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Structure-function studies of processing alpha-glucosidase-i Konasani, Venkat Rao 2018

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STRUCTURE-FUNCTION STUDIES OF PROCESSING ALPHA-GLUCOSIDASE-I by  Venkat Rao Konasani   A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Food Science)   THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver)   December 2018 © Venkat Rao Konasani, 2018 ii  The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled:  STRUCTURE-FUNCTION STUDIES OF PROCESSING ALPHA-GLUCOSIDASE-I  submitted by Venkat Rao Konasani in partial fulfilment of the requirements for the degree of Doctor of Philosophy in Food Science   Examining Committee: CHRISTINE H. SCAMAN Supervisor   EUNICE LI-CHAN Supervisory Committee Member VIVIEN MEASDAY Supervisory Committee Member  ZHAOMING XU Supervisory Committee Member KEITH ADAMS University Examiner STEPHEN G. WITHERS University Examiner  iii  Abstract Processing α-glucosidase-I (Glu-I) is an endoplasmic reticulum inner membrane-bound enzyme that plays a critical role in N-glycosylation and quality-control of protein-folding. Despite its role, details of the catalytic mechanism of Glu-I function are still unknown. The objective of this research was to address the main obstacles in studying this enzyme, namely the lack of sufficient quantities of relatively pure enzyme and a substrate to test its function. Initially, a soluble form of yeast Glu-I was expressed in Escherichia coli with a yield of 6-8 mg of Glu-I per litre of culture. After single-step purification using immobilised metal affinity chromatography, this recombinant 6xHis-tagged Glu-I showed a Km of 1.27 mM with the synthetic trisaccharide substrate α-D-Glc1,2α-D-Glc1,3α-D-Glc-O-CH3. Since the catalytic domain of Glu-I is located at the C-terminus, expression of the C-terminal domain (Cwh41Δ1-525p) was attempted, but yielded insoluble bodies. Expression of Cwh41Δ1-525p with solubility-enhancing fusion tags or the co-expression of molecular chaperones did not improve solubility. Subsequently, based on a published tertiary structure of Glu-I, I identified that Cwh41Δ1-525p lacks two α-helices of the catalytic (α/α)6 toroid domain. Therefore, the N-terminus of Cwh41Δ1-525p was extended to include the missing helices and expression of the two new constructs (Cwh41Δ1-349p and Cwh41Δ1-314p) was attempted. However, these proteins also expressed as insoluble bodies. Co-expression of the N-terminal domain (Cwhnp) improved the expression of soluble Cwh41Δ1-525p, but the expressed protein was not functional. Catalytic domain released by trypsin hydrolysis from Glu-I was 2.2 times more active than the intact Glu-I. This catalytic domain was purified using size-exclusion chromatography. Since the enzymatic hydrolysis of a glycosidic bond typically occurs with general acid and general base assistance from two amino acid side chains, generally carboxylic amino acids, site-directed mutagenesis of all six conserved carboxylic iv  residues of the catalytic domain was carried out. Glutamic acid 804 was identified as the catalytic base with the aid of nucleophile rescue. Further studies on the structure-function of yeast Glu-I will be helpful in establishing a model to study the inborn errors of metabolism involving this enzyme (CDG-IIb) in humans.    v  Lay Summary Rare diseases, which occur only in a small percentage of population, are attributed to genetic errors. Congenital disorders of glycosylation type IIb (CDG-IIb) is one such rare disease caused by an error in a gene (CWH41) that makes an enzyme called processing glucosidase-I. This enzyme removes the glucose from sugar-containing proteins, which is very important for normal development and physiology. In this work, I study how to produce a related yeast enzyme in a bacterium and purify it to study its function with the goal of establishing a model for the studies of CDG-IIb.  vi  Preface 1. Chapter 2. This chapter is based on a first author manuscript (#1) currently ready for communication to the journal. All experiments, except the synthesis of trisaccharide substrate, presented in this chapter were designed by Venkat Rao with inputs from Dr. Christine Scaman. Venkat Rao carried out the experiments, analysed the results and wrote the manuscript.   This work was also presented as a poster at a conference- The annual meeting of Society for Glycobiology (2011), 9th-12th November 2011 held in Seattle, USA.  Poster title- Synthesis of α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3 and overexpression of processing alpha-glucosidase-I. Venkat Rao Konasani, Akihiro Imamura, Todd Lowary and Christine Scaman   2. Chapter 3. This chapter is based on a first author manuscript (#2) currently in preparation. All experiments presented in this chapter were designed by Venkat Rao with inputs from Dr. Christine Scaman. Venkat Rao carried out the experiments, analysed the results and wrote the manuscript.  This work was presented at the following conference as a research poster: Glycobiology Gordon Research Conference (2013) held during 3rd -8th March 2013 at Ventura, California, USA. Poster title- Expression of the catalytic domain of processing alpha-glucosidase-I. Venkat Rao Konasani and Christine Scaman  3. Chapter 4. This chapter is a part of a manuscript (#2) currently in preparation.  vii  All experiments presented in this chapter were designed by Venkat Rao with inputs from Dr. Christine Scaman. Venkat Rao carried out the experiments, analysed the results and wrote the manuscript.   Synthetic trisaccharide substrate: Dr. Todd Lowary and Dr. Akihiro Imamura from Department of Chemistry, University of Alberta provided the trisaccharide substrate. This substrate was used to assay the activity of the enzyme preparations throughout this study.  viii  Table of Contents Abstract ................................................................................................................................... iii Lay Summary ............................................................................................................................ v Preface ...................................................................................................................................... vi Table of Contents .................................................................................................................. viii List of Tables .......................................................................................................................... xiv List of Figures.......................................................................................................................... xv List of Abbreviations ............................................................................................................xviii Acknowledgements ................................................................................................................ xxii Dedication .............................................................................................................................xxiii Chapter 1: Introduction............................................................................................................1 1.1 Introduction .............................................................................................................2 1.2 Glycosylation ...........................................................................................................3 1.2.1 Glycosylation in prokaryotes ................................................................................4 1.3 N-glycosylation pathway .........................................................................................4 1.3.1 Biosynthesis of N-linked glycans .........................................................................5 1.3.2 Generation of donor nucleotide sugars and lipid-linked monosaccharides.............8 1.4 Glycan assembly ......................................................................................................9 1.4.1 Biosynthesis of lipid-linked oligosaccharide in the cytoplasm ..............................9 1.4.2 Biosynthesis of LLO in ER ................................................................................ 10 1.4.3 Glycan processing .............................................................................................. 12 1.4.4 Alternative glycan processing/trimming pathway ............................................... 14 1.5 Glycan heterogeneity ............................................................................................. 15 ix  1.6 Congenital disorders of glycosylation .................................................................... 16 1.7 Roles of glucose residues of N-linked glycans........................................................ 16 1.8 Processing α-Glucosidase-I .................................................................................... 17 1.8.1 History ............................................................................................................... 17 1.9 Classification and Glu-I orthologs .......................................................................... 20 1.10 Purification of Glu-I ............................................................................................... 21 1.11 Biochemical properties .......................................................................................... 22 1.12 Molecular genetics of Glu-I ................................................................................... 23 1.13 Overexpression of Glu-I ......................................................................................... 24 1.14 Glu-I inhibitors ...................................................................................................... 25 1.15 Substrate specificity ............................................................................................... 27 1.16 Catalytic domain of Glu-I ...................................................................................... 28 1.17 Catalytic residues and mechanism .......................................................................... 29 1.17.1 Glu-I is an inverting glycosidase .................................................................... 29 1.17.2 Genetic mutants ............................................................................................. 31 1.17.3 Chemical modification studies of Glu-I .......................................................... 31 1.17.4 Site-directed mutagenesis studies ................................................................... 32 1.17.5 Structural insights .......................................................................................... 33 1.18 The physiological significance of Glu-I function .................................................... 34 1.19 α-Glucosidase-I and ambiguity in identification ..................................................... 36 1.20 Body of Thesis ....................................................................................................... 37 1.20.1 Lacuna ........................................................................................................... 37 1.20.2 Overall hypothesis .......................................................................................... 37 x  1.20.3 Overall objectives .......................................................................................... 37 Chapter 2: Recombinant expression and characterisation of Glu-I ......................................... 40 2.1 Summary ............................................................................................................... 41 2.2 Introduction ........................................................................................................... 42 2.3 Materials and methods ........................................................................................... 44 2.3.1 Materials ............................................................................................................ 44 2.3.2 Microbial strains and plasmid............................................................................. 44 2.3.3 Isolation of yeast genomic DNA ........................................................................ 45 2.3.4 Cloning and preparation of expression constructs of Glu-I ................................. 45 2.3.5 Expression of Glu-I in E. coli ............................................................................. 46 2.3.6 Purification of recombinant Glu-I ....................................................................... 47 2.3.7 Expression of codon optimised Glu-I ................................................................. 48 2.3.8 Synthesis of α-D-Glc1,2α-D-Glc1,3α-D-Glc–OCH3 ........................................... 49 2.3.9 Enzyme assays ................................................................................................... 52 2.3.10 Activity with other substrates ......................................................................... 52 2.3.11 Determination of Km ..................................................................................... 52 2.3.12 Other methods ................................................................................................ 53 2.4 Results ................................................................................................................... 53 2.4.1 Cloning and preparation of expression constructs of Glu-I gene ......................... 53 2.4.2 Expression and purification of Glu-I in E. coli ................................................... 54 2.4.3 Expression of codon optimised Glu-I ................................................................. 59 2.4.4 Synthesis of trisaccharide and Glu-I activity assay ............................................. 62 2.4.5 Substrate specificity ........................................................................................... 64 xi  2.5 Discussion ............................................................................................................. 65 Chapter 3: Heterologous expression of yeast Glu-I truncations in E. coli and isolation of the catalytic domain of Glu-I ....................................................................................................... 68 3.1 Summary ............................................................................................................... 69 3.2 Introduction ........................................................................................................... 70 3.3 Materials and methods ........................................................................................... 73 3.3.1 Glu-I truncations ................................................................................................ 73 3.3.2 Co-expression of molecular chaperones ............................................................. 76 3.3.3 Co-expression of N- and C-terminus .................................................................. 76 3.3.4 Trypsin hydrolysis of Glu-I ................................................................................ 78 3.3.5 Isolation of catalytic domain of Glu-I ................................................................. 78 3.4 Results ................................................................................................................... 78 3.4.1 Expression of Glu-I truncations .......................................................................... 78 3.4.2 Effect of solubility enhancing tags and co-expression of molecular chaperones on the solubility of Cwh41Δ1-525p .................................................................................... 81 3.4.3 Co-expression of the Glu-I N- and C-terminus ................................................... 87 3.4.4 Trypsin hydrolysis of Glu-I ................................................................................ 91 3.4.5 Purification of Glu-I catalytic domain using size exclusion chromatography ...... 92 3.5 Discussion ............................................................................................................. 95 Chapter 4: Identification of the catalytic base of processing α-glucosidase-I .......................... 98 4.1 Summary ............................................................................................................... 99 4.2 Introduction ......................................................................................................... 100 4.3 Materials and methods ......................................................................................... 103 xii  4.3.1 Homology and in-silico analysis....................................................................... 103 4.3.2 Site-directed mutagenesis ................................................................................. 103 4.3.3 Expression and purification of wild-type Glu-I and its mutants ........................ 105 4.3.4 The Glu-I activity of mutants ........................................................................... 105 4.3.5 Secondary structure .......................................................................................... 105 4.3.6 Nucleophile rescue of activity of Glu-I mutants................................................ 105 4.4 Results ................................................................................................................. 106 4.4.1 Homology and in-silico analysis....................................................................... 106 4.4.2 Site-directed mutagenesis ................................................................................. 106 4.4.3 Purification of wild-type and mutant Cwh41Δ1-34p ......................................... 107 4.4.4 Secondary structure analysis of mutants ........................................................... 109 4.4.5 The Glu-I activity of mutants ........................................................................... 110 4.4.6 Chemical rescue ............................................................................................... 111 4.5 Discussion ........................................................................................................... 114 Chapter 5: Overall conclusions and future perspectives ....................................................... 119 5.1 Overall Conclusions ............................................................................................. 120 5.1.1 Yeast Glu-I as a model for family 63 glycosyl hydrolases ................................ 121 5.2 Future perspectives .............................................................................................. 124 5.2.1 Expression of the catalytic domain. .................................................................. 124 5.2.2 Substrate specificity ......................................................................................... 124 5.2.3 Catalytic mechanism ........................................................................................ 125 5.2.4 Glu-I - OST - Alg10p and their interactions ..................................................... 126 Bibliography .......................................................................................................................... 127 xiii  Appendix ............................................................................................................................... 150  xiv  List of Tables Table 1.1. Inhibitors of processing alpha-glucosidase I .............................................................. 26 Table 2.1. Primers used to clone/sequence the Glu-I gene ......................................................... 46 Table 2.2 Purification of recombinant Glu-I from E. colia .......................................................... 57 Table 2.3 Km of Glu-I against different synthetic substrates ...................................................... 65 Table 3.1. Primers used for amplification of truncated Glu-I genes. ........................................... 73 Table 3.2. List of plasmids and molecular chaperones ............................................................... 76 Table 3.3. Glu-I assay of the co-expression of Cwhnp and Cwh41Δ1-525p. .............................. 89 Table 3.4. Glu-I activity assay of Cwh41Δ1-525p fraction from size-exclusion chromatography of trypsinised Cwh41Δ1-34p ..................................................................................................... 94 Table 4.1. List of mutant primers............................................................................................. 104 Table 4.2. Distances between different conserved carboxylic residues ..................................... 109 Table 4.3. Catalytic mechanism and catalytic acid/base of inverting glycoside hydrolases ....... 113  xv  List of Figures Figure 1.1. Structure of oligosaccharide.. ....................................................................................6 Figure 1.2 Types of N-linked glycans.. ........................................................................................7 Figure 1.3 Compartmentalisation of N-glycosylation.. .................................................................8 Figure 1.4. Schematic representation of glycan assembly and processing in the ER...................... 11 Figure 1.5. Schematic representation of glycan processing in Golgi apparatus.. ......................... 14 Figure 1.6 Cartoon depiction of Glu-I domains (not to scale).. ................................................... 19 Figure 1.7. Mechanism of glycoside hydrolase.. ........................................................................ 30 Figure 1.8. Structures of GH family 63 enzymes.. ..................................................................... 33 Figure 2.1. Graphical depiction of pET30a-CWH41Δ1-34.. ...................................................... 47 Figure 2.2. Schematic illustration of the synthesis of α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3.. .. 51 Figure 2.3 Restriction analysis of pET30a(+) vector and insert (CWH41Δ1-34).. ...................... 54 Figure 2.4 Restriction analysis of pET30a-CWH41Δ1-34 clones.. ............................................. 54 Figure 2.5. The screening of E. coli BL21(DE3) transformants for Cwh41Δ1-34p expression.. . 55 Figure 2.6. Elution of recombinant Glu-I from Ni-NTA column.. .............................................. 57 Figure 2.7 Size exclusion chromatography of Cwh41Δ1-34p.. ................................................... 58 Figure 2.8 SDS-PAGE analysis of Cwh41Δ1-34p.. ................................................................... 58 Figure 2.9. Ni-NTA Purification of Cwh41Δ1-34p heterologously expressed in E. coli BL21(DE3) [--] and E. coli RosettaGami®  (--)......................................................................... 59 Figure 2.10 The frequency of optimal codons for Cwh41Δ1-34p in E. coli.. .............................. 60 Figure 2.11. Purification of Cwh41Δ1-34p and Cwh41Δ1-34p-OPT expressed in E. coli BL21(DE3).. ............................................................................................................................. 61 Figure 2.12. 1HNMR analysis of the synthetic trisaccharide substrate.. ...................................... 63 xvi  Figure 2.13 Michaelis-Menten plot of Glu-I activity with the synthetic trisaccharide- α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3.. ............................................................................................ 64 Figure 3.1. Graphical depiction of Glu-I domains and positions of truncations.. ........................ 72 Figure 3.2 Graphical depiction of pGEX4T1- CWH41Δ1-525 (A) and pMALC5E- CWH41Δ1-525 (B).. .................................................................................................................................... 75 Figure 3.3 Vector map of pACYC-DUET1-CWHN-CWH41Δ1-525.. ....................................... 77 Figure 3.4. SDS-PAGE analysis of pET30a-CWH41Δ1-314 expression.. .................................. 79 Figure 3.5. SDS-PAGE analysis of pET30a-CWH41Δ1-349 expression.. .................................. 79 Figure 3.6. Purification of truncations of Glu-I by Ni-NTA chromatography.. ........................... 80 Figure 3.7. SDS-PAGE analysis of Cwh41Δ1-525p.. ................................................................ 80 Figure 3.8. SDS-PAGE analysis of pGEX4T1- CWH41Δ1-525.. .............................................. 82 Figure 3.9. SDS-PAGE analysis of pMALC5E- CWH41Δ1-280.. ............................................. 82 Figure 3.10. SDS-PAGE analysis of pMALC5E- CWH41Δ1-525.. ........................................... 83 Figure 3.11. Ni-NTA chromatography of Glu-I truncations (Cwh41Δ1-525p and Cwh41Δ1-280p) fused with solubility enhancing tags- MBP and GST.. ..................................................... 84 Figure 3.12. SDS-PAGE analysis of co-expression of Cwh41Δ1-525p with molecular chaperones GroEL and GroES.. ................................................................................................................... 85 Figure 3.13. SDS-PAGE analysis of co-expression of Cwh41Δ1-525p with molecular chaperones DnaJ and DnaK.. ....................................................................................................................... 85 Figure 3.14. SDS-PAGE analysis of co-expression of Cwh41Δ1-525p with molecular chaperone trigger factor.. ........................................................................................................................... 86 Figure 3.15. Ni-NTA chromatography of Cwh41Δ1-525p co-expressed with molecular chaperones.. .............................................................................................................................. 86 xvii  Figure 3.16. SDS-PAGE analysis of pACYC-DUET1-CWHN expression................................. 88 Figure 3.17. SDS-PAGE analysis of pACYC-DUET1- CWH41Δ1-525 expression.. ................. 88 Figure 3.18. SDS-PAGE analysis of pACYC-DUET1-CWHN-CWH41Δ1-525 expression.. ..... 89 Figure 3.19. Ni-NTA chromatography of Cwh41Δ1-525p.. ....................................................... 90 Figure 3.20. Trypsin digestion of Cwh41Δ1-34p.. ..................................................................... 91 Figure 3.21. Effect of trypsin hydrolysis on the activity of Cwh41Δ1-34p.. ............................... 92 Figure 3.22. Separation of the polypeptide fragments generated from trypsin digested Glu-I by size exclusion chromatography using HiPrep-Sephacryl-S100HR column.. ............................... 93 Figure 3.23. SDS-PAGE analysis of the fractions collected during the size exclusion chromatography using HiPrep-Sephacryl-S100HR column........................................................ 94 Figure 4.1. Homology analysis of C-terminal catalytic domain of Glu-I orthologs. .................. 107 Figure 4.2. Orientation of conserved carboxylic residues in Glu-I active site. .......................... 108 Figure 4.3. The distance between carboxylic residues.. ............................................................ 108 Figure 4.4. Circular dichroism of wild-type and mutant Cwh41Δ1-34p.. ................................. 110 Figure 4.5. Glu-I activity of mutants of Cwh41Δ1-34p.. .......................................................... 111 Figure 4.6. Nucleophile rescue of Glu-I activity of E804A mutant of Cwh41Δ1-34p.. ............. 112 Figure 4.7. Nucleophile rescue of Glu-I activity of mutants of Cwh41Δ1-34p.. ....................... 112 Figure 4.8. Schematic representation of azide rescue.. ............................................................. 114 Figure 5.1. Structures of processing α-glucosidase-I subfamily of Family 63 glycosyl hydrolases.. ............................................................................................................................. 122 Figure 5.2. Alignment of C-terminal domain of yeast Glu-I (Sc) and human MOGS (Hs)........ 123  xviii  List of Abbreviations Alg  asparagine-linked glycosylation defective mutant  bp  base pair CAI  codon adaptation index  CDG  congenital disorder of glycosylation CHO  Chinese hamster ovary cells Cnx  calnexin ConA  concanavalin A CP-DNM N-5’-Carboxypentyl-deoxynojirimycin CPTM  co- or post-translational modifications Crt  calreticulin CWH   calcofluor white hypersensitive CWH41 gene that encodes yeast processing α-glucosidase I DALI  distance matrix alignment DDQ   2, 3-dichloro-5,6-dicyanobenzoquinone DEAE  diethylaminoethyl DEPC  diethylpyrocarbonate DNA  deoxyribonucleic acid dNTP  deoxyribonucleotide triphosphate Dol-P  dolichyl-phosphate Dol-P-P  dolichol-pyrophosphate EC  enzyme commission  E. coli  Escherichia coli EDAC  3-(3-(dimethylamino) propyl) carbodiimide  EDEM endoplasmic reticulum degradation-enhancing α-mannosidase-like                                                  proteins EDTA  ethylene-diamine-tetra-acetic acid ER  endoplasmic reticulum ERAD  endoplasmic reticulum-associated degradation of protein FPLC  fast protein liquid chromatography xix  Fuc  fucose g  gravitational acceleration constant (9.8 ms-2) Gal  galactose GalNAc N-acetyl galactosamine GDP  guanosine diphosphate Glc  glucose GlcNAc N-acetyl glucosamine Glu-I  processing alpha-glucosidase-I or processing α-glucosidase-I Glu-II  processing alpha-glucosidase-II or processing α-glucosidase-II GPI  glycophosphatidylinositol 6xHis-tag 6xHistidine tag hr  hour(s) His-Trap HP Nickel-Sepharose high-performance column IMAC  immobilised metal affinity chromatography IPTG  isopropyl β-D-1-thiogalactopyranoside IUBMB International Union of Biochemistry and Molecular Biology kb  kilobases  kcat  catalytic rate constant kcat/Km specificity constant kDa  kilodaltons  Ki  enzyme-inhibitor complex dissociation constant Km  Michaelis constant LB  Luria-Bertani medium LLO  lipid-linked oligosaccharide Man  mannose MBP  maltose binding protein MCS  multiple cloning site µg  microgram µL  microlitre mg  milligram xx  min  minute(s)  ml  millilitre  mM  millimolar NEM  N-ethylmaleimide Ni-NTA Nickel linked nitrilotriacetic acid nm  nanometer nmole  nanomole NMR  nuclear magnetic resonance OD600  optical density at 600 nm ORF  open reading frame OST  oligosaccharyltransferase PAGE  polyacrylamide gel electrophoresis PCR  polymerase chain reaction pI  isoelectric point PMB   p-methoxybenzyl PMSF  phenylmethylsulfonyl fluoride  P. pastoris Pichia pastoris QC  quality control  rpm  revolutions per minute S. cerevisiae Saccharomyces cerevisiae SDS  sodium dodecyl sulfate SEC  size exclusion chromatography Sial  sialic acid TNM  tetranitromethane Tris  2-amino-2-(hydroxymethyl)-1,3-propanediol U  units of activity UDP  uridine diphosphate UGGT  UDP-Glc: glycoprotein glucosyltransferase UV  ultraviolet  xxi  Nucleotide base abbreviations A Adenine C Cytosine G Guanine T Thymine  Common amino acid abbreviations A Ala Alanine C Cys Cysteine D Asp Aspartic acid  E Glu Glutamic acid F Phe Phenylalanine  G Gly Glycine H His Histidine I Ile Isoleucine K Lys Lysine L Leu Leucine M Met Methionine N Asn Asparagine P Pro Proline Q Gln Glutamine R Arg Arginine S Ser Serine T Thr Threonine V Val Valine W Trp Tryptophan Y Tyr Tyrosine  xxii  Acknowledgements First, I would like to express my gratitude to my supervisor Dr. Christine H. Scaman for the continuous support, for her vast knowledge, immense patience, and motivation. Her guidance helped me innumerable times. I would like to thank her for the freedom she gave me regarding thinking outside the box and executing the experiments.   I would also like to acknowledge the members of my supervisory committee Dr. Eunice Li-Chan, Dr. Vivien Measday and Dr. Zhaoming Xu for their valuable guidance and feedback throughout the studies.   I would like to acknowledge exceptional technical help received from Valerie Skura and Pedro Aloise. I would also like to acknowledge Dr. Fred Rosell for his help in circular dichroism spectrophotometric measurements, and Dr. Vijay Kumar Somalinga for interesting discussions related to protein expression.   I would like to thank Dr. Todd Lowary and Dr. Akihiro Imamura (Department of Chemistry, University of Alberta) for helping with the synthesis of trisaccharide substrate.   I thank my fellow lab mates Dr. Andrea Goldson, Dr. Ingrid Elisia, Reena Mistry, Orly Varon and Zhong Peng for the interesting discussions and positive working atmosphere. I would also like to thank Arisa Thamsuaidee, Guillaume Malgorn, Cecile Boyer and Amandine Arenas for their technical assistance during my research studies. I would also like to thank Barbara Wakal for proof-reading this thesis.   I would like to thank the funding agency NSERC (grant to Dr. Scaman) for supporting my research. I would also like to acknowledge the financial support received through the awards Dr. Shruyo Nakai Memorial Award, Ulrich Freybe Memorial Award and awards from The Faculty of Land and Food Systems which helped me to pay my tuition fees.   xxiii  Dedication I would like to dedicate this thesis to a few people who have shaped me through their thoughts-  My family, my PhD supervisor Dr. Christine H. Scaman, and the late Dr. Robert G. Spiro   “Learn as if you would live forever, live as if you would die tomorrow".  1  Chapter 1: Introduction   2  1.1 Introduction N-linked glycan processing starts by removing the distal glucose residue from Glc3Man9GlcNAc2 and generating di-glucosylated glycans [1, 2]. The N-linked glycans are further modified by other processing enzymes including processing alpha-glucosidase-II, mannosidases, and glycosyltransferases [3]. The processing of N-glycans recruits molecular chaperones calnexin and calreticulin which assist the nascent proteins in achieving their native conformation. With the labyrinthine and synergistic activities of these glycoside hydrolases (GH), glycosyltransferases, and chaperones, glycoproteins acquire their native conformation and protein folding quality control is carried out. Failure to release the distal glucose, and the lack of glycan diversification, leads to severe physiological abnormalities in yeasts, plants, nematodes and humans [4-8]. Despite its physiological significance, details of the catalytic mechanism of Glu-I are still not known. Hence, this thesis research is aimed at studying the structure-function of Glu-I and furthering our understanding of the catalytic mechanism of Glu-I. The N-glycosylation pathway is well-conserved in all eukaryotes from unicellular to multicellular organisms [9]. Therefore, Glu-I from a well-studied unicellular eukaryote Saccharomyces cerevisiae was selected as the model system for this study.  Yeast Glu-I is present at the inner membrane of the endoplasmic reticulum (ER) with the catalytic domain projected into the lumen. It is composed of 833 amino acids and encoded by the CWH41 gene in yeast [10]. A soluble non-glycosylated form of this enzyme, a 94 kDa protein that lacks the 34 N-terminal residues composing the cytoplasmic and transmembrane domain, was also observed during purification from the yeast [11, 12]. Yeast and mammalian Glu-I have 24% overall identity and share similar biochemical properties [13]. Thus, yeast Glu-I can serve as an excellent model to understand the catalytic mechanism of the mammalian Glu-I counterpart. 3  1.2 Glycosylation Most proteins undergo various co- and post-translational modifications (CPTM). The nascent polypeptide may be altered by proteolytic cleavage, the formation of disulphide bonds, or covalent attachment of various chemical moieties including carbohydrate, lipid, phosphate, sulphate, and alkyl groups. These modifications may alter physicochemical properties, conformation, stability, and distribution of the target protein, which eventually affect the function [14-19]. In some cases, the attached covalent group can itself act as a functional component of the protein. Hence, CPTM of proteins are extremely important. One of the most common forms of CPTM is glycosylation, and it is also the most complicated CPTM that a protein can undergo. Glycosylation involves the synthesis of precursor glycans, transfer of these precursor glycans to nascent proteins and processing of protein-linked glycans by an array of enzymes and leads to the formation of a wide range of glycoproteins with diverse glycan chains [3]. The diversity arises from eight different amino acids that can serve as the point of glycan attachment, the thirteen sugars that can be present, and the various linkages [3]. Based on the type of linkage between the amino acid and sugar, there are five types of glycosylation: N-glycosylation, O-glycosylation, phosphoglycosylation, C-mannosylation and glypiation [3]. Among these, N-glycosylation and O-glycosylation are the most common. In N-glycosylation, the sugar moiety is linked to the asparagine nitrogen on the consensus amino acid sequence, Asn-Xxx-Ser/Thr [20, 21]. In O-glycosylation, the linkage is between the sugar and hydroxyl group of amino acids such as serine, threonine, tyrosine, proline and lysine [22]. The attachment of glycans to the polypeptide chain via a phosphodiester bond is termed phosphoglycosylation which was first detected in lysosomal serine proteases in Dictyostelium discoideum where GlcNAc-α-1-P residues are linked to serine via a phosphodiester linkage [23]. In Trypanosoma cruzi rhamnose-, xylose-, and galactose-containing phosphor 4  glycans are linked to threonine on proteins by a phosphodiester linkage [24]. C-mannosylation involves the attachment of a mannose residue to C2 of the first tryptophan of Trp-Xxx-Xxx-Trp sequence in proteins [3, 25]. Glypiation involves the attachment of a glycophosphatidylinositol (GPI) to the carboxy-terminal amino acid residue via an amide bond and is generally observed among eukaryotic cell surface glycoproteins [26]. 1.2.1 Glycosylation in prokaryotes Protein glycosylation was first reported by Neuberger [27], it has been thought to occur only in eukaryotes. However, during the last decade, there have been many reports on the existence of glycosylation in bacteria and Archaea, and interest in bacterial glycosylation has increased tremendously due to its applications in medicine such as development of carbohydrate vaccines and large-scale production of recombinant glycoproteins [28]. Bacterial N-glycosylation was first observed in Campylobacter jejuni, and O-glycosylation has been shown to occur in several other bacterial pathogens, such as Neisseria gonorrhoeae and Helicobacter pylori [29].  1.3 N-glycosylation pathway N-glycosylation is a highly orchestrated process involving sequential and coordinated reactions catalysed by glycoside hydrolases and transferases that take place in the ER and Golgi apparatus. These glycan-processing enzymes and their intricate interactions allow the formation of a plethora of carbohydrate structures, from simple high mannose N-linked glycans to complex and hybrid N-linked glycans, which contribute to the physiological and biochemical functions of the glycoproteins. N-glycosylation occurs on the amide nitrogen of an asparagine residue in the Asn-Xxx-Ser/Thr consensus sequon where Xxx can be any amino acid but proline. Although consensus sequons are essential for N-glycosylation, not all consensus sequons are glycosylated. Several 5  factors such as composition, position and the neighbouring sequons in the protein, and secondary/tertiary structure of the protein affect the glycosylation state of the protein [30-33]. In eukaryotic proteins, despite the presence of more Asn-Xxx-Ser sequons than the Asn-Xxx-Thr sequons, glycosylation occurs less frequently at Asn-Xxx-Ser [33]. Furthermore, there are exceptions to the canonical consensus sequence; in some plants, yeasts and mammals Asn-Xxx-Cys sequences act as the specific site for N-glycosylation [34-36]. N-linked oligosaccharides of eukaryotic glycoproteins fall into three different classes: high mannose, hybrid and complex (Figure 1.1 and 1.2). All these classes of N-linked glycans have a core oligosaccharide (Man3GlcNAc2) at the reducing end. High mannose N-linked glycans contain additional mannose residues α-linked to this core oligosaccharide whereas complex N-linked glycans contain other sugars such as N-acetyl glucosamine (GlcNAc), galactose (Gal), fucose (Fuc) and sialic acid (Sial) (see Figure 1.2). 1.3.1 Biosynthesis of N-linked glycans Although N-linked glycans are diverse, most of them are synthesised from a common dolichol pyrophosphate as a lipid-linked oligosaccharide (LLO) comprising three glucoses, nine mannoses and two GlcNAc residues (Figure 1.1A). Dolichol is an isoprenoid lipid that acts as a foundation for the synthesis of core oligosaccharide. The availability of dolichol is a rate-limiting factor in the assembly of glycans [37-39]. The chain length of dolichol varies in different organisms. In yeasts, it mostly consists of 15-16 isoprene units whereas in mammals 18-21 isoprene units are found [40, 41]. The sugar residues are arranged in a three antennae structure: A-antenna (α-1,3 arm), B- antenna (inner arm), and C- antenna (α-1,6 arm) [Figure 1.1B]. The core of the precursor oligosaccharide is conserved in most eukaryotes except in some protists and yeasts. Some flagellated protists (T. cruzi and Leishmania mexicana), a ciliate (Tetrahymena pyriformis), and a 6  pathogenic yeast (Cryptococcus neoformans) lack the three glucose residues on A-antenna of the N-glycan (Man9GlcNAc2) [42-44].    Figure 1.1. Structure of oligosaccharide. A. lipid-linked B. Asparagine (protein)-linked. Antennae were highlighted in boxes (a, b and c indicate antenna A, antenna B and antenna C respectively). Monosaccharide symbols in this figure follow the SNFG (Symbol Nomenclature for Glycans) system.   7   Figure 1.2 Types of N-linked glycans. Monosaccharide symbols in this figure follow the SNFG nomenclature, details of which can be found at http://www.ncbi.nlm.nih.gov/books/NBK310273/.  Some protists such as T. pyriformis lack some mannose residues but retain the glucose residues [45]. L. mexicana lacks both the glucose residues and mannose residues- Man9GlcNAc2 [46, 47]. Intra-genus and inter-genus variations in the structure of bacterial glycans are common. Campylobacter genus was reported to have 16 different structures [28]. In archaea, N-linked glycans are diverse regarding size, level of branching, and type of the linking sugar [48]. In eukaryotes, the LLO is synthesized by step-wise addition of nucleotide-activated sugars in a series of reactions catalysed by trans-membrane ER enzymes called glycosyltransferases. The first seven sugar addition reactions are carried out on the cytosolic side of ER; then the hepta-saccharide is flipped to the luminal side and undergoes further extension to form a 14-residue mature oligosaccharide precursor, Glc3Man9GlcNAc2. In eukaryotes, N-glycosylation is compartmentalised and consists of the following stages: 1. Generation of donor nucleotide sugars, 2. Glycan assembly, and 3. Glycan processing (Figure 1.3). These stages are discussed in detail in the following sections.  8   Figure 1.3 Compartmentalisation of N-glycosylation. Glycan assembly starts in the cytoplasm and ends in the ER. Processing starts immediately after transfer in the ER and as glycans move through the Golgi apparatus.  1.3.2 Generation of donor nucleotide sugars and lipid-linked monosaccharides The assembly of N-linked glycans significantly depends on the transfer of a monosaccharide donor to a glycan acceptor. These monosaccharide sugars are either delivered as activated nucleotide sugars or dolichol pyrophosphate-linked sugars (Figure 1.4).  Dolichol phosphate (Dol-P) links the glycans to the ER membrane and carries them along the membrane. De novo synthesis of Dol-P starts with the sequential addition of C5 isoprenoid units to farnesylpyrophosphate by cis-prenyltransferase followed by phosphorylation by dolichol kinase [49]. Dol-P can also be generated from dolichol pyrophosphate (Dol-PP), and by the action of glycosyltransferases on Dol-P-Man and Dol-P-Glc. Synthesis of the dolichol-linked oligosaccharide requires three different nucleotide-activated sugar building blocks: UDP-N-acetylglucosamine (UDP-GlcNAc), GDP-mannose (GDP-Man) and UDP-glucose (UDP-Glc). The synthesis of nucleotide-sugars starts from phosphorylated sugar units which are then linked with nucleotides. The synthesis of UDP-Glc starts from glucose-6-phosphate (Glc-6-P) synthesis by glucose-6-phosphatase. 9  Phosphoglucomutase then reversibly converts the Glc-6-P to Glc-1-P [50]. Finally, UDP-Glc is generated from Glc-1-P by UDP-Glc pyrophosphorylase (UGPase) [51]. Biosynthesis of UDP-GlcNAc starts with the conversion of fructose-6-P to glucosamine-6-P by glucosamine:fructose-6-P-amidotransferase [52].   Glucosamine-6-P is then N-acetylated by an acetylase to N-acetylglucosamine-6-P which is subsequently transformed to N-acetylglucosamine-1-P UDP-GlcNAc by N-acetylglucosamine-phosphomutase [50]. GDP-Man is synthesised from fructose-6-P by three enzyme-catalysed reactions: phosphomannoisomerase catalyses the isomerisation of fructose-6-P to mannose-6-P, followed by the conversion of mannose-6-P to mannose-1-P by phosphomannomutase [53], and finally the formation of GDP-Man from mannose-1-P by GDP-Man pyrophosphorylase [54]. 1.4 Glycan assembly 1.4.1 Biosynthesis of lipid-linked oligosaccharide in the cytoplasm Two kinds of glycosyltransferases are involved in the biosynthesis of LLO; glycosyltransferases that use nucleotide-activated sugars as substrates, and glycosyltransferases that utilise Dol-P-bound sugar substrates. These enzymes are encoded by the ALG (asparagine-linked glycosylation) genes. The glycan biosynthesis process is initiated by the addition of GlcNAc-P to Dol-P forming the dolichol-pyrophosphate-GlcNAc (Dol-PP-GlcNAc) [55-57] which is catalysed by a multienzyme glycosyltransferase complex: Alg7p/Alg13p/Alg14p [58]. Alg7p, a transmembrane protein, catalyses the transfer of GlcNAc from UDP-GlcNAc to Dol-P [57]. The second GlcNAc residue is transferred from UDP-GlcNAc by Alg13p/Alg14p UDP-GlcNAc transferase to form GlcNAc2-PP-dol [59]. Further extension of Dol-PP-(GlcNAc)2 occurs with the addition of mannose residues. β-1,4 mannosyltransferase, encoded by ALG1, catalyses the addition of first 10  mannose residue [60]. Subsequently, two branching mannose residues are added to the Dol-PP-GlcNAc2Man by Alg2p to form Dol-PP-GlcNAc2Man3 [61]. Dol-PP-GlcNAc2Man3 is further elongated to form the final product of cytoplasmic glycan biosynthesis Dol-PP-GlcNAc2Man5. This elongation reaction is achieved by the Alg11p [62]. The hepta-saccharide is translocated to the ER lumen by a flippase [63], and the translocated hepta-saccharide undergoes further extension in the ER lumen.  1.4.2 Biosynthesis of LLO in ER Dol-PP-GlcNAc2Man5 is extended by the addition of four mannose and three glucose residues transferred from Dol-P-Man- and Dol-P-Glc on the lumenal side of ER. Lumenal biosynthesis is initiated by the α1,3 mannosyltransferase encoded by the ALG3 locus, followed by the addition of an α1,2-linked mannose by the Alg9p α1,2 mannosyltransferase, resulting in the B-antenna of the oligosaccharide [64, 65]. The C-antenna is initiated by the Alg12p α1,6 mannosyltransferase and completed by Alg9p that adds α1,2-linked mannose [66].  While the synthesis of Man9GlcNAc2 oligosaccharide structure is occurring, Alg6p starts the glucosylation of the A-antenna [67]. Addition of the second α1,3-linked Glc residue to the LLO is catalysed by Alg8p [68]. Alg10p then catalyses the final step in LLO synthesis, the addition of α1,2-linked glucose [69].      11   Figure 1.4. Schematic representation of glycan assembly and processing in the ER. Glycan assembly is highlighted with the blue background. Various activated sugars required during glycan assembly are supplied from the cytoplasm. Processing of N-linked glycans starts immediately after the transfer of the glycan chain onto the nascent protein. Nascent glycoproteins undergo Calnexin/calreticulin (Cnx/Crt) cycle and acquire the natively folded state. Terminally misfolded proteins are targeted for degradation in the cytoplasm. Monosaccharide symbols in this figure follow the SNFG Nomenclature, details of which can be found at http://www.ncbi.nlm.nih.gov/books/NBK310273/). Abbreviations: alg- asparagine-linked glycosylation, alg1, alg2 etc.; GDP- guanosine diphosphate; UDP- Uridine diphosphate; OST- oligosaccharyl transferase; Glu-I- Processing α-glucosidase-I; Glu-II - Processing α-glucosidase-II; UGGT- UDP-glucose:glycoprotein glucosyltransferase; Cnx- Calnexin; Crt-Calreticulin; EDEM- ER degradation-enhancing α-mannosidase-like protein; ERAD- Endoplasmic-reticulum-associated protein degradation.    12  1.4.3 Glycan processing 1.4.3.1 Glycan processing in ER N-glycosylation is initiated by the transfer of the pre-assembled core oligosaccharide, Glc3Man9GlcNAc2, from Dol-PP to the polypeptide chains that are entering the ER by the oligosaccharyltransferase (OST) [70, 71]. Processing of the protein-bound oligosaccharides dictates the fate of the associated protein. Several ER-resident proteins that can bind and hydrolyse the protein-bound oligosaccharides mediate protein folding and quality control. This processing starts with a series of trimming steps immediately after the glycan transfer. Glu-I catalyses the removal of the terminal α 1,2-linked glucose [72]. Removal of terminal glucose enables the N-glycan to associate with an N-glycan-binding protein called malectin, an ER-localized type-I membrane protein [73]. Malectin may then facilitate the binding of processing α-Glucosidase-II (Glu-II) for further deglycosylation, and prevent aggregation of nascent polypeptides during the early synthesis period [73, 74]. Glu-II subsequently removes the second glucose which enables the glycoprotein to bind to the chaperones, calnexin (Cnx) and calreticulin (Crt), as well as the oxidoreductase ERp57 [75, 76]. Cnx and Crt are lectins that recognise and bind to the glycoproteins that contain monoglycosylated glycans and help them to achieve native folding [77]. ERp57 catalyses the formation of inter- and intramolecular disulfide bonds, a rate-limiting step in the folding of nascent proteins [78-80]. Following the Cnx/Crt cycle, the final glucose is removed by Glu-II which prevents further binding by Cnx and Crt [77]. If at this stage, the glycoprotein is not correctly folded, the UDP-glucose glycoprotein glucosyltransferase (UGGT) enzyme adds the terminal glucose, and the glycoprotein enters another Cnx/Crt cycle of chaperone-assisted folding [77, 81, 82]. If the target protein achieves native folding, ER α-mannosidase-I catalyses the removal of the terminal mannose residue from the B-antenna of the deglucosylated glycan on the 13  folded proteins to form the Man8GlcNAc2 core [83]. Correctly folded proteins, except ER-resident glycoproteins, are then transported to the cis-Golgi cisternae for further processing whereas terminally misfolded proteins enter the ER-associated degradation (ERAD) pathway. ER degradation-enhancing α-mannosidase-like proteins (EDEM 1, 2 and 3) interact with calnexin as well as ER α-mannosidase-I and accelerates the removal of α 1,2-linked mannose from the C-antenna of the glycan of misfolded proteins [84, 85]. These α1,2-de-mannosylated proteins are labelled as the misfolded proteins and are retro-translocated to the cytoplasm for degradation by the proteasome [86].   1.4.3.2 Glycan processing in Golgi The Golgi apparatus is home to a multitude of glycosyltransferases and glycoside hydrolases that modify glycans to create a wide array of structures (Figure 1.5). There are more than 142 glycosyltransferases in the Golgi - most of these catalyse the addition of a single sugar molecule to the glycan precursor [87]. The N-linked glycans are predominantly high mannose type and contain 8-9 mannose residues. The first glycosyl transfer reaction in the Golgi involves the transfer of a GlcNAc residue onto Man5GlcNAc2 to form GlcNAcMan5GlcNAc2. Hybrid type glycans retain the five mannose residues and undergo further extension of the GlcNAc arm with the transfer of galactose and sialic acid and other sugars. N-linked glycans lose two distal Man residues and attain a second GlcNAc to form a bi-antennary complex N-linked glycan. These complex glycans may be additionally branched up to six times, and each branch may be further extended by the addition of different sugars including Gal, GlcNAc, GalNAc, Fuc, Sial, and disaccharide units [87].  14   Figure 1.5. Schematic representation of glycan processing in Golgi apparatus. Trimming of high-mannose N-linked glycans takes place in cis-Golgi compartments by mannosidases. Formation of complex N-linked glycans and their branching and core fucosylation takes place in medial-compartments. Complex N-linked glycans undergo galactosylation, sialylation, and external fucosylation in trans- compartments. Monosaccharide symbols in this figure follow the SNFG nomenclature, details of which can be found at http://www.ncbi.nlm.nih.gov/books/NBK310273/).  1.4.4 Alternative glycan processing/trimming pathway Although glucose residues are essential for the recognition by glycan-processing enzymes, chaperones and lectins, their removal by ER processing glucosidases is indispensable for maturation of N-linked glycans. Formation of matured N-linked glycans has been detected in processing glucosidase deficient cells and cells treated with glucosidase inhibitors [88-90]. Spiro and coworkers later attributed this glucosidase-independent deglucosylation to Golgi endo α1,2 mannosidase [90, 91]. Endo α-1,2 mannosidase catalyses the hydrolysis of the α-1,2 mannosidic bond between the glucose-substituted mannose on the A-antenna and the remaining 15  oligosaccharide and releases Glc1-3Man1. Although the activity of endomannosidase has been detected in several tissue types [92, 93], it exhibits a cell type-specific expression  [94]. For example, endothelial cells, epithelia of the adrenal cortex and follicular cells lack the endomannosidase expression. In most cell types, the expression of endomannosidase cannot completely compensate for the conventional glycan processing.   1.5 Glycan heterogeneity Glycosylation is intrinsically heterogeneous. In contrast to nucleic acids and proteins, glycans are synthesised without a template. Moreover, glycosylation is a complex process involving many enzymes that are involved in assembly, transfer and processing of glycans. The intricate interplay between these enzymes, their levels of expression and presence of certain monomers may lead to mixed cell type-specific or developmental stage-specific glycosylation patterns. Glycan heterogeneity has roughly been classified into macro-heterogeneity (heterogeneity due to site occupancy) and micro-heterogeneity (heterogeneity due to glycan structure). It has been estimated that about 10-30% of the Asn-Xxx-Ser/Thr are not occupied by glycans [30]. Dol-P linked oligosaccharides appear to influence the association constant of polypeptide and OST by inducing conformational changes in the active site of OST [95]. The structure of the central amino acid in the Asn-Xxx-Ser/Thr consensus sequence also affects the occupancy rate [96]. When Xxx-position in the sequon of rabies virus glycoprotein was filled with proline, core glycosylation was completely blocked, whereas 5%, 19%, 24% and 43% of sequons were glycosylated when Xxx-position was filled with Trp, Asp, Glu and Leu respectively [97]. Moreover, efficient glycosylation was reported with the presence of hydroxyl groups as opposed to amide side chains in the region close to sequeon. The secondary structure of the polypeptide chain also affects the sequeon 16  occupancy [98, 99]. A survey of predicted secondary structures of glycoproteins indicates that about 70% of Asn-Xxx-Ser/Thr sequons are present in β-bends, 20% in β-sheets and 10% in α-helices [100]. Temporary conformations during translation or folding intermediates sometimes partially inhibit the transfer of LLO [101]. The presence of disulphide bonds in the proximity of the sequeon may also interfere with glycan transfer [102].  1.6 Congenital disorders of glycosylation Congenital disorders of glycosylation (CDG) is a term applied to a group of disorders caused by discrepancies in protein/lipid glycosylation. Since the first report in 1980, a total of 45 types of protein hypoglycosylation have been reported [103, 104]. CDGs are classified into two groups based on the biochemical pathway affected; type-I for the defects in N-linked glycan assembly and type-II for defects in N-linked glycan processing. CDGs are named depending on the type and a lower-case letter (a-z) for a subtype that indicates the chronological order of identification of the defective enzyme [105]. Among the 16 CDGs related to defects in N-glycosylation, 14 are CDG-I types, and 2 are CDG-II types [105]. Although all CDG-Is involve defects in glycosyltransferases that lead to impaired glycan assembly, phenotypic outcomes are not similar. All CDGs are autosomal recessive disorders. Complete knockout of a glycosyltransferase is lethal whereas mutations that lead to hypomorphic alleles show expression of a mutant enzyme that still retains some amount of activity. 1.7 Roles of glucose residues of N-linked glycans  Glucose residues are not a part of mature glycoproteins. Nonetheless, they play a key role in N-linked glycosylation. OST transfers N-linked glycans onto proteins and requires the presence of all three glucose residues on N-linked glycan [70]. Hence, the addition of the terminal α-1,2 17  glucose by Alg10p serves as a signal that the fully assembled LLO is ready to be transferred by OST. Removal of the terminal glucose by Glu-I frees the N-glycan from rebinding to the OST, and the unloaded OST catalyses the next LLO transfer and helps to maintain forward momentum in glycoprotein formation [106]. It has also been suggested that the glucose residues are critical for the regulation of the level of free N-linked glycan precursor as the OST has a hydrolytic activity that can release oligosaccharides from lipid [69, 107, 108]. Glucose residues also play an important role in protein quality control. Removal of the terminal glucose is essential for the binding of chaperone malectin and Glu-II, as described above [73]. Molecular chaperones Cnx and Crt cannot recognise tri- or di-glucosylated glycans. Hence, the synergistic actions of Glu-I and Glu-II to remove terminal glucose residues are essential for the regulation and quality control of N-glycosylation.   1.8 Processing α-Glucosidase-I  1.8.1 History After Neuberger et al. (1938) reported the presence of an oligosaccharide moiety on proteins, it took almost four decades to identify the structure of the oligosaccharide moiety and its processing reactions [27]. Spiro et al. (1976) proposed a partial structure of N-linked glycan using a radiolabelled study of the LLO isolated from thyroid [109]. Similar studies carried out on the LLO precursor of viral G-protein from vesicular stomatitis virus-infected Chinese hamster ovary (CHO) cells established the composition and structure of the oligosaccharide precursor [110]. Initially, the virus-infected cells were incubated with media containing radiolabelled mannose, glucosamine and galactose, and the corresponding labelled glycan structures were analysed by α-mannosidase 18  treatment and chromatography. They also observed the presence of “processing” intermediates Glc1Man9GlcNAc2 and Man9GlcNAc2, 20-30 minutes after the radiolabelling. After further structural characterisation of processing intermediates, they proposed that the “processing” steps involved removal of glucose units, as glucose was not present in the mature oligosaccharides [110]. Similar studies involving the comparison of LLOs, and oligosaccharides isolated from glycoproteins of hen oviduct membrane, established experimental evidence for glucose removal from the core oligosaccharide [111]. They also reported the presence of a membrane-bound glucosidase activity that specifically removed the glucose units with no significant degradation of the core oligosaccharide. Pulse labelling of vesicular stomatitis virus-infected HeLa cells with 3H labelled mannose and glucosamine established the view of co-translational glycan transfer [112]. Spiro and coworkers (1979) reported the occurrence of glucosidase activity in thyroid microsomes that specifically released glucose from both dolichol- and protein-linked oligosaccharides [113]. This microsomal glucosidase activity showed unusual substrate specificity, low Km value (0.7 mM) for LLO, and an optimum neutral pH. However, this study was not able to identify the α-glycosidic linkage of the glucose residues and reported that microsomal glucosidase acted on the α-1,3-linkage. Later Atkinson (1978) attributed this glucosidase activity to the ER membrane [114]. Subsequent studies involving the use of radiolabelled Glc3Man9GlcNAc2, N-linked glycan substrates and the glucosidase inhibitor 1-deoxynojirimycin, established the presence of two different glucosidase activities in the ER and termed them processing α-glucosidase-I and processing α-glucosidase-II [72, 111, 115]. Based on the cDNA sequence, Shailubhai et al. (1991) proposed Glu-I as a single polypeptide chain of 85 kDa with two adjoining domains: a membrane-bound domain that anchors the protein to the ER and a luminal domain [116]. SignalP prediction of the signal peptide cleavage site and N-terminal sequencing of soluble Glu-I isolated from yeast 19  cells established the domain organisation of the Glu-I [11]. Glu-I is a type II membrane protein, a class of proteins that has a lumenal C-terminal domain and an anchoring signal sequence in the ER membrane.  It contains a short N-terminal cytosolic tail (residues 1-10), a single transmembrane domain (residues 11-28), and a vast luminal C-terminal catalytic domain (residues 28-833) as depicted in (Figure 1.6).    Figure 1.6 Cartoon depiction of Glu-I domains (not to scale). Glu-I consists of a cytosolic tail (Yellow), a short transmembrane domain (Brown) and a large ER lumenal domain (Blue). Putative glycosylation sites, identified using NetNGlyc 1.0 server (http://www.cbs.dtu.dk/services/NetNGlyc/), are indicated with hexagons (number indicates the amino acid position).  20  1.9 Classification and Glu-I orthologs Glycoside hydrolases are classified based on their amino acid sequence similarities into 135 families on the Carbohydrate-Active enZymes (CAZy) database [117]. This classification assigns Glu-I orthologs to the GH family 63 and GH-G clan of (α/α)6-barrel catalytic domain containing glycoside hydrolases (http://www.cazypedia.org/index.php/Glycoside_Hydrolase_Family_63).  This GH family 63 was further classified into three subfamilies based on their substrate-specificity: 1. Processing α-glucosidase-I subfamily, 2. MGH subfamily, and 3. GGalase subfamily. This GH family 63 has a total of 1074 entries [118].  However, only 19 are characterised, and three structures have been reported to date. Based on reaction and type of substrate, the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NC-IUBMB) assigned the Enzyme Commission (EC) number 3.2.1.106 to Glu-I and other enzymes that hydrolyse the glycosidic bond to remove the terminal O-linked glucose from the substrate mannosyl-oligosaccharide Glc3Man9GlcNAc2.  Glu-I is present in several eukaryotes [72, 113, 119, 120]. Among these eukaryotic enzymes, yeast Glu-I has been well studied [121]. Fungal Glu-I is commonly known as Cwh41p and “processing α-glucosidase-I” whereas mammalian enzyme is often referred to as “MOGS” (mannosyl-oligosaccharide glucosidase). Surprisingly, bacteria and archaea which lack the N-linked oligosaccharides possess genes encoding proteins that are homologous to Glu-I [122, 123]. However, the biological function of these Glu-I homologs has not been precisely defined. In Escherichia coli, the YGJK gene encodes a glucosidase that shares 21% identity to the catalytic C-terminal domain of Cwh41p [124]. A Glu-I homologue has also been reported from Thermus thermophilus HB27 and Rubrobacter radiotolerans RSPS-4. However, these enzymes have a substrate preference for α-D-21  mannopyranosyl-1,2-D-glycerate (mannosylglycerate) and α-D-glucopyranosyl-1,2-D-glycerate (glucosylglycerate), respectively [125]. 1.10 Purification of Glu-I  Glu-I was first identified in crude preparations of calf thyroid microsomes [126]. Kilker et al. (1981) observed Glu-I activity in both particulate and cell-free extract of the yeast Saccharomyces cerevisiae X2180 homogenate [2]. Glu-I was partially purified from cell-free extract using ammonium sulphate precipitation followed by an anion exchange chromatography composed of DEAE-Sephadex resin. A  concanavalin A-Sepharose chromatography resin was also developed to harness the lectin property for binding of Glu-I as a glycoprotein [127]. With the information available from these initial studies of Glu-I isolation and characterisation, it was established that deoxynojirimycin acts as a Glu-I inhibitor [127]. Leveraging on the Glu-I inhibitor identification, affinity resins with deoxynojirimycin or N-(5-carboxypentyl)-1-deoxynojirimycin were prepared and used for Glu-I purification [72]. Glu-I from calf liver microsomes was purified using N-(5-carboxypentyl-1-deoxynojirimycin resin. This enzyme was reported to be a tetramer with a molecular weight between 320-350 kDa. This purified Glu-I showed optimum activity at pH 6.2 [72]. Conversely, Glu-I purified from yeast microsomal preparations by DEAE-Sephacryl gel-filtration chromatography, affinity chromatography on AH-Sepharose 4B-linked N-(5-carboxypentyl)-1-deoxynojirimycin and concanavalin A-Sepharose chromatography had a molecular mass of 95 kDa and was glycosylated [128]. Treatment of mung bean microsomal particles with 1.5% Triton X-100 released Glu-I activity. Glu-I isolated from the microsomal fraction of mung bean seedlings was a glycoprotein with one oligomannose-type oligosaccharide and a molecular weight of 97 kDa on SDS-PAGE gels [129]. Chromatography of the mung bean 22  soluble fraction on hydroxylapatite, Sephadex G-200, dextran-Sepharose and Concanavalin A-Sepharose resulted in 200-fold purification of Glu-I activity [130]. Shailubhai et al. (1987) purified Glu-I from lactating bovine mammary tissue [131]. Glu-I, purified from solubilised microsomal enzyme fraction using affinity chromatography on Affi-Gel 102 with N-(5-carboxypentyl)-1-deoxynojirimycin as ligand and DEAE-Sepharose CL-6B chromatography, showed a molecular mass of 85 kDa under reducing conditions, whereas molecular weight analyses of native enzyme using gel filtration indicated a molecular mass of 320-330 kDa, suggesting that the native enzyme was a tetrameric protein [131]. Brain tissue, like other tissues, was found to contain Glu-I. Tulsiani et al. (1990) purified a Glu-I from rat brain, and the biochemical properties were similar to the enzyme from other mammalian tissues [132]. With the advent of molecular biology techniques to manipulate genes and the availability of different hosts and expression vectors that can express a fused protein with affinity tags, there have been several reports thereafter that involved purification of recombinant Glu-I [11, 13, 119, 120, 133].   1.11 Biochemical properties Glu-I has been reported from several sources ranging from unicellular yeast to highly evolved organisms such as mammals [72, 128]. Enzymes from these diverse sources share several similar biochemical properties. Glu-I is generally made up of a single glycosylated polypeptide chain with a molecular weight ranging from 85-98 kDa  [11, 116]. The exception is Glu-I isolated from calf liver and bovine mammary gland, which were reported to be oligomers [134]. However, not all the Glu-I studied were tested for oligo-/multimer properties [72, 131]. The optimum pH of all Glu-I orthologs falls in the neutral range which distinguishes them from lysosomal glucosidases which have acidic optimum pH [128, 129, 131]. Glu-I does not depend on bivalent cations for its function 23  [72, 128]. The Km of yeast Glu-I is reported as 1.28 mM and 1.26 mM with Glc3(CH2)8OCH3 and Glc3(Man)4OCH3 respectively [133, 135]. However, Glu-I shows high affinity toward its native substrate, Glc3Man9GlcNAc2. Glu-I from A. brasiliensis exhibited a Km value of 6.1 µM when incubated with Glc3Man9GlcNAc2-(2-aminopyrimidine) [120]. C. albicans Glu-I exhibited a Km of 133 µM with 4-methylumbelliferyl--D- glucopyranoside [119].  1.12 Molecular genetics of Glu-I Ram et al. (1994), while carrying out a mutant screen to identify cell wall assembly genes in yeast S. cerevisiae, obtained a Glu-I recessive mutant in which the three glucose residues were retained on N-linked oligosaccharides of glycoproteins [136]. This mutant displayed calcofluor white hypersensitivity (CWH), showed partial resistance to the K1 killer toxin, and had reduced levels of cell wall β-1,6-glucan. Only the cells that contained the CWH41 gene released glucose from the synthetic substrate Glc3(CH2)8OCH3 while cwh41 null mutants did not [10, 136]. Moreover, comparison of the product of the CWH41 gene with human Glu-I indicated that the proteins shared significant amino acid sequence similarity. Also, a comparison of human Glu-I to the complete yeast genome showed that Cwh41p is the only gene product with such similarity; therefore, they confirmed that the CWH41 gene encoded yeast Glu-I. Kalz-Füller et al. (1995) developed PCR generated oligonucleotide probes based on the amino acid sequence of the peptides generated by the trypsin hydrolysis of pig liver Glu-I [137]. Using these oligonucleotide probes, they identified a full length human Glu-I cDNA (2881 bp) from a human hippocampus cDNA library and overexpressed it in transfected COS1 cells [137].  A similar study carried out using a mouse cDNA library resulted in the identification of a major transcript of about 3.1 Kb on a northern blot [138]. 24  A putative promoter of Glu-I was also isolated from the 5’ upstream region generated through primer extension. 1.13 Overexpression of Glu-I Kalz-Fuller et al. (1995) reported the overexpression of Glu-I cDNA isolated from human hippocampus in COS1 cells [137]. They observed a four-fold improvement in expression of Glu-I, and the enzyme was found to be a type-II membrane-bound protein [137]. Dhanawansa et al. (2002) achieved a 28-fold increase in Glu-I activity when the CWH41 gene was expressed using an episomal pHVX2 vector in S. cerevisiae [11]. Most of the expressed enzyme was soluble whereas only 10% of total Glu-I activity was associated with the membrane-bound form compared to 67% membrane-bound Glu-I activity in yeast cells that were transformed with a control pHVX2 vector [11]. The recombinant protein was glycosylated with a molecular weight of 98 kDa on SDS-PAGE. Faridmoayer and Scaman (2007) reported overexpression of the truncated form of Cwh41p that lacked 34 residues at the N-terminus (Cwh41Δ1-34p) in S. cerevisiae as a catalytically active and soluble fragment. Cwh41Δ1-34p was overexpressed 4-fold over the recombinant expression of full-length Cwh41p [12]. Barker et al. (2011) reported a yield of 4.2 mg of Glu-I per litre of culture when expressed in Pichia pastoris using the pPICZalphaB vector [133]. Expression of truncations (Cwh41Δ1-319p, Cwh41Δ1-525p) was not successful in either S. cerevisiae and P. pastoris hosts [12, 133]. In contrast, Frade-Perez et al. (2010) reported the expression of a truncated version of Candida albicans Glu-I (CaCwh41p) lacking 419 N-terminal amino acids as a His-tagged fusion protein in Escherichia coli. This Cwh41p was atypical as it showed activity against 4-methyl-umbelliferyl-α-glucoside and sensitivity to N-ethyl maleimide [119]. A variant 25  of Glu-I from A. brasiliensis that lacked 16 N-terminal amino acids was cloned and overexpressed with a glutathione S-transferase tag (GST)  [120]. 1.14 Glu-I inhibitors Glycosidase inhibitors have played a significant role in our current understanding of N-glycosylation [139]. Inhibition of processing glycosidases severely affects the maturation of various glycoproteins that interact with the Cnx/Crt pathway. Two inhibitors most frequently used to block Glu-I activity are castanospermine, which selectively inhibits Glu-I, and deoxynojirimycin, which inhibits both Glu-I and Glu-II [127, 140] [Table 1.1]. Bause et al. (1989) observed substantial inhibition of Glu-I from pig liver when incubated with 1-deoxynojirimycin (Ki≈2.1 µM), N,N-dimethyl-1-deoxynojirimycin (Ki≈0.5 µM) and N-(5-carboxypenty1)-1-deoxynojirimycin (Ki ≈ 0.45 pM) [141].  Inhibition of Glu-I has been observed to have adverse biological effects that are glycoprotein and cell-type specific. Castanospermine prevented glycoprotein processing when administered in the culture of various animal cells, and caused the production of glycoproteins with Glc3Man9GlcNAc2 type glycans [142]. Free oligosaccharide trafficking was severely perturbed in castanospermine-treated HepG2 cells. Castanospermine-treated cells showed only 20% of the total free oligosaccharides secreted into the extracellular matrix [143]. Interestingly, Glu-I inhibitors showed a drastic effect on viral glycoproteins as viruses depend on their host machinery for protein synthesis and CPTM. Viruses are known to use the host glycosylation machinery to modify their capsid proteins and depend on the Cnx/Crt pathway for protein folding. Targeting the ER alpha-glucosidases at a low level could interrupt the folding of these viral glycoproteins, and could be a potential therapeutic tool in treating viral infections, without affecting the viability of host cells 26  [144]. Several studies have reported the impairment in the assembly of virions due to failed protein folding in the presence of Glu-I inhibitors. N-butyl deoxynojirimycin suppressed the secretion of human hepatitis B viral (HBV) particles from HBV-transfected HepG cells [145]. Castanospermine and deoxynojirimycin strongly suppressed the folding of dengue viral envelope proteins in mouse neuroblastoma cells [146].  Table 1.1. Inhibitors of processing alpha-glucosidase I Source of enzyme Inhibitor Ki (µM) IC50 (µM) Reference S. cerevisiae  (microsomal) 1-Deoxynojirimycin 16 - [128] N-Methyl-deoxynojirimycin 0.3 - N-(5-Carboxypentyl)-1- deoxynojirimycin 3 - Kojibiose 55 - S. cerevisiae (soluble) Kojibiose  800 - [11] Miglitol - 22 [147] A. brasiliensis Kojibiose - 20  [120] 1-Deoxynojirimycin - 20 Nigerose - 1900 Pig liver 1-Deoxynojirimycin 2.1 - [141] N-Methyl-deoxynojirimycin 0.5 - N-(5-Carboxypentyl)-1- deoxynojirimycin 0.45 - N-7-Oxadecyl-deoxynojirimycin - 0.28 [148] ManNH2 alpha 1,2Glc - 15.7 Pig kidney Castanospermine - 0.12 [149] 6-O-Butanoyl castanospermine - 20 1-Deoxynojirimycin - 3.49 Calf Liver 1-Deoxynojirimycin  3 - [72] N-Methyl-deoxynojirimycin 0.3 - N-(5-Carboxypentyl)-1- deoxynojirimycin 8 - Mung bean seedlings Castanospermine - 1 [129] 27  1.15 Substrate specificity Glc3Man9GlcNac2 is the natural substrate of Glu-I which catalyses the removal of α1,2-linked glucose from both protein-bound and free oligosaccharides. The minimal substrate requirement of Glu-I is the tri-glucoside α-Glc1,2-α-Glc1,3-α-Glc  [135].  Glu-I is highly specific for the α 1,2 linkage in the tri-glucosylated oligosaccharide, and it does not hydrolyse the α1,2 linkage in kojibiose which is a disaccharide composed of α 1,2 glucoses  [141]; instead, kojibiose acts as an inhibitor of Glu-I activity. Mammalian and yeast Glu-I do not hydrolyse simple sugar substrates such as p-nitrophenyl α-glucoside and 4-methyl umbelliferyl α-glucoside. Glucotriose, the minimal substrate required for Glu-I activity, has an unusual combination of glycosidic bonds. The central glucose is connected to adjacent glucose residues with C1 and C2. This gives the substrate a unique conformation. Recently, based on analysis of substrate binding modes via docking of yeast Glu-I and its substrate glucotriose, Barker and Rose (2013) proposed that the intra-saccharide interaction between Glc1 and Glc3 impart a unique conformation to glucotriose [147]. This conformation of the trisaccharide is vital for the interaction with the active-site and may contribute to the substrate specificity of Glu-I [147].  In contrast to mammalian and yeast Glu-I, the E. coli Glu-I ortholog Ygjk shows relaxed substrate specificity while sharing common structural features such as the (α/α)6-barrel [124]. Ygjk preferentially hydrolyses α1,3 linkage of nigerose but also acts on trehalose, kojibiose and maltooligosaccharides.  N-glycosylation has not been established in E. coli, and the biological function of Ygjk has not been identified yet.  28  1.16 Catalytic domain of Glu-I Trypsin hydrolysis of Glu-I (85 kDa) isolated from pig liver generated a major 69 kDa peptide and three minor 60 kDa, 45 kDa and 29 kDa peptides within 10 min of incubation [141]. Further extension of incubation time to 40 min led to complete digestion of the 85 kDa protein and formed a mixture that retained 60% of initial activity and had a 29 kDa fragment as the major species. However, the authors could not attribute the retained activity to any single digested product as efforts to separate the products were not successful [141]. The 29 kDa fragment was the major stable fragment even after the extension of trypsin digestion time to >2 h suggesting that the catalytic domain is resistant to proteolysis. Further analysis of the 29 kDa fragment for the presence of N-linked glycans revealed that this fragment was non-glycosylated  [141]. In another study, Shailubhai et al. (1991) observed the release of enzymatically active 39 kDa fragment from the ER membranes following controlled trypsinisation in the presence of saponin  [116]. This 39 kDa fragment could bind to a Glu-I specific affinity resin (CP-DNJM), indicating the presence of a putative catalytic domain. A similar trypsinisation pattern of a 39 kDa fragment was observed in Glu-I from mouse, rat, guinea pig, bovine mammary glands and sheep liver [150]. To gain insight into the architecture of the active site of the enzyme, Romaniouk et al. (2004) synthesised a photoactive derivative of DNJM- 4-(ρ-azidosalicylamido)butyl-5-amido-pentyl-1-DNM . Photo-labeling of Glu-I followed by digestion of the labelled protein with V8 protease resulted in a 24 kDa labelled peptide [151]. Consistent with previous reports, yeast Glu-I also generated a 37 kDa fragment upon trypsin digestion [152]. N-terminal sequencing of the 37 kDa fragment revealed the hydrolysis site between K524 and R525 with the presence of the catalytic activity in the C-terminus. Surprisingly, this 37 kDa was 1.9 times more active than the undigested 98 kDa enzyme. 29  However, overexpression of this catalytic domain of yeast Glu-I lacking the N-terminal 524 residues was not successful in either S. cerevisiae or P. pastoris [12, 133].  1.17 Catalytic residues and mechanism 1.17.1 Glu-I is an inverting glycosidase Hydrolysis of a glycosidic bond typically takes place via general acid catalysis and requires two carboxyl residues (with a few exceptions of non-carboxylate catalytic residues in α1,2 fucosidase, sialidase and trans-sialidases) [153]. Hydrolysis of a glycosidic bond occurs through either net inversion or retention of anomeric configuration  [154]. Inverting glycoside hydrolysis occurs via a one step, single-displacement mechanism involving an oxocarbenium-ion-like transition state (Figure 1.7A), whereas retaining hydrolysis occurs through a two-step, double displacement mechanism involving a covalent glycosyl-glycosidase intermediate (Figure 1.7B). Both mechanisms typically involve glutamic or aspartic acid residues that are located 6-11 Å apart in inverting glycosidases and 5.5 Å apart in the case of retaining glycosidases. 1H-NMR spectroscopic studies of yeast and mammalian Glu-I identified Glu-I as an inverting glucosidase [155]. 30  A.  B.   Figure 1.7. Mechanism of glycoside hydrolase. A. Inverting glycosidase B. Retaining glycosidase. (Adapted from Zechel and Withers (2000) with permission. Copyright © 2000, American Chemical Society).  31  1.17.2 Genetic mutants Various studies have reported the loss of activity due to mutations in Glu-I. In humans, the compound heterozygosity involving two missense mutations (R486T and F652L) inactivated Glu-I, which was diagnosed as CDGIIb in an infant and resulted in death at the age of 74 days [8]. In yeast, G725R caused the loss of Glu-I activity and eventually led to ER stress [156]. A single point mutation involving the substitution of serine to phenylalanine at 321st position led to the inactivation of Glu-I in Lec23 CHO cells [157]. All these mutations reside in the C-terminal domain that contains the catalytic site. In contrast, Sadat et al. (2014) recently reported failed Glu-I expression in two children with compound heterozygotes carrying a nonsense mutation Q124X and two missense mutations A22E and R110H. As these missense mutations reside in the non-catalytic region, they attributed the lack of expression to altered splicing due to the mutations [158]. 1.17.3 Chemical modification studies of Glu-I Chemical modification of amino acids often provides clues about the catalytic properties of enzymes. Selective chemical modification of arginine, cysteine, tryptophan and tyrosine inactivated Glu-I from yeast and mammalian sources, and the activity was protected in the presence of deoxynojirimycin [119, 150, 152, 159]. This indicates that these amino acids may be essential for either catalytic activity, substrate binding or conformation. Pukazhenthi et al. (1993) reported the presence of a cysteine residue in/near the active site of bovine mammary gland Glu-I [150]. Faridmoayer and Scaman (2005) reported that the binding residues in yeast Glu-I are different from mammalian Glu-I based on site-specific chemical modification studies. A soluble form of yeast Glu-I was sensitive to site-specific chemical modification of histidine and tyrosine by diethylpyrocarbonate (DEPC) and tetranitromethane (TNM) and lost about 90% of its activity. 32  However, dialysis restored its function after the chemical modification was carried out on Glu-I that was pretreated with deoxynojirimycin. A similar protective effect of deoxynojirimycin was not observed when tryptophan residues were chemically modified with N-bromosuccinimide (NBS) [152]. In contrast to Glu-I from S. cerevisiae and mung bean, Glu-I from C. albicans was sensitive to N-ethylmaleimide (NEM) and completely insensitive to diethyl pyrocarbonate (DEPC) [119].  Based on the amino acid modifications studies of mammalian Glu-I, a highly conserved region between E594RHLDLRCW602 was identified and proposed to be important for substrate-binding [159].   1.17.4 Site-directed mutagenesis studies Chemical modification studies alone are not sufficient to establish the substrate binding and catalytic mechanism of an enzyme. Amino acid comparison and identification of conserved residues can provide some clues about the potential catalytic residues. Sequence homology of Glu-I orthologs revealed six highly conserved carboxylic residues (D601, D602, E613, D617, D670, and E804 of Cwh41p) [12] with only two of these carboxylic residues (E613 and D617) residing in the substrate binding region proposed by Romaniuk [159]. Site-directed mutagenesis of E613 and D617 to alanine in yeast Glu-I inactivated the enzyme  [12]. However, it was not possible to conclude whether the loss of function was due to conformational changes or disruption of substrate binding or catalysis. In a similar study carried out on the E. coli homolog of Glu-I, Ygjk, based on the structural similarities to glucoamylase (GH15) and chitobiose phosphorylase (GH65), Kurakata et al. (2008) proposed D501 and E727 as the catalytic residues [124]. Barker and Rose (2013) proposed D568 and E771 of Cwh41∆1-34p (corresponding to D601 and E804 in Cwh41p) as the catalytic residues after an investigation of the crystal structure of yeast Glu-I [147].  33   Figure 1.8. Structures of GH family 63 enzymes. A. S. cerevisiae Cwh41Δ1-34p truncated form that lacks 33 residues at N-terminus expressed in P. pastoris (4J5T) B. E. coli Ygjk (3D3I). C. Thermus thermophilus mannosylglycerate hydrolase Tt8MGH (4WVB). N-terminal domain is highlighted with the blue box. S. cerevisiae structure also contained two N-linked glycan chains. UCSF-Chimera was used to make these pictures from the PDB files.  1.17.5 Structural insights Although there are 1074 entries in GH family 63 in CAZy database, only a handful of proteins have been characterized to date. Among these only three structures: two bacterial (Ygjk and Tt8mgh) and one yeast Glu-I (Cwh41Δ1-34p) have been solved so far for the GH family 63 (Figure 1.8).  All three proteins share structural similarities and contain an (α/α)6-barrel [118, 124, 147]. Soluble Glu-I from yeast consists of two domains - the N-domain and C-terminal domain (C and C’ domain) joined by a linker helix (Figure 3.1). The C-terminal domain is (α/α)6-toroid and contains the active site [147]. Both Cwh41p and Ygjk possess the N-terminal beta-sandwich domains in 34  addition to their (α/α)6-barrel domains, and conformation of these N-terminal beta-sandwich resembles a family of carbohydrate binding molecules [147]. Conversely, Tt8mgh from Thermus thermophilus HB8 consists of a single (α/α)6-barrel catalytic domain with two additional helices and two long loops which form a homo-trimer [118].  Glucose triose is the minimum substrate cleaved by Glu-I [135]. Barker and Rose (2013) used in silco docking studies of (Glc(α1,2)Glc(α1,3)Glc(α1,3)-O-Methyl) in the binding site of yeast Glu-I and proposed a substrate binding model.  Based on this model and the high specificity of Glu-I for the α1,2 glycosidic bond, they suggested that the unique non-linear bent-back conformation of the substrate is important for the interaction with the active site. Moreover, an intra-chain stacking interaction between glucose 1 and 3 and stacking with an aromatic chain of Tyr742 further suggested this as the possible substrate binding model [147].  1.18 The physiological significance of Glu-I function Glu-I catalyses the removal of terminal glucose from N-linked glycans of glycoproteins which are important for various cellular processes. This reaction is critical for the maturation of glycoproteins as altered glycosylation states result in serious pathological conditions. In humans, loss of Glu-I leads to a severe phenotype defined as CDG IIb. The occurrence of CDG IIb is very rare, and it was first observed in a neonate with severe generalised hypotonia and dysmorphic features [8]. The clinical condition was progressive and characterised by the occurrence of hepatomegaly, hypoventilation, feeding problems, seizures, and was fatal at 74 days. The presence of the tetrasaccharide Glc(α1,2)Glc(α1,3)Glc(α1,3)Man in the patient’s urine, lack of Glu-I activity in liver tissue and cultured fibroblasts, and increased levels of endo-α1,2-mannosidase activity were attributed to the bypass of glycan processing and quality control pathways and involvement of the 35  alternative glycan processing in Golgi [160]. Sadat et al. (2014) reported severe hypogammaglobulinemia and complex developmental disorders in two siblings that are compound heterozygotes to nonsense and two missense mutations in Glu-I. Surprisingly, despite the shortened immunoglobulin half-life, these patients showed a decreased susceptibility to viral infections and have so far survived longer than the affected individual previously reported [158]. The effects of a lack of Glu-I have also been observed in culture cell-lines. Lack of Glu-I in vesicular stomatitis virus-infected Lec23 CHO cells led to the accumulation of triglucosylated oligosaccharide (Glc3Man9GlcNAc2); similar results were also observed in vesicular stomatitis virus-infected cells treated with Glu-I inhibitors castanospermine and deoxynojirimycin [161]. Knockdown of Caenorhabditis elegans Glu-I gene using RNAi led to the accumulation of free oligosaccharides, increased ER stress, and reduced life-span of mutant worms to half, despite a visibly similar appearance to wild-type worms [7]. As previously mentioned, yeast mutants generated by disruption of the CWH41 gene had a 50% reduction of β1,6 glucan levels in the cell walls, hypersensitivity to calcofluor white, and resistance to K1 killer toxin [136]. In C. albicans, the Glu-I disruption caused diminished growth rate apart from cell wall defects, in contrast to the effects reported for S. cerevisiae [4].  Glu-I is required during embryo development in Arabidopsis thaliana. Mutants that lack Glu-I activity showed decreased levels of proteins in seeds and absence of typical protein bodies [6]. Disruption of the KNF gene, that encodes Glu-I in A. thaliana, altered the shape of embryos but not the pattern of the embryo and seedling [162]. Later, Furumizu and Komeda (2008) identified a hypomorphic allele of KNF which exerted a similar phenotype to the knf-null mutants [163]. Expression of KNF is also reported to be essential for the synthesis of cellulose in Arabidopsis [164]. Analysis of the cDNA library of Gossypium hirsutum fibre secondary wall also revealed the 36  presence of two expressed sequence tags of Glu-I and differential expression during fibre secondary wall cellulose biogenesis [165]. Disruption of Glu-I in rice exhibited severe defects in root cell division and elongation resulting in a short-root phenotype [166]. The growth of leaves in Raphanus sativus seedlings was diminished by glucose trimming inhibitors [167].  1.19 α-Glucosidase-I and ambiguity in identification The term α-glucosidase is a generic name for α-glucoside hydrolases that include both “processing” and “non-processing” α-glucosidases that differ significantly in their properties. IUBMB recommends the use of mannosyl-oligosaccharide glucosidase for “processing α-glucosidase-I” (EC 3.2.1.106) and “α-glucosidase” (EC 3.2.1.20) for “non-processing” exo-α1,4-glucosidases. Despite significant differences in molecular and biochemical properties, some studies have mistakenly compared processing enzymes with the non-processing α-glucosidases. Odaci et al. (2010) compared the pH of α-glucosidase-I maltase with processing α-glucosidase-I [168]. Kaewmuangmoon and Chanchao (2013) related honey bee α-glucosidase to processing α-glucosidase-I [169]. Similarly, Li et al. (2015) and Hu et al. (2015) placed the processing α-glucosidase-I (EC 3.2.1.106) in the α-glucosidase (EC 3.2.1.20) class [170, 171].      37  1.20 Body of Thesis 1.20.1 Lacuna Many aspects of Glu-I are intriguing, including its strict substrate specificity and, its crucial role in the complex N-glycosylation pathway. From the studies on natural mutations of Glu-I, it is evident that this enzyme is essential for the development and survival of eukaryotes [7, 8, 158, 167, 172]. Despite its physiological significance, details about its catalytic mechanism are not fully known yet owing to the scarcity of methods to produce the Glu-I and lack of commercially available substrates. Overcoming these obstacles and advancing our understanding of Glu-I structure-function and mechanism are necessary steps toward fully understanding N-linked glycosylation, and toward dealing with various disease states associated with it.  The structure-function details are immensely helpful in understanding the aspects of enzyme inhibition and in designing effective mechanism-based inhibitors. Moreover, in addition to structure-function information, the establishment of methods to generate the enzyme and the substrate would greatly help in the in-vitro screening of potential inhibitors specific for Glu-I.  1.20.2 Overall hypothesis Soluble Glu-I from yeast can be used as a model to study the structure and function of a subfamily of family 63 glycoside hydrolases that include human Glu-I and to further our understanding of N-glycosylation.  1.20.3 Overall objectives To establish a robust expression and purification procedure for Glu-I and its truncations. To investigate the role of N-terminus in Glu-I function and study the catalytic mechanism and structure of Glu-I.  38  Chapter 2. Recombinant expression and characterisation of Glu-I Hypothesis- Glu-I can be expressed in a robust prokaryotic expression system Rationale- As mentioned in the introduction, Glu-I exists in both membrane-bound and soluble forms. The soluble form of Glu-I is non-glycosylated, and this circumvents the requirement of complex post-translational modifications [12]. This property can be leveraged to produce large quantities in fast-growing prokaryotic expression systems such as E. coli, with the aid of codon optimisation tools to allow efficient expression of eukaryotic proteins in a prokaryotic system. Objective #1- To carry out heterologous expression of Glu-I in E. coli, its purification and activity assay Objective #2-To carry out codon optimisation of Glu-I and expression of codon optimised Glu-I in E. coli  Chapter 3. Heterologous expression of yeast Glu-I truncations in E. coli and isolation of catalytic domain of Glu-I  Hypothesis- the N-terminal domain of Glu-I plays two significant roles of assistance in acquiring active conformation/folding and a catalytic regulatory function Rationale- The catalytic domain of Glu-I lies between residues 525 to 833, and when separated from the full enzyme, has higher activity compared to the whole enzyme [12, 152]. This intriguing increase in activity raises a question about the role of the N-terminal domain that has not been well studied. Development of methods for the isolation and characterisation of the catalytic and N-terminal domains would advance our understanding of Glu-I.   Objective# 1- To establish a method for the expression and purification of Glu-I truncations. Objective #2- To determine the effect of solubility enhancing tags and molecular chaperones on Glu-I truncations expression. 39  Objective #3- To obtain co-expression of N- and C-terminal domains of Glu-I. Objective #4- To isolate the catalytic domain of Glu-I.  Chapter 4. Identification of the catalytic base of processing α-glucosidase-I  Hypothesis- A pair of residues from the highly conserved carboxylic residues act as the catalytic pair. Rationale- Glu-I is an inverting glycoside hydrolase and catalyses the hydrolysis of α1,2-glycosidic bond via an inverting mechanism [155]. This hydrolysis reaction follows general acid-base catalysis that involves two carboxylic residues [173]. Glu-I has six highly conserved carboxylic amino acids [12]. Site-directed mutagenesis of these conserved residues and with the help of nucleophilic rescue studies can be used to confirm the catalytic pair.  Objective #1- To carry out site-directed mutagenesis of conserved carboxylic residues of Glu-I, followed by expression and purification of mutant proteins.  Objective #2- To characterise mutants and evaluate nucleophile rescue to identify the general acid/base.    40  Chapter 2: Recombinant expression and characterisation of Glu-I   41  2.1 Summary Processing α-glucosidase-I catalyses the first reaction in N-glycan processing by removing the distal α1,2-linked glucose in Glc3Man9GlcNAc2 of nascent proteins. This removal is followed by a series of other processing steps and binding of chaperones that are crucial for folding of the nascent protein. Despite its critical role in N-glycosylation and quality control of protein folding, many aspects of Glu-I function are still unknown. Lack of robust methods to produce and purify large quantities of Glu-I, and limited availability of an alternative substrate for functional assays impede biochemical, mechanistic and structural studies. To overcome these obstacles, I have developed a robust method for overexpression and purification of Glu-I. The soluble form of Glu-I (Cwh41pΔ1-34) from Saccharomyces cerevisiae has been cloned into pET30a(+) and overexpressed in Escherichia coli. Optimal expression of the 98 kDa form was attained with 1 mM IPTG concentration and induction at 25 °C. Recombinant Glu-I was obtained at 95% purity and specific activity of 3370 U/mg protein using a two-step procedure involving Ni-NTA affinity chromatography and size exclusion chromatography (SEC). The trisaccharide is the minimal requirement for assay of Glu-I activity. This avoids the requirement of complicated synthetic steps required to add extra sugars beyond the two α-oriented glycosides. Therefore, a simple substrate- α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3 was used for Glu-I activity assays. Synthesis of this trisaccharide was carried out by our collaborators- Dr. Todd Lowary and Dr. Akihiro Imamura at the University of Alberta, via a linear synthetic strategy, starting from commercially available methyl α-D-glucopyranoside to obtain α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3 with a final yield of 72%. Recombinant soluble Glu-I showed a Km of 1.27 mM with this synthetic trisaccharide. 42  2.2 Introduction Processing α-glucosidase-I (Glu-I) plays a critical role in N-linked protein glycosylation by initiating the processing of the N-linked glycan precursor Glc3Man9GlcNAc2 after the co-translational en bloc transfer of this pre-assembled 14-mer from the diphospho-dolichol precursor to an asparagine residue (Asn) in the consensus sequence (Asn-Xxx-Thr/Ser) of a nascent protein. Glu-I catalyses removal of the terminal α1,2-linked glucose residue and this removal enables processing α-glucosidase-II (Glu-II) to remove the next glucose residue to form the monoglucosylated protein that subsequently enters the calnexin and calreticulin pathway [3]. Glu-I (EC 3.2.1.106) is a type-II membrane protein bound to the ER inner membrane [137]. In Saccharomyces cerevisiae, Glu-I consists of 833 amino acids with a molecular weight of 98 kDa and encoded by the CWH41 gene [10]. It was predicted to have a short N-terminal cytoplasmic tail (amino acid residues 1-10), a transmembrane region (amino acid residues 11-28) and a major lumenal C-terminal domain (amino acid residues 29-833) [12]. The SignalP prediction tool, developed by Nielson et al. (1997), predicts a signal peptide cleavage site between residues Ala 24 and Thr 25 of the transmembrane sequence of the membrane-bound form of the yeast enzyme. Endogenous proteolytic cleavage at this site releases the soluble lumenal domain [11]. Dhanawansa et al. (2002) reported overexpression of membrane-bound Cwh41p and obtained one microgram of Glu-I per gram of wet yeast biomass after a multi-step purification involving ammonium sulphate precipitation, anion exchange, concanavalin A, and gel filtration chromatography. The soluble form of Glu-I that lacks the N-terminal 34 residues (Cwh41Δ1-34p) in S. cerevisiae yielded 45 µg of Glu-I per gram of wet yeast biomass when expressed in S. cerevisiae as a six-histidine (6xHis)-tagged protein and purified using affinity chromatography [11, 12]. Barker et al. (2011) observed an increase in expression of Cwh41Δ1-34p when 43  heterologously expressed in Pichia pastoris and obtained 4.2 mg of Glu-I per litre of the culture [133]. These methods yielded relatively low amounts of enzyme and involved multi-step purification [11, 12], or required long induction times (90 hr) for expression [133].  The soluble form of Glu-I has five putative N-glycosylation sites (Asn 42, 122, 135, 787 and 805). Asn 42 and 122 were glycosylated in soluble Glu-I (Cwh41Δ1-34p) when heterologously expressed in P. pastoris [147], whereas none of these sites was glycosylated when expressed in S. cerevisiae [12]. The fate of glycosylation of Cwh41Δ1-34p has no significant effect on function, as both the glycosylated and non-glycosylated Cwh41Δ1-34p showed similar catalytic properties with the synthetic substrate (Glc)3-(CH2)8-O-CH3. The lack of functional dependence on glycosylation can be harnessed in combination with robust prokaryotic expression systems such as Escherichia coli to develop a rapid method to attain high expression levels.  The deglucosylation steps catalysed by Glu-I and Glu-II are essential for the processing and maturation of N-linked glycans and subsequent folding of glycoproteins. The importance of Glu-I in glycan processing in fungi [5, 175], Caenorhabditis elegans [7], plants [164], and humans [8] is well documented. Impairments in the N-glycan processing alter the composition of glycan chains and subsequently affect the structure and function of the associated glycoproteins, which ultimately leads to various abnormal physiological states that include Alzheimer’s disease [176], CDGs [158, 160], and metastatic cancer progression [176, 177]. Hence, the processing glucosidase activities are critical for protein folding and ER quality control. While some eukaryotes have an alternative minor pathway that involves the deglucosylation by endo α(1,2) mannosidases, whose expression varies from tissue to tissue and cannot compensate for lack of Glu-I or Glu-II [178]. Inhibition of Glu-I has the potential to control viral amplification/disease, and tumour progression since several viral proteins rely extensively on the calnexin and calreticulin pathway for acquiring 44  native conformation [144, 179]. Understanding the function, mechanism and structure of Glu-I is essential to develop therapeutic approaches through specific inhibitors or via amelioration of activity. Lack of rapid methods to produce sufficient quantities of the enzyme and the random radiolabelling of natural substrates used for the assay have been major obstacles in the progress of these studies. Hence, the current study is aimed at the establishment of a production method using a prokaryotic expression system to obtain large quantities of the enzyme, and a procedure to synthesise a trisaccharide substrate that is simpler than other synthetic substrates previously reported to estimate the Glu-I function [120]. Glu-I has high specificity towards the α(1,2)-glycosidic bond, and a trisaccharide is the minimum length needed for its activity. Hence, a trisaccharide substrate was synthesised that can fulfil the requirements of Glu-I.  2.3 Materials and methods 2.3.1 Materials All the materials used in this study were of reagent grade. pET30a(+) plasmid was obtained from Novagen® Inc., and the glucose assay kit (GAGO-20) was obtained from Sigma (St. Louis, MO). 2.3.2 Microbial strains and plasmid Escherichia coli DH5α and BL21(DE3) strains were used for the cloning of pET30a-CWH41Δ1-34 and recombinant Cwh41Δ1-34p expression respectively. S. cerevisiae AH22 was used to obtain the genomic DNA. S. cerevisiae AH22 was grown on YPD medium. E. coli were cultured on LB broth/agar plates with or without kanamycin (50 µg/ml). pET30a(+) was used as the cloning vector as well as the E. coli expression vector.  45  2.3.3 Isolation of yeast genomic DNA Genomic DNA was isolated according to the method described by Harju et al. (2004)  . Repeated freeze-thawing of yeast cells was carried out in lysis buffer to break them, and the released genomic DNA was extracted by chloroform and subsequently precipitated by ethanol.  2.3.4 Cloning and preparation of expression constructs of Glu-I A pair of primers was designed to amplify the gene regions that encode the truncated form of Glu-I from residues 35 to 833 (designated CWH41Δ1-34) [Table 2.1]. These primers were designed to clone the gene insert in-frame with the N-terminal His-tag in the pET30a(+) vector between EcoRI and XhoI to form pET30a-CWH41Δ1-34 (Figure 2.1Error! Reference source not found.). The PCR conditions were: initial denaturation at 98 °C for 30 sec, then 30 cycles of 98 °C for 10 sec, 58 °C for 30 sec, 72 °C for 90 sec and then 72 °C for 10 min for final amplification. The PCR product was purified using Purelink® PCR Purification kit (Invitrogen). The purified PCR product and pET30a(+) plasmid were digested with EcoRI and XhoI, and the digested DNA was purified from the gel following gel electrophoresis using Purelink® Gel Extraction kit (Invitrogen). The purified digested gene insert and plasmid were ligated together with T4 DNA ligase (New England Biolabs), and the ligation reaction solution following incubation was transformed into E. coli DH5α competent cells (New England Biolabs). Positive clones were identified by colony PCR and presence of insert was confirmed by restriction analysis of recombinants with EcoRI and XhoI. Phusion® High-Fidelity DNA Polymerase (New England Biolabs Ltd.) was used to amplify the Glu-I gene whereas One Taq® master mix (New England Biolabs Ltd.) was used in colony PCR for the screening of transformants. The recombinant plasmid, called pET30a-CWH41Δ1-34, was isolated from positive transformants and sequenced using the primers listed in Table 2.1 to check the accuracy and orientation of the inserted glucosidase gene.  46  2.3.5 Expression of Glu-I in E. coli The pET30a-CWH41Δ1-34 clone with the insert properly oriented and in-frame with N-terminal His-tag was transformed into E. coli BL21(DE3) for expression studies. Three colonies were picked randomly from the transformants to screen for the expression of Glu-I on SDS-PAGE, and the colony with the highest expression was selected from these 3 clones. E. coli BL21(DE3) transformed with pET30a-CWH41Δ1-34 was cultured at 37 °C in LB medium containing 50 µg/ml of kanamycin to an absorbance of 0.6 at 600 nm. Glu-I expression was optimised by varying induction temperatures from 16 to 37 °C in the presence of 1 mM isopropyl-D-thiogalactopyranoside (IPTG). Expression level was also optimised by changing the IPTG concentration from 0.5 to 5mM, during incubation for induction of Glu-I synthesis. The expression of Glu-I was analysed on SDS-PAGE and western blot using anti-His-tag antibodies, and also by assaying activity using the synthetic trisaccharide.   Table 2.1. Primers used to clone/sequence the Glu-I gene  Primer Sequence Amplification Forward primer CCGGAATTCATGGAAGAATATCAAAAGTTCACGAATGA  Reverse primer CCGCCGCTCGAGTTAGAAGCGTCCAAGGATGTTG Sequencing  T7 promoter primer TAATACGACTCACTATAGGG T7 terminator primer GCTAGTTATTGCTCAGCGG RSP1 TAAAGAATTAGGCGAGTATC RSP2 AGTATGATTTTGACCTTGCC 47   Figure 2.1. Graphical depiction of pET30a-CWH41Δ1-34. The CWH41Δ1-34 gene was inserted between EcoRI and XhoI restriction sites and expressed as a fusion protein with N-terminal His- tag. SnapGene® was used to create this vector map.   2.3.6 Purification of recombinant Glu-I Intracellularly expressed Glu-I was isolated by first pelleting E. coli cells at 12,000 x g for 5 min. The pellet was resuspended in 4 volumes of binding buffer (20 mM phosphate buffer containing 20 mM imidazole and 500 mM NaCl), and cell lysis was carried out using ultrasonication at 20% amplitude for 20 cycles using a Qsonica™ ultrasonicator (50 W), alternating pulses and cooling on the ice every 30 sec.  The 6xHis-tagged Cwh41Δ1-34p was purified from the supernatant via immobilised metal affinity chromatography (henceforth called as Ni-NTA chromatography) using Nickel-linked nitrilotriacetic acid (Ni-NTA) column. The supernatant was first separated by centrifugation and 48  applied onto a 1 ml His-Trap HP column (GE Healthcare Ltd.) attached to an ÅKTA Purifier. The column was washed thoroughly with binding buffer followed by washing with binding buffer plus elution buffer (19:1 ratio). Bound protein was then eluted with elution buffer containing 20 mM sodium phosphate, 500 mM imidazole and 500 mM NaCl. Fractions were collected and analysed for Glu-I activity and protein concentration after imidazole was removed by exchanging into 20 mM phosphate buffer pH 6.8 and concentrated using a Centricon 10 kDa cut-off filter membrane (Millipore). Each concentrated sample was loaded onto a HiPrep™ 16/60 Sephacryl S-100 HR size exclusion column (GE Healthcare Life Sciences) that was equilibrated with a buffer that contained 20 mM phosphate buffer (pH 6.8) and 200 mM NaCl. The eluted protein was collected into two fractions. Fraction 1 contained contaminant proteins, whereas Glu-I activity was observed in Fraction 2. The fraction with Glu-I activity was concentrated to 20 mg/ml using a Centricon (Millipore) concentrator with a 10 kDa molecular weight cut-off membrane and analysed on a TGX Stain-Free gel (4%–15% acrylamide; Bio-Rad Laboratories, Hercules, CA, USA).   2.3.7 Expression of codon optimised Glu-I Cwh41Δ1-34p DNA sequence was analysed using GenScript® Rare Codon Analysis Tool software (http://www.genscript.com/cgi-bin/tools/rare_codon_analysis). Using GenScript®'s OptimumGene™ bioinformatics tool, the optimal gene was designed. GC content was adjusted to be consistent with that in E. coli, and unfavourable secondary structures and cis-acting elements were also optimised to increase the half-life of mRNA. The optimised gene was then synthesised at Genscript® and cloned into pUC57 with NdeI and XhoI restriction site modifications at the 5′ end and 3′ end, respectively. The sequences of the native Cwh41Δ1-34p gene and the optimised gene are shown in Figure A.1. Cwh41Δ1-34p gene insert was isolated from the pUC57 and ligated into the expression vector pET30a(+) which was previously digested with NdeI and XhoI. The 49  recombinant pET30a(+)-CWH41Δ1-34-OPT was initially transformed into E. coli DH5α, and positive transformants with properly oriented insert were screened by colony PCR and restriction digestion. Recombinant plasmids with proper insert were transformed into E. coli BL21(DE3), and the positive transformants were selected for expression study. Expression studies were carried out as described for the unoptimized Glu-I.  2.3.8 Synthesis of α-D-Glc1,2α-D-Glc1,3α-D-Glc–OCH3 For the synthesis of a trisaccharide substrate, our laboratory collaborated with Dr. Todd Lowary’s laboratory from The Department of Chemistry at The University of Alberta. The trisaccharide was synthesised by Dr. Akihiro Imamura using a linear synthetic strategy starting from the reducing end glucose derivative. The synthesis procedure is depicted in Figure 2.2. First, commercially available methyl α-D-glucopyranoside 1 was protected by benzylidene group on O4, and O6 positions and subsequent regioselective benzylation at the C2 position provided the glucosyl acceptor 3 in 61% yield over 2 steps.  To extend the sugar skeleton, glucosyl donors 8, 9 were synthesised using the thioglycoside derivative 6 as a key structure, which was prepared from D-glucose in 4 steps.  Benzylation of the hydroxyl groups at C2 and C3 positions with BnBr and NaH gave the non-reducing end glucosyl donor 8 in 91% yield.  The centrally located glucose residue required selective protection and deprotection of the hydroxyl group at C2 to extend the sugar chain at that position. Furthermore, it was thought that an ether-type group may be required as a protecting group on O2 to anticipate α-predominant glycosidation by virtue of the anomeric effect. In view of these reasons, the donor 9 bearing p-methoxybenzyl (PMB) group at the C2 position was designed and synthesised from compound 6 in 2 steps involving selective benzylation of O3 and subsequent introduction of PMB group on O2. The obtained donor 9 was then glycosidated with the reducing end glucosyl acceptor 3 with solvent control using diethyl ether (which is known 50  to give high α-selectivity in glycosylations) to give the desired α-glycoside 10 in 65% yield along with the corresponding β-isomer in 15% yield.  Following removal of the PMB group by treatment of 2, 3-dichloro-5,6-dicyanobenzoquinone (DDQ) gave the disaccharyl acceptor 11, which was ready for next glycosylation. The coupling of the monoglucosyl donor 8 and the disaccharyl acceptor 11 was carried out under the same conditions as that of the first glycosylation, resulting in the formation of the desired trisaccharyl framework 12 with three α-oriented glycosides in 72% yield. Finally, the concomitant removal of benzyl and benzylidene groups by hydrogenolysis furnished the targeted trisaccharide (α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3) in quantitative yield (~130 mg).   51     Figure 2.2. Schematic illustration of the synthesis of α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3. This figure was provided by Dr Akihiro Imamura, who synthesised this substrate, from Dr.Todd Lowary’s laboratory at the University of Alberta.   52  2.3.9 Enzyme assays Glu-I activity was assayed using the synthetic trisaccharide α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3. An aliquot of enzyme in 20 mM phosphate buffer, pH 6.8 was incubated with 10 nmoles of the substrate in a 5 µL total volume at 37 °C. The free glucose released by Glu-I action was measured using PGO enzyme preparation (Sigma-Aldrich chemicals catalogue# P7119).  One unit of activity corresponded to the amount of enzyme that liberated 1 nmole of glucose per min at 37 °C, pH 6.8. Enzyme assays were always carried out in duplicates, and the values are reported in mean ± range.  2.3.10 Activity with other substrates Aryl α-glucosidase activity was also determined using 100 mM p-nitrophenyl α-D-glucopyranoside as described elsewhere [13]. The Glu-I assay was also carried out with substrates such as kojibiose (0.5 and 25 mM) (Sigma-Aldrich chemicals catalogue# K4769) and methyl-umbelliferyl α-glucoside (49 µM and 10 mM) (Sigma-Aldrich chemicals catalogue# M9766) to assess the specificity.  A 4 µL aliquot of the enzyme sample was incubated with the substrate at 37 °C and released glucose was estimated using the method described above.  2.3.11 Determination of Km Glu-I (0.2 units) was incubated with varying concentrations of substrate (1 to 25 nmoles) in a total volume of 5 µL. Km was calculated using non-linear regression analysis. Graphpad® Prism® software was used for statistical analysis.  Due to the constraints of the availability of the substrate, this experiment was carried out with no replicates.   53  2.3.12 Other methods Protein was quantified using the Bradford method and the Bio-Rad protein assay reagent [181]. However, during affinity chromatography, protein amount was monitored by absorbance at 280 nm. Sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) was performed using 10.5% polyacrylamide gels [182]. Protein bands on polyacrylamide gels were visualised with Coomassie Blue staining. Western blotting was carried out using rabbit anti-His-tag IgG antibodies (Santa Cruz Biotechnology, Santa Cruz, CA) and bands on the membrane were detected by HRP conjugated anti-rabbit IgG antibodies (Santa Cruz Biotechnology, Santa Cruz, CA) and Immun-Star™ WesternC™ chemiluminescent kit (Bio-Rad Laboratories Inc.).    2.4 Results 2.4.1 Cloning and preparation of expression constructs of Glu-I gene Glu-I specific gene primers were used to amplify DNA that encodes Cwh41Δ1-34p, and the amplicon was cloned into a pET30a(+) vector as described in the materials and methods section. A pET30-CWH41Δ1-34 construct encoding 800 amino acids and an N-terminal 6xHis-tag was generated. This construct was transformed into E. coli DH5α, and positive transformants were identified by colony PCR and restriction digestion (Figure 2.3 and Figure 2.4). DNA sequencing was carried out to check the integrity and orientation of the insert in the positive transformants. These clones were further used for the expression studies. 54   Figure 2.3 Restriction analysis of pET30a(+) vector and insert (CWH41Δ1-34). Vector and insert were cut using EcoRI and XhoI and analysed on a 0.8% agarose gel. Lane 1 is molecular weight marker (Fermentas catalogue# SM0311). Lanes 2 and 3 are restriction-digested pET30a(+) and CWH41Δ1-34 respectively.    Figure 2.4 Restriction analysis of pET30a-CWH41Δ1-34 clones.  Lanes 1 and 6 are molecular weight markers (Fermentas catalogue# SM0311). Recombinant pET30a-CWH41Δ1-34 clones (Lanes 2 to 5) were digested with EcoRI and XhoI and analysed on 0.8% agarose gel. DNA bands for pET30a(+) vector (5.4 kb) and CWH41Δ1-34 (2.4 kb) were observed in all the clones tested.   2.4.2 Expression and purification of Glu-I in E. coli  Recombinant constructs (pET30a-CWH41Δ1-34) with proper insert were transformed into E. coli BL21(DE3). The expression of the constructs was carried out by IPTG induction as described in the materials and methods section. Initially, a small-scale expression of 4 transformants from each construct was carried out to screen the expression of soluble Glu-I truncations (Figure 2.5). All four transformants showed expression of soluble Cwh41Δ1-34p. Among these, the transformant 55  #3 that had the highest Cwh41Δ1-34p synthesis was selected and used for further optimisation studies. IPTG concentration (0.1 to 1 mM) and induction temperatures (16 to 37 °C) were optimised for the increased expression of soluble Glu-I. The highest soluble Glu-I production was observed when induction was carried out at 25 °C and 1 mM IPTG concentration.   Figure 2.5. The screening of E. coli BL21(DE3) transformants for Cwh41Δ1-34p expression. The cell pellet obtained from 50 µl of IPTG-induced culture was mixed with sample buffer and analysed on SDS-PAGE.  Lane 1-molecular weight marker, Lane 2-5 cell pellets of the transformants 1-4, Lane 6-empty, Lane 7 – negative control (E. coli BL21(DE3) with the empty pET30a(+) plasmid). Arrow indicates the expected size for the Cwh41Δ1-34p.  The pET30a-CWH41Δ1-34 construct expressed recombinant Cwh41Δ1-34p which retained activity (Table 2.2). The recombinant 6xHis-tagged Cwh41Δ1-34p protein was expressed intracellularly in E. coli and was isolated from crude extract by immobilised metal affinity 56  chromatography (IMAC) using an Ni-NTA column that binds the N-terminal 6xHis-tag. The 6xHis-tagged Cwh41Δ1-34p protein was eluted from the Ni-NTA column using 20 mM phosphate buffer with 500 mM each of imidazole and sodium chloride. A single major peak was observed in the Ni-NTA elution, and Glu-I activity was confined to these fractions (Figure 2.6). The presence of high levels of imidazole in the eluate caused interference with the enzyme assay. However, removal of imidazole by exchanging with 20 mM phosphate buffer pH 6.8 wholly restored the activity of S. cerevisiae Glu-I. Recombinant Cwh41Δ1-34p was observed at 100 kDa on SDS-PAGE, corresponding to the 97 kDa protein with the additional cleavage sites for thrombin and enterokinase, S-tag and 6xHis-tag (lane 6 in Figure 2.8). Ni-NTA-eluted Cwh41Δ1-34p fraction also contained a few short fragments (60 and 40 kDa) with the N-terminal 6xHis-tag, as detected on Western blot using anti-his tag antibodies (lane 8 in Figure 2.8), in the Ni-NTA eluate that may be released due to proteolysis during the sample preparation. These fragments bound to the Ni-NTA column due to the 6xHis-tag at N-terminus and were, therefore, lacking the catalytic C-terminal domain. Ni-NTA-purified recombinant Cwh41Δ1-34p after the removal of imidazole exhibited a specific activity of 3370 U per mg of protein with the synthetic trisaccharide α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3 (Table 2.2). To remove the smaller protein fragments and to further purify the Cwh41Δ1-34p from the Ni-NTA preparations, size-exclusion chromatography purification of Ni-NTA- purified Cwh41Δ1-34p was carried out. Glu-I activity was detected in fraction 2 during size exclusion chromatography ( Figure 2.7). Cwh41Δ1-34p was purified to ~97% purity based on the comparison of intensity on ImageLab® software quantification tool (Figure 2.8). Cwh41Δ1-34p contains a disulphide bond between cysteines 636 and 652 [147]. Hence, to facilitate the disulphide bond formation and to enhance the expression levels of soluble Cwh41Δ1-34p, pET30-CWH41Δ1-34 was also 57  transformed into E. coli BL21(DE3)-RosettaGami® cells which are known to enhance disulphide formation and solubility of eukaryotic proteins. However, expression in RosettaGami® did not result in a significant improvement in expression of soluble Cwh41Δ1-34p (Figure 2.9).   Figure 2.6. Elution of recombinant Glu-I from Ni-NTA column. The solid line and dashed line represent optical absorption at 280 nm and concentration of eluting buffer respectively. The unbound proteins were washed with 20 mM phosphate buffer pH 7.4 with 20 mM imidazole and 500 mM sodium chloride. In step 1 elution, 20 mM phosphate buffer pH 7.4 with 43 mM imidazole and 500 mM sodium chloride was used to wash the loosely bound contaminant proteins. In step 2, 20 mM phosphate buffer pH 7.4 with 500 mM imidazole and 500 mM sodium chloride was used to elute the bound Glu-I.   Table 2.2 Purification of recombinant Glu-I from E. colia a Results are an average ± range of two preparations. Results are reported as per gram wet weight of bacterial biomass. b Unit Glu-I activity = 1 nanomole glucose released per minute at 37°C  Purification step Activityb (Units) Protein (mg) Specific activityb (U/mg protein) Purification fold Cell homogenate 3215 ± 190 12.55 ± 0.23 255 ± 10 - Ni-NTA affinity chromatography 2290 ± 135 0.99 ± 0.14 3370 ± 370 13 58   Figure 2.7 Size exclusion chromatography of Cwh41Δ1-34p. 20 mM phosphate buffer pH 6.8 with 200 mM sodium chloride was used on HiPrep Sephacryl S-100HR column. Majority of the Glu-I activity was detected in the fraction 2.    Figure 2.8 SDS-PAGE analysis of Cwh41Δ1-34p. (Lane 1-molecular weight marker, Lane 2-whole cell extract uninduced, Lane 3- whole cell extract induced, Lane 4-insoluble fraction, Lane 5-soluble fraction, Lane 6-Ni-NTA elution), and Lane 7- Fraction 2 collected during size exclusion chromatography (SEC). Lanes 8 and 9, western blotting of Ni-NTA and SEC purified Glu-I with anti-His antibodies. Arrow indicates the bands with expected size of the Cwh41Δ1-34p. The intensity of the Cwh41Δ1-34p constitute to ~97% of the total intensity of the lane 7. The yellow boxes indicate the proteolytic products of Cwh41Δ1-34p.  59   Figure 2.9. Ni-NTA Purification of Cwh41Δ1-34p heterologously expressed in E. coli BL21(DE3) [--] and E. coli RosettaGami®  (--).   2.4.3 Expression of codon optimised Glu-I Using a T-Coffee homology alignment online tool, the ORF of the original and codon optimised Cwh41Δ1-34p gene were aligned (Figure A.1, Appendix). Codon adaptation index (CAI) is a measure of codon usage bias of a gene in a given specific host, and it ranges from 0 to 1. CAI of a gene is calculated from the CAI values of the individual codons. Codon optimisation improved the CAI from 0.64 to 0.88. In addition to codon modifications, one polyadenylation site, one Shine Delagarno-like sequence (GGRGGT) and one internal ribosome binding site (AGGAGG) were modified, and regions with high GC content were avoided. There is another alternative tool, the frequency of optimal (FOP) codons, to predict protein expression levels based on the ratio of optimised to synonymous codons [183]. FOP of a particular amino acid in a given specific host ranges from 0 to 100 where 100 indicates the highest usage frequency. Codon optimisation 60  improved the percentage of codons of CWH41Δ1-34 from 46 to 70 that have a FOP value between 91-100 (Figure 2.10). The codon optimised gene, CWH41Δ1-34-OPT, was synthesised and cloned into pUC57 by Genscript®. CWH41Δ1-34-OPT was isolated from pUC57 by restriction digestion and ligated into pre-cut pET30a(+) vector to make pET30a-CWH41Δ1-34-OPT. E. coli BL21(DE3) cells transformed with recombinant pET30a-CWH41Δ1-34-OPT expressed 6×His-tagged Cwh41Δ1-34p-OPT after a 16 hr incubation period. There was no significant increase in the expression of the soluble Cwh41Δ1-34p-OPT. However, there was a higher amount of Cwh41Δ1-34p-OPT in insoluble fraction as observed in the SDS-PAGE (lane 3 of Figure 2.11).     Figure 2.10 The frequency of optimal codons for Cwh41Δ1-34p in E. coli. (Black- unoptimised CWH41Δ1-34 and Grey- Optimised CWH41Δ1-34).  61   Figure 2.11. Purification of Cwh41Δ1-34p and Cwh41Δ1-34p-OPT expressed in E. coli BL21(DE3). A. Ni-NTA purification B. SDS-PAGE analysis of the expression of Cwh41Δ1-34p and Cwh41Δ1-34p-OPT. Lane 1 and lane 10 are molecular weight standards (kDa). Cwh41Δ1-34p-OPT samples- lane 2, Whole cell extract; lane 3, insoluble fraction; lane 4, soluble fraction; lane 5, Ni-NTA purified fraction. Cwh41Δ1-34p samples- lane 6, Whole cell extract; lane 7, insoluble fraction; lane 8, soluble fraction; lane 9, Ni-NTA purified fraction. Arrow indicates the expected size for Cwh41Δ1-34p and Cwh41Δ1-34p-OPT.   62  2.4.4 Synthesis of trisaccharide and Glu-I activity assay The synthesis of the trisaccharide substrate was successfully carried out via a linear synthetic strategy starting from a methyl glucopyranoside and the structure was verified by 1H NMR analysis (Figure 2.12). This synthetic trisaccharide partially mimics the natural substrate but lacks the Man9GlcNac2 chain. The Ni-NTA purified Cwh41Δ1-34p displayed a Km of 1.27 mM with the synthetic trisaccharide substrate.   63   Figure 2.12. 1HNMR analysis of the synthetic trisaccharide substrate. A. Experimentally determined by Dr. Akihiro Imamura at Dr. Todd Lowary’s laboratory at University of Alberta. B. Predicted 1H NMR spectra of the trisaccharide substrate.  64  2.4.5 Substrate specificity Native membrane-bound Glu-I from S. cerevisiae has high specificity for α(1,2)-linked glucose in a trisaccharide structure [11]. The recombinant soluble Cwh41Δ1-34p retained this specificity and did not hydrolyse kojibiose, p-nitrophenyl and methyl-umbelliferyl α-glucosides.   Figure 2.13 Michaelis-Menten plot of Glu-I activity with the synthetic trisaccharide- α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3. Due to the limitation in the availability of the substrate only 4 points of data were collected.    65  Table 2.3 Km of Glu-I against different synthetic substrates  Substrate  Km Reference  4-Methyl umbelliferyl α-D-glucoside N/A1 This study Kojibiose N/A2 This study (Glc)3-O-Methyl 1.27 mM This study (Figure 2.13) (Glc)3-Mannose-O-Methyl 1.26 mM Barker et al. 2011  (Glc)3-Octane-O-Methyl 1.28 mM Neverova et al. 1994  1 N/A  - no activity detected when incubated with methyl-umbelliferyl α-glucoside (49 µM and 10 mM) 2 N/A  - no activity detected when incubated with kojibiose (0.5 and 25 mM)    2.5 Discussion Cwh41Δ1-34p was successfully expressed in E. coli using pET30a(+) expression vector and 6-8 mg of enzyme was obtained per litre of culture. Codon optimisation of yeast CWH41Δ1-34 gene improved the codon adaptability index from 0.64 to 0.88. However, this increase did not lead to increased expression levels of Cwh41Δ1-34p. Despite the presence of five putative N-glycosylation sites, Cwh41Δ1-34p was heterologously expressed in E. coli as a functional protein. Previous reports also noted that glycosylation is not essential for the enzyme function [12]. Cwh41Δ1-34p was glycosylated when expressed in P. pastoris [133, 147]. However, information is not available on whether removal of the glycan chain affected the activity of Glu-I. Comparison of all three of these studies reveals that glycosylation does not significantly alter the specific activity or substrate affinity with synthetic substrates.  66  Native membrane-bound Glu-I from S. cerevisiae has a high specificity for α(1,2)-linked glucose in a trisaccharide structure [11]. The recombinant soluble enzyme retained the specificity that was observed in the original Glu-I and did not hydrolyse the kojibiose and 4-methyl umbelliferyl α-D-glucoside. However, a truncated form of C. albicans Glu-I that lacks 419 amino acids at the N-terminus heterologously expressed in E. coli was less specific and less active towards other substrates [184]. The Km value for α-D-Glc1,2α-D-Glc1,3α-D-Glc-OCH3 synthesised in this work was 1.27 mM; statistical error could not be calculated as only the single assays were carried out due to the scarcity of the substrate. The Km value of 1.27 mM is similar to the values reported for D-Glc1,2α-D-Glc1,3α-D-Glc-O(CH2)8COOCH3 of 1.28 mM [135], and 1.26 mM for the mannose containing tetrasaccharide Glc1,2α-D-Glc1,3α-D-Glc-Man-OCH3 [133]. However, a Glu-I from Aspergillus brasiliensis showed micromolar Km of 6.1 µM and 4.2 µM with Glc3Man9GlcNAc2-PA and Glc3Man4-PA respectively [120]. Similar results were observed with Glu-I isolated from mung bean seedlings when tested against the substrate with a reduced number of mannose residues [129]. Removal of up to four mannose residues from the Glc3Man5-GlcNAc2 substrate had little effect on its utilisation as a substrate for the Glu-I. The plant Glu-I acted almost as well on Glc3Man5GlcNAc as it acted on Glc3Man9GlcNAc Glc3Man5-GlcNAc. The minimum structural requirement for substrates of Glu-I has not been thoroughly investigated. However, from the current study and the previous studies that involved Glu-I activity with synthetic substrates, it is evident that the minimum structure requirement is the trisaccharide, α-D-Glc1,2α-D-Glc1,3α-D-Glc since moieties after first three glucose residues in the synthetic substrates did not significantly affect the Km of the Glu-I. Hence, the synthetic trisaccharide developed in this study can be used as a substrate for monitoring the Glu-I function. 67  The presence of imidazole interfered with Glu-I activity, and this interference was nullified when imidazole was removed, whereas imidazole irreversibly inhibited C. albicans Glu-I [184]. The specific activity of heterologously expressed Glu-I purified by a single step was 3370 U/mg which is comparable to previously reported values of 3130 U/mg and 3122 U/mg reported for Glu-I expressed in S. cerevisiae and P. pastoris respectively [13, 133]. The slight differences in specific activity of these three preparations may be a result of the different expression systems used since the enzyme from P. pastoris is glycosylated, or maybe a result of the difference in the catalytic turnover of the different substrates. Moreover, Ni-NTA eluted samples of Glu-I purified from E. coli did not show any activity with p-nitrophenyl α-D-glucopyranoside, indicating the absence of contamination of aryl glucosidases which is common in eukaryotic expression systems. I have shown that E. coli is a robust prokaryotic expression system for rapid expression of Glu-I with high specific activity. The potential of synthetic trisaccharide was investigated to be used as a substrate for the Glu-I activity assays. The recombinant enzyme had a Km of 1.27 mM with the synthetic trisaccharide; hence, the synthetic trisaccharide can be used as an alternative substrate for the determination of Glu-I activity. I have tested this expression system and the purification method more than twenty times with reproducibility. This study establishes a robust system for the heterologous expression of the soluble form of yeast Glu-I which can be used as a model to study the structure-function of GH 63 family glycoside hydrolases.   68  Chapter 3: Heterologous expression of yeast Glu-I truncations in E. coli and isolation of the catalytic domain of Glu-I   69  3.1 Summary Processing alpha-glucosidase-I (Glu-I) is a type-II membrane-bound protein that catalyses the removal of the terminal glucose from the pre-assembled glycan moieties that are co-translationally transferred to nascent proteins. Generation of a functional 37 kDa C-terminal domain during trypsin digestion exhibits the catalytic activity suggests that the catalytic domain of Glu-I is confined to a C-terminal 37 kDa fragment (Cwh41Δ1-525p) of the soluble 98 kDa protein (Cwh41p). This also suggests a potential regulatory and folding role for the N-terminal portion of the protein. In this work, different methods for expression of the C-terminal domain of Glu-I in E. coli were evaluated. Expression of Cwh41Δ1-525p alone using the pET30a(+) vector resulted in protein accumulation in inclusion bodies, even though the same system produced soluble and active Cwh41Δ1-34p. Co-expression of solubility-enhancing tags (maltose-binding protein tag and glutathione-S-transferase tag) and molecular chaperones (GroES-GroEL, DnaJ-DnaK, and Trigger factor) did not result in improvement in solubility. Co-expression of Glu-I N-terminal domain (Cwhnp) and Cwh41Δ1-525p using pACYCDuet1 dual expression vector significantly improved the solubility.  Approximately 42% of the expressed Cwh41Δ1-525p was soluble when co-expressed with Cwhnp, suggesting that the N-terminus is required for the folding of the catalytic domain. However, soluble Cwh41Δ1-525p did not hydrolyse the α1,2-glycosidic bond of trisaccharide substrate. Due to the lack of functional expression of the catalytic domain, I developed an alternative method for the isolation of the Glu-I catalytic domain from full enzyme using trypsin hydrolysis and subsequent purification by size exclusion chromatography.   70  3.2 Introduction Processing alpha-glucosidase-I is a family 63 glycoside hydrolase and plays a crucial role in glycosylation. It is a type-II membrane protein with a short cytosolic tail (M1-K10), a transmembrane region (T11-I28) and a large lumenal C-terminal domain (S29-F833). There have been several studies that reported the release of the C-terminal catalytic domain via trypsin hydrolysis of Glu-I orthologs [116, 141, 150, 152]. Glu-I (85 kDa) isolated from pig liver generated a mixture of fragments where a non-glycosylated 29 kDa fragment was a dominant species, and the mixture still retained 60% of the initial activity [141]. Similarly, trypsinisation of Glu-I from mouse, rat, guinea pig, bovine mammary glands and sheep liver generated a 39 kDa fragment [150]. Consistent with mammalian Glu-I, yeast Glu-I was also susceptible to trypsin hydrolysis and generated two catalytic domains with molecular weights of 59 and 37 kDa [152]. N-terminal sequencing of the 37 kDa fragment revealed the hydrolysis site as between lysine 524 and arginine 525. Surprisingly, this 37 kDa was 1.9 times more active than the undigested 98 kDa enzyme. Structure-function studies of this 37 kDa fragment would give more insights into the catalytic mechanism of Glu-I and reasons for the superior activity. Moreover, expression of the shorter yet functional truncations that contain the catalytic domain can bypass many hurdles that are faced during structural and catalytic/kinetic manipulations of the massive 98 kDa protein. In the past, based on the results of trypsin digestion of Glu-I, there have been a few attempts to overexpress the Glu-I truncations. However, they were not successful in either Saccharomyces cerevisiae or Pichia pastoris [12, 147]. Initially, expression studies of truncations (Cwh41Δ1-525p) were carried out before the availability of structural details of Glu-I. However, after the availability of Glu-I structural information, two more constructs (Cwh41Δ1-314 and Cwh41Δ1-349) were included in our study using the previously established Glu-I expression system in E. coli (Figure 71  3.1). Barker and Rose (2013) postulated that the lack of expression of Cwh41Δ1-525p in yeast could be because the construct begins at a long disordered loop between C-terminal α-helices CH4 and CH5 as depicted in figure 3.1, and fails to form the full domain [147].   From the trypsin hydrolysis studies, it is evident that the N-terminal domain is not required for Glu-I function. Moreover, the function of the N-terminal fragment is not known yet. Hence, I hypothesised that the N-terminal domain plays a significant role in protein formation/folding rather than being directly involved in Glu-I function. Therefore, the current study explored the possibility of generating a functional catalytic domain via heterologous expression of Glu-I truncations and co-expression of N- and C-terminal domains. Development of a method to isolate the Glu-I catalytic domain via trypsin hydrolysis was also investigated.    72   Figure 3.1. Graphical depiction of Glu-I domains and positions of truncations. (Redrawn from Barker and Rose (2013) with permission). The numbers have been modified to reflect that of full-length Glu-I.  Sandy brown area indicates N-domain, blue- C-domain and grey-C’ domain. Rectangular boxes ( ) indicate the initiation point for corresponding truncations.  73  3.3 Materials and methods 3.3.1 Glu-I truncations 3.3.1.1 Generation of Glu-I truncations The truncated gene regions were amplified using the primers listed in Table 3.1. Amplified genes were cut with EcoRI and XhoI, purified, ligated into a precut pET30a(+) vector, and transformed into E. coli DH5α. Positive clones were identified by colony PCR and presence of the insert was confirmed by restriction analysis of recombinants with EcoRI and XhoI. Phusion® High-Fidelity DNA Polymerase (New England Biolabs Ltd.) was used to amplify the Glu-I gene, whereas One Taq® master mix (New England Biolabs Ltd.) was used in colony PCR for the screening of transformants. Recombinant plasmids (pET30a-CWH41Δ1-314, pET30a-CWH41Δ1-349 and pET30a-CWH41Δ1-525) were isolated from positive transformants and sequenced using the corresponding primers listed in Table 3.1 to check the accuracy and orientation of the inserted Glu-I sequences.  Table 3.1. Primers used for amplification of truncated Glu-I genes. Primer Sequence (5’ -3’) FOR del 525 AATCCAGAATTCATGACGAACAATCTAGAAGCCAATCC FOR del 349 GGGAATTCCATATGGACTCAATTGAAAGCGTGGAGGTCAAAAG FOR del 314 GGGAATTCCATATGACTCAAAGTATTTCCACCAGGGAAC REV CWH41Δ1-34 CCGCCGCTCGAGTTAGAAGCGTCCAAGGATGTTG    74  3.3.1.2 Expression of truncated Glu-I Recombinant plasmids that contain truncated Glu-I (pET30a-CWH41Δ1-314, pET30a-CWH41Δ1-349 and pET30a-CWH41Δ1-525) were transformed into E. coli BL21(DE3), and transformants were used for the expression studies. Expression and purification of truncated Glu-I were carried out as described earlier in sections 2.2.1.3 and 2.2.1.4 in Chapter 2. 3.3.1.3 Solubility enhancing tags To enhance the solubility of the truncated forms of Glu-I, expression of Cwh41Δ1-525p was carried out with maltose binding protein (MBP) and glutathione S-transferase (GST) tags at the N-terminus. The pMALC5E vector was used to obtain the MBP-tag and pGEX4T1 vector for the GST-tag. 3.3.1.4 Molecular cloning and expression of Glu-I truncations fused with solubility tags Gene region that encodes a truncated version of Glu-I, CWH41Δ1-525, was cut from pET30a-CWH41Δ1-525 using EcoRI and XhoI and ligated with the pGEX4T1 vector that was previously digested with the same restriction enzymes (Figure 3.2A). The Cwh41Δ1-525p was expressed as a dual-tagged protein with a GST-tag on N-terminus and a 6xHis-tag on C-terminus. A PCR was carried out using a set of primers (Forward 5’-TATACATATGGAAGAATATCAAAAGTTCACG-3’, and reverse 5’-ATAGTTTAGCGGCCGCTCAATGATGATGATGATGATGGAAGCGTCCAAGGATGTGACAA-3’). The PCR product was digested with NdeI and NotI and inserted between the same restriction sites in pMALC5E by ligation (Figure 3.2B). Ligation mixtures were transformed into E. coli DH5α. Expression and purification were carried out as described in sections 2.3.5 (page 46) and 2.3.6 (page 47) in Chapter 2.   75   Figure 3.2 Graphical depiction of pGEX4T1- CWH41Δ1-525 (A) and pMALC5E- CWH41Δ1-525 (B). CWH41Δ1-525 was inserted between EcoRI and XhoI in pGEX4T1 and NdeI and NotI in pMALC5E. Vector maps were created using SnapGene® software. 76  3.3.2 Co-expression of molecular chaperones pET30a-CWH41Δ1-525 was co-expressed with molecular chaperones GroEL (60 kDa), GroES (10 kDa), DnaJ (40 kDa), DnaK (70 kDa), Trigger factor (56 kDa) and GrpE (22 kDa) in E. coli. The genes for these chaperones were obtained as plasmids from Takara Bio Inc.  (Table 3.2). Each chaperone plasmid was co-transformed along with pET30a-CWH41Δ1-525 into E. coli BL21 competent cells, and the transformed mixture was plated on LB agar with kanamycin and chloramphenicol. Positive transformants were selected and subsequently used for the expression study. Expression of Cwh41Δ1-525p was induced with 1 mM IPTG, and molecular chaperones were induced with the corresponding inducer listed in Table 3.2.  Table 3.2. List of plasmids and molecular chaperones Plasmid Chaperone Expression inducer (concentration) Selection Marker pG-KJE8 dnaK-dnaJ-grpE groES-groEL L-Arabinose (4 mg/ml) and  Tetracycline (10 ng/ml) Chloramphenicol pGro7 groES-groEL L-Arabinose (4 mg/ml) Chloramphenicol pKJE7 dnaK-dnaJ-grpE L-Arabinose (4 mg/ml) Chloramphenicol pG-Tf2 groES-groEL-tig Tetracycline (10 ng/ml) Chloramphenicol pTf16 tig (Trigger factor) L-Arabinose (4 mg/ml) Chloramphenicol  3.3.3 Co-expression of N- and C-terminus A set of primers were designed to amplify and clone the N-terminal region (CWHN). This amplified region was digested with EcoRI and NotI, and digested product was then ligated with the previously digested pACYC-DUET1 vector. The recombinant plasmid was then transformed into E. coli DH5α, and the orientation of the insert was confirmed by sequencing and restriction digestion. The recombinant plasmid pACYC-DUET1-CWHN was then transformed into E. coli 77  BL21(DE3) for expression studies. The gene region that encodes the C-terminal domain was excised from the pET30a-CWH41Δ1-525 plasmid using NdeI and Bpu1102I and inserted into the pACYCDUET-1 or pACYC-DUET1-CWHN vector that was previously digested with NdeI and Bpu1102I (Figure 3.3). This recombinant plasmid was then transformed into E. coli DH5α, and colonies chosen to perform a DNA miniprep of potential recombinant plasmids. Restriction-digestion analysis was carried out using NdeI and Bpu1102I to confirm the proper ligation of the insert. After confirmation of the orientation of the insert, the recombinant plasmids pACYC-DUET1-CWH41Δ1-525 and pACYC-DUET1-CWHN-CWH41Δ1-525 were transformed into E. coli BL21(DE3) for expression studies.  Figure 3.3 Vector map of pACYC-DUET1-CWHN-CWH41Δ1-525. Both CWHN and CWH41Δ1-525 were cloned into this dual expression vector at different multiple cloning sites- MCS-1 and MCS-2 respectively. 78  3.3.4 Trypsin hydrolysis of Glu-I  A 20-µg aliquot of Cwh41Δ1-34p was mixed with tosyl phenylalanyl chloromethyl ketone (TPCK) treated trypsin from bovine pancreas (T1426, Sigma-Aldrich® chemicals) at 1:1000 (trypsin to Cwh41Δ1-34p) ratio and incubated at 4 °C initially for 3 hr. In the control sample, in place of trypsin, an equivalent volume of 20 mM phosphate buffer was added. After the incubation, 4 µL from each of these mixtures were tested for Glu-I activity using the synthetic trisaccharide substrate.   3.3.5 Isolation of catalytic domain of Glu-I After Ni-NTA purification, Cwh41Δ1-34p was incubated with trypsin at the optimised conditions, and the hydrolysed mixture was loaded onto HiPrep-Sephacryl-S100HR size exclusion chromatography column attached to an ÄKTA purifier and pre-equilibrated with 100 mM phosphate buffer containing 500 mM sodium chloride. The 100 mM phosphate buffer containing 500 mM sodium chloride was used as the mobile phase. Fractions were collected and tested for the presence of Glu-I activity and protein concentration and analysed on SDS-PAGE.   3.4 Results 3.4.1 Expression of Glu-I truncations The Glu-I truncations Cwh41Δ1-314p and Cwh41Δ1-349p were expressed in E. coli BL21(DE3). The expressed proteins accumulated as inclusion bodies (Figure 3.4 and Figure 3.5). The supernatant, after cell lysis, was applied onto an Ni-NTA affinity column; however, no soluble protein was detected in the eluted fractions from the column (Figure 3.6). Similar to other truncations, the construct with the gene insert for the C-terminal catalytic domain of Glu-I, 79  pET30a-CWH41Δ1-525, was expressed, but the truncated protein did not bind to the Ni-NTA affinity column (Figure 3.6) and it was observed predominantly in the insoluble fraction (Figure 3.7).  Figure 3.4. SDS-PAGE analysis of pET30a-CWH41Δ1-314 expression. The yellow box indicates the expected size of the Cwh41Δ1-314p protein. All the Cwh41Δ1-314p was observed in the insoluble fraction, and no soluble Cwh41Δ1-314p was eluted from the Ni-NTA column.  A minimum of 5 µg of protein was loaded onto each lane.   Figure 3.5. SDS-PAGE analysis of pET30a-CWH41Δ1-349 expression. The yellow box indicates the expected size of the Cwh41Δ1-349p protein. All of the Cwh41Δ1-349p was observed in the insoluble fraction and no soluble Cwh41Δ1-349p was eluted from the Ni-NTA column.  A minimum of 5 µg of protein was loaded onto each lane. 80   Figure 3.6. Purification of truncations of Glu-I by Ni-NTA chromatography. The solid lines (Red, Blue, and Green) represent optical absorption at 280 nm and the dashed line represents concentration of eluting buffer (%). The unbound proteins were washed with 20 mM phosphate buffer pH 7.4 with 20 mM imidazole and 500 mM sodium chloride. In step 1 elution, 20 mM phosphate buffer pH 7.4 with 43 mM imidazole and 500 mM sodium chloride was used to wash the loosely bound contaminant proteins. In step 2, 20 mM phosphate buffer pH 7.4 with 500 mM imidazole and 500 mM sodium chloride was used to elute the bound Glu-I.   Figure 3.7. SDS-PAGE analysis of Cwh41Δ1-525p. The yellow box indicates the expected size of the Cwh41Δ1-525p protein. Most of the Cwh41Δ1-525p was observed in the insoluble fraction and no soluble Cwh41Δ1-525p was eluted from the Ni-NTA column.  A minimum of 5 µg of protein was loaded onto each lane.  81  3.4.2 Effect of solubility enhancing tags and co-expression of molecular chaperones on the solubility of Cwh41Δ1-525p The solubility of Cwh41Δ1-525p and Cwh41Δ1-280p was not enhanced when fused to solubility enhancing tags- GST and MBP (Figure 3.8, and Figure 3.9). The Ni-NTA purification of the soluble fraction did not yield any soluble Cwh41Δ1-525p or Cwh41Δ1-280p (Figure 3.10 and Figure 3.11). Co-expression of the molecular chaperones GroEL, DnaK, DnaJ and trigger factor did not improve the solubility of Cwh41Δ1-525p (Figure 3.12 Figure 3.13 and Figure 3.14). Expression of GroEL, DnaK, DnaJ and trigger factor was confirmed with the SDS-PAGE analysis after cell lysis (Figure 3.12 Figure 3.13 and Figure 3.14), while the presence of remaining chaperones- GroES and trigger factor was not clear from SDS-PAGE (Figure 3.12). The overall expression level of Cwh41Δ1-525p is lower than the level of Cwh41Δ1-525p expressed in the absence of chaperones. This inferior expression could be due to the metabolic burden of the overexpression of the molecular chaperones. When FPLC with a Ni-NTA column was carried out using the soluble fraction of the E. coli BL21(DE3) after co-expression of chaperones and Cwh41Δ1-525p, no peak was observed during the elution indicating the absence of soluble Cwh41Δ1-525p (Figure 3.15). Therefore, the expressed chaperones did not improve the solubility of the recombinant Cwh41Δ1-525p in E. coli BL21(DE3) which accumulated in the insoluble fraction.   82   Figure 3.8. SDS-PAGE analysis of pGEX4T1- CWH41Δ1-525. The yellow box indicates the expected size of the Cwh41Δ1-525p protein with the GST-tag. All of the GST-tagged Cwh41Δ1-525p (also contain a 6xHis tag) was observed in the insoluble fraction, and no soluble Cwh41Δ1-525p was eluted from the Ni-NTA column.  A minimum of 5 µg of protein was loaded onto each lane.  Figure 3.9. SDS-PAGE analysis of pMALC5E- CWH41Δ1-280. The yellow box indicates the expected size of the Cwh41Δ1-280p protein with the MBP-tag. All of the MBP-tagged Cwh41Δ1-280p (also contain a 6xHis tag) was observed in the insoluble fraction, and no soluble Cwh41Δ1-280p was eluted from the Ni-NTA column. A minimum of 5 µg of protein was loaded onto each lane. 83   Figure 3.10. SDS-PAGE analysis of pMALC5E- CWH41Δ1-525. The yellow box indicates the expected size of the Cwh41Δ1-525p protein with the MBP-tag. All of the MBP-tagged Cwh41Δ1-280p was observed in the insoluble fraction, and no soluble Cwh41Δ1-280p was eluted from the Ni-NTA column. Approximately 5 µg of protein was loaded onto each lane. 84    Figure 3.11. Ni-NTA chromatography of Glu-I truncations (Cwh41Δ1-525p and Cwh41Δ1-280p) fused with solubility enhancing tags- MBP and GST. The solid lines (Red, Black, and Green) represent optical absorption at 280 nm and the dashed line represents concentration of eluting buffer (%). The unbound proteins were washed with 20 mM phosphate buffer pH 7.4 with 20 mM imidazole and 500 mM sodium chloride. In step 1 elution, 20 mM phosphate buffer pH 7.4 with 43 mM imidazole and 500 mM sodium chloride was used to wash the loosely bound contaminant proteins. In step 2, 20 mM phosphate buffer pH 7.4 with 500 mM imidazole and 500 mM sodium chloride was used to elute the bound Glu-I.   85   Figure 3.12. SDS-PAGE analysis of co-expression of Cwh41Δ1-525p with molecular chaperones GroEL and GroES. The blue arrow indicates the expected protein band for soluble Cwh41Δ1-525p, and yellow box indicates the insoluble Cwh41Δ1-525p.    Figure 3.13. SDS-PAGE analysis of co-expression of Cwh41Δ1-525p with molecular chaperones DnaJ and DnaK. The blue arrow indicates the expected protein band for soluble Cwh41Δ1-525p.   86   Figure 3.14. SDS-PAGE analysis of co-expression of Cwh41Δ1-525p with molecular chaperone trigger factor. The blue arrow indicates the expected protein band for Cwh41Δ1-525p.   Figure 3.15. Ni-NTA chromatography of Cwh41Δ1-525p co-expressed with molecular chaperones. The solid lines (Cwh41Δ1-525p co-expressed with GroES, GroEL and trigger factor - Red, and Cwh41Δ1-525p co-expressed with DnaJ and DnaK - Blue) represent optical absorption at 280 nm and the dashed line represents concentration of eluting buffer (%). An air bubble occurred in the tubing while purifying the Cwh41Δ1-525p co-expressed with GroES and GroEL which caused a small peak during the sample loading.  87   3.4.3 Co-expression of the Glu-I N- and C-terminus Initially, expression of the Glu-I N-terminal domain (Cwhnp) and the Glu-I C-terminal catalytic domain (Cwh41Δ1-525p) was carried out separately using the pACYC-DUET1 dual expression vector in E. coli, and the expression was analysed on the SDS-PAGE. Later, co-expression of Cwhnp and the Cwh41Δ1-525p was carried out using the pACYC-DUET1. No soluble fraction of the Glu-I catalytic domain (Cwh41Δ1-525p) was observed when expressed alone in the absence of the Cwhnp, and Cwh41Δ1-525p did not bind to the Ni-NTA column (Figure 3.16 and Figure 3.17). However, a significant portion (≈ 42%) of Cwh41Δ1-525p was soluble when co-expressed with N-terminus (Figure 3.18). This soluble fraction was purified using a Ni-NTA column and tested for Glu-I activity. Soluble Cwh41Δ1-525p did not show Glu-I activity with the trisaccharide substrate when 2 µg of the protein was incubated with 10 nmoles of the substrate for overnight at 37 ˚C ( Table 3.3).  88                      Figure 3.16. SDS-PAGE analysis of pACYC-DUET1-CWHN expression. The blue arrow indicates the expected protein band for soluble Cwhnp.   Figure 3.17. SDS-PAGE analysis of pACYC-DUET1- CWH41Δ1-525 expression. The blue arrow indicates the expected protein band for soluble Cwh41Δ1-525p.  89    Figure 3.18. SDS-PAGE analysis of pACYC-DUET1-CWHN-CWH41Δ1-525 expression. Expected size for Cwh41Δ1-525p and Cwhnp are indicated by red and yellow arrow respectively. The solubility of the Cwh41Δ1-525p was improved by 42% when co-expressed with Cwhnp as observed on this gel. The soluble Cwh41Δ1-525p was detected in Ni-NTA elution and confirmed on western blot while Cwh41Δ1-34p used as a positive control.   Table 3.3. Glu-I assay of the co-expression of Cwhnp and Cwh41Δ1-525p.        Sample Absorption at 450 nm Replicates Blank 0.0866 0.0857 Positive control (Cwh41Δ1-34p ) 0.2446 0.2573 Co-expression of Cwhnp and  Cwh41Δ1-525p 0.0817 0.0776 90    Figure 3.19. Ni-NTA chromatography of Cwh41Δ1-525p. A. Cwh41Δ1-525p expressed from pACYC-DUET1-CWH41Δ1-525 in E. coli. B. Cwh41Δ1-525p co-expressed from pACYC-DUET1-CWHN-CWH41Δ1-525 in E. coli. The solid line represent optical absorption at 280 nm and the dashed line represents concentration of eluting buffer (%). The unbound proteins were washed with 20 mM phosphate buffer pH 7.4 with 20 mM imidazole and 500 mM sodium chloride. In step 1 elution, 20 mM phosphate buffer pH 7.4 with 43 mM imidazole and 500 mM sodium chloride was used to wash the loosely bound contaminant proteins. In step 2, 20 mM phosphate buffer pH 7.4 with 500 mM imidazole and 500 mM sodium chloride was used to elute the bound Glu-I. 91  3.4.4 Trypsin hydrolysis of Glu-I  Trypsin hydrolysis of Glu-I (Cwh41Δ1-34p) obtained after Ni-NTA purification generated a mixture of polypeptides (Figure 3.20). Trypsin-digested Glu-I (after 3 hr of incubation with trypsin) showed about 2.2 times the activity of the undigested enzyme (Figure 3.21). Incubation for 3 hour at 1:1000 of trypsin:Glu-I ratio was optimum for the release of the Glu-I catalytic domain. Trypsin hydrolysis was carried out in duplicates, and the values are reported in mean ± range.  Figure 3.20. Trypsin digestion of Cwh41Δ1-34p. A. Incubation of Glu-I with trypsin at 1:1000 ratio for 3 hr released a mixture of fragments. B. Time course of trypsin digestion observed up to 3 hr. Most of the Glu-I was hydrolyzed within the first hour of incubation, and the remaining Glu-I gradually disappeared as the hydrolysed products become more prominent.  92   Figure 3.21. Effect of trypsin hydrolysis on the activity of Cwh41Δ1-34p. 20 µg of Cwh41Δ1-34p was incubated for 3 hr at 4 °C with trypsin at 1:1000 ratio, and without trypsin but the equal volume of 20 mM phosphate buffer; Glu-I activity was measured using 4 µL from each sample. Error bars indicate mean ± range.   3.4.5 Purification of Glu-I catalytic domain using size exclusion chromatography Trypsin hydrolysis of Cwh41Δ1-34p released a few polypeptide fragments (Figure 3.20). The catalytic domain was purified from the mixture of polypeptides by SEC using a HiPrep-Sephacryl-S100HR column attached to a Åkta Purifier. The separation resulted in three major peaks which were collected into three different fractions (Figure 3.22). The SDS-PAGE analysis of the fractions revealed that the majority of fraction #2 majorly contained a 40 kDa fragment (Figure 3.23). Fraction #2 had activity against trisaccharide substrate suggesting that the 40 kDa fragment contained the Glu-I activity (Table 3.4).   93    Figure 3.22. Separation of the polypeptide fragments generated from trypsin digested Glu-I by size exclusion chromatography using HiPrep-Sephacryl-S100HR column. A major 40 kDa protein with Glu-I activity was present in the fraction 2. Fraction 1 consisted of a 55 kDa protein, and no protein was detected in fraction 3 which eluted after the salts as observed in the conductivity curve (blue line). The absorbance in this fraction could be due to the salts and imidazole carried forward from the Ni-NTA elution.   94   Figure 3.23. SDS-PAGE analysis of the fractions collected during the size exclusion chromatography using HiPrep-Sephacryl-S100HR column. Lane 1 and 6 are molecular weight standards. Lane 2, Ni-NTA purified Glu-I with no trypsin; Lane 3, Ni-NTA purified Glu-I digested with trypsin; Lane 4, fraction 1 collected during the size exclusion chromatography (Figure 3.22); Lane 5, Fraction 2 collected during the size exclusion chromatography. The catalytic domain (as observed in the lane 5) was successfully purified by the size exclusion chromatography using HiPrep-Sephacryl-S100HR column.   Table 3.4. Glu-I activity assay of Cwh41Δ1-525p fraction from size-exclusion chromatography of trypsinised Cwh41Δ1-34p  Sample Absorption at 450 nm (duplicate data) Substrate blank 0.041 0.04 Cwh41Δ1-525p enzyme blank 0.039 0.038 Cwh41Δ1-525p + substrate 0.273 0.271 Cwh41Δ1-34p enzyme blank 0.039 0.037 Cwh41Δ1-34p + substrate 0.273 0.280 95  3.5 Discussion In this chapter, Glu-I truncations were expressed and tested for Glu-I activity against a synthetic trisaccharide substrate to understand the catalytic domain. No Glu-I activity was detected in any of the truncated Glu-I preparations. Similar to previous studies that involved yeast expression systems [12], functional expression of Cwh41Δ1-525p was not successful in E. coli (Figure 3.7). The Cwh41Δ1-525p was not properly folded and accumulated in insoluble inclusion bodies. Based on the structural details, published after these Cwh41Δ1-525p experiments, Barker and Rose (2013) proposed that the lack of expression in yeast of the Cwh41Δ1-525p construct was due to an incomplete domain; the construct begins in a disordered loop between CH4 and CH5 of C-domain and does not include parts of C’-domain and C-domain (Figure 3.1) [147]. However, a construct that codes for a truncated form of Candida albicans Glu-I lacking 419 amino acids from N-terminus and lacking portions of C-domain and C’ region, expressed a functional protein [119]. This C. albicans truncation of Glu-I has approximately 65% homology with S. cerevisiae Glu-I [152]. Conversely, a structural analysis study suggested that the region between 419 to 525 positions is indispensable for the formation of the fully functional domain of Glu-I [147]. However, the inclusion of additional sequence on the N-terminal side did not improve the expression of Glu-I truncations. Cwh41Δ1-314p and Cwh41Δ1-349p constructs begin prior to CH1 of the C-domain. Furthermore, Cwh41Δ1-314p includes LH1 of linker region as well. Glycoside hydrolases such as trehalase and mannosyl glycerate synthase that consists of only a (α/α)6 toroid domain fold correctly into native enzymes [147]. However, the (α/α)6 toroid domain of Glu-I seems atypical and unable to fold on its own. As Glu-I significantly differs from other (α/α)6 toroid enzymes in biochemical properties and primary sequence, it may be that the yeast Glu-I requires an additional section of the N-terminal region for its native folding.  96  Molecular chaperones play an essential role in protein folding and thus in inclusion body formation. DnaK- DnaJ, grpE and GroES-EL are known to promote the proper isomerisation and folding of target proteins. The solubility of Cellvibrio gilvus β-glucosidase significantly improved upon co-expression of GroEL/ES, and the lowering of induction temperature in combination with co-expression of GroEL/ES further significantly improved the solubility from 20% to 70% of the total insoluble enzyme [185]. However, co-expression of GroES/EL chaperones did not improve the solubility of the Glu-I truncations (Figure 3.12 and Figure 3.13). Overexpression of trigger factor prevents the aggregation of the recombinant proteins and formation of inclusion bodies [186]. However, in the current work, co-expression of trigger factor did not prevent the formation of insoluble bodies of Glu-I truncations (Figure 3.14).    The solubility of Cwh41Δ1-525p significantly improved with the co-expression of the N-terminal domain- Cwhnp (Figure 3.16). The increase in the solubility could be due to the interaction of the N-terminus with the catalytic domain. The N-terminus might have assisted the C-terminal domain in folding; however, no Glu-I activity was observed in the crude-extract or the purified fractions of soluble Cwh41Δ1-525p ( Table 3.3). This lack of Glu-I activity may be because the CWH41Δ1-525 construct starts within a disordered loop that spans two helices and fails to form the full (α/α)6-barrel that is a part of the catalytic domain as postulated by Barker and Rose [147].  Confirming previous reports, the C-terminal catalytic domain of Glu-I displayed about 2.2 times more activity than the whole enzyme when it was generated by trypsin hydrolysis (Figure 3.21) [12]. Since the heterologous expression was not successful, isolation of the catalytic domain from trypsin-hydrolysed Glu-I was used as an alternative method to generate the Glu-I catalytic domain for subsequent use to study the structure-function of Glu-I. The activity of the C-terminal catalytic 97  domain also suggests that the N-terminal domain does not play a direct role in catalysis. However, it may be essential for proper assembly and folding of Glu-I, as opposed to simple (α/α)6 toroid proteins.  Even though Glu-I has 83 trypsin recognition sites in the catalytic domain, its catalytic domain (37 kDa C-terminal fragment) is mostly resistant to trypsin hydrolysis. However, due to the presence of a trypsin recognition site at Arg832 at the C-terminus, the His-tag and other fusion partners of the recombinant Glu-I at the C-terminus were hydrolysed, making un-tagged proteins hard to purify. The theoretical pI of the N-terminal and C-terminal fragments only differ by 0.2 therefore isolating the catalytic domain from this mixture via ion exchange chromatography is difficult. Initial efforts to purify the catalytic domain from the trypsin hydrolysate of the Cwh41Δ1-34p were not feasible due to the limitations of the gel-filtration chromatography column (SEC70). However, this problem was solved using a different SEC column- HiPrep-Sephacryl-S100HR column (Figure 3.22). The isolated catalytic domain showed activity against the synthetic trisaccharide substrate. However, further in-detail functional studies of this catalytic domain are needed. This study establishes a method to isolate the catalytic domain of Glu-I and opens the doors to further manipulations and to development of this domain as a model to understand the role of the N-terminal domain of Glu-I and the reasons for the superior activity of the catalytic domain.   98  Chapter 4: Identification of the catalytic base of processing α-glucosidase-I  99  4.1 Summary Processing α-Glucosidase-I is an ER membrane protein that plays a critical role in N-glycosylation and quality control of protein folding. Generally, hydrolysis of a glycosidic bond takes place via acid/base catalysis with either net inversion or retention of anomeric configuration. Glu-I acts via an inverting mechanism that involves a pair of carboxylic amino acids, where one residue acts as a nucleophile (catalytic base) and the other one as a proton donor (catalytic acid), which are typically 10 Å apart. Primary sequence alignment of the catalytic domain of Glu-I orthologs revealed six highly conserved carboxylic acid residues (D601, D602, E613, D617, D670 and E804). Single mutants (D601A, D601N, D602A, D602N, E613A, E613Q, D617A, D617N, D670A, D670N, E804A and E804Q) and double mutants (D601A-E804A and D601N-E804Q) of these conserved residues were generated using site-directed mutagenesis. Mutations of all carboxylic acid residues except D670 led to the loss of Glu-I activity. All mutants and wild-type Glu-I showed similar secondary conformations as determined by circular dichroism spectra. Addition of azide or formate as an external nucleophile for E804A restored the α1,2 glycosyl hydrolysing activity. The similar rescue was not observed with the E804Q or double mutant D601A-E804A. Therefore, this study identifies E804 as the catalytic base of Glu-I. 100  4.2 Introduction Mannosyl-oligosaccharide glucosidase (MOGS) or Glu-I belongs to glycoside hydrolase family 63 and acts on the α1,2-glycosidic bond of N-linked glycan precursors, releasing the terminal glucose residue. Hydrolysis of the glycosidic bond requires two carboxyl residues with either retention or inversion of the configuration of the anomeric carbon of the hydrolysed glycosidic bond [173]. Both inverting and retaining mechanisms typically employ a pair of carboxylic acids with unique roles. In the inverting mechanism, one carboxylic residue acts as a general base (nucleophile) and the other one as a general acid, and the substrate undergoes net inversion of anomeric configuration during catalysis. In retaining glycosidases, one carboxylic residue acts as the acid/base catalyst and the other as a nucleophile. In retaining glycosidases, the nucleophile is in close vicinity (5 Å) to the anomeric carbon. However, in inverting glycosidases, due to the involvement of a nucleophilic water molecule, the average distance between the carboxyl residues is approximately 9 Å [187]. Glu-I catalysis occurs through net inversion of anomeric configuration and releases a β-D-glucose from the N-linked glycans [155]. In yeast Glu-I, the C-terminus (F525-F833) contains the catalytic domain [152] and shares high homology with human Glu-I (48% identity) and other orthologs. There are six highly conserved carboxylic acid residues, D601, D602, E613, D617, D670 and E804, in the catalytic domain of Glu-I [12]. With the aid of amino acid modification studies on rat Glu-I, a highly conserved region between E592RHLDLRCW600 was reported to be essential for the substrate binding, and the amino acids in this region were reported to be participants in the acid-base catalysis. This highly conserved region has two carboxylic residues: E613 and D617. Site-directed mutagenesis of E613 and D617 led to the loss of function; however, they could not establish whether these were the catalytic acid and base [12]. Barker and Rose reported the structures of yeast 101  Glu-I (Cwh41Δ1-34p) and its mutant D601N [147, 188]. Due to the failed efforts of co-crystallisation and crystal soaking experiments, they performed in-silico ligand docking of Cwh41Δ1-34p using the sugars that form the natural substrate of Glu-I: α-D-glucose, α-D-glucose-(1,2)-α-D-glucose-(1,3)-α-D-glucose-(1-methyl), and inhibitors:  deoxynojirimycin, kojibiose and miglitol. Monosaccharide binding revealed two possible binding sites in the centre of the Cwh41Δ1-34p catalytic domain- Site A, and a nearby site B. Site B which is 12 Å away from site A does not possess any carboxylic acid residues which typically play an essential role in the hydrolysis of glycosidic bond. Hence, site B is not a catalytic site. However, it could potentially be a binding site for the third and non-terminal glucose of the substrate. All top hits from the docking of ligands were observed in site A including the contacts with two tryptophan residues (W748 and W822), and contacts with two carboxylic residues (D601 and E804). Based on these docking studies and site-directed mutagenesis studies, Barker et al. (2013) proposed the carboxylic residues- D601 and E804 as the catalytic residues [147]. However, there is no information on kinetic analyses of the mutations at these positions to establish the catalytic model. Hence, the current study was designed to confirm the identity of the catalytic residues, to further our understanding of Glu-I mode of action. Our approach involved the following screening criteria for potential catalytic (carboxyl) residues - • The carboxylic acid residue must be conserved and present in the catalytic domain. • The catalytic acid and base should be in 6-11 Å apart. • There should be a loss of activity when the base is replaced with another non-nucleophilic residue. • The activity of the enzyme with a non-nucleophilic base mutation should be restored by an exogenous nucleophile (nucleophile rescue). 102  Restoration of the function of a mutant enzyme by the addition of small exogenous compounds is called chemical rescue. The chemical rescue has been extensively employed to probe the mechanism of various glycoside hydrolases [189, 190]. Small compounds such as azide, formate and propionate have been extensively used in the restoration of function of glycosidase mutants. These anions act as a nucleophile and surrogate the function of the mutated residue. Using this approach, the current study aimed at the identification of glutamic acid (E804) as the catalytic base in the hydrolysis of the α1,2- glycosidic bond by Glu-I.  103  4.3 Materials and methods 4.3.1 Homology and in-silico analysis Homology analysis of orthologs of Glu-I was carried out using T-Coffee accurate alignment (http://tcoffee.crg.cat/apps/tcoffee/do:accurate) [191]. The distances between the conserved carboxylic amino acids were calculated from crystal structures of Glu-I (PDB 4j5t) using UCSF Chimera molecular visualisation software [192]. Changes in steric hindrances due to the change in the side chains of mutated residues were also examined using the calculator plugin in UCSF Chimera software.  4.3.2 Site-directed mutagenesis All six conserved carboxylic residues (D601, D602, E613, D617, D670 and E804) were mutated to either their amide counterpart or alanine. Site-directed mutagenesis was carried out using a Q5® site-directed mutagenesis kit (New England Biolabs, Canada) and pET30a-CWH41Δ1-34 as a template with the primers listed in Table 4.1 Mutant pET30a-CWH41Δ1-34 plasmids were transformed into E. coli DH5α and screened for the presence of CWH41Δ1-34 gene. Mutations were confirmed by DNA sequencing.     104  Table 4.1. List of mutant primers   Mutant primer Primer sequence (5’-3’) D601A sense AAGCGGTATGGCAGACTATCCTAG D601A anti-sense GGCAAACAATGAGTGAAG D601N sense AAGCGGTATGAATGACTATCCTAG D601N anti-sense GGCAAACAATGAGTGAAG D602A sense CGGTATGGATGCATATCCTAGAGC D602A anti-sense CTTGGCAAACAATGAGTG D602N sense CGGTATGGATAACTATCCTAGAG D602N anti-sense CTTGGCAAACAATGAGTG E613A sense AGATGTAGCAGCATTGAACGTAGAC E613A anti-sense GGTGGTTGTGCTCTAGGATAG E613Q sense ACAACCACCAGATGTAGCACAGTTGAACGTAGACGCATTAG E613Q anti-sense TGTTGGTGGTCTACATCGTGTCAACTTGCATCTGCGTAATC D617A sense ATTGAACGTAGCAGCATTAGCATGGG D617A anti-sense TCTGCTACATCTGGTGGTTG D617N sense ACCAGATGTAGCAGAATTGAACGTAAACGCATTAGCATG D617N anti-sense CATGCTAATGCGTTTACGTTCAATTCTGCTACATCTGGT D670A sense TTGCTACTGTGCAATTAGCATCG D670A anti-sense TTGTCATTTTCACTCCAGTG D670N sense GTGAAAATGACAATTGCTACTGTAATATTAGCATCGATCCAGAAGAC D670N anti-sense GTCTTCTGGATCGATGCTAATATTACAGTAGCAATTGTCATTTTCAC E804A sense TTATTGTTATGCAAATTACAGTCCGATAG E804A anti-sense CCTTGTTCTTCCCAAACTTTG E804Q sense GTTTGGGAAGAACAAGGTTATTGTTATCAGAATTACAGTCCGATAGAT GGTC E804Q anti-sense GAC CAT CTATCG GACTGT AAT TCT GAT AAC AAT AAC CTT GTT CTT CCC AAA C 105  4.3.3 Expression and purification of wild-type Glu-I and its mutants Wild-type and mutant plasmids were transformed into E. coli BL21(DE3) for expression studies. Expression was carried out as previously described in section 2.3.5 in Chapter 2. Expressed enzymes were purified using a Ni-NTA column as described in section 2.3.6 in Chapter 2.  Double mutants (D601A-E804A and D601N-E804Q) were also generated using the corresponding primers and single mutant plasmids as templates.   4.3.4 The Glu-I activity of mutants All mutants were screened for Glu-I activity by incubating with synthetic trisaccharide substrate for short (30 min) and long (12 hr) durations. The released glucose was then measured as described in section 2.3.9 in Chapter 2.  4.3.5 Secondary structure Circular dichroism (CD) measurements were performed using a Jasco J810 spectropolarimeter equipped with Peltier temperature control. All measurements were carried out at 25 ˚ C. CD spectra of native Glu-I and its mutants (2 s response time, 50 nm/min scan rate, and 5 scan average) at 2 µM concentrations in 20 mM phosphate buffer, pH 6.8, were obtained using a 0.1 cm path length cuvette. 4.3.6 Nucleophile rescue of activity of Glu-I mutants Glu-I is an inverting glycosidase and carries out the hydrolysis of glycosidic bond through general acid/base catalysis. Hence, anion rescue was carried out to identify the catalytic base using exogenous nucleophiles azide and formate. Both azide and formate were used due to their differences in restoration efficiencies [153]. As metal contaminants react with azide and interfere with the assay, metal contaminant-free azide solution was prepared using plastic spatulas and the addition of 50 µM EDTA. The nucleophile preparations were made using 20 mM phosphate buffer 106  pH 6.8, and pH was adjusted after dissolving the azide or formate. A 6 µg aliquot of each of the alanine mutants and native form of Cwh41Δ1-34p were mixed with 1M and 0.1M (final concentration) nucleophile, and assayed immediately for activity using 10 nmoles of substrate for 30 min at 37 ˚C and also after an overnight (12 hr) incubation.  Nucleophile rescue experiment was carried out in duplicates, and the values are reported as mean ±range.   4.4 Results 4.4.1 Homology and in-silico analysis Amino acid sequence alignment of the catalytic domain of all eukaryotic Glu-I orthologs (six are shown in Figure 4.1) and a prokaryotic ortholog (Ygjk; not shown in the figure) revealed six conserved carboxylic acid residues D601, D602, E613, D617, D670 and E804 (Figure 4.1). All six carboxylic residues are located in the catalytic domain (Figure 4.2). The distance between the -COO- groups of the conserved carboxylic residues, was calculated using the UCSF Chimera molecular visualisation tool (Figure 4.3). The residues E613 and D670, and D601 and D602 are separated by 8 and 9 Å respectively. The residues E804 and D601, E804 and D602, and D601 and D617 are separated by 10 Å. D617 and E804, E613 and E804, and D670 and E804 are separated by 16, 23 and 20 Å, respectively (Table 4.2). 4.4.2 Site-directed mutagenesis Initially, 12 mutants were generated for six conserved carboxylic residues (replaced by alanine and amide form) using site-directed mutagenesis. The integrity of the mutants was checked through DNA sequencing. These mutants were subsequently used for expression and 107  characterisation. Based on the structural analysis reported by Barker and Rose (2013) and the homology analyses, double mutants for D601 and E804 were also generated.   Figure 4.1. Homology analysis of C-terminal catalytic domain of Glu-I orthologs. T-Coffee alignment of eukaryotic Glu-I orthologs. The pink colour indicates the regions with high homology. Regions with conserved carboxylic residues are highlighted in boxes  4.4.3 Purification of wild-type and mutant Cwh41Δ1-34p Expression was carried out in E. coli BL21(DE3), and mutant protein was purified using AKTA-purifier system equipped with Ni-NTA column. All mutants showed the expected protein band at 100 kDa on SDS-PAGE, and the purified protein had the expected purity level as the wild-type Cwh41Δ1-34p. Purified mutants were further analysed for Glu-I activity using synthetic trisaccharide as substrate. 108   Figure 4.2. Orientation of conserved carboxylic residues in Glu-I active site. UCSF-Chimera was used to generate the picture   Figure 4.3. The distance between carboxylic residues. The dotted yellow line connects the corresponding amino acids. Amino acids were labelled in blue.  109  Table 4.2. Distances between different conserved carboxylic residues  From  To Distance (Å) D617 E804 16 E613 E804 23 E613 D670 9 D670 E804 20 D601 E804 10 D602 E804 10 D601 D617 10 D601 D613 13 D602 D601 8  4.4.4 Secondary structure analysis of mutants Expression and purification of wild-type and all mutant proteins were carried out in a similar fashion using pET30a(+) expression vector in host E. coli BL21(DE3). Absorption in the far UV region (190-250 nm) is typically due to the peptide bond. Different secondary structure elements- α-helix, β-sheet, and random coil structures lead to spectra with different shapes. Analysis of far UV CD spectra of all the mutant Glu-I enzymes was carried out, and these were compared with the spectra of wild-type Glu-I. The CD data indicated that the wildtype and mutant proteins all had similar conformations and no significant alterations from the native secondary structure were detected (Figure 4.4). Therefore, the purified mutants appeared to be properly folded and were used for the activity studies.  110  4.4.5 The Glu-I activity of mutants All mutant preparations before and after purification were screened for Glu-I activity. Four microliters of enzyme sample (6 µg of protein) were incubated with 10 mM synthetic trisaccharide substrate for overnight at 37 °C. Only D670A and D670N were active while all other mutants did not show activity after overnight incubation time (Figure 4.5).   Figure 4.4. Circular dichroism of wild-type and mutant Cwh41Δ1-34p. There were no significant changes in the secondary structure of the wild-type and mutant enzymes.       111   Figure 4.5. Glu-I activity of mutants of Cwh41Δ1-34p. Error bars indicate mean± range. 6 µg of enzyme was incubated overnight with 10 nmoles of trisaccharide substrate and the released glucose was estimated.  4.4.6 Chemical rescue Nucleophile rescue was performed to identify the catalytic base. The wild-type enzyme did not show any significant change in activity with the addition of 0.1 M azide, 1 M azide or 1 M formate (Figure 4.6). A substantial increase in activity of the mutant E804A from 0.4 nmoles to 2.3 nmoles and 2.8 nmoles of released glucose was observed upon the inclusion of 1 M azide and 1 M formate respectively when added into the reaction mixture with a 12 hr incubation with the substrate (Figure 4.6). However, a similar “rescue” effect was not observed at shorter incubation of 30 minutes. Neither formate nor azide rescued the lack of activity in the E804Q and D601A-E804A mutants (data not shown). Nucleophile rescue was also not observed with any of the other Glu-I mutants (Figure 4.7). Further kinetic analysis to determine Kcat/Km was not carried out due to the scarcity of the synthetic trisaccharide substrate.   112   Figure 4.6. Nucleophile rescue of Glu-I activity of E804A mutant of Cwh41Δ1-34p. 1. Cwh41Δ1-34p, 2. Cwh41Δ1-34p + 1M azide, 3. E804A mutant 4. E804A mutant + 1M formate, 5. E804A mutant + 1M azide. Error bars indicate ± range. 6 µg of enzyme was incubated overnight with 10 nmoles of substrate in the presence or absence of nucleophile (1 M). The data is presented in percentage of relative activity in comparison to wild-type Cwh41Δ1-34p activity determined simultaneously.    Figure 4.7. Nucleophile rescue of Glu-I activity of mutants of Cwh41Δ1-34p. Error bars indicate ± range. 6 µg of enzyme was incubated overnight with 10 nmoles of substrate in the presence or absence of azide as a nucleophile (1 M). The data is presented in percentage of relative activity in comparison to wild-type Cwh41Δ1-34p activity determined simultaneously.  113  Table 4.3. Catalytic mechanism and catalytic acid/base of inverting glycoside hydrolases   Clan GH family Mechanism Nucleophile (Base) Proton donor (Acid) Example Rescue Reference GH-G 37 Inverting  Glu (inferred) Asp (inferred) Trehalase - [193] GH-G 63 Inverting - - Glu-I E804A  1M formate 1M azide Current study GH-L 15 Inverting Glu (experimental) Glu (experimental) Glucoamylase - [194, 195] GH-L 65 Inverting Histidine (inferred) Glu Maltose phosphorylase  [196] GH-L 125 Inverting Glu (inferred) Asp (inferred) Exo alpha1,6 mannosidase - [197] GH-M 8 Inverting Glu (inferred) Glu (experimental) Chitosanase  [198] GH-M 48 Inverting Asp (experimental) Glu Cellulase (Cel48A) 0.4M azide [199]    Glu (inferred) Glu (inferred) Cellulase (Cel48F)  [200] 114   Figure 4.8. Schematic representation of azide rescue. A. Glu-I mechanism of action on synthetic trisaccharide. B. A possible mechanism of action of E804A mutant of Glu-I in the presence of azide.    4.5 Discussion Glu-I catalyses the hydrolysis of α1,2-glycosidic bond from N-linked glycans of nascent proteins that play different biological functions [37]. Despite its physiological significance, some basic characteristics of the catalytic mechanism have not been established. Glu-I is a GH family 63 glycosidase and hydrolyses the α1,2-glycosidic bond via inverting mechanism [155]. This hydrolysis reaction is carried out by a pair of carboxylic residues through acid-base catalysis [201]. Sequence homology of the C-terminal catalytic domain of eukaryotic Glu-I orthologs revealed six highly conserved carboxylic amino acids (Figure 4.1). Structural studies of Ygjk, a member of GH 115  family 63 and a hypothetical protein from E. coli K12, proposed the possible role of conserved carboxylic residues at D501 and E727 in catalysis. Based on primary sequence alignment, the D501 and E727 of Ygjk correspond to D617 and E804 of yeast [124]. However, Barker and Rose (2013) proposed D601 and E804 as the catalytic pair based on the in silico substrate docking studies of Cwh41Δ1-34p [147]. These residues are highly conserved among Glu-I from different phyla. Moreover, a recent survey of distances between catalytic carboxylic acid residues in α-inverting glycosidases revealed the mean distance as 8 ± 1.5 Å with a minimum of 6.0 and maximum of 10.2 Å [187]. When this distance parameter is applied to the conserved carboxylic residues in Glu-I, only the pairs D601-D602, D601-D617, D601-E804, D602-E804, E613-D670 have the potential to be catalytic residues (Figure 4.3). This criterion excludes the possibility of E613-E804 and D617-E804 pairs, as previously suggested [12, 124]. However, because the enzymes can change their conformation with the interaction of substrate via an induced fit mechanism, we cannot completely rule out these amino acids as catalytic residues. CD spectra analyses revealed that wild-type and mutant proteins (alanine mutants- D601A, D602A, E613A, D617A, D670A, E804A and amide mutants- D601N, D602N, E613Q, D617N, D670N, E804Q) have similar secondary structures with no major conformational changes due to site-directed mutagenesis; this eliminated the possibility of loss of function due to drastic conformational disturbances. Even though CD spectroscopy is a rapid tool to determine the secondary structure of proteins, it has limitations in detecting the subtle changes in the conformation. The amount of similar secondary structure does not translate directly to the similar tertiary structure. Hence, further structural analyses of non-functional mutants in the presence of substrate or inhibitor would be helpful to completely rule out the potential effect of conformational discrepancies. 116  With the exception of D670, mutations of all amino acids tested (D601, D602, E613, D617, D670, E804), resulted in inability to hydrolyse the trisaccharide substrate. This indicates that D670 does not play a role in either substrate binding or in catalysis. Moreover, D670 did not have any interactions with either inhibitor or substrate as revealed by in-silico docking studies [147]. The remaining residues D601, D602, E613, D617 and E804 may have either direct or indirect roles in the function of the enzyme. The loss of Glu-I catalytic function of these mutants may be due to the elimination of either the general acid and base function or may be due to the loss of carboxylic acid residues that have non-catalytic roles as observed in D55G and D324N mutations in Glu-I structural homologs glucoamylase and Ygjk respectively [202]. These details should also be taken into consideration as Glu-I shares significant structural homology with glucoamylase (DALI score 20.5) and Ygjk (DALI score 30.7) [147]. Distance matrix alignment (DALI) is a popular structure alignment tool that helps in identifying similar structures from the PDB and the DALI score is a measure of structural similarity. As similar folds often share the similar function, measurement of structural homology using tools such as DALI would be helpful in identifying functional orthologs. Ygjk is the closest ortholog of Glu-I with a few variations in domain orientation. The activity of the E804A mutant was rescued by both 1M azide and 1M formate in the presence of the synthetic trisaccharide substrate (Figure 4.6). As observed in this study and other previous studies, the addition of exogenous nucleophile such as azide can only rescue a minor percentage of the original catalytic activity. Azide and formate did not show any rescue effect on remaining non-functional single mutants (D601, D602, E613, and D617) and double mutants (D601A-E804A and D601N-E804Q). In contrast to the E804A mutant, the nucleophile rescue effect was not observed with the E804Q glutamine mutant. This could be due to the tight spatial arrangements arising from the side chain of the glutamine with no space to accommodate the nucleophile in the 117  active site, whereas the additional space formed due to the replacement of glutamic acid with the smaller alanine could have accommodated the exogenous nucleophile. The nucleophile rescue of E804A mutant allows us to assign the role of the catalytic base to the glutamic acid at 804;  this result also aligns with the previous proposals of the Glu-I catalytic residues by Barker [147]. Moreover, several inverting glycoside hydrolases have glutamic acid as the catalytic base (Table 4.3). Hydrolysis of the α1,2-glycosidic bond by nucleophile rescued E804A may take place via one of the two possible ways- 1. via direct attack of external nucleophile/catalytic base on the glycosidic bond and release of β-glucosyl azide or 2. water molecule deprotonation by an external nucleophile acting as a catalytic base. The Glu-I assay employed in the current study utilises the glucose oxidase that converts glucose to gluconic acid. This gluconic acid formation may not be possible if the reaction takes place via direct displacement by the external nucleophile. Hence, the hydrolysis of the α1,2-glycosidic bond by E804A in the presence of the external nucleophile likely takes place via the latter route where a deprotonated water molecule acts as the catalytic base. The combination of kinetic and structural studies of D601A and E804A in the presence of activated substrate and inhibitors would give clues as to the catalytic acid residue.  Structural studies involving conformational changes caused by substrate/mimic binding in combination with studies involving non-hydrolysable substrate analogues or mimics such as glycosyl fluorides that trap the enzyme intermediates would immensely help in understanding the mechanism of Glu-I catalysis.  NMR spectroscopy would be a great tool to study the properties of these mutants. However, the catalytic C-terminal domain of these enzymes must be isolated to fit within the molecular weight limitations of the traditional NMR techniques [203]. NMR spectroscopy offers some advantages over other methods such as X-ray crystallography in studying the structure-function studies which involve measurement of effects of the substrate or various inhibitors on the conformation of the 118  enzyme. NMR spectroscopy can be carried out on a protein in buffered solution rather than a static structure, and the method offers a chance to study the enzyme under conditions that most closely mimic the natural functional state.    119  Chapter 5: Overall conclusions and future perspectives  120  5.1 Overall Conclusions A robust expression method for soluble S. cerevisiae Glu-I was established using E. coli as an expression host and pET30a(+) as an expression vector. Optimal expression of Glu-I was attained with 1 mM IPTG concentration and induction at 25°C. The recombinant enzyme was purified to 95% homogeneity using Ni-NTA affinity column chromatography. The purified enzyme acted on a synthetic trisaccharide substrate and showed a specific activity of 3370 U/mg protein. Glu-I showed a Km of 1.27 mM with a synthetic substrate. Recombinant Glu-I retained strict substrate specificity and did not hydrolyse kojibiose, p-nitrophenyl and methyl-umbelliferyl α-glucosides. Codon optimisation of the CWH41 gene improved the codon adaptability index from 0.64 to 0.88. A higher codon adaptability index often improves the expression of the target protein, but in the case of Glu-I, it had no significant effect on expression levels.  Glu-I is a type-II membrane-bound protein and has the catalytic region in the C-terminal domain. The catalytic domain, generated during trypsin digestion, had 2.2 times more catalytic activity than that of the full enzyme. Expression of Glu-I truncations Cwh41Δ1-314p, Cwh41Δ1-349p and Cwh41Δ1-525p were carried out using the previously established system for the expression of Cwh41Δ1-34p and resulted in protein accumulation in inclusion bodies. Co-expression of solubility enhancing tags (MBP- tag and GST- tag) and molecular chaperones (GroES-GroEL, DnaJ-DnaK, GrEp, and Trigger factor) did not result in an improvement in solubility. Co-expression of the Glu-I N-terminal domain (Cwhnp) and Cwh41Δ1-525p using the pACYCDuet1 dual expression vector significantly improved the solubility. A significant percentage (~42%) of the expressed Cwh41Δ1-525p was soluble when co-expressed with Cwhnp compared to the completely insoluble expression of Cwh41Δ1-525p when expressed without Cwhnp. However, the 121  soluble Cwh41Δ1-525p did not hydrolyse the α1,2-glycosidic bond of the trisaccharide substrate. Due to the lack of activity of the soluble Cwh41Δ1-525p, a method was developed to generate the catalytic domain of Glu-I by trypsin hydrolysis of Cwh41Δ1-34p. The catalytic domain was active against synthetic trisaccharide substrate when cleaved off from the Cwh41Δ1-34p and purified by size exclusion chromatography.  Sequence alignment of the catalytic region of Glu-I orthologs revealed six highly conserved carboxylic acid residues (D601, D602, E613, D617, D670 and E804). Except for D670, all other carboxylic acid residues lost Glu-I activity after replacement by alanine or by their amide-forms. CD spectroscopy detected no major conformational deviations from the secondary structure of the mutants. The α1,2 glycosyl hydrolysing activity of the E804A mutant was rescued by the addition of either 1M azide or 1M formate; therefore, this study identifies E804 as the putative catalytic base of Glu-I. 5.1.1 Yeast Glu-I as a model for family 63 glycosyl hydrolases Based on the substrate specificity, GH 63 family enzymes have been classified into three sub-families- the processing α-glucosidase-I subfamily, the MGH subfamily (Glu-I from thermophilic bacteria) and the GGalase (bacterial Glu-I including E. coli Ygjk) subfamily [74]. MOGS and yeast Glu-I belong to processing α-glucosidase-I subfamily, show significant homology, and similar chemical properties. Protein homology modelling of MOGS using Phyre2 server also reveals that both the yeast and human enzymes share structural properties (Figure 5.1). Moreover, alignment of the amino acid sequences of the catalytic domain of these enzymes reveals high similarity in this region (Figure 5.2). Hence, the yeast enzyme can be used as a model for the human MOGS. The observed results or the information obtained on the structure-function of yeast 122  Glu-I may have potential to be interchangeably applied to the processing α-glucosidase-I subfamily of family 63 glycoside hydrolases that includes human Glu-I.    Figure 5.1. Structures of processing α-glucosidase-I subfamily of Family 63 glycosyl hydrolases. A. Structure of S. cerevisiae Glu-I (PDB 4j5t) B. Phyre2 predicted structure of C. albicans Glu-I C. Phyre2 predicted structure of human Glu-I. D. Superimposition of all the three structures (blue- S. cerevisiae Glu-I, purple- C. albicans Glu-I, golden- human Glu-I).   123   Figure 5.2. Alignment of C-terminal domain of yeast Glu-I (Sc) and human MOGS (Hs). The (*) indicates identical match in both the sequences, (:) indicates match of size and hydropathy, and (.) indicates match of size or evolutionary preserved hydropathy. The absence of any of these marks indicates a mismatch. C-terminal domain of Glu-I shares 26% identity, 48% similarity based on matches of amino acids with size and hydropathy, and 61% similarity based on evolutionary preserved hydropathy.  124  5.2 Future perspectives There are still many questions regarding how the Glu-I catalyses the hydrolysis of the α1,2-glycosidic bond.  5.2.1 Expression of the catalytic domain. The Glu-I truncations expressed in E. coli were not soluble. Stepwise truncations with the gradual addition of one structural element (β-sheet) at a time starting from CH4 towards the N-terminus would be another alternative to find out the region that is essential for the folding or the functional expression of the Glu-I.  Although the trypsin hydrolysis method yields the Glu-I catalytic domain from full protein, it may not always be a homogenous preparation as the region that flanks the catalytic domain has several recognition sites for trypsin. Another alternative would be to use genetic engineering to insert a protease recognition site at different intervals from 524 towards C-terminus. This would help to cleave and generate a homogenous preparation of catalytic domain efficiently. Moreover, this would also provide details about the minimal protein that is required to catalyse the hydrolysis of the glycosidic bond. However, it is not clear if the catalytic domain isolated in this manner would show enhanced activity. C-terminal sequencing of the catalytic domain obtained via trypsin hydrolysis would give information on whether the C-terminal trypsin hydrolysis is crucial for the enhanced activity of the catalytic domain, and help in designing further studies on C-terminal truncations of Glu-I. 5.2.2 Substrate specificity So far, three structures have been reported for the Glu-I orthologs. All these structural variants have an (α/α)6-barrel catalytic domain and vary in the C’ domain (Figure 3.1). Despite the presence of the (α/α)6-barrel catalytic domain, all these enzymes show different substrate 125  specificities/preferences due to sequence variation. The catalytic domain (37 kDa) isolated from yeast Glu-I that lacks part of the C’ domain (C’H1 and C’S1 to C’S4) retained activity and substrate specificity, and could not hydrolyse kojibiose or 4-methyl umbelliferyl α-glucoside (Table 2.3Error! Reference source not found.). Therefore, the remaining part of the C’ domain (C’5 to C’8) may be critical for the substrate specificity. Deletion studies of the C’ domain or the making of chimeric Glu-I with parts of C’ domain of the relatively non-specific Ygjk may give insights into substrate specificity of the Glu-I. This would also allow the possibility of tuning the specificities by making chimeric enzymes of Glu-I with broad or narrow substrate specificities.  5.2.3 Catalytic mechanism Based on the homology and structural information, catalytic mutants were generated, purified, characterised and tested for Glu-I activity. With the nucleophile rescue experiments, I have identified the glutamic acid at 804 as the catalytic base. Further kinetic analyses of the nucleophile rescued Glu-I activity need to be carried out to estimate the Kcat/Km. These kinetic parameters would help in understanding the rescue of Glu-I activity and the role of glutamic acid 804 in Glu-I function. The substrate binding model and the non-catalytic residues that are important for function still need to be identified and experimentally verified. Isolation of smaller catalytic domain opens the possibility of studying Glu-I enzyme structure using NMR spectroscopy. NMR can be a robust tool to study the enzyme and substrate or inhibitor interactions. The co-crystallisation of enzyme and substrate or inhibitor or soaking of the crystals of wild or non-functional mutant Glu-I with the substrate and inhibitors are other approaches. However, these methods are often tedious and need high amounts of the substrate or inhibitor. Moreover, the outcomes of protein crystallisation are unpredictable. As mentioned earlier in section 4.5 in chapter 4, NMR spectroscopy can be carried out on the 35 kDa C-terminal catalytic domain under 126  functional conditions in buffered solution rather than obtaining information on a static crystal structure. NMR experiments are useful for studying the chemically specific, local conformational changes of the proteins or macromolecules.  The catalytic domain generated via trypsin hydrolysis from Glu-I showed superior activity (about 2.2 times) than the full enzyme. This naturally leads to the question of why the catalytic domain shows increased activity upon release. There could be a few possibilities- conformational differences with the full enzyme, a regulatory role of N-terminus on the function of Glu-I, or the steric-hindrance of the N-terminus restricts the entry of substrate. Structural analyses of the catalytic domain in combination with synthetic trisaccharide substrate or inhibitors such as deoxynojirimycin, kinetic and substrate specificity analyses using substrates that have different alpha glycosidic bonds and hexose sugars can provide answers to the question.  5.2.4 Glu-I - OST - Alg10p and their interactions  Most protein therapeutics are glycoproteins. These glycoprotein therapeutics are required to be homogeneous glycol-forms. The LLO is made in the ER where all the OST and processing glycosidase activities are located. Alg10p that catalyses the addition of terminal glucose onto the Glc2Man9GlcNAc2 may also be involved in the protection of the glycan chain from the processing activities before the transfer on to the nascent proteins by OST. Even though all these enzymes bind and act on glycan chains, currently there is no specific information available on their interactions and how they coordinate the different reactions they carry out. Identifying what triggers the release of Alg10p, and binding of Glu-I is critical for our understanding of N-glycosylation. Research on Glu-I and its interactions with OST and Alg10p would enhance our understanding of glycosylation. The harnessing of such knowledge would greatly help to generate uniformly glycosylated therapeutics. 127  Bibliography [1] L.S. Grinna, P.W. Robbins, Glycoprotein biosynthesis. Rat liver microsomal glucosidases which process oligosaccharides., The Journal of Biological Chemistry, 254 (1979) 8814-8818. [2] R.D. Kilker, B. Saunier, J.S. 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Wider, Structure determination of biological macromolecules in solution using nuclear magnetic resonance spectroscopy, Biotechniques, 29 (2000) 1278-1282, 1284-1290, 1292 passim.   150  Appendix Sequence alignment of CWH41Δ1-34 (wildtype) and CWH41Δ1-34 codon optimised (OPT) wildtype       1 ATGGAAGAATATCAAAAGTTCACGAATGAATCTTTACTGTGG   42  OPT            1 ATGGAAGAATACCAGAAATTTACCAACGAATCGCTGCTGTGG   42  cons           1 *********** ** ** ** ** ** *****  * ******   42   wildtype      43 GCACCGTATAGATCCAATTGTTACTTTGGTATGAGGCCCAGA   84  OPT           43 GCACCGTACCGCTCTAACTGCTACTTTGGCATGCGTCCGCGC   84  cons          43 ********  * ** ** ** ******** *** * **  *    84   wildtype      85 TATGTCCATGAAAGTCCACTAATTATGGGTATCATGTGGTTC  126  OPT           85 TATGTCCATGAAAGTCCGCTGATTATGGGTATCATGTGGTTT  126  cons          85 ***************** ** ********************   126   wildtype     127 AACAGTTTGAGTCAGGATGGCTTACATTCGTTAAGACATTTT  168  OPT          127 AACAGTCTGTCCCAGGATGGCCTGCATTCCCTGCGTCACTTC  168  cons         127 ****** **   ********* * *****  *  * ** **   168   wildtype     169 GCAACGCCTCAGGATAAATTGCAAAAGTATGGTTGGGAAGTG  210  OPT          169 GCCACCCCGCAGGACAAACTGCAAAAGTACGGTTGGGAAGTT  210  cons         169 ** ** ** ***** *** ********** ***********   210    151  wildtype     211 TATGATCCAAGAATTGGTGGTAAAGAAGTTTTTATTGATGAA  252  OPT          211 TATGATCCGCGCATTGGCGGTAAAGAAGTCTTTATCGACGAA  252  cons         211 ********  * ***** *********** ***** ** ***  252   wildtype     253 AAAAATAACTTGAACTTGACTGTTTATTTTGTAAAGAGCAAG  294  OPT          253 AAGAACAATCTGAATCTGACCGTTTACTTCGTCAAGAGTAAG  294  cons         253 ** ** **  ****  **** ***** ** ** ***** ***  294   wildtype     295 AACGGGGAAAATTGGTCAGTGAGAGTTCAAGGTGAGCCTTTG  336  OPT          295 AACGGTGAAAATTGGTCCGTCCGTGTGCAGGGTGAACCGCTG  336  cons         295 ***** *********** **  * ** ** ***** **  **  336   wildtype     337 GATCCCAAGAGACCATCTACAGCATCTGTCGTATTGTACTTT  378  OPT          337 GATCCGAAACGCCCGAGCACGGCTTCTGTGGTTCTGTATTTT  378  cons         337 ***** **  * **    ** ** ***** **  **** ***  378   wildtype     379 AGTCAAAATGGTGGCGAGATAGATGGAAAATCTTCCTTAGCA  420  OPT          379 AGTCAAAACGGCGGTGAAATTGATGGCAAAAGCTCTCTGGCG  420  cons         379 ******** ** ** ** ** ***** ***   **  * **   420   wildtype     421 ATGATAGGTCATGACGGCCCTAATGACATGAAATTCTTCGGA  462  OPT          421 ATGATCGGTCACGATGGCCCGAATGACATGAAATTTTTCGGT  462  cons         421 ***** ***** ** ***** ************** *****   462    152  wildtype     463 TATTCTAAAGAATTAGGCGAGTATCATCTTACAGTAAAGGAC  504  OPT          463 TACTCCAAGGAACTGGGCGAATATCATCTGACCGTGAAAGAT  504  cons         463 ** ** ** *** * ***** ******** ** ** ** **   504   wildtype     505 AATTTTGGTCACTACTTCAAAAATCCGGAATATGAAACCATG  546  OPT          505 AACTTTGGTCACTACTTCAAGAATCCGGAATATGAAACCATG  546  cons         505 ** ***************** *********************  546   wildtype     547 GAAGTAGCACCAGGAAGTGACTGCTCTAAAACAAGTCATTTA  588  OPT          547 GAAGTTGCCCCGGGCAGTGATTGCTCCAAAACGTCACATCTG  588  cons         547 ***** ** ** ** ***** ***** *****    *** *   588   wildtype     589 TCACTTCAAATCCCGGATAAAGAAGTTTGGAAGGCTCGTGAT  630  OPT          589 TCGCTGCAGATTCCGGACAAAGAAGTGTGGAAGGCACGTGAT  630  cons         589 ** ** ** ** ***** ******** ******** ******  630   wildtype     631 GTTTTCCAATCTCTAGTTAGCGATTCGATACGTGATATACTG  672  OPT          631 GTTTTTCAAAGCCTGGTCTCAGATTCGATTCGTGACATCCTG  672  cons         631 ***** ***   ** **    ******** ***** ** ***  672   wildtype     673 GAAAAGGAAGAGACAAAGCAGCGTCCTGCTGATTTAATACCA  714  OPT          673 GAAAAAGAAGAAACCAAGCAGCGCCCGGCAGATCTGATTCCG  714  cons         673 ***** ***** ** ******** ** ** *** * ** **   714    153  wildtype     715 AGTGTTTTAACTATTAGAAATTTGTACAATTTTAATCCTGGT  756  OPT          715 TCTGTTCTGACGATCCGCAACCTGTACAACTTTAATCCGGGT  756  cons         715   **** * ** **  * **  ******* ******** ***  756   wildtype     757 AATTTTCATTATATACAAAAGACATTTGATTTGACCAAAAAA  798  OPT          757 AACTTCCACTACATCCAGAAGACCTTCGATCTGACGAAAAAG  798  cons         757 ** ** ** ** ** ** ***** ** *** **** *****   798   wildtype     799 GATGGGTTCCAATTTGATATCACTTACAATAAACTTGGCACT  840  OPT          799 GACGGTTTCCAATTCGATATCACCTACAACAAGCTGGGCACC  840  cons         799 ** ** ******** ******** ***** ** ** *****   840   wildtype     841 ACTCAAAGTATTTCCACCAGGGAACAAGTTACGGAGTTGATT  882  OPT          841 ACGCAGAGCATCTCTACGCGTGAACAAGTGACCGAACTGATT  882  cons         841 ** ** ** ** ** **  * ******** ** **  *****  882   wildtype     883 ACTTGGTCACTAAATGAGATAAACGCGCGTTTTGATAAGCAG  924  OPT          883 ACGTGGTCACTGAACGAAATCAATGCTCGCTTTGACAAACAG  924  cons         883 ** ******** ** ** ** ** ** ** ***** ** ***  924   wildtype     925 TTTAGTTTTGGAGAAGGTCCCGACTCAATTGAAAGCGTGGAG  966  OPT          925 TTTTCGTTCGGCGAAGGTCCGGATAGCATCGAAAGCGTGGAA  966  cons         925 ***   ** ** ******** **    ** ***********   966    154  wildtype     967 GTCAAAAGAAGATTTGCTTTAGAGACGCTATCAAACCTATTA 1008  OPT          967 GTGAAACGTCGCTTTGCCCTGGAAACCCTGTCAAACCTGCTG 1008  cons         967 ** *** *  * *****  * ** ** ** ********  *  1008   wildtype    1009 GGAGGAATCGGTTATTTCTATGGGAATCAACTAATTGATCGT 1050  OPT         1009 GGCGGTATTGGTTACTTCTATGGCAATCAGCTGATCGACCGT 1050  cons        1009 ** ** ** ***** ******** ***** ** ** ** *** 1050   wildtype    1051 GAAACAGAATTTGATGAGAGCCAGTTTACAGAGATCAAACTG 1092  OPT         1051 GAAACCGAATTTGATGAATCGCAATTCACGGAAATTAAACTG 1092  cons        1051 ***** ***********    ** ** ** ** ** ****** 1092   wildtype    1093 CTGAATGCAAAAGAGGAAGGTCCATTTGAACTGTTTACCAGC 1134  OPT         1093 CTGAATGCGAAGGAAGAAGGTCCGTTTGAACTGTTCACCAGT 1134  cons        1093 ******** ** ** ******** *********** *****  1134   wildtype    1135 GTTCCGAGCCGTGGCTTTTTCCCACGTGGATTCTATTGGGAT 1176  OPT         1135 GTTCCGAGCCGTGGCTTTTTCCCGCGCGGCTTTTACTGGGAT 1176  cons        1135 *********************** ** ** ** ** ****** 1176   wildtype    1177 GAAGGTTTCCATCTTCTACAAATTATGGAGTATGATTTTGAC 1218  OPT         1177 GAAGGCTTCCATCTGCTGCAGATTATGGAATATGATTTTGAC 1218  cons        1177 ***** ******** ** ** ******** ************ 1218    155  wildtype    1219 CTTGCCTTTGAAATCTTAGCGAGCTGGTTTGAAATGATCGAA 1260  OPT         1219 CTGGCCTTCGAAATCCTGGCAAGCTGGTTTGAAATGATTGAA 1260  cons        1219 ** ***** ****** * ** ***************** *** 1260   wildtype    1261 GATGATAGTGGTTGGATTGCTAGAGAAATTATACTGGGTAAT 1302  OPT         1261 GATGACTCTGGTTGGATCGCACGTGAAATTATCCTGGGCAAC 1302  cons        1261 *****   ********* **  * ******** ***** **  1302   wildtype    1303 GAGGCAAGGAGTAAAGTTCCGCAGGAATTTCAGGTGCAAAAT 1344  OPT         1303 GAAGCTCGCAGCAAAGTCCCGCAGGAATTTCAGGTGCAAAAC 1344  cons        1303 ** **  * ** ***** ***********************  1344   wildtype    1345 CCCAATATTGCTAATCCGCCAACTTTATTGCTAGCATTTAGT 1386  OPT         1345 CCGAATATTGCCAATCCGCCGACCCTGCTGCTGGCATTTTCA 1386  cons        1345 ** ******** ******** **  *  **** ******    1386   wildtype    1387 GAAATGCTTTCTAGGGCCATTGAAAACATCGGCGATTTCAAC 1428  OPT         1387 GAAATGCTGTCGCGTGCTATTGAAAACATCGGTGATTTCAAT 1428  cons        1387 ******** **  * ** ************** ********  1428   wildtype    1429 AGTGACAGCTACCACCAAGTCATGTTCAATAGTAGGACAGCC 1470  OPT         1429 AGTGACTCCTATCACCAGGTGATGTTTAACAGTCGCACCGCG 1470  cons        1429 ******  *** ***** ** ***** ** *** * ** **  1470    156  wildtype    1471 AAGTTTATGACGAACAATCTAGAAGCCAATCCTGGCTTGCTA 1512  OPT         1471 AAATTCATGACGAACAATCTGGAAGCCAATCCGGGCCTGCTG 1512  cons        1471 ** ** ************** *********** *** ****  1512   wildtype    1513 ACCGAATATGCCAAGAAAATTTATCCTAAGCTATTGAAGCAC 1554  OPT         1513 ACCGAATACGCAAAAAAGATTTATCCGAAACTGCTGAAGCAT 1554  cons        1513 ******** ** ** ** ******** ** **  *******  1554   wildtype    1555 TATAATTGGTTCAGAAAATCTCAAACAGGACTTATTGATGAA 1596  OPT         1555 TACAACTGGTTTCGTAAAAGCCAGACGGGTCTGATTGATGAA 1596  cons        1555 ** ** *****  * ***   ** ** ** ** ********* 1596   wildtype    1597 TATGAGGAAATATTGGAAGATGAAGGAATATGGGATAAGATT 1638  OPT         1597 TATGAAGAAATCCTGGAAGACGAAGGCATTTGGGATAAAATC 1638  cons        1597 ***** *****  ******* ***** ** ******** **  1638   wildtype    1639 CATAAGAACGAAGTTTATAGATGGGTTGGGCGTACCTTCACT 1680  OPT         1639 CATAAGAATGAAGTGTACCGTTGGGTTGGTCGCACCTTCACG 1680  cons        1639 ******** ***** **  * ******** ** ********  1680   wildtype    1681 CATTGTTTGCCAAGCGGTATGGATGACTATCCTAGAGCACAA 1722  OPT         1681 CACTGCCTGCCGTCTGGCATGGATGACTATCCGCGCGCTCAG 1722  cons        1681 ** **  ****    ** **************  * ** **  1722    157  wildtype    1723 CCACCAGATGTAGCAGAATTGAACGTAGACGCATTAGCATGG 1764  OPT         1723 CCGCCGGACGTGGCAGAACTGAACGTTGATGCGCTGGCCTGG 1764  cons        1723 ** ** ** ** ****** ******* ** **  * ** *** 1764   wildtype    1765 GTGGGCGTTATGACAAGATCCATGAAGCAAATTGCTCACGTG 1806  OPT         1765 GTGGGTGTTATGACCCGTAGCATGAAACAAATTGCTCATGTG 1806  cons        1765 ***** ********  *   ****** *********** *** 1806   wildtype    1807 TTGAAGTTAACACAGGACGAGCAAAGATATGCACAAATTGAG 1848  OPT         1807 CTGAAGCTGACGCAGGATGAACAACGCTATGCGCAGATCGAA 1848  cons        1807  ***** * ** ***** ** *** * ***** ** ** **  1848   wildtype    1849 CAAGAGGTGGTCGAGAATCTGGATTTGTTACACTGGAGTGAA 1890  OPT         1849 CAAGAAGTCGTGGAAAATCTGGACCTGCTGCACTGGAGTGAA 1890  cons        1849 ***** ** ** ** ********  ** * ************ 1890   wildtype    1891 AATGACAATTGCTACTGTGATATTAGCATCGATCCAGAAGAC 1932  OPT         1891 AACGATAATTGCTACTGTGATATTTCCATCGACCCGGAAGAT 1932  cons        1891 ** ** ******************  ****** ** *****  1932   wildtype    1933 GATGAGATTAGGGAGTTTGTATGTCATGAGGGTTACGTCTCC 1974  OPT         1933 GACGAAATTCGTGAATTTGTTTGTCATGAAGGTTATGTCAGC 1974  cons        1933 ** ** *** * ** ***** ******** ***** ***  * 1974    158  wildtype    1975 GTATTGCCCTTTGCATTGAAGCTAATCCCCAAAAACTCACCC 2016  OPT         1975 GTGCTGCCGTTCGCCCTGAAACTGATCCCGAAGAACTCTCCG 2016  cons        1975 **  **** ** **  **** ** ***** ** ***** **  2016   wildtype    2017 AAGCTAGAGAAAGTAGTTGCTTTGATGAGTGACCCAGAAAAA 2058  OPT         2017 AAACTGGAAAAGGTTGTCGCACTGATGTCAGACCCGGAAAAA 2058  cons        2017 ** ** ** ** ** ** **  *****   ***** ****** 2058   wildtype    2059 ATCTTTTCAGACTACGGGCTGTTATCACTATCGAGACAAGAC 2100  OPT         2059 ATTTTTAGCGATTACGGCCTGCTGTCACTGTCGCGTCAGGAT 2100  cons        2059 ** ***   ** ***** *** * ***** *** * ** **  2100   wildtype    2101 GACTATTTCGGCAAGGATGAAAACTATTGGAGAGGCCCAATT 2142  OPT         2101 GACTACTTCGGCAAGGATGAAAACTACTGGCGCGGCCCGATT 2142  cons        2101 ***** ******************** *** * ***** *** 2142   wildtype    2143 TGGATGAATATTAATTACTTGTGTCTTGACGCAATGAGATAC 2184  OPT         2143 TGGATGAACATCAATTATCTGTGCCTGGATGCGATGCGCTAT 2184  cons        2143 ******** ** *****  **** ** ** ** *** * **  2184   wildtype    2185 TACTACCCAGAGGTGATTCTCGACGTGGCTGGTGAGGCTAGC 2226  OPT         2185 TACTATCCGGAAGTCATCCTGGATGTGGCAGGCGAAGCTAGC 2226  cons        2185 ***** ** ** ** ** ** ** ***** ** ** ****** 2226    159  wildtype    2227 AATGCCAAGAAACTGTACCAAAGTTTAAAGATTAATCTCAGT 2268  OPT         2227 AACGCGAAAAAGCTGTACCAGAGCCTGAAGATCAACCTGTCT 2268  cons        2227 ** ** ** ** ******** **  * ***** ** **   * 2268   wildtype    2269 AACAACATATACAAAGTTTGGGAAGAACAAGGTTATTGTTAT 2310  OPT         2269 AACAACATCTACAAAGTGTGGGAAGAACAAGGTTACTGTTAC 2310  cons        2269 ******** ******** ***************** *****  2310   wildtype    2311 GAAAATTACAGTCCGATAGATGGTCATGGTACTGGTGCTGAG 2352  OPT         2311 GAAAACTACAGCCCGATTGATGGCCATGGTACCGGCGCGGAA 2352  cons        2311 ***** ***** ***** ***** ******** ** ** **  2352   wildtype    2353 CATTTCACAGGCTGGACAGCACTTGTTGTCAACATCCTTGGA 2394  OPT         2353 CACTTTACCGGCTGGACGGCCCTGGTGGTTAATATCCTGGGT 2394  cons        2353 ** ** ** ******** ** ** ** ** ** ***** **  2394   wildtype    2395 CGC----------------------------T---------- 2398  OPT         2395 CGTTTTCTGGAAGTCCTGTTTCAAGGTCCGCATCATCACCAC 2436  cons        2395 **                                         2436   wildtype    2399 ----TCTGA 2403  OPT         2437 CATCACTAA 2445  cons        2437      ** * 2445   

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