Science, Faculty of
DSpace
UBCV
Macdonald, Spencer
Doctor of Philosophy - PhD
University of British Columbia
Carbohydrates are abundant throughout nature, play key roles in diverse biological processes and form fundamental components of a wide range of functional materials. Their inherent complexity is a double edge sword in that it allows carbohydrates to adopt a diverse range of structures and functions, but in an industrial setting, that complexity can also make them especially difficult and costly to synthesize in a strictly defined manner. To help ensure uniform production, carbohydrate manufacturing processes are known to utilize Carbohydrate Active Enzymes (CAZymes), which have innate substrate specificities and conformational control. A class of CAZymes, known as glycoside phosphorylases (GPs) catalyze the reversible phosphorolysis of glycosidic bonds, releasing sugar 1-phosphates and have considerable potential as catalysts for the assembly of useful carbohydrates for products ranging from functional foods and prebiotics to novel materials. However, the substrate diversity of currently identified GPs is relatively small, limiting their practical applications. To address this limitation, I used a combined approach drawing upon rational, metagenomic and genomic exploration methods to discover new GPs. First, using a rational approach, I identified a new GP activity, the first reported with a β-retaining mechanism, while investigating a mechanistic oddity observed in the GH3 CAZy family. Next, I developed and deployed two metagenomic screening methodologies that targeted GP activity in the vast reservoir of uncultivated genetic diversity encoded in microbial communities inhabiting natural and engineered ecosystems. This approach yielded eight new GPs and established a screening paradigm that can be applied to an even greater range of GP activities. Lastly, a phylogenomically diverse, synthetic gene GP library was characterized, which led to the discovery of another previously undiscovered GP activity, a new biopolymer (that I named acholetin), and established a high throughput (HT) substrate specificity assay for GPs. In total, 11 new GPs were discovered, two of those representing previously unknown activities. The research presented in this dissertation will result in a better understanding of GP diversity, provide insights into how we can engineer GPs, establish a HT functional-characterization method, and provide a platform to explore genomic and metagenomic sequence space for more novel GP activities.
Discovery of New GlycosidePhosphorylases through Rational,Genomic and Metagenomic ExplorationbySpencer MacdonaldB.Sc., Wilfrid Laurier University, 2010M.Sc., The University of British Columbia, 2015A THESIS SUBMITTED IN PARTIAL FULFILLMENT OFTHE REQUIREMENTS FOR THE DEGREE OFDOCTOR OF PHILOSOPHYinThe Faculty of Graduate and Postdoctoral Studies(Genome Science and Technology)THE UNIVERSITY OF BRITISH COLUMBIA(Vancouver)July 2021© Spencer Macdonald 2021The following individuals certify that they have read, and recommend to the Faculty of Graduateand Postdoctoral Studies for acceptance, the dissertation entitled:Discovery of New Glycoside Phosphorylases throughRational, Genomic and Metagenomic ExplorationSubmitted By Spencer Macdonald in partial fulfillment of the requirements forthe degree of Doctor of Philosophyin Genome Science and TechnologyExamining Committee:Stephen G. Withers, Professor, Chemistry and Biochemistry, UBCSupervisorHarry Brummer, Professor, Michael Smith Laboratories, UBCSupervisory Committee MemberKatherine S. Ryan, Associate Professor, Chemistry, UBCUniversity ExaminerOrlando Rojas, Professor, Chemical and Biological Engineering, UBCUniversity ExaminerRobert A. Field, Professor, Chemistry, The University of ManchesterExternal ExaminerAdditional Supervisory Committee Members:Steven J. Hallam, Professor, Microbiology and Immunology, UBCSupervisory Committee MemberLindsay D. Eltis, Professor, Microbiology and Immunology, UBCSupervisory Committee MemberiiAbstractCarbohydrates are abundant throughout nature, play key roles in diverse biological processes andform fundamental components of a wide range of functional materials. Their inherent complexity isa double edge sword in that it allows carbohydrates to adopt a diverse range of structures and func-tions, but in an industrial setting, that complexity can also make them especially difficult and costlyto synthesize in a strictly defined manner. To help ensure uniform production, carbohydrate man-ufacturing processes are known to utilize Carbohydrate Active Enzymes (CAZymes), which haveinnate substrate specificities and conformational control. A class of CAZymes, known as glycosidephosphorylases (GPs) catalyze the reversible phosphorolysis of glycosidic bonds, releasing sugar1-phosphates and have considerable potential as catalysts for the assembly of useful carbohydratesfor products ranging from functional foods and prebiotics to novel materials. However, the sub-strate diversity of currently identified GPs is relatively small, limiting their practical applications.To address this limitation, I used a combined approach drawing upon rational, metagenomic andgenomic exploration methods to discover new GPs. First, using a rational approach, I identified anew GP activity, the first reported with a β-retaining mechanism, while investigating a mechanisticoddity observed in the GH3 CAZy family. Next, I developed and deployed two metagenomic screen-ing methodologies that targeted GP activity in the vast reservoir of uncultivated genetic diversityencoded in microbial communities inhabiting natural and engineered ecosystems. This approachyielded eight new GPs and established a screening paradigm that can be applied to an even greaterrange of GP activities. Lastly, a phylogenomically diverse, synthetic gene GP library was character-ized, which led to the discovery of another previously undiscovered GP activity, a new biopolymer(that I named acholetin), and established a high throughput (HT) substrate specificity assay forGPs. In total, 11 new GPs were discovered, two of those representing previously unknown activities.The research presented in this dissertation will result in a better understanding of GP diversity,iiiprovide insights into how we can engineer GPs, establish a HT functional-characterization method,and provide a platform to explore genomic and metagenomic sequence space for more novel GPactivities.ivLay SummaryGlycoside phosphorylases (GPs) are useful biocatalysts because they are practically reversible andcan be used to make desirable complex carbohydrates for a wide variety of purposes. However,only a handful of GP types are known, and therefore their applications are limited. I applied threedifferent approaches to find more GPs. The first used a rational approach to find new enzymes basedon potential phosphate possessing features. The second way involved an approach that explored themicrobial genetic diversity present in the environment. The third explored the wealth of availablescientific biological data, that has been generated over the past several decades, to identify newenzymes. From this effort I identified 11 new enzymes and developed a suite of tools that can beused for future enzyme discovery efforts, which can be applied to improve current carbohydratemanufacturing methods and help to make the next-generation of carbohydrate based materials.vPrefaceA number of sections of this thesis are partially or wholly published in press. Much of this researchwas conducted as a collaborative effort and the contributions of others are detailed below. Portionsof Chapter 2-4 are based upon previous publications but contain wording original to this thesis.These publications, for which I am an author, follow below:• Chapter 2:Macdonald, S.S., Blaukopf, M., and Withers, S.G. N -Acetylglucosaminidases from CAZyFamily GH3 Are Really Glycoside Phosphorylases, Thereby Explaining Their Use of Histidineas an Acid/Base Catalyst in Place of Glutamic Acid. J. Biol. Chem. 290, 4887-4895 (2015).◦ I conducted all experimentation except for NMR, which was performed by M. Blaukopf,and ESI-MS, which was performed by Dr. Jason Rogalski.◦ Data analysis was performed by S.G. Withers and myself with help from M. Blaukopffor NMR analysis.◦ S.G. Withers and I prepared the original draft of the manuscript and also completedreview and editing.• Chapter 3:Macdonald, S.S., Patel, A., Larmour, V.L.C., Morgan-Lang, C., Hallam, S.J., Mark, B.L.,Withers, S.G. Structural and mechanistic analysis of a β-glucoside phosphorylase identifiedby screening a metagenomic library. J. Biol. Chem. 293, 3451-3467 (2018).◦ Metagenomic libraries were generated by S.J. Hallam’s research group. BglP crystalstructures were generated by A. Patel and V.L.C. Larmour. ESI-MS was performed byAlexandra Nastasa. I conducted all other experimentation.vi◦ Bioinformatic analysis was performed by C. Morgan-Lang and myself. Structural analysiswas performed by A. Patel, V.L.C. Larmour and B.L. Mark. S.G. Withers, S.J. Hallamand I performed all other formal analysis.◦ S.G. Withers and I prepared the original draft of the manuscript, except for the X-raycrystallography sections, which were written by A. Patel and B.L. Mark. Review andediting were completed by S.G. Withers, B.L. Mark, S.J. Hallam, A. Patel and myself.• Chapter 4:Macdonald, S.S., Armstrong Z., Morgan-Lang, C., Osowiecka M., Robinson K., Hallam,S.J., Withers, S.G. Development and Application of a High-Throughput Functional Metage-nomic Screen for Glycoside Phosphorylases. Cell Chem. Biol. 26, 1001-1012.e5 (2019).◦ Metagenomic libraries were generated by Z. Armstrong and S.J. Hallam’s research group.M. Osowiecka expressed and purified metagenomic GPs. I conducted all other experi-mentation.◦ Bioinformatic analysis was performed by C. Morgan-Lang and myself. S.G. Withers andI performed all other formal analysis.◦ I prepared the original draft of the manuscript. Review and editing were completed byS.G. Withers, Z. Armstrong, K. Robinson, S.J. Hallam and myself.• Chapter 5:◦ Phylogenomic sampling, gene synthesis and GP library construction was done by theJoint Genome Institute. Synthesis of GlcNAc-β1,3-GlcNAc and Glc-β1,3-GlcNAc-α1-pNP, as well as MALDI-MS performed by Dr. Gregor Tegl. AchP crystal structureswere generated by Dr. Jose Henrique Pereira and Dr. Paul Adams. I conducted all otherexperimentation.◦ NMR analysis of GlcNAc-β1,3-GlcNAc and Glc-β1,3-GlcNAc-α1-pNP and acholetin per-formed by Dr. Feng Liu with assistance from Dr. Gregor Tegl. I performed all otherformal analysis.vii• No ethics approval was required for the research contained in this thesis.viiiContentsAbstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . iiiLay Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vPreface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . viContents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ixList of Tables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xvList of Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xviiList of Abbreviations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xxiAcknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xxviDedication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .xxvii1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.1 Carbohydrate-Active Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.2 Enzymatic Carbohydrate Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.3 Glycoside Phosphorylases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41.3.1 A Brief History of Glycoside Phosphorylases . . . . . . . . . . . . . . . . . . 61.3.2 Applications of Glycoside Phosphorylases . . . . . . . . . . . . . . . . . . . . 101.3.3 Classification of Glycoside Phosphorylases . . . . . . . . . . . . . . . . . . . 111.4 Glycoside Phosphorylase Discovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131.4.1 Experimental and Rational Exploration . . . . . . . . . . . . . . . . . . . . . 13ix1.4.2 Genomic Sequence Exploration . . . . . . . . . . . . . . . . . . . . . . . . . . 151.4.3 Functional Metagenomic Exploration . . . . . . . . . . . . . . . . . . . . . . 161.4.4 Future Outlook on Glycoside Phosphorylase Discovery . . . . . . . . . . . . . 191.5 Dissertation Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 232 Identification of a new Glycoside Phosphorylase from CAZy family GH3 . . . 252.1 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 252.2 Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262.3 Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 282.3.1 Formation of Sugar Phosphate Products . . . . . . . . . . . . . . . . . . . . 302.3.2 Determination of Kinetic Parameters . . . . . . . . . . . . . . . . . . . . . . 302.3.3 Observation of Covalent Intermediate Accumulation . . . . . . . . . . . . . . 342.3.4 Is Phosphate the Natural Nucleophile? . . . . . . . . . . . . . . . . . . . . . 352.3.5 Nag3 as a Phosphorylase and its Role . . . . . . . . . . . . . . . . . . . . . . 372.3.6 Structural and Mechanistic Implications . . . . . . . . . . . . . . . . . . . . . 382.4 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 393 Functional Metagenomic Screen Targeting Forward Phosphorolysis Activity . 413.1 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 413.2 Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 423.3 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 443.3.1 Development and Testing of the Metagenomic Screen . . . . . . . . . . . . . 443.3.2 Functional Screen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 463.3.3 Identification of Active Glycoside Phosphorylases . . . . . . . . . . . . . . . 493.3.4 Kinetic and Mechanistic Characterization of BglP . . . . . . . . . . . . . . . 513.3.5 X-ray Crystal Structure of BglP . . . . . . . . . . . . . . . . . . . . . . . . . 573.3.6 Donor Recognition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 583.3.7 Phosphate Recognition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 623.3.8 Acceptor Recognition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 643.4 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74x3.5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 774 Functional Metagenomic Screen Targeting Reverse Phosphorolysis Activity . . 794.1 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 794.2 Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 804.3 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 814.3.1 Functional Screen Validation . . . . . . . . . . . . . . . . . . . . . . . . . . . 814.3.2 Functional Screen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 834.3.3 Identification of GP activity . . . . . . . . . . . . . . . . . . . . . . . . . . . 874.3.4 Functional Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . 924.3.5 Taxonomic Classification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 974.3.6 Targeting Broader GP Activities . . . . . . . . . . . . . . . . . . . . . . . . . 994.4 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1004.5 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1035 Characterization of a Phylogenomically Diverse GP Library . . . . . . . . . . . . 1055.1 Summary and Background . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1055.2 Results and Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1065.2.1 Phylogenomic Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1065.2.2 GP Library Expression Analysis . . . . . . . . . . . . . . . . . . . . . . . . . 1065.2.3 GP Library Substrate Specificity Analysis . . . . . . . . . . . . . . . . . . . . 1095.2.4 GH65 Substrate Specificity Assay . . . . . . . . . . . . . . . . . . . . . . . . 1105.2.5 GH94 Substrate Specificity Assay . . . . . . . . . . . . . . . . . . . . . . . . 1125.2.6 GH112 Substrate Specificity Assay . . . . . . . . . . . . . . . . . . . . . . . . 1165.2.7 GH130 Substrate Specificity Assay . . . . . . . . . . . . . . . . . . . . . . . . 1195.2.8 GH13 subfamily 18 Substrate Specificity Assay . . . . . . . . . . . . . . . . . 1235.2.9 Summary of the GP Library Substrate Specificity Assay . . . . . . . . . . . . 1255.2.10 Functional Characterization of 2E04 . . . . . . . . . . . . . . . . . . . . . . . 1265.2.11 AchP Sequence Similarity Network . . . . . . . . . . . . . . . . . . . . . . . . 1265.2.12 Substrate Specificity Screen . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128xi5.2.13 Donor Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1295.2.14 Acceptor Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1305.2.15 Linkage Determination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1345.2.16 Oligomerization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1365.2.17 Core 3 Mucin-like O-Glycan Analog . . . . . . . . . . . . . . . . . . . . . . . 1395.2.18 Two-pot Large Scale Acholetin Synthesis . . . . . . . . . . . . . . . . . . . . 1395.2.19 Acholetin analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1425.2.20 Preliminary Structural Analysis . . . . . . . . . . . . . . . . . . . . . . . . . 1475.2.21 AchP and Acholetin’s Biological Role . . . . . . . . . . . . . . . . . . . . . . 1495.3 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1526 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1547 Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1577.1 General Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1577.2 Data Accessioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1577.3 Chapter 2 Experimental . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1587.3.1 Materials and Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1587.3.2 General Kinetic Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1587.3.3 Pre-steady State Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1597.3.4 1H-NMR Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1597.3.5 GlcNAc-β1-P and Glc-β1-P Isolation and Characterization . . . . . . . . . . 1597.3.6 TLC Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1607.3.7 Electrospray Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . 1607.3.8 Reactivation with Phosphate . . . . . . . . . . . . . . . . . . . . . . . . . . . 1607.4 Chapter 3 Experimental . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1617.4.1 Materials, Reagents and General Methods . . . . . . . . . . . . . . . . . . . . 1617.4.2 Plasmid Construction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1647.4.3 Protein Expression and Purification . . . . . . . . . . . . . . . . . . . . . . . 1647.4.4 HtCBP, HtCDP and Nag3 DNPGlc Phosphorolysis Assay . . . . . . . . . . . 165xii7.4.5 BCR Library Function Screen . . . . . . . . . . . . . . . . . . . . . . . . . . 1657.4.6 BCR GP Master Library Screen and Validation . . . . . . . . . . . . . . . . 1667.4.7 Full Fosmid Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1677.4.8 Contig Assembly, Open Reading Frame Prediction and Gene Identification . 1677.4.9 Kinetic Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1687.4.10 Electrospray Mass Spectrometry . . . . . . . . . . . . . . . . . . . . . . . . . 1687.4.11 Pre-Steady State Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1697.4.12 Inactivation and Reactivation of BglP . . . . . . . . . . . . . . . . . . . . . . 1697.4.13 X-Ray Crystallographic Structure Determination . . . . . . . . . . . . . . . . 1707.4.14 Reactivator Screen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1717.4.15 BglP Reactivation Product Analysis . . . . . . . . . . . . . . . . . . . . . . . 1727.5 Chapter 4 Experimental . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1737.5.1 Materials and Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1737.5.2 Metagenomic Libraries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1767.5.3 Phosphatase Inhibition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1767.5.4 Metagenomic Screen Validation . . . . . . . . . . . . . . . . . . . . . . . . . 1777.5.5 HT Metagenomic Screening Procedure . . . . . . . . . . . . . . . . . . . . . 1777.5.6 Fosmid Sequencing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1787.5.7 Sequence Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1787.5.8 ORF Prediction and Taxonomic Assignment . . . . . . . . . . . . . . . . . . 1797.5.9 Sub-Cloning Metagenomic GPs . . . . . . . . . . . . . . . . . . . . . . . . . . 1797.5.10 Heterologous Expression and Purification . . . . . . . . . . . . . . . . . . . . 1807.5.11 Functional Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . 1817.5.12 96-well Molybdenum Blue Activity Assay . . . . . . . . . . . . . . . . . . . . 1817.6 Chapter 5 Experimental . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1827.6.1 General Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1827.6.2 JGI GP Library Expression Test and Preliminary Activity Assay . . . . . . . 1827.6.3 JGI GP Library Substrate Specificity Screen . . . . . . . . . . . . . . . . . . 1837.6.4 JGI GP Library Sequence Similarity Networks . . . . . . . . . . . . . . . . . 183xiii7.6.5 Protein Expression and Purification . . . . . . . . . . . . . . . . . . . . . . . 1847.6.6 Cell Lysate Substrate Specificity Assay . . . . . . . . . . . . . . . . . . . . . 1867.6.7 Purified AchP Substrate Specificity Assay . . . . . . . . . . . . . . . . . . . . 1867.6.8 AchP Kinetics Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1877.6.9 Oligomerization analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1887.6.10 Synthesis of GlcNAc-β1,3-GlcNAc . . . . . . . . . . . . . . . . . . . . . . . . 1887.6.11 Synthesis of Glc-β1,3-GlcNAc-pNP . . . . . . . . . . . . . . . . . . . . . . . 1897.6.12 Core 3 Mucin-like O-glycan Analog Synthesis . . . . . . . . . . . . . . . . . . 1897.6.13 Acholetin Two-pot Large Scale Synthesis . . . . . . . . . . . . . . . . . . . . 190Bibliography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192AppendicesA Chapter 2 Supplemental Material . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226A.1 General Information . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226A.2 NMR Assignments of Nag3 Products . . . . . . . . . . . . . . . . . . . . . . . . . . 226A.3 Supplemental Figures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 227B Chapter 3 Supplemental Material . . . . . . . . . . . . . . . . . . . . . . . . . . . . 231C Chapter 5 Supplemental Material . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236C.1 NMR Assignments of AchP Products . . . . . . . . . . . . . . . . . . . . . . . . . . 236C.2 A. laidlawii Operon Table . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237C.3 GP Library Phylogenetic Trees . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239C.4 GP Library Expression Tests . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245xivList of Tables1.1 Summary of GPs reported in the CAZy DB arranged chronologically. . . . . . . . . . 81.2 List of GPs reported in the CAZy DB organized by CAZy family. . . . . . . . . . . . 91.3 Classification of GPs in the CAZy DB. . . . . . . . . . . . . . . . . . . . . . . . . . . 112.1 Apparent kinetic parameters for Nag3 phosphate activation. . . . . . . . . . . . . . . 332.2 Nag 3 kinetic parameters: pNPGlcNAc, pNPGlc and DNPGlc. . . . . . . . . . . . . 333.1 BglP kinetic parameters: pNPGlcNAc, pNPGlc and DNPGlc. . . . . . . . . . . . . . 533.2 BglP.Q205S kinetic parameters: pNPGlcNAc, pNPGlc and DNPGlc. . . . . . . . . . 623.3 List of substrates that failed to reactivate BglP. . . . . . . . . . . . . . . . . . . . . . 674.1 GP active ORFs identified from the molybdenum blue high-throughput metagenomicscreen. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 894.2 Amino acid percent identity between GH94 hits. . . . . . . . . . . . . . . . . . . . . 904.3 Lowest common ancestor analysis of 11 GP active fosmids. . . . . . . . . . . . . . . . 985.1 Summary of GP library expression tests. . . . . . . . . . . . . . . . . . . . . . . . . . 1085.2 GP library substrate specificity acceptor list. . . . . . . . . . . . . . . . . . . . . . . 1115.3 Known GH94 activities. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1287.1 PIPE and QuickChange PCR parameters. . . . . . . . . . . . . . . . . . . . . . . . . 1627.2 Primers and PIPE-PCR. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1627.3 Buffer Recipes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1637.4 PIPE PCR parameters. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1747.5 Partially incomplete polymerase extension (PIPE) PCR details. . . . . . . . . . . . . 174xv7.6 Molybdenum blue screen buffer recipes. . . . . . . . . . . . . . . . . . . . . . . . . . 1757.7 Protein expression and purification buffers. . . . . . . . . . . . . . . . . . . . . . . . 175B.1 BglP crystallographic statistic table. . . . . . . . . . . . . . . . . . . . . . . . . . . . 232B.2 CAZy ORFs from fosmid hits: Tabular. . . . . . . . . . . . . . . . . . . . . . . . . . 233B.3 29K06 and 31P01 annotation table. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 234C.1 A. laidlawii operon top BLAST hits. . . . . . . . . . . . . . . . . . . . . . . . . . . . 238xviList of Figures1.1 GP, GT, GH and GS general reaction scheme. . . . . . . . . . . . . . . . . . . . . . . 31.2 Hypothetical GP scheme to synthesize valuable glycans. . . . . . . . . . . . . . . . . 51.3 Metabolic efficiency advantage of GPs. . . . . . . . . . . . . . . . . . . . . . . . . . . 61.4 Catalytic mechanisms glycoside phosphorylases. . . . . . . . . . . . . . . . . . . . . . 121.5 Metagenomic library workflow. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 181.6 Example sequence similarity network (GH130). . . . . . . . . . . . . . . . . . . . . . 211.7 Alignment score vs. percent identity plot for GH130. . . . . . . . . . . . . . . . . . . 222.1 Phosphate activation of substrate cleavage by C. fimi Nag3. . . . . . . . . . . . . . . 292.2 1H NMR and TLC analysis of Nag3 products. . . . . . . . . . . . . . . . . . . . . . . 312.3 Mechanistic scheme for a retaining β-glycoside phosphorylase/glycosidase. . . . . . . 322.4 Pre-steady state burst of DNP released by Nag3. . . . . . . . . . . . . . . . . . . . . 352.5 Trapping the Nag3 covalent intermediate. . . . . . . . . . . . . . . . . . . . . . . . . 362.6 Nag3 activator screen. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372.7 Phosphate rescue of inactivated Nag3. . . . . . . . . . . . . . . . . . . . . . . . . . . 383.1 Generalized phosphorylase and hydrolase reactions. . . . . . . . . . . . . . . . . . . . 423.2 Phosphorolysis of cellobiose or cello-oligosaccharides. . . . . . . . . . . . . . . . . . . 443.3 Proposed mechanistic scheme for a retaining β-glycosidase/phosphorylase. . . . . . . 453.4 Phosphate-dependent cleavage of DNPG by HtCBP, HtCDP and Nag3. . . . . . . . 463.5 Workflow: forward phosphorolysis metagenomic screen. . . . . . . . . . . . . . . . . . 473.6 Forward phosphorolysis functional screen results. . . . . . . . . . . . . . . . . . . . . 483.7 29K06 and 31P01 fosmid ORF alignment. . . . . . . . . . . . . . . . . . . . . . . . . 49xvii3.8 Amino acid alignment of C. fimi Nag3 and BglP. . . . . . . . . . . . . . . . . . . . . 503.9 Purified BglP TLC activity assay. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 513.10 Phosphate activation of substrate cleavage by BglP. . . . . . . . . . . . . . . . . . . . 523.11 BglP pre-steady state burst phase analysis. . . . . . . . . . . . . . . . . . . . . . . . 543.12 Trapping the BglP covalent intermediate. . . . . . . . . . . . . . . . . . . . . . . . . 563.13 BglP inactivation and reactivation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 573.14 Crystal structure of BglP covalently bound to 2FGlc. . . . . . . . . . . . . . . . . . . 593.15 Phosphate activation of substrate cleavage by BglP.Q205S. . . . . . . . . . . . . . . . 613.16 Purified BglP.Q205S TLC activity assay. . . . . . . . . . . . . . . . . . . . . . . . . . 633.17 Modeling Glc1-P into the BglP active site. . . . . . . . . . . . . . . . . . . . . . . . . 643.18 Inactivation of BglP for reactivation screen. . . . . . . . . . . . . . . . . . . . . . . . 653.19 Reactivators of 2FGlc-BglP. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 663.20 BglP reactivation product analysis TLC. . . . . . . . . . . . . . . . . . . . . . . . . . 683.21 Chemical structures of BglP reactivator products. . . . . . . . . . . . . . . . . . . . . 703.22 BglP product analysis: Glc-β1-P-α1-Glc TLC and ESI-MS. . . . . . . . . . . . . . . 713.23 BglP product analysis: Glc-β1-P-α1-Glc 1H NMR. . . . . . . . . . . . . . . . . . . . 713.24 BglP product analysis: Glc-β1-N3 TLC. . . . . . . . . . . . . . . . . . . . . . . . . . 723.25 BglP product analysis: β-UMPGlc and β-UDPGlc TLC. . . . . . . . . . . . . . . . . 734.1 Molybdenum blue reaction scheme. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 814.2 Background phosphatase inhibition by molybdate. . . . . . . . . . . . . . . . . . . . 824.3 Molybdenum blue screen validation. . . . . . . . . . . . . . . . . . . . . . . . . . . . 844.4 Overview of the molybdenum blue metagenomic screen workflow. . . . . . . . . . . . 854.5 Molybdenum blue high-throughput metagenomic screen results. . . . . . . . . . . . . 864.6 Circos representation of the molybdenum blue screen fosmid hits. . . . . . . . . . . . 884.7 Amino acid sequence alignment of GH149_H, EgP1 and Pro_7066_par. . . . . . . . 914.8 Hit GP expression and lysate activity assay. . . . . . . . . . . . . . . . . . . . . . . . 934.9 Reverse and forward phosphorolysis TLC assays. . . . . . . . . . . . . . . . . . . . . 944.10 Purification and reverse phosphorolysis activity assay for GH94_F. . . . . . . . . . . 97xviii4.11 Expanded substrate compatibility assay. . . . . . . . . . . . . . . . . . . . . . . . . . 1004.12 Phosphate dependent aryl-glycoside assay. . . . . . . . . . . . . . . . . . . . . . . . . 1024.13 Hypothetical mixed linked glucan metabolism scheme. . . . . . . . . . . . . . . . . . 1045.1 GH65 acceptor specificity heat map. . . . . . . . . . . . . . . . . . . . . . . . . . . . 1105.2 GH65 sequence similarity network. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1135.3 GH94 acceptor specificity heat map. . . . . . . . . . . . . . . . . . . . . . . . . . . . 1145.4 GH94 sequence similarity network. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1155.5 GH112 acceptor specificity heat map. . . . . . . . . . . . . . . . . . . . . . . . . . . . 1175.6 GH112 sequence similarity network. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1185.7 GH130 acceptor specificity heat map. . . . . . . . . . . . . . . . . . . . . . . . . . . . 1205.8 GH130 sequence similarity network. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1215.9 GH13 subfamily 18 acceptor specificity heat map. . . . . . . . . . . . . . . . . . . . . 1235.10 GH13 subfamily 18 sequence similarity network. . . . . . . . . . . . . . . . . . . . . . 1245.11 AchP: GH94 sequence similarity network. . . . . . . . . . . . . . . . . . . . . . . . . 1275.12 E. coli cell lysate AchP donor and acceptor specificity screen. . . . . . . . . . . . . . 1295.13 Purified AchP donor and acceptor specificity screen. . . . . . . . . . . . . . . . . . . 1305.14 Reverse phosphorolysis kinetic parameters for AchP. . . . . . . . . . . . . . . . . . . 1315.15 AchP product analysis TLC. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1335.16 Reverse phosphorolysis mechanism of a β-inverting CDP contrasting Glc and Gal. . . 1345.17 GlcNAc-β1,3-GlcNAc HMBC. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1355.18 AchP reaction scheme. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1365.19 AchP degree of polymerization analysis. . . . . . . . . . . . . . . . . . . . . . . . . . 1375.20 Core 3 mucin-like O-glycan analog synthesis with AchP. . . . . . . . . . . . . . . . . 1405.21 Two-pot large scale acholetin synthesis. . . . . . . . . . . . . . . . . . . . . . . . . . . 1415.22 G25 acholetin desalting. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1435.23 Acholetin HMBC analysis. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1445.24 13C NMR of untreated acholetin compared to reducing-end reduced acholetin. . . . . 1455.25 Acholetin enzymatic degradation assay. . . . . . . . . . . . . . . . . . . . . . . . . . . 146xix5.26 Co-crystal structure of AchP and GlcNAc-β1,3-GlcNAc. . . . . . . . . . . . . . . . . 1485.27 A. laidlawii AchP operon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150A.1 1H NMR spectrum of Glc-β1-P in D2O. . . . . . . . . . . . . . . . . . . . . . . . . . 227A.2 13C NMR spectrum of Glc-β1-P in D2O. . . . . . . . . . . . . . . . . . . . . . . . . . 228A.3 1H NMR spectrum of GlcNAc-β1-P in D2O. . . . . . . . . . . . . . . . . . . . . . . . 229A.4 13C NMR spectrum of GlcNAc-β1-P in D2O. . . . . . . . . . . . . . . . . . . . . . . 230B.1 CAZy ORFs from fosmid hits: Graphical. . . . . . . . . . . . . . . . . . . . . . . . . 235C.1 GH13 annotated phylogenetic tree. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 240C.2 GH65 annotated phylogenetic tree. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241C.3 GH94 annotated phylogenetic tree. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 242C.4 GH112 annotated phylogenetic tree. . . . . . . . . . . . . . . . . . . . . . . . . . . . 243C.5 GH130 annotated phylogenetic tree. . . . . . . . . . . . . . . . . . . . . . . . . . . . 244C.6 37 °C GP library expression test. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 246C.7 25 °C GP library expression test. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 252C.8 16 °C GP library expression test. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 258C.9 Multi-angle light scattering analysis of acholetin. . . . . . . . . . . . . . . . . . . . . 264xxList of AbbreviationsAbbreviationsAA Auxiliary activitiesAchP Acholetin phosphorylaseADK Adenylate kinasAp5A P1,P5-di(adenosine-5’)pentaphosphateBCR Biochemical reactor systemBLAST Basic local alignment search toolBVR BeaverCAZy DB Carbohydrate active enzymes databaseCAZyme Carbohydrate active enzymeCBM Carbohydrate binding moduleCBP Cellobiose phosphorylaseCDP Cellodextrin phosphorylaseCE Carbohydrate esteraseChbP N,N’ -Diacetylchitobiose phosphorylaseCMP-β-KDO CMP-ketodeoxyoctonateCOSY Correlation spectroscopyCSA bis-(3’→5’)-cyclic diguanylic acidCV Column volumeDa DaltonDHB 2,5-Dihydroxybenzoic acidDNA deoxyribonucleic acidDP Degree of polymerizationdspB Dispersin BDTT DithiothreitolE-value Expect-valueeDNA Environmental DNAEFI-EST Enzyme Function Initiative-Enzyme Similarity ToolESI Electrospray ionizationFACS Fluorescence activated cell sortingxxiFPLC Fast protein liquid chromatographyGH Glycoside HydrolaseGI GastrointestinalGP Glycoside PhosphorylaseGS GlycosynthasegsChit Endochitinase from Streptomyces griseusGT GlycosyltransferaseHMBC Heteronuclear multiple bond correlationHPLC High performance liquid chromatographyHSQC Heteronuclear single quantum correlationHT High throughputIPTG Isopropyl β-D-1-thiogalactopyranosideJGI Joint Genome InstitutekDa KilodaltonLCA* Lowest common ancestorMALDI Matrix-assisted laser desorption/ionizationMALS Multiangle light scatteringMLG Mixed linked glucansMS Mass spectrometryMW Molecular weightMWCO Molecular weight cutoffNagZ N -acetylglucosaminidasesNIMS Nanostructure-initiator mass spectrometryNMR Nuclear magnetic resonanceORF Open reading framePAGE Polyacrylamide gel electrophoresisPCR Polymerase chain reactionPDB Protein Data BankPGM PhosphoglucomutasePIPE Partially incomplete polymerase extensionPL Polysaccharide lyasePNAG Poly-N -acetylglucosamineRf Retention factorRNA Ribonucleic acidrRNA Ribosomal ribonucleic acidSD Standard deviationSDS Sodium dodecyl sulfateSE Standard errorxxiispHex β-N -Acetylhexosaminidase from Streptomyces pilcatusSSN Sequence similarity networkSSU Small subunitTIM Triose phosphate isomeraseTLC Thin-layer chromatographyTOF Time of flightUV UltravioletSubstrates2FGlc-BglP 2-fluoro-glucosyl-BglPADP Adenosine diphosphateAMP Adenosine monophosphateATP Adenosine triphosphateCDP Cytidine diphosphateCMP Cytidine monophosphateCTP Cytidine triphosphateDFMUGlc 6,8-difluoromethylumbelliferyl β-glucosideDFMUGlcNAc 6,8-difluoromethylumbelliferyl β-N -acetyl glucosaminideDNP 2,4-DinitrophenolDNP2FGlc 2,4-dinitrophenyl 2-deoxy-2-fluoro-β-D-glucopyranosideDNPGlc 2,4-dinitrophenyl β-D-glucosideFru D-FructoseGal D-GalactoseGal-β1,3(4)-Gal D-GalactobioseGal-β1,4-Glc LactoseGal1-P D-Galactose 1-phosphateGalN D-GalactosamineGalNAc N -Acetyl-D-galactosamineGalNAc-α1-MU 4-Methylumbelliferyl N -acetyl-α-D-galactosamineGalNAc-α1-pNP 4-Nitrophenol N -acetyl-α-D-galactosamineGalNAc1-P N -Acetyl-D-galactosamine 1-phosphateGDP Guanosine diphosphateGlc D-GlucoseGlc-α1-P α-D-glucose 1-phosphateGlc-α1,4-Glc MaltoseGlc-β1,2-Glc SophoroseGlc-β1,3-Glc LaminaribiosexxiiiGlc-β1,4-Glc CellobioseGlc-β1,6-Glc GentiobioseGlc1-P D-Glucose 1-phosphateGlc6-P D-Glucose 6-phosphateGlcN D-GlucosamineGlcN-β1,4-GlcN ChitobioseGlcNAc N -Acetyl-D-glucosamineGlcNAc-β1,4-GlcNAc N,N’ -DiacetylchitobioseGlcNAc-1P N -Acetyl-D-glucosamine 1-phosphateGly GlycerolGMP Guanosine monophosphateGTP Guanosine triphosphateMal1-P Maltose 1-phosphateMan D-MannoseMan-β1,4-Man MannobioseMan1-P D-Mannose 1-phosphateManN D-MannosamineManNAc N -Acetyl-D-mannosamineMU 4-methylumbelliferonePLP pyridoxal 5’-phosphatepNP 4-NitrophenolpNPGlc 4-Nitrophenyl β-D-glucosidepNPGlcNAc 4-Nitrophenyl β-N -Acetyl-D-glucosamineRha L-RhamnoseUDP Uridine diphosphateUMP Uridine monophosphateUTP Uridine triphosphateXyl D-XyloseAmino acidsAla A AlanineArg R ArginineAsn N AsparagineAsp D Aspartic AcidCys C CysteineGlu E Glutamic AcidGln Q GlutaminexxivGly G GlycineHis H HistidineIle I IsoleucineLeu L LeucineLys K LysineMet M MethioninePhe F PhenylalaninePro P ProlineSer S SerineThr T ThreonineTrp W TryptophanTyr Y TyrosineVal V ValinexxvAcknowledgementsFirstly, I would like to thank my Ph.D. supervisor, Dr. Stephen Withers, for his years of support,mentorship, vision and patience. Also I would like to thank all the past and present members of theWithers lab for their friendship and support. In particular I would like to thank: Dr. HongmingChen for the many substrates he has so kindly donated over the years, which made performing muchof the research described here possible. Dr. Markus Blaukopf for his mentorship and collaborationon the Nag3 project. Dr. Feng Liu for determining countless NMR assignments, various helpfuland enlightening conversations, and collaboration on the acholetin project. Dr. Kyle Robinsonfor his friendship and many clarifying discussions and debates. Dr. Steven Hallam for his yearsof mentorship and support. Dr. Zach Armstrong, Dr. David Kwan, Dr. Gregor Tegl, Dr. RyanSweeney, Dr. Peter Rahfeld, Dr. Lyann Sim, Emily Kwan, Phil Danby, Seyed Nassari, Rhea Bains,and Jacob Wardman for helpful conversations, perspective and inspiration. Connor Morgan-Langand Ryan Karimi for their bioinformatic support. Dr. Brian Mark, Ankoor Patel and VeronicaLarmour for their collaboration on the BglP project. The JGI, in particular, Dr. Yasuo Yoshikuniand Dr. Sam Deutsch for providing the GP synthetic gene library. Dr. Jose Henrique Pereira and Dr.Paul Adams for their collaboration on the acholetin project. Dr. Jayachandran Kizhakkedathu andIrina Chafeeva for the use of their MALS. Dr. Leonard Foster, Dr. Jason Rogalski and AlexandraNastasa for use of their ESI-MS.Finally, and most gratefully, I would like to thank my partner Daniela and son Lukas for theirunending patience and support.xxviDedicationFor my mom and dad.xxviiChapter 1IntroductionCarbohydrates are abundant throughout nature, play key roles in diverse biological processes andform fundamental components of a vast range of functional materials made for human use [1]. Forinstance, polysaccharides, the most abundant biopolymers on earth, have been closely linked tohuman life for millennia and have been used for a wide variety of purposes, ranging from food, tobuilding materials, to clothing, to medicine. More recently, the renewability and carbon neutralityof bio-sourced polysaccharides has led to significant interest in their potential to replace syntheticpolymers, such as plastics, derived from fossil fuels [2]. Beyond being sustainable and environ-mentally friendly, poly- and oligo-saccharides have seen considerable biomedical applications due totheir favorable biocompatible and biodegradable properties [3, 4]. Polysaccharides, and carbohy-drates generally, have been central in the development of nano- and micro-particles for drug deliverysystems [5–11], glycan conjugated therapeutics [12–14], wound dressings [15–19] and scaffolds for tis-sue engineering and 3D bioprinting [20–26]. The diverse applications of poly- and oligo-saccharidesoriginate from the identities of their monomeric precursors and the types of glycosidic linkages thatconnect them. Together with the gamut of potential precursors and array of possible linkages car-bohydrates are able to adopt a wide range of structures and functions, with greater conceivablecomplexity than their amino and nucleic acid counterparts. With this added complexity comesadded difficulty when attempting to chemically synthesize them [27]. In the biomedical context itis vital that synthesis occurs under conditions that result in a uniform and sequence-defined car-bohydrate to achieve the desired properties of the functional material. However, due in large partto the more or less chemical equivalency of individual glycoside hydroxyl groups, chemical synthe-sis often requires multiple inefficient protection and deprotection steps to control the stereo- andregio-specificity of glycosylation [28], resulting in increased costs and diminished yields. Enzymatic1synthesis represents an advantageous alternative to achieving uniform and sequence-defined carbo-hydrates by exploiting an enzyme’s innate substrate specificity and conformational control, whichavoids the inefficiencies associated with chemical synthesis [28, 29].1.1 Carbohydrate-Active EnzymesEnzymes that act on carbohydrates are known as Carbohydrate Active Enzymes (CAZymes). Tohelp improve the understanding of this diverse group of biocatalysts a sequence-based classifica-tion system was created, known as the CAZy database (CAZy DB) (http://www.cazy.org/), wherefamilies of CAZymes are generated based on amino acid sequence similarity [30]. Since sequencedictates structure, enzymes within a family have similar structures and mechanisms. The CAZyDB describes families of structurally-related functional domains of enzymes that degrade, modifyor create glycosidic linkages. It currently encompasses 5 enzyme classes: the glycoside hydrolases(GHs), polysaccharide lyases (PLs) and carbohydrate esterases (CEs), which are involved in glycandegradation; the glycosyltransferases (GTs), which are primarily responsible for glycan biosynthe-sis; and auxiliary activities (AAs), which are redox enzymes that act in conjunction with otherCAZymes. As well, the CAZy DB covers carbohydrate-binding modules (CBMs), which assist inCAZyme catalysis through adhering to its carbohydrate substrates. Within each enzyme class arefamilies based on amino acid sequence similarities. Due to their important biotechnological andbiomedical applications [31, 32], the GHs are currently the best-characterized class of CAZymes inthe CAZy DB, containing 171 families (as of March 2021) that are responsible for the hydrolysis,transglycosylation or phosphorolysis of glycosidic linkages. In particular, the phosphorolysis andtransglycosylation activities of GHs have been exploited for the purposes of glycan assembly.1.2 Enzymatic Carbohydrate SynthesisThe focus of this dissertation will be the glycoside phosphorylases (GPs), which are a class ofCAZymes that have seen frequent use for poly- and oligo-saccharide synthesis. GPs act througha process known as phosphorolysis that cleaves the glycosidic linkage with a phosphate moleculeresulting in the release of a sugar 1-phosphate [33, 34] (Figure 1.1). Due to the roughly equivalent2Figure 1.1: Glycoside phosphorylase (GP), glycosyltransferase (GT), glycoside hydrolase (GH) andglycosynthase (GS) generalized reaction scheme. A 1,4-linkage is shown as example only (centre).free energy associated with the glycosidic linkage and the phosphoester bond of the released sugar1-phosphate, GPs can perform phosphorolysis in reverse [35]. This allows GPs to be used for car-bohydrate synthesis by adding glycosyl moieties from sugar 1-phosphates to suitable acceptors [36].Furthermore, this synthetic paradigm is amenable to large scale applications due to the relativelylow cost associated with the sugar 1-phosphate starting materials. The innate substrate specificityand conformational control offered by GPs together with their inexpensive starting materials makethese enzymes attractive tools for poly- and oligo-saccharide synthesis. As new GP activities arediscovered and the array of these useful biocatalysts continues to grow, so too does the potential togenerate novel and diverse carbohydrate-based materials.In addition to GPs, glycosyltransferases (GTs) and glycoside hydrolases (GHs) have been thesubject of considerable biochemical characterization toward developing them as tools for the purposeof glycan assembly (Figure 1.1). GTs transfer a glycosyl residue from an activated donor substrate3to an acceptor glycan or other molecule, forming a glycosidic linkage [37]. GT-catalyzed reactionsare driven toward synthesis by the free energy associated with the conversion of the activated donor,commonly a nucleoside- or lipid-phosphosugar, to the glycosidically linked product. GTs have seenuse as tools for enzymatic carbohydrate synthesis because of their particularly high regio- andstereo-specificity [38].A notable achievement was the invention of glycosynthases (GSs) [39], which are mutant variat-ions of GHs that have been engineered to catalyze the formation of the glycosidic linkage. GSs havebeen used for the efficient synthesis of many oligosaccharides and glycoconjugates [40–46]. Insteadof nucleotide-phosphosugars, as is the case with GTs, GSs use glycosyl fluoride donors of oppositeanomeric configuration to that of the desired product to carry out glycosyl transfer to appropriateacceptors [47]. GSs are mutant variants of retaining GHs that normally utilize a double-displacementmechanism involving a glycosyl enzyme intermediate to hydrolyze the glycosidic bond. The GSis made by mutating the catalytic nucleophile of a retaining GH to a non-nucleophilic residue,preventing the enzyme from forming the glycosyl enzyme intermediate [39]. The glycosyl fluoridedonor mimics the glycosyl enzyme intermediate, allowing the transfer of the glycosyl residue to anappropriate acceptor, releasing hydrogen fluoride (HF) as a byproduct. Therefore, these enzymesare able to efficiently catalyze the formation of the glycosidic bond and, because they lack thecatalytic nucleophile, are unable to hydrolyze the product. These three types of enzymes (GTs,GSs and GPs) are capable of catalyzing the formation of the glycosidic linkage (Figure 1.1) andtherefore have utility for carbohydrate synthesis.1.3 Glycoside PhosphorylasesWhile GTs and GSs are both excellent tools for the purpose of enzymatic carbohydrate synthesis,GPs are the focus of this thesis for two primary reasons. (1) The low cost of the sugar 1-phosphatestarting materials make GPs particularly suited for large scale carbohydrate synthesis applications,whereas with alternative methods, scaling can be prohibited by the high costs of starting materi-als. This factor makes GPs particularly attractive candidates for further study. Furthermore, thereversibility of GPs allows for methods to be employed that would drive the starting materials costs4Figure 1.2: Hypothetical scheme involving two GPs for the synthesis of high-value glycans. Phos-phate shown as orange circle containing "P". Coloured symbols indicate ambiguous glycosylresidues.even lower. For example, one GP could be used to degrade an inexpensive glycan to produce a poolof sugar 1-phosphates, while a second GP could be deployed to use those sugar 1-phosphates asdonors to synthesize a different, more valuable product glycan (Figure 1.2). The bottleneck in thisapproach, however, is the limited range of GPs available, which restricts the diversity of carbohy-drates that can be assembled. (2) Given the vast number of GH activities known, it seems likely thatthe relatively low (35) number of known GP activities, is a significant underestimate of the actualnumber of GP activities present in nature. This is made yet more likely given that phosphorolysis isinherently more energetically favourable than hydrolysis for the intracellular metabolism of carbo-hydrates since the released sugar 1-phosphates feed directly into the glycolysis pathway without theneed for expenditure of ATP [48, 49] (Figure 1.3). Given this metabolic efficiency advantage andconsidering the wide range of diverse glycans that are metabolized by microbes, it is not surprisingto find that as more bacterial genomes are sequenced and their gene products characterized it hasbecome apparent that microbes may be utilizing more GPs in the degradation of carbohydrates thanpreviously thought. Indeed, it is probable that numerous forms of carbohydrates are metabolizedthrough phosphorolysis, suggesting an abundance of GPs to discover in nature.5Figure 1.3: Schematic representation of the metabolic efficiency advantage of GPs. GP mediateddegradation of a glucan, Glc(-Glc)n , can bypass phosphorylation from glucokinase and thereforeconserve n ATP molecules compared to hydrolytic degradation. PGM : phosphoglucomutase, Pi:inorganic phosphate.1.3.1 A Brief History of Glycoside PhosphorylasesThe discovery of the first glycoside phosphorylase (GP) came about in the first half of the 20thcentury. Carl and Gerty Cori, a Prague-born biochemist couple, had spent their careers, whichspanned over three decades examining how the human body metabolizes glucose. Early on in theirresearch, while exploring blood glucose regulation they discovered what is now referred to as the"Cori cycle", which describes the interplay between glucose and glycogen in the liver, blood andmuscle tissue [50]. Their seminal work on the topic came in 1938 when experimenting on thecatabolism of glycogen in animal skeletal muscle tissue at the Washington University School ofMedicine in St. Louis, Missouri, USA. There, the couple identified a new intermediate of glycogenbreakdown, glucose 1-phosphate (Glc1-P), and the enzyme responsible for its formation, glycogenphosphorylase [51]. With the aid of their collaborator, Arda Green, the Coris were able to purify andchemically characterize the newly discovered enzyme [52–54] and demonstrate the reversibility of thephosphorylase catalyzed reaction to produce glycogen from malto-oligosaccharides and Glc1-P [55].This latter experiment represented the first time a biological macromolecule had been synthesizedoutside of the body in a test tube. The Cori’s discovery and subsequent characterization of glycogen6phosphorylase earned them the Nobel Prize in Physiology or Medicine in 1947 (together with B.A.Houssey), with Gerty being the first American woman to receive the honour [56]. The discovery ofglycogen phosphorylase set in motion another paramount scientific discovery when Edwin Krebs, aformer postdoc of the Cori’s, continued to study glycogen phosphorylase into the early 1950s withhis colleague, Edmond Fischer. Using muscle glycogen phosphorylase as a model, the two identifiedreversible protein phosphorylation, caused by phosphorylase kinase and phosphorylase phosphatase,as a biological regulatory mechanism [57]. The discovery earned Krebs and Fischer the Nobel Prizein Physiology or Medicine in 1992.In 1943, Michael Doudoroff discovered the second GP, sucrose phosphorylase, from a dried cellpreparation of the bacterium Pseudomonas saccharophila [58, 59]. Unlike the glycogen phospho-rylase discovered five years previous, sucrose phosphorylase acted on a disaccharide as opposed tothe glycogen polysaccharide. However, like glycogen phosphorylase, Doudoroff demonstrated thereversibility of the sucrose phosphorylase reaction by generating sucrose from fructose and Glc1-P[58]. The next GP, maltose phosphorylase, would not be discovered until 1952 [60] then over the sec-ond half of the 20th century 10 more would be discovered that acted almost exclusively by cleaving aglucose residue from the non-reducing end of a di-, oligo- or polysaccharide, producing Glc1-P. Theone exception, discovered in 1999, was a 1,3-β-galactosyl-N -acetylhexosamine phosphorylase, whichrepresented the first GP known to cleave a residue other than glucose, in this case galactose (Gal)to generate galactose 1-phosphate (Gal1-P) [61]. A year later in 2000 another non-glucose cleavingphosphorylase was reported: N -N’ -diacetylchitobiose phosphorylase, which phosphorolyzed N,N’ -diacetylchitobiose to produce N -acetylglucosamine (GlcNAc) and N -acetylglucosamine 1-phosphate(GlcNAc1-P) [62]. Since the turn of the century, 23 more GPs have been identified that act upon arange of diverse carbohydrates, including several that cleave a non-reducing end mannose (Man) toproduce mannose 1-phosphate (Man1-P), which was first reported in 2011 [63]. Also, the first andonly GP, α-1,4-glucan:maltose 1-phosphate maltosyltransferase, was reported in 2010 that releasesa non-reducing end disaccharide to liberate maltose 1-phosphate (Mal1-P) from starch [64]. A listof the 35 GPs known to date are arranged chronologically in Table 1.1 and a more comprehensivelist, arranged by CAZy family, can be found in Table 1.2.7Table 1.1: Summary of GPs reported in the CAZy DB arranged chronologically.EC Name PhosphorolysisProductCAZyFamily ReferenceFirstReported2.4.1.1 Glycogen phosphorylase Glc-α1-P GT35 [51] Cori et al. 19382.4.1.7 Sucrose phosphorylase Glc-α1-P GH13 [58] Doudoroff 19432.4.1.8 Maltose phosphorylase Glc-β1-P GH65 [60] Fitting andDoudoroff 19522.4.1.20 Cellobiose phosphorylase Glc-α1-P GH94 [65] Sih and McBee 19552.4.1.31 Laminaribiose phosphorylase Glc-α1-P GH94 [66] Goldemberg et al. 19662.4.1.30 1,3-β-Oligoglucan phosphorylase Glc-α1-P GH149 [67] Marechal 19672.4.1.49 Cellodextrin phosphorylase Glc-α1-P GH94 [68] Sheth and Alexander 19692.4.1.97 1,3-β-Glucan phosphorylase Glc-α1-P GH161 [69] Albrecht and Kauss 19712.4.1.64 Trehalose phosphorylase (inverting) Glc-β1-P GH65 [70] Marechal andBelocopitow 19722.4.1.231 Trehalose phosphorylase (retaining) Glc-α1-P GT4 [71] Kitamoto et al. 19882.4.1.211 1,3-β-Galactosyl-N -acetylhexosaminephosphorylase Gal-α1-P GH112 [61] Derensy-Dron et al. 19992.4.1.230 Kojibiose phosphorylase Glc-β1-P GH65 [72] Chaen et al. 19992.4.1.280 N,N’ -Diacetylchitobiose phosphorylase GlcNAc-α1-P GH94 [62] Park et al. 20002.4.1.216 Trehalose 6-phosphate phosphorylase Glc-β1-P GH65 [73] Andersson et al. 20012.4.1.247 1,4-β-D-Galactosyl-L-rhamnosephosphorylase Gal-α1-P GH112 [74] Nakajima et al. 20092.4.99.16 α-1,4-glucan:maltose 1-phosphatemaltosyltransferase Mal-α1-P GH13 [64] Elbein et al. 20102.4.1.281 1,4-β-Mannosyl-glucosephosphorylase Man-α1-P GH130 [63] Senoura et al. 20112.4.1.279 Nigerose phosphorylase Glc-β1-P GH65 [75] Nihira et al. 20122.4.1.282 1,3-α-D-Glucosyl-L-rhamnosephosphorylase Glc-β1-P GH65 [76] Nihira et al. 20122.4.1.319 β-1,4-Mannooligosaccharidephosphorylase Man-α1-P GH130 [77] Kawahara et al. 20122.4.1.320 1,4-β-Mannosyl-N -acetylglucosaminephosphorylase Man-α1-P GH130 [78] Nihira et al. 20132.4.1.321 Cellobionic acid phosphorylase Glc-α1-P GH94 [79] Nihira et al. 20132.4.1.329 Sucrose 6-phosphate phosphorylase Glc-α1-P GH13 [80] Verhaeghe et al. 20142.4.1.332 1,2-α-Glucosylglycerolphosphorylase Glc-β1-P GH65 [81] Nihira et al. 20142.4.1.333 1,2-β-Oligoglucan phosphorylase Glc-α1-P GH94 [82] Nakajima et al. 20142.4.1.334 1,3-α-Oligoglucan phosphorylase Glc-β1-P GH65 [83] Nihira et al. 20142.4.1.339 β-1,2-Mannobiose phosphorylase Man-α1-P GH130 [84] Chiku et al. 20142.4.1.340 1,2-β-Oligomannan phosphorylase Man-α1-P GH130 [84] Chiku et al. 20142.4.1.x β-Glucoside phosphorylase Glc-β1-P GH3 [85] Macdonald et al. 20152.4.1.x β-1,3-Mannoside phosphorylase Man-α1-P GH130 [86] Awad et al. 20172.4.1.352 Glucosylglycerate phosphorylase Glc-α1-P GH13 [87] Franceus et al. 20172.4.1.359 Glucosylglycerol phosphorylase Glc-α1-P GH13 [88] Franceus et al. 20182.4.1.374 GDP-α-D-Manp:β-1,2-D-mannosyltransferase Man-α1-P GT108 [89] Sernee et al. 20192.4.1.x β-1,4-Mannosyl-glucuronic acidphosphorylase Man-α1-P GH130 [90] Li et al. 20202.4.1.x β-1,3-N -acetylglucosaminidephosphorylase GlcNAc-α1-P GH94 Reported here 20218Table 1.2: List of GPs reported in the CAZy DB organized by CAZy family.EC Name Mechanism Substrate PhosphorolysisProductCAZyFamilyReference (first report in the scientific literature)Activity Sequence Structure2.4.1.x β-Glucoside phosphorylase β-R Glc-β1,? Glc-β1-P GH3 [85] Macdonald et al. 2015 [85] Macdonald et al. 2015 [91] Macdonald et al. 20182.4.1.7 Sucrose phosphorylase α-R Glc-α1,β2-Fru Glc-α1-P GH13 [58] Doudoroff 1943 [92] Kitao and Nakano 1992 [93] Sprogoe et al. 20042.4.99.16 α-1,4-glucan:maltose 1-phosphatemaltosyltransferase α-R Glc-(α1,4-Glc)n n≥4 Mal-α1-P GH13 [64] Elbein et al. 2010 [64] Elbein et al. 2010 [94] Syson et al. 20112.4.1.329 Sucrose 6-phosphate phosphorylase α-R Glc-α1,β2-Fru6-P Glc-α1-P GH13 [80] Verhaeghe et al. 2014 [80] Verhaeghe et al. 2014 [95] Franceus et al. 20192.4.1.352 Glucosylglycerate phosphorylase α-R Glc-α1,2-Glycerate Glc-α1-P GH13 [87] Franceus et al. 2017 [87] Franceus et al. 2017 -2.4.1.359 Glucosylglycerol phosphorylase α-R Glc-α1,2-Glycerol Glc-α1-P GH13 [88] Franceus et al. 2018 [88] Franceus et al. 2018 -2.4.1.8 Maltose phosphorylase α-I Glc-α1,4-Glc Glc-β1-P GH65 [60] Fitting andDoudoroff 1952 [96] Ehrmann and Vogel 1998 [97] Egloff et al. 20012.4.1.64 Trehalose phosphorylase (inverting) α-I Glc-α1,α1-Glc Glc-β1-P GH65 [70] Marechal andBelocopitow 1972 [98] Maruta et al. 2002 -2.4.1.230 Kojibiose phosphorylase α-I Glc-α1,2-Glc Glc-β1-P GH65 [72] Chaen et al. 1999 [99] Yamamoto et al. 2004 [100] Okada et al. 20142.4.1.216 Trehalose 6-phosphate phosphorylase α-I Glc-α1,α1-Glc6-P Glc-β1-P GH65 [73] Andersson et al. 2001 [73] Andersson et al. 2001 -2.4.1.279 Nigerose phosphorylase α-I Glc-α1,3-Glc Glc-β1-P GH65 [75] Nihira et al. 2012 [75] Nihira et al. 2012 -2.4.1.282 1,3-α-D-Glucosyl-L-rhamnosephosphorylase α-I Glc-α1,3-Rha Glc-β1-P GH65 [76] Nihira et al. 2012 [76] Nihira et al. 2012 -2.4.1.332 1,2-α-Glucosylglycerolphosphorylase α-I Glc-α1,2-Glycerol Glc-β1-P GH65 [81] Nihira et al. 2014 [81] Nihira et al. 2014 [101] Touhara et al. 20142.4.1.334 1,3-α-Oligoglucan phosphorylase α-I Glc-(α1,3-Glc)n n≥2 Glc-β1-P GH65 [83] Nihira et al. 2014 [83] Nihira et al. 2014 -2.4.1.20 Cellobiose phosphorylase β-I Glc-β1,4-Glc Glc-α1-P GH94 [65] Sih and McBee 1955 [102] Reichenbecher et al. 1997 [103] Hidaka et al. 20062.4.1.31 Laminaribiose phosphorylase β-I Glc-β1,3-Glc Glc-α1-P GH94 [66] Goldemberg et al. 1966 [104] Kitaoka et al. 2012 [105] Kuhaudomlarp et al. 20182.4.1.49 Cellodextrin phosphorylase β-I Glc-(β1,4-Glc)n n≥2 Glc-α1-P GH94 [68] Sheth and Alexander 1969 [102] Reichenbecher et al. 1997 [106] O’Neill et al. 20172.4.1.280 N,N’ -Diacetylchitobiose phosphorylase β-I GlcNAc-β1,4-GlcNAc GlcNAc-α1-P GH94 [62] Park et al. 2000 [62] Park et al. 2000 [107] Hidaka et al. 20042.4.1.321 Cellobionic acid phosphorylase β-I Glc-β1,4-Gluconic acid Glc-α1-P GH94 [79] Nihira et al. 2013 [79] Nihira et al. 2013 [108] Nam et al. 20152.4.1.333 1,2-β-Oligoglucan phosphorylase β-I Glc-(β1,2-Glc)n n≥2 Glc-α1-P GH94 [82] Nakajima et al. 2014 [82] Nakajima et al. 2014 [109] Nakajima et al. 20172.4.1.x β-1,3-N -acetylglucosaminidephosphorylase β-I GlcNAc-(β1,3-GlcNAc)n n≥1 GlcNAc-α1-P GH94 Reported here Reported here -2.4.1.211 1,3-β-Galactosyl-N -acetylhexosaminephosphorylase β-I Gal-β1,3-GlcNAc/-GalNAc Gal-α1-P GH112 [61] Derensy-Dron et al. 1999 [110] Kitaoka et al. 2005 [111] Hidaka et al. 20092.4.1.247 1,4-β-D-Galactosyl-L-rhamnosephosphorylase β-I Gal-β1,4-Rha Gal-α1-P GH112 [74] Nakajima et al. 2009 [74] Nakajima et al. 2009 -2.4.1.281 1,4-β-Mannosyl-glucosephosphorylase β-I Man-β1,4-Glc Man-α1-P GH130 [63] Senoura et al. 2011 [63] Senoura et al. 2011 [112] Nakae et al. 20132.4.1.319 β-1,4-Mannooligosaccharidephosphorylase β-I Man-(β1,4-Man)n n≥2 Man-α1-P GH130 [77] Kawahara et al. 2012 [77] Kawahara et al. 2012 [113] Ye et al. 20162.4.1.320 1,4-β-Mannosyl-N -acetylglucosaminephosphorylase β-I Man-β1,4-GlcNAc Man-α1-P GH130 [78] Nihira et al. 2013 [78] Nihira et al. 2013 -2.4.1.339 β-1,2-Mannobiose phosphorylase β-I Man-(β1,2-Man)n n≥2 Man-α1-P GH130 [84] Chiku et al. 2014 [84] Chiku et al. 2014 [114] Cuskin et al. 20152.4.1.340 1,2-β-Oligomannan phosphorylase β-I Man-β1,2-Man Man-α1-P GH130 [84] Chiku et al. 2014 [84] Chiku et al. 2014 [115] Tsuda et al. 20152.4.1.x β-1,3-Mannoside phosphorylase β-I Man-β1,3-Man Man-α1-P GH130 [86] Awad et al. 2017 [86] Awad et al. 2017 -2.4.1.x β-1,4-Mannosyl-glucuronic acidphosphorylase β-I Man-β1,4-Gluconic acid Man-α1-P GH130 [90] Li et al. 2020 [90] Li et al. 2020 -2.4.1.30 1,3-β-Oligoglucan phosphorylase β-I Glc-(β1,3-Glc)n n≥1 Glc-α1-P GH149 [67] Marechal 1967 [116] Kuhaudomlarp et al. 2018 [117] Kuhaudomlarp et al. 20192.4.1.97 1,3-β-Glucan phosphorylase β-I Glc-(β1,3-Glc)n n≥2 Glc-α1-P GH161 [69] Albrecht and Kauss 1971 [118] Kuhaudomlarp et al. 2019 -2.4.1.231 Trehalose phosphorylase (retaining) α-R Glc-α1,α1-Glc Glc-α1-P GT4 [71] Kitamoto et al. 1988 [119] Saito et al. 1998 -2.4.1.1 Glycogen phosphorylase α-R Glc-(α1,4-Glc)n n≥3 Glc-α1-P GT35 [51] Cori et al. 1938 [120] Titani et al. 1977 [121] Sprang and Fletterick 19792.4.1.374 GDP-α-D-Manp:β-1,2-D-mannosyltransferase β-I Man-(β1,2-Man)n n≥1 Man-α1-P GT108 [89] Sernee et al. 2019 [89] Sernee et al. 2019 [89] Sernee et al. 2019I and R: inverting and retaining mechanisms, respectively.This table is modified and updated from Kitaoka, 2015 [122].91.3.2 Applications of Glycoside PhosphorylasesGPs, in general, can be split into two categories: disaccharide phosphorylases, which use a donorsugar 1-phosphate in conjunction with a monosaccharide acceptor to produce a disaccharide andphosphate, and the oligosaccharide phosphorylases which use a donor sugar 1-phosphate in con-junction with a di-, tri- or oligo-saccharide to subsequently lengthen the acceptor by one glycosylresidue at a time (or two in the case of α-1,4-glucan:maltose 1-phosphate maltosyltransferase [64]),with concomitant release of phosphate. Due to tighter control over degrees of polymerization,disaccharide phosphorylases have seen more varied industrial and academic use than their oligo-saccharide counterparts.Many of the successful applications of disaccharide phosphorylases have involved multi-enzymeone-pot reactions for the synthesis of glycans, often involving two GPs [123]. To minimize startingmaterial cost, the first stage of these reactions frequently involves GP-mediated phosphorolysis of anabundant naturally occurring disaccharide, such as sucrose, cellobiose or maltose to generate a poolof donor D-glucose 1-phosphate (Glc1-P). At this stage, other non-GP enzymes, such as a series ofphosphomutases and isomerases may be deployed to convert the donor to a form that is compatiblewith the second GP. For instance from α-D-glucose 1-phosphate (Glc-α1-P) to α-D-mannose 1-phosphate (Man-α1-P), through sugar 6-phosphate intermediates [78]. The second GP is then ableto use the new pool of compatible donor along with an appropriate acceptor to produce the targetglycan [78, 124–128]. This strategy has been used to produce kilogram quantities of lacto-N -biose[127], a prebiotic linked to health benefits in humans [129–133]; gram quantities of galacto-N -biose[128], the O-linked core I structure of mucin glycoproteins and the T-antigen, a cancer biomarker[134]; Man-β1,4,-GlcNAc [78], a core component of N -linked glycans; Glc-α1,β-2-allulose [124], a‘reduced-calorie’ sweetener; and other valuable disaccharides such as nigerose (Glc-α1,3-Glc) [126]and Gal-β1,4-rhamnose [125]. Oligosaccharide phosphorylases have been primarily targeted towardthe degradation of environmentally abundant polysaccharides such as starch or cellulose, or theproduction of synthetic versions of these polymers. For example, α-glucan phosphorylases havebeen used in the production of synthetic amylose for use in nanomedical drug delivery systems [135,136] and the transformation of cellulosic biomass to starch [137], while cellodextrin phosphorylases10(CDPs) have been used to produce a variety of interesting cellulosic materials [138–140]. Combiningboth di- and oligo-saccharide phosphorylases has been used to great effect as demonstrated by Zhonget al. [141]. With sucrose, glucose and phosphate as the starting material, the group used sucrosephosphorylase in conjunction with cellobiose phosphorylase and CDP to generate prebiotic cello-oligosaccharides, with yields in the range of 100 g/L. As more and more GPs are discovered and theirspectrum of activities broadens, so too do the potential applications for these useful biocatalysts.1.3.3 Classification of Glycoside PhosphorylasesGPs are classified as members of GH and GT families in the CAZy DB (Table 1.3). Families GH65,GH94, GH112, GH130, GH149, GH161 and GT108 primarily contain GPs that react through adirect displacement reaction mechanism that inverts anomeric configuration [142–144] and thereforeare considered either α-inverters (GH65) or β-inverters (GH94, GH112, GH130, GH149, GH161,GT108) (Figure 1.4A and B). GPs belonging to families GH3 and GH13 react though a doubledisplacement mechanism [142, 143] involving a covalent glycosyl-enzyme intermediate resulting inretention of the anomeric configuration (Figure 1.4C and D). GPs that belong to GT4 and GT35operate under anomalous mechanisms. GT4 α,α-trehalose phosphorylases react in a manner thatretains anomeric stereochemistry, however, its mechanism is unclear and remains a matter of debateTable 1.3: Classification of glycoside phosphorylases in the CAZy database.CAZyFamily MechanismNo. of GPEC/familyPhosphorolysisProductGH3 β-retaining 1 Glc-β1-PGH13 α-retaining 5 Glc-α1-P or Mal-α1-PGH65 α-inverting 8 Glc-β1-PGH94 β-inverting 7 Glc-α1-P or GlcNAc-α1-PGH112 β-inverting 2 Gal-α1-PGH130 β-inverting 7 Man-α1-PGH149 β-inverting 1 Glc-α1-PGH161 β-inverting 1 Glc-α1-PGT4 α-retaining 1 Glc-α1-PGT35 α-retaining 1 Glc-α1-PGT108 β-inverting 1 Man-α1-PTotal GP activities 3511Figure 1.4: Catalytic mechanisms of α- and β-inverting glycoside phosphorylases. (A) α-inverting,(B) β-inverting, (C) α-retaining (except GT4 and GT35), and (D) β-retaining GP catalytic mech-anisms where X represents the acid/base residue. Note: Rovira et al. have proposed an alternativeβ-inverting mechanism for GH130s and GT108s GPs where the O-3 donates a proton to the leavingoxygen and in turn, an Asp acid/base residue donates a proton to O-3 [144], instead of the oxygenreceiving the proton directly from the acid/base residue as shown in (A).12[143, 145]. One proposed mechanism of GT35 glycogen phosphorylases states that it utilizes apyridoxal 5’-phosphate (PLP) co-factor which participates in the mechanism as a general acid/base.PLP donates a proton to an incoming phosphate which in turn donates a proton to the leaving O-4’from the newly formed non-reducing end of the glycogen substrate. The phosphate then performsnucleophilic attack on the C-1 of the carbonium ion, which resulted from the cleavage of the non-reducing end’s C(1)-O(1) bond, to form glucose 1-phosphate and retain the α-anomeric configuration[146–149]. Despite their mechanistic differences, these GPs are classified as either α-retainers (GH13,GT4 and GT35) or β-retainers (GH3).1.4 Glycoside Phosphorylase Discovery1.4.1 Experimental and Rational ExplorationOver the past 83 years since GPs have been known to exist, they have generally been discoveredthrough one of two means, either experimental and rational exploration or genomic sequence ex-ploration. The technology available to researchers at the time of discovery often determined whichmethod was applied. Early on and through to the end of the 20th century GPs were primarilydiscovered through experimental and rational exploration. The first three GPs to be reported wereidentified after researchers independently observed the phosphorolytic cleavage of glycogen, sucroseand maltose [59, 150, 151] in biological systems. After reporting on these phenomena, it was shortlythereafter discovered that GPs were responsible for the observed phosphorolytic cleavage, whichlead to the discovery of glycogen phosphorylase (1938), sucrose phosphorylase (1942) and maltosephosphorylase (1952) described above. The next GP would be found in 1955 after it was observedby Richard H. McBee that the organism Hungateiclostridium thermocellum (then called Clostridiumthermocellum) was able to grow on cellulose and cellobiose, but not glucose alone [152]. Recognizingthat H. thermocellum likely does not hydrolyze cellobiose to glucose, McBee together with CharlesJ. Sih, a graduate student of his at Montana State College, sought to determine whether a yetunrecognized metabolic pathway could be responsible for cellobiose utilization. While monitoring along term culture of the organism, Sih noted a significant uptake of inorganic phosphate from thegrowth medium, which led the two to identify cellobiose phosphorylase from a cellular extract, along13with the product, Glc1-P [65]. Sih speculated in his Master’s Thesis that the metabolic efficiencyadvantage (Figure 1.3) of phosphorolytically cleaving cellobiose to produce Glc1-P so to not toexpend a molecule of ATP was the explanation for his supervisor’s earlier observation [65]. In 1969,H. thermocellum was the source of a second cellulose degrading GP, cellodextrin phosphorylase [68].While H. thermocellum served as the source of the first di- and oligo-saccharide GPs that cleaveβ1,4-linked glucose, the single-cell algae Euglena gracilis provided the source of the first two GPsthat cleave β1,3-linked glucose, laminaribiose phosphorylase (1966) and 1,3-β-oligoglucan phospho-rylase (1967) [66, 67], along with the first trehalose phosphorylase (α-inverting GH65) (1972) [70]. E.gracilis has been studied extensively due, in part, to its unique ability to produce the energy storagemolecule, paramylon (linear β1,3-linked glucose chains) through photosynthesis [153]. Paramylonhas many potential applications toward human health [154] as studies have indicated its use as astimulator of the immune system [155, 156], an antitumor compound [157], a cholesterol-loweringcompound [158, 159] and a moderator of postprandial blood glucose and insulin response [159, 160].The gene sequence of the 1,3-β-oligoglucan phosphorylase was only recently reported in 2018 [116],which established the new CAZy family, GH149. In 1971, another β1,3 cleaving oligo-saccharideGP was discovered in a separate algae, Ochromonas malhamensis, and like its GH149 counterpart,the gene sequence was also only recently reported [118], which interestingly established another newCAZy family, GH161. Traditionally, paramylon has been produced through the cultivation of E.gracilis, perhaps now that these two gene sequences have been identified, more cost effective andefficient methods can be developed for the production of this valuable glycan through employingheterologously produced GPs.In 1988 a second, but mechanistically distinct trehalose phosphorylase (α-retaining GT4) wasidentified from enoki mushrooms (Flammulina velutipes) after it was observed that trehalase ac-tivity was elevated in the organism’s mycelia compared to its fruiting bodies [71]. Further investi-gation led to the discovery that, like glycogen phosphorylase, this α-retaining GT4 does not oper-ate via a double-displacement mechanism, but rather one that is likely best described as a singledisplacement mechanism as proposed by Luley-Goedl and Nidetzky [143]. Next, in 1999, a 1,3-β-galactosyl-N -acetylhexosamine phosphorylase was identified while investigating Bifidobacteriumbifidum, a normal and beneficial inhabitant of the human microbiome [61]. Followed in succes-14sion were kojibiose phosphorylase (1999) [72], N,N ’-diacetylchitobiose phosphorylase (2000) [62],trehalose 6-phosphate phosphorylase (2001) [73], α-1,4-glucan:maltose 1-phosphate maltosyltrans-ferase (2010) [64], then GDP-α-D-Manp:β-1,2-D-mannosyltransferase (2019) [89] each discoveredwhile researching carbohydrate metabolism in Thermoanaerobacter brockii, Vibrio furnissii, Lacto-coccus lactis, Mycobacterium smegmatis and Leishmania mexicana, respectively. This method ofexperimental and rational exploration for GPs has led to the discovery of 17 of the 35 known GPactivities over a 72-year span.1.4.2 Genomic Sequence ExplorationAs sequencing technology advanced into the 21st century and freely available genomic sequences be-came available, researchers began to seek out probable GP gene sequences within (mostly bacterial)genomes via sequence homology searches. The first GP to be identified through this method was1,4-β-D-galactosyl-L-rhamnose phosphorylase in 2009 [74]. Using the sequence of the only knownGH112 GP at the time, 1,3-β-galactosyl-N -acetylhexosamine phosphorylase, Nakajima et al. per-formed a homology search against the recently sequenced genome of the anaerobic cellulolytic bac-terium, Clostridium phytofermentans. They identified three homologues which were sub-cloned outof the C. phytofermentans genome into expression vectors then heterologously expressed in E. colifor functional characterization. One of the three was shown to have the novel 1,4-β-D-galactosyl-L-rhamnose phosphorylase activity, while the other two possessed the same activity as the originalqueried GH112. This process of querying sequences of known GPs against a reference database ofgenomic sequences would essentially be repeated by a handful of research groups over the next fiveyears, after which 14 new GP activities were identified [63, 74–84] (Table 1.1).Beginning in 2017, two groups began to take a different approach towards identifying new GPsthrough genomic sequence exploration by analyzing phylogenetic relationships between character-ized and uncharacterized members within a family. Awad et al. discovered a β-1,3-mannosidephosphorylase from the GH130 family in the genome of the bacterium, Zobellia galactanivorans,after it was noticed the GP did not share a significant phylogenetic relationship with any othercharacterized GH130 members [86]. This finding led the group to heterologously express the GPand biochemically characterize its activity. In that same year, Franceus et al. assembled a phyloge-15netic tree based on the known and predicted GP peptide sequences from subfamily 18 of GH13 (thesubfamily that GH13 GPs belong to, except α-1,4-glucan:maltose 1-phosphate maltosyltransferase,which is a member of subfamily three). The tree, they discovered, was composed of two majorbranches, one of which contained the known, characterized GPs and the other unexplored branchcontained no characterized GPs. By targeting a putative GP sequence in the unexplored branchwhich was flanked in the genome by genes that were predicted to use glycerate as a substrate, thegroup discovered the new GP activity, glucosylglycerate phosphorylase [87]. The following year(2018), the same research group probed the same unexplored branch for the GH13 (subfamily 18)phylogenetic tree using a distinct method than they had previously employeed. Having alreadyidentified several acceptor residues that are highly conserved throughout the subfamily they thensought out putative GP sequences in which those positions were substituted with different aminoacid residues, indicating they may act on a different substrate. The search lead the group to discoverthe novel GP activity, glucosylglycerol phosphorylase from Marinobacter adhaerens HP15 [88]. In2020, Li et al. [90] applied sequence similarity networks (SSNs), which is a tool to visualize sequencerelationships within families [161], to analyze GH130 sequence diversity and predict new functions.Through this approach they not only identified a novel β-1,4-mannosyl-glucuronic acid GP activity,but also developed a strategy to predict linkage specificity and identified several unexplored GH130meta-nodes that may contain even more new activities. The method of genomic sequence explo-ration has led to the discovery of 18 of the 35 known GP activities, with 14 of those identified overa 5 year period from 2009-2014.1.4.3 Functional Metagenomic ExplorationFunctional metagenomic exploration has yet to directly lead to the discovery of a new GP activity,however, screening paradigms now exist that can identify GP activities from CAZy families (GH) 3,13, 65, 94, 112, 130 and 149 [91, 162], which are described in Chapters 3 and 4. Metagenomic explo-ration did play a role in the discovery of 1,4-β-mannosyl-N,N’ -diacetylchitobiose phosphorylase (EC2.4.1.320) as its DNA sequence originated from a human gut metagenomic library [163]. Though,the metagenomic screen itself was not specifically targeting GP activity, but rather GH activitywhere the screening media contained phosphate. In fact, the GP activity was not identified until16three years after the original metagenomic screen during a followup study by Ladevèze et al. [164].Nevertheless, this finding established that GP activities can be detected though function-basedmetagenomic screening.Functional metagenomic exploration offers a means to search for novel biocatalysts from geneticmaterial drawn directly from the environment [165]. Metagenomic libraries provide access to thevast reservoir of uncultivated genetic diversity encoded in microbial communities inhabiting naturaland engineered ecosystems that was previously unavailable through traditional cultivation-basedmethods [165–167]. To bridge this cultivation gap, emerging sophisticated sequence- and function-based screening technologies continue to be successfully implemented to identify novel enzymes,including CAZymes, within these libraries [167–170].Sequence-based methods rely on massively parallel, high-throughput (HT) DNA sequencingtechnologies in conjunction with reference databases to functionally annotate genes from meta-genomic libraries. However, gene identification from sequence-based methods is limited to closehomologs of genes already represented in the reference database. Therefore, under the currentparadigm sequence-based methods cannot reliably assign function to novel enzymes identified frommetagenomic libraries. Function-based screening technologies can complement this limitation bytargeting enzyme function, rather than sequence similarities and therefore are able to identify novelenzymes, such as GPs. Generation of these libraries typically involves the cloning of environmentalDNA (eDNA) fragments into suitable vector backbones for heterologous expression in an exogenoushost, such as E. coli. Figure 1.5 describes the workflow of metagenomic library construction forplate-based metagenomic screens. Following the construction of a library, functional assays can beperformed on the host organisms, each carrying a unique fragment of eDNA, in a HT manner usinga reporter system which targets the desired activity. The reporter systems are highly specific to thetype(s) of activity being targeted and as a result there are a wide variety of approaches one can taketo target an enzyme of interest. However, to accommodate HT methods, most approaches couple thetarget activity to a chromo- or fluoro-genic response or another easy-to-identify phenotype. Sincethe advent of function-based metagenomic screening many methods have been successfully appliedto identify a variety of activities, including: antibiotic resistance [171], proteases [172], lipases [173],isomerases [174] and GHs [175, 176]. However, no function-based metagenomics screening approach17Figure 1.5: Metagenomic library workflow. Method used to generate microtiter plate large-insertmetagenomic large-insert libraries for sequence- and/or function-based analysis.18has been developed that targets GP activity, prior to the work presented in this thesis.1.4.4 Future Outlook on Glycoside Phosphorylase DiscoveryTypically, metagenomic screens are performed in microtiter plates or on agar, both amenable to HTscreening methods permitting up to 105 clones to be screened per experiment [177]. Fluorescence-activated cell sorting (FACS) has been recruited for metagenomic studies, which can increase thethroughput up to 109 clones per experiment [177]. Until recently, metagenomic FACS screens hadbeen limited to reporter systems that can function in vivo at the single cell level. However, newadvances have demonstrated these ultra HT metagenomic FACS screens can be done in cell lysateencapsulated in water/oil nano-droplets [178, 179] generated with micro-fluidic devices. Theseemerging methodologies will expand the future potential of FACS towards functional metagenomicscreening. For the purpose of discovering new GPs, a limiting factor remains that a stopped assay,such as the one described in Chapter 4 [162], is not amenable to the FACS method because there isno ability to add additional reagents to the droplets in a simple, reliable fashion. A continuous assay,such as the one described in Chapter 3 [91], where a fluorogenic aglycone substituent is attached tothe screening substrate is required to reliably distinguish hits using the FACS method.Over the past couple of decades there has been a massive, disorderly increase of genomic data.As sequence technologies continue to advance and reference databases continue to grow, it is becom-ing more and more challenging to keep pace by accurately assigning function to the genes containedwithin those databases. While clever methods have been developed to identify, then character-ize novel gene products using these extensive reference databases (as described above for GPs byFranceus et al. and Li et al. [87, 88, 90]), they are still only able to biochemically characterizeone (or several) gene product(s) at a time. In 2014, the Joint Genome Institute (JGI), a nationallaboratory operating under the purview of the US Department of Energy, attempted to tackle thisissue by developing a HT functional-characterization methodology [180]. As a proof-of-principle, theJGI collected a representative set of sequences belonging to CAZy family GH1 (no GPs representedin this family) from the reference database then generated a phylogenetic tree. From this tree,which represented 967 sequences, they selected a phylogenomically diverse sample of 175. Those175 GH1 gene sequence were synthesized, then cloned into an expression vector and transformed19into E. coli to generate a gene synthesis, plate-based library. The library was then assayed in aHT parallel fashion that permitted the functional characterization of the whole 175-clone libraryat once [180]. By combining sequence-rich reference databases, gene synthesis technologies andHT screening methodologies, the JGI established a broadly applicable functional-characterizationparadigm that provides a pathway toward the discovery of many new enzyme activities, includingGPs.In 2013, a group led by Gabrielle Potocki-Véronèse at IRNA Laboratories and the Universitéde Toulouse, had proposed the creation of two GH130 subfamilies and an uncharacterized clusterbased on sequence analysis, structural modelling and experimental findings [164]. One subfamilygrouped GPs with narrow specificity towards Man-β1,4-Glc together, while the other grouped thepromiscuous GPs with active sites able to accommodate complex glycan structures. However,as more research was done and new GH130 activities were identified, most of them were shownto be classified in the single uncharacterized cluster, which also contained non-GP mannosidases.Despite their 2013 classification system still being used today, the Toulouse group designed a newclassification system that utilized SSNs, the updated 2020 set of functionally characterized GH130sand a sequence list that was six-times larger than it was in 2013, and that had been supplementedwith metagenomically derived sequences [90]. The strategy allowed them to identify a novel GH130activity and identify several other meta-nodes that may also contain novel activities. This strategyinvolves the generation of a SSN like the one shown in Figure 1.6 that was generated independentlyfor this dissertation using the Enzyme Function Initiative-Enzyme Similarity Tool (EFI-EST) webportal [181–184] and the GH130 sequence list available in the CAZy DB.SSNs are generated by first performing an all-by-all BLAST of a sequence list to generate anoutput table that provides an alignment score, or Expect (E)-value, for every possible sequencepair in the data set. The alignment score is a parameter that describes the statistical significanceof a sequence pair being matched from a BLAST search by chance within the sequences beinganalyzed [185]. The lower the alignment score the more significant the match. The SSN uses theoutput table by denoting sequences as nodes and the associated alignment scores are utilized byapplying a threshold E-value, where any pair of nodes that satisfies the threshold an edge will becreated that connects the two. If an alignment score of a node pair does not satisfy the threshold20Figure 1.6: Example sequence similarity network (GH130). Amino acid SSN of 1854 unique GH130sequences from the CAZy DB with an E-value threshold of 10-90. Sequences are grouped intorepresentative nodes when they share a percent identity of 95 or greater. Sequence fragments below250 a.a. were excluded. Nodes coloured based on meta-node clustering. Functionally characterizedGH130s are represented as yellow symbols with black outlines. β-1,4-Mannosyl-glucuronic acidphosphorylase is not represented because sequence identifiers were not available.21Figure 1.7: Alignment score vs. percent identity plot for GH130. This plot is an output data setprovided by the EFI-EST after submitting a sequence set through the web portal. An E-value of10-90 was selected as the threshold value to generate the SSN in Figure 1.6.an edge will not be created and those nodes will not be linked. The general rule of thumb, atleast when rendering a SSN from a new sequence set, is to begin with an alignment score thresholdthat equates to ∼40 percent sequence identity. Determining the relationship between alignmentscore and percent identity can be achieved by plotting the two variables against each other (Figure1.7). When an appropriate threshold has been selected the SSN can be rendered to reveal theclustering pattern of the meta-nodes (Figure 1.6). The layout patterns of the nodes are determinedusing spacing algorithms that come packaged with SSN visualization software, such as Cytoscape[186]. Mapping on the functionally characterized GH130s shows that activity types primarily clustertogether in the same meta-nodes. While there isn’t a precise one-to-one mapping of activities tometa-nodes, this method can provide valuable information in terms of prioritizing the targetingof one’s search for novel activities, for instance, meta-nodes with no characterized members orlone nodes (singletons). Additionally, meta-nodes that contain multiple activities (such as thedark blue, upper right meta-node in Figure 1.6) can be isolated and more stringent thresholds canbe applied to that grouping to generate sub-meta-nodes, which can inform more precise functionalpredictions and guide biochemical characterization. As recognized by the Toulouse group, there is no22alignment score threshold that could be applied to the GH130 SSN that non-ambiguously delineatessequences based on linkage type or substrate specificity. Therefore, to overcome this limitation, theyincorporated information regarding the presence of signal peptides and active site residues involvedin catalysis and substrate recognition into their 2020 GH130 prediction strategy [90]. Using thisimproved prediction strategy, functional predictions were made for uncharacterized meta-nodes,then candidates belonging to four of those meta-nodes were biochemically characterized. One ofthe four was confirmed to not be a GP, which was predicted. The other three were confirmed topossess the linkage specificities that were predicted based on their model and one was discovered toutilize the acceptor, glucoronic acid, which represented a new GH130 activity. The Toulouse group,therefore, developed and validated a robust functional prediction strategy for GPs that incorporatessequence, functional and structural data.Future research aiming to identify new GP activities and better understand GP diversity whichrely on sequences in reference databases could benefit from a combined approach utilizing theToulouse group’s functional prediction strategy to inform the JGI’s gene synthesis selection and sub-sequent high-throughput functional characterization methods. Functional metagenomic exploration,which does not rely on pre-existing sequence databases, is also able to contribute by identifying newGP activities by providing access to the vast reservoir of uncultivated microbial diversity. Whilerational exploration that endeavours to explain and understand observed biological phenomena,such as the occurrence of a phosphorylated glyco-metabolite or an uncommon active site residue,will also continue to uncover novel GP activities. Operating in concert, these approaches have thepotential to rapidly broaden the spectrum of known GP activities, provide a more robust under-standing of this class of enzyme and close the gap between the ever expanding sequence databasesand biochemically characterized GPs.1.5 Dissertation OverviewThe aim of this thesis is to develop new tools and methodologies for GP discovery, then apply themtowards broadening the range of known GP activities. This will encompass using an experimentaland rational approach, based on endeavouring to explain a mechanistic oddity observed in the23GH3 CAZy family, to identify a new GP activity. Next, the development of two metagenomicscreening methodologies will be described that target GP activity and provide the means for futurefunctional metagenomic exploration. Lastly, a phylogenomically diverse GP library including fiveCAZy families will be assayed for substrate specificity, which resulted with the discovery of a novelGP activity, a new biopolymer and established a substrate screening paradigm to efficiently exploreGP genomic space. This thesis will lead to a better understanding of the diversity of GP activitiesin nature, insights into how we can engineer GPs to modify substrate specificity, establish a HTfunctional-characterization method, and provide a platform to explore the genomic and metagenomicsequence space for novel GP activities.Brief Overview of Research ChaptersChapter 2 details the discovery and characterization of the first GP discovered in CAZy familyGH3, which represent the first reported examples of retaining β-glycoside phosphorylases, and thefirst instance of free GlcNAc β1-phosphate in a biological context.Chapter 3 describes the development and implementation of a HT metagenomic screen target-ing GP activity (forward phosphorolysis) that identified a β-glycoside phosphorylase. Based onstructural analysis of the metagenomic GP, a residue was identified in the active site that shed lighton the simple ways in which enzyme specificity can be encoded and swapped.Chapter 4 describes the development and implementation of a HT metagenomic screen target-ing GP activity (reverse phosphorolysis) that identified several GH94s and one GH149.Chapter 5 details the functional-characterization of a phylogenomically diverse library of GPsassembled across five CAZy families and the identification of novel GP activity, β-1,3-N -acetyl-glucosaminide phosphorylase and the novel β-1,3-linked GlcNAc biopolymer, acholetin.24Chapter 2Identification of a new GlycosidePhosphorylase from CAZy family GH32.1 SummaryCarbohydrate active enzyme (CAZy) glycoside hydrolase family GH3 consists primarily of stereo-chemistry retaining β-glucosidases, but also contains a sub-family of β-N -acetylglucosaminidases.In 2000, enzymes from this sub-family were shown to use a histidine residue within a His-Asp dyadcontained in a signature sequence as their catalytic acid-base residue. Reasons for their use of Hisrather than the Glu or Asp found in other glycosidases were not apparent. Through studies ona representative member, the Nag3 β-acetylglucosaminidase from Cellulomonas fimi, I now showthat a subset of these enzymes act preferentially as glycoside phosphorylases. Their need to accom-modate an anionic nucleophile within the enzyme active site explains why histidine is used as theacid-base catalyst in place of the anionic glutamate seen in other GH3 family members. Kineticand mechanistic studies reveal that these enzymes also employ a double-displacement mechanism in-volving a covalent glycosyl-enzyme intermediate, which was directly detected by mass spectrometry.Phosphate has no effect on rates of formation of the glycosyl enzyme intermediate, but acceleratesturnover of the N -acetylglucosaminyl enzyme intermediate approximately three-fold, while accel-erating turnover of the glucosyl-enzyme intermediate several hundred fold. This chapter detailsthe identification and characterization of the first retaining β-glycoside phosphorylases and alsorepresents a new activity within the GH3 family.252.2 BackgroundThe glycoside hydrolases from CAZy family GH3 display an unusual diversity in structure, specificityand biological role and are widely distributed in bacteria, fungi and plants. This sequence-definedfamily contains β-glucosidases, N -acetylglucosaminidases (often dubbed NagZs), β-xylosidases andα-L-arabinofuranosidases. The principal initially-identified role of GH3 glycosidases was in biomassdegradation, but other roles in pathogen defense and plant and bacterial cell wall remodeling havebecome apparent. A subset of these enzymes plays a role in the recycling of peptidoglycan byremoving the non-reducing end N -acetylglucosamine from the disaccharide product released bylytic transglycosylases and muramidases (GlcNAc-anhMurNAc-peptide and its hydrated version)in gram-negative and gram-positive bacteria respectively [187]. The resultant (anhydro)-MurNAcpeptide is an activator of β-lactamase production in some gram-negative bacteria, rendering the GH3N -acetylglucosaminidases a possible therapeutic target [32, 188, 189]. Accordingly, considerableattention has been devoted to the generation of potent, selective inhibitors for this group of enzymes.Glycosidases from GH3 are retaining enzymes and cleave their substrates in an acid-base cat-alyzed two-step double displacement mechanism (Figure 1.4C and D) involving a covalent glycosyl-enzyme intermediate in which a fully conserved aspartic acid functions as the catalytic nucle-ophile. Detailed structural and mechanistic studies on several GH3 glycosidases have confirmedthat both the β-glucosidases and the N -acetylglucosaminidases follow this mechanism [190–198].This contrasts sharply with the β-hexosaminidases from CAZy family GH20, which lack an en-zymatic nucleophile and instead use the acetamide of the substrate in that role, forming an ox-azoline or oxazolinium ion intermediate [199, 200]. GH20 hexosaminidases also differ from thoseof GH3 in having both N -acetylglucosaminidase and N -acetylgalactosaminidase activity, while noN -acetylgalactosaminidase activity is seen in GH3 enzymes.While the catalytic nucleophile is fully conserved within GH3 enzymes, such is not the case for theacid/base catalyst. In some two-domain enzymes, such as that from barley [194], the nucleophile islocated in the active site within the N-terminal (β/α)8-barrel, while the acid-base protrudes into theactive site from the C-terminal domain. By contrast, most of the GH3 N -acetylglucosaminidasesare single domain enzymes for which, until recently, neither sequence alignments nor structures26suggested an obvious candidate for the acid/base catalyst. However, the structure of NagZ fromthe gram-positive organism Bacillus subtilis identified a histidine residue on a flexible loop, inconjunction with a nearby aspartic acid, as a candidate for this role [192]. Importantly the loopcontaining this Asp-His dyad was highly conserved and indeed had been identified previously as asignature sequence for N -acetylglucosaminidases within GH3 and proposed to play a role in bindingof the acetamide moiety [195].This proposal of a histidine as the acid/base was somewhat surprising since in essentially allother glycosidases this role has been played by a glutamate or aspartate, presumably in largepart to generate a more anionic environment upon proton transfer that will help stabilize theoxocarbenium ion-like transition state [201, 202]. However, as was noted previously [192, 203], asimilar replacement of the anionic Glu or Asp with a neutral residue has been seen in several othergroups of glycosidases, most notably the sialidases of GH33 and GH34 and the myrosinases of GH1.This substitution was attributed to the fact that the substrates themselves bear a negative charge,thus replacement of an anionic side chain with a neutral residue minimizes Coulombic repulsion.In the case of GH33 and GH34 sialidases the substrate has a carboxylate group at C-1 and thecatalytic nucleophile is a tyrosine residue, while in GH1 myrosinases the acid/base glutamate isreplaced by glutamine to accommodate the anionic aglycone of the substrate [204–206]. The reasonfor substitution of the acid/base Glu in GH3 N -acetylglucosaminidases by histidine is not obvious.One suggestion was that this would allow turnover of GlcNAc-MurNAc substrates from which thepeptide chain has been hydrolysed, since these bear an anionic lactyl moiety that could be involvedin substrate-assisted catalysis [192, 203]. However, subsequent studies rendered this explanationunlikely [203].Further consideration on possible reasons for this Asp-His substitution brought us to the ideathat perhaps the negative charge being accommodated lay not in the carbohydrate substrate, but inthe attacking species. The most probable such anionic nucleophile in a cellular environment wouldbe phosphate. If so, then perhaps these enzymes function preferentially as glycoside phosphory-lases rather than as glycoside hydrolases. Glycoside phosphorylases are categorized in both GT andGH families within CAZy [207]. The only GT-based phosphorylases to date are the α-retainingglycogen phosphorylases (GT35) and trehalose phosphorylases (GT4). Most other phosphorylases27are inverting enzymes with major families being the β-inverting cellobiose/cellodextrin and N,N’ -diacetylchitobiose phosphorylases (GH94) and the α-inverting GH65 phosphorylases such as maltosephosphorylase, along with a few others in GH112, GH130, GH149, GH161 and GT108 [33, 143, 207].The only retaining GH phosphorylases described are GH13 phosphorylases such as sucrose phos-phorylase, α-1,4-glucan:maltose 1-phosphate maltosyltransferase, glucosylglycerate phosphorylaseand glucosylglycerol phosphorylase, all GH13 α-cleaving/forming enzymes [58, 64, 87, 88]. Interest-ingly, at the time of this research (2015) no retaining β-glycoside phosphorylases had been reported.Therefore, should these GH3 hexosaminidases prove to function as phosphorylases through an anal-ogous retaining mechanism to that of the GH3 hydrolases, they would represent a new category ofphosphorylases.2.3 Results and DiscussionThe enzyme chosen to test this hypothesis was the GH3 β-glucosaminidase Nag3 from Cellulomonasfimi, which our laboratory had characterized previously and shown to have dual β-glucosaminidase/β-glucosidase activity and to share the conserved β-glucosaminidase signature sequence containing theAsp-His dyad noted earlier [193, 195]. All the previous studies were performed in phosphate buffer,so unwittingly phosphorolysis had been monitored rather than hydrolysis. Nag3 was expressed andpurified as described previously [193] and exchanged into 50 mM HEPES buffer (pH 7.0). It wasthen assayed at fixed concentrations of three different glycoside substrates at each of a series ofphosphate concentrations. Two glucoside substrates of quite different aglycone leaving group ability(pNP, pKa = 7 and DNP, pKa = 4) were chosen, along with one glucosaminide bearing the samepNP leaving group. As seen in Figure 2.1, a strong dependence of rate on phosphate concentrationswas observed, with rates increasing rapidly with rising phosphate at low concentrations and inhi-bition being seen at higher concentrations. This is particularly true for the glucoside substrates,where rate increases of over 100-fold are seen for aryl glucosides upon addition of phosphate. Ratesof pNPGlcNAc cleavage in the absence of phosphate are comparable to those for pNPGlc, but theactivity increases seen upon phosphate addition to pNPGlcNAc are much more modest.28ABCFigure 2.1: Phosphate activation of substrate cleavage by C. fimi Nag3. Michaelis-Menten (left) andLineweaver Burk (right) plots of initial rates of hydrolysis/phosphorolysis of (A) pNPGlcNAc: (#)1 mM, ( ) 2 mM, () 5 mM, () 10 mM and (M) 13 mM. (B) pNPGlc: (#) 5 mM, ( ) 10 mM,() 25 mM, () 35 mM and (M) 50 mM. (C) DNPGlc: (#) 0.063 mM, ( ) 0.175 mM, () 0.25mM, () 0.5 mM and (M) 1 mM. Error bars are shown on data points: where bars are not apparent,errors were smaller than the symbol size used.292.3.1 Formation of Sugar Phosphate ProductsIn order to ensure that these effects have their origins in phosphate acting as a co-substrate and notsimply as an allosteric activator, the reactions were monitored by TLC and NMR and the productsthereby identified. As can be seen in Figure 2.2, reaction of all three substrates in the presenceof increasing phosphate concentrations results in a change in the product formed from the freesugar (glucose or N -acetylglucosamine) to a product with a very similar TLC mobility to that of α-glucose 1-phosphate. This finding is consistent with the enzyme acting as a glycoside phosphorylasewithin or indeed below the physiologically relevant phosphate concentration range (20-30 mM)[208]. Confirmation of the formation of a sugar 1-phosphate in each case and determination of theanomeric stereochemistry of that sugar phosphate was achieved by isolating the sugar phosphateand analyzing it by mass spectrometry and NMR. As can be seen in Figure 2.2, the productsare the β-anomers: β-glucose 1-phosphate and β-N -acetylglucosamine 1-phosphate, with anomericstereochemistries being confirmed by the large J 1,2 couplings of 7.9 and 8.3 Hz seen for Glc-β1-P andGlcNAc-β1-P respectively. These results therefore confirm that Nag3 functions as a phosphorylasein each case and further that it is a β-retaining phosphorylase; the first such enzyme identified.Indeed, not only is this the first reported example of a retaining β-glycoside phosphorylase, butalso this would represent the first report of the occurrence of GlcNAc-β1-P itself in a biochemicalsystem. The GlcNAc-β1-P linkage occurs within the undecaprenyl-phosphate donor sugars involvedin the assembly of lipoteichoic acids, but GlcNAc-β1-P itself is not involved in their biosynthesis[209]. While I had initially assumed it would be an extremely labile derivative, thus hard to isolatedue to the potential for facile oxazoline formation, it proved to be reasonably stable, as indeedreported by Martin and co-workers for their synthetic material [210].2.3.2 Determination of Kinetic ParametersPresumably this β-retaining glycoside phosphorylase also follows a double displacement mechanismlike that of the β-retaining glycosidases, but wherein phosphate serves as the ultimate nucleophilein place of water, attacking the glycosyl-enzyme intermediate as shown in Figure 2.3. In order toconfirm this, kinetic parameters for the phosphorolysis and hydrolysis reactions were measured for30 0 0 1 5 10 25 50 100 SM Glc Glc-1-P αGlc-1-P 0 0 1 5 10 25 50 100 Glc SM GlcNAc GlcNAc-1-P 0 0 1 5 10 25 50 100 [phosphate] (mM) : GlcNAc Nag3 : - + - + - + pNPGlcNAc pNPGlc DNPGlc 100 mM 25 mM 5 mM 0 mM [phosphate] SM SM SM β-GlcNAc-1-P α-GlcNAc β-Glc-1-P α-Glc β-Glc-1-P α-Glc 1-PGlc-α-l c -GlcNAc-β l -β1-P Glc-β -Figure 2.2: 1H NMR and TLC analysis of Nag3 products. (top panels) 1H NMR spectra for Nag3and 10 mM pNPGlcNAc, pNPGlc and DNPGlc with 0, 5, 25 and 100 mM phosphate. Insets show4x amplification of the α-Glc/α-GlcNAc peak regions. Unfortunately the β-GlcNAc and β-Glcanomeric proton falls right under the large peaks from residual HOD, thus only the peaks fromα-anomers are shown. (bottom panels) TLC analysis of Nag3 hydrolysis and phosphorolysis of 10mM pNPGlcNAc, pNPGlc and DNPGlc with 0, 1, 5, 10, 25, 50 and 100 mM phosphate. The arylglycoside starting materials are labeled SM.31each substrate and these kinetic parameters are presented in Tables 2.1 and 2.2. As can be seen,values of both kcat and Km for pNPGlcNAc increase with phosphate concentration in a saturablefashion, according to an apparent Km value for phosphate of 17 mM (Table 2.1). This behaviour isconsistent with phosphate increasing the deglycosylation rate constant (pKa) up to a limiting valueof approximately 0.2 s-1. Values of kcat/Km remain constant, consistent with the fact that kcat/Kmreflects the first irreversible step, which in this case will be the formation of the glycosyl-enzymeintermediate with concomitant loss of the phenolate. This behaviour is much more extreme for theglucosides, with kcat and Km values also increasing with phosphate concentration, but with little signof apparent saturation within reasonable substrate concentrations. Again kcat/Km values remainroughly constant. Notably the Km value for DNPGlc in the absence of phosphate is remarkablylow, at approximately 3 µM, while the kcat value is also very low at 0.04 s-1. This is the expectedbehaviour for cleavage of a substrate containing an excellent leaving group by an enzyme followingsuch a double-displacement mechanism, for which the Km is given by the expression:Km =(k−1 + k2k1)(k3k2 + k3)(2.1)In this scenario, as k3 decreases relative to k2 the Km value will drop. Since DNP (pKa = 4) isa much better leaving group than pNP (pKa = 7), it is reasonable that k2 for DNPGlc will beless than that for pNPGlc. Since it seems that k3 is already the rate-limiting step, based uponOOROO[catalytic nucleophile]AOOOOR'OHOPOOOGlycosyl-enzymeintermediate[acid/base residue]E + GORHOHOYOHHOHOYOHHOHOOHYOHOHOHOOHYR' = HR' = PO32-k1k-1k2k3Pk3WG = Glc or GlcNAcY = OH or NAcR = Sugar, DNP or pNPR' = H or PO32-R-OHHAFigure 2.3: Mechanistic scheme for a retaining β-glycoside phosphorylase-glycosidase. For Nag3 k2has a similar value for pNP-glycosides where Y = OH and Y = NAc. Values of k3W = 0.05 s-1 arefound for both Y = OH and NAc. Values of k3P = 0.2 s-1 are found for Y = NAc, while k3P isgreater than 100 s-1 for Y = OH32Table 2.1: Apparent kinetic parameters for phosphate activation of substrate cleavage by Nag3.Apparent values of Km and Ki were calculated by fitting the rate data shown in Figure 2.1 to theMichaelis-Menten equation incorporating substrate inhibition. Ka and Kb values were calculatedby fitting the rate data to the equation for a Ping Pong kinetic mechanism. Equations can be foundin section 7.3.2.Substrate Km KiKb(NP-sugar)Ka(phosphate)mM mM mM mMpNPGlcNAc 17 ± 6 400 ± 200 4.7 ± 1 6 ± 2.4pNPGlc 0.3 ± 0.05 210 ± 30 64 ± 5 0.7 ± 0.06DNPGlc 3.0 ± 0.3 320 ± 50 0.43 ± 0.03 0.8 ± 0.05Table 2.2: Kinetic parameters for the reaction of Nag3 with pNPGlcNAc, pNPGlc and DNPGlc.The reactions were carried out in 50 mM HEPES buffer (pH 7.0) and the indicated concentrationof potassium phosphate (pH 7.0) at 25 °C. Values of kcat/Km for pNPGlc were measured at con-centrations of phosphate from 5 to 50 mM and ranged between 0.034 and 0.037 s-1mM-1: high Kmvalues precluded measurement of individual parameters for kcat and Km.Substrate [Pi] Km kcat kcat/KmmM mM s-1 s-1mM-1pNPGlcNAc 0 1.6 ± 0.2 0.05 ± 0.002 0.03210 2.7 ± 0.2 0.07 ± 0.002 0.02725 3.6 ± 0.2 0.09 ± 0.001 0.02650 4.3 ± 0.3 0.10 ± 0.003 0.023100 5.7 ± 0.5 0.15 ± 0.006 0.027200 7.3 ± 0.9 0.16 ± 0.01 0.022pNPGlc 0 2.0 ± 0.5 0.04 ± 0.002 0.0200.1 8.9 ± 0.6 0.43 ± 0.01 0.0480.5 45 ± 6 2.0 ± 0.1 0.0441 149 ± 18 6.4 ± 0.6 0.0432.5 190 ± 72 8.5 ± 2.7 0.045DNPGlc 0 0.003 ± 0.001 0.04 ± 0.001 130.1 0.09 ± 0.02 3.1 ± 0.1 320.5 0.20 ± 0.01 8.2 ± 0.2 401 0.23 ± 0.02 10 ± 0.3 452 0.29 ± 0.03 13 ± 0.5 455 0.28 ± 0.02 14 ± 0.4 4910 0.34 ± 0.03 16 ± 0.7 4825 0.42 ± 0.04 19 ± 0.9 4550 0.55 ± 0.06 23 ± 1.3 4233essentially identical values of kcat for pNPGlc and DNPGlc, a very low Km value is expected forDNPGlc.Interestingly, as noted earlier, kcat for pNPGlcNAc also has a similar value, thus the presenceof the acetamide has little effect on the hydrolytic deglycosylation step. However this is clearlynot the case for the phosphorolytic deglycosylation step, where k2 for phosphorolysis of the N -acetylglucosaminyl enzyme intermediate has a value of 0.2 s-1, only 4-fold higher than that forhydrolysis, while k3W for hydrolysis of the glucosyl enzyme intermediate is 8.5 s-1, thus more than200 fold higher than that for hydrolysis.Reciprocal plots of the data from which kinetic parameters for each substrate in Table 2.2 arederived, yields the three plots shown in the right hand column of Figure 2.1. These plots, with theirparallel lines, are diagnostic of a ping pong mechanism. The presence of substrate inhibition byphosphate leads to the deviations observed, which are particularly apparent at low pNP-glycosideconcentrations. This kinetic behaviour is what is predicted for a mechanism involving a covalentglycosyl enzyme intermediate, but is not seen for glycosidases due to the fact that the concentrationof the second substrate, water, cannot realistically be altered.2.3.3 Observation of Covalent Intermediate AccumulationThis interpretation of these kinetic parameters would mean that the glucosyl-enzyme intermediatehas a half-life of approximately 17 s under these conditions, suggesting that a pre-steady state ‘burst’should be observable in a standard UV-Vis spectrometer, especially if the experiment is performedat a lower temperature to slow deglycosylation further. DNPGlc was chosen as the substrateto test this since it has the best leaving group (2,4-dinitrophenolate) with the highest extinctioncoefficient. Incubation of high concentrations of Nag3 with DNPGlc in a spectrophotometer whilefollowing the absorbance at 400 nm produced a clear burst phase, followed by a steady state,as shown in Figure 2.4. The magnitude of this initial burst, once converted to molar terms usingthe extinction coefficient for DNP, corresponded perfectly with the enzyme concentration employed.Subsequent addition of phosphate (after∼ 0.5 min) resulted in a very steep increase in rate due to theacceleration of the deglycosylation step through phosphorolysis of the glycosyl-enzyme intermediate.Final leveling off of absorbance increase was due to substrate depletion.34Further confirmation of the accumulation of a glucosyl-enzyme intermediate was obtained byuse of electrospray ionization mass spectrometry to directly observe this intermediate, as shown inFigure 2.5. The mass of the enzyme incubated in the presence of DNPGlc (Figure 2.5B) is clearlyseen to be greater than that of the free enzyme (Figure 2.5A) by the mass of a glucosyl moiety,confirming the steady state accumulation of a covalent glucosyl-enzyme intermediate. Interestingly,inclusion of phosphate in this reaction mixture leads to the spectrum shown in Figure 2.5 C, where amixture of free enzyme and glucosyl-enzyme intermediate are seen, consistent with the stimulationof turnover via phosphorolysis, leading to a lower steady state concentration of the glycosyl-enzymecomplex.2.3.4 Is Phosphate the Natural Nucleophile?One possible concern might be that the reaction with phosphate is just a reflection of a general“anion rescue” analogous to that seen with designed acid/base mutants of retaining glycosidases,where azide or acetate have proven to be the most effective [201, 211]. This possibility was tested bymeasuring rates at saturating DNPGlc, for which deglycosylation is rate-limiting, in the presenceof a variety of anions. As shown in Figure 2.6 phosphate is indeed the best reactivator, by awide margin. By contrast, phosphate has never been seen to function effectively in anion rescue ofFigure 2.4: Pre-steady state burst of DNP released by Nag3. Pre-steady state burst of DNP releasedby Nag3 upon incubation at 12 °C with 250 µM DNPGlc, followed by activation with 50 mMphosphate. Solid arrow indicates addition of 33 µM Nag3. Dotted arrow indicates addition of 50mM phosphate.35Figure 2.5: Trapping the Nag3 covalent intermediate. Electrospray mass spectra of (A) Nag3, (B)Nag3 + 1 mM DNPGlc and (C) Nag3 + 1 mM DNPGlc + 50 mM phosphate incubated at roomtemperature for 1 h. The mass shift (162 Da) of the peak of the glycosyl-enzyme (calculated MW= 61107.9 Da; observed MW = 61109.0 Da) shown in (B) compared to the peak of the enzyme only(calculate. MW = 60944.9 Da; observed MW = 60947.0 Da) shown in (A) is consistent, withinerror, to a glucosyl residue (163 Da) covalently bound to Nag3.36glycosidase mutants. Phosphate would therefore seem to be the natural substrate for this enzyme,thus Nag3 should be re-classified as a phosphorylase.Further insights into the role and importance of the phosphate reaction were obtained by trap-ping the reaction intermediate as the 2-fluoroglucosyl enzyme species by reacting the enzyme withDNP2FGlc and then measuring reactivation of the purified species in the presence of phosphate.As can be seen in Figure 2.7, phosphate indeed stimulates the turnover of this species, and in aconcentration-dependent manner. However, the instability of Nag3, along with the tendency of phos-phate to stabilize it, unfortunately makes the extraction of definitive kinetic parameters difficult.Nonetheless it is clear that even the lowest phosphate concentrations studied lead to substantialreactivation.2.3.5 Nag3 as a Phosphorylase and its RoleAs is seen most clearly in Figure 2.2, once the phosphate concentration approaches 10 mM the sugarphosphate is the almost exclusive product, with essentially no hydrolysis observed, whether thesugar donor is a glucoside or an N -acetylglucosaminide. Importantly it should be noted that, evenFigure 2.6: Nag3 activator screen. Seven anionic salts were tested as activators of DNPGlc cleavageby Nag3. 50 mM D-glucuronic acid, sodium acetate, sodium citrate, sodium formate, potassiumsulfate, sodium azide or potassium phosphate were incubated with 0.1 µM Nag3 in 50 mM HEPESbuffer containing 500 µM DNPGlc for 5 minutes at 25 °C.37though artificial aryl glycosides were employed to detect and characterize this behaviour, exactlythe same product partitioning would be expected whatever the nature of the donor glucoside or N -acetylglucosaminide, since the glycosyl-enzymes formed are identical in each case. Thus, even thoughthe identity of the natural substrate for Nag3 is not clear despite considerable investigative efforts[193] repeated again here, its cleavage will occur via phosphorolysis under normal physiologicalconditions.The question now becomes, why have these enzymes evolved to function as phosphorylases, andwhat is the fate of the β-phosphate products so formed? A clue to this, at least for C. fimi, comesfrom the presence within the same operon of a gene for a β-phosphoglucomutase. This enzymewould convert the β-phosphate into the corresponding sugar-6-phosphate which would then feeddirectly into metabolism.2.3.6 Structural and Mechanistic ImplicationsOrdinarily glycoside hydrolases do not also function as phosphorylases, presumably in large partbecause the presence of the catalytic Glu and/or Asp residues provides Coulombic screening againstbinding of anions. This would be particularly important within the cellular milieu since otherwiseFigure 2.7: Phosphate rescue of inactivated Nag3. Inactive 2FGlc-Nag3 was reactivated by incuba-tion with increasing concentrations of phosphate. Samples were incubated at 25 °C in 0 mM ( ), 25mM (3), 50 mM (), 100 mM (M) and 200 mM (#) phosphate. Aliquots were assayed in HEPESbuffer containing 500 µM DNPGlc and 20 mM phosphate.38the resident glycosidases would be constantly degrading key metabolic intermediates. Indeed thisis likely one of the reasons why carboxylic amino acids have assumed these roles in most glycosi-dases. It would appear that this subset of GH3 glycosidases have broadened their repertoire toallow phosphate into the active site by replacing the acid/base catalytic glutamate with a histi-dine residue, itself activated by an adjacent, but presumably sufficiently electrostatically remote,aspartate residue. This strategy of charge-avoidance has been seen previously in, for example,some sialyltransferases, where the catalytic base is also a histidine residue to avoid electrostaticinteractions with the anionic substrate CMP-sialic acid, as shown by structural and NMR studies[212]. Even more relevant are the inverting GH65 α-glycoside phosphorylases, which are closelystructurally homologous to the inverting glucoamylases of GH15. In this case the phosphate moietyis bound in the GH65 phosphorylase at a location entirely equivalent to that of the general base,Glu-400, of a corresponding GH15 glucoamylase [97].The kinetic parameters measured reveal that phosphate has no noticeable effect on the glycosyl-ation rate constant, as reflected in kcat/Km. Only an inhibitory effect is seen, and this only athigh concentrations as seen in Figure 2.1. This would be consistent with phosphate needing tocompete for binding in the +1 site but only doing so weakly. Comparison of kcat/Km values forpNPGlcNAc and pNPGlc, which have identical leaving groups, reveals that the presence of OH orNAc at C-2 makes no real difference to the glycosylation rate constant. This was also the casefor the hydrolytic deglycosylation rate constants, where both have values of 0.05 s-1. However thefinding that phosphate only increases the deglycosylation rate constant for the N -acetylglucosaminylenzyme some 4-fold, while it accelerates the breakdown of the glucosyl enzyme many hundred-folddemands some explanation, which will be revisited in Chapter 3.2.4 ConclusionsThe discovery that Nag3 acts as a phosphorylase rather than a hydrolase provides a simple explan-ation for the use of a His-Asp dyad as acid-base catalyst rather than the standard Glu or Asp residuesby this sub-group of GH3 enzymes. Approach of an anionic phosphate to an active site bearinga deprotonated carboxylate base directly adjacent to the anomeric centre of the glycosyl enzyme39would be greatly disfavoured but is avoided in this way. The presence of the His-Asp catalyticdyad may be necessary to permit phosphorylase activity for GH3 hexosaminidases, however, astudy done by Ducatti et al. following the completion of the Nag3 research indicated that thepresence of the dyad is not the sole determining factor to impart phosphorylase activity [213].They demonstrated that a GH3 β-N -acetylglucosaminidase from H. seropedicae, which possessedthe His-Asp dyad, acted only as a hydrolase, as the presence of phosphate did not significantlyimpact the kinetic parameters of the reaction. I later confirmed this result by testing the dyadcontaining GH3 β-N -acetylglucosaminidase from E. coli (NagZ), which also failed to be impactedby the presence of phosphate. This topic will be re-examined in Chapter 3 with the discoveryand characterization of the second GH3 GP, BglP. Nevertheless, Nag3 represents the first reportedexample of a retaining β-glycoside phosphorylase, and indeed to our knowledge, this represents thefirst report of the formation of free GlcNAc-β1-P in a biological system, though chemical syntheseshave been reported. The presence of a β-phosphoglucomutase within the same operon as Nag3 inC. fimi suggests that the β-1-phosphates so produced are directly converted to their 6-phosphatesfor further metabolic conversion.40Chapter 3Functional Metagenomic ScreenTargeting Forward PhosphorolysisActivity3.1 SummaryGlycoside phosphorylases have considerable potential as catalysts for the assembly of useful gly-cans for products ranging from functional foods and prebiotics to novel materials. However, thesubstrate diversity of currently identified phosphorylases is relatively small, limiting their practicalapplications. To address this limitation, I developed a high-throughput screening approach us-ing the activated substrate 2,4-dinitrophenyl β-D-glucoside (DNPGlc) and inorganic phosphate foridentifying glycoside phosphorylase activity and used it to screen a large insert metagenomic library.The initial screen, based on release of 2,4-dinitrophenol from DNPGlc in the presence of phosphate,identified the gene bglP, encoding a retaining β-glycoside phosphorylase from the CAZy GH3 fam-ily. Kinetic and mechanistic analysis of the gene product, BglP, confirmed a double displacementping-pong mechanism involving a covalent glycosyl–enzyme intermediate. X-ray crystallographicanalysis provided insights into the phosphate-binding mode and identified a key glutamine residuein the active site that is important for substrate recognition. Substituting this glutamine for a serineswapped the substrate specificity from glucoside to N -acetylglucosaminide. In summary, presentedin this chapter is a high-throughput screening approach for identifying β-glycoside phosphorylases,which is robust, simple to implement, and useful in identifying active clones within a metagenomicslibrary. Implementation of this screen enabled discovery of a new glycoside phosphorylase and has41paved the way to devising simple ways in which enzyme specificity can be encoded and swapped,which has implications for biotechnological applications.3.2 BackgroundCarbohydrate active enzymes (CAZymes) are the biocatalysts responsible for the assembly, de-gradation and modification of glycans in biological systems [30]. They are also widely employedenzymes in industry, being used in brewing and food processing, animal feed preparation, industrialpulp and paper applications and increasingly in biofuel and bioproduct development [31, 214–216].While the use of CAZymes is cost-effective in glycan degradation, glycan assembly generally requiresthe use of expensive starting materials, such as nucleotide phosphosugars [217]. The high-cost ofthese materials makes de novo industrial-scale glycan synthesis difficult and usually non-viable.Glycoside phosphorylases (GPs) offer a potential solution to the high costs typically associated withenzymatic glycan synthesis and are increasingly being recognized and used for the biocatalysis andbiotransformation of glycans [33, 34, 143]. GPs ordinarily carry out phosphorolysis by transferringa glycosyl moiety from the non-reducing end of a di- or polysaccharide substrate onto inorganicphosphate, thereby generating a sugar 1-phosphate [142] (Figure 3.1). GPs distinguish themselvesFigure 3.1: Generalized phosphorylase and hydrolase reactions. Generalized phosphorolysis (upperpathway) and hydrolysis (lower pathway) of β1-4 linked glycans. R = H or glucosen (n = numberof glucose residues). GPs are distinguished from GHs by their use of a phosphate molecule to cleavethe glycosidic linkage thereby producing a free sugar-1-phosphate.from most CAZymes in that the hydrolytic free energy associated with the glycosidic ester linkage42of the sugar 1-phosphate product is roughly equivalent to that of the glycosidic linkage in theglycan substrate. Accordingly, the equilibrium constant for the reactions catalyzed by most GPs isclose to, or is one, thus the equilibrium position can be tipped in favour of glycoside synthesis bymanipulation of reaction conditions [164, 218, 219].Functional metagenomics screening offers a means to search for novel biocatalysts from geneticmaterial drawn directly from the environment [165]. Metagenomic libraries provide access to thevast reservoir of uncultivated genetic diversity encoded in microbial communities inhabiting naturaland engineered ecosystems [167, 220, 221]. Emerging sophisticated sequence- and function-basedscreening technologies are being deployed to identify novel enzymes, including CAZymes, withinthese libraries [167–170]. However, to our knowledge no function-based metagenomics screeningapproach has yet been developed that targets GP-encoded genes. A central challenge in designingthe needed screen is that of distinguishing phosphorylase activity from the related hydrolase activityin a high-throughput manner (Figure 3.1).Reported in this chapter is the development and implementation of such a high-throughputscreening approach to identify GP activity. By screening of a library derived from a passive minetailings biochemical reactor system (BCR) fed with lignocellulosic biomass [176], I identified apreviously unknown β-retaining GP from CAZy family GH3, and I report on its structural andmechanistic characterization. Our screening approach is based on the use of activated aryl glyco-sides as easily monitored substrates for glycoside phosphorylases when deployed in the presence ofphosphate. The BCR library is a large insert (fosmid) library with an average insert size of 40kilobases first described by Mewis and colleagues [176]. The authors used the BCR library to searchfor novel cellulose-degrading enzymes. Therefore, in testing the parallel screening approach to iden-tify GPs, I focused in this case on identifying fosmid clones encoding enzymes with the capacity ofphosphorolysing the β 1,4-glucosidic linkages of cellulose and cello-oligosaccharides.43Figure 3.2: Phosphorolysis of cellobiose or cello-oligosaccharides. Phosphorolysis of cellobiose orcello-oligosaccharides performed by the inverting phosphorylases HtCBP (R = H) and HtCDP (R= glucosen).3.3 Results3.3.1 Development and Testing of the Metagenomic ScreenThe design of the screen was based upon the notion that, since many GPs are members of CAZy GHfamilies, they might accept an activated aryl glycoside substrate, transferring the glycosyl moietyto added phosphate. This concept has not been tested previously, to our knowledge, apart from onedemonstration with a nucleotide phosphorylase [222], though the concept has parallels with previouswork on nucleotide-dependent glycosyltransferases [223]. 2,4-Dinitrophenyl glucoside (DNPGlc) wasused to test this idea since it is highly activated (pKa of 2,4-dinitrophenol = 4.0) [224] and has provedto be a near universal substrate for glucosidases. Further, the low pKa of the phenol allows direct,continuous assays to be performed at pH values down to below 4 without the need to add base in astopped manner. The choice of a glucoside was based on the cellulolytic origin of the library to bescreened, which thus might contain cello-oligosaccharide-degrading phosphorylases, especially sincea number of cellulases and β-glucosidases had already been identified within this library [176].In order to test whether glycoside phosphorylases can accept DNPGlc as a substrate in thepresence of phosphate, and catalyze glucosyl transfer to generate β-glucose 1-phosphate (Glc-β1-P), three known GPs that cleave cellobiose or cellodextrins were cloned and expressed. Two of theseGPs, cellobiose phosphorylase [225] (HtCBP) and cellodextrin phosphorylase [68] (HtCDP) fromHungateiclostridium thermocellum (formerly known as Clostridium thermocellum) belong to CAZyfamily GH94. These enzymes use a β-inverting mechanism in which phosphate attacks directlyon the sugar anomeric center, forming α-glucose 1-phosphate (Glc-α1-P) (Figure 3.2). The third44enzyme, the β-glucosidase/N -acetylglucosaminidase Nag3 from Cellulomonas fimi [85, 193], belongsto CAZY family GH3 and uses a β-retaining mechanism involving a covalent α-glycosyl-enzymeintermediate that is then attacked by phosphate, forming a β-glucose 1-phosphate (Glc-β1-P) (orβGlcNAc-1-phosphate) product (Figure 3.3).Figure 3.3: Proposed mechanistic scheme for a retaining β-glycosidase/phosphorylase. Proposedmechanistic scheme for a retaining β-glycosidase/phosphorylase. BglP and Nag3 both employ adouble-displacement β-retaining mechanism involving a glycosyl-enzyme intermediate and act aspreferential phosphorylases (k3P > k3W). Both enzymes possess the same active site residues whichact as the catalytic nucleophile (Asp) and the acid/base catalytic dyad (Asp and His).As shown in Figure 3.4A all three enzymes cleave DNPGlc in the presence of phosphate, as canbe observed by monitoring the increase in absorbance at 400 nm. Importantly, all three enzymesare only minimally active in the absence of phosphate. Confirmation that the rate stimulation isdue to phosphorylase action was provided by TLC analysis of reaction mixtures, which revealedthat all three catalysed the transfer of the glucosyl moiety of DNPGlc to inorganic phosphatethereby forming Glc1-P (Figure 3.4B). These findings therefore validate the concept of the screen:(1) Lysed extracts from metagenomic clones are initially assayed using DNPGlc in the presence ofphosphate to detect both GHs and GPs. (2) Those clones that show activity are then rescreened inthe absence of phosphate to weed out the glycosidases. (3) Since it is also possible that phosphatecould stimulate a GH activity through a specific binding effect of some sort, or simply activation45Figure 3.4: Phosphate-dependent cleavage of DNPG by HtCBP, HtCDP and Nag3. Phosphate-dependent cleavage of DNPG by HtCBP, HtCDP and Nag3. (A) HtCBP, HtCDP or Nag3 wasincubated with 2 mM DNPGlc in 0 and 50 mM phosphate for 1 h at 37 °C then absorbance wasmeasured at 400 nm. Error bars indicate standard deviation (n = 3). (B) HtCBP, HtCDP or Nag3was incubated with 20 mM DNPGlc in 0 and 50 mM phosphate and incubated for 2 h at 37 °C thensamples spotted on TLC. The glycerol visible on the TLC plate is from the enzyme storage buffer.at higher ionic strength, leading to false positives, TLC analysis of this much smaller number ofreaction mixtures is used to confirm the presence of the sugar phosphate product. (4) Fosmid clonescapable of producing Glc1-P are sequenced and analysed to identify the open reading frames (ORFs)responsible for the observed GP activity. An overview of the screening process is provided in Figure3.5.3.3.2 Functional ScreenThe BCR fosmid library was constructed using E. coli EPI300 as an expression host and contained18,048 clones in 47 x 384-well plates [176]. Following replication of the BCR master library, atotal of 880 clones failed to grow, leaving 17,168 clones to be screened. Functional screening usingDNPGlc in the presence of phosphate yielded 54 active clones with activity greater than the mean+ 4 SD, a hit rate of 0.31 % (Figure 3.6B). To distinguish clones with GP activity from those that46Figure 3.5: Overview of functional metagenomic screen targeting forward phosphorolysis. WHOLELIBRARY : Metagenomic library in 384-well plate format was screened in the presence of DNPGlcand phosphate. Clones that gave an A400 value greater than mean + 4 SD were consolidated andre-arrayed into a master library. CONSOLIDATION : Master library was screened with DNPGlc inthe presence (x3) and absence (x3) of phosphate. ANALYSIS : A Student’s t-test was performedbetween the averages of the A400 values of each clone in the absence and presence of phosphate.Clones possessing a significantly (p < 0.05) higher activity with phosphate present were analyzedfor the production of glucose 1-phosphate using TLC.47Figure 3.6: Forward phosphorolysis functional screen results. (A) Reaction underlying the functionalmetagenomic screen. (B) 400 nm absorbance measurements of 17168 metagenomic ct-testlonesassayed in the presence of phosphate. White dashed line indicates mean (0.66); blue dashed lineindicates mean + 4 SD (0.98). 54 clones displayed an A400 value greater than the mean + 4 SDcutoff, indicating cleavage of the DNPGlc substrate. (C) 400 nm absorbance measurements of 54active clones in the presence and absence of 50 mM phosphate. Each clone was assayed in triplicatewith 0 (red) or 50 mM (blue) potassium phosphate. Student’s t-test was performed and those clonesmeeting the 95 % confidence threshold and showing positive phosphate-dependent activity towardDNPGlc were selected for validation (black asterix). Error bars indicate standard deviation. (D)TLC analysis of the 12 clones displaying phosphate-dependent activity towards DNPG. Cell lysateswere treated with 20 mM DNPGlc and 0 or 50 mM potassium phosphate then incubated for 2 h at37 °C. S lane: 20 mM Glc1-P, 20 mM Glc, 20 mM DNPG.48Figure 3.7: 29K06 and 31P01 fosmid ORF alignment. ORFs were predicted using Prodigal andannotated with BLASTP. See Supplemental Table B.3 for the BLASTP functional annotations.The GH3 ORFs (bglP) are shown in red and all others in light blue.are GHs the 54 hits were re-arrayed into a 96-well master plate and rescreened both in the presenceand absence of phosphate (Figure 3.6C). Each assay condition was run in triplicate and positivephosphate dependency was determined by performing a t-test between a clone’s A400 values in 0 and50 mM phosphate. A confidence level of 95 % (p-value < 0.05) was set as a threshold to determinewhether to further validate the clone. Of the 54 clones from the master library, 12 displayed activityin the presence of phosphate exceeding the set threshold. The reaction products of these 12 cloneswere then examined by TLC analysis (Figure 3.6D). Of the 12 clones so analyzed, two were foundto produce Glc1-P in the presence of phosphate (29K06 and 31P01).3.3.3 Identification of Active Glycoside PhosphorylasesComplete sequencing of the two fosmid clones revealed they were contiguous with one another,overlapping with 23,247 bp at 100 % identity (Figure 3.7). Thirty-one open reading frames (ORFs)were predicted from the 29K06 clone and 32 from 31P01. BLASTX query of these against the CAZydatabase revealed a novel GH3 ORF (hereafter referred to as bglP) located within the overlappingregion of the two fosmids. Based on amino acid sequence analysis (Figure 3.8), bglP was predicted toencode a β-glucosidase/N -acetylglucosaminidase (NagZ), from a hexosaminidase subgroup of GH3characterized by the sequence motif [195] KH(FI)PG(HL)GX4D(ST)H. However, the motif foundin bglP had substitutions at two residues (underlined): KH(FI)PGDGX4DQH. This modified motifis also present in the only other known glycoside phosphorylase in the GH3 family, Nag3 [85, 193].In order to confirm that the activity detected was indeed associated with this gene it was subclonedinto a pET expression vector and heterologously expressed and purified. Indeed, activity assays ofpurified enzyme confirmed that BglP catalyzes the phosphate-dependent cleavage of DNPGlc seen49Figure 3.8: Amino acid alignment of C. fimi Nag3 and BglP. The conserved GH3 hex-osaminidase subgroup sequence motif is indicated (*, bold letters) and the catalytic nucle-ophile residues are denoted (•). Black shading indicated highly conserved residues and greyshading indicates conserved similar residues. Sequence alignment was generated using T-Coffee (http://tcoffee.crg.cat/apps/tcoffee/do:regular) and shading was done in BoxShade v3.21(http://www.ch.embnet.org/software/BOX_form.html) by K. Hofmann and M. Baron.50from the 29K06 and 31P01 source clones (Figure 3.9).Figure 3.9: Purified BglP TLC activity assay: Phosphate dependent cleavage of DNPGlc, pNPGlcand pNPGlcNAc. BglP was incubated with 20 mM DNPGlc, pNPGlc or saturated pNPGlcNAc in0 and 50 mM phosphate and incubated for 20 min (or 16 h for pNPGlcNAc) at room temperaturebefore being spotted on TLC. Left panel was treated with molybdate stain and right panel withp-anisaldehyde.3.3.4 Kinetic and Mechanistic Characterization of BglPThe other known GP in the GH3 family, C. fimi Nag3, has been subjected to mechanistic character-ization to probe its proposed two-step double-displacement mechanism and confirmed to produceGlc-β1-P and GlcNAc-β1-P, though no 3-D structure has been obtained. In fact, Nag3 was the firstretaining β-glycoside phosphorylase described and, like BglP, was shown to also catalyze hydrolysis,but at a reduced rate relative to phosphorolysis [85]. Comparison of their amino acid sequences(Figure 3.8) reveals a high degree of sequence similarity between BglP and Nag3 (55 % similarityand 39 % identity), making it probable that the two enzymes follow the same mechanism involvinga covalent α-glucosyl enzyme intermediate (Figure 3.3). In order to confirm this and to allowcomparison of the two enzymes, a kinetic and mechanistic analysis of BglP was performed.Kinetic parameters were determined for cleavage of three different substrates, para-nitrophenylβ-glucoside (pNPGlc), 2,4-dinitrophenyl β-glucoside (DNPGlc) and para-nitrophenyl β-N -acetyl-glucosaminide (pNPGlcNAc), both in the absence of phosphate, and in the presence of increasing51concentrations (Figure 3.10). Based upon both kcat/Km and kcat values (Table 3.1) it is apparentthat BglP prefers glucoside substrates over N -acetylglucosaminides, kcat/Km values for pNPGlcbeing approx. 10-fold higher and kcat values ∼ 100-fold higher than those of the hexosaminide. Asecond distinction of the two substrate classes is that kcat for the two glucoside substrates increasesover 10-fold as phosphate concentrations are raised, whereas increases for pNPGlcNAc are muchmore modest at 1.5 - 2-fold. Km values also increase with phosphate, by up to 20 - 30-fold for theglucosides, with the net consequence being that kcat/Km values remain approximately constant.These observations for the glucosides reflect classic kinetic behaviour for a ping-pong mechanismFigure 3.10: Phosphate activation of substrate cleavage by BglP. Phosphate activation of BglP-mediated cleavage of (A) DNPGlc, (B) pNPGlc and (C) pNPGlcNAc. Michaelis-Menten plots ofinitial hydrolysis/phosphorolysis rates of (A) 1.5 mM DNPGlc, (B) 50 mM pNPGlc and (C) 5 mMpNPGlcNAc in the presence of increasing concentrations of phosphate. Error bars indicate standarderror of the mean; where bars are not present, the error is smaller than the symbols used. Kineticparameters for cleavage of the aryl glycoside substrates at each concentration of phosphate can befound in Table 3.1.52Table 3.1: Kinetic parameters for the reaction of BglP with pNPGlcNAc, pNPGlc and DNPGlc.Substrate [Pi] Km kcat kcat/KmmM mM s-1 s-1mM-1pNPGlcNAc 0 0.3 ± 0.1 0.002 ± 0.0001 0.0065 0.4 ± 0.1 0.002 ± 0.0003 0.00510 0.6 ± 0.1 0.002 ± 0.0004 0.00425 1.1 ± 0.4 0.004 ± 0.001 0.00350 0.8 ± 0.2 0.003 ± 0.0005 0.004100 0.9 ± 0.3 0.003 ± 0.0007 0.003200 1.5 ± 0.5 0.004 ± 0.001 0.003pNPGlc 0 3 ± 1 0.22 ± 0.03 0.0650.01 5 ± 1 0.32 ± 0.02 0.0720.05 9 ± 1 0.55 ± 0.03 0.0650.5 53 ± 5 2.5 ± 0.2 0.0475 67 ± 11 3.2 ± 0.4 0.04810 55 ± 5 2.4 ± 0.2 0.045100 56 ± 7 2.3 ± 0.2 0.041DNPGlc 0 0.06 ± 0.01 0.7 ± 0.01 10.70.01 0.06 ± 0.01 0.7 ± 0.02 11.10.05 0.17 ± 0.01 1.7 ± 0.03 9.80.1 0.29 ± 0.02 3 ± 0.1 9.10.5 2.1 ± 0.2 15 ± 1 7.15 ND ND 7.625 ND ND 7.450 ND ND 7.4ND: not determined.with substrates for which the second step (cleavage of the glycosyl enzyme) is rate-limiting (Figure3.3) [226]. Introduction of an alternate, better, nucleophile (phosphate) into the reaction acceleratesthe decomposition of the glycosyl enzyme intermediate through provision of a second pathway (k3P).The net effect is to not only raise kcat values, but also to decrease the accumulation of the glycosyl-enzyme intermediate, thereby raising the Km value. By contrast the kcat/Km value, which can beexpressed in terms shown in Equation 3.1, reflects the first irreversible step and is thus unaffectedby steps occurring later in the pathway.kcatKm=k1k2k−1 + k2(3.1)53Figure 3.11: BglP pre-steady state burst phase analysis. BglP pre-steady state burst phase analysisof the cleavage of DFMUGlc (red) and DFMUGlcNAc (grey). DFMUGlc (10 µM) or DFMUGlcNAc(10 µM) were incubated in a fluorimeter pre-chilled to 12 °C. DFMU release was measured as afunction of time after the addition of 42 nM (for DFMUGlc) or 420 nM (for DFMUGlcNAc) BglPat approx. 1 min. Fluorescent signal was converted to DFMU concentration using a standard curveof free DFMU at pH 7.0.These kinetic studies therefore strongly support the presumed ping-pong mechanism for BglP. Theyalso suggest that, for the glucosides tested, and certainly for DNPGlc, the rate-limiting step is theturnover of the glucosyl-enzyme intermediate. However, the smaller effect of phosphate on kcat andKm for pNPGlcNAc might suggest that the formation of the glycosyl-enzyme intermediate remains atleast partially rate-limiting for this substrate. This implies that the presence of the acetamide moietylowers reaction rates, possibly due to different interactions with the signature loop. If this is thecase then measuring the pre-steady state phase of BglP-catalyzed hydrolysis of the glucoside and N -acetylglucosaminide substrates should reveal substantial differences between the substrates. The useof 6,8-difluoromethylumbelliferyl β-glucoside (DFMUGlc) and 6,8-difluoromethylumbelliferyl β-N -acetyl glucosaminide (DFMUGlcNAc) improved signal sensitivity and substrate reactivity (phenolpKa = 4.7) [227] over their pNP counterparts. Indeed, a clear burst phase was observed for cleavageof DFMUGlc, but not for DFMUGlcNAc, supporting the above assignment of rate-determiningsteps (Figure 3.11). Direct evidence for a two-step covalent mechanism was then sought by usingelectrospray ionization mass spectrometry to detect the covalent glycosyl-enzyme intermediate. Asseen in Figure 3.12, the BglP mass increased by 163 Da, equivalent to a glucosyl moiety, upon54incubation with DNPGlc consistent with formation of the covalent intermediate. Notably, thepresence of phosphate, which induces rapid turnover of the intermediate, resulted in disappearanceof the corresponding glycosyl-enzyme mass.Having confirmed a two-step mechanism involving a glycosyl-enzyme intermediate, I sought tolengthen the lifetime of the intermediate to allow further mechanistic and structural studies. Thiswas achieved by use of 2,4-dinitrophenyl 2-deoxy-2-fluoro-β-D-glucopyranoside (DNP2FGlc) as aslow substrate for which the deglycosylation step (k3) is much slower than the glycosylation step(k2). This arises as a consequence of inductive destabilization of the oxocarbenium ion-like transitionstates by the C-2 fluorine, which slows both steps (k2 and k3), while the incorporation of an excellent(DNP) leaving group ensures that k2 > k3, thus that the intermediate accumulates [228]. Indeed,incubation of BglP with DNP2FGlc in the absence of phosphate resulted in time-dependent, pseudo-first order inactivation of the enzyme, as shown in Figure 3.13A. The rate of inactivation varied,in a saturable manner, with the concentration of the 2-fluorosugar, allowing extraction of kineticparameters for inactivation of BglP of ki = 0.17 min-1 and Ki = 32 mM (Figure 3.13C). The 2-fluoro-glucosyl-BglP (2FGlc-BglP) covalent intermediate species was shown to be mechanistically relevantby removal of excess inactivator and then measuring rates of reactivation by assaying aliquots ofthe enzyme as a function of time in buffer containing increasing concentrations of phosphate. Ascan be seen in Figure 3.13B phosphate did indeed stimulate reactivation of the enzyme in a time-dependent fashion. The plot of kP versus [phosphate] showed no saturation behaviour, revealingthat binding of inorganic phosphate is weak. However, the slope of the line yielded a value for thesecond order rate constant for reactivation of kP/KP = 3.2 x 10-5 min-1mM-1 (Figure 3.13D). Thisabsence of saturation at phosphate concentrations of 100 mM might seem to be a concern, giventhe apparent saturation binding behaviour of phosphate seen in Figure 3.10. However, it is clearfrom the difference in maximal rates observed between DNPGlc and pNPGlc in Figure 3.10A andB that the curvature seen in these plots has its origin in changes in rate-limiting step as phosphateconcentrations increase: the glycosylation rate constant (k2) for DNPGlc is greater than that forpNPGlc.55Figure 3.12: Trapping the BglP covalent intermediate. Electrospray mass spectra of BglP, BglP +10 mM DNPGlc and BglP + 10 mM DNPGlc + 50 mM phosphate (pH 7.0). The single peak in thespectrum containing untreated BglP at 64,592.9 Da is consistent with the calculated mass of BglP(64,589.2 Da). The mass shift of 162.8 Da between the two peaks in the BglP + DNPGlc panel isconsistent with the heavier peak representing the glycosyl-enzyme intermediate (BglP + 163 Da).When phosphate is present no peak shift can be detected, presumably because phosphate inducesthe intermediate to turn over too quickly.563.3.5 X-ray Crystal Structure of BglPThe crystallography work presented in this section was performed by, and presented here in col-laboration with, Ankoor Patel, Veronica Larmour and Brian Mark at the University of Manitoba.Crystals of unliganded BglP were obtained through hanging drop vapor diffusion in a mother liquorcontaining 5 mg/mL BglP and 27 % polyethylene glycol 1000, and the structure was solved to aFigure 3.13: BglP inactivation and reactivation. BglP inactivation with DNP2FGlc. (A) BglP wasincubated with increasing concentrations of the inactivator, DNP2FGlc ( , 0 mM; , 2 mM; N, 5mM; H, 10 mM; #, 20 mM; , 25 mM; M, 50 mM; O, 75 mM) for 120 min at 25 °C. Activity wasmeasured at the indicated time points in 50 mM pNPGlc and measuring absorbance at 400 nm over5 min. (B) Reactivation of the 2FGlc-BglP intermediate with phosphate. Inactive BglP (50 mMDNP2FGlc for 210 min at 25 °C) was reactivated by incubating the 2FGlc-BglP intermediate withincreasing concentrations of phosphate ( , 0 mM; , 1 mM; N, 5 mM; H, 10 mM; #, 25 mM; ,50 mM; M, 100 mM). (C) ki and Ki plot of BglP inactivation with DNP2FGlc. kincat values plottedagainst inactivator concentration fit with the Michealis-Menten equation. ki = 0.17 ± 0.02 min-1,Ki = 32 ± 6.4 mM. (D) kp/KP plot of reactivation of the 2FGlc-BglP intermediate with phosphate.kreact values plotted against phosphate concentrations. Saturation was not achieved at 100 mMphosphate, therefore only the kp/KP (0.00032 ± 2.2 x 10-5 min-1 mM-1) could be derived.57resolution of 2.1 Å (Table B.1). BglP is a 567 a.a. protein that adopts a two-domain fold (Figure3.14A) and is monomeric in solution according to gel filtration (data not shown). The N-terminaldomain (a.a. 1-392) forms a (β/α)8 TIM barrel structure typical of many glycoside hydrolasecatalytic domains, with an active site composed of residues within loops that radiate out from theC-terminal ends of the β-strands of the barrel. The C-terminal domain (a.a. 393-562) adopts anα/β sandwich that does not participate directly in catalysis, with the exception of amino acids476-487, which pack along the edge of the TIM barrel active site to stabilize a loop that emanatesfrom β-strand 5 of the barrel and bears the sequence KH(FI)PGDGX4DQH. This loop containsthe sequence that defines the GH3 sub-group to which BglP belongs (albeit with the two alterationsnoted previously) and accommodates the general acid/base His-206 (Figure 3.14B).3.3.6 Donor RecognitionTo gain structural insights into the mechanism and specificity of BglP, crystals of the enzymewere soaked with 2,4-dinitrophenyl 2-deoxy-2-fluoro-β-D-glucopyranoside (DNP2FGlc) to form thelong-lived glycosyl-enzyme intermediate that had been characterized kinetically. The structureso determined revealed clear electron density for a 2-deoxy-2-fluoro-α-D-glucosyl moiety in a 4C1chair conformation covalently bound via C1 to Asp-288 (Figure 3.14B). This residue is conservedin GH3 enzymes and is indeed suitably positioned within the BglP active site to serve as thecatalytic nucleophile forming the key reaction intermediate. The active site contains most of thesame residues identified in other GH3 enzymes [192] as being involved in substrate recognition,such as Lys-193 and His-194 of the signature sequence. Also consistent with other members of theGH3 sub-family to which BglP belongs, the imidazole of a histidine residue (His-206), is positioned∼ 5.5 Å from the carboxylate of Asp-288 on the opposite face of the hexose ring of the boundintermediate and is thus well positioned to assist in catalysis by acting as a general acid/base.It first protonates the oxygen of the scissile bond to assist leaving group departure, followed, inthis case, by activation of an incoming phosphate that reacts with the covalent glycosyl-enzymeintermediate to generate a phosphorylated product with net retained anomeric stereochemistry(Figure 3.14B). The ∼ 5.5 Å distance between Asp-288 and His-206 of BglP is consistent withglycosidases that employ a configuration-retaining mechanism [201]. One key difference, however is58Figure 3.14: Crystal structure of BglP covalently bound to 2FGlc. (A) Overall structure of BglP.The catalytic N-terminal TIM-barrel domain (a.a. 1-392) is coloured grey, whereas the C-terminaldomain (a.a. 393-562) is coloured green. (B) 2FGlc (green carbons) covalently bound to Asp-288within the BglP active site (grey carbons). The electron density Sigma-A-weighted 2Fobs-Fcalcdensity map contoured at 0.87 e/Å3. (C) Superimposition of the BglP active site (grey) with theactive sites of NagZ enzymes from Bacillus subtilis (BsNagZ) (orange carbons) (PDB: 3NVD) [192]and Burkholderia cenocepacia (BcNagZ) (pink carbons) (PDB: 4GNV). The overlay reveals thatGln-205 in BglP is replaced by Ser residues in the NagZ enzymes, enabling them to accommodatethe C2 N -acetyl group of GlcNAc.59the identity of the residue interacting with the C2-position of the sugar, this being the amino acidflanked by His-206 and Asp-204 (the so-called His-Asp catalytic dyad) in the signature sequence. Inthe GH3 GlcNAc-cleaving enzymes, such as the NagZs, the residue at that position is a serine, whilein BglP it is a glutamine, as seen in the structural overlay of Figure 3.14C wherein the 2FGlc-BglPstructure is superimposed on the structures of two other GH3 β-N -acetylglucosaminidases (NagZfrom Burkholderia cenocepacia in complex with GlcNAc [PDB: 4GNV] and NagZ from Bacillussubtilis in complex with O-(2-acetamido-2-deoxy-D-glucopyranosylidenamino) N -phenylcarbamate(PUGNAc) [PDB: 3NVD]). The overlay reveals that Gln-205 of BglP is likely to clash with the NAcof GlcNAc based substrates, as the amide of the Gln side chain sits ∼ 2 Å from the methyl groupof the NAc from GlcNAc and PUGNAc. In the natural glucosyl intermediate, where a hydroxylgroup is present at C-2, we predict the Gln-205 amide would be 3.0-3.5 Å away from the hydroxylgroup, suggesting that Gln-205 could form a hydrogen bond with the C2 hydroxyl group of glucosidesubstrates. In both cases the interaction at that position is between an amide and a hydroxyl, butthe directionality of the interaction is inverted; a beautiful example of specificity swapping. Indeed,it had previously been speculated that the broadened substrate specificity seen for Nag3, whichalso prefers glucoside to N -acetyl-hexosaminide substrates, was due to the substitutions seen in themodified sequence motif [193]: KH(FI)PGDGX4DQH (underlined residues correspond to Asp-193and Gln-200 in Nag3). The other differing residue, Asp-198 is not close to the active site thus isunlikely to be involved in the broadened substrate specificity of the enzyme.To test whether Gln-205 indeed plays a role in discriminating between glucoside and GlcNAcsubstrates it was replaced by serine in BglP using site-directed mutagenesis to form the mutant,BglP-Q205S. Determination of the kinetic parameters for the cleavage of pNPGlc, pNPGlcNAcand DNPGlc by BglP-Q205S (Figure 3.15A-C) revealed that the Gln to Ser mutation caused thepreferred substrate to switch from pNPGlc to pNPGlcNAc as shown graphically in (Figure 3.15D).Kinetic parameters determined both in the absence and presence of 50 mM phosphate are shown in(Table 3.2). The kcat and kcat/Km values of the Q205S mutant for pNPGlcNAc are both ∼ 10-foldhigher than those of the wild type, with no obvious stimulation from phosphate. However, TLCanalysis confirms that GlcNAc1-P is the predominant product in the presence of phosphate, thusthe rate-limiting step for the mutant with the GlcNAc substrate remains the glycosylation step60Figure 3.15: Phosphate activation of substrate cleavage by BglP.Q205S. Phosphate activation ofBglP.Q205S mediated cleavage of (A) DNPGlc, (B) pNPGlc and (C) pNPGlcNAc. Michaelis-Menten plots of initial hydrolysis/phosphorolysis rates of (A) 1.5 mM DNPGlc, (B) 50 mM pNPGlcand (C) 5 mM pNPGlcNAc in the presence of increasing concentrations of phosphate. Error barsindicate standard error of the mean; where bars are not present, the error is smaller than thesymbols used. Kinetic parameters for cleavage of the sugar phosphates in 0 and 50 mM phosphatecan be found in Table 3.2. (D) Histogram displaying kcat/Km values for hydrolysis of pNPGlc andpNPGlcNAc by BglP-WT and BglP-Q205S.(Figure 3.16). The opposite situation is seen for pNPGlc, for which only kcat/Km values could beobtained since saturation was never reached, even at 50 mM substrate. The kcat/Km of the Q205Smutant is ∼10-fold lower than that of wild type, and again no significant effect of phosphate onrates was observed, pointing to rate-limiting formation of the covalent intermediate. For DNPGlc,a clear stimulation of activity by phosphate could be seen in the mutant, and once again Km (0 mMphosphate) and kcat/Km (50 mM phosphate) are ∼10-fold lower than their counterparts for the wildtype enzyme. The presence of the better leaving group (i.e. DNP) accelerates the glycosylationstep sufficiently that deglycosylation (k3) becomes rate-limiting, thus stimulation by phosphate is61observed.3.3.7 Phosphate RecognitionHoping to gain experimental insights into the structure of the product complex, we tried to obtaina crystal structure of BglP bound to 2-deoxy-2-fluoro-β-glucose 1-phosphate. Soaking BglP crystalswith this analogue (times ranging from 5 min to 1 h) either generated the same 2-fluoroglucosyl-enzyme intermediate species described above, or nothing was found bound in the active site. Wetherefore resorted to manual docking studies to predict how BglP accommodates a phosphorylgroup in its active site. The product Glc-β1-P was modeled into the active site by superimposingthe hexose ring of Glc onto the experimentally determined hexose ring of the covalently bound2-fluoroglucosyl moiety (Figure 3.17). When modelled in the energetically favoured 4C1 chair con-formation, the phosphate of Glc-β1-P sterically clashed with the side chain of Met-292. This was nottoo surprising, however, considering that retaining glucosidases are known to assist bond cleavageby distorting the substrate to position the leaving group in a pseudoaxial position, which brings theconformation of the substrate closer to that of the reaction transition state [229]. Thus, the repulsiveinteractions between Met-292 and the phosphate of Glc-β1-P, in conjunction with the favourablebinding interactions between the substrate glucosyl and phosphate moieties likely force the phos-phate of bound Glc-β1-P product into a pseudoaxial position. This is also likely to be the case fora bound oligosaccharide substrate. Interestingly, when this distortion is accounted for in our modelby distorting the Glc ring toward a 1S3 skew-boat conformation (based on a crystal structure of aTable 3.2: Kinetic parameters for the reaction of BglP.Q205S with pNPGlcNAc, pNPGlc andDNPGlc.Substrate [Pi] Km kcat kcat/KmmM mM s-1 s-1mM-1pNPGlcNAc 0 0.40 ± 0.02 0.03 ± 0.0005 0.07450 0.41 ± 0.02 0.03 ± 0.0004 0.072pNPGlc 0 ND ND 0.00550 ND ND 0.006DNPGlc 0 0.05 ± 0.02 0.03 ± 0.002 0.650 2.5 ± 0.1 1.94 ± 0.03 0.862Figure 3.16: Purified BglP.Q205S TLC activity assay: Phosphate dependent cleavage of DNPGlc,pNPGlc and pNPGlcNAc. BglP.Q205S was incubated with 20 mM DNPGlc, pNPGlc or saturatedpNPGlcNAc in 0 and 50 mM phosphate and incubated for 90 min at room temperature before beingspotted on TLC. Left panel was treated with molybdate stain and right panel with p-anisaldehyde.GH3 NagZ from Bacillus subtilis [203]), which places the phosphate group pseudoaxial, the clashwith Met-292 is relieved with no other steric interferences arising. The pseudoaxial position alsoplaces the oxygen atom of the phosphoester linkage of the product within ∼ 3.2 Å of the imidazoleof His-206 (Figure 3.17), which is predicted to be where the oxygen of the glycosidic bond of thesubstrate would reside when it is protonated during bond cleavage. If the enzyme were to employa glutamic acid residue as acid/base catalyst, as is the case for the majority of GH3 glycosidases,there would likely be significant Coulombic repulsion with the substrate phosphate. This Glu toHis substitution was proposed earlier as a modification that gave members of the subgroup of GH3enzymes the ability to carry out phosphorolysis rather than hydrolysis [85]. Recently, Ducatti etal. [213] demonstrated that while the Glu to His substitution may be necessary for phosphory-lase action, it is not itself sufficient, thus not predictive of phosphorylase activity. They showedthat a GH3 β-N -acetylglucosaminidase from Herbaspirillium seropedicae SmR1 (Hsero1941) bearingthe His-Asp dyad functions predominantly as a hydrolase and observed no activity stimulation orphosphorylated products in the presence of phosphate. They also observed that Hsero1941 showed63Figure 3.17: Modeling Glc1-P (yellow carbons) into the BglP active site (grey carbons). Repulsiveinteractions between Met-292 and the phosphate of Glc1-P, in conjunction with the favourablebinding interactions between the substrate glucosyl and phosphate moieties is predicted to forcethe phosphate of bound Glc1-P product into a pseudo-axial position, where the oxygen atom ofthe phosphoester linkage approaches within ∼ 3.2 Å of the imidazole of His-206, which is predictedto be where the oxygen of the glycosidic bond of the substrate would reside when it is protonatedduring bond cleavage. The experimentally determined 2FGlc molecule covalently bound to Asp-288is shown (green carbons).greater activity toward pNPGlcNAc (kcat = 1.2 s-1) than pNPGlc (kcat = 3.3 x 10-3 s-1), consistentwith the fact that the residue sandwiched between Asp and His in their signature sequence is indeedSer.3.3.8 Acceptor RecognitionBglP was identified based on its ability to transfer the glucosyl moiety from the aryl glycoside,DNPGlc, onto inorganic phosphate thereby generating Glc-β1-P. The use of activated substrates,such as DNPGlc, often allow for the relatively simple determination of the specificity at the -1 (or donor) sub site. But when enzymes are identified through their activity on aryl glycoside64Figure 3.18: Inactivation of BglP for reactivation screen. BglP was incubated with either 0 mM(blue #) or 25 mM (red ) DNP2FGlc and incubated at 25 °C for 210 min. Inactivation wasmeasured by monitoring pNPGlc cleavage at 0, 90, 170 and 210 min following initiation of thereaction. Data fit with single-phase decay equation.substrates it can be more challenging to determine the specificity of the +1 (or activator) sub site.Characterizing the specificity at both the -1 and +1 sub sites is critical in order to understandthe biological function of the enzyme. In the case of BglP, the -1 sub site is specific toward aglucosyl moiety, but the specificity at the +1 site remains a mystery. To investigate BglP acceptorspecificity, a reactivation screen was performed against a panel of 125 potential acceptors [230].BglP was inactivated by trapping it in its glycosyl-enzyme intermediate form through incubationwith 25 mM DNP2FGlc for 210 min at 25 °C (Figure 3.18). Excess DNP2FGlc was removed andthe inactive 2FGlc-BglP was aliquoted into a 96-well plate, each well containing a separate potentialreactivator solubilized in buffer (at 20 mM or maximum solubility). The mixtures were incubatedfor 3 h at 25 °C to allow for potential reactivation. Following this incubation window, reactivationof BglP was measured by adding pNPGlc to each well and monitoring pNP release in a plate reader.Of the 125 compounds assayed, 11 showed reactivation at a rate greater than when no potentialacceptor was present (Figure 3.19). The other 114 compounds that failed to elicit reactivation of2FGlc-BglP are listed in Table 3.3.It was expected that phosphate reactivated BglP, as this had been demonstrated in a previous ex-periment described above (Figure 3.13). With the exception of molybdate, citrate and azide, all theother reactivators contained at least one phosphate group. To ensure the reactivation observed from65Figure 3.19: Reactivators of the 2FGlc-BglP glycosyl-enzyme intermediate. Blue bars representactivation from varying concentrations of phosphate indicated on the y-axis (in mM ). Red barsrepresent reactivation from the corresponding substrates listed on the y-axis. Activities are displayedas a percentage of that seen for the reactivator which ellicited the maximum activity, phosphate (25mM). Activity was measured as change in absorbance at 400 nm per minute. The 114 compoundsthat failed to reactivate 2FGlc-BglP are listed in Table 3.3.66Table 3.3: List of substrates that failed to reactivate BglP.No reactivation of 2FGlc-BglPAcetate GlcNAc-MurNAc-L-Ala-D-Glu 4-Nitrophenyl β-D-glucuronideN -Acetylglucosamine Glucal 4-Nitrophenyl β-D-lactopyranosideN -Acetylglucosamine 6-phosphate D-Glucosamine 4-Nitrophenyl β-D-mannopyranosideN -Acetylmuramic acid D-Glucose 4-Nitrophenyl β-D-xylopyranoside1-Adamantanemethanol β-D-Glucose 1-phosphate 4-Nitrophenyl β-N -acetylglucosamineADP D-Glucose 6-phosphate 1-OctanolD-Allose Glucuronic Acid 1-PentanolAMP GTP Phenethyl alcohol1,5-Anhydro-D-glucitol 1-Hexanol PhenolAnhydro-N -acetylmuramic acid 4-Hexyloxylphenol Phenyl β-D-galactopyranosideL-(+)-Arabinose 5-Hexyne-1-ol Phenyl β-D-glucopyranosideL-(-)-Arabitol Inositol p-PhenylphenolD-(+)-Arabitol α-Lactose o-PhenylphenolArginine D-Lyxose PhloroglucinolAsparagine Maltose 1-PropanolATP Maltotriose 2-Propanol1-Butanol D-Mannitol 1,3-PropenediolCaffeic acid D-(+)-Mannose 1-PyrenemethanolD-(+)-Cellobiose 3-Mercapto-1-propanol QuercetinChitobiose 2-Mercaptoethanol RaffinoseCyclohexanol Methanol ResorcinolCysteine 2-Methoxyethanol α-L-Rhamnose2-Deoxystreptamine p-Methoxyphenol D-Ribose2-Deoxystreptamine di-protected 4-Methylumbelliferyl β-D-cellobiopyranoside Serine2-Deoxystreptamine mono-protected 4-Methylumbelliferyl β-D-galactopyranoside SorbitolL-Erythritol 4-Methylumbelliferyl β-D-glucopyranoside L-(-)-SorboseEthanediol 4-Methylumbelliferyl β-D-xyloside SucroseEthanol Myricetin Sulphate1-Ethynylcyclohexanol 1-Naphthol D-TagatoseFormate 2-Naphthol DL-ThreitolD-Fructose 4-Nitrophenyl α-D-galactopyranoside ThreonineL-(-)-Fucose 4-Nitrophenyl α-D-mannopyranoside α,α-TrehaloseGalactal 4-Nitrophenyl α-D-xylopyranoside L-TyrosineGalactitol 4-Nitrophenyl α-L-arabinopyranoside UTPD-(+)-Galactose 4-Nitrophenyl β-D-cellobioside VanadateGallic acid 4-Nitrophenyl β-D-fucopyranoside 4-VinylphenolGDP 4-Nitrophenyl β-D-galactopyranoside D-(+)-XyloseGentiobiose 4-Nitrophenyl β-D-glucopyranoside67Figure 3.20: BglP reactivation product analysis TLC. Either 2.4 mg/mL (+) or 0 mg/mL (-) BglPwas incubated with 20 mM DNPGlc and 16 mM reactivator (or 25 mM phosphate, 20 mM UDP, 10mM αGlc16-P) for 30 min at room temperature. Product spots are identified with red arrows withred letters corresponding to their proposed structures in Figure 3.21. Glc: 20 mM glucose standard.none: no reactivator. TLC mobile phase: 1-butanol:methanol:NH4OH:H2O (5:4:2:1).these substrates was not simply due to decomposition of the compounds, thereby releasing phos-phate, direct evidence of reactivation products was sought. For product identification inactivatedBglP was not required, therefore DNP2FGlc was substituted out for DNPGlc. Each reactivator(Figure 3.19), together with DNPGlc were incubated with BglP and the products (if any) analyzedby TLC (Figure 3.20).Product spots could be identified by TLC for 7 of the 11 reactivators. The generation of Glc-β1-P(Figure 3.20A) in the presence of phosphate had been established previously, as had the generationof Glc in the absence of phosphate. The three nucleoside monophosphates analyzed by TLC allshowed product spots (Figure 3.20B, C and D), as did UDP (Figure 3.20E), Glc-α1-P (Figure3.20F) and azide (Figure 3.20G). No clear product spots could be observed when either molybdate,CTP or citrate were present, however, molybdate and citrate did appear to more efficiently stimulatethe turnover of DNPGlc to Glc compared to the no reactivator control (Figure 3.20 "none"). Thismay indicate these compounds may have a positive allosteric effect on BglP hydrolysis, which mayhelp explain why they were identified in the reactivator screen (Figure 3.19). Alternatively they68may form a product that then decomposes rapidly. CTP appeared to have an inhibitory effect asno DNPGlc to Glc turnover could be detected during the 30 min incubation time. A potentialexplanation as to why it was identified in the screen is that the labile CTP may have decomposed toCDP and inorganic phosphate prior to addition to the reactivation reaction and the signal observedwas due to the presence of phosphate.Given that BglP operates through a β-retaining mechanism, I proposed that the products were aresult of the transfer of the glucosyl moiety from DNPGlc to the reactivator forming a phosphoesterbond and retaining β-conformation at the anomeric carbon of the glucose (Figure 3.21A-F). Nucle-ophilic anions, such as azide, are known to "rescue" retaining glycosidases in which the acid/baseresidue has been mutated leading to deglycosylation becoming the rate-limiting step [201]. There-fore, it was not a surprise to observe reactivation of BglP from azide. In fact, azide had previouslybeen shown to reactivate Nag3 (Figure 2.6) [85], the first characterized GH3 GP, but product anal-ysis was not done at the time. Retaining β-glucosidases belonging to other CAZy families havebeen used to generate gram quantities of 1-azido-β-D-glucose (Glc-β1-N3) [231, 232], which is ourproposed reactivator product for BglP (Figure 3.21G). To investigate further I took a closer look atthe reactivation products with Glc-α1-P, azide, UMP and UDP.Glc-β1-P-α1-GlcTo determine whether BglP is generating Glc-β1-P-α1-Glc from DNPGlc and Glc-α1-P, directevidence of the product identity was sought through electrospray ionization mass spectrometry(ESI-MS) and 1H NMR. A new reaction was initiated by incubating BglP (4 mg/mL) with 20 mMDNPGlc and 20 mM Glc-α1-P for 10 min at room temperature. BglP was removed by precipitationand the products were analyzed by TLC (Figure 3.22A) and ESI-MS (Figure 3.22B). The sameproduct spot could be observed on TLC as in Figure 3.20F and a mass peak was detected inthe ESI-MS spectrum at 421.0, which corresponds to the m/z of an anion of Glc-β1-P-α1-Glc.Confirmation of the product’s anomeric stereochemistry was achieved through 1H NMR, wheresignals were observed for both α- and β-anomeric hydrogens (Figure 3.23B) as indicated by the J 1,2couplings (phosphorus decoupled) of 3.5 and 7.8 Hz, respectively. Furthermore, by comparing thephosphorus decoupled spectrum (Figure 3.23B) to the phosphorus coupled spectrum (Figure 3.23C)69Figure 3.21: Proposed chemical structures of BglP reactivator products.70Figure 3.22: BglP product analysis: Glc-β1-P-α1-Glc TLC and ESI-MS. 20 mM DNPGlc and 20mM Glc-α1-P were incubated with 4 mg/mL BglP for 10 min at room temperature. The enzyme wasremoved and the products analyzed with (A) TLC, mobile phase: 1-butanol:methanol:NH4OH:H2O(5:4:4:1) and (B) ESI-MS in negative ion polarity mode. Masses of the highlighted peaks correspondto the m/z of an anion of the accompanying structure (MW-H). Analysis performed by Dr. FengLiu.Figure 3.23: BglP product analysis: Glc-β1-P-α1-Glc 1H NMR. (A) Chemical structure of Glc-β1-P-α1-Glc highlighting the anomeric hydrogens (α1-phosphoester linkage: blue, β1-phosphoesterlinkage: red). (B) 1H {31P} NMR spectrum of Glc-β1-P-α1-Glc (decoupled from phosphorus)zoomed on anomeric hydrogen signals. (C) 1H NMR spectrum of Glc-β1-P-α1-Glc (coupled withphosphorus). Analysis performed by, and figure prepared in collaboration with, Dr. Feng Liu.71Figure 3.24: BglP product analysis: Glc-β1-N3 TLC. 20 mM DNPGlc and 16 mM sodium azidewas incubated with either 0 (-) or 4 mg/mL (+) BglP for 10 min at room temperature. Standardsmarked in blue and the Glc-β1-N3 signal is marked with a red arrow. Azide does not produce a signal.TLC mobile phase either (A) 1-butanol:methanol:NH4OH:H2O (5:4:2:1) or (B) chloroform:methanol(5:1).I can confirm that a phosphodiester bridge links the two glucose residues through their respectiveanomeric carbons. In the phosphorus decoupled spectrum (Figure 3.23B) each anomeric hydrogensignal is seen as a doublet, split by their respective C2 hydrogens. When phosphorus is coupled(Figure 3.23C) each anomeric signal is further split by the phosphorus, producing a doublet of adoublet for each of the α- and β-anomeric signals. Taken together, the presence of the expectedmass from ESI-MS, the J 1,2 couplings of the anomeric signals from 1H NMR, and comparison ofthe phosphorus coupled and decoupled 1H NMR spectra, I can confirm that the BglP catalyzedproduct is indeed Glc-β1-P-α1-Glc (Figure 3.21F).Glc-β1-N3A standard of Glc-β1-N3 was obtained from Dr. Hongming Chen and compared against the azidereactivator product by TLC (Figure 3.24). The reaction from Figure 3.20 was replicated with apreparation of BglP that did not contain glycerol, as glycerol has a similar retention factor (Rf)to the reactivator product and thus obscured the TLC signal. With the glycerol absent, it is clearthat the Glc-β1-N3 standard runs at the same Rf as the reactivator product (Figure 3.24A). To72further validate that the standard and product possess the same Rf, the mobile phase was swappedfrom 1-butanol:methanol:NH4OH:H2O (5:4:2:1) to the less polar chloroform:methanol (5:1) eluent.Under these new conditions the two maintained their identical Rfs (Figure 3.24B), indicating thatthe standard and product are one and the same. This finding is further supported by it having beenpreviously established that β-retaining glucosidases, like BglP, are capable of producing Glc-β1-N3[231, 232] (Figure 3.21G).β-UMPGlc and β-UDPGlcScaled up reactions were prepared for UMP and UDP to analyze the products by TLC and ESI-MS. BglP (4 mg/mL) was incubated with 20 mM DNPGlc and either 20 mM UMP or UDP, thenincubated at room temperature for 10 minutes. A standard of α-UDP-Glc was found to have thesame Rf as the BglP catalyzed product when DNPGlc and UDP were starting substrates (Figure3.25). Given that the BglP product has the same Rf as the α-UDPGlc standard and BglP is knownto operate under a β-retaining mechanism, this result supports the case that the product is β-Figure 3.25: BglP product analysis: β-UMPGlc and β-UDPGlc TLC. 20 mM DNPGlc and either20 mM UMP or 20 mM UDP was incubated with 4 mg/mL BglP for 10 min at room temperature.Proposed products are marked in red. α-UDPGlc and Glc standards marked in blue. TLC mobilephase: 1-butanol:methanol:NH4OH:H2O (5:4:2:1).73UDPGlc. For further confirmation the UDP and UMP samples were analyzed by ESI-MS, however,no peaks corresponding the expected products could be identified for either product. Unfortunately,due to time restraints associated with preparing this thesis, I was unable to troubleshoot and solvethe problem to confidently identify the products seen on TLC. It should be noted that the threenucleotide monophosphates (GMP, UMP and CMP) were the best BglP reactivators (other thanphosphate) from the set of 125 analyzed (Figure 3.19) and any future characterization studies aimingto understand BglP’s biologically relevant acceptor/activity should include confirming the identifyof these products.3.4 DiscussionThe screen, developed and validated with three known β-glycoside phosphorylases, proved to berobust, simple to implement, and useful in identifying active clones within a metagenomics library.From a relatively small number of initial hits a single β-glucoside phosphorylase belonging to CAZyfamily GH3 was discovered. This represents the first reported high-throughput functional metage-nomic screen for glycoside phosphorylases, and the approach used may well be amenable to a rangeof other phosphorylases. Given successful implementation of the screening paradigm on known β-glucoside phosphorylases from GH94, and the relatively high number of initial hits, it was somewhatsurprising that GH94 enzymes were not identified in the BCR library. It is unlikely that this is dueto the activity stimulation by phosphate being masked through some other step being rate-limiting,as can be seen for the retaining enzyme studied here, since GH94 phosphorylases are inverting, witha single chemical step. A possible explanation for why no GH94s were discovered is simply thatnone are present within the BCR library. This would be consistent with the partial sequence infor-mation available from this library (from end-sequencing of fosmids), which contained no predictedGH94 ORFs. A second possible explanation is that those GH94s potentially present in the libraryhave limited expression, which in many cases is due to the E. coli host RNA polymerase’s inabilityto recognize foreign promoter sequences on the fosmid DNA [233, 234]. This limitation can becompensated for by equipping the host strain with additional sigma factors that help it recognizea wider range of promoter sequences [235], however the host strain used in this study contained74only the native E. coli sigma factors. A third possibility is that those GH94s that are expressedare not capable of cleaving DNPGlc. To overcome this issue, an alternative functional screen wouldneed to be devised that would utilize natural substrates as opposed to aryl glycosides. This couldinvolve detection of sugar phosphate products when assayed in the degradative direction (such asthe method described in Chapter 4), or oligosaccharides when assayed in the synthetic mode.The reason for the false positives (clones that showed phosphate-dependent DNPGlc cleavage,but failed to produce Glc1-P) is not known at this stage. The most probable is that these areglycosidases whose activity is enhanced by high phosphate levels either through simple salt effectsor through some form of allosteric interaction. Indeed, apart from 29K06 and 31P01, each cloneanalyzed by TLC had been previously identified through functional screens targeting glycosidehydrolases and subsequently sequenced (Supplemental Table B.2). Given that two of the non-GPGH3 ORFs were each independently found on three separate fosmids (see Supplemental Figure B.1),it is likely these are simple β-glucosidases whose activity is in some way stimulated by the presenceof phosphate.Although the screen of 17,168 fosmid clones identified only one new GP, this enzyme did turn outto be interesting, as this represents only the second β-retaining glycoside phosphorylase of any kindto be identified and characterized, and the first for which a three-dimensional structure has beendetermined. BglP seems to be specialized towards β-glucoside substrates, generating a Glc-β1-Pproduct, with much lower activities against N -acetylhexosaminides than has Nag3. Interestingly,the immediately flanking gene on the fosmid that bears BglP is a β-phosphoglucomutase. I clonedand expressed the gene and indeed confirmed that the enzyme converts Glc-β1-P to glucose 6-phosphate, but has no activity on GlcNAc-β1-P. The typical phosphate levels in bacteria of 20-30mM [208] should be sufficient for the two genes together to feed glucosyl moieties directly into theglycolytic pathway without the need for expenditure of any additional ATP. However, the identityof the natural glycan that BglP acts upon which could provide such a metabolic advantage, remainselusive.It has been established that whatever BglP’s natural substrate is, it likely possesses a non-reducing end glucosyl moiety. Reactivation studies of the 2FGlc-BglP glycosyl-enzyme intermediaterevealed a range of 10 potential acceptors that may help identify BglP’s substrate and unravel its75function in a biological context. The inorganic molecule, azide, was shown to reactivate the 2FGlc-BglP intermediate and produce a product with the donor, DNPGlc, that possess the same Rf as theexpected product, Glc-β1-N3. This however, was not considered a potential natural substrate dueto azide being a toxic compound not known to occur naturally in biological systems. Also, azide isa well known reactivator of β-retaining GHs explaining why it was produced here. Three nucleosidemonophosphates (GMP, UMP, CMP) and one nucleoside diphosphate (UDP) were also found toreactivate the intermediate and produce a product when DNPGlc was used a donor. While none ofthe products have yet been confirmed through MS or NMR, the product of DNPGlc and UDP didhave a consistent Rf with α-UDPGlc on TLC. α-UDPGlc is a well known nucleotide sugar donorsubstrate most notably used by glycosyltransferases (GT) for carbohydrate anabolism. However, toour knowledge, no GT is known to act with β-UDPGlc nor is it a known biological compound. Thereare β-anomers of CMP-ketodeoxyoctonate (CMP-β-KDO), a key intermediate in the biosynthesisof bacterial lipopolysaccharides [236], which may provide a hint towards BglP’s natural function.This was not explored further here, but should be followed up on in future studies.Given BglP’s proclivity toward the acceptor phosphate, I consider a likely explanation is thatUDP’s β-phosphate residue is accommodated as an acceptor in the active site and able to performthe deglycosylation step, resulting in the generation of the β-UDPGlc side product. The sameargument could also be made for the acceptor Glc-α1-P, which resulted in the production of Glc-β1-P-α1-Glc, which is too a compound not known to exist in biological systems. Nevertheless,its potential biological relevance was explored. Anomeric phosphodiester-bridged glycans are acommon feature of glycopolymers found in the outer membranes of bacteria, yeasts and protozoa[237], however, there are no reports of any bridged through 1,1 linkages. BglP is closely related tothe NagZ N -acetyl-hexosaminides which are involved in bacterial cell wall maintenance, specificallypeptidoglycan recycling [238]. It is an intriguing possibility to consider that BglP (and Nag3) maybe involved in maintaining a yet to be discovered cell wall feature where Glc-β1-P-α1-Glc can befound. Another intriguing possibility is a potential regulatory role for Glc-β1-P-α1-Glc. It sharesa similar structure with the regulatory molecule P1,P5-di(adenosine-5’)pentaphosphate (Ap5A),which inhibits adenylate kinase (ADK) [239], the enzyme responsible for converting AMP to ADP.Both Glc-β1-P-α1-Glc and Ap5A share a phospho-bridge linking the same type of glycan residues76through either a 1,1 or 5,5 linkage, respectively. ADK acts by transferring a phosphate residuefrom ATP and AMP to produce two molecules of ADP. Ap5A, which is a competitive inhibitorof ADK [240], essentially mimics the binding of either ATP and AMP, or two ADP, by bridgingthe two substrates/products with a phosphate residue. In the same way that Ap5A mimics ADKssubstrates, perhaps Glc-β1-P-α1-Glc is a mimic of trehalose, and therefore could potentially act asan inhibitor of an enzyme involved with trehalose biosynthesis or degradation. In a similar vein, acellulase synthase regulatory molecule, bis-(3’→5’)-cyclic diguanylic acid (also known as cellulasesynthase activator (CSA)), has a symmetrical structure similar to Ap5A and Glc-β1-P-α1-Glc.CSA forms a cyclic molecule consisting of two (or more) riboguanosine residues that possess C2’-C5’ or C3’-C5’ phosphoester-bridges [241] and acts by allosterically activating cellulose synthesis[242]. While the source organism of BglP is not known at this time, Cellulomonas fimi, whereNag3 was found, is thought to metabolize trehalose [243] as it is a known metabolite in anothermember of the Cellulomonas genus [244]. Further studies should focus on testing Glc-β1-P-α1-Glc’spotential inhibitory effect on the range of trehalose acting enzymes present in the C. fimi genome.The identity of the natural substrate of GH3 GPs still remains unconfirmed, however, the researchpresented in Section 3.3.8 should provide a starting point for further investigation which aims tobetter understand the biological role of BglP and Nag3.3.5 ConclusionsPrior to our determination of the structure of BglP bound to a glucoside analog, it was puzzlingthat BglP and Nag3, both of which appeared to be β-N -acetylglucosaminidases on the basis ofa sequence motif similar to that seen in other GH3 N -acetyl-hexosaminidases, are more activeagainst glucoside substrates than against N -acetyl-hexosaminides. Mayer et al. [193] had speculatedthat this difference in substrate specificity (in the case of Nag3) was due to residues Asp-193 andGln-200 since they differed from those in the conserved sequence motif (KH(FI)PGDGX4DQH;altered residues underlined), although at the time the group had no structures of enzymes fromthis sub-group and was unaware that Nag3 was a phosphorylase [193]. Since BglP contains thesame altered sequence motif, our structural study provided insight into the functional significance77of BglP’s homologous residues (Asp-198 and Gln-205). By comparing BglP’s structure to thatof two other NagZ structures having the fully conserved motif (KH(FI)PG(HL)GX4D(ST)H), weconcluded that Gln-205 was likely causing steric hindrance with a C2 N -acetyl group, while atthe same time the NH2 moiety of the Gln side chain was in a good position to H-bond with aC2 hydroxyl group. Indeed, by mutating Gln-205 to a serine (the amino acid present in the fullyconserved GH3 N -acetyl-hexosaminidase motif) the substrate preference was cleanly switched fromglucosides to N -acetyl-hexosaminides. Thus, through the development of a novel screen for glycosidephosphorylases, a new class of enzyme has been discovered and characterized both structurally andmechanistically. This in turn has provided new insights into simple ways in which enzyme specificitycan be encoded and swapped while opening wide a screening paradigm for the discovery of wideranging glycoside phosphorylases with biotechnological potential.78Chapter 4Functional Metagenomic ScreenTargeting Reverse PhosphorolysisActivity4.1 SummaryGlycoside phosphorylases (GPs) catalyze the reversible phosphorolysis of glycosidic bonds, releas-ing sugar 1-phosphates. To identify a greater range of these under-appreciated enzymes, I havedeveloped a high-throughput functional screening method based on molybdenum blue formation.In a proof-of-principle screen focused on cellulose-degrading GPs I interrogated ∼23,000 large in-sert (fosmid) clones sourced from microbial communities inhabiting two separate environmentsand identified seven novel GPs from Carbohydrate Active Enzyme family GH94 and one fromGH149. Characterization identified cellobiose phosphorylases, cellodextrin phosphorylases, lami-naribiose phosphorylases and a β-1,3-glucan phosphorylase. To demonstrate the versatility of thescreening method, varying substrate combinations were used to identify GP activity from familiesGH13, GH65, GH112 and GH130 in addition to GH94 and GH149. These pilot screen and substrateversatility results provide a screening paradigm platform for recovering diverse GPs from unculti-vated microbial communities acting on different substrates with considerable potential to unravelpreviously unknown degradative pathways within microbiomes.794.2 BackgroundGlycoside phosphorylases (GPs) are useful biocatalysts for the synthesis and biotransformation ofcarbohydrates. Currently associated with nine glycoside hydrolase (GH) or glycosyltransferase (GT)families in the Carbohydrate Active Enzyme (CAZy) database [30], GPs carry out phosphorolysisof the glycosidic linkage resulting in the release of a sugar 1-phosphate. Due to the roughly equiv-alent free energy associated with both the glycosidic linkage and the phospho-ester bond of thesugar 1-phosphate the Keq’s of most GPs are near 1, meaning the phosphorylase reaction is readilyreversible. The utility of GPs in biosynthesis results from their inherent capacity to transfer aglycosyl moiety from a sugar 1-phosphate donor to an acceptor glycan, resulting in the productionof a glycosidic linkage. This process, known as reverse phosphorolysis, can be used to generateglycosylated compounds or high-value di- or oligo-saccharides from inexpensive sugar 1-phosphatestarting materials [33, 34].The use of GPs in general would undoubtedly be greater if a more diverse pool of activitieswere available. In response, several research groups have successfully engineered GPs with broadersubstrate specificity or enhanced thermostability [245–248]. To complement these supervised ap-proaches and expand the pool of known GP activities, I have developed an unsupervised functionalscreen targeting the reverse phosphorolysis activity of GPs encoded in large insert (fosmid) clonessourced from environmental genomic DNA. Adapted from an assay designed by De Groeve andcolleagues [249] that has been used for GP directed evolution studies, our screen identifies GPactivity by coupling a chromogenic output, molybdenum blue formation, to the release of inorganicphosphate during glycan synthesis in a 384-well plate format.This chapter describes our initial screening results focused on GPs acting upon cellulose. Iidentified seven new GPs from the CAZy family GH94 and one from GH149 from a starting collectionof ∼23,000 fosmid clones sourced from two environments collectively encoding ∼920,000 predictedopen reading frames. The versatility of the screening method was extended with varying substratecombinations to identify GP activity from other CAZy families including GH13, GH65, GH112 andGH130.80Figure 4.1: Molybdenum blue reaction scheme for the high-throughput discovery of glycoside phos-phorylase activity from metagenomic libraries.4.3 Results4.3.1 Functional Screen ValidationThe molybdenum blue screening method utilizes the reverse phosphorolysis ability of GPs. Thisplate-based assay identifies GP activity by coupling inorganic phosphate released during glycansynthesis to the molybdenum blue reaction (Figure 4.1). Therefore, GPs can be identified byincubating fosmid clones with appropriate donor sugar 1-phosphates and acceptor glycans, thenmonitoring inorganic phosphate accumulation based on the formation of molybdenum blue, whichabsorbs strongly at 655 nm. For this approach to be effective it must be able to identify GPactivity from E. coli cell lysates. Endogenous E. coli enzymes, such as phosphatases, may act bymetabolizing the sugar 1-phosphate substrates, especially the Glc-α1-P used in our initial full-scalescreen. Therefore, it was necessary to include a phosphatase inhibitor to ensure that Glc-α1-P is notconsumed through alternative pathways. It was also important to use an inhibitor that did not alsoconstrain GP activity. Conveniently, molybdate is such an inhibitor [249, 250], and therefore wasadded to the lysate at the same time as Glc-α1-P. Molybdate can bind as a transition state analog81Figure 4.2: Background phosphatase inhibition by molybdate. (A) Inhibition was monitored bymolybdenum blue formation from E. coli cell lysate in the presence of Glc-α1-P and varying con-centrations of molybdate. Reactions performed in triplicate. Error bars not present because thestandard error was smaller than the symbols used. (B) TLC product analysis of background E. coliphosphatases acting on Glc-α1-P in varying molybdate concentrations.to the active site of sugar-1-phosphatases [251, 252], thus acting as a tight competitive inhibitor[253].GPs operate via catalytic mechanisms involving a sugar-oxocarbenium ion-like transition state,thus they do not pass through a transition state similar to that of sugar-1-phosphatases, and there-fore are not inhibited by molybdate. To demonstrate the inhibitory effect of molybdate on endoge-nous phosphatases, E. coli cell lysates were incubated with Glc-α1-P and increasing concentrationsof molybdate. After 24 h incubation, and following addition of the colour development solution(HCl and ascorbic acid), molybdenum blue concentration was highest in the lysates incubatedwith the lowest concentrations of molybdate (Figure 4.2A). Additionally, Glc-α1-P conversion toglucose and Glc6-P was shown to be inversely correlated with molybdate concentration (Figure4.2B). Molybdate appeared to not only inhibit the conversion of Glc-α1-P to Glc and phosphate,but it also appeared to inhibit conversion of Glc-α1-P to Glc6-P, presumably by endogenous α-phosphoglucomutases (αPGMs), which are also phosphotransferases, thus pass through a similartransition state to phosphatases [254]. Therefore, molybdate serves two purposes in this screen: (1)inhibit endogenous phosphatases and αPGM (but not GPs) and (2) complex with free phosphate82to facilitate the molybdenum blue reaction required to identify GP positive clones. In the full-scalemetagenomic screen, 200 mM molybdate was included along with the substrates before the celllysates are added to prevent non-GP Glc-α1-P conversion.The types of GP that can be identified are dependent on the combinations of donor and acceptorsubstrates used. By mixing and matching different substrates one can narrow or broaden the scopeof activities detected in a single screen. However, the initial full-scale screen focused on identifyingphosphorylases capable of utilizing Glc-α1-P as a donor and glucose and/or cellobiose as acceptors.These substrates were selected to target GH94 GPs, the CAZY family containing cellobiose andcellodextrin phosphorylases, in metagenomic libraries sourced from cellulose-rich environments en-riched in cellulose-degrading microbial communities. To establish whether the molybdenum bluemethod is able to identify the targeted GP activity from cell lysate in a high-throughput manner,two GH94s, cellobiose and cellodextrin phosphorylase from H. thermocellum (HtCBP and HtCDP,respectively) were used as positive controls. Lysate from E. coli (BL21) cells expressing HtCBPand HtCDP along with lysate from BL21 carrying an empty pET28 expression plasmid and lysatefrom E. coli (EPI300), the strain used in the subsequent functional metagenomic screens, wereincubated with Glc-α1-P, glucose and cellobiose for 48 h at 37 °C. Following incubation, molybde-num blue formation was induced, quenched, then absorbance was measured at 655 nm. Increasedmolybdenum blue concentration was detected for cells expressing HtCBP or HtCDP while the E.coli EPI300 lysate control remained at background levels (no lysate) (Figure 4.3). The BL21 lysatecontrol carrying an empty pET28 plasmid did show activity above the background, however, it waswell below the activity of those lysates expressing HtCBP or HtCDP.4.3.2 Functional ScreenThe two large insert (fosmid) metagenomic libraries screened for GP activity were sourced froma fecal sample of the North American beaver [255], Castor canadensis, and from a passive minetailings biochemical reactor system (BCR) fed with lignocellulosic biomass [176, 256]. These librarieswere selected on the basis of previous screening and sequencing results indicating an abundance ofcellulolytic enzyme potential. A total of 22,656 fosmid clones were screened, 4,608 from the beaverfecal library (in 12 x 384-well plates) and 18,048 from the BCR (in 47 x 384-well plates) using the83Figure 4.3: Molybdenum blue high-throughput screen validation. Small scale implementation of themolybdenum blue high-throughput metagenomic screen using known GH94s.molybdenum blue screening method outlined in the workflow shown in Figure 4.4. The positive hitthreshold was set at mean + 4 standard deviations (SD) (A655 = 1.28), which yielded 14 positivehits from the beaver fecal library (0.30 % hit rate) and 47 positive hits from the BCR library (0.26% hit rate) (Figure 4.5A). A secondary validation screen was performed by repeating the primaryscreen on a consolidated master hit plate containing the 61 positive hits from both libraries (Figure4.5B), which narrowed the hit pool to 19 positive clones (14 from BCR and 5 from BVR). Thisreduction of the total hit pool by 69 % is reflective of the low cutoff threshold employed in theprimary screen and the inherent variability of performing the molybdenum blue reaction in volumesless than 100 µL. In order to cast a wide net and maximize the likelihood of identifying weak orslow acting GPs, the cutoff threshold was set intentionally to a low value (mean + 4 SD) relativeto the group of GP-containing fosmids (described later), which had an average signal of mean + 16SD in the primary screen. The secondary validation re-screened clones exhibiting high to moderatesignal in quadruplicate allowing for the identification of potential weak GP activity, while at thesame time eliminating false positives.In the next stage of the screen with much smaller number of samples, instead of monitoringphosphate release, TLC was used to monitor the production of a product glycan as an indicatorof GP activity (Figure 4.5C and D). Cell lysates from the 19 positive clones were incubated withan acceptor and donor substrate combination, either Glc-α1-P and glucose (Figure 4.5C) or Glc-84Figure 4.4: Overview of the molybdenum blue high-throughput metagenomic screen workflow.85Figure 4.5: Molybdenum blue high-throughput metagenomic screen results for the BCR and BVRlibraries. Substrates used were Glc-α1-P, glucose, cellobiose. (A) Plot displaying the A655 of bothlibraries. Red dashed line indicates mean + 4 SD (1.28). 61 clones displayed an A655 value greaterthan the cutoff, indicating accumulation of inorganic phosphate. (B) The 61 clones were re-screenedin quadruplicate to confirm activity. Error bars indicate standard error. Red dotted line indicatesthreshold cut off. Activity was confirmed in 19 clones. (C) and (D) TLC validation of the 19positive clones using separate donor/acceptor combinations. Starting substrates for (C) Glc-α1-Pand glucose and (D) Glc-α1-P and cellobiose. Active clones are marked with an asterisk. pCC1 isbackground control.86α1-P and cellobiose (Figure 4.5D). Four of the five BVR library clones (BVR_06I15, BVR_10H18,BVR_11F19 and BVR_11G05) produced product glycans when provided with Glc-α1-P and cello-biose, while no BVR library clones produced product glycans when provided with Glc-α1-P andglucose. Of the 14 validated clones from the BCR library, seven (BCR_17C21, BCR_32A05,BCR_32B15, BCR_38M24, BCR_42I14, BCR_45N22 and BCR_46A06) produced a product gly-can from Glc-α1-P and glucose, and five (BCR_17C21, BCR_32B15, BCR_42I14, BCR_45N22and BCR_46A06) produced a product glycan from Glc-α1-P and cellobiose. Thus, of the 19 clonesassayed by TLC, 11 appeared to produce product glycans when provided with one or both of thedonor-acceptor combinations.4.3.3 Identification of GP activityFosmid DNA samples from the 11 product glycan-producing clones were sequenced uncoveringa total of 13 ORFs encoding CAZymes including 2 GH3s, 1 GH13, 2 GH16s, 7 GH94s and 1GH149 (Figure 4.6). Within the beaver fecal library three fosmids (BVR_06I15, BVR_11F19 andBVR_11G05) were contiguous with each other, mutually overlapping with 16 266 bp at 100 %sequence identity between all three. Two sequential GH94 ORFs (GH94_C and GH94_D) werepredicted within the overlapping region of the three fosmids and a GH3 ORF was predicted in themutual overlapping region of fosmids BVR_11F19 and BVR_11G05, but not in BVR_06I15. Dueto the absence of the GH3 ORF in BVR_06I15, and the fact that all three fosmids exhibited thesame activity, the GH3 was not considered to be the source of the GP activity. Furthermore, GH3suse a β-retaining mechanism that would exclude the enzyme’s utilization of Glc-α1-P as a substratefor reverse phosphorolysis. BVR_10H18, which appeared to have produced the same product glycanas the above three BVR fosmids, albeit at significantly lower levels (Figure 4.5D: lane 16), containedanother set of two sequential GH94 ORFs (GH94_B and GH94_A) and a GH3 ORF. This GH3ORF was also eliminated as a potential source of the activity for the reason stated above. BLASTPqueries against the RefSeq database provided both taxonomic and functional annotations for theputative GP ORFs (Table 4.1). Of the four GH94s found in the beaver fecal library, all returneda top hit with a percent identity score of 99 %. Based on the functional annotations of the topBLASTP hits, GH94_A and GH94_D were predicted to be N,N’ -diacetylchitobiose phosphorylases87Figure 4.6: Circos representation of the 11 fosmid sequences identified through the molybdenum bluehigh-throughput metagenomic screen. Light blue bars represent each fosmid, labelled in the outerpath (LLL_##Y**: LLL = library, ## = plate number, Y = row, ** = column). Scales arein kilobases. Coloured bars within the fosmids show the location of the CAZy ORFs, color codedby family. Ribbons in the center indicate 100 % sequence identity, red to yellow spectrum ribbonsindicate identical regions shared from BVR fosmids, and green to blue spectrum ribbons indicateidentical regions shared from BCR fosmids.88and GH94_B and GH94_C were predicted to be cellobiose phosphorylases. The identical functionalpredictions of these two pairs of ORFs is reflected in the similar CAZy ORF arrangement betweenBVR_10H18 and overlapping region of BVR_06I15, BVR_11F19 and BVR_11G05 (Figure 4.6)and in the percent identity of the pairs (GH94_A and D: 71 %, GH94_B and C: 59 %) (Table 4.2).In the BCR library, a single GH94 ORF (GH94_E) was found on BCR_32A05 and twomore sequential GH94 ORFs (GH94_F and GH94_G) with a flanking GH13 ORF were foundon BCR_38M24. GH13s, which include the sucrose phosphorylases, employ an α-retaining mecha-nism and could therefore utilize Glc-α1-P as a donor. To evaluate the potential contribution of thisORF to observed GP activity, the BCR_38M24 GH13 (subfamily 18) was cloned and expressed.Subsequent testing of the purified protein indicated no GP activity (data not shown) further im-plicating predicted GH94 ORFs encoded on BCR_32A05 and BCR_38M24. The five remainingBCR fosmids (BCR_17C21, BCR_32B15, BCR_42I14, BCR_45N22 and BCR_46A06) were con-Table 4.1: GP active ORFs identified from the molybdenum blue screen.GP ORF SourceLibrarySourceFosmid(s) Top BLAST hitPercentIdentity/SimilarityRefSeq IDFunctionalCharacterizationDi- orOligo- FunctionGH94_A BVR 10H18N,N’ -diacetylchitobiosephosphorylase [Blautiamassiliensis]99/99 WP_059086721.1 Oligo- cellodextrinphosphorylaseGH94_B BVR 10H18 cellobiose phosphorylase[Blautia sp. BCRC 81119] 99/99 WP_110102968.1 Di-laminaribiosephosphorylaseGH94_C BVR 06I15, 11F19,11G05cellobiose phosphorylase[Clostridiales bacterium 59_14] 99/99 OKZ52901.1 Di-laminaribiosephosphorylaseGH94_D BVR 06I15, 11F19,11G05N,N’ -diacetylchitobiosephosphorylase [Clostridialesbacterium 59_14]99/99 OKZ52955.1 Oligo- cellodextrinphosphorylaseGH94_E BCR 32A05 glycosyl transferase[Treponema succinifaciens] 80/88 WP_013701551.1 Di-cellobiosephosphorylaseGH94_F BCR 38M24 glycosyl transferase [Chloroflexibacterium HGW-Chloroflexi-10] 64/77 PKO13267.1 -*cellodextrinphosphorylaseGH94_G BCR 38M24 glycosyl transferase [Longilineaarvoryzae] 86/92 WP_075074767.1 Di-cellobiosephosphorylaseGH149_H BCR17C21, 32B15,42I14, 45N22,46A06hypothetical proteinLARV_03562[Longilinea arvoryzae]80/87 GAP15770.1 Di- andOligo-β-1,3-oligoglucanphosphorylaseDi-: Disaccharide phosphorylaseOligo-: Oligosaccharide phosphorylaseDi- or Oligo- activity refers to cell lysate activity assay (Figure 4.8)*: function predicted based on 4 day activity assay (Figure 4.10)89Table 4.2: Amino acid percent identity between the seven GH94 ORFs found in the molybdenumblue high-throughput metagenomic screen.GH94_A GH94_B GH94_C GH94_D GH94_E GH94_F GH94_GGH94_A 100 20 19 71 31 32 34GH94_B 100 59 20 23 19 21GH94_C 100 19 21 19 22GH94_D 100 31 33 35GH94_E 100 35 54GH94_F 100 40GH94_G 100tiguous with each other, mutually overlapping with 12 886 bp at 100 % identity between all five.The overlapping sequence contained three CAZy ORFs: 2 GH16s and a GH149. Members of theGH16 family use a β-retaining mechanism and therefore were not considered to be the source ofthe GP activity. The recently established GH149 family [116] undergoes a β-inverting mechanismmaking it the likely source of the activity seen in the five BCR fosmids.BLASTP queries of the ORFs, GH94_E, GH94_F and GH94_G, against the RefSeq databasereturned top hits with percent identities of 80, 64 and 86 %, respectively, all with the ambiguousfunctional annotation of ‘glycosyl transferase’ (Table 4.1). The GH149 ORF (GH149_H) returneda top hit with a sequence identity of 80 % and lacking a functional annotation. According to theCAZy database [30] (CAZy DB) at the time of writing, the first and only other member of theGH149 family that has been characterized is from the eukaryotic algae Euglena gracilis (RefSeq ID:AUO30192.1) and has been shown to act as a β-1,3-glucan phosphorylase [116]. While a bacterialmember of the family has been reported (Pro_7066) [116], the full sequence has not since itssource is a proprietary metagenomic database, which likely explains its absence from the CAZyDB. However, a partial peptide sequence of Pro_7066 is available from aa position 429 to 1024.Comparison of the sequence of GH149_H against the Pro_7066 partial sequence yielded a percentidentity of 49 % with a query coverage of 52 %, while a percent identity of 45 % was given whenGH149_H was aligned against the full E. gracilis β-1,3-glucan phosphorylase sequence (Figure S2).Based on the in silico analysis of sequenced GP active clones I identified 8 unique GPs affiliatedwith GH94 and GH149 ORFs. In order to identify their activities, each ORF was subcloned into apET28 expression vector, expressed with a C-terminal histidine tag and purified with immobilized90Figure 4.7: Amino acid sequence alignment of GH149_H, EgP1 (β-1,3-glucan phosphory-lase from E. gracilis) and Pro_7066_par (partial sequence of a metagenomic bacterial β-1,3-glucan phosphorylase). Black shading indicates highly conserved residues, and gray shad-ing indicates conserved similar residues. Sequence alignment was generated using T-Coffee(http://tcoffee.crg.cat/apps/tcoffee/do:regular), and shading was done in BoxShade version 3.21(http://www.ch.embnet.org/software/BOX_form.html) by K. Hofmann and M. Baron.91metal affinity chromatography to be studied in the absence of cellular lysate.4.3.4 Functional CharacterizationOur first step in characterizing the sub-cloned and expressed GH94 and GH149 ORFs identified inthe GP screen was to determine whether they are di- or oligo-saccharide phosphorylases. This pro-cess was facilitated by the combination of substrates used in screening (donor: Glc-α1-P, acceptors:glucose and cellobiose). Disaccharide phosphorylases are active in the presence of Glc-α1-P andglucose while oligosaccharide phosphorylases are active in the presence of Glc-α1-P and cellobiose.However, when both acceptors are combined as part of the same substrate solution, it is challengingto deconvolute which donor/acceptor pair is responsible for the positive hit. This can be furthercomplicated when multiple GPs are present on a single fosmid. Therefore, molybdenum blue assays(96-well format) were done using lysate from cells expressing individual GH94 and GH149 ORFs,that have been sub-cloned into expression vectors, with two separate acceptor/donor combinations(either glucose and Glc-α1-P or cellobiose and Glc-α1-P). Prior to the screen, to ensure expressionwas occurring, an expression test was performed on the soluble cell fraction from all GPs. Expressioncould be detected in the soluble cell lysate for all GPs with the exception of GH94_F, which ap-peared to be expressed only in the insoluble cell fraction, likely as inclusion bodies (Figure 4.8). Asa consequence, activity could not be detected in the GH94_F cell lysate. Cells expressing GH94_B,GH94_C, GH94_E, GH94_G and GH149_H displayed activity using the acceptor/donor combina-tion of glucose and Glc-α1-P suggesting that these GPs are disaccharide phosphorylases, while thoseexpressing GH94_A, GH94_D and GH149_H were active with the acceptor/donor combination ofcellobiose and Glc-α1-P suggesting that these GPs are oligosaccharide phosphorylases (Figure 4.8Band Table 4.1). GH149_H was active in both acceptor/donor combinations, which is consistentwith activity of the only other characterized member of the GH149 family [116].Having confirmed GP activities for seven out of eight subcloned and expressed GH94 and GH149ORFs I looked to specify which glycosidic linkage or linkages were being acted upon or formed. Thinlayer chromatography was used to determine product linkages by matching reverse phosphorolysisproducts to the starting substrates for which the linkage types of forward phosphorolysis are known(Figure 4.9). Reverse phosphorolysis products were analyzed by incubating purified soluble proteins92(GH94_F was omitted due to poor expression) with acceptor/donor combinations of Glc-α1-P andglucose, cellobiose or laminaribiose and then visualizing on TLC (Figure 4.9A). Next, the GPswere incubated with phosphate and cellobiose, cellotriose, laminaribiose or laminaritriose and theproducts visualized on TLC (Figure 4.9B). The enzymes were then grouped and analyzed based onobserved activity.In the case of GH94_A and GH94_D, which share an amino acid sequence identity of 71 %(Table 4.2), I observed a deviation between observed activity and predicted product descriptionsbased on in silico analysis. As indicated above, both proteins were originally annotated as N,N’ -diacetylchitobiose phosphorylases (disaccharide GPs). These annotations were in conflict with thecell lysate assay which predicts GH94_A and GH94_D are both oligosaccharide phosphorylases(Figure 4.8B). Reverse phosphorolysis activity was tested on both GH94_A and GH94_D withGlcNAc-α1-P and glucose, GlcNAc, cellobiose or N,N’ -diacetylchitobiose and in no cases was activityobserved (data not shown). GH94_A and GH94_D were active, however, when incubated with Glc-Figure 4.8: Hit GP expression and lysate activity assay. (A) Soluble, S, and insoluble, I, cellfractions from E. coli BL21(DE3) expressing GPs identified on active fosmids. pET lane indicatedempty pET28 expression vector. Lysozyme band marked with an asterisk. (B) Lysate activityassay of each GP in cell lysate in the presence of Glc-α1-P (grey: G1P only), Glc-α1-P and glucose(red: G1P + G1) or Glc-α1-P and cellobiose (blue: G1P + G2). pET28 indicated empty pET28expression vector. CBP is cell lysate expressing HtCBP (cellobiose phosphorylase) and CDP is celllysate expressing HtCDP (cellodextrin phosphorylase).93Figure 4.9: Reverse and forward phosphorolysis TLC assays. (A) Reverse phosphorolysis assay of 7GPs identified on active fosmids with 10 mMGlc-α1-P and 25 mM (i) D-glucose, (ii) cellobiose or (iii)laminaribiose. Red asterisks indicate products referred to in main text. (B) Forward phosphorolysisassay of 7 GPs identified on active fosmids with 50 mM phosphate and (i) 25 mM cellobiose, (ii) 10mM cellotriose, (iii) 25 mM laminaribiose or (iv) 10 mM laminaritriose. Standards: Std.1: 25 mMD-glucose, 25 mM cellobiose, 10 mM cellotriose and 10 mM Glc-α1-P. Std.2: 25 mM D-glucose, 25mM laminaribiose, 10 mM laminaritriose and 10 mM Glc-α1-P. GH94_F was not included due topoor enzyme expression.α1-P and cellobiose (Figure 4.9Aii). Both produced a product (marked with an asterisk) with aretention factor (Rf) consistent with the cellotriose (G3) standard as well as a third, more polarproduct, with an Rf slightly above that of the Glc-α1-P (G1P) standard, likely cellotetraose. Asimilar activity profile was observed when GH94_A and GH94_D were incubated with Glc-α1-Pand laminaribiose, albeit with upward shifted Rfs (Figure 4.9Aiii). The Rfs of the products marked94with an asterisk in Figure 4.9Aiii were not consistent with any of the standards and fell in betweenthe Rfs of the G3 and L3 standards. I therefore hypothesized that the asterisk marked productswere mixed linkage Glc-β1,4-Glc-β1,3-Glc tri-glucosides and the second product with the lesser Rf,a Glc-β1,4-Glc-β1,4-Glc-β1,3-Glc tetra-glucoside.Based on the reverse phosphorolysis activity profile of GH94_A and GH94_D in all accept-or/donor combinations they both appear to sequentially transfer glycosyl moieties from Glc-α1-P onto the non-reducing end of β1,3 or β1,4-linked di- or tri-glucosides forming β1,4 glycosidic linkages.These data suggest that GH94_A and GH94_D are cellodextrin phosphorylases, an already-knownactivity in the GH94 family. To confirm that GH94_A and GH94_D do act upon β1,4-linkedglucosides, forward phosphorolysis was performed on cellobiose, cellotriose, laminaribiose and lami-naritriose (Figure 4.9B). Of these four substrates, cellodextrin phosphorylases should be active onlyon the β1,4-linked cellotriose. In fact, this is what is seen for GH94_A and GH94_D, since whenincubated with phosphate, cellotriose is phosphorolyzed into cellobiose and Glc1-P (Figure 4.9Bii).GH94_D appears to be slightly active towards laminaribiose as a faint signal corresponding to glu-cose and what is believed to be Glc-β1,4-Glc-β1,3-Glc tri-glucoside was observed (Figure 4.9Biii).The latter presumably formed following the release of Glc-α1-P from the forward phosphorolysis oflaminaribiose, which in turn produces the mixed-linked tri-glucoside via reverse phosphorolysis. Itherefore concluded that GH94_A and GH94_D are cellodextrin phosphorylases (Table 4.1).GH94_B and GH94_C share an amino acid percent identity of 59 % (Table 4.2) and also sharevery similar activity profiles (Figure 4.9). In contrast to GH94_A and GH94_D, observed activityand predicted product descriptions based on in silico analysis were similar. As indicated above, bothproteins were originally annotated as cellobiose phosphorylases (disaccharide GPs) and cell lysateassays indicated this activity (Figure 4.8B). However, when incubated with Glc-α1-P and glucoseGH94_B and GH94_C produce a single product with an Rf consistent with the laminaribiose (L2)standard (Figure 4.9Ai), not cellobiose as the BLASTP annotation would suggest. For GH94_C,the Glc-α1-P is completely consumed while for GH94_B at least a portion of the donor remains.This is likely explained by the 1 hour reaction time not being sufficient to achieve equilibrium.Nonetheless, based on the reverse phosphorolysis activity profile both GH94_B and GH94_C ap-pear to be laminaribiose phosphorylases. To confirm laminaribiose phosphorylases activity, forward95phosphorolysis assays were performed with the substrates described above (Figure 4.9B). As ex-pected, GH94_B and GH94_C were only active on laminaribiose, cleaving it to glucose and Glc1-P(Figure 4.9Biii). I therefore conclude that GH94_B and C are laminaribiose phosphorylases (Table4.1).GH94_E and GH94_G share an amino acid percent identity of 54 % (Table 4.2) and also sharevery similar activity profiles (Figure 4.9). The activity observed in the cell lysate assay (Figure4.8B) indicated that both were disaccharide phosphorylases, which is in agreement with the activityobserved in the presence of Glc-α1-P and glucose (Figure 4.9Ai) and the lack of activity seen with thedisaccharide acceptors (Figure 4.9Aii and iii). When incubated with Glc-α1-P and glucose GH94_Eand GH94_G produce a single product with an Rf consistent with the cellobiose (G2) standard(Figure 4.9Ai). The observed activity is consistent with that of the cellobiose phosphorylases. Toconfirm cellobiose phosphorylase activity, forward phosphorolysis assays were performed as describedabove (Figure 4.9B). GH94_E and GH94_G were shown to be only active on cellobiose, cleavingit to glucose and Glc1-P (Figure 4.9Bi). I therefore conclude that GH94_E and GH94_G arecellobiose phosphorylases (Table 4.1).Based on the cell lysate assay (Figure 4.8B) GH149_H was shown to possess both di- and oligo-saccharide phosphorylase activity. This result was reflected in the reverse phosphorolysis profile,since GH149_H was active with all acceptor/donor combinations (Figure 4.9A). When incubatedwith Glc-α1-P and glucose, GH149_H catalyzed the production of two products consistent withthe Rfs of laminaribiose (L2) and laminaritriose (L3) along with the concomitant loss of the Glc-α1-P (G1P) signal (Figure 4.9Ai). A similar profile was seen when laminaribiose was present asthe acceptor, with product spots consistent with the Rfs of glucose (G1) and laminaritriose (L3) aswell as additional downshifted product spots presumed to be β1,3-linked oligo-glucosides (Figure4.9Aiii). With cellobiose as the acceptor, a smear of products was seen with downshifted Rfs thatwere not consistent with any of the standards, likely mixed linkage products: Glc-[β1,3-Glc]n-β1,4-Glc, 1 ≤ n ≤ 3 (Figure 4.9Aii). The forward phosphorolysis activity profile was in agreement asGH149_H was only active on the β1,3-linked substrates laminaribiose and laminaritriose (Figure4.9Biii and iv). I therefore conclude that GH149_H is a β-1,3-oligoglucan phosphorylase (Table4.1).96Despite trouble expressing GH94_F, purification was still attempted. However, a pure samplecould not be obtained. An eluted fraction from the immobilized metal affinity chromatographyfraction containing the largest overall concentration was assayed for reverse phosphorolysis activity(Figure 4.10). After a 96 h incubation at room temperature the GH94_F purification fractionexhibited cellodextrin phosphorylase activity. Given the lack of purity of the purified fraction nodefinitive conclusion can be made regarding the function of GH94_F, though it is most likely acellodextrin phosphorylase.Figure 4.10: Purification and reverse phosphorolysis activity assay for GH94_F. (A) SDSPAGE ofpurification fraction used in 96 h activity assay. GH94_F is expressed primarily in the insolublefraction. (B) Reverse phosphorolysis activity assay of GH94_F impure fraction 3. 4 µL of fraction3 or 4 µL storage buffer (Table 7.7) was incubated with (g) 0.5 µL 100 mM Glc-α1-P (G1P) and 0.5µL 250 mM glucose (G1) or (c) 0.5 µL 100 mM Glc-α1-P (G1P) and µL 250 mM cellobiose (G2)then incubated for 96 h at room temperature. G3* is presumably cellotriose and G4* presumablycellotetraose.4.3.5 Taxonomic ClassificationTo determine the taxonomic identity of the organisms encoding the characterized GPs, lowest com-mon ancestor analysis using LCA* was performed on the 11 sequenced fosmids [257] (Table 4.3).The contiguous fosmid clones BVR_06I15, BVR_11F19 and BVR_11G05 were affiliated with the97family Ruminococcaceae, specifically Gemmiger sp., an anaerobic bacterium implicated in carbo-hydrate fermentation and the production of formic- and butyric-acids [258]. The fosmid cloneBVR_10H18 was affiliated with the family Lachnospiraceae, specifically Blautia sp., another anaer-obic bacterium implicated in fermentative metabolism in the gut [259]. Both genera fall within theFirmicute phylum [259, 260], which comprises 58.4 % of the microorganisms represented in the BVRlibrary according to SSU rRNA gene sequencing [255]. The beaver’s hindgut microbial community istypical of the wider mammalian profile, being dominated by Bacteroidetes and Firmicutes [261]. Theorganisms located in the lower GI tract are expected to carry out anaerobic degradation of complexdietary carbohydrates, where GPs may be involved. The contiguous fosmid clones BCR_17C21,BCR_32B15, BCR_42I14, BCR_45N22 and BCR_46A06 and the fosmid clone BCR_38M24 wereaffiliated with the family Anaerolineaceae (Table 4.3). Members of the anaerobic Anaerolineaceaefamily within the phylum Chloroflexi have been associated with soil microbial communities involvedin regulating cadmium solubility [262] and therefore it is not unexpected they are found in a passivewastewater treatment system such as that from which the BCR library was sourced [256]. TheTable 4.3: Lowest common ancestor analysis of 11 GP active fosmids.Library Fosmid Lowest CommonAncestorTaxonomicRank GP ORFBVR06I15 Gemmiger Genus GH94_CGH94_D10H18 Blautia Genus GH94_AGH94_B11F19 Gemmiger Genus GH94_CGH94_D11G05 Gemmiger Genus GH94_CGH94_DBCR17C21 Anaerolineaceae Family GH149_H32A05 Treponema Genus GH94_E32B15 Anaerolineaceae Family GH149_H38M24 Anaerolineaceae Family GH94_FGH94_G42I14 Anaerolineaceae Family GH149_H45N22 Anaerolineaceae Family GH149_H46A06 Anaerolineaceae Family GH149_H98fosmid clone BCR_32A05 was affiliated with the family Spirochaetaceae, specifically Treponemasp., associated with several human pathogens including T. pallidum (syphilis) and T. carateum (theskin disease, pinta) [263] as well as a recently discovered species (T. ruminis) found in the bovinerumen that exhibited β-glucosidase activity [264]. More broadly, members of the Spirochaetaceaewithin the Spirochaetae phylum have long been observed in cellulose-rich anaerobic environments[265], such as sedimentary soil, and therefore it was not unexpected to see Spirochaetes representedin the BCR library.4.3.6 Targeting Broader GP ActivitiesThe success of this screening paradigm in identifying new GPs from large insert environmentalgenomic libraries lays the foundation for further screens targeting a broader range of GPs. However,to be successful, the screen must be compatible with a wider range of substrates. De Groeve andcolleagues tested a range of acceptors, such as sucrose, fructose, methyl α-glucoside, methyl β-glucoside, ethyl β-glucoside, octyl β-glucoside and phenyl β-glucoside, all of which proved to becompatible with the screening method [249]. To test the screen’s capacity to accommodate a morediverse set of donors beyond Glc-α1-P, substrate sets were selected to target GP activity from otherCAZy families with known GP activity: GH13, GH65, GH94, GH112 and GH130. Cell lysatescontaining known GPs from each of these families, expressed with the pET expression system inE. coli BL21(DE3) were assayed in 96-well plate format (Figure 4.11). For the GH13 (RefSeq ID:WP_042384192.1) and GH94 (HtCBP [91]) the same acceptor/donor combination was used as inthe large scale metagenomic screen. Different acceptor/donor combinations were used for the GH65(MalP [266]): Glc-β1-P, glucose and maltose; the GH112 (RefSeq ID: WP_013279638.1): Gal-α1-P, glucose and galactose; and the GH130 (RefSeq ID: WP_011693715.1): Man-α1-P, glucose andmannose. The GH13, GH112 and GH130 clones were synthesized by the Joint Genome Institute.Activity could be detected for all GPs in their respective acceptor/donor combinations while thelysate controls (E. coli cell lysate with no GP) remained at background levels (Figure 4.11). Theseresults establish the capacity of the GP screening method to accommodate a wider range of donorsfrom other CAZy families including GH13, GH65, GH112 and GH130. By mixing and matchingadditional acceptor/donor combinations one can narrow the target to a specific GP activity, or cast99a net as wide as possible to identify a range of GP activities encoded within an environmentalgenomic library.4.4 DiscussionThe protocol upon which our metagenomic screen was based was originally used to assay a saturationmutagenesis library at a single amino position, thus all the variants tested (94 clones) could fit onone 96-well plate [249]. For each 96-well variant library plate screened, a total of three plates wererequired: the first to replicate and grow the variant strains, the second to perform the reversephosphorolysis reaction and the third in which to carry out the molybdenum blue reaction. At thisscale, liquid handling and liquid transfer could be done manually and resource demand was limited.To scale up to accommodate tens of thousands of clones, significant adjustments were requiredto improve liquid handling and liquid transfer efficiencies and to minimize resource demand. Forexample, 96-well plates were substituted with 384-well plates and the number of plates needed perFigure 4.11: Expanded substrate compatibility assay. E. coli cell lysate expressing a GP from GH13,GH65, GH94, GH112 or GH130 was incubated with the indicated acceptor/donor combinationfollowing the 96-well molybdenum blue activity assay protocol. Control assays (-) were run withcell lysate with no GP and the indicated acceptor/donor combination.100screen was reduced from three to two. Automation was used to ensure accuracy and reproducibility,which included the unconventional use of a colony picking robot to perform the liquid transferbetween the cell lysate and the acceptor/donor plates. While De Groeve and colleagues designedtheir screen as a platform to create new GP enzymes through directed evolution [249], our screeningparadigm opens the door to investigate the wide diversity of GPs encoded in uncultivated microbialcommunities inhabiting natural and engineered ecosystems.Functional metagenomic screening provides an unsupervised method to explore the diversityof metabolic activities encoded within uncultivated microbial communities to identify new andinteresting biocatalysts. One key principle toward design of an effective metagenomic screen isknowing where to look [267]. In our proof-of-principle screen, where I targeted GH94 GPs that actupon cellulosic material, I selected two environments in which lignocellulose was a primary carbonsource. Based on previous work done on both the BCR and BVR library, it was already establishedthat they were enriched with carbohydrate processing CAZymes [176, 255] and the BCR library wasthe source of discovery of a new GP in the GH3 family [91]. A second key principle in the design of aneffective metagenomic screen is the use of a substrate (or substrates) that produce(s) a signal outputthat can be clearly differentiated from background noise. The targeted approach of identifying GPactivity by measuring the release of inorganic phosphate upon GP-mediated reverse phosphorolysisoffers three advantages over the alternative forward phosphorolysis, aryl glycoside GP metagenomicscreen [91]. First, natural substrates (sugar 1-phosphate donors and acceptor glycans) are usedas opposed to substrate analogues, such as aryl glycosides, where the aglycone substituent may beincompatible with GP active sites. Second, targeting reverse phosphorolysis avoids the complicationof deconvoluting hit pools to remove glycoside hydrolases (GHs) that are able to hydrolyze the samearyl glycoside, producing a non-productive positive signal. GHs do not utilize sugar 1-phosphates asdonors nor significantly perform the reverse reaction and therefore do not exhibit a positive signal inthe molybdenum blue screen. Finally, coupling the release of inorganic phosphate to the formationof molybdenum blue yields a better signal-to-noise ratio when compared to measuring the releaseof 2,4-dinitrophenol in the alternative screen. These three aspects may have played a role in thefailure of the previously described forward phosphorolysis screening method (Chapter 3) to identifytwo GH94 GPs (GH94_E and GH94_G) in the BCR library. GH94_E and GH94_G are both101cellobiose phosphorylases capable of phosphorolyzing the aryl glycoside substrate, 2,4-dinitrophenylβ-D-glucopyranoside (Figure 4.12) and both were identified only in the molybdenum blue screeningmethod, despite both methods being designed to target GH94s in the BCR library.Figure 4.12: Phosphate dependent aryl-glycoside assay. 0.5 mg/mL enzyme (or buffer) was incubatedwith 1 mM 2,4-dinitrophenyl β-D-glucopyranoside and 0 or 50 mM potassium phosphate (pH 7.0)and incubated for 1 h at 37 °C. Error bars indicate standard error. n = 3.The LCA* analysis of the GP-containing fosmids predicted that they originate from anaerobicmicroorganisms, which one might expect given their source environments. However, it is worthnoting that the metabolic efficiency advantage of GPs is more impactful in anaerobes than in mi-croorganisms capable of aerobic respiration. GP-mediated metabolism of glycans is inherently moreenergetically favorable than hydrolysis, because the released sugar 1-phosphates can feed into theglycolysis pathway without the expenditure of ATP [48, 49]. Organisms capable of aerobic respi-ration net 36 molecules of ATP per glucose molecule. If the glucose molecule enters the glycolysispathway already phosphorylated the organism nets 37, an increase of 2.7 %. In contrast, anaerobicorganisms only net 2 ATP per glucose molecule, therefore netting a third ATP would increase theATP produced per glucose molecule by 33 %. The potential of the metabolic efficiency advantageof GPs being more impactful in anaerobes may guide further GP screens towards libraries sourcedfrom anaerobic environments.Two unique sets of sequential GH94 ORFs were identified from four fosmids in the BVR library.Each set consisted of one laminaribiose phosphorylase and one cellodextrin phosphorylase, the latter102being able to accommodate mixed linked (1,3), (1,4)-β-glucans (MLGs) as acceptors. MLGs are adefining characteristic of plants in the order Poales (including many grasses and sedges) and Es-quisetum arvense (common horsetail) within the order Equisetales, where they exist as part of thehemicellulose meshwork of the cell wall [268]. While it is commonly known that the beaver’s dietconsists of tree bark, they are also known to forage on other non-woody herbaceous plants [269, 270],which may include grasses and horsetails in the vicinity of beaver dams. The two phosphorylases,originating from the Firmicutes, a phylum commonly represented within the mammalian micro-biome, may work in conjunction to metabolize MLGs present within the beaver’s diet. In additionto the set of GH94s, a putative phosphoglucomutase (PGM) (top BLAST hit: WP_007048272.1,86 % identity) was also discovered in the overlapping regions of BVR_06I15, BVR_11F19 andBVR_11G05 fosmids (but absent from the BVR_10H18 fosmid). In order for the MLG phospho-rolysis product, Glc-α1-P, to enter the glycolysis pathway it first must be converted to Glc6-P. APGM would catalyze the conversion of Glc-α1-P to Glc6-P without consuming an ATP molecule.As described in the previous paragraph, this MLG metabolism scheme (outlined in Figure 4.13)has the potential to increase fitness of the expressing cells through increased ATP yield duringglycolysis.4.5 ConclusionsThis chapter detailed the development and deployment of a high-throughput screening paradigm forthe recovery of GPs from large insert environmental genomic libraries. The preliminary screeningresults identified seven new GPs from CAZy family GH94 and one from GH149. Of the seven GH94s,six could be assigned functions: two cellobiose phosphorylases, two cellodextrin phosphorylases, twolaminaribiose phosphorylases with one undetermined. The single GH149 was determined to be a β-1,3-oligoglucan phosphorylase and represented the first full-length characterized bacterial GH149 toenter the public domain. These results validate gene model predictions from environmental sequenceinformation and provide more precise product descriptions based on substrate linkages. Moreover,a small-scale validation assay confirmed compatibility of the screen with additional donors beyondGlc-α1-P. This paradigm provides a platform for future screening efforts to discover a wider range103Figure 4.13: Hypothetical mixed linked glucan (MLG) metabolism scheme. GH94_D, GH94_Cand phosphoglucomutase (PGM) (black arrows) from BVR_06I15, BVR_11F19 and BVR_11G05fosmids. Orange arrow indicates the ATP consuming phosphorylation of free glucose step. Whilethe terminal reducing end glucose will always require phosphorylation, this step is avoided for allthe non-reducing end residues when GP mediated glucan metabolism is employed.of GPs from diverse environmental genomic material with potential application in the biocatalyticconversion of high-value carbohydrates.104Chapter 5Characterization of a PhylogenomicallyDiverse GP Library5.1 Summary and BackgroundDue in large part to the advent of next generation, massively parallel DNA sequencing technolo-gies, the past two decades has seen a disorderly explosion of genomic data. To keep pace with theever accumulating data generated from genomic and metagenomic sequencing studies, improvedbiochemical characterization techniques need to be developed and deployed on a large scale in orderto be able to reliable extract biologically useful information from these ballooning data sets. In themiddle of the previous decade, the Joint Genome Institute (JGI), a national laboratory operatingunder the purview of the US Department of Energy attempted to tackle this problem by developing ahigh-throughput functional characterization pipeline that involved sampling sequence databases fora phylogenetically diverse set of GH1 CAZymes that would subsequently be functionally characterized as a test case [180]. The JGI applied high-throughput methods, such as nanostructure-initiatormass spectrometry (NIMS) coupled to acoustic printing [271, 272], which allowed for parallel screen-ing of substrate specificity over a range of temperature and pH conditions. NIMS analysis was usedhere as a filtering mechanism to prioritize candidates that would go on to be further characterizedby HPLC with an expanded set of substrates. While this approach was applied to a single CAZyfamily, the analysis techniques are broadly applicable to other families as well as non-CAZymes.More recently, in 2020, Li et al. established and validated a method involving sequence similaritynetworks (SSNs) as well as other biological data to predict function of uncharacterized members ofthe GH130 family [90]. While this method is not a substitute for biochemical characterization, it105does help to target which candidates to characterize first if, for example, novel activities are beingsought after. As with the JGI’s approach, this method is also broadly applicable to other enzymefamilies.I have collaborated with the JGI to characterize a phylogenomically diverse GP library by adapt-ing the molybdenum blue screening method, presented in Chapter 4, as a broad-range substratespecificity assay. I also utilized SSNs with functionally characterized family members mapped ontoit to help predict the type of glycosidic linkage GPs may act on. I then narrowed the GP librarydown to a handful of candidates that may represent novel activities and one of those was thenselected for full characterization. This effort lead to the discovery of a new GP activity in the GH94family, the identification of a new biopolymer as well as added a new high throughput substratescreening method to the JGI’s pipeline.5.2 Results and Discussion5.2.1 Phylogenomic SamplingCandidate sequences were selected from GP-containing GH families 65, 94, 112, 130 and subfamily18 of GH13. The Joint Genome Institute (JGI) generated phylogenetic trees for each family thenselected a set of representatives that would encompass maximal sequence diversity in each of thefamilies (Supplemental Figures C.1, C.2, C.3, C.4 and C.5). To achieve this, they used an algorithmthat maximized the phylogenetic distance captured within a given set to guide sampling [273]. Wethen filtered the 125 sequence set to eliminate leader peptides, large multi-domain sequences andsequences that were predicted to contain transmembrane domains. The finalized sequences werethen codon-optimized for E. coli, synthesized, then cloned into a pET45 expression vector andtransformed into E. coli BL21(DE3) (by the JGI) and provided to the Withers lab in a 96-wellplate-based format.5.2.2 GP Library Expression AnalysisIn order to perform characterization studies such as substrate specificity assays and identify novelactivities the GPs must be expressed as soluble, functional enzymes that are active in vitro. In106the published case of the GH1 library [180] soluble expression could be detected for 105 of the 175member library, giving a soluble expression rate of 60 %. To determine the GP library’s solubleexpression rate an expression test was performed on the library under three different inductiontemperatures, 16, 25 and 37 °C. The intent of varying the induction temperature was to slow pro-tein expression, thereby potentially minimizing the formation of non-functional, insoluble inclusionbodies. To determine whether a protein expressed as a soluble or insoluble protein, SDS-PAGEwas performed on the soluble and insoluble cell fractions from each of the 125 clones in the GPlibrary. To prepare the soluble cell fraction, E. coli cells that had been grown in auto-inductionmedia [274] were lysed with lysozyme and the non-denaturing detergent, Triton X-100. The lysatewas separated by centrifugation and the supernatant (soluble cell fraction) was separated from thepellet (insoluble cell fraction). The pellet was solubilized with the harsh denaturing detergent, SDS,which permitted analysis by SDS-PAGE. For each GP, the soluble and insoluble cell lysates wererun side-by-side and if a protein band could be identified at the expected size (determined from thepeptide sequence) in the soluble cell fraction lane that clone was designated a soluble expresser. Ifa protein band could only be observed in the insoluble cell fraction lane that clone was consideredan insoluble expresser and if no band could be observed in either lane the clone was marked as ano expresser. The results of this large scale expression test are summarized in Table 5.1 and theSDS-PAGE images for each induction condition can be found in Supplemental Figures C.6, C.7 andC.8. Unfortunately the soluble expression rate of the GPs in the library was 41 % (37 °C), quite abit lower than the 60 % reported by Heins et at. [180] for the GH1 library. Furthermore, reducingthe induction temperature appeared to have the opposite effect to what was intended as indicatedby the soluble expression rates of 38, and 34 % for 25 and 16 °C, respectively.In addition to the expression test, an activity assay was also performed on each soluble fractionusing the molybdenum blue substrate specificity assay described in Chapter 4. The substrates usedin these activity tests depended on which CAZy family the GP belonged to. For GH13s and GH94s:Glc-α1-P, glucose and cellobiose, GH65s: Glc-β1-P, glucose and maltose, GH112s: Gal-α1-P, glucoseand galactose and GH130s: Man-α1-P, glucose and mannose. The results from the activity test aresummarized in Table 5.1 and presented in more detail in Supplemental Figures C.6, C.7 and C.8.The activity test was included here as a preliminary screen to establish the testing method and107Table 5.1: Summary of GP library expression tests.Well Family GenBank ID 16 °C 25 °C 37 °C Activity? Well Family GenBank ID 16 °C 25 °C 37 °C Activity?1A04 GH13 AFP32696.1 I I I 2C06 GH94 ACZ20603.1 N N N1A06 GH13 ABO68560.1 S S S 2D04 GH94 CDR30871.1 S S S 1A07 GH13 ADG91878.1 I I I 2E04 GH94 ABX81671.1 I S S1B04 GH13 ADY14271.1 I I I 2F07 GH94 BAK80394.1 N N N1B05 GH13 BAJ67724.1 S S S 2G03 GH94 ABX44201.1 I I I1B07 GH13 ABV50542.1 I I I 2G05 GH94 ABZ98496.1 N N N1B08 GH13 AFY78349.1 I N I 2H02 GH94 CBL11429.1 S S S1B12 GH13 BAB37700.1 I I I 2H03 GH94 AIQ37435.1 I S I1C04 GH13 ADL69407.1 I I I 2H04 GH94 AIG44033.1 I I I 1C06 GH13 ABE59935.1 I N I 1A09 GH112 ACZ00636.1 N N N1C07 GH13 AEV29631.1 I I I 1A10 GH112 CBL18493.1 S S S1D04 GH13 AIC48094.1 I S S 1A11 GH112 ABX40964.1 S S I1D05 GH13 BAN03569.1 I I I 1A12 GH112 CBL02866.1 I I I1D06 GH13 CAD77533.1 I I I 1B10 GH112 ADL32981.1 S S S 1D07 GH13 ADT70501.1 I I I 1B11 GH112 ADY21327.1 S S S1E05 GH13 AFE10799.1 I I I 1C09 GH112 ADY12199.1 S S S1E06 GH13 ADP96912.1 I I I 1C10 GH112 AEN95946.1 S S S1E07 GH13 ABW28639.1 I I I 1C11 GH112 ADH97994.1 I I I1F04 GH13 AGX06313.1 S S S 1C12 GH112 ACB74662.1 I I I1F06 GH13 ABO68560.1 I I I 1D11 GH112 AGJ77788.1 I I I1F07 GH13 CDM93278.1 S S S 1E09 GH112 ACV29689.1 S S S1G04 GH13 AFT95897.1 I I S 1E10 GH112 ADO77015.1 I I I1G05 GH13 ACZ08382.1 S S S 1E12 GH112 BAH10635.1 I I I1G06 GH13 CAK24900.1 I I I 1F09 GH112 AEE17762.1 S S S1G07 GH13 AEF86931.1 S S S 1F10 GH112 ABX42289.1 S S S1H02 GH13 ADY84227.1 I I I 1F12 GH112 BAB80279.1 S S S1H03 GH13 BAN99152.1 I I I 1G09 GH112 AHU89776.1 I S S1H04 GH13 BAF38859.1 S S S 1G10 GH112 ADL04976.1 S S S1H05 GH13 ADR37224.1 S S S 1G11 GH112 AGC67054.1 S S S 1H06 GH13 AHD03359.1 I I I 1G12 GH112 CBL09398.1 S S S 1B09 GH65 ADC90669.1 I I I 1H09 GH112 BAL17712.1 S S S1D08 GH65 AIL46320.1 N N N 1H10 GH112 ABG86710.1 I I I1D10 GH65 ADC27646.2 I I I 1H11 GH112 AIL73535.1 S S S 1D12 GH65 ADQ16349.1 S S S 1H12 GH112 ADH91781.1 I I I1G08 GH65 ADG12399.1 I I I 2A01 GH112 ACB74662.1 I I I1H08 GH65 ABL78553.1 I I I 2E01 GH112 ADI66539.1 S S S2A02 GH65 CCC30620.1 I I I 2F01 GH112 CBK70037.1 S S S2A05 GH65 AFW94700.1 N N N 1A03 GH130 ABI69320.1 I I I 2B01 GH65 AHX14552.1 S S S 1A05 GH130 ABW10003.1 I I I2B02 GH65 CAX21997.1 I I I 1A08 GH130 ADP77507.1 I I I2B03 GH65 ADG73177.1 I I I 1B03 GH130 AEE47952.1 I I I2C01 GH65 ABX43667.1 I S S 1C02 GH130 ACX65647.1 I I I2C02 GH65 AAK04526.1 I S S 1C03 GH130 CAD72947.1 I I I2C03 GH65 CAF33038.1 I I N 1C05 GH130 ACU74625.1 I I I2D01 GH65 BAG26361.1 S S S 1D01 GH130 AEE13905.1 I I I2D02 GH65 ADK80024.1 S N N 1D02 GH130 EDO12271.1 I I S2D03 GH65 AHL73842.1 S I I 1D03 GH130 AHF91733.1 I S S 2E02 GH65 AEB72731.1 I I I 1D09 GH130 ACK41943.1 I I I2E03 GH65 ADP39332.1 I I S 1E01 GH130 AII65745.1 I S S 2E05 GH65 AFU70794.1 S S S 1E02 GH130 ABK05267.1 S S S 2F02 GH65 AEJ61827.1 S S S 1E03 GH130 AHY54122.1 I I I2F03 GH65 CCP23819.1 I I I 1E04 GH130 ADY51890.1 I I S2G01 GH65 AHI52784.1 I S I 1E11 GH130 ACY49488.1 S I S 2H01 GH65 ABX41399.1 S I I 1F02 GH130 AIE87935.1 I I I1E08 GH94 AHC15980.1 I I S 1F03 GH130 AHE57032.1 I I S 1F01 GH94 ADC84810.1 I I I 1F05 GH130 ADY51932.1 I I I1F11 GH94 AIE87590.1 S S S 1F08 GH130 AIF25974.1 I I I2A03 GH94 BAK81609.1 I I I 1G02 GH130 AHE52113.1 S S S 2A04 GH94 CAJ26026.1 S S I 1G03 GH130 BAS15972.1 I I N2A07 GH94 CBL43852.1 N N N 1H01 GH130 ACU75655.1 S S S2B04 GH94 AER17792.1 S S S 1H07 GH130 AKM78125.1 I I I2B06 GH94 BAM05052.1 N N N 2D06 GH130 AEE96966.1 N N I2C04 GH94 AII50828.1 S S S Soluble expression rates: 34 % 38 % 41 %S: soluble expresser, I: insoluble expresser, N: no expresser, : activity detected under at least one of the threeconditions tested.Data presented here were qualitatively gathered from the expression test and activity assays presented in Supple-mental Figures C.6, C.7 and C.8.108confirm that at least a subset of GPs with compatible substrate specificities could be detected. Theset of substrates will be expanded further to better characterize GP activity in the following section.5.2.3 GP Library Substrate Specificity AnalysisThe molybdenum blue screening methodology made it possible to screen the 125 clone GP libraryfor reverse phosphorolysis activity with a range of donor and acceptor combinations. The results ofthe GP library substrate specificity analysis are described, by CAZy family, in sections that follow.Each clone in the library was assayed with up to 20 different combinations that differed dependingon which CAZy family was being analyzed. For this assay, GP expression was induced at 37 °C andall clones were analyzed, not just those that were found to be soluble expressers. The primary reasonfor including the insoluble expressers and no expressers was that their designations were based onmy qualitative assessment and I wanted to ensure I did not exclude any potential low expressing, buthighly active clones where expression could not be detected through SDS-PAGE. Additionally, thesubstrates used here were not so prohibitively costly to prevent the screening of the whole library.However, in future screens that may use a more expansive (and expensive) substrate set that requiremore wells per clone it may be useful to consider utilizing the expression data gathered in Table 5.1to narrow the focus solely on the soluble expressers.One limitation of the molybdenum blue reverse phosphorolysis detection method is that it isunable to provide definitive evidence about the glycosidic linkage being generated. In order toidentify the linkage, the product would need to be purified and analyzed by NMR, which is alaborious task and difficult to adapt to a high-throughput process. To help predict which linkageswere being generated by active GPs, I utilized sequence similarity networks (SSN), which map andcluster all members of a family based on their pairwise percent sequence identity. Functionallycharacterized GPs and uncharacterized members from the GP library were then mapped onto theSSN to reveal how each member clustered with the others. If an active, uncharacterized memberis clustered with characterized ones, where only a single linkage is well represented, I can predictthat the yet-uncharacterized member generates the same linkage. However, to confirm the linkage,biochemical characterization would still have to occur. Nevertheless, this methodology can helpus to prioritize which active GPs are most interesting to focus on first and which may represent109Figure 5.1: GH65 acceptor specificity heat map. Molybdenum blue screening assay of the GH65sin the GP library. Phosphate released from reverse phosphorolysis is coupled to the formation ofmolybdenum blue which absorbs strongly at 655 nm (A655). GPs are identified by well ID along thetop. Donor was Glc-β1-P in all cases. "LYSATE" column is a background control of E. coli solublecell lysate expressing no GP. "BUFFER" column is a molybdenum blue reaction control containingno lysate. Expression information from Table 5.1 plotted along the bottom of the heat map.previously undiscovered activities. Those active GPs that do not cluster with any characterizedmembers and/or utilize acceptors that are not known within their respective families are of particularinterest.5.2.4 GH65 Substrate Specificity AssayThe GP library contains 24 GH65s, 12 of which were designated as soluble expressers. A substratespecificity assay was performed on all 24 using the donor, Glc-β1-P, and 20 different acceptors(Figure 5.1). A full list of the acceptors and concentrations used for the GH65 screen, as well as forthe other four families, can be found in Table 5.2. Of the 24, four catalyzed glycosyl transfer to at110Table 5.2: GP library substrate specificity acceptor list.Label Name [Final] (mM)Fru D-Fructose 50Gal D-Galactose 50Gal-β1,3(4)-Gal D-Galactobiose 20Gal-β1,4-Glc Lactose 20GalN D-Galactosamine 50GalNAc N -Acetyl-D-galactosamine 50Glc D-Glucose 50Glc-α1,4-Glc Maltose 50Glc-β1,2-Glc Sophorose 5Glc-β1,3-Glc Laminaribiose 10Glc-β1,4-Glc Cellobiose 20Glc-β1,6-Glc Gentiobiose 20GlcN D-Glucosamine 50GlcN-β1,4-GlcN Chitobiose 20GlcNAc N -Acetyl-D-glucosamine 50GlcNAc-β1,4-GlcNAc N,N’ -Diacetylchitobiose 10Gly Glycerol 10Man D-Mannose 50Man-β1,4-Man Mannobiose 20ManN D-Mannosamine 50ManNAc N -Acetyl-D-mannosamine 25Rha L-Rhamnose 50UDP Uridine diphosphate 10UMP Uridine monophosphate 10UTP Uridine triphosphate 10Xyl D-Xylose 50111least one of the acceptors: 1D12, 2C01, 2D01 and 2D02. One clone (2B02) showed a higher thanbackground (LYSATE column) signal for acceptors tested, however, it showed approximately thesame signal intensity with Glc-β1-P alone. This suggests that a component of the 2B02 lysate islikely capable of hydrolyzing the donor to release phosphate, explaining the higher than backgroundA655 reading. To gain a clearer understanding of the potential product linkages generated fromthese four GH65 they were mapped on to a SSN representing the GH65 sequence space (Figure5.2).The GH65 CAZy family contains eight known GP activities represented by 20 functionallycharacterized members (as of March 2021) according to the CAZy DB. Those characterized memberswere also mapped on to the SSN to reveal potential functions of meta-nodes. 1D12 and 2D01,which appeared to be most active with the acceptor glucose are located in the same meta-nodeas several known maltose phosphorylases, suggesting these two likely carry out the same functionand therefore generate α1,4-linkages. Coincidentally, 2C01, is identical to an already characterizednigerose phosphorylase, therefore it generates an α1,3-linkage with glucose. Interestingly, however,2C01 appears to also have minor side activity with galactose, xylose, maltose and lactose. 2D02clusters in proximity to a 2-O-α-D-glucopyranosylglycerol:phosphate β-glucosyltransferase, whichgenerates an α1,2-linkage with glycerol. This same activity was confirmed by a former post-docin the Withers lab, Ryan Sweeney, who was interested in this enzyme for an unrelated project.Unfortunately, the substrate specificities of these GH65s are the same as many other known GH65s,at least when assayed with the set of substrates used here. Future exploration of this library shouldfocus on those clones designated soluble expressers, but which failed to catalyze glycosyl transfer,especially those that are found in meta-nodes with no known characterized companions.5.2.5 GH94 Substrate Specificity AssayThe GP library contains 18 GH94s, nine of which were designated as soluble expressers. A substratespecificity assay was performed on all 18 using the donor, Glc-α1-P, and 20 different acceptors(Figure 5.3). Of the 18, five catalyzed glycosyl transfer to at least one of the acceptors: 1F11,2H02, 2B04, 2D04 and 2E04. To gain a clearer understanding of the potential product linkagesgenerated from these four GH94s they were mapped on to a SSN representing the GH94 sequence112Figure 5.2: GH65 sequence similarity network. Amino acid SSN of 4085 unique GH65 sequencesfrom the CAZy DB with an E-value threshold of 10-160. Sequences are grouped into representativenodes when they share a percent identity of 95 or greater. Sequence fragments below 600 a.a.were excluded. Nodes are colour coded based on clustering. Functionally characterized GH65s arerepresented as yellow symbols (or fuchsia diamond) with black outlines. Active GH65s from theGP library are represented as black symbols with yellow outlines and are identified by their welland acceptors which elicited activity (donor: Glc-β1-P). Inactive GH65s are represented by whitesymbols with black outlines.113Figure 5.3: GH94 acceptor specificity heat map. Molybdenum blue screening assay of the GH94sin the GP library. Phosphate released from reverse phosphorolysis is coupled to the formation ofmolybdenum blue, which absorbs strongly at 655 nm (A655). GPs are identified by well ID alongthe top. Donor was Glc-β1-P in all cases. "LYSATE" column is a background control of E. colisoluble cell lysate expressing no GP. "BUFFER" column is a molybdenum blue reaction controlcontaining no lysate. Expression information from Table 5.1 plotted along the bottom of the heatmap.space (Figure 5.4).The GH94 CAZy family contains six known GP activities represented by 25 functionally character-ized members (as of March 2021) according to the CAZy DB. Those characterized members werealso mapped on to the SSN to reveal potential functional information within meta-nodes. 2B04,which was only active on disaccharide acceptors, most notably with cellobiose (Glc-β1,4-Glc) andlaminaribiose (Glc-β1,3-Glc), is located in the same meta-node as other known cellobiose phos-phorylases and therefore likely generates β1,4-linkages. None of the four remaining active GH94sare found in any large meta-nodes and each could represent a potential new activity within thisfamily. 2D04 is found in a small meta-node with a characterized laminaribiose phosphorylase. How-114Figure 5.4: GH94 sequence similarity network. Amino acid SSN of 2254 unique GH94 sequencesfrom the CAZy DB with an E-value threshold of 10-200. Sequences are grouped into representativenodes when they share a percent identity of 95 or greater. Sequence fragments below 200 a.a. andmulti-domain sequences greater than 600 a.a. were excluded. Nodes are colour coded based onclustering. Functionally characterized GH94s are represented as yellow symbols with black outlines.Active GH94s from the GP library are represented as black symbols with yellow outlines and areidentified by their well and acceptors which elicited activity (donor: Glc-α1-P). Inactive GH94s arerepresented by white symbols with black outlines.115ever, it does not appear to be particularly active using glucose as the acceptor, which would notbe expected for a laminaribiose phosphorylase. 1F11 is represented as a singleton on the SSN andactive in the presence of two disaccharide acceptors, sophorose (Glc-β1,2-Glc) and laminaribiose(Glc-β1,3-Glc). Given that 1,2-β-oligoglucan phosphorylases are known in the GH94 family, themost likely explanation is that 1F11 will generate β1,3-linkages but its representation as a singletonwarrants a closer look in future studies. 2H02 clusters as a doublet and is most active in the presenceof galactose with no activity in the presence of glucose. This clone is a strong candidate to be a newGH94 activity as there are currently none that use galactose as an acceptor. Further investigation iswarranted for this clone as well. 2E04 was selected for further characterization, which is describedin a later section (begin in 5.2.10), and as it turned out 2E04 did represent a new GH94 activity.5.2.6 GH112 Substrate Specificity AssayThe GP library contains 28 GH112s, 18 of which were designated as soluble expressers. A substratespecificity assay was performed on all 28 using the donor, Gal-α1-P, and 17 different acceptors(Figure 5.5). Of the 28, 18 catalyzed glycosyl transfer to at least one of the acceptors. To gain aclearer understanding of the potential product linkages generated from these 18 GH112s they weremapped on to a SSN representing the GH112 sequence space (Figure 5.6).The GH112 CAZy family contains two known GP activities represented by 14 functionally char-acterized members (as of March 2021) according to the CAZy DB. Those characterized memberswere also mapped on to the SSN to reveal potential functional information within meta-nodes. Forthis family I was particularly interested in identifying potential lactose (Gal-β1,4-Glc) or LacNAc(Gal-β1,4-GlcNAc) phosphorylases, which are activities yet to be discovered. Unfortunately therewere no clear candidates that may represent a new function in the family. Fifteen clones were activewith the acceptors GlcNAc and GalNAc indicating they are all likely lacto-N -biose (or galacto-N -biose) phosphorylases and therefore would generate β1,3-linkages. In fact, five of those 15 clonesshare a greater than 95 percent sequence identity with already characterized lacto-N -biose phos-phorylases, including 1H11, which is located in separate meta-node. Despite this strong evidencesupporting the generation of β1,3-linkages, it would be a relatively simple experiment to screen thisset of 15 GH112s for forward phosphorolysis activity on LacNAc in the presence of phosphate. This116Figure 5.5: GH112 acceptor specificity heat map. Molybdenum blue screening assay of the GH112sin the GP library. Phosphate released from reverse phosphorolysis is coupled to the formation ofmolybdenum blue which absorbs strongly at 655 nm (A655). GPs are identified by well ID along thetop. Donor was Gal-α1-P in all cases. "LYSATE" column is a background control of E. coli solublecell lysate expressing no GP. "BUFFER" column is a molybdenum blue reaction control containingno lysate. Expression information from Table 5.1 plotted along the bottom of the heat map.117Figure 5.6: GH112 sequence similarity network. Amino acid SSN of 268 unique GH112 sequencesfrom the CAZy DB with an E-value threshold of 10-250. Sequences are grouped into representativenodes when they share a percent identity of 95 or greater. Sequence fragments below 600 a.a.were excluded. Nodes are colour coded based on clustering. Functionally characterized GH112sare represented as yellow symbols with black outlines. Active GH112s from the GP library arerepresented as black symbols with yellow outlines and are identified by their well and acceptorswhich elicited activity (donor: Gal-α1-P). Inactive GH112s are represented by white symbols withblack outlines.118type of screen could not be coupled to molybdenum blue formation, but a simple TLC would beable to show if any of the clones were able to produce Gal1-P. Performing this experiment shouldbe a priority for future studies on this group of GPs.1B10 and 1F10 both catalyze glycosyl transfer to glucose and rhamnose and 1F10 is identical toan already characterized GH112 that generates β1,4-linkages with rhamnose, but β1,3-linkages withglucose [74]. The reason why these enzymes form different linkages depending on the acceptor is dueto the biologically relevant enantiomer of rhamnose being L-rhamnose. Structurally, L-rhamnose isquite similar to an upside-down D-glucose, resulting in this class of enzyme also accepting D-glucoseas an acceptor. However, a consequence of D-glucose binding in the +1 subsite in an upside-downorientation is that the positions of the C-3 and C-4 hydroxyl groups are inverted. Therefore, when D-glucose is the acceptor the C-3 hydroxyl is in a position to perform nucleophilic attack on the donor’sC-1, whereas with L-rhamnose it is the C-4 hydroxyl. Neither of these GPs are likely to performlactose phosphorolysis as I hoped. An interesting candidate to test further for lactose phosphorylaseactivity is 1G11, which was active with glucose, but not rhamnose. This clone did cluster separatelyfrom 1B10 and 1F10, but with another characterized Gal-β1,4-Rha phosphorylase (that was presentin the GP library and did not express) making this prospect less likely. Nevertheless, like with theLacNAc experiment described above, testing for lactose phosphorolysis should be prioritized in anyfuture studies.5.2.7 GH130 Substrate Specificity AssayThe GP library contains 25 GH130s, nine of which were designated as soluble expressers. A substratespecificity assay was performed on all 25 using the donor, Man-α1-P, and 17 different acceptors(Figure 5.7). Of the 25, four catalyzed glycosyl transfer to at least one of the acceptors. To gaina clearer understanding of the potential product linkages generated from these four GH130s theywere mapped on to a SSN representing the GH130 sequence space (Figure 5.8).The GH130 CAZy family contains six known GP activities represented by 17 functionally char-acterized members (as of March 2021) according to the CAZy DB. Those characterized memberswere also mapped on to the SSN to reveal potential functional information within meta-nodes.1E01, which is found in a cluster with characterized β1,4-mannooligosaccharide GPs and Man-119Figure 5.7: GH130 acceptor specificity heat map. Molybdenum blue screening assay of the GH130sin the GP library. Phosphate released from reverse phosphorolysis is coupled to the formation ofmolybdenum blue which absorbs strongly at 655 nm (A655). GPs are identified by well ID alongthe top. Donor was Man-α1-P in all cases. "LYSATE" column is a background control of E. colisoluble cell lysate expressing no GP. "BUFFER" column is a molybdenum blue reaction controlcontaining no lysate. Expression information from Table 5.1 plotted along the bottom of the heatmap.120Figure 5.8: GH130 sequence similarity network. Amino acid SSN of 1854 unique GH130 sequencesfrom the CAZy DB with an E-value threshold of 10-90. Sequences are grouped into representativenodes when they share a percent identity of 95 or greater. Sequence fragments below 250 a.a.were excluded. Nodes are colour coded based on clustering. Functionally characterized GH130sare represented as yellow symbols with black outlines. Active GH130s from the GP library arerepresented as black symbols with yellow outlines and are identified by their well and acceptorswhich elicited activity (donor: Man-α1-P). Inactive GH130s are represented by white symbols withblack outlines. β-1,4-Mannosyl-glucuronic acid phosphorylase is not represented because sequenceidentifiers were not available.121β1,4-GlcNAc GPs, is active when glucose, mannose, GlcNAc, mannobiose (Man-β1,4-Man) areused as acceptors and weakly on cellobiose. Analysis was performed on the 1E01 product by aformer Withers lab post doc, Gregor Tegl. On a milligram scale, Gregor used purified 1E01 withthe acceptor GlcNAc to synthesize Man-β1,4-GlcNAc, which was confirmed through NMR analysis.The product was then used as a ligand for co-crystalization analysis of a GH5 glycoside hydrolasefrom Streptomyces cattleya [275]. 1E02 showed some background donor hydrolysis explaining theelevated signal for all acceptors when compared to the "LYSATE" control. Though it appeared toshow activity above the background for mannose, fructose, mannobiose and is found in a meta-nodewith no characterized members. If fructose does act as an acceptor this clone could potentially rep-resent a new GH130 activity, however, E. coli is known to possess a mannose isomerase (EC 5.3.1.7)which performs the interconversion of mannose and fructose. Since these reactions are taking placein E. coli cell lysate there is a possibility that endogenous enzymes may act upon the acceptorsadded. In the case of 1E02 those endogenous enzymes may have converted an incompatible acceptor(fructose) to a compatible one (mannose) which provides a potential explanation to why fructoseelicited a positive signal. In order to test this hypothesis these assays would need to be redone onpurified proteins in the absence of cell lysate. If this explanation applies here then it is unlikely1E02 represents a new activity given that there are already mannooligosaccharide phosphorylasesthat act on β1,2-, β1,3- and β1,4-linkages, although β1,6-linkages still remain a possibility. 1G02demonstrated a weak signal towards mannose and fructose and is found in a meta-node with noother characterized members. The same explanation regarding the mannose isomerase may be ap-plicable here since 1G02 is also active on mannose. If this is the case and mannose is the only trueacceptor out of the set tested this clone may represent a new activity if it is forming β1,3-, β1,4-or β1,6-linked disaccharides. 1D03 showed weak activity on glucose, GlcNAc and mannose and isfound in a meta-node with no other characterized members. Future studies on this group of GH130sshould focus on product identification for 1E02, 1G02 and 1D03 and also expand the substrate baseof the specificity assay to identify the activities for the five soluble expressers that failed to showany signals.122Figure 5.9: GH13 subfamily 18 acceptor specificity heat map. Molybdenum blue screening assay ofthe GH13_18s in the GP library. Phosphate released from reverse phosphorolysis is coupled to theformation of molybdenum blue which absorbs strongly at 655 nm (A655). GPs are identified by wellID along the top. Donor was Glc-α1-P in all cases. "LYSATE" column is a background controlof E. coli soluble cell lysate expressing no GP. "BUFFER" column is a molybdenum blue reactioncontrol containing no lysate. Expression information from Table 5.1 plotted along the bottom ofthe heat map.5.2.8 GH13 subfamily 18 Substrate Specificity AssayThe GP library contains 30 GH13_18s, 10 of which were designated as soluble expressers. Asubstrate specificity assay was performed on all 30 using the donor, Glc-α1-P, and 20 differentacceptors (Figure 5.9). Of the 30, 11 demonstrated activity but donor hydrolysis obscured anyinformation that could have been gleaned from the range of acceptors tested. Nevertheless, all 30GH13_18s were mapped on to a SSN representing the GH13_18 sequence space (Figure 5.10).The GH13_18 subfamily contains four known GP activities represented by 21 functionally char-acterized members (as of March 2021) according to the CAZy DB. Those characterized memberswere also mapped on to the SSN to reveal potential functional information within meta-nodes.123Figure 5.10: GH13 subfamily 18 sequence similarity network. Amino acid SSN of 1653 uniqueGH13_18 sequences from the CAZy DB with an E-value threshold of 10-100. Sequences are groupedinto representative nodes when they share a percent identity of 95 or greater. Sequence fragmentsbelow 350 a.a. were excluded. Nodes are colour coded based on clustering. Functionally charac-terized GH13_18s are represented as yellow symbols with black outlines. Active GH13_18s fromthe GP library are represented as black symbols with yellow outlines but are not individually iden-tified because donor hydrolysis obscured the acceptor specificity result (donor: Glc-α1-P). InactiveGH13_18s are represented by white symbols with black outlines. α-1,4-Glucan:maltose 1-phosphateis not represented here because it is a member of GH13 subfamily 3, not 18.124While the SSN overlaid with the functionally characterized GH13_18 is helpful in predicting po-tential function, without any functional data from the GP library there are limited steps that canbe taken in regards to identifying potential new GP activities within this subfamily. A potentialremedy for the donor hydrolysis issue for future studies would be to reduce the incubation time tominimize the amount of phosphate released from hydrolysis and thus the formation of molybde-num blue. For this approach to be effective though, transfer to an acceptor would need to occurat a rate sufficiently higher than to water to ensure that a positive signal could be distinguishedfrom the background. If the molybdenum blue screening approach ultimately proves incompatiblewith the GH13_18s an alternative approach would be to perform the screening by TLC to identifyproduct signals. This method would greatly reduce the throughput but could be done on a limitedset of candidates, for example just the soluble expressers. Ideally, a reactivator screen (like the oneperformed on BglP in Chapter 3) would be best, but methods to trap reactive intermediates onα-retaining GPs are not readily accessible. Future studies will need to overcome or circumvent thedonor hydrolysis issue associated with the molybdenum blue screening assay. In terms of identifyingnew activities in this subfamily, the two singletons found in the GH13_18 SSN may represent the"low hanging fruit" of where to look first.5.2.9 Summary of the GP Library Substrate Specificity AssayOf the 125 clones in the GP library, 51 were designated as soluble expressers; a soluble expressionrate of 41 %. Of those 51, 31 were found to be active with at least one substrate in the specificityscreen (no GH13_18s in this count). Based on these findings there are several clones that stoodout as potential new GP activities that should be investigated further (2H02, 2D04, 2E04, 1G11,1G02 and 1D03). Beyond those, soluble expressers that did not show any activity with the setof substrates should be subjected to a wider range of acceptors in followup screens, especiallythose which are found in clusters without characterized members. Now that the molybdenum bluesubstrate specificity method has been established future screens should not only see an expanded setof acceptors applied but also alternative donors, particularly in the case of the GH94s where activitiesare known which utilize GlcNAc1-P. Additionally, further screens should involve an expanded setof candidate GPs. The SSNs produced above can inform which new GPs to add to the library to125maximize the chances of discovering new activities, particularly by selecting candidates that clusterwith no other functionally characterized members or as singletons. This substrate specificity methodwas designed to facilitate a broad scope of substrates and clones, especially when paired with roboticplate handlers, liquid handlers and plate readers. To fully take advantage of this scalability, a largeGP library containing soluble expressers paired with an expansive substrate library will need to beemployed. If this can be achieved this approach has the potential to identify a wide range of new GPactivities and help close the gap between the ever expanding wealth of uncharacterized sequencesin the databases and those that have been functionally characterized.5.2.10 Functional Characterization of 2E04The remainder of this chapter will focus on characterizing 2E04, a novel GH94 GP, which I discoveredacts as a β-1,3-N -acetylglucosaminide phosphorylase. The 2E04 gene originated from the genomeof the cell wall-less Mollicute bacterium, Acholeplasma laidlawii and was found to synthesize poly-β-1,3-linked N -acetylglucosamine (GlcNAc) using the donor α-N -acetylglucosamine 1-phosphate(GlcNAc-α1-P). To our knowledge, the resulting poly-β-1,3-N -acetylglucosamine has not previouslybeen described and therefore represents a new biopolymer. Furthermore, it completes the set ofpotential β-linked GlcNAc homo-polysaccharides (though mixed-linkage variants are also possible)together with (1) poly-β-1,4-N -acetylglucosamine, or chitin, the major component of arthropodexoskeletons, fungi cell walls and the second most abundant biopolymer on earth, and (2) poly-β-1,6-N -acetylglucosamine (PNAG), a key virulence factor required for biofilm formation by numerouspathogenic bacteria. Poly-β-1,3-N -acetylglucosamine was denoted, acholetin, a combination ofthe genus Acholeplasma and the well-known β-1,4-GlcNAc polysaccharide, chitin. Therefore, thenew β-1,3-N -acetylglucosaminide phosphorylase is referred to, hereafter, as acholetin phosphorylase(AchP).5.2.11 AchP Sequence Similarity NetworkA second SSN was constructed for the GH94 family that highlights AchP and two other A. laidlawiiGH94s rather than the GP library clones (Figure 5.11). The SSN was constructed using the sameparameters as in Figure 5.6 except the nodes were grouped at 100 percent identity instead of126Figure 5.11: AchP: GH94 sequence similarity network. Amino acid SSN of 1161 unique GH94sequences from the CAZy DB with an alignment score threshold of 10-200. Sequence fragmentsbelow 200 a.a. and multi-domain sequences greater than 600 a.a. were excluded. Nodes colouredbased on meta-node clustering. Functionally characterized GH94s are represented as yellow symbolswith black outlines and A. laidlawii GH94s are represented as black symbols with yellow outlines.Black arrow (meta-node 94-17) indicates the node with the lowest alignment score (1.1 x 10-175)with AchP.127Table 5.3: Known GH94 activities reported in the CAZy DB.EC Name Substrate Phosphorolysis Product2.4.1.20 Cellobiose phosphorylase Glc-β1,4-Glc Glc-α1-P2.4.1.31 Laminaribiose phosphorylase Glc-β1,3-Glc Glc-α1-P2.4.1.49 Cellodextrin phosphorylase Glc-(β1,4-Glc)n Glc-α1-P2.4.1.280 N,N’ -Diacetylchitobiose phosphorylase GlcNAc-β1,4-GlcNAc GlcNAc-α1-P2.4.1.321 Cellobionic acid phosphorylase Glc-β1,4-Gluconic acid Glc-α1-P2.4.1.333 1,2-β-Oligoglucan phosphorylase Glc-(β1,2-Glc)n Glc-α1-P95. Prior to this research, the GH94 family contained six known activities (Table 5.3) that arerepresented within 31 functionally characterized members. The overlaid functional annotationswere based on the list of characterized GH94s in the CAZy DB, characterized metagenomicallyderived GH94s reported in Chapter 4 (Figure 5.11: yellow symbols with black outline) and GH94sequences found in the A. laidlawii genome [276] (Figure 5.6: black symbols with yellow outline).AchP was found to be one of 23 singletons that do not share a pairwise sequence identity of over40 % with another GH94 sequence. The member that shared AchP’s lowest alignment score of 1.1x 10-175, with a pairwise score of 37 %, is marked with a black arrow in Figure 5.6 and resides asa doublet in meta-node 94-17. The member that shared the lowest alignment score (9.5 x 10-27),which also belonged to a meta-node containing functionally characterized members, is located in94-1B. However, the alignment score between these two sequences fell well short of the thresholdand therefore could provide no information toward determining AchP’s activity. Given that SSNanalysis failed to cluster AchP with any characterized GH94, but instead classified it as a singleton,I explored the possibility that AchP may represent a new activity within the GH94 family. To testthis possibility AchP was functionally characterized by analyzing its reverse phosphorolysis activityon a broader range of donors and acceptors tested previously in Section 5.2.5.5.2.12 Substrate Specificity ScreenAchP’s donor and acceptor specificity was determined using a phosphorylase screening method de-scribed previously that couples the chromogenic development of molybdenum blue to the liberationof free phosphate during reverse phosphorolysis. The initial screen of AchP was performed in the sol-uble cell lysate of E. coli, expressing the GP, along with a variety of acceptor and donor combinations128Glc1-PGlcNAc1-PGlc GlcNGlcN-β1,4-GlcNGlcNAcGlcNAc-β1,4-GlcNAcGalManXylFru RhaGlc-β1,6-GlcGlc-α1,4-GlcGlc-β1,4-GlcGlc-β1,3-GlcGlc-β1,2-GlcGal-β1,4-Glc A65501234No acceptorDONORACCEPTORFigure 5.12: E. coli cell lysate AchP donor and acceptor specificity screen. Cell lysate from E.coli expressing AchP was combined with the indicated donor and acceptor combinations. Foracceptor details see Table 5.2. Activity was monitored by coupling phosphate release from reversephosphorolysis to the formation of molybdenum blue, which absorbs strongly at 655 nm. Greyindicates corresponding donor/acceptor combination not tested.(Figure 5.12). Of the 18 combinations assayed, AchP was active with either of the donors, Glc1-Por GlcNAc1-P, in the presence of either acceptor, GlcNAc or N,N’ -diacetylchitobiose (GlcNAc-β1,4-GlcNAc), initially indicating this GP’s proclivity toward N -acetyl containing substrates. To confirmand expand on the previous substrate specificity assay, AchP was heterologously expressed, purifiedand screened, in the absence of cell lysate, against an extended set of donors and acceptors, includ-ing additional N -acetamide sugars (Figure 5.13). When incubated with Glc1-P, AchP was activein the presence of the acceptors GlcNAc, GlcNAc-β1-pNP, GalNAc and ManNAc, while no activitywas detected with the non-acetylated glucosamine and galactosamine. With GlcNAc1-P as donor,AchP was active with all acceptors that contained an N -acetyl moiety, as well as with glucose andglucosamine, but not galactose and galactosamine. Lastly, when GalNAc1-P was used as donor,activity could only be detected when GlcNAc was the acceptor.5.2.13 Donor SpecificityKinetic parameters were determined for the reverse phosphorolysis reaction with the donors, Glc1-Pand GlcNAc1-P, each in the presence of either acceptor, GlcNAc or GalNAc (Figure 5.14). Activitycould not be detected with the donor GalNAc1-P nor the acceptor glucose when using reasonablesubstrate and enzyme concentrations and therefore kinetic parameters were not determined. Basedupon both Km and kcat values it is apparent that GlcNAc1-P is the preferred donor over Glc1-P,129Glc1-PGlcNAc1-PGalNAc1-PNo acceptorGlc GlcNGlcNAcGlcNAc-β1-pNPGalGalNGalNAcGalNAc-α1-pNPGlc-β1,3-GlcGlcNAc-β1,4-GlcNAcManNAcA65500.511.52DONORACCEPTORFigure 5.13: Purified AchP donor and acceptor specificity screen. AchP donor and acceptor speci-ficity screen. Activity was monitored by coupling phosphate release from reverse phosphorolysis tothe formation of molybdenum blue, which absorbs strongly at 655 nm.as indicated by Km being 10-fold lower and kcat being 2-fold greater for GlcNAc when GlcNAc1-Pis the donor. These differences indicate the substituent group at the C-2 position of the donor’spyranose ring plays a role in the donor binding and turnover rate. Likewise, the diminished activityseen for the donor GalNAc1-P suggests that the C-4 position also plays role in donor binding.5.2.14 Acceptor SpecificityOf the 11 acceptors screened, only GlcNAc was able to stimulate activity in the presence of eachdonor assayed, albeit minimal activity with GalNAc1-P as donor (Figure 5.13). In addition toGlcNAc, activity was seen with GlcNAc-β1,4-GlcNAc, GalNAc and the aryl glycoside GlcNAc-β1-pNP with both Glc1-P and GlcNAc1-P as donors, while glucose, glucosamine, GalNAc-α1-pNPand ManNAc served as acceptor only with GlcNAc1-P as donor. In the case of Glc1-P, all activeacceptors contained an N -acetyl moiety in an equatorial configuration at the C-2 position, indicatingits potential importance in acceptor binding. Glucose and glucosamine, neither of which contain anN -acetyl group, and ManNAc, which contains an N -acetyl group in an axial configuration, servedas acceptor with GlcNAc1-P as donor but not Glc1-P. Although these acceptors do not contain C-2equatorial N -acetyl groups, the superior kinetic characteristics of GlcNAc1-P compared to Glc1-P sufficiently boosted overall activity so that phosphate liberation could be detected under theconditions assayed.1300 50 100 150 200 250 3000200400600[GlcNAc] (mM)rate (mmol • min-1 • mg-1)Km = 29.9 ±3.7 mMkcat = 24.2 ±1.6 s-1Ki = 281 ±43 mMkcat/Km = 0.8 ±0.4 mM-1 s-1r2 = 0.9902n = 30 10 20 30 40 5002004006008001000[GlcNAc] (mM)rate (mmol • min-1 • mg-1)Km = 3.5 ±0.4 mMkcat = 50.8 ±3.2 s-1Ki = 27.1 ±3.6 mMkcat/Km = 14.5 ±8.0 mM-1 s-1r2 = 0.9894n = 30 50 100 150 200 250 300 3500100200300[GalNAc] (mM)rate (mmol • min-1 • mg-1)Km = 164 ±14 mMkcat = 12.5 ±0.5 s-1kcat/Km = 0.08 ±0.04 mM-1 s-1r2 = 0.9934n = 30 50 100 150 200 250 3000200400600800100012001400[GalNAc] (mM)rate (mmol • min-1 • mg-1)Km = 71.4 ±4.1 mMkcat = 46.7 ±0.9 s-1kcat/Km = 0.7 ±0.2 mM-1 s-1r2 = 0.9952n = 3A BC DFigure 5.14: Reverse phosphorolysis kinetic parameters for AchP. Reactions were carried out with10 mM GlcNAc1-P or Glc1-P donors and varying concentrations of either GlcNAc or GalNAc.Michaelis-Menten plots are shown below.131GalNAc-α1-pNP was the only acceptor assayed that possessed an equatorial C-2 N -acetyl groupthat did not serve as acceptor with Glc1-P. A potential explanation, at least in part, being thatthe α-configuration of the aglycone substituent at the anomeric carbon resulted in steric hindrancein the active site. By contrast the GlcNAc-β1-pNP, which has a β-oriented aglycone, did serve asacceptor, as also did GalNAc-α1-pNP when the superior donor GlcNAc1-P was used. The axialhydroxyl at C-4 of GalNAc-α1-pNP also plays a role in reducing activity as borne out in the kineticanalysis (Figure 5.14). When the donor is Glc1-P the Km is ∼5-fold lower for GlcNAc compared toGalNAc as the acceptor while the kcat is 2-fold larger. A similar pattern is observed when GlcNAc1-P is the donor with a 20-fold lower Km and roughly equivalent kcat values. Additionally, when usingGlcNAc1-P, activity was detected with glucose and glucosamine as acceptors but not with galactoseand galactosamine (Figure 5.13A), further implicating the negative impact of a C-4 axial hydroxylon activity. Examination of AchP’s substrate specificities (Figure 5.13) and kinetic parameterswith varying donor/acceptor combinations (Figure 5.14) has revealed factors influencing substratebinding. For both, the donors and acceptors equatorial configurations at C-2 and C-4 were foundto be important in order to achieve optimal activity.Of the six activities described to date in the GH94 family only the N,N’ -diacetylchitobiose phos-phorylases (ChbP) are known to use GlcNAc1-P as donor, while the other five types utilize Glc1-P.The ChbPs characterized to date [62, 107] have been shown to act as disaccharide phosphorylases,able to transfer the N -acetylglucosamine moiety from GlcNAc1-P to a GlcNAc acceptor generatinga β1,4 glycosidic linkage and the product N,N’ -diacetylchitobiose. GP catalyzed reactions being re-versible, ChbP is also able to phosphorolyze N,N’ -diacetylchitobiose (GlcNAc-β1,4-GlcNAc), whichis likely its biologically relevant activity. The bacteria from which both characterized ChbPs werediscovered are native to marine environments where chitin, a polysaccharide of β1,4-linked GlcNAcand the primary structural component of the exoskeleton of marine invertebrates, is common. How-ever, ChbP have been shown to be strict disaccharide phosphorylases by their inability to utilizeGlcNAc-β1,4-GlcNAc or GlcNAc(-β1,4-GlcNAc)2 as acceptors in substrate specificity assays [107].Therefore, ChbPs do not act to degrade chitin, only disaccharides thereof. AchP on the other handis able to utilize Glc-NAc-β1,4-GlcNAc and GlcNAc(-β1,4-GlcNAc)3 as acceptors and furthermoreappears to subsequently lengthen the acceptor through iterative addition of N -acetylglucosaminyl132Glc1-PGlcNAcphosphateGlcNAc-β1,4-GlcNAcGlcNAcGlc1-PGlc-GlcNAcGlcNAc-β1,4-GlcNAcglycerolstarting material:- + - + REVERSE FORWARDGlcNAcGlcNAc-β1,4-GlcNAcGlcNAc(-β1,4-GlcNAc)3GlcNAc1-P- + - + - +AchP [3 mg/mL]:GNAc1 GNAc2 GNAc4GlcNAc1-PAcceptor:Donor:standards standardsAchP [0.1 mg/mL]:BAGlcNAcGlcNAc-β1,4-GlcNAcGlcNAc(-β1,4-GlcNAc)3GlcNAc1-PFigure 5.15: AchP product analysis TLC. (A) AchP products with GlcNAc1-P (donor) and GlcNAc(GNAc1), GlcNAc-β1,4-GlcNAc (GNAc2), or GlcNAc(-β1,4-GlcNAc)3 (GNAc4) (acceptors) wereanalyzed by TLC. Products are outlined in black boxes. (B) AchP forward phosphorolysis prod-uct analysis with GlcNAc-β1,4-GlcNAc and phosphate. Reverse phosphorolysis with Glc1-P andGlcNAc was include as a control reaction.residues (Figure 5.15A). It was initially speculated that AchP could be a chitin phosphorylase, how-ever, when forward phosphorolysis was tested on GlcNAc-β1,4-GlcNAc no substrate cleavage couldbe detected (Figure 5.15B) indicating that AchP did not act on β1,4 linkages. Although a surprisingresult at the time, it perhaps should not have been given the previous results showing AchP acceptsGalNAc, which has an inverted stereochemistry at C-4 relative to GlcNAc. For GalNAc, the axialorientation of the hydroxyl at C-4 would presumably place it in a different position, compared toGlcNAc, when situated at the +1 subsite. If AchP were producing β1,4 linkages this C-4 hydroxylwould be necessary to act as the nucleophile, attacking the anomeric carbon of the donor as shownin the reaction mechanism in Figure 5.16. Therefore, given the catalytic importance of the C-4hydroxyl it was reasonable to assume that, if β1,4 linkages were being generated, either GlcNAc orGalNAc would have failed to ellicit activity in the substrate specificity assay. However, activity wasobserved for both acceptors, leading to us to exclude the possibility that AchP was generating β1,4linkages.133Figure 5.16: Reverse phosphorolysis mechanism of a β-inverting CDP contrasting glucose and galac-tose as acceptors.5.2.15 Linkage DeterminationThe two other potential linkages AchP could be creating were β1,3 or β1,6. The possibility ofβ1,2 linkages is excluded due to the presence of the C-2 N -acetamide groups of the acceptors.To determine which linkage was in fact being made I next sought direct evidence by analyzingtwo AchP product glycans by NMR spectroscopy. The first was the GlcNAc-GlcNAc disaccharideproduct formed when GlcNAc1-P and GlcNAc were used as the donor/acceptor combination. AchPcatalyzed synthesis was performed at a 1:10 donor-to-acceptor ratio, then the product isolated by gelfiltration chromatography and analyzed by NMR. A through-bond correlation was detected betweenthe non-reducing end anomeric proton and the C-3 carbon (α and β, which are shifted ∼7 ppmdownfield as compared to unlinked GlcNAc) of the reducing end GlcNAc by Heteronuclear MultipleBond Correlation (HMBC) (Figure 5.17), providing strong evidence for a β1,3 glycosidic linkage.The second product analyzed was Glc-GlcNAc-α1-pNP produced with Glc1-P as the donor and thearyl glycoside, GlcNAc-α1-pNP as the acceptor. The Glc-GlcNAc-α1-pNP product was synthesizedon multi-milligram scale, isolated by HPLC then analyzed by NMR. The C-4 and C-6 chemicalshifts (68.22, 60.37, respectively) remained approximately equivalent compared to free GlcNAc,whereas the C-3 (82.03) signal was found to have shifted downfield, suggesting the 1,3 linkage.1343.33.43.53.63.73.83.94 .04 .14 .24 .34 .44 .54 .64 .74 .84 .95.05.15.2f2 (ppm)52545658606264666870727476788082f1 (ppm)2α2β2’6α6’45α3α3β4’3’5’5β1α1β 1’1’(β)[C-2α,H-3α][C-4,H-6][C-4,H-5α][C-5α,H-4][C-5α,H-6][C-5α,H-1α][C-3α,H-1α][C-3α,H-2α] [C-3α,H-4α][C-3α,H-1’(α)][C-3β,H-1’(β)][C-2’,H-3’][C-4’,H-3’][C-4’,H-5’][C-4’,H-6’][C-5’,H-6’][C-3’,H-2’][C-3’,H-5’][C-5’,H-4’][C-6,H-4][C-6,H-5][C-6’,H-5’]ONHAcHOHOOHOONHAcHOOHOH1’(α)6β13C chemicalshift1H chemicalshiftFigure 5.17: GlcNAc-β1,3-GlcNAc HMBC showing the β1,3-linkage. Red : Signals correspondingto the reducing end GlcNAc unit. Blue: Signals corresponding to the non-reducing end GlcNAcunit. Red & Blue: 1H-13C correlations between two GlcNAc units. Figure and analysis preparedand performed by Dr. Feng Liu.135Figure 5.18: AchP reaction scheme.Therefore, through NMR analysis, it was confirmed that AchP generates β1,3 glycosidic linkageswhen either GlcNAc1-P or Glc1-P were used as donors and was shown to synthesize both products,GlcNAc-β1,3-GlcNAc and Glc-β1,3-GlcNAc-α1-pNP. Throughout the linkage analysis there was noindication of the presence of any other glycoside linkage, demonstrating that AchP regioselectivelyforms β1,3 linkages and thus carries out the reaction shown in Figure 5.18, when using GlcNAcresidues as donor and acceptor.5.2.16 OligomerizationThe majority of GPs can be split into one of two categories: disaccharide phosphorylases whichuse a donor sugar 1-phosphate and only monosaccharide acceptors to produce a disaccharide andphosphate, or oligosaccharide phosphorylases, which use a donor sugar 1-phosphate with di-, tri-or oligosaccharide acceptors to subsequently lengthen the acceptor one glycosyl unit at a time,accompanied by a concomitant release of phosphate. Exceptions are the β-1,3-oligoglucan phosphorylases from the CAZy family GH149, which display both di- and oligosaccharide phosphorylaseactivities [116, 162]. AchP appears to also possess both activities as indicated by its ability toutilize both the monosaccharide, GlcNAc, and disaccharide, N,N’ -diacetylchitobiose, as acceptors.Furthermore, TLC analysis of the AchP reverse phosphorolysis reaction shows elongation of eachacceptor (GlcNAc, GlcNAc-β1,4-GlcNAc and GlcNAc(-β1,4-GlcNAc)3 with the donor, GlcNAc1-P)(Figure 5.15).To study the oligomerization further, the degree of polymerization (DP) of the product glycansin the presence of varying concentrations of different donors and acceptors was determined (Figure5.19). Cellodextrin phosphorylase (CDP), a well-studied oligosaccharide phosphorylase belongingto the GH94 CAZy family, carries out the reversible phosphorolysis of oligocellulose. In a character-136BA1:10 1:1 100:1Glc1-PGlcNAc1-PGalNAc1-Pmax: 13max: -max: 6max: 2max: 2max: 3max: 3max: 6max: 4DONOR:ACCEPTOR (GlcNAc)m/z m/zm/z1:10 1:1 100:1Glc1-PGlcNAc1-PGalNAc1-PDONOR:ACCEPTOR (GalNAc)max: 13max: -max: 5max: 2max: 2max: 3max: 3max: 4max: 4m/z m/zm/zFigure 5.19: AchP degree of polymerization (DP) analysis. DP was characterized using MALDI-MS with donors, Glc1-P, GlcNAc1-P or GalNAc1-P and acceptors, (A) GlcNAc or (B) GalNAc,in either 1:10, 1:1 or 100:1 donor-to-acceptor ratios. m/z peaks correspond to the acholetin+Naadducts [M+Na]. Max: largest acholetin chain detected.137ization study of CDP, product glycan DP was shown to be influenced by the relative concentrationsof donor and acceptor; the product glycans with the greatest DP were observed in reactions with thelargest donor-to-acceptor ratio [277]. To determine whether the same pattern held true for AchP,reverse phosphorylase reactions were carried out in the presence of 10 mM donor and either 100mM, 10 mM or 0.1 mM GlcNAc, giving donor-to-acceptor ratios of 1:10, 1:1 and 100:1, respectively.When GlcNAc1-P was used as donor the maximum DPs detected by MALDI-MS were 3, 6, and 13with respective donor-to-acceptor ratios of 1:10, 1:1 and 100:1. (It should be noted that MALDI-MSmay not ionize all DP’s in the same manner and could bias the results toward a certain DP range).When GalNAc1-P was donor the maximum respective DPs were 3, 4, and 6 (Figure 5.19A). Thesame set of reactions were also done substituting GalNAc for GlcNAc as the acceptor (Figure 5.19B)resulting in the same general pattern of oligomerization. Surprisingly, when Glc1-P was used asdonor, no products with a DP greater than 2 could be detected with donor-to-acceptor ratios of1:10 and 1:1. The lack of any product glycans with a DP larger than 2 indicates that, after theglucosyl moiety is transferred from Glc1-P to GlcNAc to produce Glc-β1,3-GlcNAc, AchP is unableto further oligomerize this product when Glc1-P is donor. This is consistent with the substratespecificity results described above (Figure 5.19A) where no activity could be observed with thedonor, Glc1-P, and acceptors, glucose or laminaribiose (Glc-β1,3-Glc). Also, it further implicatesthe importance of an N -acetyl moiety at the C-2 position of the glycosyl residue that would occupythe +1 subsite. Therefore, much like the GH149 β-1,3-oligoglucan phosphorylases, AchP also acts asa dual di- and oligo-saccharide phosphorylase, but only with the donors GlcNAc1-P or GalNAc1-P.Again mirroring the GH149 GP, when utilizing an alternative donor (Glc1-P in the case of AchP andGal1-P for the GH149 GP [278]) AchP can cap the non-reducing end preventing further lengtheningof the oligosaccharide. Therefore, when using Glc1-P as a donor and a monosaccharide acceptor,AchP was shown to act as a strict disaccharide phosphorylase. Much like with CDP and otheroligosaccharide phosphorylases, AchP polymerization can be controlled by adjusting the relativedonor and acceptor concentrations where a higher donor-to-acceptor ratio will result in productswith greater DP. Additionally, the ability of AchP to become a disaccharide phosphorylase whenthe donor is Glc1-P, provides another level of control to this enzyme’s glycan synthesis capabilities.1385.2.17 Core 3 Mucin-like O-Glycan AnalogSince establishing AchP’s ability to synthesize β1,3 glycosidic linkages the substrate specificityassay (Figure 5.13) was reexamined to evaluate the potential utility of the products generatedfrom the range of donor and acceptor combinations screened. A hit that stood out was with thedonor, GlcNAc1-P and the acceptor, GalNAc-α1-pNP. The resulting GlcNAc-β1,3-GalNAc moietyof the presumed product forms the core 3 mucin-type O-glycan structure [279]. The product itself,GlcNAc-β1,3-GalNAc-α1-pNP, is a useful and expensive compound which has been used for thestudy of O-glycan degradation. Enzymatic synthesis of core 3 mucin-type O-glycan structures forthese purposes is hindered by the instability of the β1,3-GlcNAc transferase responsible for its syn-thesis in vivo [279]. AchP was thus used to generate two core 3 analogs with GlcNAc1-P and either4-methylumbelliferyl N -acetyl-α-D-galactosamine (GalNAc-α1-MU) or GalNAc-α1-pNP as accep-tors to generate GlcNAc-β1,3-GalNAc-α1-MU (Figure 5.20A) and GlcNAc-β1,3-GalNAc-α1-pNP(Figure 5.20B). The presence of each product was confirmed by TLC and mass spectrometry. Toverify that the products contained the GlcNAc-β1,3-GalNAc moiety, they were combined with anendo-α-N -acetylgalactosamindase from Streptococcus pneumoniae (EngSP), which has been previ-ously shown to release the disaccharide moiety from GlcNAc-β1,3-GalNAc-α1-pNP, releasing pNP[280]. After a 30 min incubation with EngSP, release of both pNP and MU aglycones could bedetected in a spectrophotometer (Figure 5.20C and D). Only minimal aglycone release was detectedin controls containing no GlcNAc1-P and only the acceptors. These findings demonstrate that witha suitable donor and acceptor, AchP forms the core 3 mucin-type O-glycan, which can in turn becleaved by EngSP to produce a fluorogenic or colorimetric response.5.2.18 Two-pot Large Scale Acholetin SynthesisTo this point acholetin had only been synthesized on an analytical scale. To demonstrate thescalability of the reaction, acholetin synthesis was coupled to GlcNAc1-P production with the aidof an N -acetylhexosamine-1-kinase from Bifidobacterium longum JCM1217 (NahK) [281] in a two-pot reaction scheme (Figure 5.21). In the first pot, 1 g of GlcNAc was combined with 3 g ATP(molar ratio of 1 GlcNAc to 1.3 ATP) and incubated with NahK for 18 h at 37 °C (Figure 5.21A).139Figure 5.20: Core 3 mucin-like O-glycan analog synthesis with AchP. (A and B) TLC and MS anal-ysis of AchP catalyzed products with GlcNAc1-P (donor) and either GalNAc-α1-MU or Gal-NAc-α1-pNP (acceptor). (C and D) Cleavage of the GlcNAc-β1,3-GalNAc moiety from GlcNAc-β1,3-GalNAc-α1-MU or β1,3-GalNAc-α1-pNP and release of the fluoro-/chromogenic aglycone catalyzedby EngSP.140Figure 5.21: Two-pot large scale acholetin synthesis. (A) Pot one. NahK catalyzed production ofGlcNAc1-P. Sequential barium precipitation was performed to reduce ADP, AMP and inorganicphosphate concentrations following the completion of the reaction. GlcNAc1-P preparation wasprecipitated with ethanol then dried. (B) Pot two. AchP catalyzed production of acholetin. Follow-ing the completion of the reaction the acholetin sample was desalted with G25 resin (Figure 5.22)prior to lyophilization and product analysis.141Sequential barium acetate precipitations were done to remove ADP, AMP, free phosphate as wellas any unreacted ATP. After the final barium precipitation, GlcNAc1-P was crashed out of thereaction solution with ethanol then washed with acetone before being dried under vacuum. Thetotal mass of the dried GlcNAc1-P preparation was 2.7 g, however the maximum theoretical yieldof GlcNAc1-P was 1.7 g. A significant proportion of the preparation’s mass was barium acetate,which precipitated along with Glc-NAc1-P. The barium serves a purpose in the subsequent AchPreaction by precipitating free phosphate as it is released via reverse phosphorolysis, thereby drivingthe reaction toward synthesis. In the second pot, the 2.7 g GlcNAc1-P preparation was dissolvedin 10 mL of reaction buffer containing 0.6 mg/mL AchP and 0.4 mM GlcNAc. The amount ofGlcNAc added was sufficient to achieve a donor/acceptor ratio of at least 1000:1, however, it isprobable that a proportion of the 2.7 g GlcNAc1-P preparation contained GlcNAc that had notbeen phosphorylated by NahK. Therefore, the donor/acceptor ratio was not precisely known andconsequently only minimal amounts of acceptor were added in order to maximize the DP of theacholetin product. The reaction mixture was incubated at room temperature for 48 h, at whichpoint no more GlcNAc1-P could be detected by TLC (Figure 5.21B). After removing the precipitatedbarium phosphate salt and enzyme the reaction mixture was desalted on a G25 column to remove theremaining soluble barium acetate (Figure 5.22) then lyophilized to produce a white, fluffy material.The recovered product mass was 344.3 mg, a yield of 37 % from the GlcNAc starting material massof 1 g.5.2.19 Acholetin analysisTo determine the DP of the acholetin preparation multi-angle light scattering analysis was per-formed on the purified acholetin dissolved in water at a concentration of 50 mg/mL. The averagemolecular weight was determined to be 2,966 ±22 g/mol representing an average DP of 14.6 Glc-NAc residue per acholetin molecule (Supplementary Figure C.9). To confirm the homogeneity ofβ1,3 glycosidic linkages throughout, the acholetin was examined by NMR spectroscopy. HMBCanalysis revealed multiple through-bond correlations corresponding to pairs between H-1 and C-3from the adjacent unit, establishing the “HMBC” marked linkages shown on the acholetin structurein Figure 5.23. Anomerization at the reducing end and the presence of multiple GlcNAc units made1421 2 109876543 11 181920171615141312 21 282930272625242322S S SG25 Fraction:ABacholetin elutionFigure 5.22: G25 acholetin desalting. (A) UV absorbance (blue) and conductance trace (brown).Vertical red lines indicate fraction collections. (B) TLC analysis of acholetin fractions from G25column.143Figure 5.23: Acholetin HMBC analysis. Overlapping correlations were resolved with the help of 1Hand 13C COSY and HSQC experiments (Figure 5.24 and section C.1). Analysis performed by, andfigure prepared in collaboration with, Dr. Feng Liu.144Figure 5.24: 13C NMR of untreated acholetin compared to reducing-end reduced acholetin. Carbonnumbers are colour coded to correspond to the acholetin structure shown at the top. The reducingend carbons are shaded blue to indicate their shift following reduction. Analysis performed by, andfigure prepared in collaboration with, Dr. Feng Liu.assigning chemical shifts a challenge. Therefore, the same acholetin sample was reduced and the13C NMR spectra were compared to clarify the reducing end unit (which was found to be predomi-nantly α) and correctly assign the corresponding chemical shifts (5.24). No indications of any otherglycosidic linkage type were present from the NMR analysis. To further establish the homogene-ity of the β1,3 linkages, acholetin was subjected to potential digestion by an endochitinase fromStreptomyces griseus (gsChit), which hydrolytically cleaves β1,4 linkages and Dispersin B (dspB)[282], which cleaves β1,6 linkages. No degradation products were observed when acholetin was in-cubated with either of these enzymes (Figure 5.25: top), further confirming that no β1,4 or β1,6linkages were present. Activity was confirmed for gsChit and dspB by demonstrating they degradechito-oligosaccharides and PNAG, respectively (Figure 5.25: middle and bottom). Acholetin was145Figure 5.25: Acholetin enzymatic degradation assay. spHex, AchP, gsChit and dspB were incubatedwith acholetin (top), chitin (middle) or PNAG (bottom) for 2 h at room temperature. 50 mMphosphate (pH 7.0) was included in all reactions.146degraded, as were chitin and PNAG oligos, by an exolytic β-N -acetylhexosaminidase from Strepto-myces pilcatus (spHex) (Figure 5.25). This, however, was not surprising as the well characterizedspHex is known to cleave non-reducing end GlcNAc residues from a range of glycoconjugates, oligioand polysaccharides [283–285]. This establishes that acholetin is also a substrate for spHex. Insummary, from the two-pot large scale synthesis I recovered 344.3 mg of acholetin with an averageDP of 14.6 and confirmed the homogeneity of β1,3 linkages throughout. Due to the high cost ofATP used to produce GlcNAc1-P, this method may not be economical. However, the ATP expensecould be avoided if GlcNAc1-P was produced through a phosphorylase catalyzed degradation ofchitin instead, which could conceivably make this an economical process suitable to further scaling.5.2.20 Preliminary Structural AnalysisTo gain structural insights into the substrate specificities of AchP, crystals of the enzyme weresoaked with GlcNAc-β1,3-GlcNAc, GlcNAc-β1,4-GlcNAc and 2FGlc-α1-P. In all, five structureswere obtained from our collaborators in Paul Adam’s lab at the University of California, Berkeley.Those structures are the unbound apo version, AchP bound to the substrates listed above and acatalytically inactive, D513A mutant bound to GlcNAc-β1,3-GlcNAc. Unfortunately, at this time,the fully refined versions are not yet ready, however, a preliminary structure is shown in Figure 5.26of wild-type AchP with GlcNAc-β1,3-GlcNAc bound in the active site. It should be emphasizedthat this is only a preliminary structure and any findings generated from its analysis will needto be validated when the full structure is ready. In addition to GlcNAc-β1,3-GlcNAc, a sulphategroup is also found in the active site, likely in the phosphate binding site, as it appears to be inposition to perform nucleophilic attack on the anomeric carbon of the non-reducing end residue.To limit/prevent turnover of the substrate no phosphate was included as part of the crystallizationliquor, but sulphate was present.Based on AchP’s substrate specificities described above, it was apparent that the C-2 NAcgroup of the acceptor’s non-reducing end residue was necessary for binding in the +1 active site.It appears from the structure, that the non-reducing end’s NAc group interacts with R432 (Figure5.26B) suggesting a potential critical interaction to facilitate +1 subsite binding. In the substratespecificity analysis, AchP also showed a preference toward GlcNAc acceptors, over GalNAc. The147Figure 5.26: Co-crystal structure of AchP and GlcNAc-β1,3-GlcNAc (preliminary). (A) Zoomedout view of AchP structure with GlcNAc-β1,3-GlcNAc and SO4 in the active site. (B) Zoomed viewof the +1 binding site focused on the reducing end GlcNAc residue. Side chains which form polarinteractions with the reducing end residue are labelled, as is the SO4, which likely is bound to thephosphate binding site. (C) Zoomed view of the -1 binding site focused on the non-reducing endGlcNAc residue. Side chains which form polar interactions with the non-reducing end residue arelabelled, except for D513, which was hidden as to not obscure the view of the active site.148polar interaction between the GlcNAc residue’s C-4 hydroxyl and E674 may explain this preferenceas the hydroxyl configuration is inverted at C-4 for GalNAc. Both of these observations should betested in future studies by measuring activities from R432 and E674 AchP mutants. I would expectR432 mutants to be catalytically inactive because acceptor recognition would be disabled, while forE674 mutants, I would expect a reduced preference towards GlcNAc over GalNAc.In terms of donor specificity, AchP accommodated GlcNAc1-P, Glc1-P and GalNAc1-P. How-ever, no kinetic parameters could be obtained when GalNAc1-P was used as a donor at reasonableconcentrations, while they could be obtained for GlcNAc1-P and Glc1-P. This indicates the import-ance of the non-reducing end residue possessing an equatorial C-4 hydroxyl, which is shown in thestructure to form a polar interaction with W511 (Figure 5.26C). This hypothesis, as well as others,can be tested by performing future mutational analysis. Despite not having fully resolved structuraldata, the polar interactions observed between AchP side chains and the bound GlcNAc-β1,3-GlcNAcin Figure 5.26 are consistent with the substrate specificity analysis. Specifically, in regards to AchP’spreference toward both GlcNAc donors and acceptors over GalNAc and Glc, the structural analysisrevealed polar interactions with each equatorial hydroxyl and NAc group on both the reducing andnon-reducing end residues of GlcNAc-β1,3-GlcNAc, which provides a potential explanation for theGlcNAc preference.5.2.21 AchP and Acholetin’s Biological RoleA. laidlawii was first identified in 1936 from raw London (England) sewage by Laidlaw and Elford[286]. The organism is a common mycoplasma widely found throughout nature, including in human,animal and plant tissues [287, 288]. It is also a predominant contaminant of cell cultures, whichpresents significant concerns for a range of biotechnology sectors [287, 289, 290]. Mycoplasma areone of the smallest living forms of microbes and notably differ from other bacteria because they lacka cell wall [291]. Research performed on the organism in the first half of the 1970s identified a high-molecular weight polysaccharide weakly associated with the exterior surface of the plasma membrane[292, 293] that is thought to be the major antigenic determinate of A. laidlawii membranes [294].Furthermore, this polysaccharide was shown to contain exclusively GlcNAc and GalNAc residues[292, 294], however, to our knowledge the exact chemical structure (including glycosidic linkage149Figure 5.27: A. laidlawii AchP operon. Graphical representation of the A. laidlawii AchP operon. Annotations were based on myinterpretation of the top BLAST hits from a region of the A. laidlawii genome spanning 100,000 base pairs surrounding the AchPORF. The graphic above represents only a subset of the full BLAST results, which can be viewed in Supplemental Table C.1. Forsome of the annotated functions, reactions were overlaid to help elucidate a potential biochemical pathway. The overlaid reactionsare my interpretation of the ambiguous functional annotations of the top BLAST hits (see Supplemental Table C.1), therefore theycould, and likely in some cases do not reflect the actual function of the enzymes encoded by these ORFs.150type) and functional role of this polyhexosamine has never been determined. I hypothesized thatAchP is involved with the maintenance, in a bio-degradative role, of this membrane associatedpolyhexosamine. To explore this idea further I examined the genes flanking AchP in the A. laidlawiigenome (Figure 5.27). Specifically, I was most interested in identifying a candidate (or candidates)that may be involved with biosynthesis of the polyhexosamine. The AchP ORF in the genomewas flanked by five other CAZyme ORFs, a GH16, three GT4s and a GT that has not yet beenassigned to a family. I decided to first test the non-classified GT (GTnc) because the functionalannotations of the other CAZymes represented in the operon indicated they likely had other rolesand I found the possibility of identifying a new GT family too tantalizing to pass up. Unfortunately,despite significant efforts I was unable to solubly express the GTnc and therefore could not deriveany functional information. Due to time restraints of preparing this thesis I was unable to solve theexpression issues and pursue it further.Another approach I took to identify AchP and acholetin’s biological role was to isolate themembrane associated polyhexosamine from a culture of A. laidlawii and then demonstrate its phos-phorolytic cleavage by AchP. Again, despite significant effort, this ultimately proved unsuccessful,in part due to the literature method isolation used 100 L of culture and no yield was reported [292].I next attempted to devise a labelling assay where a fluorescently tagged, catalytically inactive,AchP mutant would be used to label A. laidlawii membranes by adhering to the polyhexosamine.This method was based on similar labelling assay reported by Eddenden et al. [282] where bac-terial PNAG producers were labeled with a catalytically inactive, fluorescently tagged dspB. Thisalso proved unsuccessful as in proof-of-concept studies I was not able to demonstrate that inactive,fluorescently tagged, D513A mutants of AchP were able to bind to immobilized acholetin stronglyenough to withstand washing steps. A potential reason why this failed for AchP, while it worked fordspB, is because the former is an exo-acting CAZyme and therefore only able to bind to the singlenon-reducing end of each acholetin molecule. dspB, on the other hand, is an endo-acting CAZyme,thus able to bind many sites along the PNAG polymer and multiple molecules of dspB can bindto a single PNAG molecule. These two factors make for a more localized and intense fluorescentlabelling of the polymer when an endo-acting CAZyme is used.Despite significant effort, I was unable to confirm our hypothesis regarding AchP and acholetin’s151biological role at this time. Nevertheless, ongoing studies will prioritize characterizing the GTnc,isolating a sample of of the membrane associated polyhexosamine and generation of AchP and GTncknockout-strains of A. laidlawii (a method for making A. laidlawii KO-strains is known and hasbeen published). If A. laidlawii ’s membrane associated polyhexosamine is indeed an acholetin-like β1-3-linked polysaccharide, this knowledge could have significant impact towards developingnovel agents to limit/prevent A. laidlawii contamination and further our understanding of how A.laidlawii, and Mollicutes more generally, interact with their surrounding environment.5.3 ConclusionsIn conclusion, working in collaboration with the JGI, we generated a phylogenomically diverse GPlibrary that was assayed with the molybdenum blue screening method that had been adapted as asubstrate specificity assay. The 125 member library was found to have a soluble expression rate of 41% which was significantly lower than the 60 % that was reported for the GH1 library. Despite that,each of the 125 GPs were screened with up 20 donor/acceptor combinations and mapped onto SSNs.Several active clones were identified which each have the potential to represent a novel GP activitythat will be explored in future studies. Of that group, 2E04 was selected for further biochemicalcharacterization. The enzyme turned out to represent a new activity from the GH94 family andacted on a newly discovered β1,3-linked GlcNAc bio-polymer. The 2E04 GP originated from thegenome of the cell wall-less Mollicute bacterium Acholeplasma laidlawii, which in combination withthe term, chitin, lead us to name the new bio-polymer, acholetin, and the new GP, acholetinphosphorylase (AchP). Biochemical characterization revealed that AchP prefers GlcNAc donor andacceptors, but also accommodates Glc1-P and GalNAc1-P as donors and GalNAc as an acceptor, anobservation that was supported by preliminary structural analysis. ∼350 mg of purified acholetinwith an average DP of 14.6 was synthesized using a two-pot reaction coupling NahK mediatedGlcNAc1-P production with AchP reverse phosphorolysis. The resulting compound was analyzedby NMR and shown to contain homogeneous β1,3-linkages. Ongoing studies of acholetin will focus oncharacterizing its functional properties and evaluating its potential applications in the biotechnologyand/or medical sectors. Lastly, the biological roles of AchP and acholetin were explored, and it was152speculated that AchP may be involved in the maintenance of a high-molecular weight polysaccharideweakly associated with the exterior surface of A. laidlawii ’s plasma membrane. However, despitesignificant effort I was unable to confirm our hypothesis, but doing so will remain a primary focusof ongoing studies in the Withers lab.153Chapter 6ConclusionThrough the use of rational, genomic and metagenomic exploration techniques I was successful inidentifying and characterizing 11 new GPs, two of which represented previously unknown activities.Through experimental and rational exploration of the GP landscape, I hypothesized the reasonGH3 N -acetylglucosaminidases have a conserved Asp-His dyad catalytic acid/base, as opposed to themore common glutamate or aspartate, was because they were, in fact, phosphorylases. The rationalebehind this hypothesis was that the neutral Asp-His dyad substitution avoids Coulombic repulsionfrom an incoming anionic substrate (such as phosphate), whereas, glutamate or aspartate both bearnegative charges and are therefore less likely to accommodate negatively charged substrates. I testedour hypothesis with the GH3 β-glucosaminidase Nag3 from C. fimi and discovered that it was indeeda phosphorylase that preferentially acted on a substrate possessing a non-reducing end glucosylmoiety. However, at the time, I was unable to determine Nag3’s natural substrate. Nevertheless,Nag3 represented the first GP to be discovered in the GH3 family and the first known GP to operatethrough a β-retaining mechanism. A followup study on hydrolytic GH3 N -acetylglucosaminidases,performed by Ducatti et al. [213], confirmed that while the Glu to Asp-His substitution may benecessary for phosphorylase activity, it is not itself sufficient, thus not predictive of phosphorylaseactivity. Further research into this class of enzymes and the underlying determining factors thatimpart phosphorylase over hydrolase activity is an intriguing research topic for future studies ofGH3 GPs.Prior to the research presented in this dissertation, there had been no reported functionalmetagenomic screening methods targeting GP activity. To be able to explore the vast reservoir ofuncultivated genetic diversity encoded in microbial communities inhabiting natural and engineeredecosystems I designed and deployed two functional screening methods targeting GP activities. The154first, targeted forward phosphorolysis activity and led to the discovery of the second GH3 GP, BglP,for which a crystal structure was solved of the glycosyl enzyme intermediate (2FGlc-BglP). Struc-tural analysis revealed a glutamic acid in the active site adjacent to C-2 that, that when mutatedto a serine, resulted in a substrate specificity swap from a non-reducing end Glc residue to GlcNAc,providing new insights into simple ways in which enzyme specificity can be encoded and swapped.The second functional metagenomic screening method targeted reverse phosphorolysis activity bycoupling release of phosphate to the formation of molybdenum blue. This method led to the dis-covery of seven new GH94 GPs and one new GH149 GP, the latter representing the first full-lengthcharacterized bacterial GH149 to enter the public domain. Moreover, the small-scale validationassay confirmed that this screening method is compatible with additional donors beyond Glc-α1-P.These two methods provide a platform for future screening efforts to discover a wide range of GPsfrom diverse environmental genomic material and expand the breadth of known GP activities.The molybdenum blue metagenomic screening assay was modified and adapted as a broad GPsubstrate specificity assay and used to explore a phlogenomically diverse GP library. Several inter-esting activities were identified from the 125 member synthetic gene library that warrant furtherinvestigation in future studies. Additionally, a HT substrate specificity protocol was establishedthat can be integrated into the JGI pipeline and therefore further aid in the rapid characterizationof GP libraries. The JGI pipeline could become even more powerful if it were to adopt the Toulousegroup’s strategy involving SSNs, allowing for better targeting of GPs that are most likely to repre-sent new, desired or interesting activities. Followup characterization of one of the 125 members ofthe GP library resulted in the discovery of a novel GH94 β-1,3-N -acetylglucosaminide phosphorylaseand a new poly-β1,3-linked GlcNAc biopolymer, acholetin. Exploring the potential biological rolesof AchP and acholetin lead us to hypothesized that AchP may be involved with the maintenance,in a bio-degradative role, of a membrane associated polyhexosamine, similar to acholetin, found onthe exterior surface of A. laidlawii membranes. Unfortunately, despite significant efforts to confirmour hypothesis, I was unable to isolate the membrane associated polyhexosamine. Continuing re-search in the Withers lab will strive to understand the biological roles of AchP and acholetin, inaddition to ongoing research to analyze a non-classified GT that I speculated is involved in acholetinbiosynthesis, and to generate AchP KO-strains of A. laidlawii.155Through the implementation of rational, genomic and metagenomic exploration approaches Ihave discovered and characterized new GP activities that have contributed to the better under-standing of these useful biocatalysts. In addition, this work has resulted in the development of asuite of tools that can be applied toward future GP discovery endeavours and has the potential tosignificantly expand the range of GP activities that, in turn, can be applied to make medicines,food products and the next-generation of carbon neutral, bio-sourced materials.156Chapter 7Methods7.1 General MethodsIn all instances error bars represent standard error (SE) as calculated using the equation shownbelow:SE =√∑Ni=1(xi−x¯)2N−1√N(7.1)Where {x 1, x 2,...,xN } are measured values, x¯ is the mean of those values and N is the number ofvalues measured.7.2 Data AccessioningFor Chapter 3: Fosmid sequences for 29K06 (MF625023) and 31P01 (MF625024) can be found inGenBank. Coordinate files and structure factors have been deposited to the Protein Data Bankunder accession codes 5VQD for native BglP and 5VQE for BglP bound to 2FGlc.For Chapter 4: Fosmid sequences have been deposited in GenBank under ID codes: MK411235for BCR_17C21, MK411236 for BCR_32A05, MK4112367 for BCR_32B15, MK411238 for BCR-_38M24, MK411239 for BCR_42I14, MK411240 for BCR_45N22, MK411241 for BCR_46A06,MK411242 for BVR_06I15, MK411243 for BVR_10H18, MK411244 for BVR_11F19 and MK-411245 for BVR_11G05.1577.3 Chapter 2 Experimental7.3.1 Materials and ReagentsSubstrates used in this study were either purchased from suppliers or donated by members ofthis laboratory. p-Nitrophenyl N -acetyl-β-D-glucosaminide (pNPGlcNAc) was purchased from Cal-biochem, p-nitrophenyl β-D-glucopyranoside (pNPGlc) was purchased from Sigma-Aldrich. 2,4-dinitrophenyl β-D-glucopyranoside (DNPGlc) and 2,4-dinitrophenyl 2-deoxy-2-fluoro-β-D-glucopy-ranoside (DNP2FGlc) were generously donated by H. Chen and F. Liu, respectively. Concen-tration/removal of Nag3 was done using Amicon Ultra - 0.5 mL centrifugal filters (30,000 MW)(Millipore, Darmstadt, Germany). TLC was done using TLC silica gel 60 F254 TLC plates (EMDMillipore Corporation, Billerica, MA, USA). TLC stain composition: anisaldehyde reagent (92.5 %ethanol, 4 % H2SO4, 1.5 % acetic acid, 2 % p-anisaldehyde), and molybdate reagent (2.5 % w/vammonium molybdate, 1 % w/v ceric ammonium sulfate and 10 % H2SO4).7.3.2 General Kinetic MethodsC. fimi Nag3 was expressed and purified as previously reported [193] substituting HEPES buffer(50 mM HEPES pH 7.0, 300 mM NaCl, 5 mM MgSO4, 10 % glycerol, 1 mM DTT) where sodiumphosphate buffer was previously used. The enzymatic reaction rates with the chromogenic substrateswere determined by monitoring the change in absorbance at 400 nm. Kinetic measurements wereperformed in matched 1 cm path length quartz cuvettes using a Varian Cary 300 Bio UV-visiblespectrophotometer with an automatic cell changer and temperature controller, at 25 °C in HEPESbuffer. Reactions contained potassium phosphate (pH 7.0) where specified. Hydrolysis/phosphorolysis rates were calculated by measuring absorbance changes as a function of time and convertingthese to concentration with the following extinction coefficients: 7280 M-1cm-1 (pNP) and 12460M-1cm-1 (DNP) at 25 °C, pH 7.0. Rate data was fit using the Michaelis-Menten equation incorporat-ing substrate inhibition:v0 =Vmax · [S]Km + [S](1 + [S]Ki) (7.2)158Lineweaver-Burk plots were fit to the equation for the Ping Pong mechanism:1v0=(KaVmax)(1[A])+(1Vmax)(1 +Kb[B])(7.3)Non-linear regression was performed using GraphPad Prism v6.0 or GraFit.7.3.3 Pre-steady State KineticsThe pre-steady state phase of DNPGlc binding to Nag3 was captured by performing the experimentin a spectrophotometer cuvette cooled to 12 °C and continuously monitoring absorbance at 400 nm.An assay buffer containing HEPES buffer and DNPGlc was pre-incubated in a quartz cuvette at12 °C. Once the baseline was established 40 µL of Nag3 (20 mg/mL) was added to 160 µL assaybuffer to give final concentrations of 4 mg/mL Nag3 (66 µM) and 250 µM DNPGlc. At 0.5 min,after steady state is observed, phosphate (pH 7.0) was added to a final concentration of 50 mM.7.3.4 1H-NMR Analysis10 mM substrate (pNPGlcNAc, pNPGlc or DNPGlc) was incubated with 6 mg/mL Nag3 and 0, 5,25 or 100 mM phosphate in HEPES buffer (with no glycerol) for 2 h (500 µL reaction volume). Theincubation temperature was increased to 37 °C (compared to 25 °C used for kinetic assays) to helpmaximize substrate turnover. After the incubation time Nag3 was removed with Amicon centrifugalfilters and the filtrate was freeze-dried using a SpeedVac (Savant SV 100) concentrator and dissolvedin D2O (500 µL). 1H-NMR spectra were recorded on a Bruker 400 MHz Avance spectrometerwith inverse probehead. Per sample, 16 scans employing presaturation water suppression (Brukerstandard pulse program “zgcppr”) were recorded.7.3.5 GlcNAc-β1-P and Glc-β1-P Isolation and Characterizationβ-glucose, and β-GlcNAc 1-phosphates were prepared by combining pNPGlc or pNPGlcNAc (10mM final concentration) with 1 mg/mL Nag3 and 50 mM phosphate in HEPES buffer (10 mLreaction volume). After incubation for 4 h at 25°C the samples were freeze-dried using a SpeedVac(Savant SV 100) concentrator. To purify, the dried sample was dissolved in H2O and the pH159adjusted to 5. A short column was filled with Dowex 50WX8 ion exchange resin (5 g) and theresin washed with deionized water (100 mL). The sample was applied and 10 mL fractions weretaken, which were immediately neutralized (NaOH) after collection. The collected fractions wereconcentrated (SpeedVac), pooled and applied in H2O (∼ 500 µL) to a Biogel P2 column (60x1.5cm, eluent: H2O, flow rate 0.5 mL/min, fraction size 3.5 mL). Positive fractions were identified byTLC, pooled, concentrated (SpeedVac) and analyzed by NMR. Spectra were recorded in D2O andreferenced externally to dioxane in D2O (1H: 3.75 ppm, 13C: 67.4 ppm) or H3PO4 in D2O (31P:0 ppm). High Res mass spectra were obtained on a Micromass LCT time-of-flight (TOF) massspectrometer equipped with an electrospray ion source.7.3.6 TLC AnalysispNPGlcNAc, pNPGlc or DNPGlc (10 mM final concentration) was incubated with 20 mg/mL Nag3and 0, 1, 5, 10, 25, 50 or 100 mM phosphate in HEPES buffer (with no glycerol) at 37 °C for 15 min(20 µL reaction volume). The reaction mixtures were placed on ice to "stop" reactions before beingspotted on silica TLC plates. The plates were eluted with a mobile phase of 1-butanol, methanol,ammonium hydroxide and water in a 5:4:4:1 ratio, respectively. Once dried the pNPGlcNAc platewas stained with anisaldehyde reagent, the pNPGlc and DNPGlc plates were stained with molybdatereagent and heated until the product signals became visible. Anisaldehyde reagent was used for thepNPGlcNAc plate to better visualize GlcNAc formation.7.3.7 Electrospray Mass Spectrometry1 mg/mL Nag3 (in 20 mM Tris pH 7.0, 50 mM NaCl, 2 % glycerol, 5 mM MgSO4 and 1 mMDTT) was incubated with 1 mM DNPGlc or 1 mM DNPGlc and 50 mM phosphate for 1 h at roomtemperature (50 µL reaction volume). Samples were analyzed using methods described previously[195, 295].7.3.8 Reactivation with PhosphateNag3 was inactivated by incubating 2 mg/mL of active enzyme with 75 mM of the inactivatorDNP2FGlc at 25 °C for 5 h in HEPES buffer (50 µL). Excess DNP2FGlc was diluted to ∼2 µM160by dilution with HEPES buffer and re-concentrating with Amicon centrifugal filters. Reactivationwas monitored by incubating samples of the inactive Nag3-2FGlc complex, (0.2 mg/mL) in HEPESbuffer with 0, 10, 25, 50, 100 or 200 mM phosphate at 25 °C (50 µL reaction volume) and transferring5 µL aliquots at the indicated time points from the reactivation reaction to an assay solution (195mL) containing HEPES buffer, 0.5 mM DNPGlc and 20 mM phosphate. Turnover rates werecalculated as described above.7.4 Chapter 3 Experimental7.4.1 Materials, Reagents and General MethodsAll buffers and reagents were purchased from Sigma-Aldrich, unless noted otherwise. Substratesused in this study were either purchased from suppliers or donated by members of this laboratory.p-Nitrophenyl N -acetyl-β-D-glucosaminide (pNPGlcNAc) was purchased from Calbiochem, p-nitro-phenyl β-D-glucopyranoside (pNPGlc) was purchased from Sigma-Aldrich. 2,4-Dinitrophenyl β-D-glucopyranoside (DNPGlc), 2,4-dinitrophenyl 2-deoxy-2-fluoro-β-D-glucopyranoside (DNP2FGlc),6,8-difluoromethylumbelliferyl β-D-glucopyranoside (DFMUGlc) and 6,8-difluoromethylumbelliferylN -acetyl-β-D-glucosaminide (DFMUGlcNAc) were generously donated by H. Chen and F. Liu. En-zyme quantification was performed using the Bradford method [296] or by measuring enzyme ab-sorbance at 280 nm and using a molar absorption coefficient (280) of 76945 M-1cm-1 to calculateconcentration using the Beer-Lambert law [297]. All TLC assays were done on silica gel 60 F254 TLCplates (EMD Millipore Corporation, Billerica, MA, USA). Plates were eluted with a mobile phaseof 1-butanol, methanol, ammonium hydroxide and water in a 5:4:4:1 ratio, unless otherwise stated.Following elution, plates were dried then dipped in a molybdate TLC stain (2.5 % ammoniummolybdate (w/v), 1 % ceric ammonium sulfate (w/v) and 10 % H2SO4 (v/v)) or p-anisaldehydeTLC stain (92.5 ethanol, 4 % H2SO4 (v/v), 1.5 % acetic acid (v/v), 2 % p-anisaldehyde (v/v)).Visualization of TLC plates was done by heating until the product spots became visible. PCRprimers and cycle conditions can be found in Tables 7.1 and 7.2 and buffer recipes can be found inTable 7.3.161Table 7.1: PIPE and QuickChange PCR parameters used for insert and vector amplification. SeeTable 7.2 for annealing temperatures (X ) and extension times (Y ).Temperature (°C) Time95 2 min95 30 sX 30 s72 Y min15 holdTable 7.2: Partially incomplete polymerase extension (PIPE) PCR details.Construct Primers (5’→3’) Template Template(ng/PCR)AnnealingTemp (°C)ExtensionTime (min)pET28-HtCBP.h6V F:GAAGTAATTATGGGACACCATCACCATCACCATTGAGATCCGGCTGCT pET28a 5 55 4R: TCATCAAAAAAACCGAACTTCATGGTATATCTCCTTCTTAAAGI F:TAAGAAGGAGATATACCATGAAGTTCGGTTTTTTTGA H. thermocellumgenomic DNA(ATCC 27405)5 51 2R:AGCAGCCGGATCTCAATGGTGATGGTGATGGTGTCCCATAATTACTTCAACTpET28-HtCDP.h6V F:ACTCTTAAGTTTAAACACCATCACCATCACCATTGAGATCCGGCTGCT pET28a 5 55 4R: CTCGCTGTTACTTTAGTAATCATGGTATATCTCCTTCTTAAAGI F:TAAGAAGGAGATATACCATGATTACTAAAGTAACAGCGAG H. thermocellumgenomic DNA(ATCC 27405)5 51 2R:AGCAGCCGGATCTCAATGGTGATGGTGATGGTGTTTAAACTTAAGAGTCACTATATGpET28-BglP.h6V F:GGTCTGTGGGACGCACGCCTACACCATCACCATCACC pET28-HtCBP.h6 5 56 4R: GAAAGAATGGATTCGTACATGGTATATCTCCTTCTTAAAGTI F:GAAGGAGATATACCATGTACGAATCCATTCTTTCCATCAAAC FOS_29K06 5 72 1R: AATGGTGATGGTGATGGTGTAGGCGTGCGTCCCACAGACCpET28-BglP.h6.Q205S QCF: GATTATCGGGACTCACATCTGATGTCC pET28-BglP.h6 50 60 3.5R: GGACATCAGATGTGAGTCCCGATAATC162Table 7.3: Buffer Recipes. Summary of buffers used for the functional DNPGlc metagenomic screen,expression and immobilized metal affinity chromatography purification of HtCBP, HtCDP and BglP.BufferNamePrimarycomponentsAddedImidazole(mM)Buffer Aloading50 mM HEPES (pH 7.0),100 mM NaCl,2 % glycerol,5 mM MgSO45wash 25elution 600storage 0Buffer Bloading50 mM HEPES (pH 7.0),300 mM NaCl,20 % glycerol,5 mM MgSO4,0.1 mM DTT5wash 25elution 600storage 02x assay buffer S200 mM potassium phosphate (pH 7.0),2 % triton X-100,40 mM NaCl,200 µM DNPGlc2x assay buffer H100 mM HEPES (pH 7.0),2 % triton X-100,40 mM NaCl,200 µM DNPGlc2x assay buffer P100 mM HEPES (pH 7.0),100 mM potassium phosphate (pH 7.0),2 % triton X-100,40 mM NaCl,200 µM DNPGlc1637.4.2 Plasmid ConstructionpET28-HtCBP.h6, pET28-HtCDP.h6 and pET28-BglP.h6 expression constructs were made accord-ing to the partially incomplete polymerase extension (PIPE) cloning method [298]. pET28-BglP.h6-.Q205S mutant was constructed using the QuikChange® PCR site directed mutagenesis method.HtCBP and HtCDP ORFs were amplified from H. thermocellum genomic DNA and the BglP ORFfrom FOS62_29K06 fosmid DNA. A C-terminal hexa-histidine tag was incorporated into each ORFthrough primer design. For pET28-HtCBP.h6, pET28-HtCDP.h6 and pET28-BglP.h6, heat shocktransformations were performed by combining 2 µL of the vector PCR product with 2 µL of theinsert PCR product and then 100 µL of DH5α chemically competent cells [299]. For pET28-BglP.h6.Q205S, 10 µL of PCR product (cleaned up with QIAquick PCR purification kit) was trans-formed with 100 µL of DH5α chemically competent cells. Constructs were confirmed by sequencing(Genewiz®, South Plainfield, NJ, USA) using T7 and T7term sequencing primers and subsequentlytransformed into BL21 (DE3) chemically competent cells [299].7.4.3 Protein Expression and PurificationCellulomonas fimi Nag3 was expressed and purified, as reported previously [85]. 2 L (HtCBP,HtCDP and BglP.Q205S) or 4 L (BglP) of LB media containing 25 µg/mL kanamycin was inoculatedwith 1/100 of overnight culture. Expression cultures were grown at 37 °C until OD600 = 0.5 (∼ 3h). Cells were induced with 0.5 mM IPTG and grown for an additional 3 h at 37 °C (HtCBP andHtCDP) or 18 h at 16 °C (BglP and BglP.Q205S). Cells were harvested by centrifuging at 6 000 x gfor 6 min in a Beckman Coulter Avanti® J-E floor centrifuge (JA-10 rotor) followed by re-suspension(20 mL per 1 L of original culture volume) in loading buffer A (HtCBP and HtCDP) or loadingbuffer B (BglP and BglP.Q205S) (Table 7.3). Cells were lysed with an Avestin C3 homogenizerwith an average cell pressure of 16 000 psi. The soluble fraction was isolated by centrifuging thelysate at 15 000 rpm for 30 min (JA-20 rotor). The soluble fraction was either stored at -20 °Cuntil needed or immediately purified. Purification was carried out by immobilized metal affinitychromatography (IMAC) on a GE ÄKTA FPLC equipped with a UV and conductance detector andan automatic fraction collector. Buffers used for protein purification are detailed in Table 7.3. For164each enzyme a separate 5 mL HisTrap™ FF column (GE) was equilibrated with 10 column volumesof loading buffer A (HtCBP and HtCDP) or loading buffer B (BglP and BglP.Q205S). Soluble celllysates were applied to the columns using a P-1 peristaltic pump (GE) followed by a wash step of10 column volumes (CV) of the respective loading buffers. The columns were then transferred tothe ÄKTA and washed again with 10 CV of wash buffer A (HtCBP and HtCDP) or wash bufferB (BglP and BglP.Q205S) followed by equilibration with 10 CV of the respective loading buffers.Proteins were eluted using a 20 mL gradient (0 - 100 %) of loading buffer to elution buffer withthe automatic fraction collector set to collect 1 mL fractions. Fractions were analyzed on SDSPAGE and those yielding the biggest band at 94 kDa for HtCBP, 113 kDa for HtCDP or 65 kDafor BglP and BglP.Q205S were combined and concentrated using an Amicon® Ultra-4 MWCO 30kDa centrifugal filter (Sigma-Aldrich), then dialysed against storage buffer A (HtCBP and HtCDP)or storage buffer B (BglP and BglP.Q205S) and stored at -70 °C.7.4.4 HtCBP, HtCDP and Nag3 DNPGlc Phosphorolysis AssaySpectroscopic assay: DNPGlc (2 mM) was incubated with 25 µg of purified HtCBP, HtCDP orNag3 and 0 or 50 mM phosphate in storage buffer A (200 µL reaction volume) and incubated at 37°C for 1 h. A400 measurements were performed in matched 1 cm path length quartz cuvettes using aVarian Cary 300 Bio UV-visible spectrophotometer with an automatic cell changer and circulatingwater bath. TLC assay: DNPGlc (20 mM) was incubated with 37.5 µg of purified HtCBP, HtCDPor Nag3 and 0 or 50 mM phosphate in storage buffer A (10 µL reaction volume) and incubated at 37°C for 2 h. Samples (0.5 µL) of each reaction were spotted onto TLC plates and run and visualizedas described above.7.4.5 BCR Library Function ScreenThe screening methodology was modeled on the functional metagenomic screen reported by Mewiset al [176]25. A generalized workflow of the screening method is shown in Figure 4.5. BCR li-brary plates were replicated into fresh 384-well plates (Nunc™ 384-well clear polystyrene plates,non-treated) containing 50 µL LB media per well with 100 µg/mL arabinose and 12.5 µg/mL chlo-ramphenicol using a QPix2 robot. The replicated plates were grown for 18 h at 37 °C. The assay165was performed by adding 50 µL of 2x assay buffer S (Table 7.3) and incubating at 37 °C for 6 h,then absorbance measurements were taken at 400 nm. 56 fosmid clones that displayed an A400 valuegreater than mean + 4 SD were re-arrayed to a 96-well GP master plate.7.4.6 BCR GP Master Library Screen and ValidationGP master library screen: The BCR GP master plate was replicated to six identical 96-well plates(Costar® 96 well flat bottom polystyrene plate) containing 100 µL LB media per well with 100µg/mL arabinose and 12.5 µg/mL chloramphenicol, then grown for 18 h at 37 °C. The plates werethen screened in the presence and absence of 50 mM phosphate, each condition in triplicate. Either100 µL 2x assay buffer P or 2x assay buffer H (Table 7.3) was added to each well and incubatedat 37 °C for 6 h. After the incubation period absorbance measurements were taken at 400 nm ina BioTek Synergy H1 Hybrid microtiter 96 well plate reader. A Student’s t-test was performedbetween the averages of the A400 values of each clone in the absence and presence of phosphateusing Prism GraphPad v6.0 software and according to Equation 7.4.tstat =x¯− y¯√(sp2nx) + (sp2ny)(7.4)Where x¯ is the average of the A400 readings measured in the presence of 50 mM phosphate andy¯ is the average in the absence of phosphate. nx and ny are the number of samples measured undereach condition and sp is the pooled standard deviation defined in Equation 7.5.sp2 =(nx − 1)sx2 + (ny − 1)sy2df(7.5)Where df (degrees of freedom) is described by Equation 7.6, sx2 is described by Equation 7.7and sy2 is described by Equation 7.8.df = nx + ny − 2 (7.6)sx2 =∑Ni=1(xi − x¯)2nx − 1 (7.7)166sy2 =∑Ni=1(yi − y¯)2ny − 1 (7.8)tstat values were converted to their corresponding p-values using Prism GraphPad v6.0 softwareand clones possessing a significantly (p < 0.05) higher activity with phosphate present were analyzedfurther by TLC. GP TLC validation: 2 x 100 µL of overnight cell culture from each of the 12 fosmidclones were spun down and the supernatant removed. Each cell pellet pair was resuspended with 10µL 2x assay buffer H, 2 µL 100 mM DNPGlc and 8 µL dH2O or 8 µL 125 mM potassium phosphate(pH 7.0) and incubated for 2 h at 37 °C. Samples (0.5 µL) of each reaction condition were spottedonto TLC plates and run as described above.7.4.7 Full Fosmid SequencingFosmid DNA was extracted from clones FOS62_29K06 and FOS62_31P21 using the GeneJETPlasmid Miniprep Kit (Thermo Scientific) according to the manufacturer’s instructions. Fosmidpreparations were further treated with PlasmidSafe DNase (Epicentre) to degrade contaminatingE. coli chromosomal DNA. DNA concentrations were measured with Quant-iT™ dsDNA HS AssayKit (Invitrogen) using a Qubit® fluorometer (Invitrogen). For full fosmid sequencing, 2.4 ng of eachfosmid was sent to UBC Sequencing Centre (Vancouver, Canada). Each fosmid was individuallybarcoded and sequenced using the MiSeq system.7.4.8 Contig Assembly, Open Reading Frame Prediction and GeneIdentificationAll Illumina MiSeq raw sequence data were trimmed and assembled using a python script availableon GitHub at https://github.com/hallamlab/FabFos. Briefly, Trimmomatic was used to removeadapters and low-quality sequences from the reads [300]. These reads were screened for vector andhost sequences using Burrows-Wheeler Aligner (BWA) [301], then filtered using samtools and abam2fastq script to remove contaminants. These high-quality and purified reads were assembledby MEGAHIT with k-mer values ranging between 71 and 241, increasing by increments of 10 [302].Since these libraries often had in excess of 20,000 times coverage, to prevent the accumulation of167sequencing errors interfering with proper sequence assembly, the minimum k-mer multiplicity wascalculated by 1 % of the estimated coverage of a fosmid. Outside of the python script, assemblieswhich yielded more than one contig were then scaffolded using minimus2 [303]. Parameterizedcommands can be found in both documentation on the GitHub page and in the python scriptitself. Fosmid ORFs were identified using the metagenomic version of Prodigal [304] and comparedto the CAZy database using BLASTP as part of the MetaPathways v2.5 software package [305].MetaPathways parameters: length > 60, BLAST score > 20, blast score ratio > 0.4, EValue < 1 x10-6.7.4.9 Kinetic AnalysisKinetic parameters were determined by measuring enzymatic reaction rates with the chromogenicsubstrates and monitoring the change in absorbance at 400 nm. Measurements were performed intriplicate in matched 1 cm path length quartz cuvettes using a Varian Cary 300 Bio UV-visiblespectrophotometer with an automatic cell changer and temperature controller, at 25 °C in BufferB (Table 7.3) containing the indicated concentration of substrate and potassium phosphate (pH7.0). 150 µL Buffer B was premixed with 20 µL 10x phosphate solution and 10 µL of 20x sub-strate solution. The reaction was initiated by the addition of 20 µL of 2 µM BglP or BglP.Q205Sfor DNPGlc, 20 µM BglP or BglP.Q205S for pNPGlc or 50 µM BglP or 20 µM BglP.Q205S forpNPGlcNAc reactions and the change in absorbance at 400 nm was measured for 5 min. Hydroly-sis/phosphorolysis rates were calculated by measuring absorbance changes as a function of time andconverting these to concentration with the following extinction coefficients: 7280 M-1cm-1 (pNP)and 12460 M-1 cm-1 (DNP) at 25 °C, pH 7.0. Substrate final concentrations: DNPGlc: 2.5, 10, 100,500, 1000 and 1500 µM. pNPGlc: 0.5, 1, 2.3, 3.6, 5, 10 and 25 mM. pNPGlcNAc: 0.1, 0.25, 0.5, 1,2, 5 and 10 mM. Concentrations of phosphate were chosen to encompass apparent Km values wherepossible. Non-linear regression was performed using GraphPad Prism v6.0.7.4.10 Electrospray Mass Spectrometry1 mg/mL BglP (in 20 mM Tris, pH 7.0, 300 mM NaCl, 10 % glycerol (v/v), 5 mM MgSO4 and0.1 mM DTT) was mixed, on ice, with 10 mM DNPGlc or 10 mM DNPGlc and 50 mM sodium168phosphate (pH 7.0). 10 µL reaction samples were immediately diluted 1 in 1000 (5 % acetonitrile,0.1 % formic acid, pH 2.2). Samples were analyzed using methods described previously [85, 195,306].7.4.11 Pre-Steady State KineticsPre-steady state phases of the reaction of BglP with 6,8-difluoromethylumbelliferyl β-D-gluco-pyranoside (DFMUGlc) and (separately) 6,8-difluoromethylumbelliferyl N -acetyl-β-D-glucosaminide(DFMUGlcNAc) were monitored in a temperature controlled fluorimeter (Varian Cary Eclipse flu-orescence spectrophotometer) cooled to 12 °C. Excitation of 353 nm (slit length 2.5 nm) was usedand the fluorimeter was set to continuously monitor emission at 451 nm (slit length 5 nm) for theduration of the experiment. A solution containing 980 µL of pre-chilled (12 °C) assay buffer (50mM HEPES pH 7.0, 300 mM NaCl, 10 % glycerol (v/v), 5 mM MgSO4, 0.1 mM DTT) and either10 µM DFMUGlc or DFMUGlcNAc in a 4.5 mL plastic cuvette (Fisherbrand™ Disposable PlasticCuvette, UV-VIS, CLR SIDE, Methacrylate) was used to establish a baseline. After approx. 1 min,20 µL of BglP (2.1 µM for DFMUGlc or 21 µM for DFMUGlcNAc) was added to the assay bufferand the reaction was allowed to proceed for 15 min. The fluorescence signal intensity was convertedto DFMU concentration using a standard curve of the fluorophore in the same assay buffer.7.4.12 Inactivation and Reactivation of BglPInactivation: In individual 50 µL reactions 5 µM BglP was combined with 0, 2, 5, 10, 20, 25, 50 and75 mM DNP2FGlc in Buffer B (Table 7.3) and incubated in a 25 °C circulating water bath. At 0, 10,30, 60, 90 and 120 min after beginning incubation 5 µL of each reaction was transferred to a matched1 cm path length quartz cuvette containing 50 mM pNPGlc in 195 µL Buffer B and the changein absorbance at 400 nm was measured over 5 min. Absorbance was converted to concentrationusing the extinction coefficient for pNP (7280 M-1 cm-1). Rates were measured at each time point,then plotted against time and fitted to a first order decay expression using Prism GraphPad v6.0to give apparent rate constants (kinact) for inactivation at each concentration of DNP2FGlc. kinactwere then plotted versus DNP2FGlc concentration and fit to a Michaelis Menten-like expressionto yield values of ki and Ki. Reactivation: In 50 µL, 100 µM BglP was incubated with 50 mM169DNP2FGlc for 210 min at 25 °C. Inactivation was confirmed by monitoring pNPGlc hydrolysisthroughout the duration of the incubation period. Following inactivation, excess DNP2FGlc wasdiluted to ∼ 0.1 µM by sequential dilutions with reactivation (RA) buffer (50 mM HEPES pH 7.0,300 mM NaCl, 10 % glycerol (v/v), 5 mM MgSO4, 0.1 mM DTT) and re-concentration steps withAmicon® Ultra – 0.5 mL centrifugal filters (30,000 MW) (Millipore, Darmstadt, Germany). Afterthe final re-concentration step inactive BglP was adjusted to 100 µL final volume with RA buffer.Inactive BglP (5 µL) was incubated at 25 °C with 45 µL RA buffer supplemented with 0, 1, 5, 10,25, 50 or 100 mM sodium phosphate pH 7.0. At 0, 12, 25, 60, 98, 130 and 170 min after beginningincubation 5 µL of each reaction was transferred to a quartz cuvette containing 50 mM pNPGlc in195 µL of RA buffer. Rates were measured at each time point, then plotted against time and fittedto a first order increase expression to give apparent rate constants (kreact) for reactivation at eachconcentration of phosphate. kreact were then plotted versus phosphate concentration and fit to aMichaelis Menten-like expression to yield values of kp and Kp.7.4.13 X-Ray Crystallographic Structure DeterminationThe methods in this section were performed by members of the Mark Lab at the University of Man-itoba. To produce BglP in quantities sufficient for crystallization trials, E. coli BL21-Gold (DE3)harboring the pET28-BglP.h6 expression plasmid were grown in 500 mL of LB media supplementedwith 35 µg/mL kanamycin at 37 °C to an OD600 of 0.5-0.6, then induced with 1 mM IPTG andgrown for an additional 18-20 hrs at 16 °C with shaking. The culture was pelleted by centrifugation,resuspended in lysis buffer (50 mM HEPES pH 7.0, 500 mM NaCl, 10 % glycerol (v/v) and 5mMimidazole), and lysed using a French pressure cell press (Aminco). The lysate was centrifuged at17,000 x g for 1 h at 4 °C. The supernatant was mixed with 1.5 mL of Ni-NTA resin (Qiagen) for 1 hat 4 °C, then poured into a gravity column and washed with 25 mL of lysis buffer, followed by 25 mLof lysis buffer supplemented with 25 mM imidazole. Recombinant BglP was eluted from the washedresin using lysis buffer supplemented with 250 mM imidazole and dialyzed overnight against 500 mMNaCl, 50 mM Tris-Cl (pH 7.5), 10 % glycerol (v/v), and 1 mM DTT. The protein was concentratedand loaded onto a size exclusion gel filtration column (Superdex 75) pre-equilibrated with dialysisbuffer. Fractions containing BglP were pooled and concentrated using an Amicon Ultra Centricon170spin cartridge (Merck Millipore). When concentrated to 5 mg/mL, BglP crystallized after one dayin a mother liquor composed of 27 % PEG 1000 and 100 mM MES (pH 6.5), by hanging-dropvapor diffusion. To obtain a complex of BglP with DNP2FGlc, crystals of the protein were soakedfor 1 h with the ligand at a final concentration of 20 mM. Both native and ligand-bound BglPwere subsequently flash-cooled in liquid N2 using the mother liquor above as a cryosolution. X-raydiffraction data were collected in-house at 100 K using a Rigaku MicroMax HF X-ray generatorand R-AXIS IV++ image plate detector. Data were indexed using MOSFLM[307] and scaled usingAimless [308]. A molecular replacement (MR) search model was generated by SCULPTOR [309] us-ing the crystal structure of a family GH3 N -acetylglucosaminidase (PDB ID: 3BMX) and a pairwisesequence alignment of the search model with BglP generated in GENEIOUS 8.1.7 [310]. MR wascarried out using PHENIX.PHASER [311] to generate initial phase estimates for reflections collectedfrom a crystal of BglP bound to 2FGlc, followed by model building using PHENIX.AUTOBUILD[311] and iterative model improvement using COOT [312] and PHENIX.REFINE [311]. A modelof 2FGlc covalently linked to BglP was fitted into its ascribed density using COOT and restrainedduring refinement using geometric restraints generated by PHENIX eLBOW [311]. Initial phaseestimates for native BglP were obtained using the refined 2FGlc-bound model from which 2FGlcand solvent had been removed as an MR search model in PHENIX.PHASER followed by iterativemodel building and refinement using COOT and PHENIX.REFINE. A model of bound 2FGlc1-Pwas built based on the GlcNAc-MurNAc substrate bound to a GH3 NagZ from Bacillus subtilis(PDB ID: 4GYJ) and 2-deoxy-2-fluoro-α-D-glucose 1-phosphate (gfp_msd.pdb) from the HIC-updatabase [313]. MurNAc was replaced in the GlcNAc-MurNAc substrate with the phosphate groupof gfp while maintaining the distorted conformation of GlcNAc and pseudoaxial orientation of theglycosyl ester linkage connecting C1 of the sugar to phosphate. The 2-acetamide group of GlcNAcwas then replaced with a fluorine to complete the model, which was fitted into the active site usingthe experimentally determined BglP-2FGlc complex as a guide.7.4.14 Reactivator ScreenInactivated BglP was prepared by incubating 380 µl enzyme (13.5 mg/ml) with 20 µl 500 mMDNP2FGlc (final concentration 25 mM) for 210 min at 25 °C. BglP inactivation was confirmed171by monitoring pNPGlc hydrolysis throughout the duration of the incubation period (Figure 3.18).Following inactivation, excess DNP2FGlc was diluted to ∼0.1 µM by sequential dilutions with BufferB (Table 7.3) and re-concentration steps with Amicon centrifugal filters (30 kDa cutoff). After thefinal re-concentration step inactive BglP was diluted to 1 ml final volume with Buffer B and theenzyme concentration was determined to be 3 mg/ml using a Bradford assay [296]. The reactivatorspecificity assay was performed in a 96 well plate (Costar® 96 well flat bottom polystyrene plate)by combining 5 µl of inactive BglP, 8 µl 5x assay buffer (100 mM HEPES pH 7.0, 1.5 M NaCl, 50% glycerol, 25 mM MgSO4, 0.5 mM DTT), 16 µl activator (to a final concentration of 20 mM or25 % saturation) and 35 µl dH2O per well and incubated at 25 °C for 3 h. After incubation 8 µlof 225 mM pNPGlc was added to each well and the release of the pNP was monitored in a BioTekSynergy H1 Hybrid microtiter 96 well plate reader by measuring continuous change in absorbanceat 400 nm over 60 min.7.4.15 BglP Reactivation Product AnalysisIn order to avoid potential interference during product analysis from buffer components, mainlyglycerol, BglP was exchanged into a new buffer (25 mM HEPES, pH 7.0, 50 mM NaCl) using 0.5mL Amicon® Ultra Centrifugal Filters (30 kDa cutoff). This new preparation of BglP was used forsubsequent reactivation product analysis except for the initial TLC product screen (Figure 3.20)where the BglP storage buffer contained 20 % glycerol.Glc-β1-P-α1-GlcESI-MS : 10 µL of BglP (∼20 mg/mL), 10 µL DNPGlc (100 mM), 10 µL Glc-α1-P (100 mM)and dH2O were combined and incubated for 10 min at room temperature. BglP was removed byincubating the reaction mixture at 70 °C for 5 min then separating the precipitate by centrifugation.The mass spectrum was obtained on a HCTultra PTM Discovery System by electrospray ionizationmass spectrometry (ESI-MS) with detection in negative ion polarity mode. 1H NMR: FollowingESI-MS analysis, the sample prepared above was purified by preparative TLC (mobile phase: 1-butanol:methanol:NH4OH:H2O (5:4:4:1)) lyophilized, then re-solubilized in D2O. The sample wasthen analyzed by 1H NMR spectroscopy with and without 31P decoupling, on a Bruker Avance 400172spectrometer with inverse broadband probe.Glc-β1-N31 µL of BglP (∼20 mg/mL), 1 µL DNPGlc (100 mM), 1 µL sodium azide (80 mM) and 2 µLbuffer (25 mM HEPES, pH 7.0, 50 mM NaCl) were combined and incubated for 10 min at roomtemperature. A no-enzyme control was also prepared, with 1 µL buffer in place of BglP. 0.5 µLof the reaction and the control were loaded on 2 separate TLC plates next to Glc-β1-N3 (10 mM),Glc (10 mM), DNPGlc (20 mM) and sodium azide (16 mM) standards. TLC plates were run witheither 1-butanol:methanol:NH4OH:H2O (5:4:2:1) or chloroform:methanol (5:1) mobile phases.β-UMPGlc and β-UDPGlc35 µL of BglP (glycerol removed) (∼20 mg/mL), 40 µL DNPGlc (100 mM), 40 µL UDP or UMP(100 mM) and 85 µL water were combined then incubated for 10 min at room temperature. BglPwas precipitated and removed by incubating for 5 min at 70 °C and centrifugation (60 s at 14,000x g). This sample was analyzed by TLC then preparative TLC was done to purify the productspot, however, MS analysis failed to produce an expected signal corresponding either UDP-Glc orUMP-Glc’s expected m/z.7.5 Chapter 4 Experimental7.5.1 Materials and ReagentsEnzyme quantification was performed using the Bradford method [296]. All thin layer chromatogra-phy (TLC) assays were done on silica gel 60 F254 TLC plates (EMD Millipore Corporation, Billerica,MA, USA). Plates were eluted with a mobile phase of 1-butanol, methanol, ammonium hydroxideand water in a ratio of 5:4:1:1, 5:4:4:1 or 5:2:4:1. Following elution, plates were dried then dippedin a molybdate TLC stain (2.5 % ammonium molybdate (w/v), 1 % ceric ammonium sulfate (w/v)and 10 % H2SO4 (v/v)) Visualization of TLC plates was done by heating until the product spotsbecame visible. PCR primers and cycle conditions can be found in Tables 7.4 and 7.5, and bufferrecipes can be found in Tables 7.6 and 7.7. Circos figure generated using RCircos [314, 315].173Table 7.4: PIPE PCR parameters used for insert and vector amplification. See Table 7.5 for an-nealing temperatures (X ) and extension times (Y ).Temperature (°C) Time95 2 min95 30 sX 30 s72 Y min15 holdTable 7.5: Partially incomplete polymerase extension (PIPE) PCR details.Construct Primers (5’→3’) Template AnnealingTemp (°C)ExtensionTime (min)pET28-GH94_A.h6V F: GAAATTGAATTCTATATGTCACACCATCACCATCACCAT pET28-HtCBP.h6 58 4R: CAAAATACCCATATCTCATGGTATATCTCCTTCTTAAAGTTAI F: AAGAAGGAGATATACCATGAGATATGGGTATTTTGATGAAG BVR_10H18 55 1.5R: GGTGATGGTGATGGTGTGACATATAGAATTCAATTTCTGpET28-GH94_B.h6V F: GGATCAAAGTGGAACTGATGCACCATCACCATCACCAT pET28-HtCBP.h6 58 4R: TATCCTGAAATTTTATATATTTCATGGTATATCTCCTTCTTAAAGI F: GGAGATATACCATGAAATATATAAAATTTCAGGATAAAAATGG BVR_10H18 60 1.5R: GGTGATGGTGATGGTGCATCAGTTCCACTTTGATCCpET28-GH94_C.h6V F: TCACGGTGATACTCCACGCGCACCATCACCATCACCATTG pET28-HtCBP.h6 62 4R: CAAACCCGATGTTGGTCATGGTATATCTCCTTCTTAAAGTTAAACI F: GGAGATATACCATGACCAACATCGGGTTTGCCGGG BVR_06I15 70 1.5R: CAATGGTGATGGTGATGGTGCGCGTGGAGTATCACCGTGATCpET28-GH94_D.h6V F: TCACGGTGATACTCCACGCGCACCATCACCATCACCATTG pET28-HtCBP.h6 62 4R: CAAACCCGATGTTGGTCATGGTATATCTCCTTCTTAAAGTTAAACI F: AGGAGATATACCATGCAGTATGGTTATTTTGACGAC BVR_06I15 55 1.5R: CAATGGTGATGGTGATGGTGCATTGTCAGAACGATCTCGTpET28-GH94_E.h6V F: ATGTTGTCGCTGAAATAGACCACCATCACCATCACCATTG pET28-HtCBP.h6 62 4R: CAAAAAAGCCAAATTTCATGGTATATCTCCTTCTTAAAGTTAAACI F: GGAGATATACCATGAAATTTGGCTTTTTTGATGAC BVR_32A05 55 1.5R: CAATGGTGATGGTGATGGTGGTCTATTTCAGCGACAACATpET28-GH94_F.h6V F: CCAGGTGGAAGCCTGGATACACCATCACCATCACCATTG pET28-HtCBP.h6 62 4R: CGAAATAACCGAAATTCATGGTATATCTCCTTCTTAAAGTTAAACI F: GGAGATATACCATGAATTTCGGTTATTTCGACGACG BVR_38M24 63 1.5R: CAATGGTGATGGTGATGGTGTATCCAGGCTTCCACCTGGpET28-GH94_G.h6V F: AAGTCCACGCCAGCCTTTACCACCATCACCATCACCATTG pET28-HtCBP.h6 62 4R: CAAAATGGCCGTATTTCATGGTATATCTCCTTCTTAAAGTTAAACAAAATTI F: GGAGATATACCATGAAATACGGCCATTTTGATGAT BVR_38M24 63 1.5R: CAATGGTGATGGTGATGGTGGTAAAGGCTGGCGTGGACTTpET28-GH149_H.h6V F: CTGTTTCGGGAATCGAACACCATCACCATCACCATTG pET28-HtCBP.h6 62 4R: GGGCGTGAAGTCGCGTTCATGGTATATCTCCTTCTTAAAGTTAAACI F: CTTTAAGAAGGAGATATACCATGAACGCGACTTCACGCCC BCR_17C21 67 3R: CAATGGTGATGGTGATGGTGTTCGATTCCCGAAACAGATACCTCC174Table 7.6: Molybdenum blue screen buffer recipes.Buffer Name Concentration CompoundLysisBuffer50 mM HEPES pH 7.01 mM EDTA0.5 % Triton X-1004 mM MgSO450 mM NaCl1 mg/mL LysozymeSubstrateBuffer20 mM MES pH 6.5200 mM Sodium Molybdate100 mM D-Glucose100 mM D-Cellobiose10 mM α-D-Glucose 1-phosphateDevelopmentSolution A0.25 % Ascorbic Acid0.7 N HClDevelopmentSolution B0.36 % Ascorbic Acid0.375 N HClDevelopmentSolution C0.24 % Ascorbic Acid0.25 N HClQuenchingSolution A2 % Acetic Acid100 mM Sodium CitrateQuenchingSolution B3 % Acetic Acid150 mM Sodium CitrateTable 7.7: Protein expression and purification buffers.Buffer Name Compounds Imidazole (mM)Loading Buffer 50 mM HEPES (pH 7.0),300 mM NaCl, 10 % glycerol,5 mM MgSO4, 0.1 mM DTT5Wash Buffer 25Elution Buffer 250Storage Buffer 01757.5.2 Metagenomic LibrariesTwo fosmid libraries were constructed from environmental genomic DNA sourced from beaver fecesand a passive mine tailings bioremediation system i.e. biochemical reactor system (BCR), respec-tively using the pCC1 copy control vector system in host strain Escherichia coli EPI300. The beaverfecal library was constructed as described by Armstrong et al. [255] and the BCR [256] library wasconstructed as described by Mewis et al. [176].7.5.3 Phosphatase InhibitionOvernight cultures of E. coli EPI300 carrying pCC1 were grown in LB media supplemented with100 µg/mL arabinose and 12.5 µg/mL chloramphenicol. 1.5 mL of cells were pelleted at 12 000 x gfor 1 min in a micro-centrifuge. The supernatant was removed and the cells were lysed by adding0.2 mL of lysis buffer (Table 7.6) and incubating at 37 °C for 1 h then centrifuged at 12 000 x gfor 10 min. For the molybdenum blue assay : 30 µL of lysate supernatant was transferred to 170µL of a substrate buffer (20 mM MES pH 6.5, 10 mM Glc-α1-P) containing 0, 10, 25, 50, 100 or200 mM sodium molybdate and incubated for 24 h at 37 °C. For each molybdate concentration, 50µL was transferred (in 3 replicates) to a 96-well plate. To normalize the molybdate concentrationin all reactions to 85 mM prior to colour development 50 µL of 170, 162, 149, 128, 85 or 0 mMsodium molybdate was added to the 0, 10, 25, 50, 100 and 200 mM substrate and lysate mixture,respectively. To induce the molybdenum blue reaction 100 µL of development solution B (Table7.6) was added to each well and the plate was incubated for 5 min at room temperature. The colourdevelopment was quenched by addition of 100 µL quenching solution B (Table 7.6) to each well.Absorbance at 655 nm was measured with a BioTek Synergy H1 Hybrid microtiter plate reader.For TLC assay : 7.5 µL of E. coli EPI300 lysate supernatant was incubated in a final volume of 50µL of 20 mM MES pH 6.5, 50 mM Glc-α1-P and 0, 10, 25, 50, 100 or 200 mM final concentrationof sodium molybdate then incubated for 24 h at 37 °C. To ensure consistent colorimetric responsesfrom all reactions, molybdate concentrations were normalized to 85 mM prior to loading on TLC,50 µL of 170, 162, 149, 128, 85 or 0 mM sodium molybdate was added to the 0, 10, 25, 50, 100 and200 mM TLC reactions, respectively. To ensure consistent Rfs of the Glc, Glc-α1-P and Glc6-P176standards, the 25 mM standard solutions were prepared containing 85 mM sodium molybdate. 0.5µL of each reaction was spotted on the TLC plate.7.5.4 Metagenomic Screen ValidationA 384-well control plate containing 3 replicates of four E. coli strains (EPI300 carrying pCC1,BL21(DE3) carrying empty pET28, BL21(DE3) carrying pET28-HtCBP.h6 and BL21(DE3) carry-ing pET28-HtCDP.h6) were grown in LB media (80 µL per well) supplemented with 100 µg/mLarabinose and 12.5 µg/mL chloramphenicol for the EPI300 cells and auto-induction media (80 µLper well) supplemented with 25 µg/mL kanamycin. The pET28-HtCBP.h6 and pET28-HtCDP.h6were constructed as described previously [91]. The control plate was then assayed following thescreening procedure outlined below.7.5.5 HT Metagenomic Screening ProcedureMetagenomic libraries were replicated as described by Mewis et al. [176] in 384-well plates (NUNC)containing 80 µL LB growth media + 100 µg/mL arabinose + 12.5 µg/mL chloramphenicol per well.Following incubation at 37 °C for 18 h, the replicated libraries were centrifuged (Beckmam CoulterAvanti® J-E Centrifuge) for 10 min at 4000 x g in a JS-5.3 swinging bucket rotor. Library plateswere transferred to a Beckman Coulter Biomek® FX robot outfitted with a 96 channel pipette headthat was used to remove the growth media. A single set of 96 x 250 µL MBP® BioRobotix™ Tipswas used to aspirate the 80 µL of growth media. A rinsing step in dH2O was included betweenplates to minimize contamination. To ensure the cell pellet was undisturbed the head depth was setto reach no deeper than 1 mm above the floor of the plates. To lyse the cells, 50 µL of lysis buffer(Table 7.6) was dispensed into each well using a Genetix QFill3 microplate filler then incubated for18 h at 37 °C in a humidity box. Plates were centrifuged (as described above) for 20 min at 4000 xg to pellet the insoluble cell debris following lysis.A Genetix Qpix2XT colony picking robot equipped with an automatic plate stacker was usedto transfer ∼3 µL of the lysate to a second 384-well plate containing 50 µL per well of substratebuffer. To transfer 3 µL of lysate to the substrate buffer a 96 honeycomb pin replicating headwas used. The surface tension of the lysate allowed for volume transfer by performing sequential177dips into the lysate and then substrate plates. The robot was programed to perform 5 sequentialtransfers from the lysate to the substrate buffer, followed by a sterilization protocol with 1 % bleachbefore proceeding to the subsequent set of wells. Each transfer consisted of a 2 second dip intothe source plate followed by 3 rapid successive dips into the destination plate. Following volumetransfer, the substrate plates were incubated for 48 h at 37 °C in humid conditions. After the 48h incubation period, 25 µL of development solution A (Table 7.6) was added to each well using aGenetix QFill3 microplate filler then incubated at room temperature for 30 min. 20 µL of quenchingsolution A (Table 7.6) was added to each well and the absorbance was measured at 655 nm witha BioTek Synergy H1 Hybrid microtiter plater reader. Positive clones (defined by having an A655value greater than mean + 4 SD) were selected and retested 4 times to confirm activity.For TLC validation, 19 positive clones and a background control of E. coli EPI300 carryingthe empty pCC1 fosmid background were grown in 8 mL of LB media supplemented with 100µg/mL arabinose and 12.5 µg/mL chloramphenicol for 18 h at 37 °C. Cultures were pelleted andresuspended in 200 µL lysis buffer (Table 7.6), incubated for 1 h at 37 °C then spun again at 12,000x g for 10 min in a micro-centrifuge. The soluble cell lysates (supernatants) were transferred to newmicrofuge tubes. Reverse phosphorolysis reactions were performed by combining 5 µL soluble celllysate with 25 µL 40 mM MES (pH 6.5) and 400 mM sodium molybdate, 5 µL water, 5 µL 200mM Glc-α1-P and 10 µL 250 mM glucose or cellobiose then incubated for 16 h at 37 °C. 0.25 µL ofeach reaction mixture was spotted on a TLC plate and run for 2 h in a mobile phase of 1-butanol,methanol, ammonium hydroxide and water in a ratio of 5:4:1:1.7.5.6 Fosmid SequencingFosmid DNA sequencing was done as described previously [91]. In brief, fosmid DNA was extractedusing a GeneJet plasmid miniprep kit (Thermo Fischer Scientific) then treated with PlasmidSafeDNase (Epicenter) before being individually barcoded and sequenced using the MiSeq system.7.5.7 Sequence AssemblyFosmid sequence reads were individually assembled using FabFos (https://github.com/hallamlab-/FabFos) as previously described [91]. In brief, 2 x 150bp reads were first aligned against a refer-178ence of vector backbone and E. coli sequences using BWA MEM v0.7.13 with default parameters,trimmed with Trimmomatic v0.35 and assembled using MEGAHIT v1.0.6 (k=71, 81, 91, 101, 111,121, 131, 141, 147). High sequencing coverage was accounted for by increasing the minimum k-mermultiplicity, based on an estimated coverage, to limit the number of erroneous k-mers introducedinto the assembly graph.7.5.8 ORF Prediction and Taxonomic AssignmentOpen reading frame prediction and gene identification was done as described previously [91]. In brief,Prodigal v2.6.3, as part of MetaPathways v2.5.1 [305], with default parameters was used to generateconceptually translated ORFs. These were then queried against the RefSeq protein database usingthe BLASTP [316] webtool hosted by the NCBI. LCAStar (or LCA*) was used to assign taxonomyby minimizing entropy changes to the taxonomic distribution i.e. lowest common ancestor for eachORF encoded on individual fosmid clones. The original LCA* codebase implemented by Hanson etal. [257] relied on files formatted specifically for the MetaPathways pipeline. A Python 3 compatibleversion (available at https://github.com/cmorganl/LCAStar release v0.1) was forked and changedto accept minimal inputs: a BLAST-output table, the NCBI taxonomic tree, and a tabular filemapping NCBI numerical taxonomic identifiers to organism name and rank. This was necessary toquery the most recent version without storing and indexing a large database.7.5.9 Sub-Cloning Metagenomic GPsSequencing of the positive fosmid hits from the functional screen identified eight unique GH94 andGH149 ORFs. Each of the eight were cloned into pET28a to generate expression constructs contain-ing a C-terminal histidine tag using the partially incomplete polymerase extension (PIPE) cloningmethod [298]. PCR parameters and primer sequences are listed in Tables 7.4 and 7.5. For all con-structs, heat shock transformations were performed by combining 2 µL of the vector PCR productwith 2 µL of the insert PCR product and then 100 µL of DH5α chemically competent cells [299].Constructs were confirmed by sequencing (Genewiz®, South Plainfield, NJ, USA) using T7 andT7term sequencing primers and subsequently transformed into BL21(DE3) chemically competentcells [299].1797.5.10 Heterologous Expression and PurificationA total of 1 L (GH94_A - GH94_G) or 2 L (GH149_H) auto-induction media [274] containing25 µg/mL kanamycin was inoculated with 1/100 of overnight culture of BL21(DE3) harboringpET28.GH94_A.h6 – pET28.GH94_G.h6 or pET28.GH149_H.h6 plasmids. Expression cultureswere grown at 30 °C for 24 h. Cells were harvested by centrifugation at 6000 x g for 6 min ina Beckman Coulter Avanti® J-E floor centrifuge (JA-10 rotor) followed by resuspension (20 mLper original culture volume) in loading buffer (50 mM HEPES pH 7.0, 300 mM NaCl, 10 % v/vglycerol, 5 mM MgSO4, 0.1 mM DTT, 5 mM imidazole). Cells were lysed using an Avestin C3homogenizer with an average cell pressure of 16,000 p.s.i. Soluble cell fractions were either storedat -20 °C until needed or immediately purified. Purification was carried out by immobilized metalaffinity chromatography on a GE Healthcare ÄKTA FPLC equipped with a UV and conductancedetector and an automatic fraction collector. Separate 5-mL HisTrap™ FF columns (GE Healthcare)were equilibrated with 10 column volumes (CV) of loading buffer. Soluble cell lysates were appliedto the columns using a P-1 peristaltic pump (GE Healthcare) followed by a wash step of 10 CVof loading buffer. The columns were then transferred to the ÄKTA and washed with 10 CV ofwash buffer (50 mM HEPES pH 7.0, 300 mM NaCl, 10 % v/v glycerol, 5 mM MgSO4, 0.1 mMDTT, 25 mM imidazole) followed by equilibration with 10 CV loading buffer. Proteins were elutedusing a 4 CV gradient (0-100 %) of loading buffer to elution buffer (50 mM HEPES pH 7.0, 300mM NaCl, 10 % v/v glycerol, 5 mM MgSO4, 0.1 mM DTT, 250 mM imidazole) with the fractioncollector set to collect 1 mL fractions. Fractions were analyzed by SDS PAGE, and those fractionsyielding the largest bands at (92 kDa for GH94_A, 104 kDa for GH94_B, 99 kDa for GH94_C,90 kDa for GH94_D, 94 kDa for GH94_E, 91 kDa for GH94_F, 92 kDa for GH94_G or 130 kDafor GH149_H) were combined individually and concentrated using an Amicon® Ultra-4 MWCO30-kDa centrifugal filter (Sigma), then dialyzed against storage buffer (50 mM HEPES pH 7.0, 300mM NaCl, 10 % v/v glycerol, 5 mM MgSO4, 0.1 mM DTT) and stored at -70 °C.1807.5.11 Functional CharacterizationFor forward phosphorolysis 4 µL of 0.5 mg/mL purified enzyme (diluted with 50 mM HEPES pH7.0, 300 mM NaCl, 2 % v/v glycerol, 5 mM MgSO4, 0.1 mM DTT) was incubated with 0.5 µLof 500 mM sodium phosphate (pH 7.0) and 0.5 µL 250 mM cellobiose, 100 mM cellotriose, 250mM laminaribiose or 100 mM laminaritriose then incubated for 1 h at room temperature. 0.25 µLof reaction volume was spotted on TLC and run for 30 minutes in a mobile phase of 1-butanol,NH4OH, MeOH and H2O in a 5:2:4:1 ratio. For reverse phosphorolysis 4 µL of 0.5 mg/mL purifiedenzyme was incubated with 0.5 µL 100 mM Glc-α1-P and 0.5 µL acceptor (250 mM glucose, 250mM cellobiose, 250 mM laminaribiose) then incubated for 1 h at room temperature. TLC performedas described for forward phosphorolysis. GH94_F was omitted because a sufficiently pure samplecould not be produced through heterologous expression.7.5.12 96-well Molybdenum Blue Activity AssayIn sterile 96-deep well plates (Axygen Scientific® P-DW-20-C) 1.5 mL of auto-induction media[274] containing 25 µg/mL kanamycin for pET28 vectors or 100 µg/mL carbenicillin for pET45 vec-tors was inoculated with 15 µL of overnight culture of BL21(DE3) harboring pET28.GH94_A.h6– pET28.GH94_G.h6, pET28.GH149_H.h6, pET28-HtCBP.h6, pET28-HtCDP.h6 or pET28 plas-mids; each in duplicate (Figure 4.8), or pET28-HtCBP.h6, malP-pET28, pET45-h6.GH13-1A06,pET45-h6.GH112-1B10 or pET45-h6.GH130-1E02 (Figure 4.11). The deep well plates were incu-bated for 18 h at 37 °C then centrifuged (Beckman Coulter Avanti® J-E Centrifuge) for 10 minat 4000 x g in a JS-5.3 swinging bucket rotor. The media was poured off and the cell pellets wereresuspended in 250 µL of lysis buffer (Table 7.6) then incubated for 3 h at 37 °C. The plates werecentrifuged again as described above to collect the insoluble cell debris. The supernatant containingthe soluble cell fraction was transferred to a new 96-well plate. For the plate containing the Figure4.8 clones, 250 µL of 1 % sodium dodecyl sulfate (SDS) was added to the insoluble cell debris pelletand incubated for 1 h at 37 °C. To test protein expression, 10 µL of each soluble cell fraction and theSDS solubilized cell debris were loaded on SDS PAGE. The soluble cell fractions from all clones wereused to test for activity using donor and acceptor combinations described below. 10 µL of soluble181cell lysate was transferred to a new 96-well plate containing 20 µL lysis buffer (Table 7.6) and 170µL of 20 mM MES pH 6.5, 200 mM sodium molybdate, 10 mM donor and 100 mM acceptor(s) orno acceptor (Glc-α1-P only) then incubated for 2 h at 37 °C. Donor/acceptor combinations for allclones from Figure 4.8 were either Glc-α1-P and glucose, Glc-α1-P and cellobiose or Glc-α1-P only.For the Figure 4.11 clones: pET28-HtCBP.h6 and pET45-h6.GH13-1A06: Glc-α1-P, glucose andcellobiose; malP-pET28: Glc-β1-P, glucose and maltose; pET45-h6.GH112-1B10: Gal-α1-P, glucoseand galactose or pET45-h6.GH130-1E02: Man-α1-P, glucose and mannose. To initiate molybde-num blue formation, 50 µL of the substrate lysate mixture was transferred to a new 96-well platefollowed by addition of 150 µL development solution C (Table 7.6). The colour development platewas incubated for 5 min at room temperature before the reaction was stopped by adding 100 µL ofquenching solution B (Table 7.6). Absorbance at 655 nm was measured with a BioTek Synergy H1Hybrid microtiter plate reader.7.6 Chapter 5 Experimental7.6.1 General MethodsEnzyme quantification was performed using the Bradford method [296]. TLC assays were performedon silica gel 60 F254 TLC plates (EMD Millipore Corporation, Billerica, MA, USA) in a mobile-phase of BuOH:MeOH:NH4OH:H2O 5:4:4:1 unless otherwise stated and stained with molybdateTLC stain (2.5 % ammonium molybdate (w/v), 1 % ceric ammonium sulfate (w/v) and 10 %H2SO4 (v/v)) or p-anisaldehyde TLC stain (92.5 % ethanol, 4 % H2SO4 (v/v), 1.5 % acetic acid(v/v), 2 % p-anisaldehyde (v/v)). Visualization of TLC plates was done by heating until the productspots became visible.7.6.2 JGI GP Library Expression Test and Preliminary Activity AssayThe expression and activity test for the GP master library was done as described above in sec-tion 7.5.12 for pET45 vectors. For the activity assay, donor/acceptor combinations are as follows:GH13_18 and GH94: 10 mM Glc-α1-P, 50 mM glucose, 50 mM cellobiose; GH65: 10 mM Glc-β1-P, 50 mM glucose, 50 mM maltose; GH112: 10 mM Gal-α1-P, 50 mM glucose, 50 mM galactose;182GH130: 10 mM Man-α1-P, 50 mM glucose, 50 mM mannose. At time points 3, 24 and 48 hmolybdenum blue reactions were performed.7.6.3 JGI GP Library Substrate Specificity ScreenThe GP library was replicated and the soluble cell fraction was prepared as described above insections 7.5.12 and 7.6.2. Substrate solutions were prepared that contained 20 mM MES pH 6.5,200 mM sodium molybdate, 10 mM donor (either Glc-α1-P, Glc-β1-P, Gal-α1-P or Man-α1-P) andone of the acceptors listed at the indicated concentration in Table 5.2. 7.5 µL of soluble cell lysateof each GP clone was transferred to x number of wells of a fresh 96-well plate (x = the numberof donor/acceptor combinations tested for that particular clone). 42.5 µL of each of the substratesolutions prepared according to Table 5.2 were added to each well so that each GP clone was mixedwith each substrate solution. The plates were then incubated at 37 °C for 3 h. Molybdenum blueformation was initiated by addition of 150 µL development solution C (Table 7.6). The plate wasthen incubated for 5 min at room temperature before the reaction was stopped by adding 100 µLof quenching solution B (Table 7.6). Absorbance at 655 nm was measured with a BioTek SynergyH1 Hybrid microtiter plate reader. The A655 readings were then visualized as a heat map using theComplexHeatmap package [317] in RStudio.7.6.4 JGI GP Library Sequence Similarity NetworksGP library SSNs: Sequences from each CAZy family (or subfamily in the case of GH13_18) wereobtained from the CAZy DB in 2020. These sets of sequences were uploaded to Enzyme Func-tion Initiative’s Enzyme Similarity Tool (EFI-EST) [181–184] to generate the Sequence SimilarityNetworks (SSNs). Sequence length restrictions were applied where noted to eliminate fragmentsbelow a certain length and/or multi-CAZy domain fusions above a certain length. The alignmentscore thresholds were set to achieve pairwise sequence identity cut offs of approximately 40 %, anddiffered between families. Sequences were grouped into representative nodes when they shared apercent identity of 95 or greater. The SSN graphics were produced in Cytoscape (v3.8.2) [186] andannotated in Adobe Illustrator.AchP GH94 SSN : The set of 3,077 GH94 sequences was retrieved from CAZy DB filtered for183identical sequences which reduced the set to 2,308 sequences. This set was uploaded to EFI-ESTto generate the SSN data set. A sequence length restriction was applied that eliminated fragmentsbelow 600 a.a. and multi-CAZy domain fusions above 1500 a.a., which reduced the set to 1,161sequences. The alignment score threshold was set to 10-200, translating to a pairwise sequenceidentity cut off of approximately 40 %. The resulting SSN contained 1,161 nodes, 76,904 edges andthe graphic was produce in Cytoscape (v3.8.2).7.6.5 Protein Expression and PurificationAchP (ABX81671.1): The DNA encoding the AchP gene was synthesized and inserted into thepET45b expression plasmid by the US Department of Energy Joint Genome Institute. 2 L of LBmedia containing 100 µg/mL carbenicillin was inoculated with 20 mL of overnight culture of E.coli BL21(DE3) harbouring the pET45-h6.AchP plasmid. The culture was grown for 3 h at 37 °C,IPTG was added to a final concentration of 0.5 mM, the temperature was reduced to 30 °C and theculture was grown for a further 18 h. Cells were harvested by centrifugation at 6000 x g for 10 minin a Beckman Coulter Avanti® J-E floor centrifuge (JA-10 rotor) followed by resuspension in 40 mLloading buffer (50 mM HEPES pH 7.0, 300 mM NaCl, 10 % v/v glycerol, 50 mM MgSO4, 0.1 DTT,5 mM imidazole). Cells were lysed using an Avestin C3 homogenizer with an average cell pressureof 16,000 psi. The soluble cell lysate fraction was isolated by centrifuging the crude cell lysate at15,000 rpm for 30 min (JA-20 rotor). AchP purification was carried out by immobilized metal affinitychromatography on a GH Healthcare AKTA FPLC equipped with a UV and conductance detector,an automatic fraction collector and two inlet pumps. Pump A was equilibrated with loading bufferpump B with elution buffer (50 mM HEPES pH 7.0, 300 mM NaCl, 10 % v/v glycerol, 50 mMMgSO4, 0.1 mM DTT, 250 mM imidazole). A 5-mL HisTrap™ FF column (GE Healthcare) wereequilibrated with 10 column volumes (CV) of loading buffer. The soluble cell lysate was applied tothe column using a P-1 peristaltic pump (GE Healthcare) followed by a wash step of 10 CV loadingbuffer. The HisTrap™ column was transferred to the AKTA and washed with 10 CV of 8.2 % pump B(25 mM imidazole). AchP was eluted using a 4 CV gradient (8.2-100 %) of loading buffer to elutionbuffer with the fraction collector set to collect 1 mL fractions. Fractions were analyzed by SDSPAGE and the fractions containing the largest bands at 98 kDa were combined and concentrated184using Amicon® Ultra-4 MWCO 30-kDa centrifugal filter (Sigma). The concentrated protein wasdiluted with storage buffer (50 mM HEPES pH 7.0, 300 mM NaCl, 10 % v/v glycerol, 50 mMMgSO4, 0.1 DTT) in multiple cycles until the imidazole concentration was approximately 1 mM.Final AchP concentration was 12 mg/mL (48 mg total yield) and stored at -70 °C.NahK [281, 318]: NahK was expressed and purified as described above for AchP, with thefollowing modifications. Expression culture was 3 L. Fraction concentration was done with a 10kDa centrifugal filter. Final NahK concentration was 6 mg/mL (12 mg total yield).AchP (for crystallography): The gene was expressed from pET45b using E. coli BL21 (DE3)cells in Terrific Broth media with 0.5mM IPTG used for induction. The seleno-methionine proteinwas expressed using the same E. coli strain in PASM-5052 media [274]. Expressions were at 0.5L scale in 2 L baffled flasks for 2 days at 20 °C and 180 rpm. At harvest the cells were pelletedand stored at -80 °C until thawed for protein purification. The protein was purified using the sameprotocol whether it was native or seleno-methionine. Cell pellets were removed from the -80 °Cfreezer and re-suspended by stirring with a stir bar in 25 mM HEPES, pH 7.4, 100 mM NaCl, 1mM DTT (base buffer) + 0.1 mg/ml lysozyme, 3 ug/ml DNaseI, 1x Novagen EDTA free proteaseinhibitor. The cells were lysed by multiple passes in the Emulsiflex C3 emulsifier. The lysate wasclarified by centrifugation at 40,000 x g for 40 min. The clarified lysate had imidazole added to30 mM prior to loading onto a 5 mL Histrap column on the AKTA purifier HPLC instrument.The HisTrap column was equilibrated with base buffer prior to loading of the lysate. The boundprotein was eluted with a 0-45 % B gradient in 20 CV. Buffer B was the same as base buffer withthe addition of 0.5 M imidazole. The elution peak was analyzed by SDS-PAGE and the cleanestfractions were pooled prior to dialysis against 1 L of 25 mM HEPES, pH 7.4, 25 mM NaCl, 1 mMDTT for 2 x 1 hour at 4 °C. At the end of dialysis the protein showed no precipitation and wasloaded onto a 5 mL Q ion exchange column on the FPLC. The bound protein was eluted with a0-45 % B gradient in 24 CV. Buffer B was the same as the dialysis buffer but NaCl was 1 M. Theelution fractions were analyzed by SDS-PAGE and the cleanest fractions pooled, concentrated andinjected onto a 10 x 300 Superdex size exclusion column (SEC). The column was pre-equilibratedwith base buffer prior to injecting the protein. The SEC column was run at 0.5 mL/min and theinjection volumes were ∼1 mL. The cleanest fractions, as determined by SDS-PAGE analysis, were185pooled. The protein was concentrated with a 50 kDa centrifugal concentrator to 10-16.5 mg/mLprior to being used for crystallization trials.7.6.6 Cell Lysate Substrate Specificity Assay1.5 mL of auto-induction media [274] containing 100 µg/mL carbenicillin was inoculated with 15 µLof overnight culture (LB media) of E. coli BL21(DE3) harboring the pET45-h6.AchP plasmid. Theculture was incubated for 18 h at 37 °C then transferred to a 1.5 mL microfuge tube. The culturewas centrifuged, media poured off and the cell pellets were resuspended in 250 µL of lysis buffer(50 mM HEPES pH 7.0, 1 mM EDTA, 0.5 % Triton X-100, 4 mM MgSO4, 50 mM NaCl, and 1mg/mL lysozyme). The cell lysis mixture was incubated for 3 h at 37 °C then centrifuged again tocollect the insoluble cell debris. The supernatant containing the soluble cell lysate was transferredto a new 1.5 mL microfuge tube. 10 µL of soluble cell lysate was transferred to a 96-well (Costar®96-well flat-bottom polystyrene)plate containing 20 µL lysis buffer and 170 µL substrate solution(20 mM MES pH 6.5, 200 mM sodium molybdate, 10 mM donor (Glc1-P or GlcNAc1-P) andacceptor (50 mM glucose (Glc), 50 mM glucosamine (GlcN), 20 mM chitobiose (GlcN-β1,4-GlcN),50 mM N-acetylglucosamine (GlcNAc), 10 mM N,N’ -diacetylchitobiose (GlcNAc-β1,4-GlcNAc), 50mM D-galactose, 50 mM D-mannose (Man), 50 mM D-xylose (Xyl), 50 mM D-fructose (Fru), 50mM L-rhamnose (Rha), 20 mM gentiobiose (Gen), 50 mM maltose (Mal), 20 mM cellobiose (Cel),10 mM laminaribiose (Lam), 5 mM sophorose (Sop) or 20 mM β-lactose (bLac)) then incubated for3 h at 37 °C. To initiate molybdenum blue formation, 50 µL of the substrate lysate mixture wastransferred to a new 96-well colour development plate followed by addition of 150 µL developmentsolution (0.24 % ascorbic acid and 0.25 N HCl). The color development plate was incubated for 5min at room temperature before the reaction was stopped by adding 100 µL of quenching solution(3 % acetic acid and 3 % citrate). Absorbance at 655 nm was measured with a BioTek Synergy H1Hybrid microtiter plate reader.7.6.7 Purified AchP Substrate Specificity AssayThirty-six donor and acceptor combinations were done with purified AchP in a 25 µL reactionvolume. 10 µL buffer (200 mM HEPES pH 7.0, 200 mM NaCl, 10 mM MgSO4, 400 mM sodium186molybdate) was combined with 5 µL water, 2.5 µL donor (100 mM Glc1-P, 100 mM GlcNAc1-Por 100 mM GalNAc1-P) and 2.5 acceptor (500 mM Glc, 400 mM GlcN, 500 mM GlcNAc, 100 mMGlcNAc-β1-pNP, 500 mM Gal, 500 mM GalN, 500 mM GalNAc, 100 mM GalNAc-α1-pNP, 100 mMGlc-β1,3-Glc, 100 mM GlcNAc-β1,4-GlcNAc, 500 mM ManNAc or no acceptor) in a 200 µL PCRtube. 5 µL of purified AchP (0.2 mg/mL) was added to each reaction which were then incubatedfor 3 h at room temperature. Each 25 µL reaction was transferred to a sperate well of a 96-wellplate. 75 µL of development solution was added to each well then incubated for 5 min at roomtemperature before the reaction was stopped by adding 50 µL of quenching solution. Absorbanceat 655 nm was measured with a BioTek Synergy H1 Hybrid microtiter plate reader.7.6.8 AchP Kinetics AnalysisKinetic parameters for AchP reverse phosphorolysis were determined using the phosphate releasemethod described previously [116]. In brief, phosphate release was coupled to the formation ofmolybdenum blue, which can be quantified by measuring absorbance at 655 nm. Phosphate con-centration was determined using a standard curve ranging between 0 and 10 mM phosphate. Kineticparameters for AchP were determined with four donor and acceptor combinations: (A) Glc1-P andGlcNAc, (B) Glc1-P and GalNAc, (C) GlcNAc1-P and GlcNAc, and (D) GlcNAc1-P and GalNAc.Donor concentration was held constant at 10 mM while the concentrations of the acceptors werevaried (as described below). Reactions were initiated by adding 5 µL of 0.25 mg/mL AchP to 10 µL2x buffer (200 mM HEPES pH 7.0, 200 mM NaCl, 10 mM MgSO4 and 400 mM sodium molybdate),2.5 µL of 100 mM donor, 2.5 µL of 10x acceptor and 5 µL water. Reactions were stopped at thetimes (t) indicated below by boiling for 5 min, then 20 µL from each reaction was transferred toa 96-well plate. Molybdenum blue formation was initiated by adding 90 µL of color developmentsolution (0.1 N HCl and 0.24 % sodium ascorbate) then incubated for 30 min at room temperature.The color development reaction was stopped by adding 90 µL of quenching solution (68 mM sodiumcitrate and 2 % acetic acid). Absorbance at 655 nm was measured with a BioTek Synergy H1 Hybridmicrotiter plate reader. All reactions were performed in triplicate and acceptor concentrations werechosen to encompass apparent Km value where possible. (A) 10 mM Glc1-P; 0, 1, 5, 10, 25, 50,100, 200 and 300 mM GlcNAc; t = 2 min 45 s. (B) 10 mM Glc1-P; 0, 5, 25, 50, 100, 150, 200,187300 and 380 mM GalNAc; t = 5 min. (C) 10 mM GlcNAc1-P; 0, 0.1, 0.25, 0.5, 1, 2, 5, 10, 25 and50 mM GlcNAc; t = 3 min. (D) 10 mM GlcNAc1-P; 0, 5, 10, 25, 50, 100, 150, 200 and 300 mMGalNAc; t = 3 min. Non-linear regression was performed using GraphPad Prism version 6.0. For(A) and (C) data were fit using the Michaelis-Menten equation incorporating substrate inhibition(Equation 7.2). For (B) and (D) data were fit using the standard Michaelis-Menten equation:v0 =Vmax · [S]Km + [S](7.9)7.6.9 Oligomerization analysisAcholetin oligomerization was assayed with 10 mM Glc1-P, GlcNAc1-P or GalNAc1-P as donorsand GlcNAc or GalNAc as acceptor with donor/acceptor ratios of 1:10, 1:1 and 100:1. 0.5 µL of100 mM donor was combined with 1 µL 5x acceptor (500 mM, 50 mM or 0.5 mM), 2.5 µL reactionbuffer A (100 mM HEPES pH 7.0, 100 mM NaCl, 5 mM MgSO4) and 0.5 µL water. Reactionswere initiated by adding 0.5 µL AchP (6 mg/mL) then incubating at room temperature for 3 hfor those containing Glc1-P and GalNAc1-P and 30 min for those with GlcNAc1-P. Reactions werestopped by diluting 50x with super DHB MALDI matrix (Fluka) then analyzed by MALDI-TOFMS (Bruker Autoflex).7.6.10 Synthesis of GlcNAc-β1,3-GlcNAcSynthesis performed by Dr. Gregor Tegl. GlcNAc1-P and GlcNAc were dissolved in buffer (HEPES,pH 7.0), the enzyme AchP was added and the reaction mixture was incubated at 37 °C. Final reac-tion conditions: Glc1P (15 mg/mL) and GlcNAc (40 mg/mL), AchP (0.05 mg/mL), 50 mM HEPES.Reaction progress was monitored by TLC (BuOH:MeOH:NH4OH:H2O 5:4:4:2, p-anisaldehyde stain-ing). After GlcNAc1-P was fully consumed, the enzyme was removed from the reaction mixtureby ultrafiltration (MWCO 10 kDa, Satorius). The product was isolated by gel filtration chro-matography (eluent: H2O; Bio-gel P2, BioRad), fractions containing pure product were pooled andlyophilized. The pure dimer was characterized by MALDI-TOF MS (Bruker Autoflex) and NMR(Bruker AV-400 MHz spectrometer; solvent: D2O). NMR assignments in section C.1.1887.6.11 Synthesis of Glc-β1,3-GlcNAc-pNPSynthesis performed by Dr. Gregor Tegl. Glc1-P and GlcNAc were dissolved in buffer (HEPES, pH7.0), AchP was added and the reaction mixture was incubated at 37 °C. Final reaction conditions:15 mg/mL Glc1-P and 40 mg/mL GlcNAc, 0.05 mg/mL AchP, 50 mM HEPES. Reaction progresswas monitored by TLC (BuOH:MeOH:NH4OH:H2O 5:4:4:2, p-anisaldehyde staining). After Glc1-Pwas fully consumed, the enzyme was removed from the reaction mixture by ultrafiltration (MWCO10 kDa, Satorius) and product was isolated by gel filtration chromatography (eluent: H2O; Bio-gelP2, BioRad). The fraction containing pure product was lyophilized and characterized by MALDI-TOF MS (Bruker Autoflex) and NMR (Bruker AV-400 MHz spectrometer; solvent: D2O). NMRassignments in section C.1.7.6.12 Core 3 Mucin-like O-glycan Analog SynthesisAchP was used to synthesize T-antigen core 3 analogs, GlcNAc-β1,3-GalNAc-MU and GlcNAc-β1,3-GalNAc-pNP using GlcNAc1-P as donor and either 4-methylumbelliferyl N -acetyl-α-D-galactosamine(GalNAc-MU) or 4-nitrophenyl N -acetyl-α-D-galactosaminide (GalNAc-pNP) as acceptor. ForGlcNAc-β1,3-GalNAc-MU, 2 µL of 100 mM GlcNAc1-P (Carbosynth) was combined with 1 µL 100mM GalNAc-MU (prepared by Dr. Hongming Chen), 15 µL reaction buffer A (100 mM HEPES pH7.0, 100 mM NaCl, 5 mM MgSO4). For GlcNAc-β1,3-GalNAc-pNP, 5 µL of 100 mM GlcNAc1-Pwas combined with 2.5 µL 100 mM GalNAc-pNP (prepared by Dr. Hongming Chen), 37.5 µLreaction buffer A. Reactions were initiated by addition of 2 µL (MU reactions) or 5 µL (pNP re-actions) of AchP (6 mg/mL) and incubated at room temperature for 1 h. Reaction progress wasmonitored by TLC with a mobile phase of EtOAc:MeOH:H2O 7:2:1 and visualized with molybdateTLC stain or under UV light and analyzed by mass spectrometry. Both GlcNAc-β1,3-GalNAc-MUand GlcNAc-β1,3-GalNAc-pNP were assayed as potential substrates for the core 3 T-antigen cleav-ing enzyme endo-α-N -acetylgalactosaminidase from Streptococcus pneumoniae R6 (EngSP)47,57.For GlcNAc-β1,3-GalNAc-MU, in a 96-well plate 2 µL of the AchP reaction was combined with48 µL reaction buffer A and initiated by adding 0.4 µL spGH101 (10 mg/mL). Release of MUwas monitored continuously in a BioTek Synergy H1 Hybrid microtiter plate reader (Ex. 365 nm,189Em. 440 nm). For GlcNAc-β1,3-GalNAc-pNP, in a 96-well plate 10 µL of the AchP reaction wascombined with 39 µL reaction buffer A and reaction initiated by adding 1 µL EngSP (10 mg/mL).Release of pNP was monitored continuously in a BioTek Synergy H1 Hybrid microtiter plate reader(absorbance 400 nm).7.6.13 Acholetin Two-pot Large Scale SynthesisGlcNAc1-P synthesis: In 60 mL, 1 g GlcNAc, 3 g ATP were dissolved in Tris, pH 7.0 buffer withMgCl2. NahK was added and the reaction was incubated at 37°C for 18 h. Final reaction conditions:GlcNAc (75 mM), ATP (100 mM), MgCl2 (10 mM), NahK (0.08 mg/mL) and 967 mM Tris, pH7.0. ATP, ADP, AMP and free phosphate was precipitated by the addition of barium acetate.6.7 mL of 1 M barium acetate was added, the slurry was centrifuged, then the supernatant wastransferred to clean 50 mL Falcon tubes. This barium acetate precipitation process was repeatedtwice more. GlcNAc1-P was precipitated from the supernatant by adding 267 mL of ethanol followedby centrifugation. The pellet was then washed with 333 mL acetone then dried under vacuum for 18h. Total mass of the dried pellet was 2.66 g, which is composed primarily of GlcNAc1-P (maximumtheoretical yield was 1.36 g) and barium acetate.Acholetin synthesis: In 10 mL, 2.66 g of the GlcNAc1-P/barium acetate product was dissolvedin 10 mM HEPES, pH 7.0 buffer with 0.4 mM GlcNAc. The donor/acceptor ratio was set roughlyto 1000:1 to maximize the degree of polymerization (DP) of the acholetin product. AchP wasadded to a final concentration of 0.6 mg/mL to initiate the reaction. After addition of AchP, thereaction began to turn cloudy as the phosphate released from reverse phosphorolysis formed aninsoluble barium salt. The reaction was incubated at room temperature for 48 h. The precipitatedbarium phosphate salt was removed by centrifugation, the supernatant was then incubated at 70 °Cfor 5 minutes to precipitate the AchP, which was also removed by centrifugation. The product wasdesalted on either a Sephadex G-25 column with a bed height of 33 cm or a PD-10 Desalting Column(GE Healthcare). The 33 cm column was attached to the AKTA (same as above) and equilibratedwith degassed water. 4.4 mL of the acholetin mixture was injected onto the column with a 5 mL/minflowrate and the fraction collector set to collect 2 mL fractions after 20 mL post injection. Fractions4-13 were individually lyophilized. The combined mass was 151.5 mg which equates to a 344.3190mg yield from the whole 10 mL reaction. NMR analysis was performed on fraction 4, the fractioncontaining the highest proportion of high molecular weight acholetin oligomers. Fraction 4 wasdissolved in D2O and subjected to 1H, H-H COSY, HSQC and HMBC NMR analyses. The samesample was also reduced by NaBH4 (followed by quenching with acid, 4x treatment with MeOHand removal of solvent) and subjected to the same NMR analyses to identify the reducing endGlcNAc unit. A second desalting preparation was done by applying 1 mL of the acholetin mixtureto a PD-10 Desalting Column according to the manufacturer’s instructions with water used as themobile phase. The desalted 3.5 mL elution fraction was lyophilized then resuspended in water toa final concentration of 50 mg/mL. This material was analyzed by Multi-Angle Light Scattering(Supplement Figure C.9). NMR assignments in section C.1.191Bibliography1. 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Microbiol. 73, 6444–6449 (2007).225Appendix AChapter 2 Supplemental MaterialA.1 General InformationNMR spectra were recorded on a Bruker Avance 400 MHz in D2O and referenced externally todioxane in D2O (1H: 3.75 ppm, 13C:67.4 ppm) or H3PO4 in D2O (31P: 0 ppm). High Res massspectra were obtained on a Micromass LCT time-of-flight (TOF) mass spectrometer equipped withan electrospray ion source.A.2 NMR Assignments of Nag3 ProductsSection A.2 written by Dr. Markus Blaukopf.GlcNAc1-P : 1H (400 MHz, D2O) δ 4.93 (app t, 1 H, J 1,P =J 1,2 =8.3 Hz, H-1), 3.91 (dd, 1 H,J 6a,6b =12.3 Hz, J 6a,5 =2.1 Hz, H-6a), 3.72 (dd, 1 H, J 2,1 =8.3 Hz, J 2,3 =10.4 Hz, H-2), 3.69 (dd,1 H, J 6b,6a =12.3 Hz, J 6b,6a =6.6 Hz, H-6b), 3.54 (dd, 1 H, J 3,2 =10.4 Hz, J 3,4 =8.8 Hz, H-3), 3.49(ddd, 1 H, J 5,4 =9.8 Hz, J 5,6a =2.1 Hz, J 5,6b =6.7 Hz, H-6b), 3.38 (dd, 1 H, J 4,3 =9.2 Hz, J 4,5=9.5 Hz, H-4), 2.05 (s, 3 H, NHAc). 13C (100 MHz, D2O), δ 175.88 (NHAc), 96.48 (d, J 1,P = 5.1Hz, C-1), 77.14 (C-5), 74.83 (C-3), 70.91 (C-4), 61.96 (C-6), 57.32 (d, J 2,P =7.7 Hz, C-2), 23.20(NHAc). 31P:(161.9 MHz, D2O) δ 0.88 HRMS (m/z ): TOF-MS ES [MH]- calc for C8H15NO9P,300.0484 found: 300.0486.Glc1-P : 1H (400 MHz, D2O): δ 4.83 (app t, J 1,P =J 1,2 = 7.7 Hz, H-1), 3.85 (dd, 1 H, J 6a,6b =12.3Hz, J 6a,5 =2.1 Hz, H-6a), 3.61 (dd, 1 H, J 6b,6a =12.3 Hz, J 6b,6a =6.7 Hz, H-6b), 3.46 (t, 1 H, J 3,2=J 3,4 =9.2 Hz, H-3), 3.43 (m, 1 H, H-5), 3.28 (t, 1 H, J 4,3 =J 4,5 =9.4 Hz, H-4), 3.24 (dd, 1 H,J 2,3 =9.3 Hz, J 2,1 =7.9 Hz, H-2). 13C (100 MHz, D2O): δ 97.92 (d, J 1,P = 5 Hz, C-1), 77.09 (C-5),76.18 (C-3), 74.93 (d, J 2,P = 7.0 Hz, C-2), 70.56 (C-4), 61.81 (C-6). 31P: (161.9 MHz, D2O): δ 1.35.226HRMS (m/z ): TOF-MS ES [MH]- calc for C6H12NO9P, 259.0219 found: 259.0216.A.3 Supplemental Figures3.23.43.63.84.04.24.44.64.8 ppm3 .2 43 .2 63 .2 63 .2 83 .3 13 .4 13 .4 23 .4 33 .4 43 .4 43 .4 63 .4 73 .4 93 .5 03 .5 63 .5 73 .5 93 .6 13 .6 23 .6 43 .6 63 .8 33 .8 43 .8 63 .8 74 .0 14 .7 74 .8 24 .8 34 .8 51 .9 52 .0 91 .0 30 .4 11 .0 00 .6 67 6 5 4 3 2 1 ppm126a 6b53 4Glucose-beta-1-phosphateFigure A.1: 1H NMR spectrum of Glc-β1-P in D2O.2276065707580859095100 ppm6 1. 817 0. 567 4. 907 4. 977 6. 197 7. 099 7. 909 7. 9513 456Glucose-beta-1-phosphate2Figure A.2: 13C NMR spectrum of Glc-β1-P in D2O.2282.53.03.54.04.55.0 ppm2 .0 53 .3 63 .3 83 .4 03 .4 73 .4 83 .4 93 .4 93 .5 03 .5 13 .5 23 .5 23 .5 53 .5 53 .5 73 .6 73 .6 93 .7 03 .7 03 .7 23 .7 23 .7 33 .7 53 .9 03 .9 03 .9 33 .9 44 .9 24 .9 44 .9 62 .5 51 .0 32 .0 52 .1 91 .0 00 .7 77 6 5 4 3 2 1 ppmGlcNAc-beta-1-phosphateFigure A.3: 1H NMR spectrum of GlcNAc-β1-P in D2O.229180 160 140 120 100 80 60 40 20 ppm2 3. 205 7. 295 7. 366 1. 967 0. 917 4. 837 7. 149 6. 469 6. 511 75 .8 813456GlcNAc-beta-1-phosphateNHAcNHAc2Figure A.4: 13C NMR spectrum of GlcNAc-β1-P in D2O.230Appendix BChapter 3 Supplemental Material231Table B.1: BglP crystallographic statistic table.Crystal BglP BglP-2FGlc(PDB ID: 5VQD) (PDB ID: 5VQE)X-ray source Rigaku R-AXIS IV++ Rigaku R-AXIS IV++Crystal geometrySpace group C2221 C2221Unit cell (Å) a=80.52 b=84.15 c=161.19; a=80.39 b=83.95 c=161.27;α=β=γ=90° α=β=γ=90°Crystallographic dataWavelength (Å) 1.5418 1.5418Resolution range (Å) 54.72-2.10 (2.16-2.10)* 33.12-1.89 (1.96-1.89)Total observations 310027 (27157) 292494 (16456)Unique reflections 38928 (2647) 43229 (2693)Multiplicity 10.1 (10.3) 6.8 (6.1)Completeness (%) 94.6 (99.9) 98.4 (96.1)Rmerge 0.112 (0.590) 0.073 (0.235)CC1/2 0.99 (0.95) 0.99 (0.96)I/σI 15.7 (4.0) 15.0 (6.9)Wilson B-factor (Å2) 13.89 16.79Refinement statisticsReflections in test set 1964 (208) 1996 (196)Non-hydrogen atoms:Protein atoms 4305 43342FGlc atoms - 11Water 468 366Glycerol 12 -Rwork/Rfree 0.21/0.26 0.17/0.21Root mean square deviationBond lengths/angles (Å/°) 0.002/0.57 0.005/0.81Ramachandran plotFavoured/allowed/outliers (%) 95.45/3.45/1.09 96.03/3.25/0.72Average B factor (Å2) 16.30 20.31Macromolecules 15.88 19.77Ligands 23.41 13.04Solvent 19.92 26.94*Values in parentheses refer to the highest resolution shell232Table B.2: CAZy ORFs found on the 12 fosmids displaying phosphate dependent turnover ofDNPGlc. All fosmids, except for 29K06 and 31P01, were previously identified in functionalmetagenomic screens of the BCR library targeting glycoside hydrolase activity. CMUC, 6-chloro-methylumbelliferyl cellobioside; DNPC, 2,4-dinitrophenyl cellobioside; NP, not published. Accessionnumbers indicate top BLAST hit for each ORF. The top hit for BglP (AEI40672.1) is a predicitedβ-N-acetylglucosaminidase from Paenibacillus mucilaginosus (44 % sequence similarity and 64 %sequence identity). A graphical representation of these 12 fosmids can be found in SupportingFigure B.1.FosmidCloneProduceGlc1-P?OriginalSubstrateReference(Fosmid Clone ID)CAZyORFsTop BLASTAnnotation(Accession)FOS62_03A16 No DNPC Mewis et al. 2013 (D1) GH16 AEW47952.1GH3 AEW47953.1FOS62_10P15 No DNPC NP GH3 AFU63317.1FOS62_15C24 No DNPC Mewis et al. 2013 (E2) GH16 AEW47952.1GH3 AEW47953.1FOS62_15N09 No DNPC Mewis et al. 2013 (F2)GH1 AEW47962.1GT28 ACD83006.1CE9 ACD04780.1CE9 ACD04780.1FOS62_19A10 No DNPC NP -FOS62_26C23 No CMUC NP GH1 AEV28543.1FOS62_29K06 Yes DNPGlc Current Study GH3 AEI40672.1FOS62_31P01 Yes DNPGlc Current Study GH3 AEI40672.1FOS62_38A06 No CMUC NPGH3 AEW47953.1GH16 AEW47952.1GH16 AEW47952.1FOS62_41D24 No CMUC NP GH3 AHF25061.1FOS62_48I14 No DNPC NP GH3 AHF25061.1FOS62_48J10 No DNPC NP GH3 AHF25061.1233Table B.3: 29K06 and 31P01 annotation table. ORF # map to Figure 3.7. ORFs 1-32 were foundon 31P01 and 11-41 were found on 29K06. BLASTP annotations represent annotations of top hit.bglP ORF in bold.ORF # BLASTP Functional Annotations1 hypothetical2 NTP phosphatase3 hypothetical4 phosphoesterase [RecJ]5 endopeptidase [Clp] ATP-binding regulatory subunit [ClpX]6 protease proteolytic subunit [ClpP]7 trigger factor8 cupin9 cytosine deaminase-like metal-dependent hydrolase10 MATE efflux family protein11 MATE efflux family protein12 phosphate-binding protein13 phosphate ABC transporter, permease [PstC]14 phosphate ABC transporter, permease [PstC]15 amino-acid transporter subunit ATP-binding component ABC superfamily16 PhoU family transcriptiona lregulator17 hypothetical18 ArsR family transcriptional regulator19 putative membrane protein20 xylose isomerase domain-containing protein21 N -acetylneuraminate lyase22 oxidoreductase, short chain dehydrogenase/reductase family protein23 cation diffusion facilitator family transporter24 hypothetical25 hypothetical26 hypothetical27 putative sugar ABC transporter sugar-binding protein28 binding-protein-dependent transport systems inner membrane component29 binding-protein-dependent transport systems inner membrane protein30 (bglP) Beta-hexosamidase A, glycoside hydrolase family 3 [GH3]31 beta-phosphoglucomutase, partial32 hypothetical33 hypothetical34 hypothetical35 hypothetical36 Ferriciron ABC transporter, permeaseprotein37 fe(3+) ions import ATP-binding protein [FbpC]38 hypothetical39 kDPGKHG aldolase40 2-dehydro-3-deoxy gluconatekinase41 transcriptional regulator, IclR family protein234Figure B.1: Graphical representation of CAZy ORFs from the 12 fosmids displaying phosphatedependent turnover of DNPGlc. 29K06 and 31P01 each contain the BglP ORF. Fosmids containingsimilar ORF configurations are grouped and CAZy ORFs are colour coded. Further details can befound in Supplemental Table B.2.235Appendix CChapter 5 Supplemental MaterialC.1 NMR Assignments of AchP ProductsSection C.1 written by Dr. Feng Liu.GlcNAc-β1,3-GlcNAc1H NMR (400 MHz, D2O) σ 5.11 (d, J1α,2α = 3.4 Hz, 1H, H-1α), 4.66 (d, J1β,2β = 8.3 Hz, 1H,H-1β), 4.58 (d, J1’(β),2’(β) = 8.3 Hz, 1H, H-1’(β)), 4.57 (d, J1’(α),2’(α) = 8.2 Hz, 1H, H-1’(α)), 3.95(ddd, J1α,2α = 3.4, J2,NH = 10.5, J2,2 = 10.5 Hz, 1H, H-2α), 3.96-3.86 (m, 4H, H-6 αnd H-6’),3.93-3.87 (m, 1H, H-3α), 3.91-3.88 (m, 1H, H-2β), 3.91-3.85 (m, 1H, H-5α), 3.76-3.65 (m, 3H, H-2α,H-2’(α) αnd H-2’(β)), 3.75-3.72 (m, 1H, H-3β), 3.62-3.54 (m, 1H, H-3’), 3.59-3.50 (m, 1H, H-4α αndH-4β), 3.51-3.43 (m, 3H, H-5β, H-5’(α) αnd H-5’(β)), 3.50-3.44 (m, 1H, H-4’), 2.07 (s, 3H, CH3),2.07 (s, 3H, CH3), 2.02 (s, 3H, CH3), 2.01 (s, 3H, CH3).13C NMR (101 MHz, D2O) σ 174.43 (C=O), 174.40 (C=O), 174.00 (C=O), 173.72 (C=O),101.13 (C-1’(β)), 101.10 (C-1’(α)), 95.00 (C-1β), 90.87 (C-1α), 81.41(C-3β), 78.86(C-3α), 75.76(C-5’(β)), 75.69 (C-5’(α)), 75.41 (C-5β), 73.26 (C-3’(α)), 73.24 (C-3’(β)), 71.09 (C-5α), 69.73 (C-4’), 68.50(C-4α), 68.48(C-4β), 60.74 (C-6β), 60.59 (C-6α), 60.53 (C-6’), 55.70 (C-2’(β)), 55.67 (C-2’(α)), 55.57(C-2β), 52.97 (C-2α), 22.30 (CH3), 22.24 (CH3), 22.06 (CH3).β1,3 linkage was determined by HMBC H-1’, C-3.Glc-β1,3-GlcNAc-β1-pNP1H NMR (400 MHz, D2O) σ 8.27 (d, J = 9.3 Hz, 1H), 7.21 (d, J = 9.3 Hz, 1H), 5.36 (d, J1,2 = 8.5Hz, 1H, H-1), 4.54 (d, J1’,2’ = 7.8 Hz, 1H, H-1’), 4.19 (dd, J1,2 = 8.5 Hz, J2,3 = 10.4 Hz, 1H, H-2),3.97 (dd, J5,6 = 2.2 Hz, J6a,6b = 12.6 Hz, 1H, H-6a), 3.93 (dd, J2,3 = 10.4 Hz, J3,4 = 8.4 Hz, 1H,236H-3), 3.92 (dd, J5’,6’a = 2.1 Hz, J6’a,6’b = 12.3 Hz, 1H, H-6’a), 3.83 (dd, J5,6b = 5 Hz, J6a,6b = 12.6Hz, 1H, H-6b), 3.77-3.71 (m, 1H, H-5), 3.74 (dd, J5’,6’b = 5.6 Hz, J6’a,6’b = 12.1 Hz, 1H, H-6’b), 3.68(dd, J3,4 = 8.4 Hz, J4,5 = 9.7 Hz, 1H, H-4), 3.53-3.39 (m, 3H, H-3’, H-4’, H-5’), 3.33 (dd, J1’,2’ =7.8 Hz, J2’,3’ =9.3 Hz, 1H).13C NMR (101 MHz, D2O) σ 174.96, 161.62, 126.08, 116.52, 103.08 (C -1’), 98.35 (C-1), 82.03(C-3), 75.95 (C-5), 75.80 (C-5’), 75.44 (C-3’), 72.87 (C-2’), 69.41 (C-4’), 68.22 (C-4), 60.58 (C-6’),60.37 (C-6), 54.28 (C-2), 22.11 (CH3).β1,3 linkage was suggested by the 7-8 ppm downfield shift of the C-3 signal.Acholetin*Non-reducing end unit is marked with ’ ”. Unit next to non-reducing end is marked with ”, all otherunits, except for the reducing end, are marked with ’.1H NMR (400 MHz, D2O) σ 5.32-5.28 (m, 1H, H-1a), 4.6-4.51 (m, all b-H-1’s), 4.00-3.94 (H-2),3.99-3.91 (H-3), 3.98-3.93 (m, H-5), 3.95-3.88 (m, H-6a, H-6’a, H-6”a, H-6’ ”a), 3.86-3.77 (m, H-3’),3.86-3.79 (m, 1H, H-3”), 3.82-3.73 (m, H-6b, H-6’b, H-6”b, H6’ ”b), 3.76-3.67 (m, H-2’), 3.70 (dd,1H, J = 8,10 Hz, H-2’ ”), 3.59 (dd, 1H, J = 8,10 Hz, H-3’ ”), 3.59-3.53 (m, 1H, H-4), 3.57-3.43 (m,H-2”, H-4’, H-4”, H-4’ ”, H-5’, H-5”, H-5’ ”).13C NMR (75 MHz, D2O) σ 174.69(C=O), 174.28(C=O), 174.05(C=O), 173.95(C=O), 101.82(C-1”), 101.14(C-1’ ”), 100.75(C-1’), 92.88(C-1α), 80.99(C-3”), 80.00(C-3’), 78.70(C-3α), 75.93(C-5’ ”),75.39(C-5’,C-5”), 73.37(C-3’ ”), 72.07(C-5α), 69.97(C-4’ ”), 68.48(C-4α,C-4’,C-4”), 60.82(C-6,C-6’,C-6”,C-6’ ”), 58.67(C-2”), 55.84(C-2’ ”), 55.03(C-2’), 53.36(C-2α), 22.63(CH3), 22.45(CH3), 20.23(CH3).β1,3 linkage was determined by HMBC H-1, C-3’, H-1’, C-3’, H-1”, C-3’, and H-1’ ”, C-3”C.2 A. laidlawii Operon Table237Table C.1: A. laidlawii operon top BLAST hits.Begin End Strand Top BLAST Annotation3 305 + MULTISPECIES: 6-phosphofructokinase323 1693 + pyruvate kinase1564 1848 - hypothetical protein2028 3908 + surface-anchored NAD dependent sugar epimerase, putative3926 5065 + hypothetical protein5101 8061 + putative DNA/RNA helicase8045 8425 + DNA mismatch repair protein MutT8420 8998 - serine recombinase site-specific9249 10076 + adenine-specific DNA methyltransferase10069 10839 + MULTISPECIES: DNA methyltransferase10836 11687 + type II restriction endonuclease11725 11973 + hypothetical protein12097 12690 + hypothetical protein12927 13019 + hypothetical protein13088 15082 + hypothetical protein15114 15440 + hypothetical protein15510 15761 + hypothetical protein15931 16176 + integral membrane protein16297 17790 + conserved surface-anchored protein Band 7 family18250 18561 + toxin-antitoxin system, toxin component, RelE family18579 19616 + addiction module antidote protein19635 20663 + hypothetical protein20669 21454 - hypothetical protein21757 23898 + DNA helicase24147 25595 + ABC transporter substrate-binding protein25601 26545 + ABC transporter permease26545 27414 + aBC-type transport system permease component27468 30287 + hypothetical protein30384 31553 + glucokinase, ROK family protein31567 34023 + O-glycosyl hydrolase, family 16 [ (GH16)34026 35399 + phosphoglucomutase35404 37932 + putative uncharacterized protein (GH94) [AchP]38285 38845 + integral membrane protein38852 39160 + integral membrane protein39399 40049 + hypothetical protein40033 40755 + hypothetical protein40854 42044 + hypothetical protein42204 43382 + hypothetical protein43375 45153 + Bipolar DNA helicase45156 45368 + hypothetical protein45753 46748 + UDP-glucose 4-epimerase47218 47838 + MarR family transcriptional regulator47856 48968 + aminotransferase DegT48965 49585 + glycosyl transferase possibly involved lipopolysaccharide synthesis [ (GT4)49587 50561 + uncharacterized protein50558 51232 + HAD superfamily hydrolase51246 52445 + putative glucosyltransferase [ (GT4)52447 53460 + UDP-glucose 4-epimerase [53468 54577 + capsular polysaccharide biosynthesis protein Cap8F54579 55709 + MULTISPECIES: UDP-N-acetylglucosamine 2-epimerase55702 56775 + putative sugar transferase [GTnc]56756 57928 + glucosyltransferase [ (GT4)57915 59024 + integral membrane protein59051 60169 + UDP-N-acetylglucosamine 2-epimerase60171 61601 + integral membrane protein61591 62901 + phosphoglucosamine mutase62904 64286 + MULTISPECIES: N-acetylglucosamine-1-phosphate uridyltransferase64529 65866 + NADH oxidase66078 67778 - phosphoenolpyruvate carboxykinase [67904 68557 + esterase68760 70436 + hypothetical protein70972 71943 + hypothetical protein71927 72847 + hypothetical protein73064 73525 + hypothetical protein73549 74493 - L-lactate dehydrogenase74610 75386 - integral membrane protein75663 76988 + trk system potassium uptake protein TrkG76996 77664 + trk system potassium uptake protein TrkA77762 78979 + hypothetical protein79166 81121 + excinuclease ABC subunit B81123 81401 + hypothetical protein81421 83178 + peptidase M3B, oligoendopeptidase F83230 83973 + hypothetical protein83970 84725 + hypothetical protein84664 85389 + putative phosphoesterase domain-containing protein85392 86264 + MULTISPECIES: EDD domain protein, DegV family86261 86989 + hypothetical protein87064 89469 + surface-anchored signal transduction protein89466 90866 + surface-anchored signal transduction protein91210 91953 + hypothetical protein91955 92893 + hypothetical protein92974 93885 + hypothetical protein93993 94682 + hypothetical protein94712 96262 + hypothetical surface-anchored protein96278 98059 + hypothetical protein98056 98649 + hypothetical protein98733 99365 + hypothetical protein99406 101607 + hypothetical protein101675 102493 - mechanosensitive ion channel102549 102872 - ABC transporter substrate-binding protein 238C.3 GP Library Phylogenetic Trees239Figure C.1: GH13 annotated phylogenetic tree. Phylogenetic tree was generate by the JGI and used to select the GPs to synthesize.I subsequently annotated the tree to reflect the expression and activity results and proximity to characterized GPs.240Figure C.2: GH65 annotated phylogenetic tree. Phylogenetic tree was generate by the JGI and used to select the GPs to synthesize.I subsequently annotated the tree to reflect the expression and activity results and proximity to characterized GPs.241Figure C.3: GH94 annotated phylogenetic tree. Phylogenetic tree was generate by the JGI and used to select the GPs to synthesize.I subsequently annotated the tree to reflect the expression and activity results and proximity to characterized GPs.242Figure C.4: GH112 annotated phylogenetic tree. Phylogenetic tree was generate by the JGI and used to select the GPs to synthesize.I subsequently annotated the tree to reflect the expression and activity results and proximity to characterized GPs.243Figure C.5: GH130 annotated phylogenetic tree. Phylogenetic tree was generate by the JGI and used to select the GPs to synthesize.I subsequently annotated the tree to reflect the expression and activity results and proximity to characterized GPs.244C.4 GP Library Expression Tests245Figure C.6: 37 °C GP library expression test (1/6).continued on next page...246Figure C.6: 37 °C GP library expression test (2/6).continued on next page...247Figure C.6: 37 °C GP library expression test (3/6).continued on next page...248Figure C.6: 37 °C GP library expression test (4/6).continued on next page...249Figure C.6: 37 °C GP library expression test (5/6).continued on next page...250Figure C.6: 37 °C GP library expression test (6/6).251Figure C.7: 25 °C GP library expression test (1/6).continued on next page...252Figure C.7: 25 °C GP library expression test (2/6).continued on next page...253Figure C.7: 25 °C GP library expression test (3/6).continued on next page...254Figure C.7: 25 °C GP library expression test (4/6).continued on next page...255Figure C.7: 25 °C GP library expression test (5/6).continued on next page...256Figure C.7: 25 °C GP library expression test (6/6).257Figure C.8: 16 °C GP library expression test (1/6).continued on next page...258Figure C.8: 16 °C GP library expression test (2/6).continued on next page...259Figure C.8: 16 °C GP library expression test (3/6).continued on next page...260Figure C.8: 16 °C GP library expression test (4/6).continued on next page...261Figure C.8: 16 °C GP library expression test (5/6).continued on next page...262Figure C.8: 16 °C GP library expression test (6/6).263Figure C.9: Multi-angle light scattering analysis of acholetin synthesized by the two-pot large scalereaction (1/6).continued on next page...264Figure C.9: Multi-angle light scattering analysis of acholetin synthesized by the two-pot large scalereaction (2/6).continued on next page...265Figure C.9: Multi-angle light scattering analysis of acholetin synthesized by the two-pot large scalereaction (3/6).266
Thesis/Dissertation
eng
Genome Science and Technology
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Withers, Stephen G.
Discovery of new glycoside phosphorylases through rational, genomic and metagenomic exploration
Text