Science, Faculty of
Chemistry, Department of
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UBCV
Golisch, Benedikt
Master of Science - MSc
University of British Columbia
The human gut microbiota (HGM) is a complex microbial community that possesses a huge capacity for carbohydrate degradation in contrast to humans and is dominated by the phylum Bacteroidetes comprising Bacteroides and Prevotella. While Bacteroides species are associated with people consuming protein/fat-rich diets, members of Prevotella including Prevotella copri are highly abundant in people consuming plant-based diets. Here, I present a comprehensive genetic, biochemical, and structural-biological investigation of a recently identified gene cluster of P. copri upregulated in response to mixed-linkage β-glucan (MLG), a polysaccharide commonly found in cereals. The MLG utilization locus (MLGUL) contains an SGBP-A that interacted with MLG specifically and an SGBP-B that recognized 1,4-linked β-glucans promiscuously. The tertiary structure of SGBP-A revealed an aromatic binding platform and flexible loop that interact with cellopentaose or potential MLG-breakdown products. Similarly, a structural model suggested a binding platform for the SGBP-B. Furthermore, the MLGUL encodes a GH5_4, a highly specific, endo-acting MLGase, that produced oligosaccharides by preferential hydrolysis of β-1,3-linkages. The product analysis and tertiary structure revealed that β-1,4-linked glucose units in the negative subsites and occupation of the +2 subsite facilitate this cleavage. In addition, the MLGUL includes an exo-acting β-glucosidase from the GH3 family that degraded longer oligosaccharides into glucose and cellobiose. The latter will be further cleaved to glucose and glucose 1-phosphate by a predicted cellobiose phosphorylase, a member of GH94. Together, these enzymological and biochemical studies support a model for MLG degradation by P. copri in which SGBPs bind substrate at the cell surface to facilitate cleavage by the endo-MLGase GH5_4 and import by the TonB-dependent transporter. The generated oligosaccharides will be sequentially cleaved by a GH3 β-glucosidase and a GH94 cellobiose phosphorylase, in the periplasm and cytosol, respectively, generating glucose and glucose 1-phosphate as final products for the metabolism of P. copri. Thus, the work provides insights into the molecular mechanism of MLG utilization by P. copri and might indicate factors affecting the fitness of P. copri within the HGM. This knowledge might advance our understanding of diet-host-microbe interactions and may result in innovative therapeutic approaches for prevalent diseases linked to the human gut.
Functional characterization of a mixed-linkage beta-glucan utilization locus from the human gut microbe Prevotella copri by Benedikt Golisch B.S., Humboldt Universität zu Berlin, 2020 A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE in The Faculty of Graduate and Postdoctoral Studies (Chemistry) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) August 2023 © Benedikt Golisch, 2023 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the thesis entitled: Functional characterization of a mixed-linkage beta-glucan utilization locus from the human gut microbe Prevotella copri submitted by Benedikt Golisch in partial fulfillment of the requirements for the degree of Master of Science in Chemistry Examining Committee: Dr. Harry Brumer, Professor, Dept. of Chemistry, Dept. of Biochemistry and Molecular Biology, Dept. of Botany, UBC Supervisor Dr. Reinhard Jetter, Professor, Dept. of Chemistry, Dept. of Botany, UBC Supervisory Committee Member Dr. David Perrin, Professor, Dept. of Chemistry, UBC Supervisory Committee Member Dr. Chris Orvig, Professor, Dept. of Chemistry, UBC Additional Examiner iii Abstract The human gut microbiota (HGM) is a complex microbial community that possesses a huge capacity for carbohydrate degradation in contrast to humans and is dominated by the phylum Bacteroidetes comprising Bacteroides and Prevotella. While Bacteroides species are associated with people consuming protein/fat-rich diets, members of Prevotella including Prevotella copri are highly abundant in people consuming plant-based diets. Here, I present a comprehensive ge-netic, biochemical, and structural-biological investigation of a recently identified gene cluster of P. copri upregulated in response to mixed-linkage β-glucan (MLG), a polysaccharide commonly found in cereals. The MLG utilization locus (MLGUL) contains an SGBP-A that interacted with MLG spe-cifically and an SGBP-B that recognized 1,4-linked β-glucans promiscuously. The tertiary struc-ture of SGBP-A revealed an aromatic binding platform and flexible loop that interact with cello-pentaose or potential MLG-breakdown products. Similarly, a structural model suggested a binding platform for the SGBP-B. Furthermore, the MLGUL encodes a GH5_4, a highly specific, endo-acting MLGase, that produced oligosaccharides by preferential hydrolysis of β-1,3-linkages. The product analysis and tertiary structure revealed that β-1,4-linked glucose units in the negative sub-sites and occupation of the +2 subsite facilitate this cleavage. In addition, the MLGUL includes an exo-acting β-glucosidase from the GH3 family that degraded longer oligosaccharides into glucose and cellobiose. The latter will be further cleaved to glucose and glucose 1-phosphate by a predicted cellobiose phosphorylase, a member of GH94. Together, these enzymological and biochemical studies support a model for MLG degradation by P. copri in which SGBPs bind substrate at the cell surface to facilitate cleavage by the endo-MLGase GH5_4 and import by the TonB-dependent transporter. The generated oligosaccharides will be sequentially cleaved by a GH3 β-glucosidase iv and a GH94 cellobiose phosphorylase, in the periplasm and cytosol, respectively, generating glu-cose and glucose 1-phosphate as final products for the metabolism of P. copri. Thus, the work provides insights into the molecular mechanism of MLG utilization by P. copri and might indicate factors affecting the fitness of P. copri within the HGM. This knowledge might advance our understanding of diet-host-microbe interactions and may result in innovative thera-peutic approaches for prevalent diseases linked to the human gut. v Lay Summary Cereals including oats and barley contain mixed-linage β-glucans (MLGs) which have health-promoting effects on blood glucose, insulin, and cholesterol levels. While humans cannot degrade certain carbohydrates, bacteria from the human gut employ enzyme machineries for the metabolism of various carbohydrates. Therefore, species such as Prevotella copri that thrive in people consuming plant-carbohydrate-rich diets are of interest to find novel treatments for com-mon diseases such as diabetes, obesity, or inflammatory bowel disease. Today, our knowledge about the specific mechanism of MLG utilization in P. copri is scarce since detailed biochemical analyses are required to uncover the role of the involved proteins. This work investigates the com-plementary interaction of enzymes degrading MLGs and distinct carbohydrate-binding proteins from P. copri. Furthermore, it analyses key differences to species associated with diets enriched in protein and fat to explore the potential of P. copri for treatments of common diseases. vi Preface Chapter 1: Introduction Chapter 1 was written by me, Benedikt Golisch. My supervisor, Prof. Harry Brumer, re-viewed and helped revise it. A former postdoctoral researcher in our lab, Dr. Jonathon Briggs, analyzed the transcriptome of P. copri in response to various carbohydrates. The subchapter “Pol-ysaccharide utilization loci encode protein machineries for efficient carbohydrate uptake” re-phrases a paragraph from our review article discussing the general mechanism of PULs: Golisch, B., Lei, Z. H., Tamura, K. and Brumer, H. (2021) Configured for the human gut microbiota: Molecular mechanisms of dietary β-glucan utilization. ACS Chemical Biol-ogy. 16, 2087-2102 Chapter 2: Extracellular MLG hydrolysis by the specific PcGH5_4 Chapter 2 was written by me, Benedikt Golisch. My supervisor, Prof. Harry Brumer, and a postdoctoral researcher in our lab, Dr. Rosa Lorizolla Cordeiro, reviewed and helped revise it. I cloned, expressed, and purified the discussed protein constructs of PcGH5_4. Furthermore, I con-ducted the biochemical analysis investigating the substrate range and kinetic parameters of poly-saccharide digestion using the BCA assay. Product analysis and oligosaccharide degradation mon-itored by HPAEC-PAD and MS were performed by me. Initially, conditions for crystallization of the protein were screened by Dr. Cordeiro and me using the facilities and the equipment of our collaborators, the group of Prof. Filip Van Petegem (Life Science Institute, UBC). Then, I opti-mized the conditions that displayed crystal growth and diffracted the crystals under the guidance of Dr. Cordeiro. The protein crystal structure was solved by me roughly and an in-depth refinement of the structure was done by Dr. Cordeiro. vii Chapter 3: Downstream degradation of oligosaccharides mediated by PcGH3 and PcGH94 Chapter 3 was written by me, Benedikt Golisch. My supervisor, Prof. Harry Brumer, and Dr. Rosa Lorizolla Cordeiro reviewed and helped revise it. Dr. Cordeiro helped clone PcGH3V23-R772 while all other protein constructs were cloned by me. After I tested the expression, Dr. Cor-deiro and I purified the different constructs. Biochemical analysis of the PcGH3 was conducted by me. MS-analyses of long oligosaccharides were performed by the mass spectrometry service in the Chemistry Department. Chapter 4: Carbohydrate recognition by extracellular SGBPs Chapter 4 was written by me, Benedikt Golisch. My supervisor, Prof. Harry Brumer and Dr. Rosa Lorizolla Cordeiro reviewed and helped revise it. The chapter includes experiments per-formed, designed, and analyzed by me supervised by Prof. Harry Brumer. A postdoctoral re-searcher in our group, Dr. Jonathon Briggs, cloned the construct PcSGBP-AD33-K595(+IGSG). Another postdoctoral researcher, Dr. Cordeiro, optimized the crystallization conditions of PcSGBP-A and helped me in collecting diffraction data. The collected data was then processed and refined by Dr. Cordeiro. Chapter 5: Conclusion and future perspectives Chapter 5 was written by me, Benedikt Golisch. My supervisor, Prof. Harry Brumer, re-viewed and helped revise it. viii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary .................................................................................................................................v Preface ........................................................................................................................................... vi Table of Contents ....................................................................................................................... viii List of Tables ............................................................................................................................... xii List of Figures ............................................................................................................................. xiii List of Schemes ........................................................................................................................... xvi List of Abbreviations ................................................................................................................ xvii Acknowledgments ...................................................................................................................... xxi Dedication ................................................................................................................................. xxiii Chapter 1: Introduction ................................................................................................................1 1.1 The human gut and the human gut microbiota ............................................................... 1 1.2 Prevotella copri – keystone species of the HGM ........................................................... 3 1.3 Mixed-linkage β-glucan – unique polysaccharide in the human diet ............................. 5 1.4 Polysaccharide utilization loci encode protein machineries for efficient carbohydrate uptake …………………………………………………………………………………………..9 1.5 Mixed-linkage β-glucan utilization locus (MLGUL) in P. copri ................................. 13 1.6 Aims of the Thesis ........................................................................................................ 15 Chapter 2: Extracellular MLG hydrolysis by the specific PcGH5_4......................................16 2.1 Introduction ................................................................................................................... 16 2.2 Results and discussion .................................................................................................. 16 ix 2.2.1 PcGH5_4 is an endo-acting 1,3/1,4-β-glucanase ...................................................... 16 2.2.2 PcGH5_4 preferentially hydrolyzes β-1,3-linkages in MLG .................................... 21 2.2.3 Specific subsite arrangement for the cleavage of MLG-oligosaccharides ................ 24 2.2.4 Structural features of the MLG-specific PcGH5_4 .................................................. 30 2.2.5 Conserved negative subsites stabilize β-1,4-linked carbohydrates in the active site 34 2.2.6 Unique positive subsites of PcGH5_4 with potential for substrate discrimination .. 38 2.2.7 Catalytic comparison with other MLGases ............................................................... 41 2.3 Conclusion .................................................................................................................... 43 Chapter 3: Downstream degradation of oligosaccharides mediated by PcGH3 and PcGH94........................................................................................................................................................44 3.1 Introduction ................................................................................................................... 44 3.2 Results and discussion .................................................................................................. 45 3.2.1 PcGH3 and periplasmic oligosaccharide breakdown ............................................... 45 3.2.2 A complementary path for cellobiose degradation via PcGH94 .............................. 54 3.3 Conclusion .................................................................................................................... 60 Chapter 4: Carbohydrate recognition by extracellular SGBPs ..............................................61 4.1 Introduction ................................................................................................................... 61 4.2 Results and discussion .................................................................................................. 62 4.2.1 PcSGBP-A – an MLG-specific SGBP-A .................................................................. 62 4.2.1.1 Specific binding of MLG by PcSGBP-A .......................................................... 63 4.2.1.2 Structural features of PcSGBP-A without ligand and in complex with cellopentaose ..................................................................................................................... 68 4.2.1.3 Carbohydrate binding interactions of PcSGBP-A ............................................ 71 x 4.2.1.4 Structural conservation of PcSGBP-A .............................................................. 75 4.2.1.5 Role of SGBP-A in MLG utilization ................................................................ 76 4.2.2 PcSGBP-B – an auxiliary binding protein ................................................................ 79 4.2.2.1 Promiscuous binding of 1,4-β-glucans ............................................................. 79 4.2.2.2 Potential function and role of PcSGBP-B ......................................................... 81 4.3 Conclusion .................................................................................................................... 84 Chapter 5: Conclusion and future perspectives ........................................................................85 5.1 Summary of the results ................................................................................................. 85 5.2 Future experiments........................................................................................................ 86 5.3 Outlook ......................................................................................................................... 88 References .....................................................................................................................................90 Appendices ..................................................................................................................................100 Appendix A Materials and methods ....................................................................................... 100 A.1 Cloning, expression, and purification of recombinant proteins .............................. 100 A.1.1. Cloning and sequencing .................................................................................. 100 A.1.2. Expression and purification ............................................................................ 102 A.2 Biochemical characterization .................................................................................. 103 A.2.1. Substrates .................................................................................................... 103 A.2.2. Bicinchoninic acid assay ............................................................................. 104 A.2.3. para-Nitrophenol (pNP) substrate assay ..................................................... 106 A.2.4. Carbohydrate analysis by HPAEC-PAD .................................................... 106 A.2.5. Native oligosaccharide hydrolysis .............................................................. 109 A.2.6. Thermal shift assay ..................................................................................... 109 xi A.2.7. Affinity gel electrophoresis......................................................................... 110 A.2.8. Isothermal titration calorimetry .................................................................. 111 A.3 Oligosaccharide production and purification .......................................................... 111 A.4 Crystallization, X-ray diffraction data collection and data processing ................... 112 xii List of Tables Table 2.1 Kinetic data for the digestion of various polysaccharides by the endo-active PcGH5_4C21-E406. ........................................................................................................................... 21 Table 2.2 Kinetic data for the digestion of shorter oligosaccharides by the endo-active PcGH5_4C21-E406 ............................................................................................................................ 28 Table 2.3 Structural homologs of PcGH5_4T41-E406 ...................................................................... 32 Table 2.4 Activity of catalytic mutants of PcGH5_4 on barley MLG. ......................................... 33 Table 3.1 Sequence similarity of Prevcop_05090 towards crystallized members of GH94. ....... 54 Table 4.1 Thermodynamic parameters of the protein-carbohydrate interaction of the SGBPs .... 67 Table 4.2 Closest structural homologs of PcSGBP-AD33-K595(+IGSG) ............................................. 76 Table A.1 List of the used protein constructs, primers, and vectors. .......................................... 101 Table A.2 Crystallographic data and refinement statistics table ................................................. 113 xiii List of Figures Figure 1.1 Comparison of different carbohydrate representations for a heptaoligosaccharide. ..... 5 Figure 1.2 Overview of representative complex carbohydrate structures and their respective sources............................................................................................................................................. 6 Figure 1.3 Genetic arrangement of PULs from certain Bacteroides species. ............................... 11 Figure 1.4 Model of the stepwise hydrolysis of a polysaccharide mediated by proteins of a generic PUL. ................................................................................................................................. 12 Figure 1.5 Genetic organization of MLGULs............................................................................... 14 Figure 2.1 Structure of the cloned and expressed constructs of PcGH5_4. .................................. 17 Figure 2.2 SDS-PAGE gels of the purified PcGH5_4 constructs ................................................. 18 Figure 2.3 Initial screen for the hydrolytic activity of GH5_4 on various polysaccharides. ........ 19 Figure 2.4 Specific activity of PcGH5_4C21-E406 on barley MLG in different buffers and different temperatures. ................................................................................................................................. 20 Figure 2.5 Michaelis-Menten kinetics for the hydrolysis of different polysaccharides. .............. 20 Figure 2.6 Product analysis of the digestion of barley MLG by PcGH5_4C21-E406 via HPAEC-PAD............................................................................................................................................... 22 Figure 2.7 Amounts of generated products during the digestion of barley MLG by PcGH5_4C21-E406. ................................................................................................................................................ 23 Figure 2.8 HPAEC-PAD analysis of the digestion of oligosaccharides by PcGH5_4C21-E406. ..... 25 Figure 2.9 Initial rate kinetics for the hydrolysis of various oligosaccharides by PcGH5_4C21-E406........................................................................................................................................................ 27 Figure 2.10 Alignment of selected oligosaccharides to the subsites of PcGH5_4. ...................... 27 xiv Figure 2.11 Mass spectra of products generated from the digestion of cellopentaose by PcGH5_4C21-E406. ........................................................................................................................... 30 Figure 2.12 Crystal structure of PcGH5_4T41-E406. ........................................................................ 31 Figure 2.13 Selected crystallization conditions of the catalytic mutants, PcGH5_4T41-E406(E197A) and PcGH5_4T41-E406(E331A). ........................................................................................................... 34 Figure 2.14 Structural comparison of PcGH5_4T41-E406 with two selected closest structural homologs. ...................................................................................................................................... 35 Figure 2.15 Overlay of the crystal structure of PcGH5_4T41-E406 with carbohydrate ligands in the negative subsites. .......................................................................................................................... 36 Figure 2.16 Superposition of potential carbohydrate ligands and the crystal structure of PcGH5_4T41-E406. ........................................................................................................................... 41 Figure 2.17 Hydrolysis of MLG by the endo-acting PcGH5_4 in comparison to BoGH16. ....... 42 Figure 3.1 Structure of the cloned and expressed constructs of PcGH3. ...................................... 45 Figure 3.2 Purification and stability analysis of PcGH3............................................................... 46 Figure 3.3 Specific activity of PcGH3V23-R772 on pNP-glycosides. .............................................. 47 Figure 3.4 Digestion of oligosaccharides by PcGH3V23-R772. ....................................................... 49 Figure 3.5 Mass spectra and HPLC chromatograms of purified MLG-oligosaccharides generated by PcGH5_4C21-E406. ...................................................................................................................... 50 Figure 3.6 AlphaFold model of the tertiary structure of the PcGH3V23-R772 ................................. 51 Figure 3.7 Predicted conformation of the active site of PcGH3V23-R772. ....................................... 52 Figure 3.8 Hydrolysis of MLG-derived oligosaccharides by the exo-acting PcGH3 in comparison to BoGH3MLG. ............................................................................................................................... 53 Figure 3.9 Structure of the cloned and expressed constructs of PcGH94. .................................... 55 xv Figure 3.10 SDS-PAGE gels of the expression tests for the constructs of PcGH94. ................... 56 Figure 3.11 Structural model of PcGH94 obtained by AlpaFold ................................................. 57 Figure 3.12 Predicted three-dimensional structure of the active site of PcGH94. ........................ 59 Figure 4.1 Structure of the cloned and expressed constructs of PcSGBP-A. ............................... 62 Figure 4.2 SDS-PAGE gels of the purified PcSGBP-A constructs .............................................. 63 Figure 4.3 Binding affinities of MLGUL encoded SGBPs to polysaccharides. ........................... 65 Figure 4.4 Polysaccharide binding by PcSGBP-AP50-K595 and PcSGBP-BF42-E332 monitored by ITC. ............................................................................................................................................... 66 Figure 4.5 Three-dimensional crystal structure of PcSGBP-AD33-K595(+IGSG). ............................... 69 Figure 4.6 Crystal structure of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose. ........... 70 Figure 4.7 Contact sites of two asymmetric units in the crystal structure of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose. .................................................................................... 71 Figure 4.8 Carbohydrate binding site of PcSGBP-AD33-K595(+IGSG). .............................................. 74 Figure 4.9 Structural model for the SusC-SusD interaction. ........................................................ 78 Figure 4.10 Structure of the cloned and expressed constructs of PcSGBP-B. ............................. 80 Figure 4.11 SDS-PAGE gels of the purified PcSGBP-B constructs ............................................ 80 Figure 4.12 Selected crystallization conditions for the constructs, PcSGBP-BF42-E332 and PcSGBP-BP136-E332. ....................................................................................................................... 81 Figure 4.13 Model of the structure of PcSGBP-B. ....................................................................... 82 Figure 5.1 Cellular model for the MLG uptake encoded by the MLGUL of P. copri. ................ 86 xvi List of Schemes Scheme 2.1 Koshland double-displacement mechanism for the retaining hydrolysis of a β-1,4-glucan. ........................................................................................................................................... 33 xvii List of Abbreviations APS: Ammonium persulfate B. fluxus: Bacteroides fluxus B. ovatus: Bacteroides ovatus B. theta: Bacteroides thetaiotaomicron B. uniformis: Bacteroides uniformis BCA: Bicinchoninic acid BoGH16: Member of GH16 from an MLGUL of B. ovatus BoGH3MLG: Member of GH3 from an MLGUL of B. ovatus BoGH3XyG: Member of GH3 from a xyloglucan utilization locus of B. ovatus BoSGBP-A: SGBP-A from an MLGUL of B. ovatus BoSGBP-B: SGBP-B from an MLGUL of B. ovatus BSA: Bovine serum albumin BtSGBP-A: SGBP-A from a predicted PUL of B. theta CAZyme: Carbohydrate active enzyme CIHR: Canadian Institutes of Health Research ddH2O: Sterile ultra-pure water ESI: Electrospray ionization F32EG5: Member of GH5_4 from Caldicellulosiruptor sp. F32 G3G: Laminaribiose G3G3G: Laminaritriose G3G3G3G: Laminaritetraose xviii G3G4G: 32-β-D-glucosyl-cellobiose G3G4G4G: 33-β-D-glucosyl-cellotriose G4G: Cellobiose G4G3G: 31-β-D-cellobiosyl-glucose G4G3G4G: 32-β-D-cellobiosyl-cellobiose G4G4G: Cellotriose G4G4G3G: 31-β-D-cellotriosyl-glucose G4G4G4G: Cellotetraose G4G4G4G4G: Cellopentaose G6G: Gentibiose GH##: Glycoside hydrolase family ## GH: Glycoside hydrolase GH5_4: Glycoside hydrolase family 5, subfamily 4 Glc: Glucose Gly-OH: Glycine hydroxide HEPES: 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid HGM: Human gut microbiota HPAEC-PAD: High-performance anionic-exchange chromatography with pulsed amperometric detection HPLC: High-performance liquid chromatography HTCS: Hybrid two-component system ID: Sequence identity IMAC: Immobilized metal chelate affinity chromatography xix IPTG: Isopropyl-D-thiogalactoside ITC: Isothermal titration calorimetry kDa: Kilodalton MES: 2-(N-Morpholino)ethanesulfonic acid MLG: Mixed-linkage β-glucan MLGUL: Mixed-linkage β-glucan utilization locus MW: Molecular weight NaCit: Sodium citrate NaOAc: Sodium acetate NaPi: Sodium phosphate OD600: Optical density at 600 nm P. copri: Prevotella copri PbGH5_4: Member of GH5_4 from Prevotella bryantii PcGH3: Member of GH3 from an MLGUL of P. copri PcGH5_4: Member of GH5_4 from an MLGUL of P. copri PcGH94: Predicted member of GH94 from an MLGUL of P. copri PcSGBP-A: SGBP-A from an MLGUL of P. copri PcSGBP-B: SGBP-B from an MLGUL of P. copri PDB-ID: Alphanumeric protein structure identifier within the Protein Data Bank PFAM: Protein Families Database pNP: para-Nitrophenol PpGH5_4: Member of GH5_4 from Paenibacillus pabuli PUL: Polysaccharide utilization locus xx r.m.s.d.: Root-mean-square deviation SDS-PAGE: Sodium dodecyl sulfate - polyacrylamide gel electrophoresis SEC: Size-exclusion chromatography SGBP: Surface glycan-binding protein SusC: TonB-dependent transporter of the starch utilization system SusD: SGBP-A of the starch utilization system SusR: Transcriptional regulator of the starch utilization system TBDT: TonB-dependent transporter TEMED: Tetramethyl ethylenediamine TEV: Tobacco etch virus Tris: Tris(hydroxymethyl)aminomethane UV-Vis: Ultraviolet-Visible XXXG: Xyloglucan heptamer unit, Xyl3Glc4, where "X" represents a xylose unit attached to glu-cose, and "G" an unsubstituted glucose residue Xyl: Xylose ZgGH5_4: Member of GH5_4 from Zobellia galactanivorans xxi Acknowledgments First and foremost, I would like to thank my supervisor, Prof. Harry Brumer, who offered me this great opportunity for my academic, professional, and personal growth. Over the last three years, he guided me through the ups and downs of my work in a very motivating, inspiring, un-derstanding, and supportive way. The advice and experiences I obtained will help me become a scientist who thinks critically, investigates in-depth, works together with fellow researchers, and can smile about the joys of life and science. I would also like to acknowledge my committee mem-bers, Prof. Reinhard Jetter and Prof. David Perrin, and the chair of my examination, Prof. Chris Orvig, who took the time to critically read this thesis. Great thanks go out to my fellow lab members for their open ears to my questions about genetic, biochemical, and structural biological analyses. Their help and inspiring suggestions pushed my project forward. Thus, I want to thank especially former and current members of our PUL team. Dr. Julie Grondin patiently introduced me to the main techniques in biochemistry, the expertise of Dr. Rosa Lorizolla Cordeiro helped in crystallizing the proteins, in solving their three-dimensional structure and in writing this thesis, and the exchange of ideas and methods with Zhenhuan Lei and Dr. Deepesh Panwar widened my horizons. Apart from that, I would like to emphasize that my time in Vancouver would have been much more monotonous without our din-ner parties, hikes, table tennis sessions, and discussions about love and life. I want to appreciate our collaboration with the lab of Prof. Filip van Petegem at the Life Science Institute (UBC). The possibility to use their equipment and to receive professional feed-back created the basis for the structural analysis. xxii I would also like to highlight that part of the research described in this thesis was performed using beamline CMCF-ID at the Canadian Light Source, a national research facility of the Univer-sity of Saskatchewan, which is supported by the Canada Foundation for Innovation (CFI), the Natural Sciences and Engineering Research Council (NSERC), the National Research Council (NRC), the Canadian Institutes of Health Research (CIHR), the Government of Saskatchewan, and the University of Saskatchewan. Furthermore, some data included in this thesis was obtained by use of the Stanford Synchrotron Radiation Lightsource, SLAC National Accelerator Laboratory, which is supported by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Contract No. DE-AC02-76SF00515. The SSRL Structural Molecular Biology Pro-gram is supported by the DOE Office of Biological and Environmental Research, and by the Na-tional Institutes of Health, National Institute of General Medical Sciences (P30GM133894). The contents of this thesis are solely the responsibility of the authors and do not necessarily represent the official views of NIGMS or NIH. At this point, I want to appreciate The University of British Columbia for supporting my first two years with a Four-Year Fellowship allowing me to grow independently of financial concerns. Fur-thermore, I want to thank CIHR for generously funding this project. In my opinion, the investment in the education of young scientists contributes significantly to the progress of our society. Last, I am very grateful for the love and unending support from Annika, my family, and close friends. Without their support, this adventure never would have started. xxiii Dedication This thesis is dedicated to all who do not have the chance of an academic career. 1 Chapter 1: Introduction 1.1 The human gut and the human gut microbiota The human gut is home to trillions of microorganisms including bacteria, fungi, and viruses collectively forming a complex ecosystem [1] making humans holobionts [2]. Predominantly found in the large intestine due to slower flow rates, milder pH conditions, and an anaerobic envi-ronment [3], the microbial community often referred to as human gut microbiota (HGM) com-prises at least a similar number of microbial cells as there are human host cells [1]. Thus, the complexity of the HGM can be described in terms of species diversity and the influence of host-mediated factors. Within the HGM, Bacteroidetes (e.g., the genera Bacteroides, and Prevotella) and Firmicutes (e.g., Clostridium, Roseburia, and Lactobacillus) dominate other phyla including Actinobacteria (e.g., Bifidobacterium), Proteobacteria (e.g., Escherichia), Fusobacteria (e.g., Fusobacterium), and Verrucomicrobia (Akkermansia) which contribute to the remaining 10% of bacteria [3]. However, this composition is influenced by various host-related factors including diet, host genetics, medical treatments (e.g., antibiotics), as well as age, sex, and stress [4]. Among these factors, host health, genetics, and diet exert the most substantial influence on the HGM composi-tion [5]. For instance, the diversity of species increases in children after switching from breast milk to a more diverse diet [3, 6] and alterations among dominant species of the HGM induced by seasonal changes occur in traditional hunter-gatherer communities [7]. Due to the complexity, a specific definition of a healthy HGM composition remains elusive [4]. However, numerous studies have recognized the importance of high taxa diversity, high microbial gene richness, and stable microbiome functional cores as beneficial for human health [8, 9]. In other words, a highly diver-sified gut microbiome prevents an imbalance within the ecosystem and maintains the overall gut homeostasis. 2 The influence of the HGM on this homeostasis is a result of intricate interactions involving diet, microbial species, and human host cells. Unfortunately, the human genome only possesses a small arsenal of enzymes capable of degrading cooked starch, which comprises amylose and am-ylopectin [10]. Conversely, numerous enzymes present in the microbial genome enable the HGM to thrive on indigestible complex polysaccharides [10, 11]. As a result, components of the diet, including complex carbohydrates, are partially or completely fermented by the HGM in the lower gastrointestinal tract. The main products of this digestion include short-chain fatty acids, im-munostimulants, neurohormones, and vitamins, which are released partially into the gut lumen [4, 5, 12]. There, short-chain fatty acids including acetate, butyrate, propionate, or succinate decrease the luminal pH and serve as a main energy source for the colonocytes contributing to up to 10% of the caloric intake [4, 13, 14]. Furthermore, acetate, butyrate, propionate stimulate the secretion of small molecules such as peptide YY that modulate postprandial glucose levels. This upregula-tion improves glucose metabolism, insulin secretion, food intake, and energy expenditure [4, 15]. Moreover, the combination of various nutrients in the human gut enhances the formation of a thick intestinal mucus layer serving as a gut barrier. The natural barrier formed by the HGM prevents the colonization of the gut by pathogenic bacteria or fungi and decreases the risk of the invasion of pathogenic viruses [4, 16, 17]. Thus, a healthy HGM helps to maintain and regulate the overall gut climate. Despite the growing knowledge of the composition and function of the HGM in healthy people, many interactions within the HGM remain unclear. However, the concept of a healthy HGM ultimately implies that modulation of the HGM affects the host’s health. Therefore, research efforts have intensified to explore potential probiotic and prebiotic treatment strategies to address 3 prevalent diseases such as diabetes, obesity, inflammatory bowel disease, and colorectal cancer [18, 19]. 1.2 Prevotella copri – keystone species of the HGM Dietary factors have a significant impact on the composition of the HGM and hence dif-ferences between populations following Western diets and so-called non-Western diets have been observed [20]. Western diets are usually characterized by a high protein and fat content, whereas non-Western diets are rich in plant carbohydrates. However, other terminologies exist to further differentiate these diets for instance according to certain lifestyles such as in traditional hunter-gatherer or preindustrialized agrarian communities, or with regard to the ingredients, such as a plant-fiber-based vegetarian lifestyle [21, 22]. In 2011, Arumugam and coworkers mapped general differences in the HGM and identified distinct clusters of bacteria termed enterotypes [23]. These enterotypes, namely enterotype I dominated by Bacteroides species, enterotype II dominated by Prevotella species, and enterotype III dominated by Ruminococcus species, exhibit relative stabil-ity even during short-term dietary interventions [23]. Each enterotype represents a combination of complementary species that enable distinctive routes for energy production within the human gut [24]. As a result, enterotypes tend to align with specific diets. Plant-carbohydrate-rich diets coin-cide with an increased abundance of Prevotella whereas Bacteroides species are associated with diets consisting of animal proteins and unsaturated fats [21]. In fact, vegetarians contain an in-creased level of Prevotella species whereas Bacteroides strains are highly prevalent in omnivores [22]. Furthermore, the replacement of Prevotella strains with Bacteroides strains in people that migrated from Thailand to the US emphasizes the influence of distinct diets on the composition of the HGM [25]. 4 So far, research focus has traditionally been distributed towards several Bacteroides spe-cies including Bacteroides thetaiotaomicron (B. theta), Bacteroides ovatus (B. ovatus), and Bac-teroides uniformis (B. uniformis) [26-28]. But in recent years, more and more studies have recog-nized the significance of Prevotella for the understanding of the microbial ecosystem in the human gut [21, 22, 29, 30]. Meanwhile, the variation in microbial composition between people consuming Western diets and people following non-Western diets might point towards specific switches in the HGM whose modulation might cause, aggravate, and prevent common diseases associated with the human gut. In the case of the genus Prevotella, several studies investigated Prevotella copri as a model organism [29]. P. copri is the most abundant Prevotella species and comprises four different clades [9]. First described in 2007, the rod-shaped Gram-negative cells of P. copri require anaerobic con-ditions for growth. The authors observed acid production upon growth on several carbohydrates including glucose and cellobiose and identified succinic and acetic acid as the main products of digestion [31]. Despite these observations, the beneficial role of P. copri in the human gut is highly debated [21, 29]. While P. copri improves glucose metabolism and insulin sensitivity in mouse models by generating succinate as a substrate for intestinal glucogenesis, other studies suggested a correlation of P. copri with inflammatory conditions, glucose intolerance, and insulin resistance [21]. Indeed, different P. copri strains display opposite effects and their prevalence depends on specific diets [32]. Thus, correlation studies can provide initial insights but a comprehensive un-derstanding of the molecular processes and interactions related to the organism is required to com-pletely entangle the specific role of P. copri in the context of human health [29]. Despite that, increased abundance and broad diversity of P. copri strains in non-Western populations indicate a strong connection between P. copri and fiber-rich diets [21, 33]. In fact, the 5 enzyme repertoire encoded by the genome of P. copri reflects the enormous potential for carbo-hydrate utilization, enables P. copri to grow on various polysaccharides, and serves as a basis for further investigations [21, 30]. 1.3 Mixed-linkage β-glucan – unique polysaccharide in the human diet A diverse diet includes a significant amount of complex carbohydrates. Polysaccharides, in particular, possess a complex structure due to the diverse monosaccharide building blocks, two configurations of the anomeric linkage, alpha and beta, different glycosidic bond regiochemistry, possibility for branching, and various non-carbohydrate substituents. To express these structural features, various chemical notations are commonly used (Figure 1.1) [34]. Figure 1.1 Comparison of different carbohydrate representations for a heptaoligosaccharide. 6 Figure 1.2 Overview of representative complex carbohydrate structures and their respective sources. Different monosaccharides and substituents are provided in the left bottom corner. 7 The structure of the polysaccharide also depends on the individual species and varies among terrestrial plants, algae, fungi, and bacteria (Figure 1.2). The organisms mainly employ carbohydrates as structural components within the cell wall or for energy storage. Since oxidation of glucose by molecular oxygen releases – 2823 kJ/mol energy, starch consisting of polysaccha-rides containing α-1,4- and α-1,6-linked glucose units, plays a fundamental role in human nutrition [35]. However, multiple other polysaccharides present in the human diet remain undigested by human-encoded enzymes and are therefore called dietary fiber. Regarding their potential health benefits, national health agencies worldwide advise a daily consumption of 20 – 40 g of dietary fiber [36, 37]. Mixed-linkage β-glucans (MLGs), consisting of β-1,4- and β-1,3-linked glucose units (Fig-ure 1.2), resist digestion in the upper gastrointestinal tract. In nature, these carbohydrates serve as key matrix glycans in the cell walls of monocotyledonous plants from the taxonomic Family Po-aceae and a few other species [38, 39]. For instance, MLGs contribute to around 75% of the total cell wall material of the endosperm of oats and barley [40]. Remarkably, the β-1,3-linkages in MLG introduce kinks into the polymer chain which prevents the carbohydrates from self-assembly observed in the pure β-1,4-glucan, cellulose [41]. The distribution of the linkages is random and varies from species to species but β-1,3-linkages almost never occur in series. Therefore, the pol-ysaccharide can be considered as a progression of β-1,4-linked oligosaccharides which are con-nected through β-1,3-glycosidic linkages. The most common β-1,4-linked oligosaccharides are cellotriose (G4G4G, where β-glycosyl residues are indicated by a “G”) and cellotetraose (G4G4G4G) whereas longer cellooligosaccharides account for only 10% [42]. Thus, the ratio of 8 β-1,4- to β-1,3-glycosidic bonds usually lies within 2.2 – 2.6 : 1 for cereals [42] and ratios of cel-lotriose to cellotetraose units range from 3.0 − 4.5 (wheat), 1.8 − 3.5 (barley), 1.9 − 3.0 (rye), and 1.5 − 2.3 (oats) [41]. The unique structure of MLGs also results in various human health benefits. In the lower gastrointestinal tract, the polysaccharide, MLG, is still intact and can encapsulate other food com-ponents [41]. Thus, the viscous carbohydrate slows down the digestive attack of enzymes on the food matrix. Furthermore, the viscosity affects the diffusion and adsorption of glucose leading to constant glucose uptake and extending the feeling of satiety [41]. For instance, decreased post-prandial blood glucose levels were observed in people consuming barley or rye kernel breakfasts in contrast to people eating breakfasts consisting of white-wheat bread [43]. Similarly, oat β-glu-can significantly improved the insulin response within the first 30 min after consumption [44]. The intake of MLGs also has implications for blood cholesterol levels. The viscous poly-saccharides enclose bile acids in the upper gastrointestinal tract which stimulates bile acid biosyn-thesis. As a major substrate for the bile acid biosynthesis, cholesterol is therefore consumed and its concentration in the blood decreases [45, 46]. This blood cholesterol-lowering property of MLGs can be considered health-promoting in the context of cardiovascular diseases. Thus, gov-ernment agencies such as the European Food Safety Authority approved the advertisement of ce-real products as heart risk lowering suggesting beneficial effects at a daily intake of 3 g barley or oat β-glucan per quantified portion [36]. Apart from that, MLGs play further roles within the hu-man gut serving as antioxidants, immunomodulators, and anti-tumor compounds [47]. These effects might therefore present a promising starting point for the development of low-cost, low-risk, non-invasive treatment strategies for metabolic disorders such as diabetes and obesity, as well as for cardiovascular and inflammatory diseases. 9 1.4 Polysaccharide utilization loci encode protein machineries for efficient carbohydrate uptake The structural variety of carbohydrates presents an immense challenge for microbial deg-radation. However, the Gram-negative phylum Bacteroidetes in particular comprises species that are principal utilizers of complex carbohydrates. Their genomes incorporate specific gene clusters called polysaccharide utilization loci (PULs) [48]. PULs encode protein machineries that recog-nize, hydrolyze, and transport distinct poly- and oligosaccharides. PULs also typically include transcriptional regulators which interact with specific carbohydrates and subsequently modulate the expression of the corresponding PUL [49]. In the late 1980s, the archetypal PUL, a starch utilization system of B. theta, was identified comprising eight different genes BT3698 – 3705 named SusA – G/SusR (Figure 1.3) [50, 51]. Based on the cellular model of the starch utilization system, the presence of homologous proteins of SusC and SusD became the core identifier for PULs [52, 53]. The homologous SusC/SusD pair usually comprises a TonB-dependent transporter (TBDT) and a surface glycan-binding protein (SGBP) (Figure 1.3) [54]. Additionally, PULs in-corporate auxiliary SGBPs required for the binding, and carbohydrate-active enzymes (CAZymes) such as glycoside hydrolases (GHs) from various families (GH##, where ## is an integer that in-dicates the enzyme family) for specific cleavage of distinct carbohydrates. The size of PULs or in other words the number and role of encoded proteins varies according to the complexity of the substrate [41]. Degradation of polysaccharides via PUL systems usually includes the following steps (Figure 1.4): (1) At the cell surface, SGBPs recognize polysaccharides bringing them in close contact with extracellular CAZymes. (2) Hydrolysis of the polysaccharide by endo-acting CAZymes generates oligosaccharides. 10 (3) The oligosaccharides are then imported into the periplasm via the pair of SusC/SusD homologs. (4) In the periplasm, exo-acting CAZymes degrade the oligosaccharides producing mono-saccharides or disaccharides which are eventually transported to the cytosol. (5) Transcriptional regulators recognize specific oligosaccharides in the periplasm and in-itiate and regulate the expression of PUL genes. This concerted interaction enables rapid carbohydrate breakdown and import. Notably, the proteins on the cell surface associate to form a utilisome that brings endo-acting CAZymes, SGBPs and the TBDT in close contact and hence facilitates the carbohydrate utilization [55]. The distinct structure and function of the protein machineries provided the basis for de-tailed metagenomic, biochemical, and structural analyses of PULs from various organisms of the HGM [41]. These analyses revealed that PULs containing similar proteins can nevertheless exhibit different substrate specificities. For instance, B. uniformis, B. fluxus, and B. theta express similar PULs (Figure 1.3). While the PULs from all three species utilize laminarin, PULs from B. theta and B. uniformis also utilize yeast β-glucan and only B. uniformis barley MLG [56]. This illustrates how even small changes within PULs affect the overall function of a specific PUL and why more detailed biochemical data is required to complement the information obtained from metagenomic analyses. 11 Figure 1.3 Genetic arrangement of PULs from certain Bacteroides species. (a) The genetic organization of the archetypal starch PUL from B. theta encodes CAZymes (i.e., SusA, SusB, and SusG), SGBPs (SusD and SusF), transport proteins (i.e., SusC), and transcriptional regulators such as SusR [51]. (b) Similar Bacteroides PULs show different substrate specificities depending on the encoded GHs, SGBPs, and regulators such as hybrid two-component systems (HTCSs) [56]. b) a) 12 Figure 1.4 Model of the stepwise hydrolysis of a polysaccharide mediated by proteins of a generic PUL. Carbo-hydrate utilization is a concerted action of the different proteins incorporated in PULs. The polysaccharide is bound by the SGBPs (1) and hydrolyzed into oligosaccharides by endo-acting enzymes (2) which are then imported into the periplasm via the TBDT (3). In the periplasm, exo-acting CAZymes degrade the oligosaccharides further (4) while transcriptional regulators (SusR; HTCS) sense specific carbohydrates and modulate the expression of the proteins included in the PUL (5). Protein anchors are indicated as wobbled lines. 13 The concerted action of proteins encoded by PULs seems to be an evolutionary response to the immense competition within the HGM for nutrients found in the human diet. While fungi rely on the secretion of various CAZymes and other bacteria on extracellular carbohydrate utiliza-tion systems such as cellulosomes, PULs enable members of Bacteroidetes to break down and import complex carbohydrates rapidly [27]. However, individual species have been shown to share particular degradation products with their microbial community modulating the fitness of neigh-boring species and the overall butyrate production [57]. For instance, B. ovatus produces and re-leases CAZymes into the extracellular environment generating free inulin fragments that can be utilized by other Bacteroides species [58]. This shows how the increasing knowledge about PULs from Bacteroidetes improves our understanding of the ecosystem found in the human gut. Hence, certain PULs might indicate keystone species, define ecological niches of distinct species, or ex-plain specific interspecies interactions in the ecosystem. 1.5 Mixed-linkage β-glucan utilization locus (MLGUL) in P. copri In 2017, Tamura et. al. described a PUL from B. ovatus (BACOVA_2740−02745; Figure 1.5) which exclusively accounts for the growth on MLGs [59]. Interestingly, other members of the Bacteroidetes phylum contain syntenic PULs (Figure 1.5) according to PUL predictions in the PUL database [52]. Hence, the reported PUL might serve as a marker to predict MLG utilization. 14 Figure 1.5 Genetic organization of MLGULs. The characterized MLGUL from B. ovatus [59] and syntenic PULs from selected Bacteroides species [52] have a different genetic structure from the MLGUL of P. copri. Two years later in 2019, Fehlner-Peach et al. reported the growth of P. copri on MLG and other polysaccharides [30]. Following this observation, Dr. Jonathon Briggs, a former postdoctoral fellow in the Brumer group, investigated the transcriptome of the P. copri strain DSM 18205 after growth on various polysaccharides. Indeed, P. copri upregulated 20 different PULs and associated CAZyme clusters including a single PUL (Prevcop_05090 – 05102) in response to MLG (Figure 1.5). Sequence comparison performed by BlastP [60] predicted that the PUL contains the characteristic SGBP-A/TBDT pair, three different GHs, another SGBP, and a transcriptional regulator, a hybrid 15 two component system (HTCS). This overall structure resembles the MLG utilization locus (MLGUL) in B. ovatus. But three major differences distinguish the PULs from each other. First, the MLGUL from B. ovatus contains an endo-acting member of GH16 (BoGH16) whereas the MLGUL of P. copri encodes a predicted member of GH5 subfamily 4 (PcGH5_4). This difference might affect the hydrolysis of the long polysaccharide and might play a significant role in the downstream degradation of the oligosaccharides. Second, an additional enzyme from GH94 (PcGH94) is upregulated upon growth on MLG and might be required for the complete degrada-tion of oligosaccharides to glucose. And third, a low sequence similarity between PcGH5_4 and other members of GH5_4 and between PcSGBP-B and characterized SGBPs make a detailed struc-ture-function analysis necessary to understand the utilization of MLG entirely. 1.6 Aims of the Thesis This work deciphered the specific molecular mechanism of MLG utilization by the human gut microbe P. copri and addressed molecular differences between the MLGUL of P. copri and B. ovatus. Therefore, I characterized the gene cluster, Prevcop_05090 – 05102, through detailed ge-netic, biochemical, and structural biological investigations focusing primarily on three parts: (i) a detailed structure-function analysis of the predicted endo-acting enzyme PcGH5_4, (ii) the outline of the downstream degradation of the carbohydrates by PcGH3 and PcGH94, and (iii) the characterization of two respective SGBPs. Subsequent chapters will provide a comprehensive analysis of the experimental findings and elu-cidate the molecular functions of the five proteins in the context of MLG utilization. The Appendix contains detailed information regarding the experimental methods employed throughout this study. 16 Chapter 2: Extracellular MLG hydrolysis by the specific PcGH5_4 2.1 Introduction The utilization of complex polysaccharides typically involves extracellular endo-acting en-zymes that can bind the long carbohydrate chain and hydrolyze distinct linkages along the back-bone generating smaller oligosaccharides. Notably, the genomic and transcriptomic data pertaining to the MLGUL of P. copri indicated the presence of a member belonging to GH5_4. Enzymes within GH5_4 comprise endo-active cellulases, xyloglucanases, and β-1,3/1,4-glucanases that use the classical Koshland double-displacement mechanism. Today, over 120 members of GH5_4 have been characterized highlighting their broad distribution across bacteria and eukaryotes [61]. However, predicting the substrate specificity based on sequence analysis alone lacks in accuracy and level of detail due to the broad structural variety within GH5_4. Furthermore, species such as B. ovatus contain an endo-acting member of GH16 in the MLGUL [59]. Thus, an in-depth struc-ture-function analysis is necessary to investigate the role and function of PcGH5_4 in the context of MLG utilization. 2.2 Results and discussion 2.2.1 PcGH5_4 is an endo-acting 1,3/1,4-β-glucanase Sequence comparisons by BlastP [60] suggested that the upregulated gene, Prevcop_05101, encodes a member of GH5_4. Signal peptide analysis performed by SignalP [62] and Psipred [63] indicated that the 406 amino acid-long protein contains a signal peptide consti-tuted by 20 N-terminal amino acid residues (Figure 2.1). After anchoring the protein to a lipid molecule at C21 via a diacylglyceryl transferase, the signal peptidase II cleaves the signal peptide 17 [64]. Thus, PcGH5_4 might be located at the outside of the cell where it comes near diverse poly-saccharides. To investigate the potential for polysaccharide degradation, four different constructs (Fig-ure 2.1), PcGH5_4C21-E406 (46.5 kDa), PcGH5_4T41-E406 (44.5 kDa), PcGH5_4T41-E406(E197A) (44.5 kDa), and PcGH5_4T41-E406(E331A) (44.5 kDa), were successfully cloned, expressed and purified (Figure 2.2). The construct PcGH5_4C21-E406 was used for biochemical characterization while crys-tallization experiments were performed with the truncated PcGH5_4T41-E406 where a disordered linker was excluded. Figure 2.1 Structure of the cloned and expressed constructs of PcGH5_4. 18 Figure 2.2 SDS-PAGE gels of the purified PcGH5_4 constructs, PcGH5_4C21-E406 (46.5 kDa), PcGH5_4T41-E406 (44.5 kDa), PcGH5_4T41-E406(E197A) (44.5 kDa), and PcGH5_4T41-E406(E331A) (44.5 kDa). The molecular weight (MW) of the bands of the protein ladder is indicated in kDa on the left. An initial substrate screen showed that PcGH5_4C21-E406 is a β-glucanase with high activity on barley MLG (Figure 2.3). While low activities on some β-1,4-linked glucans including xyloglu-can, glucomannan, hydroxyethyl cellulose, and carboxymethyl cellulose were observed, the en-zyme did not hydrolyze the common food additive xanthan gum E415 [65] that also contains β-1,4-linked glucose units in its backbone (Figure 1.2). The enzyme displayed no activity on 1,3-linked β-glucans such as curdlan, laminarin, or yeast β-glucan, and did not cleave pustulan, a β-19 1,6-linked glucan. Similarly, no hydrolytic activity was detected for xylan, a polysaccharide with xylose monosaccharide units in its backbone (Figure 2.3). Figure 2.3 Initial screen for the hydrolytic activity of GH5_4 on various polysaccharides. (a) Specific activities of PcGH5_4C21-E406 on different polysaccharides at 37 °C; (b) Comparison of the specific activities of non-MLG sub-strates. Structures of the individual polysaccharides are provided in Figure 1.2. Michaelis-Menten kinetics at the optimal pH of 4.5 and temperature of 50 °C (Figure 2.4, Figure 2.5) confirmed that MLG is the preferred substrate of PcGH5_4C21-E406 with a kcat of 1,490 ± 67 s-1 and a Km of 0.056 ± 0.0092 mg/mL (Table 2.1). Thus, the specificity, as determined by the ratio of kcat/Km values of the enzyme for barley MLG was ca. 560 times higher than for xyloglucan and ca. 440 times higher than for glucomannan. Even for other linear polysaccharides, carboxymethyl cellulose and hydroxyethyl cellulose, substrate specificity was significantly lower in comparison to MLG. MLG is therefore the main substrate for the PcGH5_4. b) a) 20 Figure 2.4 Specific activity of PcGH5_4C21-E406 on barley MLG in different buffers and different temperatures. (a) The pH-profile was determined at 37 °C in 40 mM buffer: sodium citrate (NaCit), sodium phosphate (NaPi), gly-cine-sodium hydroxide (Gly-OH), 2-(N-morpholino)ethanesulfonic acid (MES). (b) Temperature profiles were ob-tained in 40 mM NaCit buffer (pH 4.5). Figure 2.5 Michaelis-Menten kinetics for the hydrolysis of different polysaccharides. (a) Degradation of barley MLG and (b) xyloglucan, glucomannan, hydroxyethyl cellulose, and carboxymethyl cellulose by PcGH5_4C21-E406 in 40 mM NaCit buffer (pH 4.5) at 50 °C. b) a) a) b) 21 Table 2.1 Kinetic data for the digestion of various polysaccharides by the endo-active PcGH5_4C21-E406. Polysaccharide kcat (s-1) Km (mg mL-1) kcat/Km (s-1 mg-1 mL) Assay Barley MLG 1,490 ± 67 0.056 ± 0.0092 27,000 BCA Xyloglucan 21 ± 1.8 0.44 ± 0.071 48 BCA Glucomannan 14 ± 1.0 0.23 ± 0.039 61 BCA Hydroxyethyl cel-lulose 1.74 ± 0.064 0.50 ± 0.039 3.5 BCA Carboxymethyl cellulose 1.18 ± 0.051 0.19 ± 0.023 6.2 BCA 2.2.2 PcGH5_4 preferentially hydrolyzes β-1,3-linkages in MLG Monitoring the degradation of MLG over time via HPAEC-PAD showed that PcGH5_4C21-E406 works in an endo-dissociative mechanism generating longer oligosaccharides first. To distin-guish the small oligosaccharides produced in the limit digest, HPAEC-PAD was performed using a second gradient (Figure 2.6). The comparison with standard solutions confirmed that cellobiose (G4G), cellotriose (G4G4G), and G3G4G4G concentrations accumulate over time (Figure 2.6). At the beginning of the digestion, PcGH5_4C21-E406 mainly produced cellotriose, cellotetraose (G4G4G4G), and cellopentaose (G4G4G4G4G) which indicated specific hydrolysis of β-1,3-link-ages. Later on, the concentration of cellobiose increased while the amounts of cellotetraose and cellopentaose decreased suggesting enzymatic degradation of β-1,4-linkages. 22 Figure 2.6 Product analysis of the digestion of barley MLG by PcGH5_4C21-E406 via HPAEC-PAD. (a) Profile of produced oligosaccharides over time and (b) chromatograms of the degradation using gradient II for better separation of smaller oligosaccharides. Chromatograms from reactions are colored in blue and standard solutions are indicated in grey. a) b) 23 To further validate that PcGH5_4C21-E406 hydrolyzed β-1,3-linkages between the cellotriose and cellotetraose blocks in MLG, the amounts of the products, glucose, cellobiose, cellotriose, and cellotetraose, were determined from the digestion profile (Figure 2.7). Then, ratios of cellotriose to cellotetraose were calculated assuming that nearly all of the cellobiose generated after 2 h of digestion originated from degraded cellotetraose. The calculated ratios varied between 1.8 and 3.0 which corresponds to reported ratios of cellotriose to cellotetraose blocks in barley MLG [42] pointing towards a preference for hydrolytic cleavage of β-1,3-linkages. Figure 2.7 Amounts of generated products during the digestion of barley MLG by PcGH5_4C21-E406. (a) Time-dependent production and hydrolysis of selected oligosaccharides; (b) ratio of [G4G4G] to [G4G4G4G]*. Since side reactions generating G4G such as the degradation of G4G4G4G4G were negligible during the first 2 h of degradation (Figure 2.6), [G4G4G4G] was estimated using the equation: [G4G4G4G]* = ([G4G]/2) + [G4G4G4G]. a) b) 24 2.2.3 Specific subsite arrangement for the cleavage of MLG-oligosaccharides Since PcGH5_4C21-E406 hydrolyzed β-1,3- and β-1,4-linkages, an in-depth analysis of po-tential cleavage sites was performed using short oligosaccharides. HPAEC-PAD analysis of the oligosaccharides (Figure 2.8) confirmed that PcGH5_4C21-E406 was unable to hydrolyze cellobiose (G4G), cellotriose (G4G4G) and G3G4G4G. Furthermore, the data revealed that the enzyme did not cleave MLG-trisaccharides, G3G4G and G4G3G. However, cellotetraose and two of the MLG-tetrasaccharides, G4G4G3G and G4G3G4G, were hydrolyzed. The hydrolysis of G4G4G3G gen-erated glucose (Glc), cellobiose, laminaribiose (G3G), and cellotriose whereas hydrolysis of G4G3G4G resulted in cellobiose exclusively. Degradation of cellotetraose produced cellobiose but minor amounts of cellotriose were also detected suggesting a potential side reaction. The cleav-age of β-1,4-linkages was also observed for cellopentaose (G4G4G4G4G) and generated cellobi-ose and cellotriose. However, PcGH5_4C21-E406 did not deplete laminarioligosaccharides, G3G, G3G3G, and G3G3G3G (Figure 2.8). The observed oligosaccharide digestions underlined that PcGH5_4 is able to hydrolyze β-1,3- and β-1,4-linkages. Moreover, the data revealed that at least four glucose molecules are required for sufficient substrate recognition and that β-1,3-linkages have to occur isolated within the carbohydrate to be hydrolyzed. 25 Figure 2.8 HPAEC-PAD analysis of the digestion of oligosaccharides by PcGH5_4C21-E406. The enzyme only hy-drolyzed carbohydrates containing more than three glucose units which are either cellulose- or MLG-derived oligo-saccharides. The cross indicates blank solutions without enzyme. To further specify the site of hydrolysis within a carbohydrate motif, enzymatic subsites have been defined labeling the monosaccharide units adjacent to the point of hydrolytic cleavage subsites -1 and +1. Subsite numbers towards the reducing end of the carbohydrate are positive and increase away from the point of cleavage (i.e., +2, +3, …) while subsites towards the non-reducing end of the carbohydrate are numbered negatively and decrease away from the point of cleavage (i.e., -2, -3) [66]. Michaelis-Menten kinetics of the oligosaccharide degradations (Figure 2.9, Table 2.2) indicated that the cleavage of the β-1,3-linkage in the MLG-tetraose, G4G3G4G, was around 26 eight times faster than the cleavage of the central β-1,4-linkage in cellotetraose, G4G4G4G. Fur-thermore, the hydrolysis of cellotetraose into cellobiose, where the tetrasaccharide was bound in subsites -2 to +2 was ca. seven times more efficient than a -3 to +1 subsite configuration, generat-ing glucose and cellotriose (Table 2.2, Figure 2.10). Interestingly, PcGH5_4T41-E406 hydrolyzed the β-1,3-linkage and the central β-1,4-linkage of the MLG-tetraose, G4G4G3G. The catalytic effi-ciency of cleaving the β-1,3-linkage was similar to the hydrolysis of the β-1,4-linkage since subsite -3 was occupied by a glucose molecule. Nevertheless, the rate of hydrolysis of G4G4G3G bound in subsites -3 to +1 was more than 20 times less specific than the hydrolysis of G4G3G4G bound in subsites -2 to +2 (Table 2.2, Figure 2.10). This suggested that the +2 subsite contributes more to the hydrolysis of β-1,3-linkages than subsite -3. Hence, the data highlighted the strong prefer-ence of PcGH5_4 towards β-1,3-linkages and pointed out that the contribution of the +2 subsite changes the kinetic reaction parameters drastically. Whereas the enzyme hydrolyzed G4G3G4G and G4G4G3G, the third MLG-tetrasaccha-ride, G3G4G4G, remained intact. This revealed that neither preferred hydrolysis of the β-1,3-link-age upon binding the substrate in subsites -1 to +3 nor cleavage of β-1,4-linkages upon binding of the substrate in a -2 to +2 or -3 to +1 subsite configuration are possible (Figure 2.8, Figure 2.10). Thus, the enzyme requires the occupation of subsite -2 by a glucose unit that is linked via a β-1,4-glycosidic bond to the glucose molecule in subsite -1. 27 Figure 2.9 Initial rate kinetics for the hydrolysis of various oligosaccharides by PcGH5_4C21-E406. (a) Michaelis-Menten kinetics and (b) the expansion of initial rate kinetics of oligosaccharide degradation by PcGH5_4C21-E406 indi-cated the preference for cleavage of β-1,3-linkages. Figure 2.10 Alignment of selected oligosaccharides to the subsites of PcGH5_4. The binding of substrates that resulted in hydrolytic cleavage are indicated by a green tick. a) b) 28 Table 2.2 Kinetic data for the digestion of shorter oligosaccharides by the endo-active PcGH5_4C21-E406 (Figure 2.9). Oligosaccharide Product kcat (s-1) Km (μM) kcat/Km (s-1 M-1) Assay G4G NDa NDa NDa NDa HPAEC-PAD G3G NDa NDa NDa NDa HPAEC-PAD G3G4G NDa NDa NDa NDa HPAEC-PAD G4G3G NDa NDa NDa NDa HPAEC-PAD G4G4G NDa NDa NDa NDa HPAEC-PAD G3G4G4G NDa NDa NDa NDa HPAEC-PAD G4G4G3G G4G + G3G 6.8 ± 0.17 1,700 ± 130 4,000 HPAEC-PAD G4G4G3G G4G4G + G 2.7 ± 0.20 500 ± 140 5,400 HPAEC-PAD G4G3G4G G4G + G4G 291 ± 8.1 2,600 ± 200 110,000 HPAEC-PAD G4G4G4G G4G + G4G 37.89 ± 0.096 2,220 ± 16 17,000 HPAEC-PAD G4G4G4G G4G4G + G 1.8 ± 0.24 700 ± 310 2,600 HPAEC-PAD G3G3G3G NDa NDa NDa NDa HPAEC-PAD G4G4G4G4G G4G4G + G4G 328 ± 5.4 720 ± 41 460,000 HPAEC-PAD a ND, not detected.29 The kinetic data also showed that the occupation of more than four subsites increased the substrate specificity of the hydrolysis. For instance, cellopentaose was a 27 times better substrate than cellotetraose (Table 2.2). Since the transition state activation energy (ΔGǂ) can be expressed as the sum of the free energy of individual subsites, the difference (ΔΔGǂ) between individual transition state activation energies (ΔGǂn+1 and ΔGǂn) of the homologous substrates with chain length n and n+1 describes the contribution of an individual subsite to the transition state stabili-zation. This contribution of the additional glucose unit to the stabilization of the transition state was calculated from the kcat/Km values of the degradation of cellotetraose and cellopentaose using the equation: ΔΔGǂ = ΔGǂn+1 – ΔGǂn = – RT ln[(kcat/Km)n+1/(kcat/Km)n] [67] The calculated difference in transition state activation energy ΔΔGǂ of the digestion of cellopen-taose to cellobiose and cellotriose vs the degradation of cellotetraose to cellobiose was – 8.5 kJ/mol. Similarly, a difference ΔΔGǂ of – 13 kJ/mol was calculated for the digestion of cel-lopentaose to cellobiose and cellotriose in comparison to the degradation of cellotetraose to cel-lotriose and glucose. Nevertheless, it is unclear which subsites were responsible for the energetic stabilization due to two potential subsite binding modes (Figure 2.10). Digestion of cellopentaose by PcGH5_4C21-E406 in [18O] water (Figure 2.11) revealed that incorpo-ration of [18O] mainly occurred at the anomeric position of cellotriose. Therefore, PcGH5_4C21-E406 binds the oligosaccharide in subsites -3 to +2 preferentially and hydrolyzes the linking glyco-sidic bond (Figure 2.10). Therefore, three well-defined negative subsites might be present. 30 Figure 2.11 Mass spectra of products generated from the digestion of cellopentaose by PcGH5_4C21-E406. Prod-ucts of the digestion of cellopentaose (a) in [16O] water and (b) in [18O] water displayed a shift of the isotopic distri-bution for cellotriose indicating that hydrolytic cleavage mainly occurs in a three-plus-two configuration. 2.2.4 Structural features of the MLG-specific PcGH5_4 To examine the molecular basis of substrate recognition and conversion, a detailed struc-ture-function analysis of PcGH5_4 was required. The truncated construct PcGH5_4T41-E406 crys-tallized in 0.2 M lithium chloride containing 19% polyethylene glycol 3,350 and 5 mM cel-lotetraose giving rod-shaped crystals that were frozen using 30% ethylene glycol as cryoprotectant. The protein crystal structure was solved to a resolution of 1.6 Å by molecular replacement using an Alphafold [68] model of the construct PcGH5_4T41-E406 (Table A.2). The tertiary structure has a) b) 31 the space group P1211 and contains four molecules in the asymmetric unit (Figure 2.12). The mol-ecules within the asymmetric unit adopt an (α/β)8-barrel fold consisting of eight inner β-sheets enclosed by eight longer α-helices (Figure 2.12) which is typical for members of GH5_4 [69]. Despite these structurally conserved features, variations along the loop regions connecting the α-helices and β-sheets result in low sequence similarity of PcGH5_4T41-E406 and its closest structural homologs (Table 2.3) and complicate the prediction of substrate specificity. Although the protein was cocrystallized with cellotetraose, no electron density that might represent a carbohydrate moi-ety was observed. This may have resulted from a tight crystal packing or the inability to stabilize short carbohydrates in the active site since PcGH5_4C21-E406 hydrolyzed cellotetraose to cellobiose (Figure 2.8). Figure 2.12 Crystal structure of PcGH5_4T41-E406. (a) Four chains of PcGH5_4T41-E406 are coordinated within the asymmetric unit. (b) All of the four chains adopt the (α/β)8-barrel fold typical for enzymes of GH5_4 containing two catalytic residues E197 and E331 (gray) in the center. a) b) 32 Table 2.3 Structural homologs of PcGH5_4T41-E406 according to a DALI search[70, 71]. Enzyme family PDB-ID Organism Predominant ac-tivity Z score r.m.s.d. % ID Ref. GH5_4 6Q1I Clostridium longisporum glucomannanase 46.6 1.6 33 [72] GH5_4 6MQ4 Acetivibrio cellulolyticus glucomannanase 46.4 1.6 35 [72] GH5_4 2JEQ Paenibacillus pabuli xyloglucanase 45.9 1.7 35 [73] GH5_4 5OYD Cellvibrio japonicus Ueda107 xyloglucanase 45.8 1.7 36 [69] GH5_4 6PZ7 Clostridium acetobutyli-cum ATCC 824 glucomannanase 45.6 1.6 35 [72] GH5_4 5D9M Prevotella bryantii B14 β-1,3/1,4-glucanase 41.4 1.9 32 [74] Structural comparison with the five closest structural homologs (Table 2.3) determined via a DALI search [70, 71] indicated two conserved glutamic acid residues E197 and E331 located on loop 4 and loop 7 respectively (Figure 2.12) that may serve as catalytic residues in the classical Koshland double-displacement mechanism (Scheme 2.1). E197 may act as the acid/base during hydrolysis while E331 may serve as the nucleophile attacking the anomeric carbon. Indeed, site-directed mutagenesis revealed that both of the catalytic residues were required for the hydrolysis of barley MLG (Table 2.4). Recently, both catalytic mutants were cocrystallized with several oli-gosaccharides including G4G3G4G, an MLG-hexamer, and MLG-heptamer (Figure 2.13) and will be diffracted in the near future. 33 Scheme 2.1 Koshland double-displacement mechanism for the retaining hydrolysis of a β-1,4-glucan. Table 2.4 Activity of catalytic mutants of PcGH5_4 on barley MLG. Protein construct Mutant Specific activity on barley MLG (μmol min-1 mg-1) Relative activity (%) PcGH5_4T41-E406 No 720 ± 47 100 PcGH5_4 T41-E406(E197A) Acid/base 0.071 ± 0.0094 0.01 PcGH5_4 T41-E406(E331A) Nucleophile 0.003 ± 0.0043 0 34 Figure 2.13 Selected crystallization conditions of the catalytic mutants, PcGH5_4T41-E406(E197A) and PcGH5_4T41-E406(E331A). 2.2.5 Conserved negative subsites stabilize β-1,4-linked carbohydrates in the active site The crystal structure of PcGH5_4T41-E406 revealed a ca. 30 Å long substrate-binding cleft that is formed by the loops and narrows towards the catalytic residues (Figure 2.14). Regarding the structural homologs, several amino acid residues including N56, W85, H151, N196, Y277, and W370 are conserved within the active site (Figure 2.14). These highly conserved residues were found exclusively within the negative subsites. Superposition of the crystal structure of PcGH5_4T41-E406 with a cellotetraose ligand that was crystallized with a GH5_4 member from Caldicellulosiruptor sp. F32 (PDB-ID: 5H4R; [75]) revealed three well-defined negative subsites (Figure 2.15). The residues H151, H152, N196, and W370 may interact with the glucose unit in the -1 subsite via hydrogen bond interactions and hydrophobic stacking interactions. In addition, residues N56, W370, and N372 might stabilize the glucose units from the oligosaccharide in sub-site -2. Here, hydrogen bond interactions might also occur between the flexible residue E380 and the C2 and C3 hydroxyl group of the glucose. Meanwhile, the solvent-exposed aromatic residue W85 might interact with the glucose unit at subsite -3 (Figure 2.15). The alignment suggested π-stacking interactions between the electrons of the aromatic ring and the carbohydrate. Individual 35 mutations of residues corresponding to N56, H152, and N372 to alanine (Figure 2.15) in an MLG-degrading GH5_4 from the marine bacterium Zobellia galactanivorans (ZgGH5_4; PDB-ID: 6GL2; [76]) decreased the activity on barley MLG 56 times for N77A, five times for H156A, and six times for N358A. Figure 2.14 Structural comparison of PcGH5_4T41-E406 with two selected closest structural homologs. In addition to the catalytic residues (orange), several amino acid residues in the binding site of PcGH5_4T41-E406 are conserved (violet). Corresponding residues of GH5_4 members from Acetivibrio cellulolyticus (PDB-ID: 6MQ4; [72]) and Pae-nibacillus pabuli (PDB-ID: 2JEQ; [73]) are indicated in green and magenta respectively. 36 Figure 2.15 Overlay of the crystal structure of PcGH5_4T41-E406 with carbohydrate ligands in the negative sub-sites. (a) Superposition of a cellotetraose ligand (red; PDB-ID: 5H4R; [75]) with the crystal structure of PcGH5_4T41-E406 revealed three conserved negative subsites. The three residues, N77, H156, and N358, that were mutated in ZgGH5_4 (PDB-ID: 6GL2; [76]) are colored brown. (b) The negative subsites might accommodate a xyloglucan oligosaccharide, XXXG (yellow, PDB-ID: 5D9M; [74]), that was superposed with the active site of PcGH5_4T41-E406. Furthermore, H374 might adopt a similar conformation as H365 in the crystal structure of PpGH5_4 (PDB-ID: 2JEQ; a) b) 37 [73]) stabilizing xylose. All catalytic residues of PcGH5_4T41-E406 are highlighted in orange, conserved residues in the active site are colored in violet, and residues that are not conserved but might interact with the carbohydrate are indicated in cyan. PcGH5_4 showed high activity on MLG and low activity on xyloglucan similar to previ-ously characterized structural homologs including ZgGH5_4 and a GH5_4 from Prevotella bry-antii (PbGH5_4; Table 2.3) [74]. To identify structural constraints for the branched polysaccharide xyloglucan, the crystal structure of PcGH5_4T41-E406 was superposed with the xyloglucan hepta-saccharide XXXG (Xyl3Glc4, where “X” represents a xylose unit attached to glucose, and “G” an unsubstituted glucose residue) from the complex of PbGH5_4 with XXXG·XXXGXXXG (PDB-ID: 5D9M; [74]) (Figure 2.15). Similar to PbGH5_4, the binding cleft of PcGH5_4T41-E406 only tightens around the catalytic residues but widens farther away. Thus, residues in the negative sub-sites might interact with side branches. For instance, hydrogen bond interactions between E68 and xylose may stabilize the carbohydrate in the side pocket at subsite -1 (Figure 2.15). In subsite -3, the xylose branch at the glucose unit appears to clash with the electron density of H373. But this residue is present in two different conformations and one of them might stabilize the substrate. In fact, in the structure of GH5_4 from Paenibacillus pabuli (PpGH5_4; PDB-ID: 2JEQ; [73]), a histidine residue adopts a conformation that stabilizes the xylose side branch in this position (Fig-ure 2.15). In summary, the active site contains three conserved negative subsites that form a binding platform for the recognition of β-1,4-linked glucans. Multiple residues may interact with the glu-cose moieties through hydrogen bond networks and aromatic stacking interactions and therefore 38 promote interactions with carbohydrates containing cellotriose and cellotetraose blocks. Further-more, the data supports the kinetic analysis underlining that a β-1,4-linkage between subsite -2 and -1 is required for the hydrolysis of oligosaccharides. The drastic transition state stabilization after the addition of one glucose unit to cellotetraose is, therefore, most likely the result of interactions in subsite -3 and explains why cellopentaose is predominantly bound and cleaved in a three-plus-two configuration. However, the structural features in the negative subsites do not explain the low activity of PcGH5_4 on 1,4-linked β-glucans such as xyloglucan. Therefore, a close look at the positive subsites might indicate the reasons for the substrate preference. Notably, the structural overlay with cellotetraose (Figure 2.15) showed how the glucose unit in subsite +1 may be dis-torted with regard to the orientation of β-1,4-linked glucose moieties bound in subsites -3 to -1 and hence might indicate why hydrolyses of long chains of β-1,4-linked carbohydrates such as xy-loglucan or cellulose derivatives were less preferred. 2.2.6 Unique positive subsites of PcGH5_4 with potential for substrate discrimination The active site groove tightens at subsites +1 and +2 due to two amino acid residues W204 and W280. Interestingly, none of the amino acid residues within the positive subsites are conserved among the five closest structural homologs identified by the DALI search [70, 71] (Figure 2.16). However, the MLG-specific PbGH5_4 has a similar configuration of the positive subsite. Super-position of the xyloglucan ligand XXXG and XXXG·XXXGXXXG bound in the positive subsite of PbGH5_4 (PDB-ID: 5D9N, 5D9M; [74]) with the crystal structure of PcGH5_4T41-E406 revealed that the glucose backbone of xyloglucan may interact with W204 and W280 at subsite +1 and +2 through stacking interactions (Figure 2.16). In addition, the active site of PcGH5_4T41-E406 forms a pocket at the +1 subsite which may accommodate an additional xylose residue at subsite +1. 39 Previous studies have highlighted that the size of this pocket is crucial for activity on xyloglucan [77] and it seems that the sidechain in the +1 subsite levers the backbone of the carbohydrate out of the binding cleft (Figure 2.16). However, cellotetraose from the complex of PbGH5_4 (PDB-ID: 5D9O; [74]) fits tightly into the active site of PcGH5_4T41-E406 (Figure 2.16). In comparison to the xyloglucan ligands, the glucose unit in subsite +1 is flipped 180° along the C1-C4 axis. Hence, the hydroxyl group at C3 of the glucose in subsite +1 is positioned in close proximity to the catalytic residues. Since the enzyme PbGH5_4 from the complex with cellotetraose does not contain any amino acid residue in subsite +4 that could promote interactions with a carbohydrate, no clear electron density was observed for the reducing glucose residue in the structure of PbGH5_4 in complex with cel-lotetraose. By comparison, the presence of the aromatic residue W248 in the crystal structure of PcGH5_4T41-E406 suggested a stabilizing stacking interaction of the carbohydrate in subsite +4 (Fig-ure 2.16) that might promote tight substrate binding. In fact, a previous study by Meng et al. showed that mutation of the residues E252 and R280 (Figure 2.16) of a GH5_4 specific for MLG (F32EG5) from Caldicellulosiruptor sp. F32 (PDB-ID: 5H4R), corresponding to the amino acid residues W248 and Q278 in PcGH5_4T41-E406, displayed lower activity on barley MLG or carbox-ymethyl cellulose [75]. For the double mutant E252A and R280A, the specific activity on barley MLG decreased to 30% whereas the activity on the 1,4-linked carboxymethyl cellulose only de-creased to 70% for E252A and 45% for R280A. Similarly, the double mutant W203A/Y282W of F32EG5 where W203 corresponds to W204 in PcGH5_4T41-E406 and Y282 to W280 respectively (Figure 2.16) was eight times less active on both linear substrates highlighting the importance of the amino acid residues in subsite +1 and +2. 40 a) b) c) 41 Figure 2.16 Superposition of potential carbohydrate ligands and the crystal structure of PcGH5_4T41-E406. (a) Cellotetraose (dark green; PDB-ID: 5D9O; [74]) and the xyloglucan oligosaccharides, (b) XXXG (light green; PDB-ID: 5D9N; [5]) and (c) XXXGXXXG (yellow; PDB-ID: 5D9M; [5]), were modeled into the positive subsites of PcGH5_4T41-E406. The relative orientation of the carbohydrates in the active sites are represented by solid lines and the positions of the carbon or oxygen atom closest to the catalytic residues are indicated. Catalytic residues are highlighted in orange and residues in the active site groove that are not conserved but might be responsible for carbohydrate interactions are colored in cyan. Mutation of three residues of F32EG5 (pink; PDB-ID: 5H4R; [75]), W203, E252, and R280, displayed in (a) decreased the activity on MLG suggesting a similar importance of corresponding residues of PcGH5_4. Thus, a unique set of amino acid residues located in the subsites +1 and +2 forms a narrow binding cleft suggesting better stabilization of carbohydrates comprising an isolated β-1,3-linkage. In addition, the presence of W248 in the +4 subsite might point towards a better fit of longer MLG-derived carbohydrates. Moreover, W280 located in subsite +2 might be responsible for the signif-icant increase in the rate of hydrolysis of G4G3G4G vs G4G4G3G and a potential aromatic stack-ing might mitigate the energetic loss of the unoccupied -3 subsite. 2.2.7 Catalytic comparison with other MLGases The data showed that PcGH5_4 is an extracellular enzyme that is well-equipped for the endo-hydrolysis of MLG. The ability of PcGH5_4 to cleave 1,3- and 1,4-linkages complies with activities observed for other characterized GH5_4 members [74-76]. However, the observed pref-erence for the hydrolysis of β-1,3-linkages clearly distinguishes the PcGH5_4 from classical MLG degrading enzymes which are members of GH16 [59, 78]. For instance, BoGH16 from the MLGUL of B. ovatus only hydrolyzes β-1,4-linkages that succeed a β-1,3-linkage generating 42 MLG-triose, G4G3G, and MLG-tetraose, G4G4G3G, in the limit-digest (Figure 2.17) [59]. Inter-estingly, PcGH5_4 can digest the MLG-tetraose whereas BoGH16 is unable to hydrolyze the main products of enzymatic digestion of PcGH5_4, cellotriose (G4G4G), and cellotetraose (G4G4G4G). Furthermore, the substrate specificity of PcGH5_4C21-E406 for barley MLG (Table 2.1) was 100 times higher than the reported kcat/Km of BoGH16 (238 s-1 mg-1 mL) and over 300 times higher than other MLG-degrading GH16 enzymes from B. theta (72 s-1 mg-1 mL) or B. uni-formis (78 s-1 mg-1 mL) [56, 59]. These significant catalytic differences might play a crucial role for the fitness of P. copri in the competitive environment of the human gut where rapid MLG utilization might provide an advantage over other species such as B. ovatus. Figure 2.17 Hydrolysis of MLG by the endo-acting PcGH5_4 in comparison to BoGH16. Cleavage sites are in-dicated in red. 43 2.3 Conclusion The enzyme, PcGH5_4, is likely located on the outer cell membrane and therefore deter-mines the initial step of carbohydrate utilization. The presented data revealed that PcGH5_4 is highly specific towards the kinked 1,3/1,4-linked β-glucan MLG since it hydrolyzes isolated β-1,3-linkages preferentially. Product analysis by HPAEC-PAD confirmed that the enzyme is an endo-acting β-glucanase that cleaves the polysaccharide in a dissociative manner. Furthermore, analyses of oligosaccharide digestions highlighted the importance of a specific subsite arrange-ment where a β-1,4-glycosidic bond has to link the glucose units in subsite -2 and -1 and where the occupation of subsite +2 increases the stabilization of the carbohydrate drastically. The crystal structure of PcGH5_4 revealed that the enzyme is highly adapted for MLG breakdown. The conserved negative subsites might promote the specific recognition and subse-quent cleavage of β-1,4-linked glucans, the narrow binding cleft containing the conserved catalytic residues might improve the hydrolysis of linear polysaccharides, and residues located in the posi-tive subsites may discriminate against carbohydrates without β-1,3-linkages. Overall, the enzyme represents a specificity determinant for the carbohydrate utilization mediated by the MLGUL of P. copri and therefore indicates a high degree of specialization by P. copri. Moreover, the mode of action and catalytic efficiency of PcGH5_4 clearly distinguishes the MLGUL of P. copri from the MLGUL of B. ovatus, a competitor in the human gut, suggesting that convergent evolution occurred within the Bacteroidetes genus. Furthermore, this might ex-plain the excellent growth of P. copri on MLG. 44 Chapter 3: Downstream degradation of oligosaccharides mediated by PcGH3 and PcGH94 3.1 Introduction PULs encode a variety of CAZymes located on the outer cell membrane, the periplasm and cytosol. While extracellular enzymes usually process the conversion of polysaccharides to oligo-saccharides, intracellular enzymes degrade the oligosaccharides into their monosaccharide build-ing units. Notably, two genes, Prevcop_05090 and Prevcop_05096, were upregulated in response to MLG in addition to the endo-acting PcGH5_4 and were predicted to encode two GHs, PcGH3 and PcGH94. Enzymes belonging to GH3 comprise a broad variety of exo-activities including β-D-glucosidases, α-L-arabinofuranosidases, β-D-xylopyranosidases, or N-acetyl-β-D-glucosamini-dases and their structural diversity complicates clustering into subfamilies [79]. Nevertheless, all members of GH3 hydrolyze glycosidic bonds via the classic Koshland double-displacement mech-anism [80]. Interestingly, members of GH94 follow a different mechanism for the cleavage of glycosidic bonds. Instead of water, phosphate attacks the anomeric carbon directly while an aspar-tic acid residue serves as the proton donor for the glycosidic oxygen atom [81]. The acid-catalyzed hydrolysis inverts the anomeric configuration and generates 1-phosphate glycosides. In contrast to the MLGUL of P. copri, the MLGUL of B. ovatus only contains a single BoGH3MLG for the complete breakdown of oligosaccharides, produced by the extracellular BoGH16, to glucose. Therefore, this chapter analyzes the mechanism behind the oligosaccharide digestion by PcGH3 and evaluates the potential and the necessity of the additional PcGH94 in-cluded in the MLGUL of P. copri. 45 3.2 Results and discussion 3.2.1 PcGH3 and periplasmic oligosaccharide breakdown According to a BlastP search [60], the gene Prevcop_05096 encodes a protein whose 772 amino acid long sequence resembles other members of GH3. Regarding sequence analysis per-formed by SignalP [62] and Psipred [63], the 19 N-terminal residues constitute a signal peptide (Figure 3.1) that might be cleaved by the membrane-bound serine protease signal peptidase I sug-gesting that the enzyme is localized within the periplasm. To further understand the role and func-tion of that enzyme for MLG utilization, a biochemical and structural analysis was performed. The PcGH3 construct, PcGH3V23-R772 (86.0 kDa), that excluded the N-terminal signal pep-tide and the three following amino acid residues was cloned, expressed, and purified. The profile of the size-exclusion chromatography (SEC) indicated that the protein dimerized in solution (Fig-ure 3.2). Regarding the Thermal Shift Assay, the protein was most stable in the pH range of 5.0 – 7.0 (Figure 3.2). Figure 3.1 Structure of the cloned and expressed constructs of PcGH3. 46 Figure 3.2 Purification and stability analysis of PcGH3. (a) SEC profile and (b) SDS-PAGE gel of the dimer peak of PcGH3V23-R772 (86.0 kDa) indicated that the enzyme dimerized in solution. Masses of the calibration standards a) c) b) 47 (carbonic anhydrase, BSA, ALS dehydrogenase, β-amylase, ferritin, and thyroglobulin) are displayed in the chroma-togram. The MW of the protein ladder bands are provided on the left in kDa. (c) A Thermal Shift Assay of the protein in different buffers showed higher melting temperatures in the pH range from 5.0 to 7.0. To identify the specificity of the enzyme towards different glycosidic linkages and mono-saccharide building units, the activity of PcGH3V23-R772 was screened against seven para-nitrophe-nol (pNP) glycosides (Figure 3.3). The enzyme showed much higher specific activity on pNP-β-glucose in comparison to pNP-α-glucose. Apart from pNP-β-glucose, PcGH3V23-R772 hydrolyzed pNP-β-xylose with lower activity but did not cleave pNP linked to other stereoisomers of glucose, e.g., galactose and mannose. Since xylose, an aldopentose, can occur in the pyranose conformation where the hydroxyl group of C5 forms the hemiacetal, the corresponding pNP-β-glycoside only lacks the C6 atom and the corresponding hydroxyl group in comparison to glucose. This vacancy might destabilize the pNP-β-xylose in the active site and cause low activity. Hence, the substrate screen suggested that PcGH3 serves as a traditional β-glucosidase in the MLGUL. Figure 3.3 Specific activity of PcGH3V23-R772 on pNP-glycosides. (a) The initial substrate screen on various pNP- a) b) 48 glycosides showed that the enzyme was specific for the cleavage of β-linked glucose units. (b) pH profile of the di-gestion of pNP-β-glucose indicated high activities around pH 6.0. After determining the pH profile for PcGH3V23-R772 using pNP-β-glucose (Figure 3.3), the ability of the enzyme to degrade various oligosaccharides was investigated (Figure 3.4). Interest-ingly, the enzyme hydrolyzed the β-1,4-linkage in cellobiose (G4G) as well as the β-1,3-linkage in laminaribiose (G3G), and the β-1,6-linkage in gentibiose (G6G) producing glucose. However, the digestion patterns indicated that PcGH3V23-R772 digested the β-1,3-linkage preferentially whereas cellobiose and gentibiose were still present after 25 min of digestion. Apart from the disaccharide digestion, product analysis revealed that hydrolysis of longer β-1,4-linked oligosaccharides such as cellotriose (G4G4G), cellotetraose (G4G4G4G), and cello-pentaose (G4G4G4G4G) released the monosaccharide glucose (Figure 3.4). Furthermore, the pres-ence of oligosaccharides with chain length n–1 in the chromatograms indicated that the enzyme cleaved the substrates in an exo-acting manner. Interestingly, when compared to the cellobiose digestion containing the same amount of PcGH3V23-R772, degradation of longer oligosaccharides by the enzyme occurred significantly faster. No long cellooligosaccharides were observed after 10 min of digestion indicating an efficient cleavage. Similarly, purified MLG oligosaccharides generated by PcGH5_4C21-E406 (Figure 3.5) were rapidly hydrolyzed into glucose and cellobiose. Thus, the β-glucosidase, PcGH3, hydrolyzes products of the MLG degradation by PcGH5_4C21-E406 and therefore complements the initial degradation step. However, the low activity on cellobiose suggests that an alternative pathway for cellobiose degradation might exist.49 Figure 3.4 Digestion of oligosaccharides by PcGH3V23-R772. (a) Digestion of 1 mM cellobiose (G4G), laminaribiose (G3G), gentibiose (G6G), and longer cel-looligosaccharides and (b) degradation of MLG-oligosaccharides at various concentrations by 0.008 mg/mL GH3 highlighted poor activity on G4G.a) b) 50 Figure 3.5 Mass spectra and HPLC chromatograms of purified MLG-oligosaccharides generated by PcGH5_4C21-E406. So far, the protein has not been crystallized and therefore the structural analysis relied on a prediction of the three-dimensional structure of PcGH3V23-R772 by AlphaFold [68]. A structural comparison between the model and the closest structural homologs provided further evidence for 51 the role of PcGH3 in the context of MLG degradation. A DALI search [70, 71] identified a GH3 member from a xyloglucan PUL in B. ovatus (BoGH3XyG; PDB-ID: 5JP0; [80]) as the closest structural homolog with over 60% sequence identity. The structure might contain three distinct domains after the N-terminal His6-tag: an (α/β)8-barrel domain stretching from residue 25 to 414, followed by an α/β-sandwich domain ranging from residue 415 to 653, and a C-terminal fibron-ectin type-III-like domain including residues 654 to 774 (Figure 3.6). Furthermore, a loop might point towards the active site after dimerization and therefore might be important for the function of PcGH3. Figure 3.6 AlphaFold model of the tertiary structure of the PcGH3V23-R772 [68]. (a) The predicted structure of PcGH3V23-R772 comprises an (α/β)8-barrel domain (green), an α/β-sandwich domain (violet), and the fibronectin type-III-like domain (orange). (b) To mimic the structure of the dimer, a second peptide chain was modeled revealing a loop pointing towards the active site of PcGH3. a) b) 52 Similar to other members of GH3, the active site might be a pocket located in the center of the (α/β)8-barrel but might comprise additional residues from loops of the neighboring α/β-sand-wich domain. The catalytic residues D309 and E527 might be conserved (Figure 3.6), may serve as nucleophile and acid/base respectively, and may be essential for hydrolysis via the classical Koshland double-displacement mechanism [80]. The overlay of the model of PcGH3V23-R772 with the crystal structure of BoGH3XyG in complex with a single glucose molecule revealed that other residues in the active site might be conserved and might interact with glucose units forming a -1 subsite (Figure 3.6, Figure 3.7). In addition, the superposition suggested that two tryptophan resi-dues, W310 and W453, might be conserved, narrow the entry to the active site pocket, and may stabilize carbohydrates in a +1 subsite via π-stacking interactions (Figure 3.7). This “coin-slot” conformation might also influence the substrate specificity of PcGH3V23-R772 by preventing α-gly-cosides such as pNP-α-glucose to enter the tight pocket. Figure 3.7 Predicted conformation of the active site of PcGH3V23-R772. (a) Superposition of glucose bound in the active site of BoGH3XyG (red; PDB-ID: 5JP0; [80]) with an AlphaFold model of PcGH3V23-R772 indicated a distinct -1 and +1 subsite. (b) The overlay of the active site residues of the modelled PcGH3V23-R772 (green and violet) with the b) a) 53 respective residues of the crystal structure of BoGH3XyG (cyan; PDB-ID: 5JP0; [80]) suggested structural conservation of the active site pocket. The biochemical data and the structural comparison suggested that PcGH3 cleaves single glucose molecules from the non-reducing end of oligosaccharides and hence acts as an exo-acting β-glucosidase. However, the enzyme displayed low activity on cellobiose which is consistent with observed activities of another GH3 member from the MLGUL of B. ovatus (BoGH3MLG; BA-COVA_02745; ID: 61%) that was 30 times less specific towards cellobiose than to cellotriose or cellotetraose and 20 times less specific towards laminaribiose [59]. Since the extracellular BoGH16 hydrolyzes MLG leaving a β-1,3-linkage between the reducing glucose unit and the fol-lowing unit (Figure 2.17), degradation by BoGH3MLG starting from the non-reducing end eventu-ally results in laminaribiose (G3G) [59]. PcGH5_4 however cleaves β-1,3-linkages preferentially leading to oligosaccharides that are degraded to cellobiose inevitably by PcGH3. Thus, questions remain regarding the alternative routes for cellobiose degradation. Figure 3.8 Hydrolysis of MLG-derived oligosaccharides by the exo-acting PcGH3 in comparison to BoGH3MLG. Cleavage sites are indicated in red. 54 3.2.2 A complementary path for cellobiose degradation via PcGH94 The transcriptome of P. copri growing in MLG-enriched media showed upregulation of the gene Prevcop_05090 that occurs isolated from other genes encoded by the MLGUL (Figure 1.5). Considering signal peptide analysis using SignalP [62] and Psipred [63], the protein does not contain a signal peptide and therefore is most likely located in the cytosol. Regarding a sequence comparison by BlastP [60] against the protein data bank, high sequence similarity to cellobiose phosphorylases from GH94 suggested a similar activity of Prevcop_05090 (Table 3.1). Despite all efforts, the cloned protein constructs of Prevcop_05090 (Figure 3.8) could not be expressed and purified in amounts required for biochemical or structural biological characteri-zation. Therefore, the role of the PcGH94 remains mostly unexplored and might be an interesting target for future investigations. Nevertheless, the high sequence similarity to various members of GH94 (Table 3.1) allowed the structural prediction by AlphaFold [68] and comparison of the model with existing protein crystal structures. Table 3.1 Sequence similarity of Prevcop_05090 towards crystallized members of GH94.Enzyme family PDB-ID Organism Activity/func-tion % ID Alignment length Mis-matches Ref. GH94 3QDE Acetivibrio ther-mocellus YM4 Cellobiose phos-phorylase 64 829 274 [81] GH94 2CQT Cellvibrio gilvus ATCC13127 Cellobiose phos-phorylase 61 840 304 [82] GH94 3RRS Cellulomonas uda DSM 20108 Cellobiose phos-phorylase 59 840 314 To be published 55 Figure 3.9 Structure of the cloned and expressed constructs of PcGH94. His6-tags of different vectors are indicated in different colors. The construct PcGH94K5-M831 excluded four amino acid residues of the native protein that might be disordered. 56 Figure 3.10 SDS-PAGE gels of the expression tests for the constructs of PcGH94. Expression tests in small vol-umes (a) and large volumes (b) for different constructs of PcGH94 did not show overexpression. The smeared band in the gel of PcGH94M1-M831 in pET28a resulted from overloading. Cells were cultivated in autoinduction media or Luria-Bertani broth (LB). For the latter, overexpression was initiated by adding isopropyl-D-thiogalactoside (IPTG). The MW of the bands of the protein ladder are provided in kDa on the left. a) b) 57 A DALI search [70, 71] revealed high similarity to two published cellobiose phosphorylase crystal structures (PDB-ID: 3QDE, 2CQT; [81, 82]). Therefore, the structure of PcGH94 might also dimerize and comprise four different domains: the N-terminal region (residues 1 – 282), a helical linker domain (residues 283 – 317), an (α/α)6-barrel domain (residues 324 – 757) and a region (residues 318 – 323 and 758 – 831) adopting a β-sandwich fold (Figure 3.10). Figure 3.11 Structural model of PcGH94 obtained by AlpaFold [68]. (a) The predicted structure of PcGH94 com-prises an N-terminal region (green), an (α/α)6-barrel domain (orange) connected via a helical linker (blue), and a C-terminal β-sandwich domain (yellow). The arrows in the box indicate the relative orientation of the protein in three-dimensional space. (b) The dimer of the predicted structure shows a loop containing a short α-helix that might narrow the entrance to the active site similar to a GH94 from Cellvibrio gilvus (grey; PDB-ID: 2CQT; [81, 82]). a) b) 58 The overlay with the crystal structure of GH94 from Acetivibrio thermocellus (AtGH94; PDB-ID: 3QDE; [81]) revealed two conserved subsites involved in substrate recognition and tran-sition state stabilization (Figure 3.10). The active site pocket may completely enclose the -1 subsite while subsite +1 may be situated at the entry of the cavity. In the model, the back wall created by W484 confines the space of the active site which might impede the binding of substrates with bulky substituents at C4 or C5 (Figure 3.11). Moreover, the tight conformation of the +1 subsite formed by the conserved residues might facilitate substrate discrimination via a distinct hydrogen bond network (Figure 3.11). Positioned right between subsites -1 and +1, the amino acid residue D486 might serve as catalytic acid during hydrolysis. The superposition of the modelled tertiary structure of PcGH94 with the crystal structure of AtGH94 revealed that the active site pocket may contain conserved residues (R347, H675, Q722, T741, T743) that might stabilize phosphate via hydrogen bond networks in the interior of the (α/α)6-barrel domain. Thus, the protein might com-pensate for the charges on the phosphate molecule and might decrease the distance between phos-phate and subsite -1 enabling a direct nucleophilic attack on the anomeric carbon of the glucose unit bound in the -1 subsite (Figure 3.11). However, the ambiguity of the position of amino acids such as Q502 between the predicted structure of PcGH94 and observed tertiary structures of other members of GH94 highlighted the limitations of modeling three-dimensional protein structures (Figure 3.11). Since proteins are not rigid, some amino acids may adopt manifold conformations, and therefore a profound understand-ing of the protein activity requires further structural analysis. 59 Figure 3.12 Predicted three-dimensional structure of the active site of PcGH94. (a) The overlay of the active site residues of PcGH94 (orange) with the residues of a crystal structure of AtGH94 (violet; PDB-ID: 3QDE; [81]) and cellobiose (cyan) from a complex with a GH94 from Cellulomonas uda (PDB-ID: 3S4A; to be published) suggest structural conservation of the active site. (b) The relative position of phosphate (PDB-ID: 3QDE; [81]) and cellobiose (PDB-ID: 3S4A; to be published) in the active site of GH94. The arrow indicates the proposed direction of the nucle-ophilic attack and the dotted line represents the interaction between the proton-donating amino acid D486 and the oxygen of the glycosidic bond. According to the sequence and structural model, Prevcop_05090 encodes a cellobiose phosphorylase that may generate α-D-glucose 1-phosphate and D-glucose from cellobiose without ATP involvement. This alternative digestion route is energetically efficient and might promote rapid growth of P. copri on carbohydrates containing β-1,4-linked glucose units. Interestingly, enzymes of GH94 have been predicted to occur in several Bacteroides species such as B. uniformis whereas others such as B. ovatus do not contain cellobiose phosphorylases and therefore exclu-sively rely on members of GH3 for the complete decomposition of MLG [52] highlighting the uniqueness of the MLGUL of P. copri. b) a) 60 3.3 Conclusion After extracellular degradation of β-1,3-linkages of MLG by PcGH5_4 and transport of the oligosaccharides into the periplasm, additional carbohydrate breakdown is required. Therefore, the MLGUL of P. copri incorporates an exo-acting glycoside hydrolase, PcGH3. Enzymes belonging to GH3 are commonly found in various PULs (Figure 1.3) and have a broad substrate specificity [79]. The enzyme localized in the periplasm dimerized in solution and hydrolyzed different β-glycosidic linkages in oligosaccharides. Indeed, the comparison of the predicted structure of PcGH3 with active sites of other members of GH3 revealed a binding pocket that might accom-modate a glucose unit in subsite -1. The narrow entry to the active site pocket might constitute subsite +1 and might be responsible for the specificity of the enzyme. Thus, PcGH3 acts as a classic exo-acting enzyme degrading oligosaccharides that are produced by PcGH5_4 and im-ported into the periplasm. However, the low activity on cellobiose suggests the involvement of additional enzymes in the complete degradation of MLG In comparison to other species such as B. ovatus, P. copri upregulated a protein that dis-plays high sequence similarity to characterized cellobiose phosphorylases and hence might play a unique role within the MLGUL. The structural model of PcGH94 indicated a conserved active site that may stabilize cellobiose. But in contrast to ordinary GHs, phosphorylases catalyze the nucle-ophilic attack of phosphate onto the carbohydrate. This process might point towards an energetic benefit for P. copri since glucose and glucose 1-phosphate are produced without an ATP. Further-more, the poor ability of GH3 enzymes to cleave cellobiose hints towards an essential involvement of PcGH94 in MLG utilization. 61 Chapter 4: Carbohydrate recognition by extracellular SGBPs 4.1 Introduction PULs include genes for various non-enzymatic proteins. Some of these additional proteins are localized on the outer cell membrane and show binding affinities to carbohydrates. These proteins termed SGBPs are essential components in canonical Bacteroidetes PULs and increase the local carbohydrate concentration at the cell surface. In this way, they guide distinct carbohydrates to-wards enzymes that hydrolyze the glycosidic bonds specifically [41]. Generally, two distinct types of SGBPs have been identified, SGBPs-A, which are SusD-hom-ologs and SGBPs-B. The SusD-homologs possess a globular structure and can interact with TBDTs (SusC-homologs) to facilitate carbohydrate transport [83, 84]. SGBPs-B, however, typi-cally contain several domains that are not conserved across PULs and primarily capture carbohy-drates on the cell surface [85]. This carbohydrate recognition occurs on binding platforms that in some cases allow promiscuous binding of complex carbohydrates. The following chapter evaluates the role that the two non-catalytic proteins encoded by the MLGUL, Prevcop_05097 and Prevcop_05098, play in the context of MLG utilization. Therefore, the binding affinities towards different carbohydrates and distinct features of both proteins ena-bling binding and discrimination of potential substrates were investigated. 62 4.2 Results and discussion 4.2.1 PcSGBP-A – an MLG-specific SGBP-A According to sequence analysis, the MLGUL of P. copri contains a gene, Prevcop_05098, that encodes a SusD-homolog, PcSGBP-A. The assessment of signal peptide boundaries by Sig-nalP [62] and Psipred [63] indicated that the native protein includes a 29 amino acid long, N-terminal signal peptide. In nature, signal peptidase II might cleave this peptide and a diacylglyceryl transferase might anchor the protein to the outer cell membrane. Therefore, this sequence was excluded from the constructs of PcSGBP-A (Figure 4.1). In addition, a disordered N-terminal re-gion was deleted in construct PcSGBP-AP50-K595 based on a structural model predicted by Al-phaFold [68]. However, most of this linker was included in construct PcSGBP-AD33-K595(+IGSG) which contained four additional amino acid residues, IGSG. Both constructs, PcSGBP-AP50-K595 (64.7 kDa) and PcSGBP-AD33-K595(+IGSG), were successfully cloned, expressed, and purified (Figure 4.2) and the N-terminal His6-tag of PcSGBP-AD33-K595(+IGSG) was cleaved during protein purifica-tion using TEV protease resulting in a protein with the molecular weight of 64.6 kDa. Figure 4.1 Structure of the cloned and expressed constructs of PcSGBP-A. 63 Figure 4.2 SDS-PAGE gels of the purified PcSGBP-A constructs, PcSGBP-AP50-K595 (64.7 kDa) and PcSGBP-AD33-K595(+IGSG) after TEV-cleavage (64.6 kDa). The MW of the bands of the protein ladder are indicated in kDa on the left. 4.2.1.1 Specific binding of MLG by PcSGBP-A Binding affinities of PcSGBP-AP50-K595 to various polysaccharides were assessed by affin-ity gel electrophoresis (Figure 4.3). Retardation of PcSGBP-AP50-K595 was observed in gels con-taining barley MLG, xyloglucan, glucomannan, and hydroxyethyl cellulose whereas gels contain-ing lentinan, laminarin, xylan, galactomannan, galactan, yeast β-glucan, and 1,4-β-mannan did not affect the migration of the protein. The results indicated that PcSGBP-AP50-K595 recognized poly-saccharides that contained backbones of β-1,4-linked glucose units (Figure 1.2) specifically and did not bind other β-glucans comprising a β-1,3-linked backbone and β-1,6-linked side branches. 64 Furthermore, polysaccharides without glucose molecules in their backbone such as xylan, galac-tomannan-, galactan-, and 1,4-β-mannan (Figure 1.2) did not interact with PcSGBP-AP50-K595. Nev-ertheless, binding affinity towards glucomannan, a polysaccharide that contains glucose and man-nose moieties in its backbone structure (Figure 1.2), suggested that the glucose units are the essen-tial feature for recognition. To further validate and quantify the observed interactions between PcSGBP-AP50-K595 and the four polysaccharides, barley MLG, xyloglucan, glucomannan, and hydroxyethyl cellulose, car-bohydrate binding was monitored using isothermal titration calorimetry (ITC) (Figure 4.4, Table 4.1). The ITC data revealed that the binding of PcSGBP-AP50-K595 to the 1,4-linked β-glucans was relatively weak. The interaction between PcSGBP-AP50-K595 and barley MLG showed the smallest dissociation constant and the largest energetic stabilization. The comparison of dissociation con-stants indicated that barley MLG was bound with nine times higher affinity than xyloglucan, ca. 20 times higher affinity than glucomannan, and more than 300 times higher affinity than hydrox-yethyl cellulose, a water-soluble cellulose derivative. However, no binding affinity was observed for the interaction with cellotetraose, one of the major products of degradation of barley MLG with PcGH5_4, suggesting that the PcSGBP-A only recognizes longer carbohydrates. The ITC analysis confirmed the observed binding affinities to 1,4-linked β-glucans and suggested that the binding preference towards barley MLG resulted from the kinked structure which might promote specific protein-carbohydrate interactions. Nevertheless, other structural dif-ferences between the analyzed carbohydrates might account for the drastic difference in the bind-ing affinity of xyloglucan, glucomannan, and hydroxyethyl cellulose. These structural intricacies such as xyloglucan side branches, mannose units in glucomannan and substitution of hydroxyl groups in the cellulose derivative might destabilize the interactions on the protein surface. 65 Figure 4.3 Binding affinities of MLGUL encoded SGBPs to polysaccharides. For affinity gel electrophoresis, acrylamide gels containing polysaccharides were loaded with PcSGBP-AP50-K595 (SGBP-A), PcSGBP-AP50-K595 after TEV-cleavage (SGBP-A_TEV), PcSGBP-BF42-E332 (SGBP-B), PcSGBP-BP136-E332 (SGBP-B_s), and bovine serum albumin (BSA), a non-interacting standard, used to compare migration behavior. Multiple bands in the BSA containing lane indicate dimerization of the protein. The concentration of polysaccharides in the gel is given in w/v and structures of the individual polysaccharides are provided in Figure 1.2. 66 Figure 4.4 Polysaccharide binding by PcSGBP-AP50-K595 and PcSGBP-BF42-E332 monitored by ITC. The upper graph displays the recorded heat changes fol-lowing each injection of polysaccharide (barley MLG, xyloglucan, glucomannan, and hydroxyethyl cellulose) or oligosaccharide (cellotetraose) into protein and the bottom graph shows the integrated heats. 67 Table 4.1 Thermodynamic parameters of the protein-carbohydrate interaction of the SGBPs encoded by the MLGUL in P. copri observed by ITC at 25 °C. Carbohydrate Kd (μM) ∆G (kJ/mol) ∆H (kJ/mol) T∆S (kJ/mol) n PcSGBP-AP50-K595 PcSGBP-BF42-E332 PcSGBP-AP50-K595 PcSGBP-BF42-E332 PcSGBP-AP50-K595 PcSGBP-BF42-E332 PcSGBP-AP50-K595 PcSGBP-BF42-E332 PcSGBP-AP50-K595 PcSGBP-BF42-E332 Barley MLGa 15 ± 4.2 210 ± 34 -28 -21 -17 ± 1.1 -23 ± 2.5 11 -2 1.0 1.0 Xyloglucanb 140 ± 24 270 ± 82 -22 -20 -15 ± 1.2 -100 ± 28 7.0 -80 1.0 1.0 Glucomannana 290 ± 28 630 ± 60 -20 -18 -22 ± 1.5 -120 ± 11 -2 -10 1.0 1.0 Hydroxyethyl cellulosea 4,700 ± 830 3,000 ± 490 -13 -14 -180 ± 28 -120 ± 16 -170 -110 1.0 1.0 Cellotetraose NDc - NDc - NDc - NDc - NDc - a Data was fitted assuming a 1:1 binding model to a pentamer (PcSGBP-A) or hexamer (PcSGBP-B) unit within the polysaccharide. The size of the binding site was estimated according to the crystal structure of PcSGBP-A, the predicted structure of PcSGBP-B, and previous studies on SGBPs [86, 87]. b Data was fitted assuming a 1:1 binding model to a xyloglucan octamer unit XXLGXXLG that approximates the amounts of glucose, xylose, and galactose in the used xyloglucan. The interaction was proposed according to previous studies on SGBPs [86, 87]. c ND, data not determinable.68 4.2.1.2 Structural features of PcSGBP-A without ligand and in complex with cellopen-taose Construct PcSGBP-AD33-K595(+IGSG) (Figure 4.1) crystallized in 0.1 M tris(hydroxyme-thyl)aminomethane (Tris)-HCl buffer (pH 8.5) containing 26% polyethylene glycol 8000 and was frozen using 30% ethylene glycol as cryoprotectant. After diffraction, the three-dimensional struc-ture of the protein was solved to a resolution of 2.05 Å by molecular replacement using an Al-phafold model [68] (Table A.2). The asymmetric unit of PcSGBP-AD33-K595(+IGSG) contains a single protein that consists of 570 amino acid residues and has a molecular weight of 64.6 kDa. The protein adopts a globular protein fold including the α-helix rich tetratricopeptide repeat motif (PFAM PF07980) (Figure 4.5) typical for SusD-like proteins [86]. Unfortunately, no electron density was observed in the crystal structure of PcSGBP-AD33-K595(+IGSG) for the first 18 amino acid residues constituting the disordered linker (Figure 4.1) and the four C-terminal residues, IGSG. Furthermore, the position of a ten amino acid-long loop starting at residue T267 could not be determined. 69 Figure 4.5 Three-dimensional crystal structure of PcSGBP-AD33-K595(+IGSG). (a) The structure reveals the typical SusD-like structure including four tetratricopeptide repeat motifs. The exact location of a ten amino acid long loop that could not be determined is indicated by a dashed line. (b) The overlay of the structure (blue) with the closest structural homolog, SGBP-A from B. ovatus (gray; BoSGBP-A; PDB-ID: 6DMF; [86]), shows the conservation of the α-helix rich region. To solve the complete structure of the PcSGBP-A, multiple crystallization screens were performed using over 300 buffers and several oligosaccharides. The construct PcSGBP-AD33-K595(+IGSG) crystallized in 0.1 M Tris-HCl buffer (pH 8.5) containing 28% polyethylene glycol 8000 and 5 mM cellotetraose and obtained crystals were diffracted using 30% ethylene glycol as cryo-protectant. Then, the tertiary structure was determined to a resolution of 1.5 Å using the previously solved structure as a template for molecular replacement (Table A.2). Similar to the previously obtained structure, a single protein occupies the asymmetric unit displaying a SusD-like overall conformation. However, the electron density indicates that a small oligosaccharide, cellopentaose, b) a) 70 is present generating a protein-carbohydrate complex (Figure 4.6). Cellopentaose resulting from an impurity in the cellotetraose solution appears to have crystallized locally with PcSGBP-AD33-K595(+IGSG) since multiple positions along the crystal were diffracted but only one of them contained the ligand in the protein crystal structure. Apart from the first 18 amino acid residues, the electron density of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose allows modeling of the whole protein structure. The position of the four C-terminal residues, IGSG, in the tertiary structure suggests that the residues are involved in interactions with proteins in neighboring asymmetric units and hence facilitate crystallization and diffraction (Figure 4.7). Moreover, the crystal structure of the complex of PcSGBP-AD33-K595(+IGSG) displays a stabilized configuration of the loop starting at residue T267 (Figure 4.5) which might interact with a protein of a neighboring asymmetric unit. Figure 4.6 Crystal structure of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose. (a) The tertiary structure of PcSGBP-AD33-K595(+IGSG) (green) in complex with cellopentaose (orange) confirms that the protein-carbohydrate a) b) 71 interactions mostly occur in a region characterized by loops and short α-helices. (b) The electron density suggests that the interaction with cellopentaose stabilizes the flexible loop that could not be modeled in the structure without ligand. Figure 4.7 Contact sites of two asymmetric units in the crystal structure of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose. The four additional amino acid residues, IGSG, at the C-terminus are highlighted in blue, a loop that might facilitate crystallization is indicated in red, and the cellopentaose ligand is colored in orange. 4.2.1.3 Carbohydrate binding interactions of PcSGBP-A The crystal structure of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose reveals a flat carbohydrate binding site formed by amino acid residues located on short α-helices and loop regions (Figure 4.6). Although the crystallization condition contained 5 mM cellotetraose, the elec-tron density suggests that cellopentaose is bound and therefore carbohydrate binding likely re-quires at least five glucose residues. Notably, all of the glucose units display the 4C1 conformation and are linked via β-1,4-glycosidic bonds. 72 Two exposed, aromatic residues, W86 and Y363, stack towards the third and fifth glucose unit of cellopentaose respectively (all glucose residues are numbered according to their position away from the reducing end) (Figure 4.8). Furthermore, the structure indicates hydrophobic inter-actions between D69 and the first glucose unit, between S293 and the third glucose, and between W362 which is located periplanar to Y363 and the fifth glucose unit. Apart from these hydrophobic interactions, hydrogen bond interactions play a significant role in stabilizing the carbohydrate on the protein surface. N87 interacts with the C3 hydroxyl group of the second glucose unit and R104 stabilizes the C5 oxygen atom of the reducing glucose residue (Figure 4.8). Interestingly, a loop containing Q289 and D290 adopts a different conformation in the protein-carbohydrate complex structure compared to the structure without ligand which enables Q289 to form a hydrogen bond with the C3 hydroxyl group of the second glucose unit and D290 with the hydroxyl group at C6 of the third glucose residue. In comparison to the tertiary structure without carbohydrate, the ligand-protein complex reveals a conformational change of the ten amino acid-long loop starting at T267 (Figure 4.6). An aromatic residue, Y269, located on the loop stacks to the glucose unit at the reducing end and forms a hydrogen bond to the C2 hydroxyl group of the second glucose unit (Figure 4.8). Similarly, loops in a BtSusD from B. theta rearrange upon binding of α-glucan oligosaccharides [88]. Thus, the flexible loop contributes to the general shape of the substrate binding site and might enhance interactions with longer carbohydrates. Interestingly, both crystal structures of PcSGBP-AD33-K595(+IGSG) display another exposed aromatic residue, W366, that is located close to the binding site but does not interact with cello-pentaose. The structures reveal that W366 is not positioned along the straight line of the binding 73 site formed by other residues including W86 and Y363 (Figure 4.8). In addition, amino acid resi-dues that might stabilize a glucose unit stacked to W366 are absent. Hence, the crystal structure of the complex of PcSGBP-AD33-K595(+IGSG) and cellopentaose was superposed with an MLG-nonamer from a complex with SGBP-A from the MLGUL of B. ovatus (BoSGBP-A; PDB-ID: 7NOX; [87]). In fact, W366 might interact with the seventh glucose unit via π-stacking (Figure 4.8). Further-more, a similar hydrogen bond pattern seems to stabilize the carbohydrate on the protein surface such as R104 which may form hydrogen bonds with the C1 and C2 hydroxyl groups of the first glucose unit and N87 that might interact with the C3 hydroxyl group at the second glucose unit which shows a similar conformation as in cellopentaose. Hence, the binding interactions and exact positions of interacting residues may result in a slightly altered binding configuration for MLG. The structural insights showed how PcSGBP-A binds β-1,4-linked carbohydrates. On the protein surface, a combination of π-stacking, van-der-Waals, and hydrogen bond interactions sta-bilizes the carbohydrate and might therefore minimize the impact of solvation and desolvation. Moreover, the protein adopts a specific conformation where a flexible loop stabilizes cellopentaose and therefore might accommodate β-1,4-linked polysaccharides. This flexible loop and the pres-ence of an additional aromatic residue, W366, adjacent to the binding site might therefore promote the recognition of the kinked polysaccharide structure of MLG and may account for the specific binding behavior regarding barley MLG. 74 Figure 4.8 Carbohydrate binding site of PcSGBP-AD33-K595(+IGSG). (a) Overlay of the binding site of the PcSGBP-AD33-K595(+IGSG) (green; blue) with corresponding residues of BoSGBP-A (pink; PDB-ID: 6DMF; [86]) shows how a) b) c) 75 cellopentaose (orange) interacted at the protein surface. (b) The superposition with an MLG-nonamer ligand from BoSGBP-A structure (yellow; PDB-ID: 7NOX; [87]) suggests possible interactions between W366 and the seventh glucose unit counted from the reducing end. (c) The relative orientation of the carbohydrates in the binding site is shown and indicated by solid lines. 4.2.1.4 Structural conservation of PcSGBP-A Previous studies indicated that variations within the carbohydrate-binding region affect substrate specificity [86]. Interestingly, BoSGBP-A from the MLGUL, the closest structural hom-olog of PcSGBP-AD33-K595(+IGSG) identified by a DALI search [70, 71] (Table 4.2), displayed similar binding behavior towards barley MLG and xyloglucan [86]. The comparison of the structure of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose with the complex of BoSGBP-A and cel-lohexaose revealed that the residues R72, W86, N87, R104, and D290 in the carbohydrate-binding site are conserved (Figure 4.8) whereas D69, Q289 and Y363 of PcSGBP-AD33-K595(+IGSG) are sub-stituted by E60, N280 and W350 in BoSGBP-A respectively. Tamura et al. showed that the mutants W77A and W350A of BoSGBP-A corresponding to W86 and Y363 in PcSGBP-AD33-K595(+IGSG) extinguished the ability to bind barley MLG whereas mutation of W353, corresponding to W366, to alanine just decreased the affinity to barley MLG [86]. Furthermore, the crystal structure of BoSGBP-A indicated that the reducing glucose unit of cellohexaose interacts with the residue Y266 located on a loop corresponding to the flexible loop in PcSGBP-AD33-K595(+IGSG). Notably, the mutant Y266A decreased binding affinity to barley MLG [86]. However, the structure of BoSGBP-A without ligand revealed that the loop adopts the same conformation as in the protein crystal structures of carbohydrate-protein complexes and therefore might indicate a rigid protein structure that stabilizes MLG better than pure β-1,4-linked glucans. 76 Thus, similar binding sites of PcSGBP-A and BoSGBP-A might result in a similar carbohydrate binding preference and might point towards a similar strategy of both species for substrate recog-nition at the cell surface. Table 4.2 Closest structural homologs of PcSGBP-AD33-K595(+IGSG) according to a DALI search [70, 71].Protein family PDB-ID Organism Function Z score r.m.s.d. % ID Ref. SGBP-A 6DMF Bacteroides ovatus ATCC 8483 SusD-like protein from MLGUL 51.2 1.5 45 [86] SGBP-A 5J5U Flavobacterium john-soniae UW101 Chitin-binding protein 45.2 1.8 35 [89] 4.2.1.5 Role of SGBP-A in MLG utilization In Bacteroidetes, SGBPs-A enhance the carbohydrate uptake into the periplasm via inter-actions with a corresponding TBDT (SusC-homolog) [41, 88]. In fact, the structural similarity of PcSGBP-AD33-K595(+IGSG) to other characterized SusD-like proteins (Table 4.2) might point towards a close assembly with the TBDT encoded by the MLGUL. Superposing the crystal structure of PcSGBP-AD33-K595(+IGSG) with the crystal structure of a SusC-SusD homolog pair from a predicted PUL of B. theta (BtSGBP-A; PDB-ID: 5FQ6; [90]) suggested that PcSGBP-AD33-K595(+IGSG) may be positioned on top of the transport protein with the binding site facing inwards (Figure 4.9). Similar to the BtSGBP-A, all four tetratricopeptide repeat motifs point to the outside and thus stabilization of the extracellular lid may occur via hydrogen bond networks. In the overlay, clashes between the proteins result from the difference in the variable binding region of the PcSGBP-AD33-77 K595(+IGSG). Nevertheless, the distinct arrangement suggested that PcSGBP-AD33-K595(+IGSG) might enable oligosaccharide import via the “pedal-bin” mechanism proposed for SusC-SusD homolog pairs. Without ligand, SGBP-A hinges away from the SusC-like protein allowing substrate recog-nition. Then, the protein lid closes and the oligosaccharide is transported through the SusC-hom-olog [83, 90]. Interestingly, the data revealed that PcSGBP-A undergoes a structural change and recognizes oligosaccharides containing five glucose units. Therefore, smaller digestion products of MLG generated by PcGH5_4 such as cellotetraose may not be transported whereas longer oli-gosaccharides might be captured and imported via the complementary interaction of PcSGBP-A and the SusC-homologous TBDT directly after the degradation of MLG by PcGH5_4. This con-certed interaction of the extracellular proteins might allow the rapid import of oligosaccharides into the periplasm and therefore might play a significant role in MLG utilization at the cell surface. 78 Figure 4.9 Structural model for the SusC-SusD interaction. (a) The crystal structure of PcSGBP-AD33-K595(+IGSG) (green) in complex with cellopentaose (orange) was superposed with the three-dimensional structure of a SusC-SusD homolog pair from B. theta (PDB-ID: 5FQ6; [90]). The BtSusC-homolog is colored in blue and BtSGBP-A in light orange. The structural similarity of both SGBPs-A suggested that the SusC-SusD complex of P. copri might show similar interactions. (b) The surface representation of the SusC-homolog of B. theta (blue; PDB-ID: 5FQ6; [8]) aligned to PcSGBP-AD33-K595(+IGSG) (green) in complex with cellopentaose showed how carbohydrate ligands might be encap-sulated and captured by the protein complex. a) b) 79 4.2.2 PcSGBP-B – an auxiliary binding protein 4.2.2.1 Promiscuous binding of 1,4-β-glucans In addition to the non-catalytic binding protein, PcSGBP-A, the MLGUL of P. copri com-prises a gene, Prevcop_05097, that was highly upregulated. Sequence analysis by BlastP [60] in-dicated 37.3% sequence similarity of Prevcop_05097 to a characterized SGBP-B from the MLGUL of B. ovatus (BoSGBP-B) suggesting that Prevcop_05097 encodes an SGBP-B as well. According to signal peptide predictions by SignalP [62] and Psipred [63], PcSGBP-B contains a 34 amino acid-long signal peptide that might be cleaved by signal peptidase II and might allow anchoring of the protein to the outer cell membrane at C35. Hence, this sequence was deleted from the protein constructs. Tertiary structure prediction by AphaFold [68] suggested that a disordered region links the protein consisting of three modules to the lipid molecule (Figure 4.10) and there-fore this linker was excluded from the protein constructs as well. Two constructs, PcSGBP-BF42-E332 (34.4 kDa) and PcSGBP-BP136-E332 (24.0 kDa) without the N-terminal domain, were success-fully cloned, expressed, and purified (Figure 4.11). At first, binding affinities of the protein to various polysaccharides were elucidated using native affinity gel electrophoresis (Figure 4.3). Similar to PcSGBP-A, PcSGBP-BF42-E332 interacted with 1,4-linked β-glucans including barley MLG, xyloglucan, glucomannan, and hydroxyethyl cellulose but did not recognize 1,3-linked β-glucans. Furthermore, the gels containing xylan, ga-lactan, galactomannan, and β-1,4-linked mannan showed no retention of the protein. Thus, the data indicated that PcSGBP-B binds 1,4-linked β-glucans specifically. 80 Figure 4.10 Structure of the cloned and expressed constructs of PcSGBP-B. Figure 4.11 SDS-PAGE gels of the purified PcSGBP-B constructs, PcSGBP-BF42-E332 (34.4 kDa) and PcSGBP-BP136-E332 (24.0 kDa). The MW of the bands of the protein ladder are indicated in kDa on the left. 81 ITC analyses underscored the observed affinities of PcSGBP-B to 1,4-linked β-glucans (Figure 4.4, Table 4.1). The binding interaction towards barley MLG was the strongest while hy-droxyethyl cellulose displayed the weakest binding. However, dissociation constants of barley MLG, xyloglucan, and glucomannan were not significantly different highlighting that PcSGBP-B binds 1,4-linked β-glucans promiscuously. This intriguing result suggests that PcSGBP-B serves as a primary sensor that increases the local 1,4-linked β-glucan concentration on the cell surface. 4.2.2.2 Potential function and role of PcSGBP-B To identify structural features responsible for the promiscuous binding behavior, both con-structs, PcSGBP-BF42-E332 and PcSGBP-BP136-E332, were crystallized in various buffers. Crystals of PcSGBP-BF42-E332 were found in more than 60 conditions and PcSGBP-BP136-E332 crystallized in more than 30 conditions (Figure 4.12). Unfortunately, the structure could not be determined by molecular replacement since all of the crystals did not diffract well. Thus, the structural analysis of PcSGBP-B relied on a model obtained from AlphaFold [68] (Figure 4.13). Figure 4.12 Selected crystallization conditions for the constructs, PcSGBP-BF42-E332 and PcSGBP-BP136-E332. 82 Figure 4.13 Model of the structure of PcSGBP-B. (a) The predicted model generated by AlphaFold [68] suggests a multimodular three-dimensional structure of PcSGBP-B composed of three individual domains (blue, green, orange). (b) Superposition of the predicted structure of the C-terminal domain of PcSGBP-B with the C-terminal domain of BoSGBP-B (gray; PDB-ID: 6E57; [86]) reveals a binding platform comprising three structurally conserved aromatic residues. However, the relatively low sequence similarity suggests that the protein might adopt an alternative structure. (c) The overlays of the modeled C-terminal domain of PcSGBP-B with cellopentaose (blue) and an MLG-heptamer (green) from complexes with BoSGBP-B (PDB-ID: 6E57, 6E9B; [86]) indicated that all of the three aromatic residues might interact with the carbohydrate. a) b) c) 83 The predicted structure of PcSGBP-B revealed three modules connected via loops. How-ever, only module C displays coherent and exposed aromatic residues. Indeed, an overlay of the C-terminal domain of PcSGBP-B with a corresponding domain of the closest structural homolog, BoSGBP-B, revealed three exposed tryptophan residues, W238, W275, and W276, that might form a potential binding platform for carbohydrate recognition (Figure 4.13). Carbohydrate stabilization would occur through π-stacking interactions between the flat binding site and the carbohydrate. Thus, the flat binding site might accommodate structural intricacies such as the kinked structure of MLG or the mannose containing backbone of glucomannan and therefore might lead to the promiscuous binding behavior. Furthermore, side branches of xyloglucan might increase the con-tact site towards the binding platform which do not interfere with those sterically challenging car-bohydrate branches. In contrast to the more closely related SGBPs-A, PcSGBP-B showed only low sequence similarity to BoSGBP-B underlining the enormous diversity among those PUL-associated binding proteins. Nevertheless, the observed binding affinities and the structural prediction suggested that PcSGBP-B is a carbohydrate-binding protein involved in MLG utilization. While previous studies indicated that the presence of an SGBP-A is vital for the fitness of species of Bacteroidetes [86], SGBPs-B may serve more as auxiliaries and might associate with the SusD/SusC-homolog com-plex forming a utilisome as recently described by White et al. [55]. The vicinity and potential interaction of the different proteins involved in this multi-protein complex might be a key factor in facilitating the specific MLG utilization at the cell surface. 84 4.3 Conclusion The MLGUL of P. copri encodes two distinct carbohydrate-binding proteins, PcSGBP-A (Prevcop_05098) and PcSGBP-B (Prevcop_05097). Both proteins might be anchored to the outer cell membrane and bind glycans that contain β-1,4-linked glucose units in their backbone specifi-cally. PcSGBP-A is a specific MLG binding protein whereas PcSGBP-B binds the 1,4-linked β-glucans promiscuously. The obtained crystal structure of PcSGBP-AD33-K595(+IGSG) in complex with cellopentaose shows a typical SusD-like structure comprising a carbohydrate binding site. The region contains conserved residues enabling substrate recognition via hydrophobic interactions and hydrogen bonds. Interestingly, the ligand cellopentaose stabilizes a loop containing the interacting residue Y269. In the structure without ligand, the loop is too flexible to be modeled revealing how the protein structure accommodates the ligand. Apart from that, the structural similarity to known SGBPs-A suggested an association of PcSGBP-A with the TBDT encoded by the MLGUL. Thus, carbohydrate import might occur via the proposed “pedal-bin” mechanism where the SGBP-A or the “lid” opens up enabling substrate recognition. Upon binding, conformational rearrangements might result in a “lid”-closing followed by carbohydrate transport. In comparison to SGBPs-A, the structural prediction of PcSGBP-B revealed a multi-mod-ular protein architecture. The modeled C-terminal domain of PcSGBP-B contains three aromatic residues generating a potential carbohydrate binding platform resembling the C-terminal domain of BoSGBP-B. Therefore, PcSGBP-B might serve as an auxiliary binding protein in the MLGUL recognizing 1,4-linked β-glucans at the cell surface. Thus, both SGBPs may enhance the rapid and efficient carbohydrate utilization on the outside of the cell and thus might result in the outstanding performance of P. copri on MLG. 85 Chapter 5: Conclusion and future perspectives 5.1 Summary of the results In this work, I performed an in-depth biochemical and structural biological characterization of the MLGUL of P. copri. By investigating the three enzymes encoded by the MLGUL, namely PcGH5_4, PcGH3, and PcGH94, along with two non-catalytic proteins, PcSGBP-A and PcSGBP-B, I was able to elucidate a model for the molecular mechanisms underlying the degradation of the complex carbohydrate MLG (Figure 5.1). On the cell surface, the non-catalytic binding protein, PcSGBP-B, captures 1,4-linked β-glucans promiscuously via a potential binding platform. This binding event increases the local concentration of potential substrates. Then, the endo-acting PcGH5_4 hydrolyzes MLG specifically via the classic Koshland double-displacement mechanism typical for members of this GH family. The enzyme prefers cleavage of β-1,3-linkages connecting two blocks of β-1,4-linked glucose units since the β-1,4-linked glucan motif might interact with the conserved negative subsites and positioning of the kink of the MLG structure close to the catalytic residues might allow tight binding in the positive subsites. The generated longer MLG-oligosaccharides are recognized by the PcSGBP-A through a binding site constituted by aromatic residues and a flexible loop. These oligosaccharides are then imported into the periplasm via the concerted action of the PcSGBP-A/TBDT complex. Close proximity between the extracellular proteins as in observed utilisomes might accelerate the rapid breakdown and subsequent transport. In the periplasm, the exo-acting β-glucanase, PcGH3, degrades the oligosaccharides producing glucose and cellobiose. Notably, the enzyme shows only low activity on cellobiose and therefore a predicted cellobiose phosphorylase, PcGH94, is upregulated and colocalized to the other genes generating glucose and glucose 1-phosphate. 86 Figure 5.1 Cellular model for the MLG uptake encoded by the MLGUL of P. copri. PcSGBPs interact with MLG at the cell surface. PcGH5_4 hydrolyzes the carbohydrate into oligosaccharides that can be imported through the PcSGBP-A/TBDT complex. In the periplasm, PcGH3 degrades the oligosaccharides into mono- and disaccharides and PcGH94 cleaves cellobiose after the transport into the cytosol. The genetic location and sequence similarity sug-gested that the HTCS served as a transcriptional regulator for the MLGUL recognizing specific breakdown products. 5.2 Future experiments To deepen the understanding of the specific carbohydrate-protein interactions, the compar-ative structural analysis of PcGH5_4 requires further research. The next step will include data collection at the Canadian Light Source for the obtained crystal complexes of the catalytic mutants of PcGH5_4 with G4G3G4G, an MLG-hexamer, and MLG-heptamer (Figure 2.13). Furthermore, 87 GH5_4-specific inhibitors including N-bromoacetylglycosylamines and 1-fluoro-2-deoxyglyco-sides will reveal the distinct binding conformation of the protein. Especially, the less conserved positive subsites are of interest because they might determine the preference for hydrolysis of β-1,3-linkages. Similarly, the catalytic activity of the mutant W248A that can be cloned will show if this residue contributes significantly to MLG degradation. To elucidate the ecological impact of degrading MLG via a member of GH5_4 or a GH16, growth studies should be performed comparing the ability of P. copri and B. ovatus to utilize cellotriose and cellotetraose. A major growth difference will support the hypothesis of an ad-vantage of P. copri in MLG digestion and might explain the abundance of P. copri in people con-suming carbohydrate-rich diets. Today, only one close structural homolog has been identified for PcGH3. Therefore, crys-tallization might advance the understanding of the whole enzyme family. Despite our efforts, prob-lems in the expression and purification of the PcGH94M1-M831 constructs occurred. Recloning the protein into a vector containing a C-terminal His6-tag might solve these issues and allow biochem-ical characterization and crystallization. Regarding PcSGBP-A, the individual contribution of the aromatic residues W78, Y267, Y355, and W358 for the carbohydrate binding can be obtained by site directed mutagenesis. Sim-ilarly, an alanine scan of the three tryptophan residues constituting the binding platform of PcSGBP-B and a crystal structure of the C-terminal domain of PcSGBP-B might help to determine the structure-function relationship of MLG binding. 88 5.3 Outlook Overall, my comprehensive characterization provides valuable insights into the intricate machinery of MLG degradation that evolved in P. copri and highlights the potential of the organ-ism to degrade complex plant-carbohydrates rapidly. Thus, this data might be the starting point for the investigation of other PULs within P. copri. Studying these PULs holds significant importance since we obtain insights into the diversity of the carbohydrate breakdown and utilization mecha-nisms which are vital for survival and proliferation in a highly competitive environment. Thus, the studies might indicate adaptations of P. copri to specific environmental pressures and might ex-plain the fitness of P. copri within the HGM. Furthermore, by understanding the mechanism of carbohydrate utilization, future studies might identify and analyze specific interspecies interactions and might define the role of P. copri within the ecosystem where competition for resources let generalists and specialists evolve. Hence, the characterized MLGUL might help to explain the abundance of P. copri in people consuming non-Western/plant-carbohydrate-based diets. The increasing knowledge of the metabolism in P. copri helps pinning down the specific host-microbe interactions and their effects on human health. Distinct PUL repertoires might indi-cate specific dietary preferences of distinct microorganism. Therefore, comparative studies on commensal and pathogenic species might be able to elucidate their role in nutrient acquisition, colonization, and virulence. Thus, this work contributes to the identification of growth stimulants of P. copri which might be a major factor in modulating the HGM of people suffering under con-ditions caused by metabolic diseases, such as diabetes, inflammatory bowel disease, and colorectal cancer. This may result in novel low-cost, low-risk, non-invasive therapeutic treatments of these intricate and partially elusive diseases. 89 Finally, the detailed analysis of the MLGUL might have implications for biotechnological applications. 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Cloning and sequencing Genes, Prevcop_05090 (PcGH94), Prevcop_05096 (PcGH3), Prevcop_05097 (PcSGBP-B), Prevcop_05098 (PcSGBP-A), and Prevcop_05101 (PcGH5_4), were analyzed for signal pep-tides using SignalP [62] and Psipred [63] and for disordered regions using structural models cre-ated by AlphaFold [68]. Signal peptides and N-terminal, disordered linkers were excluded from several constructs of PcSGBP-A, PcSGBP-B, PcGH5_4, and PcGH3. Primers were designed ac-cordingly (Table A.1) and the genes were amplified from gDNA of P. copri DSM 18205 via PCR using Q5® high-fidelity polymerase (New England Biolabs®). Constructs were cloned into pMCSG53 vectors through ligation independent cloning (LIC) [91] and transformed into E. coli DH5α-cells. After cultivation in Luria-Bertani broth (LB), plasmids were extracted, purified, and validated via colony PCR and Sanger sequencing (Genewiz®). Then, plasmids were transformed into E. coli BL21 (DE3) cells. In a similar way, PcGH94M1-M831 was cloned into pET28a using the restriction enzymes NdeI and high fidelity HindIII as well as T4 DNA ligase (New England Bi-olabs®). PcGH5_4 active site mutants, PcGH5_4T41-E406(E197A) and PcGH5_4T41-E406(E331A), were generated using the Q5® Site-Directed Mutagenesis Kit (New England Biolabs®) and the plasmid of the clone PcGH5_4T41-E406. 101 Table A.1 List of the used protein constructs, primers, and vectors. Genetag Protein construct Forward primer sequence (5’→3’)a Reverse primer sequence (5’→3’)a Vector Prevcop_05090 PcGH94M1-M831 TACTTCCAATCCAATGCAATGGCAGA AAAAAAGTATG TTATCCACTTCCAATGTTATTACAT CTCAACCACTAC pMCSG53 PcGH94K5-M831 TACTTCCAATCCAATGCAAAGTATGG TCATTTTGATG TTATCCACTTCCAATGTTATTACAT CTCAACCACTAC PcGH94M1-M831 GACCATATGGCAGAAAAAAAGTATGG TCATTTTGATGACG GACAAGCTTTTACATCTCAACCACT ACTTCATGCTTGCC pET28a Prevcop_05096 PcGH3V23-R772 TACTTCCAATCCAATGCAGTTATCAAG AGCGACCCTAAGATTGAGG TTATCCACTTCCAATGTTATTATCG GTTGGCAGTAGACCACTGATAAG pMCSG53 Prevcop_05097 PcSGBP-BF42-E332 TACTTCCAATCCAATGCATTCTTCACC GTATCAGAAGATGACGATCC TTATCCACTTCCAATGTTATTATTC AATAGTTACTTTCTTTACGGT GTAAC CATCAC pMCSG53 PcSGBP-BP136-E332 TACTTCCAATCCAATGCACCAGCCCTG GGTACCAATGCC TTATCCACTTCCAATGTTATTATTCA ATAGTTACTTTCTTTACGGTGTAACCA TCAC pMCSG53 Prevcop_05098 PcSGBP-AP50-K595 TACTTCCAATCCAATGCACCTAATGCG CAGTTGACAACATCTTTG TTATCCACTTCCAATGTTATTACTTA ATGTAGCCCTTGCCGTAAGG pMCSG53 PcSGBP-AD33-K595(+IGSG) TACTTCCAATCCAATGCCGATATGATG GAGCG TTATCCACTTCCAATCTTAATGTAGC CCTTGCCG pMCSG53 Prevcop_05101 PcGH5_4C21-E406 TACTTCCAATCCAATGCATGCGAA-GCAAG TTATCCACTTCCAATGTTATTATTCTT TCCCAAAGATTC pMCSG53 PcGH5_4T41-E406 TACTTCCAATCCAATGCAACTGCCCAG TTATCCACTTCCAATGTTATTATTCTT TCCCAAAGATTC pMCSG53 PcGH5_4T41-E406(E197A) CGGTACCAATgcaGTTCACATCAGGG GCGAAAGCCAAGCGACTG pMCSG53 PcGH5_4T41-E406(E331A) CGTAATAGGAgcaTGGGGAGTAACC ATACCCAGTCCTTTGTTG pMCSG53 a Restriction sites and complementary sequences for ligation independent cloning with vector pMCSG53 are indicated in bold. Mutated nucleotides are written in small caps.102 A.1.2. Expression and purification Cells were cultivated in LB at 37 °C and overexpression was initiated at an optical density of 0.6 (OD600) via the addition of isopropyl-D-thiogalactoside (IPTG) to a final concentration of 1 mM. Then, the solution was stirred at 200 rpm at 16 °C for 48 h. Cells containing the plasmids for PcGH3 and PcGH94 were also cultivated in autoinduction media at 37 °C to an optical density of 0.6 and incubated afterwards at 16 °C for 48 h. After centrifugation at 4367 g for 30 min, cell pellets were resuspended in binding buffer (20 mM sodium phosphate (NaPi); pH 7.4; 500 mM NaCl; 20 mM imidazole) and stored at – 20 °C. Frozen cell pellets were thawed and the cells were lyzed afterwards using a Sonic Dismem-brator F55 Ultrasonic Homogenizer (Fisher Scientific). After centrifugation at 4367 g for 45 min, the supernatant was loaded onto a 5 mL Ni2+ HisTrapTM HP column (Cytiva) and purified using an NGCTM chromatography system (Biorad). The column was washed with 40 mL of binding buffer and then eluted using gradient elution. The concentration of elution buffer (20 mM NaPi; pH = 7.4; 500 mM NaCl; 500 mM imidazole) changed from 0 to 100% within 100 mL. As soon as the protein started to elute, the respective gradient was held. Fractions were collected and purity was confirmed by SDS-PAGE. Pure fractions were pooled and concentrated using a VivaspinTM centrifugal filter (Cytiva). For the cleavage of the His6-tag of PcSGBP-AD33-K595(+IGSG), the protein solution containing binding and elution buffer was exchanged after the first immobilized metal chelate affinity chro-matography (IMAC) with cleavage buffer (20 mM NaPi; pH 7.4; 500 mM NaCl) using a Vi-vaspinTM centrifugal filter (Cytiva). To the concentrated protein solution, 400 μg of tobacco etch virus (TEV) protease were added and the solution was incubated overnight at 4 °C. On the next 103 day, PcSGBP-AD33-K595(+IGSG) was separated from the cleaved His6-tag by a second IMAC using cleavage buffer. Purity of protein fractions was confirmed by SDS-PAGE and the pure fractions were combined and concentrated. After IMAC, concentrated protein solutions were transferred into an Eppendorf tube and centrifuged for 10 min at 21100 g. The supernatant was injected into a 1 mL loop attached to the NGCTM chromatography system (Biorad). Then, the protein was purified by size-exclusion chro-matography (SEC) using Superdex 75 prep grade or Superdex 200 prep grade resin (Cytvia) and storage buffer (20 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES); pH 7.0; 100 mM NaCl). Purity of selected fractions was examined by SDS-PAGE. Then, pure fractions were combined and concentrated. The protein concentration was determined with a Cary 60 UV-Vis spectrophotometer (Ag-ilent Technologies) at λ = 280 nm. Protein yields usually were around 65 mg per L of culture ex-cept for PcGH3 and PcGH94. PcGH3V23-R772 expressed in lower yields. Finally, protein aliquots were flash frozen in liquied nitrogen and stored at – 70 °C. A.2 Biochemical characterization A.2.1. Substrates All used polysaccharides (barley MLG, carboxymethyl cellulose, curdlan, potato galactan, carob galactomannan, konjac glucomannan, hydroxyethyl cellulose, lentinan, laminarin from Laminaria digitata, β-1,4-linked mannan, pustulan, xanthan gum, xylan, xyloglucan, yeast β-glu-can), oligosaccharides (cellobiose, cellotriose, cellotetraose, cellopentaose, laminaribiose, lami-naritriose, laminaritetraose, gentibiose, 31-β-D-cellobiosyl-glucose (G4G3G), 32-β-D-glucosyl-cellobiose (G3G4G), 31-β-D-cellotriosyl-glucose (G4G4G3G), 32-β-D-cellobiosyl-cellobiose 104 (G4G3G4G), 33-β-D-glucosyl-cellotriose (G3G4G4G)), and para-nitrophenol (pNP) glycosides (pNP-α-glucose, pNP-β-glucose, pNP-α-galactose, pNP-β-galactose, pNP-β-mannose, pNP-α-xy-lose, and pNP-β-xylose) were purchased from Megazyme International (Bray, Ireland), Acros Or-ganics (Morris Plains, NJ, USA), Amresco (Solon, OH, USA), and Sigma Aldrich (St. Louis, MO, USA). Longer MLG-oligosaccharides were produced according to the protocol described in Ap-pendix A.3. A.2.2. Bicinchoninic acid assay Enzyme activity on polysaccharides was determined by the bicinchoninic acid (BCA) re-ducing sugar assay [92]. The reaction volume was set to 100 μL and a solution of carbohydrate and buffer was equilibrated at the desired reaction temperature (Appendix A.2.4.2). Then, the re-action was started by the addition of 10 μL enzyme solution composed of PcGH5_4C21-E406 at the desired concentration, buffer, and 0.1 mg/mL bovine serum albumin (BSA). To terminate the re-action after 6 min, 100 μL of BCA solution were added to the reaction mixture. Then, the solution was heated to 80 °C for 20 min and 100 μL of solution were transferred to a 96-well plate after-wards. The absorbance was measured at λ = 562 nm with an Epoch Microplate Spectrophotometer (Agilent BioTek). All reactions were performed in the linear range of the enzyme. The amounts of generated carbohydrate were calculated using a glucose standard curve (10 – 150 μM). Subtraction of absorbance values for blanks mitigated the effect of buffer or sub-strate on the observed absorbance. For blanks, 10 μL of buffer containing 0.1 mg/mL BSA were injected instead of the enzyme solution. The characteristic kcat and Km values were obtained by fitting the data to the Michaelis-Menten model in OriginPro 2019 Version 9.6.0.172. 105 A.2.2.1. Initial substrate screen The hydrolytic activity on various polysaccharides at 37 °C was assessed by incubating 10 μL of 5 mg/mL polysaccharide (barley MLG, xyloglucan, glucomannan, hydroxyethyl cellu-lose, carboxymethylcellulose, xylan, curdlan, laminarin from Laminaria digitata, yeast β-glucan, and xanthan gum) or 2.5 mg/mL polysaccharide (pustulan) with 80 μL 50 mM NaPi buffer (pH 6.0) and 10 μL PcGH5_4C21-E406 (0.0001 – 0.1 mg/mL). A.2.2.2. pH- and temperature profile To determine optimal reaction conditions for PcGH5_4, reactions were performed in so-dium citrate buffer (NaCit; pH 3.0 – 6.0), 2-(N-morpholino)ethanesulfonic acid buffer (MES; pH 5.5 – 6.0), NaPi buffer (pH 6.0 – 8.0), glycine sodium hydroxide buffer (Gly-OH; pH 9.0 – 10) or at temperatures ranging from 20 – 70 °C. 10 μL of 5 mg/mL barley MLG were added to 80 μL 50 mM buffer and 10 μL of 0.0001 mg/mL PcGH5_4C21-E406 were injected. A.2.2.3. Michaelis-Menten kinetics For the determination of kcat and Km, all reactions were performed in 40 mM NaCit buffer at pH 4.5. 90 μL of polysaccharide solution (0 – 1.2 mg/mL) were equilibrated at 50 °C and the reaction was initiated by the addition of 10 μL enzyme solution containing 0.0001 – 0 .1 mg/mL PcGH5_4C21-E406. A.2.2.4. Specific activity of catalytic mutants To compare the hydrolytic activity of PcGH5_4T41-E406, PcGH5_4T41-E406(E197A), and PcGH5_4T41-E406(E331A) on barley MLG, 10 μL of polysaccharide (5 mg/mL) were incubated with 10 μL enzyme (0.0001 – 0.2 mg/mL) at 37 °C in 40 mM NaCit buffer (pH 4.5). 106 A.2.3. para-Nitrophenol (pNP) substrate assay The chromogenic pNP substrate assay was employed to monitor the activity of exo-acting enzymes such as PcGH3. All experiments were performed at 37 °C in an end-point assay setup with a fixed reaction volume of 100 μL containing 1 mM of pNP-glycoside. 10 μL of PcGH3V23-R772 (0.08 mg/mL) were added to initiate the reaction which was stopped after 10 min by the addi-tion of 100 μL of 1 M Na2CO3 solution. 100 μL of solution were transferred and the absorbance of released pNP was measured on an Epoch Microplate Spectrophotometer (Agilent BioTek) at λ = 405 nm. The substrate screening was performed in 40 mM NaPi buffer (pH 6.2) and the pH profile was determined using 40 mM NaCit, NaPi, and glycine-hydroxide (Gly-OH) buffers at dif-ferent pHs. A.2.4. Carbohydrate analysis by HPAEC-PAD High-performance anionic-exchange chromatography with pulsed amperometric detection (HPAEC-PAD) was used to identify and quantify carbohydrates that were products of enzymatic degradation. The carbohydrates were separated on a 3 × 250 mm Dionex ™ CarboPac™ PA200 IC column with a 3 × 50 mm guard column using a Dionex ICS-6000 system. This system was coupled to an AS-AP autosampler with a temperature-controlled sample tray configured for se-quential injection. The system was operated through Chromeleon 7 control software and obtained chromatograms were processed afterwards using the same software and OriginPro 2019 Version 9.6.0.172. 10 μL of the sample were injected into the column and separated using three gradients. Solvents used for carbohydrate separation were ddH2O (solvent A), 1.0 M NaOH (solvent B), and 1.0 M sodium acetate (NaOAc; solvent C). The three optimized gradients were designed for different resolution ranges and were as follows: 107 Gradient I: 0 – 15 min; 10% B, 1 – 30% C linear gradient; 15 - 15.1 min; 10 – 50% B, 30 – 50% C linear gradients; 15.1 – 16 min, 50 – 10% B, 50 – 1% C exponential gradients; 16 – 20.1 min, 10% B, 1% C; Gradient II: 0 – 20 min; 10% B, 1 – 8% C linear gradient; 20 – 20.1 min; 10 – 50% B, 8 – 50% C linear gradients; 20.1 – 21 min, 50 – 10% B, 50 – 1% C exponential gradients; 21 – 25.1 min, 10% B, 1% C; Gradient III: 0 – 14 min; 10% B, 1 – 5.9% C linear gradient; 14 – 14.1 min; 10 – 50% B, 5.9 – 50% C linear gradients; 14.1 – 15 min, 50 – 10% B, 50 – 1% C exponential gradients; 15 – 20 min, 10% B, 1% C. A.2.4.1. Polysaccharide digestion profile The digestion profile of barley MLG by PcGH5_4C21-E406 was determined by HPAEC-PAD. 800 μL 10 mg/mL of polysaccharide and 100 μL 400 mM NaCit (pH 4.5) were incubated with 100 μL 0.002 mg/mL PcGH5_4C21-E406 in enzyme buffer containing 40 mM NaCit buffer (pH 4.5) and 0.1 mg/mL BSA at 37 °C. At desired time intervals, 100 μL of the reaction solution were transferred into 100 μL heated NaCit buffer (40 mM, pH 4.5) and the solution was heated to 90 °C for 10 min. 20 μL of this solution were diluted to 100 μL in ddH2O, the solution was filtered using a Costar® Spin-X® centrifuge tube filter containing a 0.22 μm pore CA membrane, and loaded on the column. Gradient I was used to monitor the production of longer and shorter oligo-saccharides whereas gradient II was used to separate shorter oligosaccharides with the same MW. The generated oligosaccharides were quantified using a standard curve (1 – 175 μM). Glucose, cellobiose, cellotriose, and cellotetraose were diluted to the desired concentration in 0.84 mM NaCit buffer (pH 4.5) and separated using gradient II. 108 A.2.4.2. Oligosaccharide limit-digestion Limit digestion of different oligosaccharides (G4G, G4G4G, G4G4G4G, G4G4G4G4G, G3G, G3G3G, G3G3G3G, G3G4G, G4G3G, G3G4G4G, G4G3G4G, and G4G4G3G) by PcGH5_4 was monitored by HPAEC-PAD. The reaction was initiated by mixing 10 μL PcGH5_4C21-E406 (0.2 mg/mL) with 90 μL of 100 μM oligosaccharide in 40 mM NaCit buffer (pH 4.5). The mixture was incubated at 37°C for 30 min and the reaction was stopped by heating the solution at 95 °C for 10 min. Thereafter, 10 μL of 1 mM D-mannitol were added as internal stand-ard, the solution was filtered (Costar® Spin-X® centrifuge tube filter) and separated by HPAEC-PAD using gradient III. A.2.4.3. Michaelis-Menten kinetics For Michaelis-Menten kinetics, reactions were performed in 40 mM NaCit buffer (pH 4.5) at 37 °C. 90 μL of oligosaccharide (1 μM – 10 mM) were incubated with 10 μL of PcGH5_4C21-E406 in enzyme buffer at the required concentration and the reaction was stopped after 30 min by heating the solution to 95 °C for 10 min. Then, 10 μL of 1 mM D-mannitol were added as internal standard, the solution was filtered (Costar® Spin-X® centrifuge tube filter) and gradient III was used to separate the oligosaccharides on the column. Carbohydrate amounts were calculated from standard curves for cellobiose or cellotriose (0.5 – 100 μM) diluted in 40 mM NaCit buffer (pH 4.5). Characteristic kcat and Km values for the digestions were determined from a fit using the Michaelis-Menten model in OriginPro 2019 Version 9.6.0.172. A.2.4.4. Oligosaccharide digestion profile To monitor the degradation of the oligosaccharides (G4G, G4G4G, G4G4G4G, G4G4G4G4G, G3G, and G6G) by PcGH3V23-R772, hydrolysis was performed on 1 mM oligosac-charide in 40 mM NaPi buffer (pH 6.0) at 37 °C. The total reaction volume was set to 600 μL and 109 the reaction was started by adding 60 μL of 0.08 mg/mL PcGH3V23-R772 into the solution. After selected time intervals, aliquots of 100 μL were heated to 95 °C for 10 min. Then, 10 μL of 1 mM D-mannitol, the internal standard, were added, the solution was filtered (Costar® Spin-X® centri-fuge tube filter) and degradation products were separated and detected by HPAEC-PAD using gradient III. Degradation of MLG-oligosaccharide was performed following a similar protocol. A final con-centration of 180 μM MLG6 and 99 μM MLG7 was digested and the resulting product separated via HPAEC-PAD using gradient I. A.2.5. Native oligosaccharide hydrolysis To determine the regiospecificity of PcGH5_4, cellopentaose hydrolysis was performed in [18O] water (97%; Cambridge Isotope Laboratories) as described by McGregor et al. [74]. 0.5 μL PcGH5_4C21-E406 (0.02 mg/mL), 0.5 μL 400 mM NaCit buffer (pH 4.5), and 11 μL [18O] water were mixed and put on ice. As soon as 0.5 μL 10 mM cellopentaose were added, the solution was heated to 37 °C. After 10 min, the reaction was terminated by boiling the solution at 95 °C for 10 min. Controls were performed using 11 μL [16O] water. The incorporation of [18O] was detected via mass spectrometry using a Waters® 2695 Separation Module connected to a Waters® Micro-mass ZQ Detector. A.2.6. Thermal shift assay To elucidate the stability of PcGH3 in various buffers, a thermal shift assay was performed using Multiplate 96-Well PCR Plates. The reaction volume was set to 25 μL comprising 4 μM of protein and 10 × SYPROTM Orange protein gel stain (Invitrogen). The PcGH3V23-R772 sample was diluted in different buffers and analyzed using a 7500 Fast Real-Time PCR System (Applied Bio-systemsTM). To observe temperature-dependent fluorescence, samples were heated according to 110 the following profile: 10 °C for 5 min; 10 °C for 1 min; heating to 95 °C with a 1% increase per minute; cooling to 10 °C for 10 min. Obtained data sets were processed and plotted using OriginPro 2019 Version 9.6.0.172. A.2.7. Affinity gel electrophoresis The polysaccharides (barley MLG, xyloglucan, glucomannan, hydroxyethyl cellulose, ga-lactan, galactomannan, lentinan, laminarin, and xylan) were dissolved in ddH2O at 60 °C to give a concentration of 10 mg/mL. However, 100 mg of yeast β-glucan, curdlan and β-1,4-linked man-nan were dissolved in 10 mL 10% NaOH. The solution was neutralized to pH 7 with 100% acetic acid and diluted to concentrations of 5 mg/mL for yeast β-glucan and β-1,4-linked mannan, and 4 mg/mL curdlan. 7.5 mL of gelling solution (1 × running buffer, 1 mg/mL polysaccharide, 10% acrylamide/ bisacrylamide) were prepared using the polysaccharide solutions, 5 × running buffer (250 mM Tris-base, 125 mM boric acid, pH 8.7), and 30% acrylamide/bisacrylamide solution (ratio 37:5.1) (Bio-Rad). 5 μL tetramethyl ethylenediamine (TEMED) and 37.5 μL ammonium persulfate (APS) were added to start the polymerization reaction. 4.5 mL of the solution were transferred into glass frames and covered with 1 mL 1-propanol. After 30 min, the 1-propanol was removed and 1.5 mL stacking gel poured on the polysaccharide gels. For the stacking gel, 1.75 mL ddH2O, 0.5 mL 5 × running buffer, and 0.25 mL 30% acrylamide/bisacrylamide solution were mixed, and the polymerization was initiated by adding 5 μL TEMED and 37.5 μL APS. 6 μL of protein (2 mg/mL) were diluted in 54 μL ddH2O and mixed with 15 μL of loading dye (272 mM Tris-base, 121 mM boric acid, 1.5 mM bromphenol blue, 50% glycerol, pH 8.7). 10 μL of that solution were loaded on the gel and gel electrophoresis was performed at 100 V for 180 min. Then, protein bands were stained with Coomassie Brilliant Blue stain. 111 A.2.8. Isothermal titration calorimetry Isothermal titration calorimetry was performed with a MicroCal PEAQ-ITC (Malvern Pan-alytical) at 25 °C. Proteins were diluted in buffer containing 20 mM HEPES, 100 mM NaCl, at pH 7.0, to concentrations of 80 – 150 μM. Polysaccharide solutions at concentrations of 2.5 – 5 mg/mL or 1 mM cellotetraose were titrated against protein. Preceded by a 0.2 μL injection, 2 μL of carbohydrate solution were injected 18 times every 150 s and stirred at 750 rpm. Recorded heat changes were analyzed using MicroCal PEAQ-ITC Analysis Software Version 1.21 (Malvern Panalytical). Binding affinities were calculated assuming a 1:1 binding model. A.3 Oligosaccharide production and purification To obtain MLG-oligosaccharides with more than five glucose units, 900 μL of 400 mM NaCit buffer pH 4.5, 100 μL of ddH2O, and 8 mL of 10 mg/mL barley MLG were equilibrated at 37 °C. The degradation of MLG started after the addition of 1 mL of 0.002 mg/mL PcGH5_4C21-E406 in enzyme buffer (40 mM NaCit; pH 4.5; 0.1 mg/mL BSA) and was terminated after 10 min by heating the solution to 90 °C for 10 min. 30 mL ice-cold ethanol were added and the solution was stored at – 20 °C overnight. The solution was centrifuged at 4367 g for 1 h and the superna-tant was transferred to a round bottom flask. Solvents were evaporated using a Rotovapor® RII (BUCHI). The oligosaccharides were dissolved in 100 μL ddH2O and separated on a size-exclu-sion column using Bio-Gel P2 resin (Biorad). Fractions were collected and oligosaccharide dis-tribution was determined by mixing 50 μL of solution with 50 μL of BCA solution. After heating to 80 °C, the absorbance was measured at λ = 562 nm. The purity of fractions was verified by HPAEC-PAD using gradient I and gradient II. Pure fractions were combined, lyophilized and dissolved in ddH2O. Then, the oligosaccharides were diluted to a concentration of 100 μM and 112 analyzed on an Agilent 1100/1200/1260/1290 LC Q-TOF system using a high resolution ESI-method without a column (HRESI_+_Accuarcy_ms_no column.m). Amounts of purified oligosaccharide were determined after the degradation of 2.5 μL of oligosac-charide with 0.08 mg/mL PcGH3V23-R772 in a 100 μL reaction. The digestion was performed for 4 h in 40 mM NaPi (pH 6.0) at 37 °C. Generated glucose was quantified via HPAEC-PAD using gradient I and a glucose standard curve (5 – 200 μM). A.4 Crystallization, X-ray diffraction data collection and data processing Crystallization conditions were screened using solutions from IndexTM, Crystal ScreenTM, PEG/Ion ScreenTM and Additive ScreenTM (Hampton Research). For screening, sitting drops were plated by a mosquito® LCP robot (SPT Labtech) while hanging drops were prepared by hand in volumes of 2 or 10 μL for condition optimization. Conditions showing crystal growth were altered using varying concentrations of polyethylene glycol. Crystals were frozen in liquid nitrogen using different cryoprotectants including ethylene glycol, polyethylene glycol and isopropanol. The crystals were diffracted at the Stanford Synchrotron Radiation Lightsource [93, 94] and the Canadian Light Source [95] and the collected data sets were autoprocessed on the basis of XDS [96]. Molecular replacement and refinement were performed in house using Phenix [97] and CCP4 [98, 99]. Solved structures were validated in Coot [100] and the R-factor and the Rfree were calculated [99]. The crystallographic parameters are shown in Table A.2. 113 Table A.2 Crystallographic data and refinement statistics table PcGH5_4T41-E406 PcSGBP-AD33-K595(+IGSG) PcSGBP-AD33-K595(+IGSG) cellopentaose Data collection Beamline 12-2 (SSRL) 12-2 (SSRL) CMCF-ID (CLS) Wavelength (λ) 0.979460 0.979460 0.953720 Resolution range (Å) 38.43 - 1.6 (1.657 - 1.6) 33.56 - 2.05 (2.123 - 2.05) 42.03 - 1.5 (1.554 - 1.5) Space group P1211 P212121 P212121 Unit cell a, b, c (Å) 97.008 51.56 157.057 68.877 76.849 99.918 68.96 77.41 100.09 α, β, γ (°) 90 101.501 90 90 90 90 90 90 90 Total reflections 1,380,408 (136,175) 177,899 (17,254) 1,110,954 (116,427) Unique reflections 196,882 (19,223) 33,783 (3,340) 86,193 (8,470) Redundancy 7.0 (7.1) 5.3 (5.2) 12.9 (13.7) Completeness (%) 97.79 (96.03) 99.31 (99.70) 99.84 (99.88) I/σI 17.80 (2.29) 9.01 (2.41) 18.38 (4.81) Wilson B-factor (Å2) 18.13 27.54 R-meas 0.06272 (0.9094) 0.1338 (0.7698) 0.1068 (1.102) CC (1/2) 0.999 (0.819) 0.999 (0.946) 0.999 (0.914) Molecules in AU 4 1 1 Refinement R-work 0.1880 (0.2525) 0.1857 (0.2659) 0.1518 (0.1767) R-free 0.2135 (0.2975) 0.2248 (0.3107) 0.1658 (0.1912) Number of non-hydrogen atoms All 12951 4528 5078 Macromolecules 12216 4317 4516 Ligands 20 2 63 Water 715 209 499 RMS deviations Bond lengths (Å) 0.006 0.007 0.006 Bond angles (°) 0.87 0.88 0.960 Ramachandran statistics Favored (%) 97.47 96.44 97.45 Allowed (%) 2.33 3.38 2.55 Outliers (%) 0.21 0.19 0 Average B-factor (Å2) All 22.10 30.26 19.93 Macromolecules 21.62 30.23 18.95 Carbohydrate ligands 34.09 50.21 30.36 Solvent 30.05 30.73 27.44
Thesis/Dissertation
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Chemistry
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Brumer, Harry
Functional characterization of a mixed-linkage beta-glucan utilization locus from the human gut microbe Prevotella copri
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