S.; Bellenger, J.-P.; Cabana, H., Characterization of combined cross-linked enzyme aggregates from laccase, versatile peroxidase and glucose oxidase, and their utilization for the elimination of pharmaceuticals. Sci. Total Environ. 2014, 481, 90-99. 278. Nguyen, L. T.; Yang, K.-L., Combined cross-linked enzyme aggregates of horseradish peroxidase and glucose oxidase for catalyzing cascade chemical reactions. Enzyme Microb. Technol. 2017, 100, 52-59. 162 Appendices Appendix A : Supporting Information for Chapter 2: Two Fusarium Copper Radical Oxidases with High Activity on Aryl Alcohols A.1 Supporting Tables Table A1. Pairwise sequence ID/similarity matrix between FgrAAO and FoxAAO and other AA5_2 enzymes. Sequence similarity is highlighted in turquoise and sequence identity is highlighted in blue. 1 2 3 4 5 6 7 8 9 10 1. FgrAAO (XP_011322138) 91.3 58.0 55.4 54.7 78.2 79.1 54.8 42.2 35.9 2. FoxAAO (XP_018246910) 95.1 57.3 54.7 53.9 82.8 84.2 54.0 41.2 35.9 3. FgrGalOx (AAO95371) 72.1 70.6 77.7 90.0 59.2 59.1 76.9 37.6 38.8 4. FoxGalOx (AHA90705) 68.1 67 86.9 81.3 60.7 60.6 94.9 36.7 39.5 5. FsaGalOx (AIR07394) 67.7 66.4 92.6 91.0 59.9 59.4 80.2 35.4 39.2 6. FsuAA5 (ADG08187) 81.7 84.7 71.9 76.1 74.7 96.9 60.6 35.8 36.8 7. FveAA5 (ADG08188) 82.3 85.4 72.0 75.3 74.4 98.4 60.3 35.8 36.8 8. FsuGalOx (AJE27923) 68.6 67.3 86.5 97.5 90.6 76.1 75.9 36.2 39.3 9. CgrAlcOx (EFQ30446) 57.2 57.1 52.1 50.5 49.2 50.2 50.1 50.7 33.4 10. CgrAAO (EFQ27661) 51.1 50.9 54.4 56.8 55.4 54.1 54.3 56.1 47.3 163 Table A2. Initial activity screens* of FgrAAO and FoxAAO Substrate Specific Activity (μmol min-1 mg-1) FgrAAO FoxAAO Carbohydrates D-Galactose (300 mM) 2.99 ±0.06 2.51 ±0.10 D-Lactose (300 mM) 2.59 ±0.03 2.29 ±0.42 Raffinose (300 mM) 1.41 ±0.04 1.24 ±0.23 D-Mannose (300 mM) 9.40 ±0.37 5.68 ±0.25 Melibiose (300 mM) 2.00 ±0.02 0.60 ±0.04 D-Fructose (300 mM) 3.00 ±0.07 3.33 ±0.12 D-Xylose (300 mM) 0.30 ±0.03 0.09 ±0.07 D-Glucose (300 mM) 0.15 ±0.01 0.09 ±0.00 Sucrose (300 mM) 0.22 ±0.01 0.25 ±0.00 D-Ribose (300 mM) n.m.§ n.m.§ L-Arabinose (300 mM) n.m.§ n.m.§ Xyloglucan (2.5 mg mL-1) 0.01 ±0.00 n.m.§ Carab Galactomannan (2.5 mg mL-1) 0.09 ±0.02 0.04 ±0.01 Levan (2.5 mg mL-1) 0.01 ±0.00 n.m.§ Inulin (2.5 mg mL-1) n.m.§ n.m.§ Fructo Oligos (2.5 mg mL-1) 0.03 ±0.00 n.m.§ Beta Mannan (2.5 mg mL-1) n.m.§ n.m.§ Polyols Glycerol (300 mM) 1.53 ±0.02 1.24 ±0.02 Sorbitol (300 mM) 0.09 ±0.00 0.07 ±0.00 Galactitol (300 mM) 0.04 ±0.00 0.03 ±0.00 Mannitol (300 mM) 0.11 ±0.00 0.07 ±0.00 Diols 1,3-Propanediol (300 mM) 0.26 ±0.00 0.20 ±0.00 1,2-Propanediol (300 mM) 0.23 ±0.00 0.15 ±0.00 1,4-Butanediol (300 mM) 0.34 ±0.00 0.25 ±0.01 Aldehyde Methyl Glyoxal (5 mM) 0.25 ±0.04 0.07 ±0.01 Primary alcohols Ethanol (300 mM) 0.06 ±0.00 0.04 ±0.00 Methanol (300 mM) 0.05 ±0.00 0.03 ±0.00 1-Butanol (300 mM) 0.09 ±0.00 0.05 ±0.00 Hexanol (300 mM) 0.03 ±0.00 0.08 ±0.00 2-Phenylethanol (1 mM) 0.01 ±0.00 n.m.§ Secondary alcohols 1-Phenylethanol (1 mM) n.m.§ n.m.§ Aryl alcohols Benzyl alcohol (5 mM) 1.23 ±0.03 1.23 ±0.02 Cinnamyl alcohol (5 mM) 4.90 ±0.13 5.00 ±0.08 4-methoxybenzyl alcohol (5 mM) 0.89 ±0.02 0.98 ±0.02 3-methoxybenzyl alcohol (5 mM) 12.21 ±0.09 16.41 ±0.25 Coniferyl alcohol (5 mM) n.m.§ n.m.§ Veratryl alcohol (5 mM) 20.13 ±0.42 25.19 ±0.89 4-hydroxybenzyl alcohol (5 mM) 1.05 ±0.09 0.45 ±0.10 Vanillyl alcohol (5 mM) 0.01 ±0.00 n.m.§ Furans HMF (5 mM) 3.00 ±0.09 3.46 ±0.13 DFF (5 mM) 0.91 ±0.04 0.98 ±0.18 HMFCA (5 mM) 1.82 ±0.07 2.64 ±0.35 164 FFCA (5 mM) n.m.§ n.m.§ *Measurements were performed in triplicate at 25 ˚C in 50 mM sodium phosphate buffer at pH 7.5 using the HRP/ABTS assay. Activities were monitored using concentrations indicated within parentheses for each substrate. §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 1 nmole of purified enzyme. 165 Table A3. Comparison of catalytic parameters of FgrAAO and FoxAAO with other enzymes acting on galactose, fructose and mannose* Enzyme Galactose Fructose Mannose References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) FgrAAO 1700 ± 150 21 ± 1.0 12 460 ± 91 9.7 ± 0.55 21 450 ± 74 36 ± 1.2 79 This work FoxAAO 1600 ± 150 23 ± 1.2 14 510 ± 82 16 ± 0.72 31 620 ± 57 19 ± 0.35 31 This work FgrGalOx 102 ±6.4 1059 ±18.9 10400 ±680 2.48 ±0.512 22.8 ±3.57 9.2 ±2.4 ND ND ND [102] R330Ka 895 ±85.9 208 ±10.8 232 ±25 1.81 ±0.111 80.4 ±5.01 75.1 ±9.1 ND ND ND [102] M1a 43 ±2 1376 ±35 32 000 ±1700 ND ND ND ND ND ND [99] M3b 54 ±7 1800 ±400 31 ±7 ND ND ND ND ND ND [104] Des 3-2a ND ND 1100 ±300 ND ND ND ND ND ND [96] H1b ND ND ND ND ND ND ND ND 35 ± 3 [103] C383Na 390 ±38 410 ±17 1100 ±150 ND ND ND ND ND ND [97] C383S, Y436H, V494Aa 19 ±2 1301 ±46 68500 ±7600 ND ND ND ND ND ND [100] V494Aa 53 ±8 1119 ±66 21100 ±3400 ND ND ND ND ND ND [100] CgrAAO ND ND 13.1 ±0.8 ND ND ND ND ND ND [48] * When comparing catalytic constants, please note that different assay temperatures, generally in the range 25 ˚C - 35 ˚C, may have been used for each enzyme, as described in the original publication cited. ND = not determined. M1 = S10P, M70V, P136, G195E, V494A, N353D; M3 = W290F, R330K and Q406T, Des 3-2 = Q326E, Y329K and R330K, H1 = R330K 166 Table A4. Comparison of catalytic parameters of FgrAAO and FoxAAO with other enzymes acting on benzyl alcohol and other aryl alcohols* Enzyme Benzyl alcohol Cinnamyl alcohol References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) FgrAAO 86 ± 12 37 ± 2.8 430 ND ND 3000 This work FoxAAO 30 ± 19 31 ± 4.2 1000 ND ND 3200 This work FgrGalOx ND ND 424 ±2 ND ND ND [213] CgrAlcOx 10.69 ±0.04 94 ±1 140000 0.06 ±0.003 93 ±1 1600000 [46] CgrAAO 27 ±0.9 54.5 ±0.6 2020 ± 70 ND ND ND [48] 3-methoxybenzyl alcohol 4-methoxybenzyl alcohol References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) FgrAAO 6.5 ± 1.5 52 ± 1.5 8000 59 ±7.2 32 ±1.8 540 This work FoxAAO 6.7 ± 0.59 34 ± 1.7 5100 51 ± 12 34 ± 2.9 290 This work FgrGalOx ND ND 4172 ±14 ND ND 449 ±2 [213] CgrAlcOx ND ND ND ND ND ND [46] CgrAAO 21 ±0.8 140 ±2 6600 ± 300 24 ±1.3 48 ±1 2000 ± 120 [48] * When comparing catalytic constants, please note that different assay temperatures, generally in the range 25 ˚C - 35 ˚C, may have been used for each enzyme, as described in the original publication cited. ND = not determined. 167 Table A5. Comparison of catalytic parameters of FgrAAO and FoxAAO with other enzymes acting on HMF and its derivatives* Enzyme HMF DFF HMFCA FFCA References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) FgrAAO 14 ± 2.6 29 ± 1.7 2100 10 ± 2.9 3.6 ± 0.27 360 4.4 ± 0.90 3.8 ± 0.33 860 ND ND ND This work FoxAAO 17 ± 4.9 26 ± 2.6 1500 3.7 ± 1.0 1.4 ± 0.13 380 3.1 ± 0.31 5.2 ± 0.14 1700 ND ND ND This work CgrAAO 6.5 ±0.3 126 ± 1.5 1940 ± 90 ND ND ND 26.9 ±3 28.3 ±1.3 1100 ± 100 ND ND ND [48] Bacterial HMFO 1.4 9.9 7100 1.7 1.6 940 73 8.5 120 ND ND <10 [38] PerAAO 1.6 ± 0.2 0.33 ± 0.01 220 ±42 3.3 ± 0.2 0.52 ± 0.01 158 ± 9.2 ND ND ND ND ND ND [218] MtGLOX 20.2 ± 9.0 15.9 982 ND ND ND ND ND ND ND ND ND [216] PciGLOX1 15.66 ± 2.35 1.59 ± 0.12 101.66 ± 0.01 4.38 ± 0.1 0.54 ± 0.24 124.39 ± 0.01 ND ND ND 0.85 ± 0.14 0.03 ± 0.01 38.55 ± 0.01 [217] PciGLOX2 5.87 ± 2.04 0.56 ± 0.09 96.04 ±0.01 0.21 ± 0.04 4.80 ± 0.24 23400 ± 100 ND ND ND 1.40 ± 0.39 2.02 ± 0.03 1400 ± 10 [217] PciGLOX3 6.35 ± 1.32 0.75 ± 0.07 118.35 ± 0.01 8 ±0.05 1.28 ± 0.09 7300 ± 10 ND ND ND 0.61 ± 0.58 0.04 ± 0.01 72.03 ± 0.01 [217] *When comparing catalytic constants, please note that different assay temperatures, generally in the range 25 ˚C - 35 ˚C, may have been used for each enzyme, as described in the original publication cited. ND = not determined. 168 Table A6. Assignments of 1H chemical shifts (ppm) of substrates and products of FgrAAO oxidation of 1-O-Methyl D-fructose as determined by 1D and 2D NMR experiments recorded at 400MHz in D2O. The peaks are referenced to internal acetone (2.22 ppm). Compound H-1 H-3 H-4 H-5 H-6 -OMe 3 3.77 (S) 3.93 (D) 3.87 - 3.84 (M) 3.82 (M) 3.76 (DD) 3.28 (S) 8 5.17 (S) 4.09 (D) 3.87 - 3.85 (M) 3.97 (M) 3.77 (S) 3.37 (S) 2 3.68 (DD) 4.17 (D) 3.87 -3.83 (M) 4.05 (T) 3.72 (DD) 3.32 (S) 7 3.70 (DD) 4.17 (S) 3.64 (S) 4.17 (S) 4.95 (D) 3.30 (S) 1 3.72 (DD) 4.10 (M) 3.96 (M) 3.96 (M) 3.75 (DD) 3.32 (S) Table A7. Assignments of 13C chemical shifts (ppm) of compounds substrates and products of FgrAAO oxidation of 1-O-Methyl D-fructose as determined by 1D and 2D NMR experiments recorded at 400MHz in D2O. The peaks are referenced to internal acetone (30.89 ppm). Compound C-1 C-2 C-3 C-4 C-5 C-6 -OMe 3 61.4 101.0 68.9 70.2 69.6 64.4 49.1 8 90.0 99.7 68.5 70.3 69.4 64.7 49.8 2 60.3 104.4 77.4 81.6 75.6 63.3 49.6 7 59.7 104.7 77.7 83.2 76.9 91.6 49.3 1 58.3 108.9 80.7 83.9 78.0 61.9 48.8 169 Table A8. Assignments of 1H and 13C chemical shifts (ppm) of FgrAAO oxidation of raffinose as determined by 1D and 2D NMR experiments recorded at 400MHz in D2O. The peaks are referenced to internal acetone (2.22 ppm). Ring System δ Multiplicity Coupling Constant (J) Ring System δ 1H (ppm) (Hz) 13C (ppm) Galactose (Product) Galactose (Product) H1 3.55 or 3.64 M n.a. C1 n.a. H2 3.55 or 3.64 M n.a. C2 n.a. H3 4.54 DD 2.98; 8.02 C3 66.41 H4 5.27 D 2.54 C4 99.96 H5 3.89 M n.a. C5 70.54 H6 6.19 D 3.00 C6 125.11 Galactose (Substrate) Galactose (Substrate) H1 4.99 D 3.63 C1 99.09 H2 3.83 M n.a. C2 69.10 H3 3.89 M n.a. C3 70.07 H4 4.00 D n.a. C4 69.82 H5 3.95 T 6.28 C5 71.64 H6/H’6 3.76 M n.a. C6 61.72 Glucose (Product) Glucose H1 5.39 D 3.91 C1 92.70 H2 3.55 M n.a. C2 n.a. H3 3.76 M n.a. C3 73.22 H4 3.55 M n.a. C4 69.98 H5 4.00 D n.a. C5 71.98 H6/H’6 4.05; 3.89 M; M n.a. C6 67.94 Glucose (Substrate) Glucose (Substrate) H1 5.42 D 3.83 C1 92.73 170 H2 3.55 M n.a. C2 71.58 H3 3.76 M n.a. C3 73.27 H4 3.55 M n.a. C4 70.02 H5 4.05 M n.a. C5 72.02 H6/H’6 4.05; 3.70 M; M n.a. C6 66.50 Fructose (Product) Fructose (Product) H1/H’1 3.70 M n.a. C1 70.21 H3 4.14 M n.a. C2 104.39 H4 3.55 M n.a. C3 69.34 H5 3.64 M n.a. C4 74.67 H6 5.12 D 7.36 C5 73.40 C6 89.09 Fructose (Substrate) Fructose (Substrate) H1/H’1 3.64 M n.a. C1 62.01 H3 4.21 M n.a. C2 104.43 H4 4.05 M n.a. C3 76.95 H5 3.89 M n.a. C4 74.60 H6/H’6 3.83; 3.76 M; M n.a. C5 81.97 C6 63.09 171 A.2 Supporting Figures FgrAAO --GVGKWGPTLDFPVIPVAGAVEPVSGKVVIWSAYRYDAFQGTNPRGGFTLTSIWDPKTN FoxAAO AKGLGKWGPTLDFPVIPVAGAVEPVSGKVVIWSAYRYDAFQGTTPRGGFTLTSIWDPKTN FgrGalOx --GLGRWGPTIDLPIVPAAAAIEPTSGRVLMWSSYRNDAFGGSP--GGITLTSSWDPSTG CgrAlcOx --NVGKWGPMVKFPVVPVAVALVPETGNLLVWSSG--WPNRWTTAGNGKTYTSLYNVNTG FgrAAO VISNRNVTNNKHDMFCPGISMDGEGQIVVTGGNDAKKTTILNP---NGE-WVPGPDMQIA FoxAAO VISNRNVSNNHHDMFCPGISMDGEGQIVVTGGNDAKKTTILMP---DGN-WVPGPDMQIA FgrGalOx IVSDRTVTVTKHDMFCPGISMDGNGQIVVTGGNDAKKTSLYDS---SSDSWIPGPDMQVA CgrAlcOx NISDAIVQNTQHDMFCPGTSLDADGRIIVTGGSSAAKTSVLDFKKGESSPWTPLSNMQIS FgrAAO RGYQSSATTSDGRVFTIGGSWSGPRGGKNGEIYDPKARTWTSLPKCLVGPMLTKDKEGVY FoxAAO RGYQSSATCSDGRVFTIGGSWSGPRGGKNGEIYDPKAKTWTSLPKCLVGPMLTKDKEGVY FgrGalOx RGYQSSATMSDGRVFTIGGSWSGGVFEKNGEVYSPSSKTWTSLPNAKVNPMLTADKQGLY CgrAlcOx RGYQSSCTTSEGKIFVIGGSFSG-AGTRNGEVYDPKANTWTKLAGCPVKPLVMQRG---M FgrAAO KADNHAWLFGWKKGSVFQAGPSTAMNWYYTTRGTQGDTKAAGTRRKNGRVDPDSMNGNCV FoxAAO KADNHAWLFGWKKNSVFQAGPSTAMNWYYTTRGTQGDTKAAGTRRKNGRIDPDSMNGNVA FgrGalOx RSDNHAWLFGWKKGSVFQAGPSTAMNWYYTSG--SGDVKSAGKRQSNRGVAPDAMCGNAV CgrAlcOx FPDSHAWLWSWKNGSVLQAGPSKKMNWYDTKG------TGSNTPAGLRGTDEDSMCGVSV FgrAAO MYDALDGKILTYGGATSYQQAPATANAHVLAIAEPGAIAQTYLVGNNGAGNYARVFHTSV FoxAAO MFDALNGKILTFGGATSYQQAPATANAHVLTIDEPGAIAQTALVGNNGAGIHARVFATSV FgrGalOx MYDAVKGKILTFGGSPDYQDSDATTNAHIITLGEPGTSPNTVFASNG--LYFARTFHTSV CgrAlcOx MYDAVAGKIFTYGGGKGYTGYDSTSNAHILTLGEPGQAVQVQKLANG---KYNRGFANAV FgrAAO VLPDGNVFITGGQSYSNPFTDTNAQLTPEMYIPTTHEFKTQQPNTIPRTYHSMSLLLPDA FoxAAO ILPDGNVFITGGQSYSDPFTDTNAQLEPEMFISSSNTFTKQQTNTIPRTYHSMSLLLPDA FgrGalOx VLPDGSTFITGGQRRGIPFEDSTPVFTPEIYVPEQDTFYKQNPNSIVRVYHSISLLLPDG CgrAlcOx VMPDGKIWVVGGMQKMWLFSDTTPQLTPELFDPATGSFTPTTPHTVPRNYHSTALLMADA FgrAAO TVFNGGGGLCGS-CSSNHFDAQIYTPQYLLDGN-GNFATRPKITAVSATTAKIGSTITVT FoxAAO TVFNGGGGLCGG-CKTNHFDAQIFTPQYLLDGN-GNLATRPKITAVSATTAKVGSTITVT FgrGalOx RVFNGGGGLCGD-CTTNHFDAQIFTPNYLYNSN-GNLATRPKITRTSTQSVKVGGRITIS CgrAlcOx TIWSGGGGLCGANCKENHFDGQFWSPPYLFEADGVTPAKRPVIQSLSDTAVRAGAPITIT FgrAAO ANSA-IKSASLIRYGTATHTVNTDQRRIPLALTGAG-TNKYSFKIPNDSGIALPGYWMLF FoxAAO ANSA-IKSASLIRYGTATHVVNTDQRRIPLALTGAG-TNKYSFKIPNDSGIALPGYWMLF FgrGalOx TDSS-ISKASLIRYGTATHTVNTDQRRIPLTLTNNG-GNSYSFQVPSDSGVALPGYWMLF CgrAlcOx MQDAGAYTFSMIRVSATTHTVNTDQRRIPLDGQDGGDGKSFTVNVPNDYGVAIPGYYMLF FgrAAO VLNNAGVPSVASTIKVTV FoxAAO VLNNAGVPSVASTIKVTI FgrGalOx VMNSAGVPSVASTIRVTQ CgrAlcOx AMNEAGVPCVAQFFKVTL Figure A1. Sequence alignment of catalytic domains of Fusarium graminearum AAO (FgrAAO), Fusarium oxysporum AAO (FoxAAO), Fusarium graminearum GalOx (FgrGalOx) and Colletotrichum graminicola AlcOx (CgrAlcOx). Predictive signal peptides and N-terminal modules have been removed. Conserved active-site catalytic residues and residues involved in substate recognition and stability are highlighted in green and yellow, respectively. 172 Figure A2. SDS-PAGE of FgrAAO and FoxAAO and N-deglycosylation study. A. Aliquot of purified FgrAAO (4 μg) (lane 1) and purified FoxAAO (4 μg) (lane 2). B. FgrAAO and FoxAAO (3 μg 1 to 6) were N-deglycosylated under denaturing conditions with PNGaseF. 1: PNGaseF, 2: native FgrAAO (3 μg), 3: FgrAAO (3 μg) + pNGaseF, 4: native FoxAAO (3 μg), 5: FoxAAO (3 μg) + pNGaseF. All gels were stained by Coomassie blue. MW = molecular weight markers, as indicated. Figure A3. pH-rate profiles. A. FgrAAO; B. FoxAAO. pH-rate profiles were determined using the coupled HRP-ABTS assay with 1M galactose as the substrate at each pH value. Black squares for phosphate citrate buffer, red circles for sodium phosphate buffer and blue triangles for glycine NaOH buffer. Error bars represent standard deviations over three replicates. 5 6 7 8 9 10012345 Phosphate-citrate Sodium-phosphate Glycine-NaOHSpecific Activity (µmol/min/mg enz)pHA. 5 6 7 8 9 10012345 Phosphate-citrate Sodium-phosphate Glycine-NaOHSpecific Activity (µmol/min/mg enz)pHB. 173 Figure A4. Temperature-rate profiles. A. FgrAAO; B. FoxAAO. Temperature-rate profiles were determined using the coupled HRP-ABTS assay. 50 mM sodium phosphate, pH 7.5 and 1M galactose were incubated at each temperature for at least 10 min prior to the addition of HRP, ABTS and purified enzyme. Error bars represent standard deviations over three replicates. Figure A5. Temperature stability. A. FgrAAO; B. FoxAAO. Activity values were determined using the coupled HRP-ABTS assay with 1 M galactose as the substrate. The enzyme was pre-incubated at each temperature, maintained by a gradient thermocycler: 30 °C (black square), 39 °C (red circle), 49 °C (blue triangle), 60 °C (green triangle). The reactions were performed at 30 °C. Error bars represent standard deviations over duplicate measurements. 0 10 20 30 40 50 60 70 8002468101214161820Specific Activity (µmol/min/mg enz)Temperature0 10 20 30 40 50 60 70 8002468101214161820Specific Activity (µmol/min/mg enz)TemperatureA. B. 0 500 1000 1500 2000 2500 3000 3500020406080100120Time (min) 30oC 39oC 49oC 60oCResidual Activity (%)0 500 1000 1500 2000 2500 3000 3500020406080100120Residual Activity (%)Time (min) 30oC 39oC 49oC 60oCA. B. 174 175 176 177 178 Figure A6. Initial-rate kinetics. Panels A-L: FgrAAO, panels M-X: FoxAAO. Individual substrates are indicated in the x-axis labels. Values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. For cinnamyl alcohol, which did not display saturation kinetics for either enzyme due to solubility limitations, kcat/KM values were obtained from the slope of linear fittings. Benzyl alcohol with FoxAAO (panel Q) displayed apparent substrate inhibition. The data was fit both with the classical Michaelis-Menten equation and the substrate inhibition variant, Vo/[E]T = kcat[S]/(KM + [S] + [S]2/Kis). Due to high errors on the individual constants kcat, KM, and Kis in the latter case, kcat and KM values from the former are reported in Table 2.1. 179 Figure A7. 1H-NMR analysis of the oxidation of HMF by FgrAAO and FoxAAO. A. FgrAAO; B. FoxAAO. 1H NMR spectra (400 MHz, 1:9 D2O:phosphate buffer, 50 mM, pH 7.5) of reaction product profiles after 16.5 h incubation with FgrAAO (top, pink) and FoxAAO (bottom, blue) in the presence of catalase and HRP for HMF at 10 mM showing full conversion to DFF ( ) and the hydrated form, DFFhyd( ), as well as FFCA ( ). 180 Figure A8. 1H-NMR analysis of the oxidation of DFF by FgrAAO and FoxAAO. A. FgrAAO; B. FoxAAO. 1H NMR spectra (400 MHz, 1:9 D2O:phosphate buffer, 50 mM, pH 7.5) of reaction product profiles after 16.5 h incubation with FgrAAO (top, pink) and FoxAAO (bottom, blue) in the presence of catalase and HRP for DFF ( and ) at 10 mM showing partial conversion to FFCA ( ). 181 Figure A9. 1H-NMR analysis of the oxidation of HMFCA by FgrAAO and FoxAAO. A. FgrAAO; B. FoxAAO. 1H NMR spectra (400 MHz, 1:9 D2O:phosphate buffer, 50 mM, pH 7.5) of reaction product profiles after 16.5 h incubation with FgrAAO (top, pink) and FoxAAO (bottom, blue) in the presence of catalase and HRP for HMFCA ( ) at 10 mM showing almost full conversion to FFCA ( ). 182 183 184 Figure A10. Complete assignment of proton and carbon nuclei from 1H,13C HSQC of fructose derivatives. A. Methyl β-D-fructopyranoside. B. Methyl β-D-fructofuranoside. C. Methyl α -D-fructofuranoside. Each black crosspeak corresponds to a 1JC,H-coupling interaction from the HSQC experiment. 185 186 187 Figure A11. Selective assignment of proton and carbon nuclei from 1H,13C HMBC of fructose derivatives. A. Methyl β-D-fructopyranoside. B. Methyl β-D-fructofuranoside. C. Methyl α-D-fructofuranoside. Each black crosspeak corresponds to a 2JC,H- or 3JC,H -coupling interaction from the HMBC experiment. 188 Figure A12. Analysis of oxidation reaction components of FgrAAO of methyl β-D-fructopyranose from 1H,13C HSQC zoomed into 3.6-4.6 ppm region. Each black crosspeak corresponds to a 1JC,H-coupling interaction from the HSQC experiment. 189 Figure A13. Analysis of oxidation reaction components of FgrAAO of methyl β-D-fructofuranose from 1H,13C HSQC zoomed into 3.6-4.6 ppm region. Each black crosspeak corresponds to a 1JC,H-coupling interaction from the HSQC experiment. 190 Figure A14. Analysis of oxidation reaction components of FgrAAO of methyl α-D-fructofuranoside from 1H,13C HSQC. Each black crosspeak corresponds to a 1JC,H-coupling interaction from the HSQC experiment. Two carbon peaks at 63.1 ppm and 72.7 ppm have been attributed to an impurity that is present in either the HRP, catalase or buffer solutions. 191 192 193 Figure A15. Selective assignment of proton and carbon nuclei from 1H,13C HMBC of oxidation reaction components of FgrAAO with fructose derivatives. A. Methyl β-D-fructopyranoside. B. Methyl β-D-fructofuranoside. C. Methyl α-D-fructofuranoside. Each black crosspeak corresponds to a 2JC,H- or 3JC,H -coupling interaction from the HMBC experiment. Two carbon peaks at 63.1 ppm and 72.7 ppm have been attributed to an impurity that is present in either the HRP, catalase or buffer solutions. 194 195 196 Figure A16. Analysis of oxidation reaction components of FoxAAO of fructose derivatives from 1H,13C HSQC. A. Methyl β-D-fructopyranoside. B. Methyl β-D-fructofuranoside. C. Methyl α -D-fructofuranoside. Each black crosspeak corresponds to a 1JC,H-coupling interaction from the HSQC experiment. Two carbon peaks at 63.1 ppm and 72.7 ppm have been attributed to an impurity that is present in either the HRP, catalase or buffer solutions. 197 198 199 Figure A17. Selective assignment of proton and carbon nuclei from 1H,13C HMBC of oxidation reaction components of FoxAAO with fructose derivatives. A. Methyl β-D-fructopyranoside. B. Methyl β-D-fructofuranoside. C. Methyl α-D-fructofuranoside. Each black crosspeak corresponds to a 2JC,H- or 3JC,H -coupling interaction from the HMBC experiment. Two carbon peaks at 63.1 ppm and 72.7 ppm have been attributed to an impurity that is present in either the HRP, catalase or buffer solutions. 200 Figure A18. Selective assignment of proton and carbon nuclei from 1H,13C HSQC and HMBC of oxidation product FgrAAO with 2,5-anhydro-mannitol. Each teal and blue crosspeak corresponds to a 1JC,H-coupling interaction from the HSQC experiment while each red crosspeak corresponds to a 2JC,H- or 3JC,H -coupling interaction from the HMBC experiment. Two carbon peaks at 63.1 ppm and 72.7 ppm have been attributed to an impurity that is present in either the HRP, catalase or buffer solutions. 201 Figure A19. Selective 1H 1D-TOSCY experiment of oxidized raffinose by FgrAAO. A. New proton peaks in the proton region between 5.0-6.5 ppm were targeted based on the full 1D-NMR spectrum. Spectra corresponding to individual spin systems are displayed in various colours with B. the galactose ring of oxidized raffinose in red. C. the fructose ring in oxidized raffinose in blue. Irradiations from 1H 1D-TOSCY experiments are displayed with black stars above corresponding peaks. 202 Figure A20. 1H 2D-TOSCY experiment oxidized raffinose by FgrAAO. Each peak indicates protons that are coupled through magnetization transfer. Selective assignment of crosspeaks originating from the same spin system are shown in matching colours specific to each monosaccharide with oxidized/unoxidized galactose shown in dark blue and light blue respectively; oxidized/unoxidized fructose shown in dark red and magenta respectively; glucose in orange. 203 Figure A21. Complete assignment of proton and carbon nuclei from oxidized raffinose by FgrAAO using 1H,13C HSQC experiments. Each crosspeak corresponds to a 1JC,H-coupling interaction from the HSQC experiment. Red crosspeaks correspond to unoxidized substrate, blue crosspeaks correspond to oxidized product and purple crosspeaks correspond to signals overlapping from substrate and product. Black crosspeaks are from impurities associated with the buffer, HRP and catalase mixture. The notation stated the sugar ring and its corresponding atoms(s) below it. Abbreviations used: Gal, galactose; Glu, glucose, Fru, fructose; ox, Oxidized. 204 Figure A22. Selective assignment of proton and carbon nuclei from 1H,13C HMBC of oxidation reaction components of FgrAAO with raffinose. Each crosspeak corresponds to a 2JC,H- or 3JC,H -coupling interaction from the HMBC experiment. Crosspeaks for C5-H6 on the galactose ring in the product and C6-H5 on the fructose ring in the oxidized product are highlighted in blue and red, respectively. Two carbon peaks at 63.1 ppm and 72.7 ppm have been attributed to an impurity that is present in either the HRP, catalase or buffer solutions.205 Figure A23. Stereochemistry determination of glycerol oxidation by FgrAAO and FoxAAO. A. Chromatogram of L/D-glyceraldehyde-hydrazone. B. Glyceraldehyde-hydrazone composition after FgrGalOx oxidation of glycerol. C. Glyceraldehyde-hydrazone composition after CgrAlcOx oxidation of glycerol. D. Glyceraldehyde-hydrazone composition after FgrAAO oxidation of glycerol. E. Glyceraldehyde-hydrazone composition after FoxAAO oxidation of glycerol. Peaks are labelled with retention times. 206 Figure A24. Representative mass spectrum of glycerol oxidation by FgrGalOx and CgrAlcOx. A. L-glyceraldehyde-hydrazone from FgrGalOx oxidation (peak at 2.37, Figure B23). B. D-glyceraldehyde-hydrazone from CgrAlcOx oxidation (peak at 2.83, Figure B23). 207 Appendix B : Supporting Information for Chapter 3: Biochemical survey of Copper Radical Oxidase specificity in Auxiliary Activity Family 5 B.1 Supporting Tables Table B1. Key amino acid residues in selected AA5_2 genes compared to other characterized members. Enzyme Organism GenBank/JGI mycocosm Accession Active Site Amino Acids Distal Amino Acids Modularity Radical Stabilization Substrate Recognition/Active Site Cavity Shape Catalytic Efficiency FgrGalOx Fusarium graminearum AAO95371 W290 F194 Q326 Y329 R330 Q406 P463 C383 CBM 32-CAT CgrAlcOx Colletorichum graminicola EFQ30446 F W G F M T L C CAT CgrAAO Colletorichum graminicola EFQ27661 Y F E W R T V C PAN 1-CAT FgrAAO Fusarium graminearum XP_011322138 W F E Y K Q P N CBM 32-CAT Y1 Eurotium rubrum XP_040635673 W F K W R D P Y CBM32-CAT Y11 Melanomma pulvis-pyrius KAF2798557 W F G F R A P N CBM32-CBM32-CAT Y12 Niesslia exilis jgi|Nieex1|772836 W F E W R D A S CBM32-CAT Y13 PfeGalOx Penicillium fellutanum jgi|Penfe1|382062 W Y Q Y R Q P S CBM32-CAT Y14 PhuRafOx Pseudozyma hubeiensis XP_012186969 Y F A W R S G C UNK-CAT Y15 Setosphaeria turcica XP_008020456 W F Q L R V G C UNK-CAT Y16 StoAA5 Stagonospora sp. OAK97814 W Y E F R D G C CAT Y17 Talaromyces marneffei XP_002148539 F F G Y R N V T UNK-CAT Y18 AsyAlcOx Aspergillus sydowii XP_040706357 F Y H D R V P C UNK-CBM32-CAT 208 Y19 Torpedospora radiata jgi|Torra1|425635 W Y E F R Q P C CBM32-CAT Y2 UmaRafOx Ustilago maydis XP_011389156 Y F A W R S G C UNK-CAT Y20 Metarhizium acridum XP_007812971 W Y Q Y R Q P N CBM32-CAT Y21 Leptosphaeria maculans XP_003841328 W F S F H G G C CAT Y22 Acremonium strictum jgi|Acrst1|1377707 W F E S K R P N CBM32-CAT Y23 CcaAA5 Corynespora cassiicola PSN67470 W A A N E H G C CAT Y24 Myrothecium inundatum jgi|Myrin1|117638 W F A Y K A N C CBM32-CAT Y25 Niesslia exilis jgi|Nieex1|192818 W F Q F R Q P C CBM32-CAT Y29 MreGalOx Mytilinidion resinicola XP_033570565 W F Q F R Q P N CBM32-CBM32-CAT Y3 Loramyces macrosporus jgi|Lorma1|509879 Y F E W R T V C UNK-CAT Y35 GciAlcOx Glomerella cingulata jgi|Gloci1|1901294 F W L F M T L C CAT Y36 PruAA5_2 (PruAlcOx) Penicillium rubens CAP96757 W Y D Y R E P S CBM32-CAT Y4 Colletotrichum incanum KZL84074 W F R F A D Q N CAT Y40 FoxAlcOx Fusarium oxysporum f. sp. conglutinans race 2 54008 FOPG_18201 W F D S K A Q C CBM32-CAT Y41 FoxGalOxB Fusarium oxysporum FOTG_04629 W M Q F R Q P C UNK-CBM32-CAT Y42 ExeGalOx Exophiala xenobiotica KIW55415 W F Q F R Q P C CBM32-CBM32-CAT Y5 PorAlcOx Magnaporthe oryzae XP_003719369 F F G L Y T L C WSC-CAT Y6 Colletotrichum zoysiae jgi|Colzo1|587911 Y F G Y - T V C UNK-CAT 209 Y7 Amniculicola lignicola KAF1999280 W Y D Y R Q P N CBM32-CBM32-CAT Y8 AflAlcOx Aspergillus flavus KAF7627372 W Y L Y H E P S UNK-CBM32-CAT Y9 Didymella exigua XP_033446128 W F Q F R E P C CBM32-CBM32-CAT Y10 Cladonia grayi jgi|Clagr3|10990 W F E Y R N V C CBM32-CAT Y26 Bisporella sp. jgi|Bissp1|110239 W Y Q F R Q P N UNK-CBM-CAT Y27 Xanthoria parietina jgi|Xanpa2|1578647 W F R Y R Q P N CBM32-CAT Y28 Pseudovirgaria hyperparasitica XP_033597585 W F G V R E P N CBM32-CBM32-CAT Y30 Ampelomyces quisqualis KAF1914598 Y F E W R T V C UNK-CAT Y31 Usnea florida jgi|Usnflo1|819633 W F Q Y R Q P C CBM32-CAT Y32 Nectria haematococca XP_003039318 W F E Y R R P C CBM32-CAT Y33 Penicillium rubens CAP97212 W F - L R D P Y CBM32-CAT Y34 Claviceps purpurea CCE30181 W F Q Y R Q P D CBM32-CAT Y37 Phialophora attae KPI34764 W Y K Y R D I C UNK-CAT 210 Table B2. Pairwise sequence ID/similarity matrix between characterized AA5_2 enzymes. Sequence similarity is highlighted in turquoise and sequence identity is highlighted in blue. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 1. FoxGalOx_2 (FOTG_04629) 33.8 32.4 33.3 39.6 36.8 36.4 26.3 22.6 31.5 25.9 24.9 23.2 22.1 39.7 31.4 29.6 22.5 27.9 2. AflAlcOx (KAF7627372) 46.1 48.2 50.9 43.8 38.9 49.4 36.2 32.3 45.2 35.2 36.2 33.7 31.8 51.6 42.9 28.2 31.7 35.5 3. PruAA5_2A (PruAlcOx) (CAP96757) 43.3 63.6 56.1 40 36.8 48.6 37.1 34.6 42.8 35.5 36.1 34.5 33.8 49 43.3 26 32.5 30.9 4. PfeGalOx (jgi|Penfe1|382062) 44.8 65.9 72.1 40.6 36.6 49.6 36.4 33.7 43.1 36 36.3 35.6 32.6 47.5 45.3 26 32.1 35.2 5. ExeGalOx (KIW55415) 51.2 58.2 52.8 55.3 52.7 43.7 35 32.7 37.9 33.8 33.9 32.4 29.8 46.5 39.1 32 31.3 35.9 6. MreGalOx (XP_033570565) 48.7 53.4 50.2 49.8 66.7 42.3 35.1 31.3 35.3 34.7 34.4 32.3 29.6 40.8 36 32.7 30.5 32.2 7. FoxAlcOx (FOPG_18201) 46.2 65.6 64.6 64.6 55.1 52.6 39.6 37.4 43.3 35.7 37.6 36.2 35 58.4 67.6 28.2 35.8 36 8. PorAlcOx (XP_003719369) 36.9 52.6 54.2 53.6 46.2 47.6 52.2 57.3 37.7 37.7 38.3 38.4 38.5 37.9 40 26 55 33.9 9. GciAlcOx (jgi|Gloci1|1901294) 31.5 46.2 49.9 49.9 42.3 41.4 50.4 68.8 31.7 36.4 36.4 46.9 44.5 33.9 36.9 23.4 79.4 29.8 10. AsyAlcOx (XP_040706357) 43 63.6 61.2 59.5 53.4 51 60 53.2 46.8 32.8 33.2 33.4 31.8 39.2 35.8 27.5 29.9 32.7 11. UmaRafOx (XP_011389156) 35.7 51.6 52.6 53.9 47.6 48.2 51.6 54.6 52.7 51.1 80.5 43.2 39.9 36.2 35.1 43.2 33.4 35.5 12. PhuRafOx (XP_012186969) 35.9 53.1 53.9 54.1 49.5 48.7 52.9 55.5 53 52 88.2 44.5 41 36 36.3 42.5 33.4 36.7 13. StoAA5 (OAK97814) 32 46.5 50.2 48.4 42.1 43.3 49.6 53.5 65.3 46.1 57.5 58.9 62.7 34.4 36.4 27.1 41.3 36.9 14. CcaAA5 (PSN67470) 30.3 44.4 47.7 45.9 39.5 39.2 46.8 52.7 63.2 45.3 54.3 55.4 75.6 32.7 35.1 24.1 40.1 34 15. FgrGalOx (AAO95371) 48.5 67.6 66.4 65.2 58 51.9 71.9 53.8 48.5 57.9 54.8 55.6 50.5 47.6 58 30 37.6 38.8 16. FgrAAO (XP_011322138) 40.7 57.3 60.4 60.9 49.6 46.6 74.6 56.3 54.5 52.3 53.2 53.9 53.3 50.9 72.1 27.6 42.2 35.9 17. CgrRafOx (EFQ36699.1) 44.3 42.1 39.8 41.1 48.6 48 40.9 39.4 34.6 41.4 52.2 52 36 34.4 40.9 37.8 25.2 32.8 18. CgrAlcOx (EFQ30446) 31.6 46.2 46.9 48.1 41.5 40.3 48.5 67.5 88.4 45.3 50.2 49.8 60.8 59.1 52.1 57.2 34.7 33.4 19. CgrAAO (EFQ27661.1) 40.7 54.6 51.3 54.7 51.1 49.2 52.2 50.4 44.6 50.5 53.3 54.9 48.7 47.4 54.4 51.1 47.3 47.3 211 Table B3. Purification yields of expressed AA5_2 proteins from 400 mL of BMMY culture after three days of methanol feeding. Enzyme Purification Yield mg mg L-1 UmaRafOx 16 40 PorAlcOx 0.9 2.2 AflAlcOx 17 43 PfeGalOx 8.4 21 PhuRafOx 2.4 6.0 StoAA5 2.2 5.5 AsyAlcOx 26 65 CcaAA5 3.9 9.8 MreGalOx 2.0 5.0 GciAlcOx 21 53 PruAlcOx 14 35 FoxAlcOx 2.8 7.0 FoxGalOxB 3.0 7.5 ExeGalOx 3.1 7.8 Table S4. Initial activity screens* of produced AA5_2 on galactose, raffinose and benzyl alcohol Enzyme Specific Activity (μmol min-1 mg-1) Galactose (300 mM) Raffinose (300 mM) Benzyl Alcohol (30 mM) UmaRafOx 0.19±0.01 2.78±0.27 1.11±0.07 PorAlcOx 0.44±0.02 0.97±0.45 4.44±0.09 AflAlcOx 15.1±4.3 n.m.§ 48.8±1.2 PfeGalOx 41.4±4.7 27.8±1.0 11.4±2.3 PhuRafOx 0.04±0.02 1.89±0.45 1.00±0.09 AsyAlcOx 0.30±0.02 0.06±0.01 3.71±0.08 MreGalOx 4.08±0.48 4.53±0.14 0.38±0.01 GciAlcOx 2.27±0.12 26.9±0.9 213±25. PruAlcOx 50.1±0.6 20.2±0.9 35.5±2.7 FoxAlcOx 0.79±0.03 0.15±0.03 2.02±0.19 FoxGalOxB 68.0±1.0 50.3±4.3 6.50±0.22 ExeGalOx 13.8±1.0 6.52±0.15 3.45±0.27 *Measurements were performed in triplicates at 30 ˚C in 100 mM sodium phosphate buffer at pH 8.0 using the HRP/ABTS assay. Activities were monitored using concentrations indicated within parentheses for each substrate. §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 0.39 nmole of purified enzyme. 212 Table B5. Initial activity screens* of ExeGalOx and MreGalOx Substrate Specific Activity (μmol min-1 mg-1) ExeGalOx MreGalOx Carbohydrates D-Galactose (300 mM) 20.7 ±0.42 5.73 ±1.72 D-Lactose (300 mM) 6.27 ±0.45 1.93 ±0.20 Raffinose (300 mM) 9.06 ±1.45 4.28 ±1.06 D-Mannose (300 mM) 0.01 ±0.00 0.01 ±0.00 Melibiose (300 mM) 9.00 ±0.80 4.67 ±0.50 D-Fructose (300 mM) 0.08 ±0.00 n.m.§ D-Xylose (300 mM) 0.12 ±0.01 0.01 ±0.00 D-Glucose (300 mM) 0.01 ±0.00 0.01 ±0.00 Sucrose (300 mM) n.m.§ 0.02 ±0.02 D-Ribose (300 mM) n.m.§ 0.01 ±0.00 L-Arabinose (300 mM) 0.68 ±0.09 0.08 ±0.02 Polyols Glycerol (300 mM) 1.42 ±0.01 0.09 ±0.02 Sorbitol (300 mM) 0.37 ±0.00 0.01 ±0.00 Galactitol (300 mM) 0.16 ±0.00 0.01 ±0.00 Mannitol (300 mM) NA 0.01 ±0.00 Diols 1,2-Propanediol (300 mM) 0.23 ±0.03 n.m.§ 1,3-Propanediol (300 mM) 0.68 ±0.03 0.01 ±0.00 1,4-Butanediol (300 mM) 0.66 ±0.01 0.01 ±0.00 1,5-Pentanediol (300 mM) NA n.m.§ 1,6-Hexanediol (300 mM) NA n.m.§ Aldehyde Methyl Glyoxal (5 mM) n.m.§ n.m.§ Glycolaldehyde Dimer (5 mM) NA NA Primary alcohols Ethanol (300 mM) 0.30 ±0.01 n.m.§ Methanol (300 mM) 0.12 ±0.01 n.m.§ 1-Butanol (300 mM) 0.39 ±0.02 n.m.§ Hexanol (2.5 mM) 0.38 ±0.04 0.01 ±0.00 Octanol (2.5 mM) n.m.§ n.m.§ Aryl alcohols Benzyl alcohol (30 mM) 0.72 ±0.02 0.02 ±0.01 Cinnamyl alcohol (2.5 mM) 0.40 ±0.00 n.m.§ 4-methoxybenzyl alcohol (2.5 mM) 0.40 ±0.01 0.02 ±0.02 3-methoxybenzyl alcohol (2.5 mM) 0.81 ±0.02 0.04 ±0.01 Coniferyl alcohol (5 mM) n.m.§ n.m.§ Veratryl alcohol (2.5 mM) 1.19 ±0.02 0.05 ±0.01 4-hydroxybenzyl alcohol (2.5 mM) 0.06 ±0.01 n.m.§ Vanillyl alcohol (2.5 mM) 0.01±0.00 0.02 ±0.02 Furans HMF (2.5 mM) 0.26 ±0.05 n.m.§ HMFCA (2.5 mM) 0.31 ±0.04 n.m.§ DFF (2.5 mM) n.m.§ n.m.§ FFCA (2.5 mM) 0.02 ±0.01 n.m.§ Furfural (2.5 mM) NA n.m.§ 213 *Measurements were performed in triplicate at RT in 100 mM buffer in different buffers at different pH using the HRP/ABTS plate assay. Activities were monitored using concentrations indicated within parentheses for each substrate. §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 0.17 nmole of purified ExeGalOx enzyme and of purified MreGalOx enzyme. 214 Table B6. Initial activity screens* of PfeGalOx and FoxGalOxB Substrate Specific Activity (μmol min-1 mg-1) PreGalOx FoxGalOxB Carbohydrates D-Galactose (300 mM) 182 ±11.6 60.7 ±5.77 D-Lactose (300 mM) 69.7 ±11.2 16.2 ±1.16 Raffinose (300 mM) 93.0 ±0.75 25.9 ±1.34 D-Mannose (300 mM) 15.4 ±1.27 0.37 ±0.09 Melibiose (300 mM) 26.7 ±11.5 59.7 ±3.46 D-Fructose (300 mM) 23.0 ±4.04 0.35 ±0.02 D-Xylose (300 mM) 114 ±22.0 0.61 ±0.06 D-Glucose (300 mM) 4.55 ±0.46 0.03 ±0.02 Sucrose (300 mM) 0.44 ±0.08 0.01 ±0.00 D-Ribose (300 mM) 45.8 ±2.40 0.85 ±0.09 L-Arabinose (300 mM) 9.02 ±0.49 2.13 ±0.07 Polyols Glycerol (300 mM) 193 ±44.9 5.07 ±0.22 Sorbitol (300 mM) 28.0 ±2.36 2.63 ±0.07 Galactitol (300 mM) 5.29 ±0.30 1.04 ±0.06 Mannitol (300 mM) 19.6 ±1.91 NA Diols 1,2-Propanediol (300 mM) 11.4 ±1.29 2.44 ±0.22 1,3-Propanediol (300 mM) 6.21 ±1.76 4.88 ±0.42 1,4-Butanediol (300 mM) 1.33 ±0.18 5.57 ±0.37 1,5-Pentanediol (300 mM) 0.66 ±0.03 4.83 ±0.15 1,6-Hexanediol (300 mM) 4.28 ±0.44 3.68 ±0.23 Aldehyde Methyl Glyoxal (5 mM) 0.43 ±0.14 0.61 ±0.02 Primary alcohols Ethanol (300 mM) 0.43 ±0.14 3.76 ±0.57 Methanol (300 mM) 1.25 ±0.02 0.89 ±0.05 1-Butanol (300 mM) 0.91 ±0.07 3.80 ±0.10 Hexanol (2.5 mM) 0.24 ±0.07 2.24 ±0.31 Octanol (2.5 mM) 0.44 ±0.17 2.00 ±0.11 Aryl alcohols Benzyl alcohol (30 mM) 20.3 ±1.59 5.84 ±0.20 Cinnamyl alcohol (2.5 mM) 6.76 ±1.02 3.60 ±0.43 4-methoxybenzyl alcohol (2.5 mM) 2.39 ±0.21 1.78 ±0.71 3-methoxybenzyl alcohol (2.5 mM) 3.76 ±0.15 6.66 ±0.30 Coniferyl alcohol (5 mM) 0.15 ±0.08 n.m.§ Veratryl alcohol (2.5 mM) 2.34 ±0.21 10.2 ±0.21 4-hydroxybenzyl alcohol (2.5 mM) 0.05 ±0.01 0.10 ±0.12 Vanillyl alcohol (2.5 mM) 0.01 ±0.00 n.m.§ Furans HMF (2.5 mM) 1.51 ±0.12 3.34 ±0.20 HMFCA (2.5 mM) 1.31 ±0.19 3.31 ±0.32 DFF (2.5 mM) 0.43 ±0.18 0.52 ±0.03 FFCA (2.5 mM) 0.30 ±0.11 0.02 ±0.00 Furfural (2.5 mM) 0.17 ±0.03 n.m.§ *Measurements were performed in triplicate at RT in 100 mM buffer in different buffers at different pH using the HRP/ABTS plate assay. Activities were monitored using concentrations indicated within parentheses for each substrate. 215 §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 0.2 nmole of purified PfeGalOx enzyme and 0.13 nmole of purified FoxGalOxB enzyme. 216 Table B7. Initial activity screens* of UmaRafOx and PhuRafOx Substrate Specific Activity (μmol min-1 mg-1) UmaRafOx PhuRafOx Carbohydrates D-Galactose (300 mM) 0.17 ±0.01 0.24 ±0.02 D-Lactose (300 mM) 0.04 ±0.01 0.08 ±0.02 Raffinose (300 mM) 1.77 ±0.56 2.21 ±0.13 D-Mannose (300 mM) 0.01 ±0.00 n.m.§ Melibiose (300 mM) 0.67 ±0.06 0.41 ±0.03 D-Fructose (300 mM) 0.04 ±0.01 0.05 ±0.01 D-Xylose (300 mM) 0.01 ±0.00 n.m.§ D-Glucose (300 mM) n.m.§ n.m.§ Sucrose (300 mM) n.m.§ n.m.§ D-Ribose (300 mM) 0.02 ±0.00 n.m.§ L-Arabinose (300 mM) 0.08 ±0.01 0.12 ±0.02 Polyols Glycerol (300 mM) 3.22 ±0.15 5.41 ±0.58 Sorbitol (300 mM) 0.23 ±0.02 0.42 ±0.05 Galactitol (300 mM) 0.01 ±0.00 n.m.§ Mannitol (300 mM) 0.07 ±0.01 0.11 ±0.01 1,2-Propanediol (300 mM) 0.28 ±0.05 0.37±0.06 Diols 1,3-Propanediol (300 mM) 0.47 ±0.17 1.23 ±0.08 1,4-Butanediol (300 mM) 0.32 ±0.11 0.23 ±0.09 1,5-Pentanediol (300 mM) 0.02 ±0.00 0.01 ±0.00 1,6-Hexanediol (300 mM) 0.02 ±0.00 0.01 ±0.00 Aldehyde Methyl Glyoxal (5 mM) 0.03 ±0.05 0.01 ±0.00 Glycolaldehyde (5 mM) 1.40 ±0.26 1.03 ±0.30 Primary alcohols Ethanol (300 mM) 0.01 ±0.00 0.02 ±0.01 Methanol (300 mM) 0.02 ±0.00 0.03 ±0.00 1-Butanol (300 mM) 0.01 ±0.00 0.01 ±0.00 Hexanol (2.5 mM) 0.01 ±0.00 n.m.§ Octanol (2.5 mM) n.m.§ 0.02 ±0.01 Aryl alcohols Benzyl alcohol (30 mM) 0.46 ±0.16 0.40 ±0.04 Cinnamyl alcohol (2.5 mM) 0.23 ±0.05 0.39 ±0.04 4-methoxybenzyl alcohol (2.5 mM) 0.04 ±0.01 0.06 ±0.04 3-methoxybenzyl alcohol (2.5 mM) 0.15 ±0.06 0.23 ±0.02 Coniferyl alcohol (5 mM) 0.01 ±0.00 0.01 ±0.01 Veratryl alcohol (2.5 mM) 0.13 ±0.00 0.14 ±0.03 4-hydroxybenzyl alcohol (2.5 mM) 0.02 ±0.01 0.02 ±0.02 Vanillyl alcohol (2.5 mM) n.m.§ 0.03 ±0.01 Furans HMF (2.5 mM) 0.12 ±0.02 0.03 ±0.01 HMFCA (2.5 mM) 0.07 ±0.01 0.01 ±0.01 DFF (2.5 mM) 0.01 ±0.01 0.04 ±0.00 FFCA (2.5 mM) n.m.§ n.m.§ Furfural (2.5 mM) n.m.§ 0.05 ±0.03 217 *Measurements were performed in triplicate at RT in 100 mM buffer in different buffers at different pH using the HRP/ABTS plate assay. Activities were monitored using concentrations indicated within parentheses for each substrate. §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 1.12 nmole of purified UmaRafOx enzyme and 0.23 nmole of purified PhuRafOx enzyme. 218 Table B8. Initial activity screens* of AflAlcOx and PruAlcOx Substrate Specific Activity (μmol min-1 mg-1) AflAlcOx PruAlcOx Carbohydrates D-Galactose (300 mM) 65.0 ±1.41 79.9 ±29.4 D-Lactose (300 mM) 7.07 ±0.37 12.2 ±2.12 Raffinose (300 mM) 12.0 ±1.36 13.5 ±3.73 D-Mannose (300 mM) 0.14 ±0.09 2.87 ±0.76 Melibiose (300 mM) 17.4 ±1.39 8.41 ±2.32 D-Fructose (300 mM) 2.64 ±0.01 9.10 ±3.36 D-Xylose (300 mM) 3.34 ±1.63 3.70 ±1.05 D-Glucose (300 mM) 0.07 ±0.01 0.29 ±0.04 Sucrose (300 mM) 0.28 ±0.02 n.m.§ D-Ribose (300 mM) 0.45 ±0.12 4.67 ±1.78 L-Arabinose (300 mM) 1.95 ±0.55 3.20 ±0.13 Polyols Glycerol (300 mM) 35.7 ±1.50 119 ±12.7 Sorbitol (300 mM) 1.46 ±0.09 9.50 ±2.18 Galactitol (300 mM) 3.10 ±0.04 3.44 ±0.36 Diols 1,2-Propanediol (300 mM) 14.5 ±0.21 38.3 ±2.38 1,3-Propanediol (300 mM) 9.51 ±0.24 34.8 ±2.71 1,4-Butanediol (300 mM) 30.9 ±6.72 18.8 ±2.43 1,5-Pentanediol (300 mM) 25.0 ±0.67 17.8 ±1.04 1,6-Hexanediol (300 mM) 25.0 ±0.25 14.0 ±3.01 Aldehyde Methyl Glyoxal (5 mM) 2.47 ±0.26 4.57 ±1.92 Primary alcohols Ethanol (300 mM) 1.51 ±0.09 3.50 ±0.21 Methanol (300 mM) 1.22 ±0.04 2.54 ±0.46 1-Butanol (300 mM) 25.0 ±2.27 12.9 ±2.08 Hexanol (2.5 mM) 0.46 ±0.11 0.95 ±0.22 Octanol (2.5 mM) n.m.§ 0.07 ±0.02 Aryl alcohols Benzyl alcohol (30 mM) 248 ±16.6 61.5 ±5.43 Cinnamyl alcohol (2.5 mM) 22.4 ±1.02 26.2 ±2.57 4-methoxybenzyl alcohol (2.5 mM) 12.1 ±0.53 8.56 ±1.30 3-methoxybenzyl alcohol (2.5 mM) 45.5 ±2.82 23.9 ±2.85 Coniferyl alcohol (5 mM) n.m.§ 4.57 ±1.92 Veratryl alcohol (2.5 mM) 9.80 ±0.32 2.61 ±0.38 4-hydroxybenzyl alcohol (2.5 mM) 2.48 ±1.16 2.51 ±3.70 Vanillyl alcohol (2.5 mM) n.m.§ n.m.§ Furans HMF (2.5 mM) 52.8 ±3.92 28.7 ±11.2 HMFCA (2.5 mM) 8.58 ±0.89 21.7 ±2.17 DFF (2.5 mM) 0.16 ±0.01 0.46 ±0.03 FFCA (2.5 mM) n.m.§ 0.03 ±0.02 Furfural (2.5 mM) NA NA *Measurements were performed in triplicate at RT in 100 mM buffer in different buffers at different pH using the HRP/ABTS plate assay. Activities were monitored using concentrations indicated within parentheses for each substrate. §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 0.2 nmole of purified AflAlcOx enzyme and 1.0 nmole of purified PruAlcOx enzyme. 219 Table B9. Initial activity screens* of FoxAlcOx and AsyAlcOx Substrate Specific Activity (μmol min-1 mg-1) FoxAlcOx AsyAlcOx Carbohydrates D-Galactose (300 mM) 0.83 ±0.22 0.33 ±0.11 D-Lactose (300 mM) 0.11 ±0.01 0.25 ±0.12 Raffinose (300 mM) 0.08 ±0.02 0.02 ±0.03 D-Mannose (300 mM) 0.26 ±0.03 0.01 ±0.01 Melibiose (300 mM) 0.03 ±0.01 n.m.§ D-Fructose (300 mM) 0.35 ±0.07 0.76 ±0.29 D-Xylose (300 mM) n.m.§ n.m.§ D-Glucose (300 mM) 0.05 ±0.01 0.01 ±0.01 Sucrose (300 mM) 0.02 ±0.00 n.m.§ D-Ribose (300 mM) n.m.§ n.m.§ L-Arabinose (300 mM) 0.01 ±0.00 n.m.§ Polyols Glycerol (300 mM) 0.15 ±0.03 5.47 ±2.24 Sorbitol (300 mM) 0.01 ±0.00 4.56 ±0.29 Galactitol (300 mM) 0.01 ±0.00 0.76 ±0.02 Mannitol (300 mM) 0.01 ±0.00 3.66 ±0.56 Diols 1,2-Propanediol (300 mM) 0.05 ±0.00 1.04 ±0.16 1,3-Propanediol (300 mM) 0.06 ±0.01 1.83 ±0.42 1,4-Butanediol (300 mM) 0.16 ±0.08 1.60 ±0.44 1,5-Pentanediol (300 mM) 0.06 ±0.00 3.98 ±1.22 1,6-Hexanediol (300 mM) 0.03 ±0.00 7.41 ±1.15 Aldehyde Methyl Glyoxal (5 mM) 0.81 ±0.04 0.09 ±0.16 Glycolaldehyde Dimer (5 mM) 0.69 ±0.27 NA Primary alcohols Ethanol (300 mM) 0.02 ±0.00 0.44 ±0.10 Methanol (300 mM) 0.01 ±0.00 0.01 ±0.01 1-Butanol (300 mM) 0.02 ±0.00 1.76 ±0.32 Hexanol (2.5 mM) 0.01 ±0.00 0.12 ±0.11 Octanol (2.5 mM) n.m.§ n.m.§ Aryl alcohols Benzyl alcohol (30 mM) 1.85 ±0.37 11.6 ±5.83 Cinnamyl alcohol (2.5 mM) 0.10 ±0.06 3.24 ±0.73 4-methoxybenzyl alcohol (2.5 mM) 0.25 ±0.03 3.13 ±1.18 3-methoxybenzyl alcohol (2.5 mM) 0.10 ±0.02 4.20 ±0.37 Coniferyl alcohol (5 mM) n.m.§ n.m.§ Veratryl alcohol (2.5 mM) 0.21 ±0.01 1.22 ±0.19 4-hydroxybenzyl alcohol (2.5 mM) 0.02 ±0.00 n.m.§ Vanillyl alcohol (2.5 mM) n.m.§ n.m.§ Furans HMF (2.5 mM) 0.09 ±0.00 0.45 ±0.04 HMFCA (2.5 mM) 0.09 ±0.01 0.27 ±0.03 DFF (2.5 mM) 0.05 ±0.01 n.m.§ FFCA (2.5 mM) 0.01 ±0.00 n.m.§ Furfural (2.5 mM) n.m.§ n.m.§ 220 *Measurements were performed in triplicate at RT in 100 mM buffer in different buffers at different pH using the HRP/ABTS plate assay. Activities were monitored using concentrations indicated within parentheses for each substrate. §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 0.20 nmole of purified FoxAlcOx enzyme and 0.20 nmole of purified AsyAlcOx enzyme. 221 Table B10. Initial activity screens* of PorAlcOx and GciAlcOx Substrate Specific Activity (μmol min-1 mg-1) PorAlcOx GciAlcOx Carbohydrates D-Galactose (300 mM) 1.71 ±0.89 0.82 ±0.04 D-Lactose (300 mM) 1.16 ±0.90 0.40 ±0.02 Raffinose (300 mM) 3.00 ±0.65 61.21 ±7.27 D-Mannose (300 mM) 2.95 ±0.99 n.m.§ Melibiose (300 mM) 0.25 ±0.18 1.18 ±0.09 D-Fructose (300 mM) 3.14 ±0.90 6.15 ±1.78 D-Xylose (300 mM) 1.03 ±0.52 0.68 ±0.04 D-Glucose (300 mM) 3.04 ±0.15 0.44 ±0.03 Sucrose (300 mM) 0.29 ±0.36 0.81 ±0.04 D-Ribose (300 mM) 0.66 ±0.42 1.37 ±0.06 L-Arabinose (300 mM) 0.52 ±0.17 0.61 ±0.03 Polyols Glycerol (300 mM) 9.76 ±1.27 318 ±17.3 Sorbitol (300 mM) 10.2 ±1.38 207 ±9.06 Galactitol (300 mM) 3.62 ±0.84 80.9 ±1.86 Mannitol (300 mM) 2.69 ±0.55 NA Diols 1,2-Propanediol (300 mM) 11.2 ±1.69 393 ±11.7 1,3-Propanediol (300 mM) 11.8 ±0.74 576 ±31.0 1,4-Butanediol (300 mM) 12.0 ±0.18 698 ±42.3 1,5-Pentanediol (300 mM) 10.9 ±0.58 705 ±135 1,6-Hexanediol (300 mM) 9.94 ±0.70 657 ±71.4 Aldehyde Methyl Glyoxal (5 mM) 2.45 ±1.45 n.m.§ Primary alcohols Ethanol (300 mM) 5.87 ±0.80 496 ±32.0 Methanol (300 mM) 2.07 ±0.90 78.6 ±3.50 1-Butanol (300 mM) 6.32 ±0.82 532 ±17.6 Hexanol (2.5 mM) 4.06 ±0.87 428 ±4.66 Octanol (2.5 mM) 3.67 ±1.48 180 ±10.6 Aryl alcohols Benzyl alcohol (30 mM) 11.5 ±0.81 569 ±31.1 Cinnamyl alcohol (2.5 mM) 5.75 ±1.58 541 ±15.6 4-methoxybenzyl alcohol (2.5 mM) 6.06 ±0.80 319 ±17.9 3-methoxybenzyl alcohol (2.5 mM) 7.46 ±0.49 402 ±0.00 Coniferyl alcohol (5 mM) n.m.§ n.m.§ Veratryl alcohol (2.5 mM) 4.49 ±0.43 160 ±11.6 4-hydroxybenzyl alcohol (2.5 mM) 1.00 ±0.15 10.2 ±1.76 Vanillyl alcohol (2.5 mM) n.m.§ n.m.§ Furans HMF (2.5 mM) 6.81 ±1.17 690 ±18.6 HMFCA (2.5 mM) 10.2 ±0.92 272 ±29.0 DFF (2.5 mM) 1.20 ±0.12 n.m.§ FFCA (2.5 mM) 1.14 ±0.09 n.m.§ Furfural (2.5 mM) n.m.§ NA *Measurements were performed in triplicate at 25 ˚C in 50 mM sodium phosphate buffer at pH 7.5 using the HRP/ABTS assay. Activities were monitored using concentrations indicated within parentheses for each substrate. 222 §No activity detected with a specific activity limit of detection of 9 x 10-4 μmol min-1 mg-1 using 0.23 nmole of purified PorAlcOx enzyme and 0.03 nmole of purified GciAlcOx enzyme. 223 Table B11. Comparison of catalytic parameters of FoxAlcOx, AflAlcOx, PruAAA5_2A (PruAlcOx), PfeGalOx, ExeGalOx, MreGalOx and FoxGalOxB with other enzymes acting on galactose* * ND not determined Enzyme Galactose References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) FoxGalOxB 24 ±1.3 81 ±1.0 3400 This work ExeGalOx 25 ±4.0 130 ±3.7 5200 This work MreGalOx 64 ±4.8 32 ±0.7 500 This work AflAlcOx 980 ± 61 400 ±10 410 This work PruAlcOx 110 ±18 220 ±8.6 2000 This work PfeGalOx 28 ±6.7 140 ±4.5 5000 This work FoxAlcOx 2000 ±650 8.3 ±1.7 4.1 This work FgrGalOx 102 ±6.4 1059 ±18.9 10400 ±680 [102] R330K 895 ±85.9 208 ±10.8 232 ±25 [102] M1 43 ±2 1376 ±35 32 000 ±1700 [99] M3 54 ±7 1800 ±400 31 ±7 [104] C383N 390 ±38 410 ±17 1100 ±150 [97] C383S 34 ±3.6 1100 ±30 32000 ±4500 [97] CgrAAO ND ND 13.1 ±0.8 [48] FgrAAO 1700 ± 150 21 ± 1.0 12 [249] FoxAAO 1600 ± 150 23 ± 1.2 14 [249] 224 Table B12. Comparison of catalytic parameters of PorAlcOx, GciAlcOx, FoxAlcOx, AflAlcOx, PruAA5_2A (PruAlcOx), PfeGalOx, UmaRafOx, PhuRafOx, AsyAlcOx with other enzymes acting on benzyl alcohol, glycerol, butanol and 1,4-butanediol* Enzyme Benzyl alcohol Glycerol References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) PorAlcOx 8.6 ±0.15 1.3 ± 0.10 6600 8.5 ±0.32 57 ± 8.1 1500 This work GciAlcOx 350 ±20 3.6 ±0.8 97000 NA NA NA This work FoxAlcOx 230 ±70 24 ±4.5 100 NA NA NA This work AflAlcOx 410 ±11 19 ± 1.9 21000 NA NA NA This work PruAlcOx 110 ±5.0 9.2 ±1.8 12000 170 ±3.0 34 ±1.7 4900 This work PfeGalOx NA NA NA 140 ±1.1 69 ±1.6 2000 This work UmaRafOx NA NA NA 2900 ±720 130 ±24 45 This work PhuRafOx NA NA NA 2900 ±390 56 ±3.8 19 This work AsyAlcOx 79 ±22 17 ±1.8 220 700 ±80 38 ±2.0 54 This work FgrGalOx ND ND 424 ±2 NA NA NA [213] CgrAlcOx 0.69 ±0.04 94 ±1 140000 104 ± 4 96 ± 2 920 [46] CgrAAO 27 ±0.9 54.5 ±0.6 2020 ± 70 605 ± 34 58.7 ± 0.9 97 ± 5.6 [48] FgrAAO 86 ± 12 37 ± 2.8 430 2200 ± 360 17 ± 1.2 7.7 [249] FoxAAO 30 ± 19 31 ± 4.2 1000 2700 ± 403 24 ± 1.6 8.8 [249] Enzyme Butanol 1,4-Butanediol References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) KM (mM) kcat (s-1) kcat/KM (M-1.s-1) PorAlcOx 7.9 ±0.09 4.6 ± 0.30 1700 9.5 ±0.51 4.3 ± 0.56 2200 This work GciAlcOx NA NA NA 330 ±13 3.9 ±0.4 85000 This work CgrAAO 0.68 ± 0.04 96 ± 1 140000 1.2 ± 0.04 97 ± 1 83000 [48] * NA not assessed; ND not determined 225 Table B13. Comparison of catalytic parameters of GciAlcOx and AflAlcOx with other enzymes acting on HMF Enzyme HMF References KM (mM) kcat (s-1) kcat/KM (M-1.s-1) GciAlcOx 0.09 ±0.01 240 ±16 2700000 This work AflAlcOx 190 ± 20 5.0 ± 1.3 38000 This work CgrAAO 6.5 ±0.3 126 ± 1.5 19400 ± 90 [48] FgrAAO 14 ± 2.6 29 ± 1.7 2100 [249] FoxAAO 17 ± 4.9 26 ± 2.6 1500 [249] Bacterial HMFO 1.4 9.9 7100 [38] PerAAO 1.6 ± 0.2 0.33 ± 0.01 220 ±42 [218] MtGLOX 20.2 ± 9.0 15.9 982 [216] PciGLOX1 15.66 ± 2.35 1.59 ± 0.12 101.66 ± 0.01 [217] PciGLOX2 5.87 ± 2.04 0.56 ± 0.09 96.04 ±0.01 [217] PciGLOX3 6.35 ± 1.32 0.75 ± 0.07 118.35 ± 0.01 [217] 226 B.2 Supporting Figures 227 Figure B1: (A-B) Active site residues of FgrGalOx involved in catalysis and/or substrate binding investigated in this study. (A) Surface representation (B) Sticks representation (C-J) Sequence similarity networks at an alignment score cut-off of 10-550 of 623 catalytic modules from the AA5_2 subfamily with their corresponding amino acid variability for a specific position in FgrGalOx. For each panel, predicted native signal peptides and additional N-terminal modules have been removed. Each node is coloured according to its variability assessed by inspecting the multiple sequence alignment used to generate the phylogenetic tree and the SSNs in Figure 3.1. Figure B2. SDS-PAGE of recombinantly produced enzyme in P. pastoris and N-deglycosilation study. Aliquot of purified enzymes were N-glycosylated under denaturing conditions with pNGaseF. A. 1: pNGase, 2: native UmaRafOx (2.5 μg), 3: UmaRafOx (2.5μg) + pNGase, 4: native PhuRafOx, 5: PhuRafOx (2.5μg) + pNGase, 6: native AlfAlcOx (2.5 μg), 7: AlfAlcOx (2.5 μg) + pNGase, 8: native PruAA5_2A (PruAlcOx) (2.5 μg), 9: PruAA5_2A (PruAlcOx) (2.5 μg) + pNGase. B. 1: native ExeGalOx (2.5 μg), 2: ExeGalOx (2.5 μg) + pNGase, 3: native MreGalOx (2.5 μg), 4: MreGalOx (2.5 μg) + pNGase. C. 1: native GciAlcOx (2.5 μg), 2: GciAlcOx (2.5 μg) + pNGase. D. 1: pNGaseF, 2: native PorAlcOx (2.5 μg), 3: PorAlcOx (2.5 μg) + pNGase. E. 1: pNGase, 2: native FoxAlcOx (2.5 μg), 3: FoxAlcOx (2.5 μg) + pNGase. F. 1: native FoxGalOxB (2.5 μg), 2: FoxGalOxB (2.5 μg) + pNGase. G. 1: pNGase, 2: native PfeGalOx (2.5 μg), 3: PfeGalOx (2.5 μg) + pNGase, H. 1: pNGase, 2: native AsyAlcOx (2.5 μg), 3: AsyAlcOx (2.5 μg) + pNGase. I. 1: native StoAA5 (2.5 μg), 2: StoAA5 (2.5 μg) + pNGAse, 3: native CcaAA5 (2.5 μg), 4: CcaAA5 (2.5 μg) + pNGase. All gels were stained by Coomassie blue. MW = molecular weight markers, as indicated. 228 Figure B3. ABTS radical standard curve. A. Carry UV-VIS. B. BioTek Epoch microplate spectrophotometer. 229 Figure B4. Proof-of-concept experiment for medium through-put HRP-ABST plate assays using AflAlcOx. A. pH profiles. B. Substrate activity screens. C. Temperature stability assay. Left panels correspond to experiments performed using Cary 60 UV-VIS spectrometer while the right panel corresponds to the experiments performed using the BioTek Epoch microplate spectrophotometer. 230 231 Figure B5. pH-rate profiles. pH-rate profiles were determined using the coupled HRP-ABTS assay with either 300 mM galactose, 30 mM benzyl alcohol and 300 mM glycerol as the substrate at each pH value. Panels are labelled by corresponding enzyme. Black squares for phosphate citrate buffer, red circles for sodium phosphate buffer and blue triangles for glycine NaOH buffer. Error bars represent standard deviations over three replicates. 232 233 234 Figure B6. Temperature stability. Activity values were determined using the coupled HRP-ABTS assay with either 300 mM galactose, 30 mM benzyl alcohol and 300 mM glycerol as the substrate. Panels are labelled by corresponding enzyme. The enzyme was pre-incubated at each temperature, maintained by a gradient thermocycler: 30 °C (black square), 39 °C (red circle), 49 °C (blue triangle), 60 °C (green triangle), 69 °C (purple diamond) and 79 °C (yellow side triangle). The reactions were performed at RT. Error bars represent standard deviations over duplicate measurements. Graphs in the left column contain the full data over 48 hours while on the right column is a zoomed in graph showing the first 500 min. 235 Figure B7. ExeGalOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 236 Figure B8. MreGalOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Melibiose that did not display saturation kinetics due to substrate inhibition, therefore kcat/KM values were obtained from the slope of linear fittings. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 237 Figure B9. FoxGalOxB kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Melibiose that did not display saturation kinetics due to substrate inhibition, therefore kcat/KM values were obtained from the slope of linear fittings. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 238 Figure B10. PfeGalOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 239 Figure B11. UmaRafOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 240 Figure B12. PhuRafOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 241 Figure B13. AflAlcOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 242 Figure B14. PruAA5_2A/AlcOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 243 Figure B15. FoxAlcOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 244 Figure B16. PorAlcOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 245 Figure B17. GciAlcOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 246 Figure B18. AsyAlcOx kinetics. A. Specific activities. Measurements were performed in triplicate at RT in 50 mM buffer using the HRP/ABTS assay. Activities were monitored using 300 mM for carbohydrates, polyols, diols and primary alcohols, 30 mM for benzyl alcohol and galactitol, 5 mM for methyl glyoxal and glycolaldehyde dimer, 2.5 mM for aryl alcohols and furans. B. Michaelis-Menten kinetics. Initial-rate values were measured in triplicate at each substrate concentration. Error bars represent the standard deviation from the mean values. The individual kcat and KM values were determined by performing a non-linear fitting analysis of the standard Michaelis-Menten equation to the data using OriginLab 9.55. Individual substrates are indicated in the x-axis labels and the enzyme under study is annotated in each graph. 247 Figure B19. 1H-NMR analysis of the oxidation of HMF, HMFCA, DFF and FFCA by GciAlcOx. A.HMF (magenta); B. HMFCA (black). C. DFF (red) and D. FFCA (blue). 1H NMR spectra (600 MHz, 12:1 D2O: buffer, 50 mM) of reaction product profiles after 17 h incubation with GciAlcOx in the presence of catalase and HRP with 10 mM substrate. Reactions with HMF, HMFCA and DFF showed full consumption of substrate to produce FFCA ( ) and FDCA ( ). Reactions with FFCA ( ) produced partial conversion to FDCA ( ). 248 Figure B20. 1H-NMR analysis of the oxidation of HMF, HMFCA, DFF and FFCA by PorAlcOx. A.HMF (black); B. HMFCA (blue). C. DFF (red) and D. FFCA (pink). 1H NMR spectra (600 MHz, 12:1 D2O: buffer, 50 mM) of reaction product profiles after 17 h incubation with PorAlcOx in the presence of catalase and HRP with 10 mM substrate. Reactions with HMF ( ) showed 51% consumption of substrate to DFF ( ) and its hydrate form DFFhyd( ) and FFCA ( ). Reactions with HMFCA and DFF showed full consumption of substrate to produce FFCA ( ) and FDCA ( ). Reactions with FFCA ( ) produced partial conversion to FDCA ( ). 249 Figure B21. 1H-NMR analysis of the oxidation of HMF, HMFCA, DFF and FFCA by PruAA5_2A (PruAlcOx). A.HMF (black); B. HMFCA (blue). C. DFF (red) and D. FFCA (pink). 1H NMR spectra (600 MHz, 12:1 D2O: buffer, 50 mM) of reaction product profiles after 17 h incubation with PruAA5_2A/AlcOx in the presence of catalase and HRP with 10 mM substrate. Reactions with HMF ( ) showed 100% consumption of substrate to DFF ( ) and its hydrate form DFFhyd( ) and FFCA ( ). Reactions with DFF showed partial conversion to FFCA ( ). Reactions with HMFCA shower 100% conversion to FFCA ( ) while reactions with FFCA ( ) shower low conversion to FDCA ( ). 250 Figure B22. 1H-NMR analysis of the oxidation of HMF, HMFCA, DFF and FFCA by AflAlcOx. A.HMF (black); B. HMFCA (blue). C. DFF (red) and D. FFCA (pink). 1H NMR spectra (600 MHz, 12:1 D2O: buffer, 50 mM) of reaction product profiles after 17 h incubation with AflAlcOx in the presence of catalase and HRP with 10 mM substrate. Reactions with HMF ( ) showed 100% consumption of substrate to DFF ( ) and its hydrate form DFFhyd( ) and FFCA ( ). Reactions with DFF showed partial conversion to FFCA ( ). Reactions with HMFCA shower 100% conversion to FFCA ( ) while reactions with FFCA ( ) shower no conversion. 251 Figure B23. 1H-NMR analysis of the oxidation of HMF, HMFCA, DFF and FFCA by FoxGalOxB. A.HMF (black); B. HMFCA (blue). C. DFF (red) and D. FFCA (pink). 1H NMR spectra (600 MHz, 12:1 D2O: buffer, 50 mM) of reaction product profiles after 17 h incubation with FoxGalOxB in the presence of catalase and HRP with 10 mM substrate. Reactions with HMF ( ) showed 62% consumption of substrate to DFF ( ) and its hydrate form DFFhyd( ) and FFCA ( ). Reactions with HMFCA and DFF showed 17% and 12%, respectively, consumption of substrate to produce FFCA ( ). Reactions with FFCA ( ) produced partial conversion to FDCA ( ). 252 253 Figure B24. Stereochemistry determination of glycerol oxidation by PorAlcOx, GciAlcOx, PruAlcOx, AflAlcOx, PfeGalOx, FoxGalOxB, AsyAlcOx, UmaRafOx and PhuRafOx. A. Chromatogram of L/D-glyceraldehyde-hydrazone. B. Glyceraldehyde-hydrazone composition after PorAlcOx oxidation of glycerol. C. Glyceraldehyde-hydrazone composition after GciAlcOx oxidation of glycerol. D. Glyceraldehyde-hydrazone composition after PruAA5_2A (PruAlcOx) oxidation of glycerol. E. Glyceraldehyde-hydrazone composition after AflAlcOx oxidation of glycerol. F. Glyceraldehyde-hydrazone composition after PfeGalOx oxidation of glycerol. G. Glyceraldehyde-hydrazone composition after FoxGalOxB oxidation of glycerol. H. Glyceraldehyde-hydrazone composition after AsyAlcOx oxidation of glycerol. I. Glyceraldehyde-hydrazone composition after UmaRafOx oxidation of glycerol. J. Glyceraldehyde-hydrazone composition after PhuRafOx oxidation of glycerol. Peaks are labelled with retention times. 254 Figure B25. Molecular docking of galactose (cyan) in FgrGalOx (PDB ID 1GOF)56 (A) and a Phyre2 homology model of ExeGalOx (B) using AutoDock Vina as implemented in Chimera. Copper (dark orange), W290 (yellow), Y329/F329 (green), and R330 (magenta). Ligand coordination is indicated, specifying the distances (black). 255 Figure B26. Molecular docking of galactose (cyan) in FgrGalOx (PDB ID 1GOF)56 (A) and a Phyre2 homology model of ExeGalOx (B), FoxAlcOx (C) and AflAlcOx (D) using AutoDock Vina as implemented in Chimera. Copper (dark orange) and R330/K330/H330 (magenta). Ligand coordination is indicated, specifying the distances (black). 256 Figure B27. Molecular docking of galactose (cyan) in FgrGalOx (PDB ID 1GOF)56 (A) and a Phyre2 homology model of PfeGalOx (B), PruAlcOx (C), AflAlcOx (D) and FoxGalOxB (E) using AutoDock Vina as implemented in Chimera. Copper (dark orange), W290 (yellow), Y329 (green), R330/H330 (magenta), F494 (brown) and F194 (red). Ligand coordination is indicated, specifying the distances (black). 257 Appendix C : Supporting Information for Chapter 4: Cross-linked Enzyme Aggregates of a Copper Radical Alcohol Oxidase for Increased Potential in Biocatalysis C.1 Supporting Tables Table C1. Effect of precipitant agents on CgrAlcOx activity Precipitant CgrAlcOx Residual Activity (%) Buffer 100 Methanol 10 Ethanol 46 1-Propanol 104 2-Propanol 122 tert-Butanol 99 Acetone 93 Acetonitrile 89 Sat. (NH4)2SO4 125 DMF 1 DMSO 1 C.2 Supporting Figures Figure C1. Effect of glutaraldehyde concentration on CgrAlcOx activity. Measurements were performed in triplicate at 25 °C in 50 mM sodium phosphate buffer pH 8.0 using the HRP/ABTS assay. Activities were monitored using 3 mM benzyl alcohol and reactions were started with the addition of 5.5 pmol of purified CgrAlcOx preincubated with 0 – 100 mM glutaraldehyde. 258 Figure C2. Effect of shaking and cross-linker concentration on CLEA/combi-CLEA activity. CgrAlcOx CLEA (0.8 nmol CgrAlcOx plus 3.3 nmol HRP and 25 nmol of soluble catalase, black square and red circle) or combi-CLEA (0.8 nmol CgrAlcOx cross-linked with 3.3 nmol HRP and 25 nmol catalase, blue triangle and yellow triangle) prepared with varying concentrations of glutaraldehyde cross-linker were incubated with 1 mM benzyl alcohol for 3 hours. Half the CLEA reactions were shaken at 250 rpm and performed in triplicate (black circle and red triangle), while others were not shaken and performed once (red circle and yellow triangle). 259 Figure C3. Influence of CLEA preparation protocols on benzyl alcohol. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx prepared with varying concentrations of glutaraldehyde as cross-linker were incubated with 1 mM benzyl alcohol for 5 hours. Half the CLEA particles were lyophilized (blue and yellow triangle) and half the CLEA reactions were shaken at 250 rpm (red circle and yellow triangle). All experimental reactions were run in triplicate and benzaldehyde production was quantified by HPLC. 260 Figure C4. Influence of two-enzyme combi-CLEA preparations on benzyl alcohol conversion. CLEA particles were prepared with combinations of 0.8 nmol CgrAlcOx, 3.3 nmol HRP and 25 nmol catalase. Reaction with combi-CLEA made of CgrAlcOx and catalase (blue), made of CgrAlcOx and HRP (purple) and made of catalase and HRP (green) were incubated with 1 mM benzyl alcohol in water (dark colour) or in 50 mM sodium phosphate buffer, pH 7.0 (light colour) for 10 mins. Reactions were run using combi-CLEA with the addition of the third enzyme (solid bars) and control reactions without any added soluble enzyme (dashed bars) were also performed. Reactions with CgrAlcOx CLEA (0.8 nmol CgrAlcOx plus 3.3 nmol HRP and 25 nmol of soluble catalase, black and grey) and with combi-CLEA (0.8 nmol CgrAlcOx cross-linked with 3.3 nmol HRP and 25 nmol catalase, red) were also performed. All experimental reactions were run in duplicate while controls were performed once. 261 Figure C5. Reusability of CLEA and combi-CLEA. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 3.3 nmol HRP and 25 nmol of soluble catalase (black and grey dashed bars) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 3.3 nmol HRP and 25 nmol catalase (dark red and light red dashed bars) prepared with 1 mM glutaraldehyde cross-linker were incubated with 1 mM benzyl alcohol in either water (black and dark red) or 50 mM sodium phosphate buffer, pH 7.0 (grey and light red) for 1 hour. Control reactions with all enzymes not cross-linked but with either only CgrAlcOx precipitated (black and grey solid columns) or all three enzymes precipitated (dark red and light red solid columns) were also performed with 1 mM benzyl alcohol. After 1 hour, all reactions were centrifuged, supernatant removed and another 1 mL of 1 mM benzyl alcohol was incubated for another 60 min (cycle 2). All experimental reactions were run in duplicate while controls were performed once. For the control reactions, enzymes in square brackets were precipitated. 262 Figure C6. Substrate loading effect on CLEA/combi-CLEA percent conversion. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 3.3 nmol HRP and 25 nmol of soluble catalase (black and grey dashed bars) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 3.3 nmol HRP and 25 nmol catalase (dark red and light red dashed bars) prepared with 1 mM glutaraldehyde cross-linker were incubated with 1 mM, 10 mM and 30 mM benzyl alcohol in either water (black and dark red) or 50 mM sodium phosphate buffer, pH 7.0 (grey and light red) for 10 min. Control reactions with all enzymes not cross-linked but with either only CgrAlcOx precipitated (black and grey solid columns) or all three enzymes precipitated (dark red and light red solid columns) were also performed. All experimental reactions with 1 mM benzyl alcohol and combi-CLEA with 10 mM and 30 mM benzyl alcohol were run in duplicate while others were performed once. For the control reactions, enzymes in square brackets were precipitated. 263 Figure C7. Influence of pH and ferricyanide concentration on CLEA / combi-CLEA percent conversion of 1 mM benzyl alcohol. A. Reactions were performed in 50 mM sodium phosphate buffer at pH 6; B. at pH 7.0; C. at pH 8.0 or in D. water. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase (solid lines) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 25 nmol catalase (dashed lines), were incubated with 1 mM benzyl alcohol with either 1 mM (pink, light blue, light purple, light green) or 10 mM (red, dark blue, dark purple, dark green) K3Fe(CN)6 was present. Benzaldehyde production was quantified by HPLC after 10 min and 30 min. All experiments were performed in duplicate. 264 Figure C8. Effect of ferricyanide concentration in water and buffer on CLEA / combi-CLEA percent conversion of 10 mM benzyl alcohol. Reactions were performed with either A. 1 mM K3Fe(CN)6 monitored at 10 min and 30 min B. 10 mM K3Fe(CN)6 monitored at 10 min and 30 min and C. 1 mM (blue), 10 mM (green) and 100 mM (purple) K3Fe(CN)6 at 30 min. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase (solid lines and bars) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 25 nmol catalase (dashed lines and bars), were incubated with 10 mM benzyl alcohol in either 50 mM sodium phosphate buffer pH 7.0 (dark blue, dark green) or water (light blue and light green). Benzaldehyde production was quantified by HPLC. All experiments were performed in duplicate. 265 Figure C9. Effect of ferricyanide and enzyme on CLEA / combi-CLEA percent conversion of 10 mM benzyl alcohol. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase (solid bars) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 25 nmol catalase (dashed bars), were incubated with 10 mM benzyl alcohol in either 50 mM sodium phosphate buffer pH 7.0 (light and dark blue) or water (grey and black) with either 10 mM (gray and light blue) or 100 mM (black and dark blue) K3Fe(CN)6 for 4 hours. Benzaldehyde production was quantified by HPLC. All experiments were performed in duplicate. 266 Figure C10. Effect of ferricyanide concentration in water and buffer on CLEA / combi-CLEA percent conversion of 30 mM benzyl alcohol over time. CgrAlcOx CLEA made of 0.8 nmol CgrAlcOx plus 25 nmol of soluble catalase (solid bars) or combi-CLEA made of 0.8 nmol CgrAlcOx cross-linked with 25 nmol catalase (dashed bars), were incubated with 30 mM benzyl alcohol in either 50 mM sodium phosphate buffer pH 7.0 (light and dark blue) or water (grey and black) with either 30 mM (gray and light blue) or 300 mM (black and dark blue) K3Fe(CN)6 for 30 min or 17 hours. Benzaldehyde production was quantified by HPLC. All experiments were performed in duplicate. 267 Figure C11. Investigation of product inhibition on CgrAlcOx activity in solution. CgrAlcOx (0.8 nmol), catalase (25 nmol) and K3Fe(CN)6 (5 mM (black square and blue triangle) or 50 mM (red circle or purple diamond)) were added to 5 mM benzyl alcohol in 50 mM sodium phosphate buffer, pH 7.0 and percent conversion was monitored over time by HPLC. 5 mM benzaldehyde (PhCHO) was added to trials at time 0 (black square and red circle). 268 Figure C12. Effect of ferricyanide concentration in water and buffer on CgrAlcOx activity in solution. CgrAlcOx (0.8 nmol), catalase (25 nmol) and K3Fe(CN)6 (10 mM (black square and red circle) or 100 mM (blue triangle or purple diamond) were added to 10 mM benzyl alcohol in either water (black square and blue triangle) or in 50 mM sodium phosphate buffer, pH 7.0 (red circle and purple diamond). and percent conversion was monitored over time by HPLC.
Thesis/Dissertation
eng
Chemistry
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Brumer, Harry
Investigating catalytic diversity in copper radical oxidases for potential bioproduct applications
Text