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Engineering and Production of Glucooligosaccharide
Oxidases for Site-specific Activation of Cellulose and
Hemicellulose Substrates
by
Maryam Foumani Alhaeri
A thesis submitted in conformity with the requirements
for the degree of Doctor of Philosophy
Graduate Department of Chemical Engineering and Applied Chemistry
University of Toronto
© Copyright by Maryam Foumani Alhaeri 2015
ii
Engineering and Production of Glucooligosaccharide Oxidases for
Site-specific Activation of Cellulose and Hemicellulose Substrates
Maryam Foumani Alhaeri Doctor of Philosophy
Department of Chemical Engineering and Applied Chemistry
University of Toronto
2015
Abstract
Canada has an extensive supply of residual biomass, which comprises plant
polysaccharides that represent a renewable resource for the production of biochemicals,
polymers and fuels. Enzymatic oxidation of plant oligo- and poly-saccharides can alter
the characteristics of these compounds, enhancing their application in food products;
enzymatic oxidation could also facilitate site-specific chemical derivatizations of
carbohydrates, leading to new bio-based polymers.
A glucooligosaccharide oxidase from Sarocladium strictum (GOOX) with reported
activity on oligosaccharides comprising up to seven glucose units, was engineered in this
study using a genetic engineering approach to extend the activity of this enzyme on a
wider range of plant polysaccharides and oligosaccharides. For the first time, in addition
to the previously reported cello- and malto-oligosaccharides, activity of wild type GOOX
on xylooligosaccharides was reported. The catalytic efficiency of two mutant enzymes
(GOOX-Y300A, GOOX-Y300N) on cellooligosaccharides and xylooligosaccharides was
2-fold higher than the wild-type enzyme. Notably, the binding affinity of these mutants
iii
towards oligosaccharides had decreased, which was correlated to reduced substrate
inhibition by cellooligosaccharides. The GOOX-W351F mutant showed enhanced
activity and affinity on galactose compared to wild-type GOOX, suggesting that this
mutation reduces steric hindrance between galactose and substrate binding amino acids
within the substrate binding cleft of the enzyme.
Wild type GOOX exhibited low activity on polysaccharides including konjac
glucomannan, and barley β-glucan, and weak activities on carboxy-methyl cellulose,
regenerated amorphous cellulose, microcrystalline cellulose (Avicel), and xyloglucan.
The specific activity was improved by up to 56 %, 55 % and 30 % for crystalline
cellulose (Avicel), regenerated amorphous cellulose (RAC) and glucomannan,
respectively, by constructing fusions between GOOX and various carbohydrate binding
modules (CBMs). Binding capacities of the fusion proteins on crystalline and amorphous
cellulose, as well as glucomannan, β-glucan, and xyloglucan also increased by over 10-
fold as determined by SDS-polyacrylamide gel electrophoresis and affinity gel
electrophoresis. The immobilized fusion enzyme on a solid cellulose surface remained
stable and active. This finding is anticipated to broaden applications of GOOX as an
immobilized enzyme used in cellulose-based biosensing devices.
In addition to enzyme engineering and biochemical characterizations, three applications
of GOOX enzymes were evaluated: 1) production of oxidized oligosaccharides as sugar
standards, 2) synthesis of plant oligosaccharides and polysaccharides with enhanced
nutraceutical value, and 3) substitution of glucose oxidase used in baking applications.
iv
Acknowledgments
I would like to express my deepest and sincere gratitude to Prof. Emma R. Master for her
kind and firm supervision, motivations for taking unpredictable science routes, and her
continuous supports all through this exciting journey. Her dedication to provide an
energetic environment for students to explore and discover new findings and enthusiasm
towards research never cease to amaze me.
I am also very grateful to all the members of the Master lab for being excellent labmates,
making my graduate study a rewarding and enjoyable experience. In particular, Dr. Thu
Vuong for his kind assistance and guidance toward my research, and Dr. Dragica
Jeremic, Julie-Anne Gandier, Ruoyu Yan, and Mabel Wong for being great friends and
emotionally supportive throughout my study. I am also thankful to all the BioZone
members especially Endang Susilawati (Susie), Melanie Duhamel and Angelika Duffy
for their supports.
I am very thankful to my committee members Professor Bradley Saville and Professor
Ning Yan for their valuable feedback and insightful inputs to my research throughout the
course of my PhD.
I would like to extend my heartfelt gratitude to my parents, Nasrin Moeini and Mostafa
Foumani for their inspiration, support and unconditional love throughout my life, and to
my husband, Mohammad Reza, for his gracious patience, considerate guidance, and
never-ending kindness and support. I am especially thankful to my 5-year old daughter,
Parnian, who showed an exceptional independence towards the end of my work while
caring for her little sister, Parimah, in my absence at the family times. I am also grateful
for my siblings, Fatemeh, Javad, Mohammad, and Mahdi for being the kindest and most
supportive siblings I could ever know. Finally, I am thankful to my wonderfull friends
and neigbours, especially Dr. Fatemeh Akbarian, Lili Zahedi, Hoda Mofidi and Zahra
Choolaei for being by my side through the difficult times and energizing those moments.
v
Table of Contents
Table of Contents ............................................................................. v
List of Tables .................................................................................. ix
List of Figures .................................................................................. x
List of Appendices ......................................................................... xii
List of Abbreviations .................................................................... xiii
Chapter 1 : Overview ....................................................................... 1
Chapter 2 : Literature review ........................................................... 3 2.1 Oxidation of sugars and polysaccharides ...............................................................................3 2.2 Examples of oxidoreductases from selected Auxiliary Activity (AA) families .....................4 2.3 Oligosaccharide oxidases within AA7 ...................................................................................8
2.3.1 Oligosaccharide oxidase from Microdochium nivale, MnCO .........................................8 2.3.2 Cello-oligosaccharide oxidase from Paraconiothyrium sp., PCOX ................................9 2.3.3 Chitooligosaccharide oxidase from Fusarium graminearum, ChitO ............................10 2.3.4 Gluco-oligosaccharide oxidases from Sarocladium strictum T1, GOOX-T1 ...............10
2.4 Current strategies for enzyme engineering ...........................................................................13 2.4.1 Mutagenesis: Random versus rational design................................................................13 2.4.2 Gene shuffling ...............................................................................................................15 2.4.3 Fusion proteins ..............................................................................................................15
2.5 Carbohydrate binding modules .............................................................................................16 2.5.1 Contributions in cellulose/hemicellulose active enzymes .............................................17 2.5.2 Industrial applications of CBMs ....................................................................................18 2.5.3 Effects of CBMs on neighbouring modules ..................................................................18 2.5.4 Important factors for designing CBM fusion proteins ...................................................20
2.6 Applications of carbohydrate oxidase in food industry ........................................................21 2.6.1 Oxidoreductases in baking applications ........................................................................21 2.6.2 Enzyme utilization for production of prebiotics ............................................................24
2.7 Research Hypotheses and Specific Research Objectives .....................................................26
vi
Chapter 3 Improved GOOX activity on mono and oligosaccharides
using site-directed mutagenesis ..................................................... 29 3.1 Introduction ..........................................................................................................................30 3.2 Materials and Methods .........................................................................................................32
3.2.1 Fungal strain and materials ............................................................................................32 3.2.2 Cloning of the GOOX-encoding gene ...........................................................................32 3.2.3 Site-directed mutagenesis ..............................................................................................33 3.2.4 Recombinant protein expression ...................................................................................34 3.2.5 Enzyme purification ......................................................................................................34 3.2.6 Enzymatic assays and kinetics analyses ........................................................................35 3.2.7 Deglycosylation .............................................................................................................36 3.2.8 Substrate docking ..........................................................................................................36 3.2.8 Nucleotide sequence accession number.........................................................................37
3.3 Results and Discussion .........................................................................................................37 3.3.1 Variations of GOOX ......................................................................................................37 3.3.2 Production of recombinant protein ................................................................................39 3.3.3 Novel substrate specificity.............................................................................................40 3.3.4 Improvement of substrate specificity ............................................................................45
3.4 Conclusions ..........................................................................................................................47
Chapter 4 Enhanced binding and activity of GOOX towards
polysaccharides through CBM fusions .......................................... 50 4.1 Introduction ..........................................................................................................................51 4.2 Materials and Methods .........................................................................................................53
4.2.1 Materials ........................................................................................................................53 4.2.2 Construction of fusion enzymes ....................................................................................53 4.2.3 Recombinant expression of fusion proteins in Pichia pastoris .....................................54 4.2.4 Purification of recombinant enzymes ............................................................................55 4.2.5 Specific activity on oligosaccharides, soluble polysaccharides and insoluble cellulose
substrates ................................................................................................................................56 4.2.6 Cellulose binding ...........................................................................................................57 4.2.7 Quartz crystal microbalance with dissipation (QCM-D) ...............................................58 4.2.8 Affinity gel electrophoresis ...........................................................................................59
vii
4.2.9 Temperature stability .....................................................................................................59 4.2.10 Nucleotide sequence accession number.......................................................................60
4.3 Results and Discussion .........................................................................................................60 4.3.1 Recombinant protein production ...................................................................................60 4.3.2 Improved binding to polymeric substrates ....................................................................63 4.3.4 Specific activity on polymeric substrates ......................................................................67 4.3.5 Immobilization of GOOX through CtCBM3 .................................................................70 4.3.6 Effect of CBM on thermostability .................................................................................72
4.4 Conclusions ..........................................................................................................................74
Chapter 5 Application trials of wild-type and engineered GOOX 75 5.1 GOOX in the production of sugar standards ........................................................................76
5.2.1 Introduction ...................................................................................................................76 5.2.2 Materials and Methods ..................................................................................................77
5.2.2.1 NMR analysis of oxidized products ..................................................................................... 77 5.2.2.2 Mass spectrometric analysis of oxidized products ............................................................... 78
5.2.3 Results and Discussion ..................................................................................................79 5.2.3.1 Confirming the regioselectivity of gluco-oligosaccharide oxidases ..................................... 79 5.2.3.2 Efficient oxidation of cellooligos. and Impact of chain length on GOOX activity .............. 82
5.2 Application of GOOX CBM fusions in the synthesis of plant oligosaccharides with
enhanced nutraceutical value ......................................................................................................84 5.2.1 Introduction ...................................................................................................................84 5.2.2 Materials and Methods ..................................................................................................86
5.2.2.1 Oxidation of xylooligosaccharides ....................................................................................... 86 5.2.2.2 Prebiotic Assay ..................................................................................................................... 87
5.2.3 Results and Discussion ..................................................................................................88 5.2.3.1 Small-scale fermentation on xylooligosaccharides and aldouronic acid .............................. 88 5.2.3.2 Cultivation of B. longum on oxidized and non-oxidized xylooligosaccharides ................... 89
5.2.4 Conclusions ...................................................................................................................91 5.3 A Mutant gluco-oligosaccharide oxidase is suitable to replace glucose oxidase for baking
applications .................................................................................................................................92 5.3.1 Introduction ...................................................................................................................92 5.3.2 Materials and Methods ..................................................................................................96
5.3.2.1 Activity assays ...................................................................................................................... 96 5.3.2.2 Oxidation Reactions for H2O2 inactivation study ................................................................. 97
viii
5.3.2.3 Detection of gluconic acid by HPLC .................................................................................... 97 5.3.2.4 Monitoring the oxygen content using Oxygraph .................................................................. 98
5.3.3 Results and Discussion ..................................................................................................98 5.3.3.1 GOOX-Y300A shows higher oxidation of oligosaccharides ................................................ 98 5.3.3.2 H2O2 inactivation .................................................................................................................. 99
5.3.4 Conclusions .................................................................................................................102
Chapter 6 : Conclusions ............................................................... 103
Chapter 7 : Future directions ....................................................... 109 7.1 Incorportaion of GOOX-oxidized oligosaccharides in an LPMO standard assay. ............109 7.2 Effect of debranching enzyme on prebiotic activity of polysaccharides ............................109 7.3 Additional value of a CBM fusion GOOX-Y300A for baking application ........................110
References .................................................................................... 112
Appendix 1: Supplemental information for chapter 3 ................. 126
Appendix 2: Supplemental information for chapter 4 ................. 128
Appendix 3: Supplemental information for chapter 5 ................. 131
ix
List of Tables
Table 2.1 Brief reviews of common enzyme engineering strategies ..............................................16
Table 3.1 List of oligonucleotide primers used for gene amplification and site-directed
mutagenesis. ...........................................................................................................................33
Table 3.2 Amino acid substitutions in GOOX in comparison with GOOX-T1 .............................38
Table 3.3 The effect of deglycosylation with PNGaseF on enzyme activity .................................40
Table 3.4 Kinetics parameters of wild-type and mutant GOOX enzymes. ....................................41
Table 3.5 Docking parameters of oligosaccharides with GOOX enzymes. ...................................44
Table 4.1 Specific activity of wild-type and CBM fusion GOOX on oligosaccharides. ................62
Table 4.2 Binding of wild-type GOOX and CBM fusions to insoluble cellulose. .........................63
Table 4.3 Kinetics parameters of wild-type GOOX and its CBM fusions on cellotetraose. ..........66
Table 4.4 Specific activity of the wild-type GOOX and CBM fusions on polysaccharides. .........67
Table 4.5 The half life of fusion and wild-type GOOX at 45°C. ...................................................73
Table 5.1 Growth rate of B. longum cultivations. ..........................................................................90
Table 5.2 Specific activities of GOOX-Y300A and GO on selected mono and oligosaccharides. 99
Table 5.3 Amount of Gluconic acid produced by GOOX-Y300A or GO in the presence of various
concentrations of H2O2. ........................................................................................................101
x
List of Figures
Figure 2.1 Oxidation mechanism of GOOX ...................................................................................11
Figure 2.2 GOOX substrate binding and active sites .....................................................................12
Figure 3.1 The structural model of GOOX.....................................................................................31
Figure 3.2 Conformational changes of S388 upon substrate binding .............................................43
Figure 3.3 Docking of monosaccharides to GOOX .......................................................................47
Figure 3.4 The biding site for GOOX-T1. ......................................................................................49
Figure 4.1 Schematic representation of wild-type GOOX and GOOX fusions..............................62
Figure 4.2 Affinity gel electrophoresis (AGE) of wild-type GOOX and CBM fusions .................64
Figure 4.3 Specific activity of wild-type GOOX and CBM fusions on polysaccharides.. .............69
Figure 4.4 Frequency - dissipation plot of enzymes binding to cellulose ......................................71
Figure 5.1 NMR spectra of cellobiose and xylobiose oxidation ....................................................80
Figure 5.2 MS/MS spectra and fragmentation of GOOX oxidized cellotriose ..............................81
Figure 5.3 Positive ion ESI-MS spectra of four cello-oligosaccharide samples ............................83
Figure 5.4 Structure of compounds used in prebiotic assay ...........................................................85
Figure 5.5 Viable cell count of B. longum preliminary cultures ....................................................88
Figure 5.6 Viable cell count of B. longum cultures ........................................................................90
Figure 5.7 Prosed mechanisms for GOOX benefits in baking applications. ..................................95
Figure 5.8 Proposed mechanism for GOOX reinforcing the protein network in dough ................95
Figure 5.9 H2O2 tolerance of GOOX-Y300A and GO .................................................................101
Figure S3.1 Multiple sequence alignment of GOOX homologues…………………………….. 126
Figure S3.2 Stability of wild-type and mutant GOOX at 37°C………………………………....127
Figure S3.3 SDS-PAGE of deglycosylated GOOX-VN and mutant enzymes………………….127
Figure S4.1 Purified wild-type and fusion GOOX proteins on 10% SDS-PAGE……………....128
Figure S4.2 Binding of wild-type GOOX and CBM fusions to insoluble cellulose…….........…128
xi
Figure S4.3 Specific activity of wild-type GOOX and CBM fusions on konjac glucomannan...129
Figure S4.4 Adsorbed mass of GOOX and CtCBM3_GOOX on cellulose-coated sensors ……129
Figure S4.5 Cellobiose oxidation of enzyme-bound sensors……………………………………130
Figure S4.6 Thermostability of proteins at 45°C………………………………………………..130
Figure S5.1 Behaviour of GOOX-Y300A in the presence or absence of H2O2………….….......132
Figure S5.2 - Log of GluA concentration produced versus H2O2 concentration………………..132
xii
List of Appendices
Appendix 1: Supplemental information for chapter 3 Appendix 2: Supplemental information for chapter 4 Appendix 3: Supplemental information for chapter 5
xiii
List of Abbreviations
AA7/9- Carbohydrate active enzymes with Auxiliary Activity from family 7 or 9 BMGY- Buffered complex medium containing glycerol BMMH- Buffered minimal medium containing methanol and histidine BSA- Bovine serum albumin CAZy- Carbohydrate active enzyme database CBM- Carbohydrate binding module CDH- Cellobiose dehydrogenase CelK- Clostridium thermocellum cellobiohydrolase ChitO- Chitooligosaccharide oxidase from Fusarium graminearum CMC- Carboxymethyl cellulose DP- Degree of polymerization FAD- Flavin adenine dinucleotide GH61- Glycosyl hydrolase from family 61 (re-named to AA9) GluM- Glucomannan from Konjac GO- Glucose oxidase from Aspergillus niger HPAEC- High-performance anion-exchange chromatography HRP- Horseradish peroxidase GOOX- Glucooligosaccharide oxidase from Sarocladium strictum strain CBS 346.70 GOOX-T1- Glucooligosaccharide oxidase from Sarocladium strictum strain T1 LPMO- Lytic polysaccharide monooxygenase MnCO- Carbohydrate oxidase from Microdochium nivale MRS- Bacterial growth medium so-named by its inventors: de Man, Rogosa and Sharpe NAG- N-acetyl-glucosamine NMR- Nuclear magnetic resonance PCOX- Cello-oligosaccharide oxidase from Paraconiothyrium sp. PDB- protein database of The Research Collaboratory for Structural Bioinformatics PI- Prebiotic index POX- Pyranose oxidase QCM-D- Quartz crystal microbalance with dissipation RAC- Regenerated amorphous cellulose SDS-PAGE- Sodium dodecyl sulfate polyacrylamide gel electrophoresis TCAG- Center for Applied Genomics TEMPO- 2,2,6,6-tetramethylpiperidine-1-oxyl XOS- Xylooligosaccharide FAEXynZ- Feruloyl esterase domain of a xylanase from Clostridium thermocellum YNB- Yeast nitrogen base without amino acids 4-AA- 4-aminoantipyrine
1
Chapter 1 : Overview
Production of biochemicals and functionalized fibres for value-added bio-products from
biomass can help to offset the price of biofuel while providing sustainable, and
environmentally friendly replacements for petroleum based materials. Enzymatic
modification of the plant oligo- and polysaccharides of biomass can alter the
characteristics of these materials to meet the requirements for the downstream
applications (e.g. reactivity, solubility, compatibility with other biopolymers). By using
an enzymatic approach, it is also possible to catalyze regio-selective and stereo-specific
modifications, while retaining the degree of polymerization and/or crystallinity of the
substrate. Moreover, routine molecular biology techniques can be applied to fine-tune
enzyme activities to broaden substrate range and increase enzyme stability. Lastly, since
enzyme reactions are typically performed in aqueous solutions at intermediate pH and
temperatures below 80°C, they represent a class of “green” catalysts with minimal safety
considerations.
Following a summary of my scholarly contributions below, Chapter 2 will review the
main literature and concepts pertinent to my PhD thesis, and will end by stating my
specific research hypotheses and objectives.
Summary of Scholarly Contributions
Peer-reviewed Publications
2
Foumani M, Vuong TV, and Master ER. Altered substrate specificity of the gluco-
oligosaccharide oxidase from Acremonium strictum, Biotechnology and Bioengineering.
2011, 108(10): 2261-2269.
Foumani M, Vuong TV, and Master ER. Oligosaccharide oxidase derived from
Acremonium strictum and uses thereof, Patent WO 2012/116431 A1, 2012.
Vuong T, Vesterinen A, Foumani M, Juvonen M, Seppälä J, Tenkanen M, Master ER.
Xylo- and cello-oligosaccharide oxidation by gluco-oligosaccharide oxidase from
Sarocladium strictum and variants with reduced substrate inhibition. Biotechnology for
Biofuels. 2013, 6: 148.
Foumani M, Vuong TV, MacCormick B, Master ER. Enhanced polysaccharide binding
and activity on linear β-glucans through addition of carbohydrate-binding modules to
either terminus of a glucooligosaccharide oxidase. PLOS ONE J. Accepted.
Manuscripts in Preparation
Vuong TV, MacCormick B, Master ER, Foumani M. Gluco-oligosaccharide oxidase
variants as suitable substitutes to glucose oxidase for baking applications.
Anticipated Manuscript
Vuong TV, Foumani M, Gudmundsson M, Master ER. Assessing enzymatic oxidation of
cellulosic substrates through XPS and fluorescence detection of carboxylic functionality.
3
Chapter 2 : Literature review
2.1 Oxidation of sugars and polysaccharides
Oxidation of hydroxyl groups to carbonyls can enhance the gelation, antiflocculation,
adhesion, thickening, and metal sequestration potential of polysaccharides (de Nooy et al.
1997; da Silva Perez et al. 2003). It can also alter the rheology of corresponding
polymers, and be performed as an initial step to subsequent esterification or amination of
hydroxyl groups. Pursuant to these objectives, chemicals such as 2,2,6,6-
tetramethylpiperidine-1-oxyl (TEMPO) have been used to oxidize primary hydroxyl
groups to uronic acids (Isogai and Kato 1998; da Silva Perez et al. 2003; Ciriminna and
Pagliaro 2010). Sodium periodate has also been used to oxidize C2 and C3 positions of
cyclic sugars, thereby introducing dialdehydes into polysaccharides (Kristiansen et al.
2010), whereas halide ions including I- and Br- have been used to further oxidize
aldehydes at positions C1 and C6 to aldonic acids (Diehl et al. 1974; Parikka et al. 2012).
However, chemical methods can compromise the polymerization and/or crystallinity of
the starting material, which is problematic when derivatizing oligosaccharide and
nanocrystalline substrates (Isogai et al. 2009; Saito et al. 2010).
Alternatively, carbohydrate oxidases can facilitate regio-selective oxidation of highly
functionalized carbohydrates without arduous protection/deprotection steps. Mild
reaction requirements also mean that loss in the degree of polymerization and
crystallinity of oligo- and poly-saccharide substrates can be minimized. Carbohydrate
oxidases (EC 1.1.3) can catalyze the oxidation of the primary hydroxyl (C6), secondary
hydroxyls (C2, C3 or C4) or anomeric carbon hydroxyl (C1) to an aldehyde, ketone or a
4
lactone (then carboxylic acid), respectively, with concomitant reduction of molecular
oxygen to H2O2 (van Hellemond et al. 2006). These enzymes were recently categorized
as auxiliary activities in the carbohydrate-active enzyme database (CAZy; Levasseur et
al. 2013). In the following paragraphs some of the enzymes from AA families will be
briefly reviewed.
2.2 Examples of oxidoreductases from selected Auxiliary Activity (AA) families
The most versatile branch of AA enzymes is perhaps AA3, which contains enzymes
belonging to the glucose-methanol-choline (GMC) oxidoreductase superfamily. These are
FAD containing enzymes include cellobiose dehydrogenase (CDH, EC 1.1.99.18,
AA3_1), a hemoflavoenzyme, oxidizing cellobiose and higher cellodextrin at the
anomeric position. The occurrence of CDH in wood degrading fungi, as well as oxidation
mechanism and structure-functional relationships, have been thoroughly reviewed for
CDH enzymes (Henriksson et al. 2000; Zamocky et al. 2006). Recent evidence suggests
that CDHs are physiological partners for polysaccharide monooxygenases, playing
important roles in oxidative cellulose decomposition (Langston et al. 2011; Phillips et al.
2011). From an applied perspective, the electron transfer by the CDH cytochrome domain
as well as its electrochemical properties make it suitable in biosensors and enzymatic
biofuel cell applications (Ludwig et al 2013).
The model enzyme for the GMC superfamily, glucose-1-oxidase (GO, EC 1.1.3.4,
AA3_2), oxidizes the anomeric hydroxyl of glucose, yielding a lactone that can be
hydrolyzed to form the corresponding acid (van Hellemond et al. 2006). GO has been
studied in detail given its importance in diagnostic reagents, biosensors, baking, and other
5
applications (Bankar et al. 2009). Unlike GO, aryl alcohol oxidase (AaO, EC 1.1.3.7,
AA3_2) is an extracellular flavoenzyme which is thought to be involved in fungal
degradation of lignin, providing H2O2 required for ligninolytic peroxidases (Hernández-
Ortega et al. 2012). Phylogenetic analyses of GMC oxidoreducases reveal that close to
the AaO cluster are pyranose dehydrogenases. Pyranose dehydrogenases (PDH, EC
1.1.99.29, AA3_2) have broader substrate specificity and regioselectivity than GO,
catalyzing the oxidation of C1, C2, or C3 hydroxyls of mono di- and tri-saccharides to
form the corresponding lactone or keto sugars (Peterbauer and Volc, 2010).
Unlike PDH, pyranose 2-oxidases (POX, EC 1.1.3.10, AA3_4) target C2/3 hydroxyls
(Giffhorn 2000; Kujawa et al. 2006) and can utilize molecular oxygen as an electron
acceptor. POX shows a hydride transfer mechanism for FAD reduction with a conserved
histidine as the catalytic base for deprotonation of the substrate; a feature that is also
conserved among all the above-mentioned GMC oxidoreducases (Wongnate et al. 2013).
The crystal structure of POX reveals a size exclusion mechanism for substrate binding
(Hallberg et al. 2004) similar to that observed in GO (Wohlfahrt et al. 1999). As a result,
the application of these enzymes is likely limited to the oxidation of mono- and di-
saccharides.
Whereas AA3 enzymes are flavoproteins, carbohydrate oxidases in family AA5 are
copper radical oxidases. Family AA5 comprises two subfamilies, the glyoxal oxidases in
subfamily AA5_1 and galactose oxidases in subfamily AA5_2. Rather than oxidizing
hydroxyl groups at C1, C2 or C3 positions, galactose 6-oxidase (GaOx, EC 1.1.3.9,
AA5_2) oxidizes the primary C6 hydroxyl of galactose along with galactose containing
oligosaccharides and polysaccharides (Whittaker 2005). The biochemistry and production
6
of GaOx has been studied in detail (Whittaker 2005; Spaduit et al. 2010) and extensive
analyses of its structure have revealed a comparatively shallow active site, explaining the
activity of this enzyme on galactopyranosyl units of galactoglucomannans, in addition to
monosaccharides (D/L-galactose) and oligosaccharides with terminal galactopyranosyl
units (Firbank et al. 2001; Parikka and Tenkanen 2009; Parikka et al. 2010). Studies with
GaOx reveal that a key benefit to enzymatic oxidation of polysaccharides is regio-
selectivity along with reduced loss in the degree of polymerization of oligomeric and
polymeric substrates (Parikka et al. 2010; Parikka et al. 2012).
Comparatively new classes of AA enzymes that act on polysaccharides include the lytic
polysaccharide monooxygenases (LPMOs), which are now classified as AA families 9,
10, 11, and 13 in the CAZy database. LPMOs are copper dependent enzymes involved in
oxidative cleavage of polysaccharides resulting in corresponding oxidized
oligosaccharides. These enzymes play a key role in lignocellulose degradation as they are
found in the genome of most plant cell wall degrading fungi (Morgenstern et al. 2014). In
addition, they show a synergistic boosting effect with hydrolytic enzymes and CDH,
however these latter enzymes are not necessary for the action of LPMOs (Dimarogona et
al. 2013; Vaaje-Kolstad et al. 2013).
Whereas fungal LPMOs from family AA9 (formerly GH61) target crystalline cellulose as
well as hemicellulose (Agger et al. 2014), bacterial LPMOs from family AA10 (formerly
CBM33) that have been characterized to date are selective towards cellulose and chitin
(Book et al. 2014). LPMOs from AA9 and AA10 families share a conserved ß-sandwich
fold as well as conserved histidine residues involved in copper binding within the
substrate binding site (Hemsworth et al. 2013; Book et al. 2014). However, studies on
7
the surface electrostatic potential of enzymes from families AA9 and AA10 reveal
significant differences in charge distribution particularly within the substrate binding
region (Book et al. 2014). While the anomeric C1 is the most favourable oxidation site
for LPMOs in general, AA9 enzymes have been also reported to oxidize hydroxyl groups
at the C4 position (Isaken et al. 2014).
Most recently, two additional LPMO families were discovered, namely AA11
(Hemsworth et al. 2014) and AA13 (Vu et al. 2014), which are specific towards chitin
and starch, respectively. These new classes share the conserved histidine brace for copper
binding in the active site and both require an electron donor such as ascorbic acid or
CDH. Similar to previously discovered LPMOs, C1 hydroxyl groups are oxidized by
AA11 and AA13 enzymes, although action of AA11 enzymes at the non-reducing C4 has
also been reported (Hemsworth et al. 2014).
Because LPMOs cleave glycosidic linkages in targeted polysaccharide, their action
ultimately reduces the degree of polymerization of the starting oligo- and
polysaccharides. By contrast, oligosaccharide oxidases from family AA7 oxidize the
anomeric carbon of existing reducing ends in oligomeric and polymeric substrates.
Despite the significance of the AA7 enzymes for carbohydrate oxidation, this enzyme
family is comparatively less well characterized. Given the importance and relevance of
this enzyme family to the context and scope of the present study, these enzymes will be
reviewed in detail in the following section.
8
2.3 Oligosaccharide oxidases within AA7
Oligosaccharide oxidases from family AA7 belong to the growing vanillyl-alcohol
oxidase (VAO) flavoenzyme family (Leferink et al. 2008), targeting the C1 hydroxyl of a
broad range of oligosaccharides, including cello-, -xylo-, and malto-oligosaccharides.
Examples of characterized AA7 enzymes include a carbohydrate oxidase from
Microdochium nivale (MnCO) (Xu et al. 2001), a cello-oligosaccharide oxidase from
Paraconiothyrium sp. (PCOX) (Kiryu et al. 2008), a chito-oligosaccharide oxidase from
Fusarium graminearum (ChitO) (Heuts et al. 2007), and a gluco-oligosaccharide oxidase
(EC 1.1.3.-) from Sarocladium strictum T1 (GOOX-T1) (Lin et al. 1991; Lee et al. 2005).
The above-mentioned enzymes are flavoproteins, with unique bi-covalent linkages to the
flavin adenine dinucleotide (FAD) cofactor, providing a relatively high redox potential
for these enzymes. Like other flavin carbohydrate oxidases that target the anomeric
carbon hydroxyl (C1), oligosaccharide oxidases are thought to mediate oxidoreductase
activity through two half-reactions: 1) oxidation of the reducing sugar to the
corresponding lactone, then 2) spontaneous hydrolysis of the lactone product to the
corresponding acid (Huang et al. 2005; van Hellemond et al. 2006).
2.3.1 Oligosaccharide oxidase from Microdochium nivale, MnCO
The gene encoding MnCO contains one intron and the coding region shows low
similarity to other FAD-containing carbohydrate oxidases other than AA7 family
enzymes (Xu et al. 2001). MnCO was recombinantly expressed in Aspergillus oryzae and
was well characterized in terms of specificity and kinetics parameters (Kulys et al. 2001a;
Xu et al. 2001). The crystal structure of the enzyme was also solved by Duskova et al.
9
(2009). Among the many different oligosaccharides, MnCO prefers tetrameric dextrins,
revealing that four α-(1→4) linked glucose units make a favourable interaction with the
substrate-binding pocket. Notably, this enzyme was the first reported AA7 enzyme that
can oxidize polysaccharides, including starch and carboxymethyl cellulose with 0.8 %
and 9 % activity relative to cellobiose, respectively (Xu et al. 2001). Compared to glucose
oxidase, the reactivity of this enzyme is lower on glucose, however the substrate
specificity of MnCO is broader towards mono- and di-saccharides (Kulys et al. 2001a).
Thus, this enzyme has been tested to replace glucose oxidase in biosensors (Kulys et al.
2001b) and baking applications (Schneider et al. 2003).
2.3.2 Cello-oligosaccharide oxidase from Paraconiothyrium sp., PCOX
PCOX efficiently oxidizes β-(1→4) linked sugars, such as cellooligosaccharides,
xylobiose and lactose at an optimal pH of 5.5 (Kiryu et al. 2008). Like MnCO, PCOX is
active on a wide range of sugar types ranging from glucose and galactose to xylose and
arabinose. Moreover, the results of Kiryu et al. (2008) indicate that the enzyme is capable
of oxidizing xylooligosaccharides in addition to cellooligosaccharides. This enzyme has
been mainly used to produce lactobionate, a lactose derivative with potential
nutraceutical benefits including prebiotic activity (Murakami et al. 2008). Despite the
wide substrate specificity of this enzyme, very little information is available from the
genetic perspective. Neither the nucleotide sequence of the gene encoding PCOX, nor the
amino acid sequence of this protein has been published, which limits our ability to study
this protein based on sequence homology to other enzymes.
10
2.3.3 Chitooligosaccharide oxidase from Fusarium graminearum, ChitO
ChitO catalyzes the oxidation of hydroxyl moieties at the C1 position of
chitooligosaccharides. Mutagenesis studies using the ChitO encoding gene reveal that the
conversion of the active site residue Q268 to R268 affects the recognition of N-acetyl
groups present on chitooligosaccharide substrates (Heuts et al. 2007). Interestingly, the
corresponding amino acid position in most other AA7 oligosaccharide oxidases,
including GOOX-T1, is arginine. Accordingly, arginine at this position is predicted to
hinder the formation of favourable interactions between branched molecules such as
glucosamine with GOOX-T1 active sites (Heuts et al. 2007). More recently, Ferrari et al.
(2015) created a set of ChitO variants with completely different substrate tolerance than
the wild-type enzyme. By combining single mutants with altered substrate preference, a
variant with activity towards lactose, cellobiose and maltose was generated although with
lower oxidation efficiency than those of GOOX and MnCO on these substrates. Notably
the engineered ChitO retained 40% of its catalytic efficiency towards
chitooligosaccharides.
2.3.4 Gluco-oligosaccharide oxidases from Sarocladium strictum T1, GOOX-T1
The GOOX-T1 gene consists of 1500-bp of coding sequence and one short 53-bp intron.
The deduced protein sequence is 499 amino acids and the predicted protein has a
molecular mass of 55.2 kDa. The predicted molecular weight of GOOX-T1 is lower than
the reported 61 kDa that was obtained from size exclusion chromatography. This
discrepancy is consistent with the N-glycosylation detected at Asn341 and Asn305
(Huang et al. 2005). Indeed, post-translational modification of GOOX-T1 might be
11
required for functional protein folding since recombinant expression of this enzyme in
E.coli was not successful; it is also possible that E. coli does not support complete
incorporation of the required FAD cofactor. By contrast, the recombinant GOOX
expression has been demonstrated in Pichia pastoris with yields reported as high as 300
mg per liter of cultivation medium (Lee et al. 2005).
GOOX-T1 is shown to function best at 37 °C and pH 8, and remain stable up to 50 °C. It
also shows a wide range of pH stability from pH 4 to pH 12 (Lin et al. 1991; Fan et al.
2000). The impact of temperature and pH on GOOX-T1 activity was studied extensively
using cello- and malto-oligosaccharides (Fan et al. 2000). In their study, Fan et al (2000)
revealed that the oxidation of maltose was highest between pH 9 to 10.5 and the Km of
this reaction was also highest at pH 10. Fan et al. (2000) also demonstrated that the
activation energy of GOOX is similar at pH 7 and pH 10, suggesting that the catalytic
mechanism of GOOX is retained within this pH range.
Figure 2.1 Oxidation mechanism of GOOX. The tyrosine residue (Y429) is a catalytic base and along with aspartic acid (D355) initiates the hydride transfer from the substrate, e.g. glucose to reduce the FAD cofactor, which is bi-covalently linked to GOOX through a cysteine (C130) and a histidine (H70) residue. The image was generated using ChemSketch.
12
Among tested carbohydrates and derivatives, GOOX-T1 oxidizes both α-linked and β-
linked glucose substrates, including lactose, malto-oligosaccharides and cello-
oligosaccharides (Lin et al. 1991; Fan et al. 2000; Lee et al. 2005). The highest catalytic
efficiency of native GOOX-T1 is observed with cellotriose (Lee et al. 2005). Notably,
this GOOX did not oxidize xylose, galactose, or many other sugars (Lin et al. 1991).
The crystal structure of GOOX-T1 is resolved by Huang et al. (2005) and was proposed
that Tyr429 initiates sugar oxidation by proton abstraction from the C1 hydroxyl,
followed by H1 hydride transfer to the N5 position of the FAD cofactor (Figure 2.1)
(Huang et al. 2005). Notably, the FAD is covalently bound by two amino acids, His70
and Cys130; this unique configuration is predicted to modulate the oxidative potential of
the FAD cofactor. Residues predicted to be involved in substrate binding and catalytic
Figure 2.2 GOOX substrate binding and active sites. A) Residues predicted to be involved in GOOX substrate binding B) Residues involved in FAD bi-covalent linkages and oxidation of substrate through FAD reduction. The predicted hydrogen bonds are shown with gray dashed-lines. The figure was generated using PyMOL.
13
mechanism of GOOX are illustrated in Figure 2.2. The crystal structure of GOOX-T1
further reveals that the enzyme possesses an open carbohydrate-binding groove, allowing
the accommodation of oligosaccharide substrates (Lee et al. 2005; Huang et al. 2008).
2.4 Current strategies for enzyme engineering
Several strategies have been developed for enzyme engineering including random
mutagenesis, site-directed mutagenesis, recombination, and fusion proteins. The
following paragraphs briefly review common protocols for each method, along with
corresponding advantages and disadvantages; the information is also summarized in
Table 2.1.
2.4.1 Mutagenesis: Random versus rational design
Random mutagenesis is a technique to obtain an improved enzyme by introducing
random mutations into the corresponding gene and then screening the recombinant
enzymes for mutants with desirable characteristics. A key requirement for this approach
is easy assessment of enzyme activity, for example via a colorimetric detection of
reaction products, This approach also benefits from the ability to express the target
enzyme in E. coli. In this way, activity measurements are less confounded by differences
in levels of recombinant protein expression between transformants. This method does not
require any information about the protein sequence nor the crystal structure of the
protein. The technique is usually applied to enhance thermostability or other
characteristics that are not easily predicted from protein sequence or structural
information. Several reviews have been published on the techniques used for random
mutagenesis (Cadwell and Joyce 1992; Cadwell and Joyce 1994; Bloom et al. 2005) and
14
consistently report the advantages of error-prone PCR over other methods. The approach
typically generates mutants through error-prone PCR, which applies a low fidelity Taq
polymerase. Notably, the error rate of the Taq polymerase used for this purpose is the
highest of the known DNA polymerases, approximately 2×10-5 compared to the error rate
of 4×10-7 for commercially available high fidelity polymerases. Additional changes to the
PCR reaction conditions can further increase the error rate, for example, increasing or
unbalancing the concentration of deoxynucleosides (dNTPs), addition of MnCl2, and
increasing the concentration of MgCl2 or Taq polymerase (Cadwell and Joyce 1992).
In contrast to random mutagenesis, site-directed mutagenesis requires knowledge of the
sequence and ideally the crystal structure of the protein. This method is usually used
when active sites of the enzyme are known and the engineering goal is to alter substrate
specificity. For instance, site-directed mutagenesis can be used to change a few amino
acid residues that are thought to prevent productive interactions with a target compound.
Molecular modeling software is typically used to help reveal amino acid residues
involved in substrate-enzyme interactions, and to compare the active sites of homologous
enzymes. For instance, Visual Molecular Dynamics (VMD) is a freely accessible
software (Humphrey et al. 1996) that provides tools for structural superimposition of
homologous proteins and compares the 3-dimenstional positioning of substrate
interacting residues; this tool is commonly used for proposing mutants with altered
specificity. Among the reported methods for site directed mutagenesis, non-PCR based
methods are more reliable since the primers are designed such that only the parental
plasmids are amplified in a linear fashion, preventing error propagation during successive
rounds of thermal cycling. Moreover, high accuracy is maintained through the use of
15
PfuTurbo DNA polymerase, to reduce random errors. Currently, the most cited
commercial kit for site-directed mutagenesis is QuikChange from Stratagene.
2.4.2 Gene shuffling
Gene shuffling is another approach to improve the functionality of a protein.
Recombining structurally similar proteins generate even more changes to the protein
sequence than random mutagenesis (Drummond et al. 2005). Yet, like random
mutagenesis, this approach is particularly valuable when the change required for a
desirable function is not predictable from the protein sequence or structure; it also
depends on an easy screen for protein function. Gene shuffling can be combined with
site-directed mutagenesis to produce a consensus gene sequence that comprises the most
frequent nucleotide residues identified from an alignment of genes corresponding to
homologous proteins (Lehmann and Wyss 2001; Lehmann et al. 2002; Steipe 2004).
2.4.3 Fusion proteins
Fusion proteins can be generated using a combination of PCR and recombination to fuse
domains from different proteins. For instance, the Green florescence protein (GFP) is a
common tag, which is fused to various proteins for quantitative bio analysis and easy
detection and localization studies (Remington 2011). Similarly, carbohydrate binding
modules (CBM) from naturally occurring glycoside hydrolases have been fused to
catalytic domains of other enzymes to increase the specific activity of the enzyme
towards cellulose and other polymeric carbohydrates (Shoseyov et al. 2006). Examples of
the CBM fused proteins will be reviewed in the following section.
16
Table 2.1 Overview of common enzyme engineering strategies
Approach Typical uses Advantages Disadvantages References
Random mutagenesis
When the characteristic under study is not predictable from protein sequence or structure
No information about sequence or structure is required
An easy screening method is required Ideally, targeted protein can be functionally expressed in E.coli
Cadwell and Joyce 1992 Bloom et al. 2005 Cadwell and Joyce 1994
Site-directed mutagenesis
When rational design is possible by structure-function correlations
Any expression system can be applied
A few mutations usually results in moderate changes to the function
Bhat 1996 Costa et al. 1996 Braman et al. 1996
Gene-shuffling When the function under study is not predictable or conserved among the sequence or structure of homologous proteins
Results in more significant changes
Benefits from sequence information for homologous proteins that score high on desired function.
Drummond et al. 2005 Steipe 2004 Lehmann and Wyss 2001 Lehmann et al. 2002
Fusion proteins When a tag is required for detection, binding or improved specificity
Specific choices are available depending on the application
The fusion module might change the conformation or functionality of the protein
Remington 2011 Shoseyov et al. 2006
2.5 Carbohydrate binding modules
Carbohydrate active enzymes that act on high molecular weight polysaccharides
including insoluble cellulose fibrils often contain carbohydrate binding modules (CBMs)
that can promote functional association of the enzyme and targeted substrate (Shoseyov
et al. 2006). To date, CBMs have been classified into more than 70 families based on
amino acid sequence similarities (www.cazy.org; 2015). These modules have been
further grouped into three types based on folding and substrate specificity (Boraston et al.
2004; Gilbert et al. 2013). Type A CBMs possess a flat binding site, which is thought to
17
associate with surfaces presented by crystalline cellulose. The planar architecture of the
binding site shows no or little affinity towards soluble carbohydrates (Zhang et al. 2012).
By contrast, Type B CBMs contain a cleft or a groove architecture that is better suited to
the conformation of amorphous cellulose or oligosaccharide chains. The depth of the
binding groove varies among Type B CBMs; it can be shallow or deep enough to
accommodate the entire width of a pyranose ring, as in CBM4-2 from C. fimi (Boraston
et al. 2002; Christiansen et al. 2009). Finally, lectin-like Type C CBMs have binding sites
that form several hydrogen bonds with sugar molecules typically having less than three
monosaccharide units (Notenboom et al. 2002).
2.5.1 Contributions in cellulose/hemicellulose active enzymes
The varied contributions and significance of CBMs on cellulolytic enzymes was recently
reviewed (Várnai et al. 2014). Notably, recent genome sequences indicate that the
majority of predicted cellulolytic enzymes lack CBMs and that CBMs are somewhat
enriched among cellobiohydrolases (Palonen et al. 2004). Whereas certain CBMs can
increase non-productive binding to lignin present in lignocellulose substrates (Várnai et
al. 2014; Palonen et al. 2004), CBMs can also improve the performance of cellulolytic
enzymes on insoluble substrates, particularly when presented with low substrate
concentrations (Tomme et al. 1988; Boraston et al. 2003; Costaouëc et al. 2013; Várnai et
al. 2013). Similar positive effect of CBMs is observed in mananases (Hagglund et al.
2003; Mizutani et al. 2012) and in some cases for xylanases (Lehtiö et al. 2003; Meng et
al. 2015) as truncated versions of these enzymes lacking the corresponding CBMs show
significant reduction of binding and activity towards insoluble mannans and xylans,
respectively. Moreover, CBMs from thermophiles can increase the thermostability of
18
carbohydrate-active enzymes (Charnock et al. 2000; Jun et al. 2009). Further relevant
examples will be reviewed in section 4.1.
2.5.2 Industrial applications of CBMs
CBMs have been applied broadly, and new applications continue to be reported. In
medical science, the affinity of CBMs contained in grass and dust allergens towards
oxidized cellulose was used to treat allergies and asthma (Shani et al. 2011). In terms of
process design, a family 3 CBM has been developed as an easy and cost effective tag for
protein purification (Guerreiro et al. 2008). To provide a tag free purification system, a
formic acid recognition site for chemical cleavage (Ramos et al. 2010) or an intein region
to excise and re-join the remaining protein, was designed between CBMs and target
proteins; this method was successfully applied for protein purification in E. coli and P.
pastoris (Wan et al. 2011). In the detergent industry, the effect of CBMs on enzyme
affinities has been used to improve the performance of laundry powders, where chimeric
amylases, proteases, lipases, and oxidoreductases are employed (Osten et al. 2000 a;
Osten et al. 2000 b).
2.5.3 Effects of CBMs on neighbouring modules
Recent reports on CBMs indicate that these modules can affect both the activity and
thermostability of the cognate catalytic module. In most cases, the activity of the
neighbouring domain is improved, harnessing the affinity of CBMs towards soluble and
insoluble polysaccharides. The significant decrease in binding affinity, and often catalytic
activity, of glycoside hydrolases upon genetic truncation of CBMs from the catalytic
module also reveals the boosting function of CBMs on enzyme activity (Ali et al. 2001;
19
Ali et al. 2005). The enhancement of activity has been attributed to: 1) a proximity effect,
2) a targeting function and 3) a disruptive function (Boraston et al. 2004). Firstly, through
binding to the substrate, CBMs would increase the local concentration of the polymeric
substrate relative to the active site of the enzyme. Secondly, the selectivity of the CBM
towards specific polymers, allows targeted binding to the substrate of interest from a
mixture of various polymers; this would be particularly relevant for enzyme applications
on plant biomass given the heterogeneity and complexity of the polysaccharides present.
Lastly, the potentially disruptive function of CBMs on cellulose could increase
amorphous structures within an otherwise crystalline substrate, although this role of
CBMs does not appear to be universal.
Besides enhancing the activity, varying effects of CBM fusion on the temperature
stability of associated enzymes have been reported. For instance, Jun et al. (2009) show
that appending a xylan specific CBM from Thermotoga maritima to xylanase 2 from
Hypocrea jecorina improves the thermostability and substrate affinity of the enzyme. In
another study, fusion of a family 42 CBM from Aspergillus kawachii to a feruloyl
esterase from A. awamori was shown to increase enzyme stability and affinity towards
arabinoxylan (Koseki et al. 2010). However, while fusion of a family 6 CBM from
Clostridium stercorarium Xy1A to Bacillus halodurans Xy1A does not affect enzyme
stability (Mangala et al. 2003), fusion of a family 22 CBM to B. halodurans C-125 family
10 xylanase decreases the thermostability of the enzyme (Mamo et al. 2007). The range
in effects of CBMs on protein stability is further exemplified by Kataeva et al. (2001),
who show that the native CBM4 of CelK from C. thermocellum increases the
20
thermostability of the catalytic module, which is not retained when substituting CBM4
for a CBM6 encoded by the same organism.
2.5.4 Important factors for designing CBM fusion proteins
Construction of chimeric enzymes is simple in theory; however, many challenges arise
when they are generated in practice. Several factors need to be considered when
designing the CBM-fusion enzymes. First, the type of CBM should be selected in a way
that imparts substrate selectivity to the fused enzyme. Ye et al. (2011) report that several
CBMs from different families (3,4,6,9) and types (A, B and C) were fused to C.
thermocellum cellodextrin phosphorylase (CtCDP) and only one of them enhanced
CtCDP activity on amorphous cellulose; the other fusion enzymes either showed less or
similar activity to the wild-type enzyme.
Second, the positioning of the CBM at the N-terminus or C-terminus of the catalytic
module should be thoughtfully selected. Notably, while family 4 CBMs are often located
at the N-terminus of GH9 glycosyl hydrolases, family 3 CBMs are typically located at the
C-terminus of this enzyme family (Kataeva et al. 2001). Moreover, Kateava et al.
reported that the fusion of a family 4 cellulose specific CBM to the C-terminus of
feruloyl esterase domain of a xylanase from C. thermocellum (FAE XynZ) did not
significantly affect the domain function, whereas positioning this CBM at the N-terminus
increased FAE XynZ affinity towards acid swollen cellulose (Kataeva et al. 2001).
Therefore, it appears that the CBM should be fused to the terminus that places the module
in closer proximity to the enzyme active site while not obstructing substrate accessibility
to the catalytic module.
21
Third, the stability towards proteases, and flexibility of the linker sequence, which is used
to connect the CBM to the catalytic module, should be carefully considered. In some
cases, the linker sequence affects the performance of the chimeric enzyme. For example,
Dias et al. report that the thermostability of C. thermocellum xylanase Xyn10B was
retained after removing the CBM22 domain and leaving the linker sequence, while
removing the whole linker-CBM22 sequence resulted in reduced thermostability of the
enzyme (Dias et al. 2004).
Lastly, if the microbial origin of the CBM or catalytic domain of the recombinant protein
is different from the expression host, codon optimization could be beneficial (Daly and
Hearn 2005). Codon optimization can be performed commercially, as a means to enhance
the expression of the full-length protein. For instance, codon optimization for yeast
resulted in 10.6 fold increase in protein expression of human glucocerebrosidase in P.
pastoris (Sinclair and Choy 2002). Similar strategies also resulted in increased expression
of α-amylase in P. pastoris (Tull et al. 2001).
Having reviewed the protein elements and engineering strategies most pertinent to this
study of carbohydrate oxidases, section 2.6 will review existing applications of
carbohydrate oxidase activity.
2.6 Applications of carbohydrate oxidase in food industry
2.6.1 Oxidoreductases in baking applications
In baking applications, carbohydrate oxidases, such as glucose oxidase from Aspergillus
niger (GO) and oligosaccharide oxidase from M. nivale (MnCO) have been used to
enhance the quality of the dough. Several authors have demonstrated the use of GO in
22
flour (Vemulapalli et al. 1998; Rasiah et al. 2005; Bonet et al. 2006; Hanft and Koehler
2006; Dagdelen and Gocmen 2007; Decamps et al. 2013) this enzyme converts glucose
constituents of the dough into gluconic acid, while reducing molecular oxygen to H2O2.
The H2O2 is shown to be the active reagent of a GO treatment, as it oxidizes different
positions of wheat proteins, i.e. gluten, including free thiols in cysteine residues to form
disulfide bonds, and phenolic tyrosine residues to generate di-tyrosines through oxidative
coupling (Hanft and Koehler 2006). These cross-linkages reinforce the protein network of
the dough (Rasiah et al. 2005), thereby reducing stickiness and enhancing machinability
of the dough. H2O2 also tends to generate a dry surface on the dough by oxidation and
gelation of water-soluble pentosans (Hanft and Koehler 2006). The water intake by
pentosans also improves freshness and softness of the baked products upon long storage
(Bonet et al. 2006).
Despite the advantages of GO to baking applications, its effectiveness is limited by the
selectivity of the enzyme towards glucose and susceptibility to inactivation by H2O2. An
early study by Kleppie (1966) suggests that in acidic pH, less than 0.01 M H2O2 oxidizes
methionine residues that are close to or in the active cleft of reduced GO. More
specifically, the methionine residues were shown to oxidize to methionine sulfoxide,
where the amount of methionine sulfoxide measured increased as the enzyme was
exposed to higher concentrations of H2O2 e.g. 0.1- 0.2 M, which in turn resulted in a
dramatic drop in enzyme performance by over 80%. Kleppie (1966) further highlighted
that GO in its reduced form is inhibited at least 100 times more readily than when in its
oxidized form. Hachimori et al. (1964) also studies the effect of H2O2 on several proteins
23
at alkaline pH, and report that in all cases, a tryptophan residue is oxidized by H2O2,
albeit to different extents.
The mechanism and kinetics of GO inhibition by H2O2 was studied in more detail by Bao
et al. (2003). Based on corresponding kinetics profiles in the presence of increasing H2O2
concentration, a competitive inhibition mechanism was suggested where the inhibition
constant was reported to be equal to the Michaelis constant KM. This result implies that
the affinity of GO in the reduced form to H2O2 is almost similar to that of oxygen. This
finding is consistent with their earlier report on competitive inhibition of immobilized
GO by H2O2 where the inhibition constant was also similar to the apparent Michaelis
constant (Bao et al. 2001).
Greenfield et al. already highlighted the importance of GO inactivation by H2O2 in 1975.
In that study, they demonstrate that the stability of immobilized GO in the presence of
H2O2 is inferior in a continuous operation than in storage tests. Therefore, they concluded
that for industrial usage of GO, it will be necessary to either directly reduce H2O2
inactivation or else indirectly minimize enzyme inactivation through GO immobilization
and/or addition of catalase. Much later, Yoshimoto et al. (2004) introduced a liposomal
capsulated GO system to decompose the H2O2 and protect GO from inactivation by H2O2.
The fluorescence properties of tryptophan residues and the FAD cofactor of GO at high
conversion of glucose revealed that the tertiary structure of the free enzyme is disordered,
while the encapsulated GO is protected (Yoshimoto et al. 2006).
Alternatives to GO are oligosaccharide oxidases, showing broader substrate specificity
and higher activity on oligosaccharides. Schneider et al. (2003) studied the rheology of
24
gluten content of the dough after addition of an oligosaccharide oxidase from M. nivale
(MnCO) and found that the enzyme increases the elastic modulus of the gluten, thereby
increasing the dough elasticity in a dose-dependent manner. They also studied the dough
consistency in terms of dough stickiness and firmness. The results indicated that the M.
nivale carbohydrate oxidase provides excellent dough consistency at 200-300 units/kg, as
evaluated using a scoring system by a skilled baker (Schneider et al. 2003). Despite the
better performance of MnCO compared to GO as a result of its wider substrate specificity
(Kulys et al. 2001a), this enzyme was reported to be also inactivated in the presence of
H2O2 (Nordkvist et al. 2007); however, the mechanism and extent level of H2O2
inactivation of MnCO was not reported.
2.6.2 Enzyme utilization for production of prebiotics
Carbohydrate active enzymes have been harnessed to produce prebiotic compounds.
Prebiotics were first described as “nondigestible food ingredients that beneficially affect
the host by selectively stimulating the growth and/or activity of one or a limited number
of bacteria in the colon, thus improving host health” (Gibson 1997). This definition was
later refined to include benefits from selective growth of microorganisms in other
sections of the gastrointestinal system: “a selectively fermented ingredient that allows
specific changes, both in the composition and/or activity in the gastrointestinal
microbiota that confers benefits.” (Gibson et al. 2004)
Carbohydrates with known prebiotic activity include but are not limited to
fructooligosaccharides, xylooligosaccharides (Kondepudi et al. 2012), konjac
glucomannan (Al-Ghazzewi et al. 2007), β-glucan, and arabinoxylan (Crittenden et al.
25
2002). As reviewed by Panesar et al. (2013) most enzymes used in prebiotic production
are glycoside hydrolases, which hydrolyze starting polysaccharides into a series of
oligosaccharides (Panesar et al. 2013). Very few studies have been done to harness other
enzyme types for prebiotic production, or to investigate the effect of those enzyme
treatments on the prebiotic activity of the product. For instance, oligosaccharide oxidases
such as GOOX-T1, MnCO and PCOX have been used to oxidize lactose into lactobionic
acid, which is a lactose derivative and potential prebiotic (Lin et al. 1996; Nordkvist et al.
2007; Murakami et al. 2008). However, the prebiotic potential of products prepared
using oligosaccharide oxidases has not been directly evaluated.
The prebiotic activity of a substrate can be studied both in vivo and in vitro. In the former
approach, often mice are fed with the potential prebiotic substrate and fecal samples are
collected for microbial analysis and to document fecal frequency as well as water content
(Wu et al. 2011). For in vitro studies, potential prebiotics are tested for their ability to
selectively stimulate the growth of bacteria correlated with healthy digestion, such as
species of Bifidobacteria and Lactobacilli, as compared to bacteria correlated with
unhealthy digestion including species of Clostridia and Bacteroides. In this case,
bacterial growth is typically measured in terms of viable cell counts or optical density
(Crittenden et al. 2002).
The prebiotic index (PI) was developed as a quantitative score to better compare the
prebiotic activity of different carbohydrates. Specifically, PI is defined as the portion of
healthy microflora subtracted by that of the unhealthy bacteria (Palframan et al. 2003).
For instance, when Bifidobacteria (Bif) and Lactobacilli (Lac) are considered as healthy
26
bacteria and Bacteroides (Bac) and Clostridia (Clos) constitute unhealthy bacteria, the
following equation describes PI correspondingly.
𝑃𝐼 = �𝐵𝑖𝑓𝑇𝑜𝑡𝑎𝑙
� + �𝐿𝑎𝑐𝑇𝑜𝑡𝑎𝑙
� − �𝐵𝑎𝑐𝑇𝑜𝑡𝑎𝑙
� − (𝐶𝑙𝑜𝑠𝑇𝑜𝑡𝑎𝑙
)
,where Bif, Lac, Bac and Clos are numbers of corresponding bacteria at sample time
divided by their numbers at inoculation; and Total refers to total bacteria numbers at
sample time divided by numbers at inoculation. Although PI has been used for systematic
analysis of commercial and novel prebiotics, a recent review by Bindels et al. (2015)
argues the selectivity criteria for prebiotics, and challenges our previous understanding of
healthy over unhealthy gut microorganisms. Thus, it is likely that new measures will be
developed in the coming years to better define and regulate prebiotic products.
In light of studies to date concerning carbohydrate oxidases, opportunities for protein
engineering, and existing applications of these enzymes, section 2.7 describes the
research hypotheses and specific research objectives addressed through my PhD thesis.
2.7 Research Hypotheses and Specific Research Objectives
The main objective of my PhD thesis was to study the enzymatic oxidation of plant oligo-
and polysaccharides by a glucooligosaccharide oxidase from Sarocladium strictum
(GOOX). This enzyme has reported activity on glucose and its oligomers with up to
seven sugar units. Thus, a specific aim of my PhD thesis was to engineer the GOOX
enzyme to expand its substrate specificity towards a broader range of plant
polysaccharides and oligosaccharides. My research hypotheses were:
1. Site-directed mutagenesis of the non-conserved residues in the active site of
27
GOOX will alter the sugar specificity of this enzyme so as to act not only on
glucose but also on xylose and galactose.
2. Fusion of selected carbohydrate binding modules to the GOOX catalytic domain
will extend its activity and binding capacity towards longer chain substrates e.g.
polysaccharides.
3. The GOOX-oxidized oligosaccharides can be directly used as sugar standards for
lytic polysaccharide monooxygenases (LPMOs), such as enzymes from family
AA9 (formerly GH61). Also, the enzymatic oxidation of oligo- and
polysaccharides will improve their nutraceutical value in various food
applications. In addition, the preferred oxidation of oligosaccharides versus
monosaccharides by GOOX makes this enzyme advantageous over commonly
used glucose oxidase for baking applications.
To test the above mentioned hypotheses, the following approaches were used:
1. Site-directed mutagenesis of the GOOX
a) The gene encoding GOOX protein was recovered from S. strictum RNA and
the non-conserved residues predicted to play an important role for substrate
specificity were mutated using the QuikChange site directed mutagenesis
technique. The wild type and mutant GOOXs were expressed in Pichia
pastoris and purified using an affinity chromatography method.
b) Enzyme kinetics study was performed using selected mono-, di- and tri-
saccharides to assess the effect of mutations on kinetics parameters.
28
2. CBM fusion to the GOOX
a) Selected CBM genes from Clostridium thermocellum, CBM3, CBM11, and
CBM44, were appended to both N- and C-terminus of the GOOX via natural
and synthetic linkers generating six fusion proteins that were expressed and
purified in P. pastoris.
b) The effect of the CBM on the binding capacity of protein fusions was assessed
using affinity gel electrophoresis; specific activities and thermostabilities were
compared using the standard GOOX assay, which detects H2O2 production.
3. Application of the GOOX-oxidized products
a) To confirm whether GOOX-oxidized oligosaccharides can be used as sugar
standards relevant to LPMO characterizations, the oxidized products were
characterized using NMR and mass spectroscopy to assess the oxidation
position and percent conversion.
b) To assess the prebiotic activity of plant oligosaccharides and polysaccharides
before and after GOOX treatment, Bifidobacterium. longum was grown on
oxidized and non-oxidized oligo- and poly-saccharides and the growth was
measured using viable cell count and optical density. The growth rates were
calculated and compared for each carbohydrate source.
c) To assess the advantage of GOOX over glucose oxidase in baking applications,
selected mono- and oligosaccharides were treated with both enzymes and
specific activities were measured. The inactivation of each enzyme by H2O2
was also compared.
29
Chapter 3 Improved GOOX activity on mono and oligosaccharides using
site-directed mutagenesis
Parts of this chapter are published in:
Foumani M, Vuong T.V, and Master E.R. 2011. Altered substrate specificity of the
gluco-oligosaccharide oxidase from Acremonium strictum, Biotechnology and
Bioengineering, 108(10): 2261-2269.
Contributions: Design of the study; performing the experiments, data collection and
analyses corresponding to strain cultivation, gene cloning, site-directed mutagenesis,
recombinant protein expression and purification, deglycosylation assay, specific activity
measurements, and kinetics studies; and manuscript preparation.
30
3.1 Introduction
Carbohydrate oxidases (EC 1.1.3) can catalyze the oxidation of hydroxyl groups to an
aldehyde, ketone or a lactone (then carboxylic acid), with concomitant reduction of
molecular oxygen to H2O2 (van Hellemond et al. 2006). Given the ease of detecting
H2O2, several carbohydrate oxidases, including glucose oxidase (GO) and pyranose
oxidase (POX), have been widely applied in clinical biosensors. Galactose oxidase
(GaOX) is also used to oxidize the primary C6 hydroxyl groups of polysaccharides
containing terminal galactose units (e.g. galactoglucomannan, galactomannan, and
xyloglucans) to alter the rheology of these compounds (Parikka et al. 2010).
By contrast, oligosaccharide oxidases that oxidize C1 hydroxyl groups of β-1,4-linked
sugars could be used to derivatize xylan and cellulosic substrates. Gluco-oligosaccharide
oxidase from Sarocladium strictum, previously Acremonium strictum, strain T1 (GOOX-
T1) (Lin et al. 1991) is an example of such enzyme. The crystal structure of GOOX-T1
reveals that similar to CDH and GaOX (Firbank et al. 2001; Hallberg et al. 2003), the
enzyme possesses an open carbohydrate-binding groove, allowing the accommodation of
oligosaccharide substrates (Figure 3.1A).
A screen of more than 50 carbohydrates and derivatives show that GOOX-T1 oxidizes
both α-linked and β-linked glucose substrates, including lactose, malto-oligosaccharides
and cello-oligosaccharides (Lin et al. 1991; Fan et al. 2000; Lee et al. 2005). The catalytic
efficiency of native GOOX-T1 is highest with cellotriose (Lee et al. 2005); however, this
GOOX did not oxidize either xylose, and galactose, or many other sugars (Lin et al.
1991).
31
Figure 3.1: The structural model of GOOX built by the Swiss-Model Workspace using the X-ray structure of GOOX-T1 (PDB ID: 2AXR). (A) The gross structure of GOOX with the active site containing the intermediate analogue 5-amino-5-deoxy-cellobiono-1,5-lactam (ABL) and the FAD cofactor. (B) The location of key residues Y300, W351, and N388 in relation to the ABL and the FAD cofactor (for neatness, the side chain of W300, N388 and a part of FAD are shown). Hydrogen bonds are shown as dashed lines.
While the catalytic mechanism of GOOX has been characterized, residues that affect the
substrate preference of this enzyme are still unknown. Given the limited arsenal of
biocatalysts that can be applied for oxidative modification of plant oligosaccharides,
GOOX variants with gained activity on xylose, galactose, and/or mannose containing
substrates would constitute a valuable set of new industrial enzymes. Accordingly, this
article reports the purification and substrate specificity of a gluco-oligosaccharide oxidase
from an S. strictum strain, and the improvement of its substrate specificity through site-
directed mutagenesis.
32
3.2 Materials and Methods
3.2.1 Fungal strain and materials
Sarocladium strictum (previously Acremonium strictum) type strain CBS 346.70 was
obtained from the American Type Culture Collection (ATCC) No.34717. 4-
aminoantipyrine (4-AA), 5-bromo-4-chloro-3-indolyl phosphate (BCIP), biotin, histidine,
imidazole, nitroblue tetrazolium (NBT) and phenol, as well as alkaline phosphatase-
linked anti-Rabbit IgG conjugates, anti-Myc antibodies, and horseradish peroxidase were
purchased from Sigma. Glucose, galactose, mannose, arabinose, N-acetyl-glucosamine,
xylose, maltose, and cellobiose were purchased from Sigma, while cellotriose,
mannobiose, mannotriose, xylobiose, and xylotriose were purchased from Megazyme. All
carbohydrates obtained from Sigma and Megazyme were between 95-99% pure.
3.2.2 Cloning of the GOOX-encoding gene
S. strictum was grown on 1 g mL-1 food-grade wheat bran at 27°C for 5 days, harvested
by filtration through Miracloth (Calbiochem), and then flash-frozen using liquid nitrogen.
Total RNA was extracted from the ground sample using the RNeasy Plant Mini Kit
(Qiagen). The full-length cDNA encoding the GOOX protein was isolated using the Long
Range 2Step RT-PCR Kit (Qiagen). Briefly, reverse transcription at 42°C for 90 min was
followed by PCR using Pfu DNA polymerase (Agilent Technologies), gene-specific
primers (Lee et al. 2005), and 35 cycles of 93°C for 30 s, annealing at 56°C for 40 s, and
extension at 72°C for 2 min. The PCR product was purified using the QIAquick PCR
Purification Kit (Qiagen), and then sequenced at the Centre for Applied Genomics
(TCAG, the Hospital for Sick Children). The GOOX encoding gene was cloned into the
33
pPICZαA expression vector (Invitrogen), downstream of an α-factor secretion signal,
using EcoRI and XbaI and T4 DNA ligase (Invitrogen).
3.2.3 Site-directed mutagenesis
Chito-oligosaccharide oxidase, ChitO, (accession no.: XP_391174) from Fusarium
graminearum and a carbohydrate oxidase from Microdochium nivale, MnCO, (accession
no.: CAI94231-2) were aligned to GOOX (accession no.: ADI58761) using the Megalign
program (DNASTAR-Lasergene) (Figure S3.1). Amino acids that were predicted to
participate in substrate binding, and that varied between the enzymes analyzed, were
selected for site-directed mutagenesis. Mutations Y300A, Y300N and W351F were
introduced using mutagenic primers (Table 3.1). PCR was performed for 14 cycles of
95°C for 30 s; 55°C for 1 min; and 68°C for 5 min, using the QuikChange method
(Agilent Technologies). The mutations were confirmed by sequencing (TCAG, the
Hospital for Sick Children).
Table 3.1 List of oligonucleotide primers used for gene amplification and site-directed mutagenesis.
Primer Sequence
EX1* GCTTCATGGATCCAGGAATTCAACTCAATCAACGCCTG
EX2* TTCAAGTCTAAATCATCTAGATAGGCAATGGGCTCAAC
Y300A-F CAACACCTACTTGGCCGGTGCTGACC
Y300A-R GGTCAGCACCGGCCAAGTAGGTGTTG
Y300N-F CAACACCTACTTGAACGGTGCTGACC
Y300N-R GGTCAGCACCGTTCAAGTAGGTGTTG
W351F-F GCGGCTGGTTCATCCAATGGGACTTC
W351F-R GAAGTCCCATTGGATGAACCAGCCGC
*From Lee et al. (2005) for gene amplification; others for site-directed mutagenesis.
34
3.2.4 Recombinant protein expression
Mutated plasmids were transformed into Pichia pastoris GS115 according to the
manufacturer’s instructions (Invitrogen, Pichia Expression version G). Transformants
were selected on buffered minimal methanol medium containing histidine (BMMH, 100
mM potassium phosphate, pH 6.0; 1.34 % yeast nitrogen base without amino acids
(YNB); 4 x 10-5 % biotin; 0.5 % methanol, 0.004% histidine), and then screened for
protein expression by immuno-colony blot using nitrocellulose membranes (0.45 µm,
Bio-Rad), anti-Myc antibodies, alkaline phosphatase-linked anti-Rabbit IgG conjugates,
and BCIP/NBT solution. Positive transformants were grown overnight in 100 mL of
buffered minimal glycerol medium containing histidine (BMGH, 100 mM potassium
phosphate, pH 6.0; 1.34 % YNB; 4 x 10-5 % biotin; 1 % glycerol, 0.004% histidine) at
30°C with continuous shaking at 300 rpm. The cells were harvested by centrifugation at
1,500 × g for 10 min and suspended in 300 mL of BMMH medium in 1 L- flasks to
OD600 ~ 1. Cultures were grown at 30°C and 300 rpm for 3 days and 0.5 % methanol was
added every 24 h to induce recombinant protein expression. Levels of recombinant
extracellular protein expressions were monitored every 24 h by activity and SDS-PAGE.
3.2.5 Enzyme purification
Supernatants from methanol-induced cultures of P. pastoris expressing the secreted
recombinant proteins were harvested by centrifugation at 6,000 × g for 10 min and
filtration through 0.22 μm Sterivex filter units (Millipore). Cleared supernatants were
concentrated approximately 150 times using Centricon concentration units (Millipore).
Each recombinant protein was purified using a new Ni-NTA resin (Qiagen). Fractions
35
were eluted with 250 mM imidazole and the buffer was replaced by 40 mM Tris-HCl (pH
8.0) using Vivaspin6 concentration units (GE Healthcare). Protein concentration
measurements were performed using the Pierce BCA assay (Thermo Scientific) and
enzyme purity was verified by SDS-PAGE.
In-gel trypsin digestion with sequencing-grade trypsin (Promega), followed by tandem
mass spectrometry was performed to confirm the identity of each protein sample. Tryptic
fragments were analyzed using the Applied Biosystems/MDS Sciex API QSTAR XL
Pulsar System coupled with an Agilent nano HPLC (1100 series) (The Advanced Protein
Technology Centre, the Hospital for Sick Children). Proteomic data were analyzed using
Scaffold Viewer (www.proteomesoftware.com).
3.2.6 Enzymatic assays and kinetics analyses
A chromogenic assay was used to measure H2O2 production (Lin et al. 1991). Reactions
contained 0.1 mM 4-AA, 1 mM phenol, 0.5 U horseradish peroxidase, 40 mM Tris-HCl
(pH 8.0), and different substrates were initiated by adding 0.2 µg of enzymes to the 250
μL reaction mixture. The production of H2O2 was coupled to the oxidation of 4-AA by
horseradish peroxidase and detected at 500 nm. Reactions were incubated at 37°C for 15
min to measure the specific activity of GOOX on 10 mM of monosaccharide or 1 mM of
oligosaccharide.
Kinetics parameters were determined with a wide range of substrate concentrations: 0.1
mM to 300 mM glucose, 1 mM to 1500 mM xylose, 1 mM to 600 mM galactose, 1 mM
to 600 mM N-acetyl-glucosamine (NAG), 0.1 mM to 300 mM maltose, 5 µM to 1.5 mM
cellobiose, 10 µM to 3.5 mM cellotriose, 20 µM to 40 mM xylobiose, and 20 µM to 50
36
mM xylotriose. At least 12 substrate concentrations were included to obtain kinetics
parameters for each substrate. Initial rates were obtained by measuring reaction products
every 30 s for 15 min at 37°C and pH 8.0, and kinetics parameters were calculated using
the Michaelis-Menten equation (GraphPad Prism5 Software).
The enzyme stability under activity assay conditions was evaluated in triplicate by
incubating 0.6 µg of each enzyme preparation in 40 mM Tris-HCl buffer (pH 8.0) for 0,
5, 15, 25, 35, and 60 min at 37°C. Residual enzyme activity was measured at 37oC for 15
min at pH 8.0 using 10mM maltose and 0.2 µg of protein.
3.2.7 Deglycosylation
Approximately 2 µg of purified enzyme was treated with Peptide -N-Glycosidase F from
Flavobacterium meningosepticum, also known as PNGaseF (New England Biolabs) using
denaturing and native conditions. Samples that were deglycosylated using denaturing
conditions were analysed by SDS PAGE, while samples deglycosylated using native
conditions were used to evaluate the impact of glycosylation on enzyme activity. The
activity of enzymes was measured on 10 mM maltose at 37oC for 15 min. N-
glycosylation was predicted by NetNGlyc (http://www.cbs.dtu.dk/services/NetNGlyc/)
while O-glycosylation was predicted by OGPET (http://ogpet.utep.edu/OGPET/).
3.2.8 Substrate docking
The structural model of GOOX from S. strictum type strain CBS 346.70 expressed in
Pichia was built by our post doctoral fellow co-authored in this work Dr. Vuong based on
the X-ray structure of GOOX from S. strictum strain T1, GOOX-T1 (PDB ID: 2AXR)
using the Swiss-Model Workspace (Arnold et al. 2006). The structures of glucose,
37
cellobiose, cellotriose, xylose, xylobiose, xylotriose and galactose were obtained from the
protein database of The Research Collaboratory for Structural Bioinformatics (PDB ID:
2FVY, 3ENG, 1UYY, X, 1B3W, 1UX7 and 2J1A, respectively). The program
AutodockTools 1.5.2 ran on Python 2.5 (http://autodock.scripps.edu/) was used to prepare
the oligosaccharides and the enzyme for docking. All hydrogen atoms were added and the
non-polar hydrogens were merged for all ligands and protein. A number of degrees of
torsions of each oligosaccharide were set up to evaluate different thermodynamic
properties. A Lamarckian genetic algorithm (Morris et al. 1998) with different number of
energy evaluations and a population size of 150 individuals were applied for docking.
The program, Autogrid 4, which pre-calculates grip maps of interaction energies, was
used to prepare the grid files, and then docking simulation was performed by Autodock 4
(http://autodock.scripps.edu/). After docking, free energies of binding ∆Gb and
dissociation constants Kd were reported.
3.2.8 Nucleotide sequence accession number
The cloned gene encoding GOOX from Sarocladium strictum type strain CBS 346.70 has
been deposited in the GenBank database under accession number GU369974.
3.3 Results and Discussion
3.3.1 Variations of GOOX
The GOOX gene cloned from Sarocladium strictum type strain CBS 346.70 encoded a
mature protein containing 474 amino acids, which is the same length as a previously
reported GOOX isolated from S. strictum strain T1 (hereafter GOOX-T1) (Lin et al.
1991; Lee et al. 2005). However, there were 15 amino acid substitutions between the two
38
proteins, 13 resulting from differences in corresponding wild-type gene sequences, and 2
(A38V and S388N) resulting from random mutations introduced during the construction
of the expression system (Table 3.2). The GOOX obtained in this study shares 97%
sequence identity with the reported GOOX-T1 (Lin et al. 1991), and it has a similar fold
to GOOX-T1.
Table 3.2 Amino acid substitutions in GOOX in comparison with GOOX-T1
No. Amino acid position
Amino acid in On protein surface*
Distance to sugar O1 (Å)
GOOX GOOX-T1 a
1 23 E D Yes 28.0
2 38 V A No 29.7
3 99 D N Yes 33.7
4 126 T S No 14.3
5 135 I V No 15.0
6 159 V I Yes 24.4
7 175 K E Yes 25.1
8 235 E Q No 22.2
9 259 Y F No 18.3
10 269 V I No 23.2
11 332 S Q No 26.7
12 366 S A Yes 21.2
13 367 H V Yes 20.5
14 388 N S No 11.3
15 435 D T Yes 24.1
*Determined by 20% accessible surface area a reported by Lin et al. (1991)
39
3.3.2 Production of recombinant protein
The recombinant expression of GOOX in P. pastoris GS115 was highest after three days
of incubation with 0.5 % methanol. Proteins were purified to more than 95 %
homogeneity by affinity chromatography; similar to previous reports of recombinant
GOOX-T1 expression by P. pastoris (Huang et al. 2008). Approximately 1.5 mg L-1 of
purified GOOX was recovered, and after confirming that one freeze-thaw cycle did not
affect enzyme activity, the purified enzyme was stored as 20 µL aliquots (~ 4 µg) at -
80°C. The enzyme remained fully active following pre-incubation at 37°C for 60 min
(Figure S3.2).
The deduced molecular mass of the mature protein with a c-myc epitope and a
polyhistidine tag is approximately 56 kDa (Protean, DNASTAR-Lasergene), which is
less than the electrophoretic molecular weight of purified GOOX (~70 kDa) (Figure
S3.3). By comparison, the reported molecular weight of GOOX-T1 determined by size
exclusion chromatography is approximately 61 kDa (Lee et al. 2005). Recombinant
proteins expressed in P. pastoris GS115 can be N-glycosylated with high-mannose-type
structures containing 8 to 14 mannose residues (Hirose et al. 2002; Blanchard et al.
2007). And NetNGlyc predicted three N-glycosylation sites in GOOX, including N305,
N341, and N394, which are all located in exposed loop regions. Still, the molecular
weight of deglycosylated GOOX was ~60 kDa, (Figure S3.3), suggesting that other post-
translational modifications, including O-glycosylation and/or phosphorylation, probably
occurred (Cereghino and Cregg 2000; Letourneur et al. 2001; Boraston et al. 2003).
Notably, deglycosylation of GOOX under native conditions did not cause a detectable
loss in enzyme activity (Table 3.3).
40
Table 3.3 The effect of deglycosylation with PNGaseF on enzyme activity
Enzyme Activity (nmol min-1)*
Glycosylated Deglycosylated
GOOX-VN 3.5 3.4
W351F 3.1 3.2
Y300A 4.1 3.9
Y300N 2.6 2.8
*Enzyme activity was measured in duplicate with 10 mM maltose following 15 min at 37oC.
3.3.3 Novel substrate specificity
GOOX oxidase activity was evaluated using glucose, xylose, galactose, NAG, mannose,
and arabinose. Glucose, xylose, galactose, and NAG were oxidized by the recombinant
GOOX, and the highest catalytic efficiency was observed using glucose (Table 3.4).
Previous analyses of GOOX-T1 did not detect activity on xylose, galactose or NAG, and
activity was limited to glucose and oligosaccharides with reducing end-glucosyl residues
(Lin et al. 1991; Fan et al. 2000). To check whether GOOX can oxidize oligomers of C5
sugars, the enzyme was then tested for oxidation of xylo-oligosaccharides. GOOX
oxidized xylo-oligosaccharides as efficiently as cello-oligosaccharides (Table 3.4), and
the catalytic efficiency of GOOX on these oligosaccharides was over two orders of
magnitude higher than that of the corresponding monomers. These findings show that
GOOX has broader substrate specificity than GOOX-T1, and GOOX oxidizes C6 and C5
mono- and oligomeric sugars.
41
Table 3.4 Kinetics parameters of wild-type and mutant GOOX enzymes.
Substrate Enzyme kcat
(min-1) Km
(mM) kcat/Km
d (mM-1min-1)
Specific activitya (µmol mg-1min-1)
From Vmax Defined substrate
concentrationb Glucose GOOX 449 ± 6c 17.4 ± 0.7 25.9 ± 1.1 7.4 2.6 Glucose W351F 337 ± 5 31.0 ± 1.3 10.9 ± 0.5 5.6 1.3 Glucose Y300A 793 ± 14 8.1 ± 0.4 98 ± 5 13.1 7.2 Glucose Y300N 649 ± 6 3.11 ± 0.12 209 ± 8 10.7 8.3 Xylose GOOX 315 ± 9 105 ± 10 3.0 ± 0.3 5.2 0.4 Xylose W351F 277 ± 4 288 ± 10 0.96 ± 0.04 4.6 0.2 Xylose Y300A 680 ± 8 51.8 ± 1.9 13.1 ± 0.5 11.2 1.6 Xylose Y300N 700 ± 10 32.0 ± 1.7 21.7 ± 1.2 11.5 2.3
Galactose GOOX 429 ± 8 132 ± 7 3.3 ± 0.2 7.1 0.5 Galactose W351F 394 ± 4 36.1 ± 1.3 10.9 ± 0.4 6.5 1.3 Galactose Y300A 798 ± 15 96 ± 5 8.3 ± 0.5 13.2 1.2 Galactose Y300N 705 ± 5 110 ± 2 6.4 ± 0.1 11.6 1.0
NAGe GOOX 485 ± 12 340 ± 17 1.4 ± 0.1 8.0 0.2 NAG W351F 470 ± 40 950 ± 120 0.5 ± 0.1 7.7 0.1 NAG Y300A 768 ± 13 92 ± 5 8.3 ± 0.4 12.7 1.1 NAG Y300N 680 ± 7 56.0 ± 1.9 12.2 ± 0.4 11.2 1.5
Mannose GOOX ND ND ND ND ND Mannose W351F ND ND ND ND ND Mannose Y300A ND ND ND ND 0.1 Mannose Y300N ND ND ND ND 0.2 Maltose GOOX 360 ± 5 2.81 ± 0.16 128 ± 8 5.9 1.5 Maltose W351F 323 ± 4 5.0 ± 0.2 65 ± 3 5.3 0.8 Maltose Y300A 625 ± 7 11.0 ± 0.5 57 ± 3 10.3 0.8 Maltose Y300N 624 ± 6 19.6 ± 0.7 31.8 ± 1.1 10.3 0.5
Cellobiose GOOX 375 ± 11 0.07 ± 0.01 6000 ± 800 6.2 5.4 Cellobiose W351F 344 ± 5 0.083 ± 0.005 4100 ± 60 5.7 5.1 Cellobiose Y300A 823 ± 16 0.25 ± 0.02 3400 ± 300 13.6 10.7 Cellobiose Y300N 684 ± 11 0.38 ± 0.02 1800 ± 100 11.3 8.2 Cellotriose GOOX 361 ± 12 0.085 ± 0.010 4200 ± 500 6.0 5.1 Cellotriose W351F 315 ± 6 0.11 ± 0.01 2900 ± 300 5.2 4.6 Cellotriose Y300A 670 ± 10 0.25 ± 0.02 2700 ± 200 11.0 9.0 Cellotriose Y300N 599 ± 5 0.44 ± 0.01 1400 ± 30 9.9 6.9 Xylobiose GOOX 529 ± 5 0.098 ± 0.004 5400 ± 50 8.7 7.9
42
Table 3.4 (Continued) Kinetics parameters of wild-type and mutant GOOX enzymes.
Substrate Enzyme kcat
(min-1) Km
(mM) kcat/Km
d (mM-1min-1)
Specific activitya (µmol mg-1min-1)
From Vmax Defined substrate
concentrationb Xylobiose W351F 478 ± 4 0.35 ± 0.01 1400 ± 40 7.9 5.9 Xylobiose Y300A 797 ± 7 5.11 ± 0.15 156 ± 5 13.2 2.1 Xylobiose Y300N 718 ± 8 4.83 ± 0.17 149 ± 6 11.8 1.8 Xylotriose GOOX 498 ± 7 0.10 ± 0.01 5100 ± 500 8.2 7.5 Xylotriose W351F 473 ± 4 0.31 ± 0.01 1500 ± 50 7.8 6.0 Xylotriose Y300A 832 ± 7 3.15 ± 0.11 260 ± 10 13.7 3.2 Xylotriose Y300N 718 ± 11 4.3 ± 0.2 170 ± 10 11.8 2.0
a 0.2 µg of enzyme was used in each reaction. b Reactions contained either 10 mM of monosaccharide or 1 mM of oligosaccharide substrate; ND- not detected. c Standard deviations (n=3). d Standard deviations (SD) for kcat /Km values were calculated using following formula: SD (kcat /Km) = kcat / Km *(SQRT((SD(Km)/Km)^2 + (SD(kcat)/kcat)^2)). e NAG, N-acetylglucosamine.
The broader substrate range of GOOX detected in the current study compared to previous
reports using GOOX-T1 could be the result of different assay conditions, in particular the
enzyme concentration. While reactions for kinetics analyses of GOOX proceeded for up
to 15 min and included substrate concentrations over 500 mM, oxidation of xylose,
galactose and NAG by GOOX was detected after 3 min using 10 mM of each sugar,
which were the reaction conditions previously used to screen GOOX-T1 activity (Lin et
al. 1991). However, the enzyme concentration was not specified in Lin et al. (1991), and
so it is possible that lower enzyme dose was used in that study. Furthermore, earlier
substrate screens were performed using the native GOOX-T1 (Lin et al. 1991) while in
the current study the recombinant GOOX was used to measure specific activities. This is
important in light of the fact that different kcat values were obtained by native and the
43
recombinant GOOX-T1 on maltose (531 min-1 and 361 min-1, respectively) (Lin et al.
1991; Lee et al. 2005), and the discrapency was explained by the various processing
methods for proteins in P. pastoris and S. strictum (Lee et al. 2005).
Alternatively, novel substrate specificity of GOOX could be due to amino acid
substitutions in this enzyme. Most substitutions are located on the protein surface or far
from the oxidation site (Table 3.2); however, N388 is positioned on the same β16-sheet
as conserved residues Q384 and Y386, which are predicted to participate in substrate
binding (Huang et al. 2005). The side chain of N388 is located near the predicted -2
subsite, within 6.2 Å from the substrate. When comparing the X-ray structures of
precursor and mature galactose oxidase from Fusarium spp., Firbank et al. (2001) showed
that the Cα of Tyr290 moved by 6.3 Å and the loop containing this residue could shift up
to 8 Å (Firbank et al. 2001).
Figure 3.2 Conformational changes of S388 upon substrate binding. (A) X-ray structure of GOOX-T1 (PDB ID: 1ZR6); the backbone of S388 formed two H-bonds (dashed lines) to that of G349. (B) X-ray structure of GOOX-T1 in the presence of a substrate analog (PDB ID: 2AXR); the side chain of S388 formed a weak H-bond to the G349 backbone while the side chain of D387 H-bonded to the backbone of S388. (C) The position of N388 in the structural model of GOOX.
44
While general loop movement was not observed when comparing GOOX-T1 structures
before and after inhibitor binding, the side chain of S388 in GOOX-T1 turned
significantly upon substrate binding to form a weak H-bond with the G349 backbone of
the β15-sheet (Figure 3.2) (Huang et al. 2005). Accordingly, the beneficial effect of the
S388N substitution on GOOX activity might be due to the potential of Asn to stabilize
substrates that contain fewer hydroxyl groups and/or to stabilize the β16-sheet for
substrate binding.
Docking analysis determined that the computational Kd for xylose was two times higher
than that for glucose, suggesting that low activity on xylose, which does not possess an
exocyclic CH2OH, might be due to weak binding of this substrate by GOOX (Table 3.5).
Table 3.5 Docking parameters of oligosaccharides with GOOX enzymes.
GOOX Y300A Y300N
Docked
energy ∆Gb
(kcal/mol)
Kd
(µM)
Distance to
Y429
Oη(Å)*
Docked
energy ∆Gb
(kcal/mol)
Kd
(µM)
Distance to
Y429 Oη(Å)
Docked
energy ∆Gb
(kcal/mol)
Kd
(µM)
Distance to
Y429
Oη(Å)
Glucose -5.2 150 2.8 -5.0 210 2.9 -5.1 190 2.8
Cellobiose -6.8 10 2.9 -6.1 35 2.7 -6.4 20 2.9
Cellotriose -6.8 10 2.8 -6.6 14 2.8 -6.7 12 2.7
Xylose -4.7 360 3.1 -4.6 430 2.9 -4.6 410 3.2
Xylobiose -6.2 27 2.6 -6.0 39 2.8 -5.9 45 3.2
Xylotriose -7.5 3.2 3.0 -5.7 63 2.9 -5.9 50 3.2
Galactose -5.1 190 2.6 -4.9 250 2.8 -5.0 230 2.9
45
*Distance between the Oη atom of Y429 and the O1 atom of oligosaccharides. This distance is 2.8 Å in the crystal structure of GOOX-T1 and an inhibitor (PDB ID: 2AXR).
The Km values for di- and tri-saccharides obtained experimentally, as well as the
corresponding Kd values derived from the docking models, are an order of magnitude
lower than the Km and Kd values for monosaccharides (Table 3.4, Table 3.5). These
results support the presence of two glycosyl-binding subsites in the carbohydrate-binding
groove of GOOX, which was also predicted by the X-ray structure of GOOX-T1 (Huang
et al. 2005).
3.3.4 Improvement of substrate specificity
The catalytic activity of GOOX on monosaccharides and oligosaccharides was further
improved through site-directed mutagenesis. Amino acids targeted for this analysis were
chosen by 1) referencing the published structure of GOOX-T1 (Huang et al. 2005), and 2)
identifying amino acids in GOOX that participate in substrate-binding, which consistently
differ from corresponding residues in ChitO from F. graminearum and MnCO from M.
nivale.
Y300 and W351 are located at the -2 glucosyl-binding subsite (Figure 3.1B), and likely
stabilize oligosaccharide binding through stacking interactions. Y300 is substituted by
alanine in ChitO and asparagine in MnCO while W351 is substituted by phenylalanine in
MnCO. Since MnCO is distinguished by its activity on galactose, xylose and to some
extent on mannose (Xu et al. 2001), altering the polarity and/or size of Y300 and W351
could increase the activity of GOOX on sugars with an axial OH4 group or that lack an
exocyclic CH2OH group. Accordingly, Y300N, Y300A and W351F substitutions were
46
generated in GOOX, and 3 mg L-1, 4 mg L-1 and 1.3 mg L-1 of each purified protein was
recovered, respectively. The mutant enzymes remained active after a one hour- pre-
incubation at 37°C (Figure S3.2).
The catalytic activity (kcat) of Y300A and Y300N mutant enzymes on all tested
monosaccharides and oligosaccharides was approximately two times higher than that of
GOOX (Table 3.4). These two mutant enzymes also gained low activity on mannose
(Table 3.4). However, the loss in hydrophobic interactions at the -2 subsite also increased
the Km and Kd values for oligosaccharides, reducing overall catalytic efficiency. These
results suggest that Y300 affects substrate positioning relative to the catalytic Y429
residue and the FAD cofactor, and that Y300 contributes to stacking interactions with
substrates containing more than two units.
The W351F mutation slightly reduced the catalytic activity of GOOX on all substrates.
Like Y300A and Y300N mutations, the W351F mutation also increased the Km values of
GOOX with oligomeric substrates (Table 3.4). These results are consistent with both
Y300 and W351 participating in stabilizing stacking interactions with penultimate
reducing sugars of oligomeric substrates, which also explains why the impact of these
mutations on Km is similar with di- and tri-saccharides (Table 3.4). Notably, the W351F
mutation also increased the Km values of GOOX with glucose and xylose, but decreased
the Km of GOOX with galactose, resulting in higher catalytic efficiency with this
substrate (Table 3.4).
47
Figure 3.3 Docking of monosaccharides to GOOX. Docking positions of glucose (A), xylose (B) and galactose (C); and the side chains of Y300 and W351 were shown. The O4 atom of galactose (circled) pointed to the benzene ring of W351, and their distance was 3.1Å.
Docking studies showed that while glucose and xylose binding at the active-site was not
restricted, the axial OH4 group of galactose points directly towards the benzene ring of
tryptophan (Figure 3.3), suggesting that the indole structure hinders GOOX binding of
sugars with axial OH4 groups.
3.4 Conclusions
This study demonstrates that GOOX with different substrate specificity were produced by
different strains of S. strictum, widening the application of GOOX from S. strictum for
the oxidation of mono- and oligo-saccharides. In addition to glucose, maltose and cello-
oligosaccharides, the new GOOX oxidized xylo-oligosaccharides, galactose, and N-
acetylglucosamine, which were not detected in GOOX-T1 from previous studies. Y300A
and Y300N substitutions increased the catalytic activity of GOOX on all substrates, and
gained low activity on mannose. Rational engineering approaches are now being applied
to decrease the Km of GOOX and its mutant enzymes on oligomeric substrates. In
particular, given the consistency between computational docking analyses and
experimental data reported in the current study, docking analyses will be used to predict
48
the effect of selected amino acid substitutions on the binding affinity, conformation, and
orientation of substrates bound by GOOX and variant enzymes. It is anticipated that
resulting carbohydrate oxidases will constitute important tools for the production of new
materials from plant fibre.
3.5 A followed-up study on the structure-function analysis of GOOX
Given the interesting outcomes of the site-directed mutagenesis work presented in this
chapter, a followed up study was designed to further unravel the role of amino acid
residues within the GOOX substrate binding site (Vuong et al. 2013). Since this analysis
was not a main part of my PhD thesis, I will only briefly summarize the main highlights
here.
The additional amino acids targeted for single mutation included Y72 and Q384 located
near the -1 substrate binding site, E247 and W351 located near the -2 substrate binding
site, and Q353, which is positioned between these two subsites (Figure 3.4). While most
substitutions were to alanine, some were mutated to related residues.
All mutations resulted in higher Km values than the wild-type GOOX, confirming the
important role of the targeted residues for substrate binding by GOOX. Notably, Y72 and
Q353 mutants lost oxidation activity on monosaccharides and the latter also lost activity
on xylooligosaccharides, suggesting the role of Q353 for substrate preference of this
enzyme.
49
Figure 3.4 The biding site for GOOX-T1. Residues for mutation in relation to the substrate analog, 5-amino-5-deoxy-cellobiono-1,5-lactam (ABL); the movement of residues in the absence of ABL (PDB ID: 1ZR6, green) compared with the presence of ABL (PDB ID: 2AXR, cyan) are indicated by arrows.
Moreover, it was confirmed that the two random mutations on S388 and A38 do not
explain the differences in substrate preference of GOOX and GOOX-T1. Yet, the
destabilizing effect of S388N proposed in this study was validated, as the back mutation
of N388S resulted in a more thermostable mutant consistent with GOOX-T1
thermostability. In addition, it was shown that the Y300A mutants generated by the
current study exhibits reduced substrate inhibition; this characteristic was also shown for
alanine substitution of W351 residue, suggesting these mutants as ideal candidates for
oxidizing oligosaccharides when present at high substrate concentrations (Vuong et al.
2013).
50
Chapter 4 Enhanced binding and activity of GOOX towards polysaccharides
through CBM fusions
Parts of this chapter are submitted in:
Foumani M, Vuong T.V, MacCormick B, Master E.R. 2015. Enhanced polysaccharide
binding and activity on linear β-glucans through addition of carbohydrate-binding
modules to either terminus of a glucooligosaccharide oxidase. PLOS ONE J.
Contributions: Design of the study; performing the experiments, data collection and
analyses corresponding to construction of CBM fusion genes, recombinant protein
expression and purifications, cellulose binding and affinity gel electrophoresis
experiments, specific activity measurements, kinetics studies, and thermal stability
assays; as well as manuscript preparation.
51
4.1 Introduction
Carbohydrate oxidases can facilitate regio-selective oxidation of sugars and
polysaccharides, and were recently categorized as auxiliary activities (AA) in the CAZy
(Levasseur et al. 2013). Among these enzymes, oligosaccharide oxidases from family
AA7 are comparatively less well characterized. The corresponding carbohydrate oxidases
target the C1 hydroxyl of a broad range of oligosaccharides, including cello-, -xylo-,
chito- and malto-oligosaccharides. Examples of characterized AA7 enzyme include a
gluco-oligosaccharide oxidase (EC 1.1.3) from Sarocladium strictum T1 (GOOX-T1)
(Lee et al. 2005) and a gluco-oligosaccharide oxidase from Sarocladium strictum CBS
346.70 (GOOX), which was discussed in previous chapter (Foumani et al. 2011). More
examples have been reviewed in section 2.2. These flavoenzymes are likely to share a
conserved FAD-binding domain, while having different substrate binding domains.
GOOX-T1 and GOOX were previously shown to function best at 37 °C and pH 8, and
remain stable up to 50 °C and 45 °C, respectively (Lin et al. 1991; Fan et al. 2000;
Foumani et al. 2011). Both enzymes oxidize maltose, lactose and cello-oligosaccharides
(Lee et al. 2005; Foumani et al. 2011); GOOX was later shown to also oxidize xylo-
oligosaccharides and xylan (Vuong et al. 2013; Vuong and Master 2014). Given the
precedence for GOOX activity on oligomers of glucose, the aim of the current study was
to evaluate the potential of selected CBMs to increase GOOX activity on plant
polysaccharides, particularly β-glucans.
In a recent study, Telke et al. (2012) found that fusing CBM3, CBM4 or CBM30 to a
family GH9 endoglucanase increased enzyme activity on different cellulose preparations
52
by roughly 10-fold. Similarly, Voutilainen et al. (2014) attached various cellulose
binding modules from both microbial and fungal origins to a fungal family GH7
cellobiohydrolase. In all cases, the CBM addition increased enzyme binding to cellulose
as well as thermostability, where the addition of a bacterial CBM3 from Clostridium
thermocellum (CipA) led to highest activity gains. Fusions of a xylan specific CBM from
family 22 to a xylanase from Bacillus halodurans also showed increased activity on
insoluble xylan while thermostability of the fusion enzyme was reduced (Mamo et al.
2007). Moreover, a xylan-binding module from family CBM22A was recently appended
to the N terminus of GOOX, and the resulting fusion protein retained activity after
immobilization to xylan-coated surfaces (Vuong and Master 2014).
In this study, we evaluate the potential of selected CBMs to increase the binding capacity
and activity of GOOX towards insoluble celluloses and β-glucans presented at low
concentrations. Specifically, three characterized CBMs were fused to either the N-
terminus or C-terminus of GOOX, namely 1) the Type-A CtCBM3 from C. thermocellum
CipA, which can bind cellulose (Lehtiö et al. 2003; Wan et al. 2011), 2) the Type-B
CtCBM11 from C. thermocellum Lic26A-Cel5E, which can bind β-(1→3), (1→4)-glucan
(Carvalho et al. 2004), and 3) the Type-B CtCBM44 from C. thermocellum Cel9D-
Cel44A, which can bind xyloglucan, glucomannan and β-(1→3), (1→4)-glucan
(Najmudin et al. 2006). These CBMs were chosen for this analysis since corresponding
binding affinities have been characterized; being sourced from a thermophilic organism
also presented possibility to confer thermostability to GOOX as previously observed for
cellulolytic enzymes (Telke et al. 2012; Voutilainen 2014). Although the number of CBM
fusion studies has been increased in recent years, we believe this work is unique in terms
53
of wide selection of CBMs appended to both terminus of a single domain non-hydrolytic
enzyme via several types of linkers. Notably, to our knowledge this work is among the
first studies examining the potential of CBMs from family 11 and 44 in fusion constructs
to enhance the behaviour of a catalytic module other than their native cognates.
4.2 Materials and Methods
4.2.1 Materials
All chemicals were reagent grade with high purity, and purchased from Sigma (Canada)
unless otherwise specified. Cellobiose (99% pure) was purchased from BioShop Inc.
(Canada). Cello-oligosaccharides, barley β-glucan, konjac glucomannan, xyloglucan from
tamarind seed, carboxymethyl cellulose, and hydroxyethyl cellulose, all with min 95%
purity, were purchased from Megazyme (Ireland). Mixed xylo-oligosaccharides (DP 2-7)
was obtained from Cascade Biochemicals (USA). Nanocrystalline cellulose was a kind
gift from Dr. Y. Boluk (University of Alberta). Regenerated amorphous cellulose was
produced from Sigmacell cellulose Type 20 after the cellulose was wetted with water,
dissolved in phosphoric acid, and regenerated in water as described previously (Zhang et
al. 2006). The CBM encoding genes from Clostridium thermocellum, CtCBM3,
CtCBM11, and CtCBM44 were purchased from Nzytech (Portugal). The same genes
were codon optimized for expression in Pichia pastoris and were synthesized by DNA
2.0 (USA).
4.2.2 Construction of fusion enzymes
To construct C-terminal CBM fusion proteins, genes encoding CtCBM3, CtCBM11 and
CtCBM44 with native N-terminal linkers (CtCBM3: PTNTPTNTPTNTP, CtCBM44:
54
PPPY) or no linker sequence (CtCBM11) were separately cloned into the XbaI restriction
site of pPICZαA-GOOX, a plasmid for recombinant expression of GOOX in P. pastoris
(Foumani et al. 2011). Three genes were synthesized by DNA 2.0 to produce
corresponding N-terminal CBM fusion proteins. In this case, a DNA sequence encoding
a TP linker (SRGGGTATPTPTPTPTP) was inserted between genes encoding CtCBM3,
CtCBM11 or CtCBM44, and the gene encoding GOOX (Figure 4.1). All plasmid
constructs were sequenced at the Center for Applied Genomics (the Hospital for Sick
Children, Toronto, Canada) before being transferred to P. pastoris for recombinant
protein production.
4.2.3 Recombinant expression of fusion proteins in Pichia pastoris
All plasmids were transformed into P. pastoris GS115 according to the manufacturer's
instructions (Invitrogen, Pichia Expression version G). P. pastoris transformants were
selected on buffered minimal methanol medium containing histidine (BMMH, 100 mM
potassium phosphate pH 6.0; 1.34 % YNB; 4×10−5 % biotin; 0.5 % methanol, 0.004%
histidine), and then screened for protein expression using an overlay activity assay.
Briefly, the assay mixture (0.3% agarose, 2% cellobiose, 50 mM Tris-HCl pH 8.0, 2 mM
phenol, 0.4 mM 4-AA, and 15 U/mL horseradish peroxidase) was maintained in a 40°C
water bath, and 10 mL of the solution was gently poured on top of each BMMH plate
containing the transformant colonies. After solidifying at room temperature for 15 min,
plates were transferred to 37°C for 60 min to induce the chromogenic reaction between 4-
AA and H2O2. Transformants with highest activity were then selected for liquid
cultivation.
55
Selected P. pastoris transformants were grown overnight in 100 mL of buffered minimal
glycerol medium (BMGY, 1% yeast extract; 2% peptone; 100 mM potassium phosphate
pH 6.0; 1.34% YNB; 4×10−5 % biotin; 1% glycerol) at 30 °C with continuous shaking at
250 rpm. The cells were harvested by centrifugation at 1,500 × g for 10 min and
suspended in 200 mL of BMMH medium supplemented with 1% casamino acid in 1-L
flasks to OD600 ∼2. Cultures were grown at 27 °C and 250 rpm for 4 days and 0.5%
methanol was added every 24 h to induce recombinant protein expression. To minimize
proteolysis of the secreted recombinant protein, 2 µM leupeptin was added to the culture
medium every 24 h.
4.2.4 Purification of recombinant enzymes
Culture supernatants containing the recombinant protein were harvested by centrifugation
at 6,000 × g for 10 min and filtered through 0.22 µm PES filter membrane (GE water and
process technologies, USA). The culture supernatants were concentrated and buffer
exchanged into binding buffer (100 mM potassium phosphate pH 8, 300 mM NaCl, and 5
mM imidazole) using a Jumbosep centrifugal device (Pall Corp, USA). Resulting
concentrates were incubated separately with Ni-NTA resin (Qiagen, Germany) and eluted
with 250 mM imidazole; the protein solution was exchanged to 40 mM Tris-HCl pH 8
using Vivaspin 20 concentration units (GE healthcare, UK). Protein concentrations were
measured using the Pierce BCA assay (Thermo Scientific, Canada) and confirmed using
SDS-PAGE densitometry, where the band density of purified protein and a dilution series
of bovine serum albumin (BSA) were determined using ImageJ (http://rsbweb.nih.gov/ij/)
(Schneider et al. 2012).
56
4.2.5 Specific activity on oligosaccharides, soluble polysaccharides and insoluble
cellulose substrates
A chromogenic assay was used to detect and measure H2O2 production (Lin et al. 1991).
To measure activity on oligosaccharides, reactions contained 3 pmol of enzyme, 0.1 mM
4-AA, 1 mM phenol, 0.5 U horseradish peroxidase, 50 mM Tris–HCl (pH 8.0), and 0.1
mM of each substrate; 20 mM CaCl2 was also included in reaction mixtures as CtCBMs
from family 3, 11 and 44 show conserved calcium binding sites (Najmudin et al. 2006;
Viegas et al. 2008; Yaniv et al. 2011). Reactions without enzyme served as negative
controls. In all cases, the final reaction volume was 250 µL. The production of H2O2 was
coupled to the reduction of 4-AA by horseradish peroxidase and detected at 500 nm.
One unit of the GOOX activity corresponded to the formation of 1 µmol of the product
per min. Reactions were incubated at 37°C for 10 min and absorbance was measured
every 1 min. H2O2 (0-0.04 mM) was used to generate a standard curve. All experiments
were performed as triplicates.
To determine kinetics parameters on cellotetraose, initial rates of reactions were
measured using the above assay with eight substrates concentrations from 0.01-1 mM.
Kinetics parameters were then calculated using the Michaelis–Menten equation
(GraphPad Prism5 Software).
The standard reaction described above was also used to measure enzyme activity on
soluble polysaccharides. Since GOOX activity significantly differed on each test
polysaccharide, enzyme and substrate concentrations were adjusted so that activity data
could be collected within the linear range of the standard curve for the assay.
57
Accordingly, between 7.7 pmol and 30 pmol (∼ 0.5-2 µg) of enzyme was added to the
assay mixture, and reactions contained 0.1% to 0.5 % (w/v) of test polysaccharides.
Specifically, reactions containing 7.7 pmol of enzyme were with 0.5% xyloglucan,
whereas 30 pmol of enzyme were used with 0.1% konjac glucomannan, 0.3% barley β-
glucan, 0.3% carboxymethyl cellulose, and 0.5% hydroxyethyl cellulose. All reactions
were incubated at 37°C for 30 min and the absorbance was measured every 5 min.
Similarly, to measure enzyme activity on insoluble substrates, between 7.7 pmol and 30
pmol of enzyme was added to the standard assay mixture, and reactions contained 0.2%
and 0.5 % of test polysaccharides. Specifically, reactions containing 7.7 pmol of enzyme
were with 0.5% of microcrystalline cellulose (Avicel pH-101), 0.5% of nanocrystalline
cellulose, and 0.5% of oat spelt xylan, whereas 30 pmol of enzyme were used with 0.2 %
of regenerated amorphous cellulose. All reactions were incubated with mixing (500 rpm)
at 37°C for up to 24 h using an Eppendorf thermomixer equipped with an adaptor for 96-
well plates. To avoid evaporation, microplates were sealed using an adhesive sheet. For
each time point (0, 2, 4, 6, and 24 h) the entire 250 µL reaction was collected, centrifuged
to precipitate the insoluble fraction, and then 150 µL of the supernatant was used to read
absorbance at 500 nm.
4.2.6 Cellulose binding
The binding of wild-type GOOX and CBM fusions to microcrystalline cellulose (Avicel
pH-101) and regenerated amorphous cellulose was determined by mixing 10 µg of
enzyme with 0.5 mg of the cellulose sample in 250 µL of a buffer solution (20 mM CaCl2,
0.05% Tween 20, 50 mM Tris-HCl pH 8). Following incubation for 2 h at 4˚C with
58
vigorous shaking at 1,400 rpm, mixtures were centrifuged to recover supernatants
containing the unbound protein fraction and pellets containing the bound protein fraction.
Supernatant samples were concentrated to 20 µL by vacuum centrifugation while pellets
were washed three times with the buffer solution before being extracted for 10 min at
100˚C with 20 µL of a denaturing solution (10% SDS and 10% β-mercaptoethanol). The
bound and unbound fractions were then analyzed by SDS-PAGE and the proteins were
visualized by Coomassie blue staining.
4.2.7 Quartz crystal microbalance with dissipation (QCM-D)
QCM-D experiments were performed by the co-authors Dr. Vuong and Ben MacCromick
with cellulose-coated sensors (QSX 334, Q-Sense, Sweden) using the Q-Sense E4
instrument (Q-Sense, Sweden). Briefly, this instrument detects adsorption of materials to
the sensor surface by measuring changes in oscillation frequency and dissipation values
dictated by the mass and viscosity of the bound material (Vuong and Master 2014). The
flow rate was kept constant at 0.05 mL/min and the temperature was maintained at 25 °C.
The changes in areal mass (ng/cm2) were obtained using the Voigt model of the Q-Tools
software (Q-sense, Sweden). All enzyme and substrate solutions were prepared in a
reaction buffer of 50 mM Tris-HCl pH 8.0. The sensors were equilibrated with 50 mM
Tris-HCl pH 8.0 for approximately 16 h, and then 1.5 µg/mL of CtCBM3_GOOX or the
wild-type GOOX was flowed over the coated sensors until the frequency and dissipation
values stabilized. The protein solutions were then replaced by the equilibration buffer to
rinse away unbound materials, and after washing, the equilibration buffer was replaced
by 0.5 mM cellobiose and the flow-through was collected. Following 40 min of reaction
between bound GOOX enzymes and cellobiose, corresponding quartz sensors were
59
removed from the QCM-D and washed 3 times with the equilibration buffer before being
incubated for 24 h with 800 µL of 0.5 mM cellobiose. In this way, reaction products
were allowed to accumulate in the reaction mixture, which facilitated product detection.
The cycle wash and batch incubation of GOOX-immobilized sensors with 0.5 mM
cellobiose was repeated 3 times. The presence of H2O2 in the flow-through from the
QCM-D, as well as in the sensor incubation solution, was detected using the standard
chromogenic assay.
4.2.8 Affinity gel electrophoresis
Binding of GOOX and fusion constructs to konjac glucomannan, barley β-glucan,
xyloglucan from tamarind seed, and carboxymethyl cellulose was examined by native
affinity gel electrophoresis as described by Freelove et al. (2001) with minor
modifications. Briefly, the native polyacrylamide gels prepared for these analyses
contained 7.5 % (w/v) acrylamide in 25 mM Tris, 250 mM glycine buffer (pH 8.3), and
0.01 % of the test polysaccharide. Approximately 5 µg of GOOX and each fusion
construct were loaded onto the gels and then run at 90 V for 2 h at room temperature.
Relative binding affinities were inferred from the migration distance of the fusion
proteins and wild-type GOOX on gels with and without the test polysaccharides. BSA (5
µg) was also used as a reference protein for these analyses.
4.2.9 Temperature stability
To investigate the potential of each CBM to increase the temperature stability of GOOX,
0.5 µg of each fusion protein or wild-type GOOX was incubated for up to 4 h at 45°C,
before being cooled to room temperature to measure residual enzyme activity using the
60
GOOX standard assay and 1 mM cellobiose as substrate. All experiments were performed
in triplicate. The half-life was measured by plotting the logarithm of percent remaining
activity versus the incubation time using Microsoft Excel (v 14.1.4).
4.2.10 Nucleotide sequence accession number
The genes encoding N-terminal CtCBM3_GOOX, CtCBM11_GOOX,
CtCBM44_GOOX and C-terminal GOOX_CtCBM3, GOOX_CtCBM11, and
GOOX_CtCBM44 have been deposited in the GenBank database under accession
numbers: JX181765, JX181766, JX181767, JX181768, JX181769, and JX181770,
respectively.
4.3 Results and Discussion
4.3.1 Recombinant protein production
When using the standard Pichia expression protocol to produce N-terminal CBM fusions,
degradation products that corresponded to the size of the wild-type GOOX and CBMs
separately were observed by SDS-PAGE. This observation implied proteolysis of the N-
terminal fusion proteins at the linker site. P. pastoris is expected to release extracellular
serine, cysteine and asparatic proteases (Shi et al. 2003). The activity of these proteases
depends on the pH, where serine protease activity is highest above pH 7, cysteine-type
protease activity is highest between pH 5-7, and aspartic protease activity is highest
below pH 5 (Shi et al. 2003).
In cases where proteolysis of recombinant proteins is observed, optimization of
expression parameters can significantly improve protein expression. For example,
61
lowering the induction temperature can reduce cell lysis and thereby minimize the release
of extracellular proteases; casamino acids can compete with the recombinant protein for
extracellular protease action and thereby protect the expressed protein from degradation
(Shi et al. 2003); daily addition of protease inhibitors can also increase protein expression
in P. pastoris (Kurokawa et al. 2002). Accordingly, in the present work the above
parameters were evaluated as a means of improving the functional, recombinant
expression of intact GOOX fusions.
Lowering the induction temperature to 15°C led to protein aggregation. However, slight
reduction of temperature to 27°C along with supplementation of 1% casamino acid and
daily addition of 2 µM leupeptin, significantly improved the expression of soluble, intact
fusion proteins. When using the improved cultivation conditions, the yield of the C- and
N- terminal constructs were similar, suggesting that the codon optimization used for N-
terminal constructs did not significantly increase recombinant protein expression in P.
pastoris. Moreover, the susceptibility of N-terminal fusion proteins to proteolysis at the
linker site suggests that in this case, the natural linkers used in the C-terminal constructs
were more resistant than the synthetic TP linker to secreted proteases.
The average yield of fusion and wild-type enzymes was approximately 5 mg/L and the
Ni-NTA purification system recovered wild-type and fusion proteins to over 95 % purity,
as judged by SDS-PAGE (Figure S4.1). The specific activities of the fusion proteins on
cellobiose were comparable or higher than that of wild-type GOOX with the exception of
GOOX_CtCBM3, which was approximately 80% of the wild-type activity (Table 4.1).
62
Figure 4.1 Schematic representation of wild-type GOOX and GOOX fusions. Schematic representation of carbohydrate binding modules fused to the amino and carboxyl terminal end of glucooligosaccharide oxidase from Sarocladium strictum. Natural linkers were used in C-terminal fusions while TP linkers were used in N-terminal fusions. The linker sequences are shown in the connecting lines.
Table 4.1 Specific activity of wild-type and CBM fusion GOOX on oligosaccharides.
Enzymes a Cellobiose Cellotetraose Cellohexaose Mixed XOS
GOOX wild-type 244 ± 7 b 197 ± 8 150 ± 2 229 ± 5 CtCBM3_GOOX 304 ± 4 253 ± 7 194 ± 3 289 ± 1 CtCBM11_GOOX 287 ± 1 235 ± 6 178 ± 6 274 ± 4 CtCBM44_GOOX 325 ± 6 256 ± 12 201 ± 4 317 ± 4 GOOX_CtCBM3 200 ± 8 166 ± 12 126 ± 0 185 ± 2 GOOX_CtCBM11 344 ± 24 279 ± 12 213 ± 4 327 ± 2 GOOX_CtCBM44 291 ± 3 239 ± 11 183 ± 4 271 ± 3
a All CBM fusions led to statistically significant differences in specific activities compared to wild-type GOOX as determined using a two-tailed t-test (P < 0.05) b The unit for specific activity is U/ µmol measured on 0.1 mM oligosaccharides. Standard deviations represent three biological replicates.
63
4.3.2 Improved binding to polymeric substrates
GOOX fusion to selected CBMs significantly improved GOOX binding to tested
polysaccharides, and binding selectivity was observed according to the appended CBM.
For instance, CtCBM3 fusion increased GOOX binding to crystalline cellulose (Avicel)
and regenerated amorphous cellulose by more than 10-fold (Table 4.2, Figure S4.2). As
expected, fusion to CtCBM11 and CtCBM44 increased the affinity of GOOX towards
soluble polysaccharides, including β-glucan, glucomannan, and xyloglucan (Figure 4.2);
fusion of GOOX to CtCBM11 or CtCBM44 also promoted enzyme binding to
regenerated amorphous cellulose (Table 4.2). Overall, binding results were consistent
with previous studies that confirmed CtCBM11 and CtCBM44 affinity towards β-glucan,
lichenan, hydroxyethyl cellulose, glucomannan and oat spelt xylan, along with CtCBM44
binding to xyloglucan (Carvalho et al. 2004; Najmudin et al. 2006; Viegas et al. 2008).
However, to our knowledge, binding of CtCBM11 and CtCBM44 to regenerated
amorphous cellulose has not been previously reported.
Table 4.2 Binding of wild-type GOOX and CBM fusions to insoluble cellulose.
Enzyme Portion bound (%) Avicel RAC
GOOX wild-type 10 10 CtCBM3_GOOX 60 95 CtCBM11_GOOX 5 65 CtCBM44_GOOX 10 75 GOOX_CtCBM3 55 95 GOOX_CtCBM11 5 70 GOOX_CtCBM44 10 95
The bound and unbound fractions were analyzed by SDS-PAGE (Figure S4.2), and the percentage of bound protein was calculated by measuring protein band density using ImageJ. Approximately, 5-10% uncertainty level is expected for the numbers given above.
64
In most cases, binding was not affected by CBM positioning at the N-terminus or C-
terminus of GOOX (Figure 4.2, Table 4.2). The exception was CtCBM44 fusions, where
C-terminal constructs showed slightly better binding to regenerated amorphous cellulose
than corresponding N-terminal constructs. Notably, the N-terminus of CtCBM44
contains a PKD domain, which would extend the linker region when CtCBM44 is fused
to the C-terminus of GOOX.
Figure 4.2 Affinity gel electrophoresis (AGE) of wild-type GOOX and CBM fusions. Purified proteins were subjected to AGE using a 7.5 % (w/v) polyacrylamide gel containing A: no polysaccharides, B: β-glucan, C: xyloglucan, D: glucomannan, E: carboxymethyl cellulose. The final concentration of each polysaccharide was 0.01 %. Bovine serum albumin (BSA) was used as a reference.
65
4.3.3 Activity on oligosaccharides
With the exception of C-terminal CtCBM3 fusion, all CBM fusions generated herein
increased GOOX activity on 0.1 mM cellobiose, cellotetraose and cellohexaose, as well
as mixed xylooligosaccharides with a DP of 2-7 (XOS) (Table 4.1). Similar results were
previously reported for GOOX activity on cello-oligosaccharides and xylo-
oligosaccharides after fusion to xylan-binding CtCBM22A (Vuong and Master 2014).
Subsequent kinetics analyses performed herein revealed that increased activities were
best explained by slight but statistically significant increases in kcat for all N-terminal
fusions as well as the C-terminal CtCBM11 construct (Table 4.3). Likewise, the reduced
activity of GOOX-CtCBM3 on all cellooligosaccharides could be explained by a
decrease in kcat.
The increase in kcat values on oligosaccharides observed for the N-terminal fusions
constructed herein was even greater in earlier studies where CtCBM22A was fused to the
N-terminus of GOOX (Vuong and Master 2014). In all cases, the same artificial TP
linker was encoded between the N-terminal CBM and GOOX. However in the case of
the earlier CtCBM22A fusion, the TP linker was extended towards the CBM by a 9-
amino acid loop (AVAGTVIEG), whereas the TP linker was extended by only 1 to 3
amino acids in the GOOX fusions generated herein. As previously proposed, CBM
addition (Vuong and Master 2014) or mutation (Vuong et al. 2013) at the N-terminus of
GOOX could cause a conformational change in the proximal FAD-binding domain,
thereby affecting the redox potential of the enzyme. We reasoned that a more flexible
linker would likely have a greater impact on the conformation of the FAD-binding
domain, which could explain the nearly 2-fold increase in kcat values upon CtCBM22A
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fusion (Vuong and Master 2014) compared with approximately 0.5-fold increase in kcat
values observed in the current study (Table 4.3). By contrast, the slight increase in kcat
values for the C-terminal fusion of CtCBM11 construct was correlated to the specificity
of CBMs from family 11 towards short oligosaccharides including cellotetraose (Viegas
et al. 2008), which is anticipated to promote functional associations between the substrate
and substrate binding site that is positioned at the C-terminal end of GOOX. The
decrease in kcat observed for GOOX-CtCBM3 is more difficult to rationalize. However,
since kcat is a function of both the catalytic rate constant and rate constant for dissociation
of the product, it is conceivable that product release is reduced upon C-terminal
positioning of the cellulose-binding CtCBM3 and corresponding linker sequence.
The addition of CtCBM22A to GOOX did not affect the Km value for oligosaccharides
(Vuong and Master 2014). Likewise in this study, no statistically significant changes to
the Km values were observed for cellotetraose upon CBM fusions, suggesting that the
CBMs used herein do not compete with -1 or -2 substrate binding subsites of GOOX
(Foumani et al. 2011).
Table 4.3 Kinetics parameters of wild-type GOOX and its CBM fusions on cellotetraose.
Enzymes kcat (min -1) Km (mM) kcat/Km b (min -1. mM-1) GOOX wild-type 250 ± 20 a 0.07 ± 0.02 3400 ± 700 CtCBM3_GOOX 300 ± 20 * 0.10 ± 0.02 3100 ± 700 CtCBM11_GOOX 310 ± 20 * 0.10 ± 0.02 3100 ± 700 CtCBM44_GOOX 380 ± 30 * 0.12 ± 0.03 3100 ± 700 GOOX_CtCBM3 150 ± 10 0.05 ± 0.01 2900 ± 600 GOOX_CtCBM11 360 ± 30 * 0.11 ± 0.02 3200 ± 700 GOOX_CtCBM44 290 ± 20 0.09 ± 0.02 3300 ± 700 a Standard deviations (n=3), b Standard deviations (SD) for kcat/Km values were calculated using following formula: SD kcat/Km= kcat/Km × [SQRT((SD(Km)/Km)2 + (SD(kcat)/kcat)2)]. * Statistically significant increase compared to the wild type as evaluated by t-test (p < 0.05).
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4.3.4 Specific activity on polymeric substrates
Specific activities were then tested using both soluble and insoluble polysaccharides.
Oxidation by wild-type GOOX or GOOX fusions was not detected on nanocrystalline
cellulose, total oat spelt xylan, or hydroxyethyl cellulose. However, activity of wild-type
GOOX was detected for the first time on konjac glucomannan, barley β-glucan,
carboxymethyl cellulose, regenerated amorphous cellulose, xyloglucan from tamarind
seed, and Avicel (Table 4.4). In the case of glucommanan, it is likely that the reducing
end glucose is mainly oxidized, given the low activity of GOOX on mannose (Foumani et
al. 2011).
Table 4.4 Specific activity of the wild-type GOOX and CBM fusions on polysaccharides.
Specific activitya (U/mmol)
GluM b(0.1%) RAC c (0.2%) β-Glu d (0.3%) CMC e (0.3%) XG f (0.5%) Avicel (0.5%)
GOOX wild-type 4,100 ± 300 g 290 ± 30 1,520 ± 90 600 ± 100 118 ± 3 79.2 ± 0.3
CtCBM3_GOOX 4,800 ± 100 * 420 ± 10 * 1,700 ± 100 630 ± 60 120 ± 20 123± 7 *
CtCBM11_GOOX 5,100 ± 100 * 430 ± 40 * 1,300 ± 200 560 ± 50 118 ± 6 86 ± 4
CtCBM44_GOOX 5,400 ± 200* 410 ± 50 * 1,700 ± 200 500 ± 100 112 ± 5 97 ± 8
GOOX_CtCBM3 4,100 ± 300 320 ± 20 1,500 ± 200 620 ± 50 110 ± 10 100 ± 4 *
GOOX_CtCBM11 5,000 ± 400 * 400 ± 30 * 1,500 ± 400 500 ± 90 120 ± 20 84 ± 7
GOOX_CtCBM44 5,400 ± 200 * 460 ± 60* 1,600 ± 200 500 ± 200 113 ± 3 99± 4 *
a Substrate concentrations were optimized to measure initial rates of reaction and are indicated in parentheses. bglucomannan from konjac, c regenerated amorphous cellulose, d β-glucan from barley, e carboxymethyl cellulose, f xyloglucan from tamarind seed.g Standard deviations (n=3) * Statistically significant improvements compared to wild-type GOOX as determined using a two-tailed t-test for two samples with unequal variance (p < 0.05)
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In experiments using polysaccharides, substrate concentrations were optimized to
measure initial rates of reaction. While different substrate concentrations complicate
comparisons between substrates, this analysis permitted comparisons relative to wild-type
GOOX for a given substrate, which was the main objective of the study. Relative to
wild-type GOOX, CBM fusions led to statistically significant improvements in GOOX
activity on 0.5% Avicel, 0.2% regenerated amorphous cellulose, and 0.1% konjac
glucomannan (Figure 4.3). For example, CtCBM3 and CtCBM44 fusions increased
GOOX activity on Avicel by up to 56%, where the highest enhancement was observed
with the N-terminal CtCBM3 fusion. Consistent with this trend, Telke et al. (2012) and
Voutilainen et al. (2014) also report greatest enhancement of activity on Avicel upon
fusion of CtCBM3 to an endoglucanase from Alicyclobacillus acidocaldrious (Cel9A),
and cellobiohydrolase from Talaromyces emersonii (Cel7A), respectively. Comparable to
specific activities reported for oligosaccharides (Table 4.1), all CBM fusions with the
exception of GOOX-CtCBM3 increased GOOX activity on regenerated amorphous
cellulose and glucomannan by up to 55% and 30%, respectively (Figure 4.3).
The improvement to GOOX activity was comparable to previous studies reporting 20-
50% increased performance of a cellodextrin phosphorylase on regenerated amorphous
cellulase upon addition of a CBM9 (Ye et al. 2011), and the 10-80% improved activity of
Cel9A Alicyclobacillus acidocaldrious on β-glucan upon fusion of a CBM from families
3,4 and 30 (Telke et al. 2012). Highest improvement was observed herein with fusion to
CtCBM44, where impacts were approximately two times higher when using 0.1%
glucomannan compared to 0.3% (Figure S4.3). This result, along with highest activity
improvement on Avicel and regenerated amorphous cellulose, is consistent with previous
69
reports showing greatest impact of CBMs on glycoside hydrolase activity when using
insoluble substrates or relatively low concentrations of soluble polysaccharides (Bolam et
al. 1998; Ali et al. 2001; Boraston et al. 2003; Várnai et al. 2013).
Figure 4.3 Specific activity of wild-type GOOX and CBM fusions on polysaccharides. A: crystalline cellulose (Avicel, 0.5%), B: regenerated amorphous cellulose (RAC, 0.2%), and C: glucomannan from konjac (0.1%). Substrate concentrations were optimized to measure initial rates of reaction. One unit corresponds to 1 µmol of product per min. Error bars represents standard deviations; n=3. The dotted line represents the specific activity of wild-type GOOX.
Changes in specific activity values were generally consistent with the selectivity of the
respective CBM. For instance, the increased specific activity of CtCBM3_GOOX on
Avicel compared to wild-type GOOX was consistent with improved binding of this
fusion protein to Avicel (Table 4.2), as well as the previously reported affinity of
CtCBM3 towards microcrystalline cellulose (Lehtiö et al. 2003). Similarly, the highest
specific activity of GOOX_CtCBM44 on glucomannan was correlated to the relatively
high binding of this construct on glucomannan (Figure 4.2D) and the previously reported
affinity of CtCBM44 towards this polysaccharide (Najmudin et al. 2006).
Even though CBM fusion improved GOOX binding to all tested polysaccharides, it did
not increase GOOX oxidation of barley β-glucan, carboxymethyl cellulose or xyloglucan.
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Whereas barley β-glucan comprises mixed β-1,3- and β-1,4-linkages and is unbranched,
xyloglucan and carboxymethyl cellulose have β-1,4-linked glucose backbones that are
substituted with branching sugars or carboxymethyl groups, respectively. An early study
of gluco-oligosaccharide oxidases did not detect GOOX-T1 activity on β-1,3-linked
glucose of laminaribiose (Lin et al. 1993), and a more recent study showed reduced
GOOX activity on branched xylo-oligosaccharides and anionic xylo-oligosaccharides
(Vuong et al. 2013). It is therefore likely that the mixed-linkage backbone structure of
barley β-glucan, and branching groups in carboxymethyl cellulose and xyloglucan,
restrict functional interactions between these substrates and the -1 and -2 subsites of
GOOX (Huang et al. 2005; Vuong et al. 2013).
4.3.5 Immobilization of GOOX through CtCBM3
In addition to enhancing activity on cellulose, we postulated that CtCBM3 fusion could
promote GOOX immobilization to cellulosic surfaces, which is relevant to several
applications including the use of enzymes in biosensing materials. Given the
comparatively high activity of CtCBM3-GOOX, additional comparative analyses were
restricted to CtCBM3-GOOX and wild-type GOOX, this time using a QCM-D equipped
with cellulose-coated piezoelectric crystal sensors.
In this analysis, protein adsorption is observed as a decrease in oscillation frequency of
the sensor, whereas an increase in dissipation reflects a more viscoelastic surface layer
(Hook et al. 1998). Accordingly, the higher change in frequency (∆f) observed using
CtCBM3-GOOX confirms enhanced binding of this enzyme to cellulose compared to the
wild-type GOOX (Figure 4.4; Figure S4.4). Considering corresponding differences in
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molecular weight, ∆f values corresponded to a molar adsorption ratio of the fusion and
wild-type enzymes of approximately 1.6. Notably, the slight increase in slope (∆D/∆f)
of the D-f plot depicting CtCBM3-GOOX binding suggests a more viscoelastic surface
layer was formed by the fusion enzyme (Figure 4.4).
Figure 4.4 Frequency - dissipation plot of enzymes binding to cellulose. Changes in frequency (Δf) and dissipation (ΔD) during 1.5 µg/mL enzyme addition (1), 50 mM Tris-HCl pH 8 buffer washing (2) and 0.5 mM cellobiose addition (3) in the experiments with CtCBM3_GOOX (green, triangle) and wild-type GOOX (red, square). The total running time is 210 min. Linear fitting for cellulose binding of the CBM fusion (dashed arrow) and the wild-type (solid arrow) were analyzed by GraphPad Prism 5.
After extensive washing to remove loosely bound enzyme, cellobiose was passed over
the sensors coated with immobilized CtCBM3-GOOX or GOOX. The addition of
cellobiose resulted in negligible changes to frequency and dissipation (Figure 4.4),
indicating that the enzymes remained bound to cellulose in the presence of the soluble
substrate. Moreover, the activity of immobilized CtCBM3-GOOX and GOOX was
confirmed by measuring H2O2 after batch incubation of recovered sensors with 0.5 mM
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cellobiose. Given that measured activities remained stable after repeated cycles of
washing and incubation of the sensors with cellobiose (Figure S4.5), this analysis
demonstrated that fusion of GOOX to CtCBM3 could have advantages beyond increased
activity on cellulose, including one-step purification of active enzyme from culture media
based on cellulose adsorption, as well as biosensing and biofuel cell applications
involving mixed sugars.
4.3.6 Effect of CBM on thermostability
Earlier studies of wild-type GOOX confirmed its stability at 40 °C, but loss in over 30%
and 90% activity after 1 h at 45°C and 50°C, respectively (Foumani et al. 2012).
Therefore, in the current study, the half life of wild-type GOOX and CBM fusions were
compared at 45 °C. Despite selecting CBMs from a thermophilic bacterium, their fusion
to GOOX had a moderate impact on the temperature stability of corresponding fusion
proteins (Table 4.5, Figure S4.6). In particular, C-terminal fusion of CtCBM44 and N-
terminal fusion of CtCBM3 and CtCBM11 increased the half-life of GOOX at 45 °C by
10-40% and the C-terminal fusion of CBM3 decreased the half-life by 15%.
From an applied perspective, the half-lives under the reaction conditions (i.e. 37 °C), or at
room temperature, are expected to be much longer and the impact of CBM fusion could
be different in those situations.
It is interesting to note that the impact of CBM fusion on GOOX stability was more
readily explained by the nature of the linker region rather than CBM family. For
example, the rigidity of the short linker and connecting loops discussed earlier might
73
restrict heat-induced conformational shifts in the N-terminal catalytic domain of GOOX,
which could explain the relative thermostabilities of N-terminal fusions (Table 4.5).
Table 4.5 The half-life of fusion and wild-type GOOX at 45°C.
Enzymes Half-life (min)a GOOX wild-type 119 ± 4 b CtCBM3_GOOX 132 ± 2 * CtCBM11_GOOX 132 ± 2 * CtCBM44_GOOX 127 ± 4 GOOX_CtCBM3 104 ± 9 GOOX_CtCBM11 111 ± 10 GOOX_CtCBM44 170 ± 20 *
a. Values were obtained by plotting the log (% residual activity) versus incubation time (min). b Standard deviations (n=3) * Statistically significant improvements compared to wild-type GOOX as determined using a two-tailed t-test for two samples with unequal variance (p < 0.05)
This notion was further evident when comparing linker sequences of C-terminal fusions.
Specifically, the higher half-life of GOOX-CtCBM44 compared to other C-terminal
fusions could be explained by the PPPY linker sequence between GOOX and CtCBM44,
which likely adopts a more rigid conformation compared to the longer linker of CtCBM3
(PTNTPTNTPTNTP) and the connecting loop of CtCBM11 (SRAVGE). The potential
impact of linker sequences on thermostability is interesting in the context of an earlier
report by Dias et al. (2004), who showed that the thermostability of C. thermocellum
xylanase Xyn10B was retained after removing the CBM22 domain and leaving the linker
sequence, whereas removing the whole linker-CBM22 sequence reduced the
thermostability of the enzyme. Moreover, cellulase fusions with either a flexible
polyglycine linker or a rigid alpha-helix linker showed that the rigid linker significantly
74
enhanced enzyme activities and thermostability (Zou et al., 2012). Further examples
showing variable effects of CBM fusion on the temperature stability of associated
enzymes have been reviewed in section 2.4.3.
4.4 Conclusions
A selection of CBMs with affinity towards different β-glucans were appended to either
the C-terminus or N-terminus of GOOX, and fusion proteins with similar or higher yield
than the wild-type enzyme were successfully expressed and purified from P. pastoris. All
N-terminal fusion proteins as well as the C-terminal CtCBM11 fusion showed higher
catalytic activity on tested oligosaccharides than wild-type GOOX, suggesting a
beneficial conformational change to the FAD binding domain. In addition, unchanged Km
values confirmed that the fused CBMs did not compete with the GOOX subsites for
oligosaccharide binding. Similar to activity studies, thermostability of fusion constructs
was dictated by the nature of the linker sequence rather than CBM type, where more rigid
linkers resulted in more stable fusion proteins, underscoring the relevance of linker
selection to fusion protein design. Finally, regardless of positioning, CBM fusion
promoted GOOX binding to cellulosic and hemicellulosic polysaccharides, and GOOX
remained active when immobilized to cellulose through CtCBM3. This result highlights
that CBM fusion, in particular the N-terminal CtCBM3, could facilitate applications of
GOOX in cellulose-based biosensing devices.
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Chapter 5 Application trials of wild-type and engineered GOOX
Section 5.1 of this chapter are published in: Vuong T, Vesterinen A, Foumani M, Juvonen M, Seppälä J, Tenkanen M, Master E.R,
Xylo- and cello-oligosaccharide oxidation by gluco-oligosaccharide oxidase from
Sarocladium strictum and variants with reduced substrate inhibition. Biotechnology for
Biofuels, June 2013.
Contributions to the above article: Expression and purification a GOOX variant enzyme
used in this study, performing preliminary mass spectrometry experiments to validate the
method and to design sample preparation; Preparation of GOOX-oxidized
cellooligosaccharides (DP: 2-5) for the NMR and mass spectrometry analysis; and
preparing the corresponding sections in the manuscript.
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The wild-type and engineered GOOX have been evaluated for the production of oxidized
carbohydrate standards required to better characterize the action of enzymes relevant to
cellulose conversion to sugars (e.g. LPMO enzymes); GOOX and GOOX variants were
also evaluated for their utility in two applications relevant to food industries. While the
results are promising, further experiments are needed to validate and implement these
applications. Accordingly, the following sections will describe analyses completed
through the course of my PhD, and Chapter 6 (Future Directions) will summarize our
next steps.
5.1 GOOX in the production of sugar standards
5.1.1 Introduction
Recent characterizations of lytic polysaccharide monooxygenases (LPMOs) from
auxiliary activity family 9 (AA9), has revealed new routes to enzymatic biomass
conversion (Horn et al. 2012). LPMO activity involves the oxidative cleavage of
polysaccharide chains, and while the mechanism of LPMO action is not fully resolved, it
is clear that these enzymes cleave glycosidic bonds while oxidizing the adjacent carbon.
Thus, the products of LPMOs activity are mainly oxidized at the anomeric carbon,
although oxidization of the non-reducing end has also been reported (Quinlan et al. 2011;
Beeson et al. 2012; Westereng et al. 2013). C1 oxidation produces oligosaccharides that
lack a reducing end. Instead, under physiological conditions, the lactone spontaneously
converts to an aldonic acid. As a result, cleavage of glycosidic linkages by LPMOs
cannot be monitored using regular glucose-detecting assays.
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New assays have been developed to monitor LPMOs activity, and chemical routes were
applied to generate oxidized-oligosaccharides as standards. For instance, Westereng et al.
(2013) used iodine oxidation in a methanol:potassium hydroxide mixture to selectively
oxidize the C1 carbon of the cellooligosaccharides to serve as standards in an LPMO
assay (Westereng et al. 2013) and other reports applied similar techniques using bromine
for oxidation of the anomeric carbon in oligosaccharides (Diehl et al. 1974). However,
since the oxidation is not complete, these chemical techniques usually involve costly
purification steps; moreover they require handling of toxic regents such as methanol.
In this study, we hypothesize that GOOX can efficiently oxidize oligosaccharides at the
anomeric position, thus providing a cleaner method to generate oxidized sugar standards
for LPMO activity assays. Accordingly, the present project investigates the structure of
the GOOX-oxidized cellooligosaccharides with degree of polymerization from two to
five and studies the efficiency of oxidation by detecting the non-oxidized compounds.
5.1.2 Materials and Methods
5.1.2.1 NMR analysis of oxidized products
Reaction mixtures containing 10 mM cellobiose or 10 mM xylobiose, and 160 nM
GOOX or GOOX-Y300A, in 50 mM Tris HCl (pH 8.0) were incubated overnight at 37oC.
Oxidized products were analyzed at Aalto University, Prof. Seppälä Lab, by proton
nuclear magnetic resonance (1H NMR) using a Bruker 400 MHz NMR Spectrometer
(Bruker Ultrashield 400 Plus, USA). Samples were measured by our collaborator, Arja-
Helena Vesterinen, directly in the reaction solvent with water suppression using 10%
78
deuterium oxide as a co-solvent for deuterium lock. The peaks were identified using the
estimation program of ChemBioDrawUltra 12.0 (CambridgeSoft).
5.1.2.2 Mass spectrometric analysis of oxidized products
Reaction mixtures containing 1 mM of cello-oligosaccharides, from cellobiose to
cellohexaose, and 160 nM GOOX or GOOX-Y300A, in 50 mM Tris HCl (pH 8.0) were
incubated overnight at 37oC. To characterize oxidized products, 100 µL of each reaction
mixture was diluted in 900 µL of MilliQ-water, and diluted samples were purified at
University of Helsinki, by Minna Juvonen in Prof. Tenkanen’s lab and fractionated using
a Hypersep porous graphitized carbon column (Thermo Scientific, MA, USA), following
the protocols of Packer et al. (Packer et al. 1998) and Chong et al. (Chong et al. 2011) but
with modifications. Neutral sugars were eluted using 40 % acetonitrile, and a mixture of
50% acetonitrile and 0.05 % TFA was used to elute acidic sugars. Collected fractions
were dried with nitrogen gas for 20 min and then freeze-dried overnight.
Positive ion mass spectrometric analyses were performed using an Agilent XCT Plus
model ion trap mass spectrometer (Agilent Technologies, Waldbronn, Germany)
equipped with an electrospray source. For ESI-MS-analysis, freeze dried samples were
dissolved in 20 µL of MilliQ-water, and 6 µL of each sample was diluted in 100 µL of
methanol-water-formic acid solvent (50:49:1 (v:v:v)). Sample solutions were introduced
into the ES source at a flow rate of 5 µL/min via a syringe pump. The drying gas
temperature was set to 325 °C; drying gas flow was 4 L/min; the nebulizer pressure was
79
15 psi, and the ES capillary voltage was set to 3164 V. Ions were collected in the m/z
range of 50 to 1000.
5.1.3 Results and Discussion
5.1.3.1 Confirming the regioselectivity of gluco-oligosaccharide oxidases
To date, very few studies have confirmed the position of hydroxyl groups oxidized by
family AA7 gluco-oligosaccharide oxidases. Lee et al. used 13C and 1H NMR to confirm
that GOOX-T1 targets the hydroxyl group of the anomeric carbon; however, only
maltose was used in their analysis (Lee et al. 2005). Since gluco-oligosaccharide oxidase
activity is higher on cello-oligosaccharides and xylo-oligosaccharides than malto-
oligosaccharides (Foumani et al. 2011), 1H NMR was used here to evaluate the effect of
sugar type and chain length on the regio-selectivity and catalytic efficiency of GOOX
enzymes.
The disappearance of the H1 doublet signals from the reducing end of α- and β-glucose
units of cellobiose is consistent with oxidation at the anomeric C1 position (Figure 5.1A)
(Nouaille et al. 2009). Similarly, the peak height for the H1 signals from the reducing end
of α- and β-xylose units of xylobiose was decreased in oxidized xylobiose samples
(Figure 5.1B). Ring opening at the anomeric position was also revealed by the detection
of H2 and H3 signals at 4.05 ppm and 3.95 ppm in case of oxidized cellobiose, and at
4.01 ppm and 3.81 ppm, respectively in case of oxidized xylobiose (Higham et al. 1994;
Nouaille et al. 2009). The signals for the corresponding lactone were not observed
probably due to the relatively long oxidation reaction (24 h); similar observations were
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reported after overnight incubation of Phanerochaete chrysosporium CHD with
cellobiose (Higham et al. 1994).
Figure 5.1 NMR spectra of cellobiose (A) and xylobiose (B) oxidation. (A): From top to bottom are the spectra of cellobiose, cellobiose that was oxidized by wild type GOOX, and cellobiose oxidized by GOOX_Y300A; CB red. alpha and CB red. beta: H1 signals due to reducing α-glucose and reducing β-glucose units of cellobiose, correspondingly; CBA-H2 and CBA-H3: H2 and H3 signals of the cellobionate molecule. (B): From top to bottom are the spectra of untreated xylobiose and GOOX oxidized xylobiose; XB red. alpha and XB red. beta: H1 signals due to reducing α-xylose and reducing β-xylose units of xylobiose, correspondingly; XBA: Overlapped signals of the xylobionate molecule. 10 mM cellobiose and 10 mM xylobiose were used in oxidation reactions.
ESI-MS/MS analyses also indicated the enzymatic oxidation of cellotriose at the
anomeric carbon. In the positive ionization mode, the acidic fraction of oxidized
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cellotriose only produced glycosidic bond cleavage fragments, generating B- and Y-ions
(Figure 5.2A); cross ring cleavage fragmentation was not observed. Since neutral reducing
oligosaccharides usually form cross ring cleavage fragments from reducing ends if a
sodium cation is present (Hofmeister 1991; Asam and Glish 1997), oxidation of the
anomeric carbon seemed to change the fragmentation behaviour of sodium cationized
cellotriose. In the negative mode, B and C-ions from glycosidic bond cleavage were the
most abundant fragment ions (Figure 5.2B).
Figure 5.2 MS/MS spectra and fragmentation of GOOX oxidized cellotriose. (A): ESI MS/MS in the positive ionization; the m/z ratio of the precursor, [M+Na]+, is 543. (B): ESI MS/MS in the negative ionization; the m/z ratio of the precursor, [M-H]-, is 519. Fragment ions were named according to Domon and Costello (Domon and Costello 1988).
The molecular masses of Y- and Z-ions increased by 16 Da, compared to the un-oxidized
control sample (data not shown), supporting that the oxidation reaction occurred in the
reducing glucose. Cross ring cleavage fragmentation was also observed in the negative
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mode. For instance, a peak at the m/z ratio of 383 was generated from oxidized
cellotriose (m/z 519) by the loss of 136 Da from cross ring cleavage of the oxidized
monosaccharide unit, leading to the formation of a 2,4A3-ion (Fig. 5.2B).
Additional, indirect evidence, from colorimetric assays, for the oxidation at C1 is that no
activity was detected on D-glucose derivatives lacking a C1 hydroxyl group, including
1,5-anhydroglucitol (D-glucose with -H instead of -OH at C1) and methyl-β -D-
glucopyranoside (D-glucose with -OCH3 instead of -OH at C1)
5.1.3.2 Efficient oxidation of cellooligos. and Impact of chain length on GOOX activity
Mass spectrometric analysis of oxidized cello-oligosaccharides from cellobiose to
cellopentaose revealed a 16 Da increase in m/z values of the acidic fraction (Figure
5.3M-P) compared to the control, un-oxidized samples (Figure 5.3A-D), confirming that
in all cases, the oxidation by GOOX introduced a single oxygen atom to all the
oligomeric substrates. The oxidation of different cello oligosaccharides was efficient, but
not complete at the tested concentrations, as can be seen from small amount of un-
oxidized oligosaccharides detected in the neutral fraction (Figure 5.3I-L).
The current analyses confirmed that the GOOX oxidize mono- and oligo-saccharides
only at the reducing anomeric position suggesting that GOOX production of oxidized
cello oligosaccharides would be an efficient way to generate oxidized carbohydrate
standards to facilitate the characterization of the C1-oxidizing enzymes of family AA9.
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Figure 5.3 Positive ion ESI-MS spectra of four cello-oligosaccharide samples. G2: Cellobiose; G3: Cellotriose; G4: Cellotetraose; G5: Cellopentaose. (A)-(H): Un-oxidized cello-oligosaccharide samples; (I)-(P): GOOX oxidized cello-oligosaccharide samples; (A)-(D) and (I)-(L): Neutral fractions: (E)-(H) and (M)-(P): Acidic fractions.
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5.2 Application of GOOX CBM fusions in the synthesis of plant oligosaccharides
with enhanced nutraceutical value
5.2.1 Introduction
Plant oligo- and poly-saccharides have been used to enhance the nutritional value of
different food products (Broekaert et al. 2011). For instance, certain oligosaccharides
have immunostimulatory activity, serving as a potent stimulator of the immune system;
or prebiotic activity, promoting a healthy digestive system (Barreteau 2006). Panesar et al.
(2013) review a comprehensive list of prebiotics currently used in the food industry, and
summarize those enzymes used in prebiotic production. Notably, most enzyme treatments
used to date hydrolyze starting polysaccharides to a series of oligosaccharides (Panesar et
al. 2013).
Previous studies have shown that among prebiotic carbohydrates, acidic oligosaccharides
such as those from pectin are correlated to comparatively high prebiotic activity
(Mandalari et al. 2007). Specific nutritional benefits include metabolic resistance and
promotion of mineral absorption, which have been attributed to the acidic functionality of
these oligosaccharides (Schaafsma 2008). These benefits are exemplified in lactobionic
acid, produced by oxidation of lactose using oligosaccharide oxidases such as GOOX-T1,
MnCO and PCOX (Lin et al. 1996; Nordkvist et al. 2007; Murakami et al. 2008). This
compound is a potential prebiotic that also enhances intestinal absorption of calcium
(Brommage et al. 1993) without contributing to calcification risk; it is also resistant to
digestive enzymes, which makes it available to be fermented by the intestinal flora
(Schaafsma 2008). In particular, the increase in the mineral uptake is believed to occur in
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the large intestine by the passive absorption. Unlike the active mineral absorption that
depends mainly on vitamin D to transport the minerals across the cells and takes place in
the upper gastrointestinal tract, the passive mineral uptake occurs along the length of
small and large intestine by paracellular diffusion as a result of gradient in mineral
concentrations (Padma Ishwarya and Parbhasankar 2014). Thus, the potential of the acid
to make a strong complex with minerals leaves the acidic prebiotic compound in complex
with mineral in the large intestine where the prebiotic can be fermented to fatty acids,
which is predicted to lower the pH of contents in the large intestine and thereby increase
the solubility of minerals, promoting their passive absorption.
Aldouronic acids from xylan, which are acidic xylooligosaccharides (XOS), have also
been shown to relieve iron deficiency (Kobayashi et al. 2011). However these acidic
XOSs, which comprise xylooligosaccharides substituted with glucuronic acid by α-1,2
bonds (Figure 5.4), do not show prebiotic activity (Ohbuchi et al. 2009) even though
neutral XOSs are known to have prebiotic function (Okazaki et al. 1990; Aachary and
Prapulla 2011; Kondepudi et al. 2012).
Figure 5.4 Structure of compounds used in prebiotic assay. A) xylooligosaccharides (XOS), B) GOOX-CBM44 oxidized XOS, and C) aldouronic acid used in this study.
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In this study, we hypothesized that the loss in prebiotic activity observed for aldouronic
acids from xylan reflect the branching structure of corresponding oligosaccharides, rather
than the acidity of the molecule. To test this hypothesis, we oxidized neutral XOSs using
a glucooligosaccharide oxidase (GOOX), which selectively produces an aldonic acid at
the reducing end of the molecule (Figure 5.4); both aldouronic acids from xylan and the
GOOX-treated XOSs were then tested using an in vitro prebiotic activity assay. In short,
GOOX treatment of XOS did not reduce the prebiotic potential of XOSs, as evaluated by
cultivation of B. longum. By contrast, the aldouronic acids from xylan were not utilized
by this strain of bifidobacteria. The growth rate of B. longum on GOOX-treated XOSs
and neutral XOSs was similar, and yields of B. longum were slightly higher after 24h
cultivation on GOOX-treated XOSs than neutral XOSs. These findings suggest that
GOOX oxidation of XOS can support bifidobacteria proliferation characteristic of
prebiotic activity, while also providing additional benefits previously correlated to the
acidic functionality.
5.2.2 Materials and Methods
5.2.2.1 Oxidation of xylooligosaccharides
Mixed xylooligosaccharide (DP 2-7, 95% pure) was purchased from Cascade
biochemical (USA), while aldouronic acid was obtained from Megazyme (Ireland). The
mixed xylooligosaccharide was oxidized using the GOOX-CBM fusion that was shown
in Section 4.3.3 to be most active towards XOS, namely GOOX-CBM44. Approximately
4 µg/ml of the enzyme was added to 1% w/v of the filter-sterilized substrate in final
volume of 10 mL, and the reaction was incubated overnight at 37°C. To confirm the
87
oxidation, small-scale reactions were performed in parallel and characterized using the
standard chromogenic GOOX assay described in section 3.2.6 to detect the production of
H2O2. Following enzymatic oxidation, GOOX-CBM44 was de-activated by incubation at
50 °C for 1 h.
5.2.2.2 Prebiotic Assay
Prebiotic assays were performed in Professor Elena Comelli’s lab in the Department of
Nutritional Sciences at the University of Toronto; all prebiotic experiments were
performed under the supervision of Dr. Amel Taibi, a post-doctoral fellow in Professor
Cornelli’s laboratory. Bifidobacterium longum NCC2705 was pre-cultured anaerobically
in a glove box (Coy Laboratories, Midland, MI, USA) containing an atmosphere of 90%
N2, 5% CO2, and 5% H2 (v/v). The pre-cultures were grown on MRS broth (1% Casein
peptone, 1% Meat extract, 0.5% Yeast extract, 0.1%g Tween-80, 0.2% K2HPO4, 0.5%
Na-acetate, 0.2% (NH4)2 citrate, 0.02% MgSO4-7H2O, and 0.005% MnSO4-H2O) at pH
6.5 supplemented with 0.5% glucose. Prior to fermentation, the cells from the pre-culture
were counted under the microscope using a Hemocytometer (Fisher scientific). A volume
corresponding to 5×105 CFU was centrifuged, and the cells were re-suspended in MRS
broth lacking glucose, and supplemented with the 0.5% of defined carbohydrates, i.e.
oxidized and non-oxidized XOS. Amended carbohydrates were the sole carbon source,
and 0.5 % glucose was used as the positive control. A final volume of 1-2 mL was used
for preliminary small-scale cultivations whereas 5 mL cultures were prepared for
studying the effect of enzyme treatment. The cultures were grown anaerobically at 37°C
under stationary conditions in either a gas-pack system (AnaeroGen TM, Oxoid) for
preliminary cultivation, or in a glove box for 5-mL cultures. An aliquot of each culture
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was harvested at each time point to determine cell growth using viable colony counts.
Viable cell counts were determined by spreading 100 µL of 10-fold serially diluted
cultures onto solid MRS agar plates containing 0.5% glucose, and plates were incubated
in the glove box at 37°C for 24-48 h.
5.2.3 Results and Discussion
5.2.3.1 Small-scale fermentation on xylooligosaccharides and aldouronic acid
Neutral xylooligosaccharides promoted the growth of B. longum almost to the same level
as glucose, while aldoronic acid was not digested by this strain (Figure 5.5). The inability
of B.longum to grow on this compound was expected and consistent with the analysis of
Ohbuchi et al (2009). Based on this result, only xylooligosaccharides before and after
oxidation with GOOX-CBM44 were used for the subsequent cultivations.
Figure 5.5 Viable cell count of B. longum preliminary cultures. The growth media was MRS supplemented with xylooligosaccharides or aldouronic acid. Glucose was served as positive control and MRS media without a sugar was severed as the blank. Data points represent cultivation of one culture per compound.
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5.2.3.2 Cultivation of B. longum on oxidized and non-oxidized xylooligosaccharides
The viable cell count results indicate slightly higher population of B.longum on GOOX-
CBM44 oxidized XOS vs the non-oxidized substrate at 24 h measurements (Figure 5.6).
However, as shown in Figure 5.6, the growth profile is different throughout the
cultivation time. For instance, the cultures grown on XOS shows higher population than
those grown on enzyme treated XOS after 2 h of incubation while this correlation is not
maintaed at 5 h or 24 h time points.Thus, to quantitavely compare this profile the growth
rate was calculated in the exponential phase using the following equation:
𝐾 =(log𝑁2 − log𝑁1) × 2.3
𝑇2 − 𝑇1
, where N1 is the population (colony forming unit, CFU) at T2 (h), and N1 is the
population at T1 (h).
As shown in Table 5.1, the growth rate of B. longum in the exponential phase from T1 =
2.5 h to T2 = 7.5 h is higher on glucose and is slightly higher on GOOX-CBM44 treated
XOS vs non-treated XOS. However, a paired t-test with p value of 0.05 revealed that the
difference between growth rates on treated and non-treated XOS was not statistically
significant. These data confirm that the acidic functional group does not reduce the
prebiotic activity of XOS as evaluated using B. longum . In the case of aldouronic acid,
also know as acidic XOS, glucoronic acid substituents are at terminal or branching
positions of the xylose backbone. It is possible then, that the main reason for the loss of
prebiotic value could be limited ability to remove glucuronic acids from the
xylooligosaccharide, rather than inability to metabolize acidic xylooligosacccharides in
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general. In this case, GOOX-CBM44 oxidized XOS could be a viable route to enhancing
the nutritional value of XOSs, for example through promoting mineral absorption.
Figure 5.6 Viable cell count of B. longum cultures. The growth media was MRS supplemented with xylooligosaccharides (XOS) with and without GOOX-CBM44 treatment. Glucose was served as positive control. The error bars indicate standard deviation of three biological replicates.
Table 5.1 Growth rate of B. longum cultivations.
Substrates Growth rate, K (h-1)
XOS 0.51 ± 0.02 a
GOOX-CBM44 treated XOS 0.53 ± 0.09
Glucose 0.60 ± 0.05
a The values are measured at exponential phase between 2.5 to 7.5 h. standard deviations represents the growth rates of the three biological replicates.
1.00E+07
1.00E+08
1.00E+09
1.00E+10
0 5 10 15 20 25 30
Log
(cfu
/ml)
Time (h)
XOS XOS+GOOX Glucose
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5.2.4 Conclusions
This study compared the growth profile of B. longum on XOS before and after oxidation
by GOOX-CBM44, which introduces a carboxyl groups at the reducing end of the
oligosaccharide. This analysis did not reveal statistically significant differences between
the growth rates of B. longum on the two XOS samples implying that the introduction of
the acidic groups at the reducing end of the xylooligosaccharide does not reduce the
digestibility of XOS by B. longum.
The acidic XOS prepared in this work by GOOX-CBM44 treatment maintains the
bifidobacteria proliferation characteristic of XOS while potentially increasing intestinal
mineral absorption potential. This finding presents a new opportunity for GOOX
application in the preparation of value added prebiotics.
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5.3 A Mutant gluco-oligosaccharide oxidase is suitable to replace glucose oxidase for
baking applications
5.3.1 Introduction
In baking applications, additives have been widely used to improve the texture, volume,
flavor, and shelf life of the packed goods. Most additives contain enzymes, including
amylase, hemicellulase, cellulase, and carbohydrate oxidases, the action of which results
in improved rheological and handling properties of the dough (Sharma and Singh 2010).
Unlike amylase, xylanase and cellulase, which reduce the molecular weight of
polysaccharides present in the dough, oxidases target hydroxyl groups of corresponding
sugars, thereby producing aldonic acids.
Glucose oxidase (GO) has been widely used as a carbohydrate oxidase in baking
applications (Vemulapalli et al. 1998; Rasiah et al. 2005; Bonet et al. 2006; Hanft and
Koehler 2006; Dagdelen and Gocmen 2007; Decamps et al. 2013) to reduce stickiness
and enhance machinability and stability of the dough while retaining the softness of
baked products after long storage (Bonet et al. 2006). However, the effectiveness of GO
is limited to the selectivity of this enzyme towards glucose, and the generally low
concentration of glucose in cereal flour (Schneider et al. 2003).
Briefly, cereal flour from various grains generally comprise carbohydrates, dietary fibre,
and protein as their main ingredients. The primary constituent of the carbohydrate
fraction is starch, which accounts for 60-70% of the flour (Fišteš et al. 2014); free sugars
including glucose, maltose, and sucrose account for less than 2% (MacArthur and
Dappolonia 1979). The composition of dietary fibres such as cellulose, arabinoxylan, and
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ß-(1→3), (1→4)-glucan originated mainly from brans and germs (Koehler and Wieser
2013) is different among the grains ranging from 12-20% of whole grain flour and 1-8%
of the white flour from wheat, rye, barley, corn, sorghum and rice (Nyman et al. 1984).
While these percent compositions reflect the polysaccharide content in cereal flour, the
actual fibre content in the baking ingredients is anticipated to be higher because prebiotic
dietary fibres are increasingly used in bakery for their nutritional and technical benefits to
the backed products (Padma Ishwarya and Parbhasankar 2014). In addition, the use of
hydrolytic enzymes in baking processes reflects the presence of corresponding short
oligosaccharides in the dough mixtures. This wide range of carbohydrates present in
cereal flour suggests that an oxidative enzyme with wider carbohydrate specificity is
advantages over GO for dough enhancement.
Accordingly, oxidoreductases that act on oligosaccharides have been shown to perform
better than GO in baking applications in terms of introducing aldonic acids and H2O2 to
the dough mixture (Schneider et al. 2003). More specifically, MnCO used in a baking
trial improved the machinability of the dough in a dose-dependent manner. As well, a
better consistency of the dough was exhibited upon addition of this enzyme (Schneider et
al. 2003). Despite the promising characteristics of MnCO, the performance of this
enzyme in terms of H2O2 inactivation has not been studied in detail.
GO is inactivated in the presence of low concentrations of H2O2, i.e. less than 0.01 M, as
described in section 2.5.1. This compound is the co-product of most enzymatic sugar
oxidations and is an antimicrobial factor, maintaining dough freshness. However, it has
also been shown to oxidize amino acids such as methionine and tryptophan that reside
94
near the active site of carbohydrate oxidases, thereby inactivating these enzymes
(Hachimori et al. 1964; Kleppie 1966). Earlier reports show that immobilized GO under
continuous operation is still inactivated by H2O2 (Greenfield et al. 1975) whereas
liposomal encapsulation of GO (Yoshimoto et al. 2004) can protect the enzyme from
H2O2 inactivation by decomposition of this compound at the lipid membrane, keeping the
concentration of H2O2 low inside and outside of the liposome capsules.
In this work, a mutant gluco-oligosaccharide oxidase, GOOX-Y300A, with higher
specific activity on glucose (as discussed in section 3.3.4) and reduced substrate
inhibition than the wild-type GOOX (Foumani et al., 2011), was investigated for its
potential to replace GO, benefiting the baking applications with a similar mechanism
(Figure 5.7, Figure 5.8) but likely with higher efficiency. GOOX-Y300A is capable of
oxidizing oligosaccharides including maltose-based oligosaccharides, which are the
components of starch, as well as glucose-based and xylose-based oligosaccharides
(Foumani et al., 2011), which are constituents of fibre in flour. GOOX-Y300A in fact,
shows higher rates of activity on oligosaccharides than the corresponding
monosaccharides. In particular, this enzyme is expected to benefit the properties of dough
commonly prepared from flour with the aid of hydrolytic enzymes (e.g. cellulases,
xylanases or amylases), which is expected to have relatively high oligosaccharide content.
This study compares GO and GOOX-Y300A for their specific activities on
oligosaccharides and their inactivation by H2O2.
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Figure 5.7 Prosed mechanisms for GOOX benefits in baking applications. Proposed effects are similar to those of other oxidative enzymes used in baking (Rasiah et al. 2005; Bonet et al. 2006). The figures were taken from www.cornishpasties.com/breads and www.feastsforallseasons.com/english-muffins.
Figure 5.8 Proposed mechanism for GOOX reinforcing the protein network in dough; similar to that proposed for GO (Rasiah et al. 2005). The figure was generated using ChemSketch.
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5.3.2 Materials and Methods
Cellobiose, maltose, xylose, glucose, and gluconic acid were reagent grade with over
95% purity and purchased from BioShop Inc. (Canada); cellotetraose (>95% pure) was
obtained from Megazyme (Ireland), whereras mixed xylooligosaccharides (DP-2-7, 95%
pure) were obtained from Cascade Biochemicals (United State). Glucose oxidase from A.
niger (GO) was purchased from Sigma (United state); this product exhibits less than 10
U/mg catalase activity. GOOX-Y300A was prepared as previously described in section
3.2.3 (Foumani et al., 2011).
5.3.2.1 Activity assays
Since different preparation methods were used for GOOX-Y300A and GO, the
concentration of both enzymes was re-measured in parallel using Pierce BCA Protein
Assay (Thermo Scientific). Specific activities of enzymes were then tested on mono- and
oligo-saccharides.
To determine the GOOX-Y300A activity, the standard chromogenic GOOX assay
described in section 3.2.6 was used to measure the production of H2O2. To initiate
oxidation by GOOX, 6 pmol of enzyme was added to the 250 µL reaction mixture (0.1
mM 4-AA, 1 mM phenol, 0.5 U horseradish peroxidase) in 50 mM Tris–HCl buffer (pH
8). Oxidations by GO were performed in the same reaction mixture as above except with
the buffer being 50 mM sodium acetate (pH 5) and were initiated by adding 6 pmol of
enzyme to the same reaction mixture as above. In both cases, reactions were incubated at
37 °C for 15 min and specific activities were separately measured on 10 mM xylose,
maltose and cellobiose as well as 1 mM of glucose, cellotetraose and mixed
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xylooligosaccharides. One unit of the enzyme activity was correlated to the production of
1 µmol of the product per minute.
5.3.2.2 Oxidation Reactions for H2O2 inactivation study
To measure the effect of H2O2 on the enzyme activities, 0.1 μg of GOOX-Y300A or GO
were added to 250 μL mixtures of 1 mM glucose in either 50 mM Tris-HCl buffer (pH 8)
for GOOX-Y300A, or 50 mM sodium acetate buffer (pH 5) for GO reactions, containing
various concentrations from 0-0.2 M of H2O2. The reactions were incubated at 25 °C for
5h, and then were stopped by removing the enzyme using Nanosep centrifugal device
(Pall Corp). The extent of the oxidation was evaluated by analyzing the amount of
gluconic acid produced using HPLC.
5.3.2.3 Detection of gluconic acid by HPLC
The oxidized product (i.e. gluconic acid) was diluted 5-10 times and then 5 μL were
analyzed using an ICS5000 HPAEC equipped with a pulsed amperometric detector
(Dionex, USA) and CarboPac PA1 column coupled to a guard column. The eluents were
0.1 M NaOH (eluent A) and 1 M NaOAc (eluent B), and elution was performed at 30 °C
with a constant flow rate of 0.25 mL/min using a linear gradient of 0% to 10% eluent B
over 10 min, followed by a linear gradient of 10% to 30% eluent B over 15 min, and
finally from 30% to 100% eluent B over 5 min. Reconditioning of the column was
achieved by running the initial conditions (i.e. eluent A) for 10 min. This resulted in a
total runtime of 30 min.
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Chromatograms were recorded and the peak area corresponding to gluconic acid was
calculated using Chromeleon 7.2 (Dionex, USA); gluconic acid solutions (0.01-0.5 mM)
were used as standards to correlate the peak area with the gluconic acid concentration.
5.3.2.4 Monitoring the oxygen content using Oxygraph
Oxygen concentrations in reaction mixtures were measured using an Oxygraph
(Hansatech, USA) equipped with a 10 mL chamber and attached to a circulating water
bath to maintain reaction temperatures at 25 °C. Briefly, 500 μL of the reaction buffer
containing 10 mM glucose with 0-2 mM H2O2 were equilibrated to 25 °C, and then the
reaction was initiated by adding 10 μL of either GO or GOOX containing approximately
16 pmol of the enzymes to the reaction chamber using a Hamilton syringe. The oxygen
content was measured continuously and enzyme action was evaluated by determining
oxygen consumption rates using the Oxygraph Plus software (Hansatech, USA).
5.3.3 Results and Discussion
5.3.3.1 GOOX-Y300A shows higher oxidation of oligosaccharides
Specific activity comparison shows wider substrate specificity for GOOX-Y300A
compared to GO. The oxidation activity of GO on maltose, cellotetraose, and
xylooligosaccharides were over two orders of magnitude less than that measured using
similar molar quantities of GOOX-Y300A. The activity on xylose and cellobiose was
more than 30 times lower with GO whereas on glucose, the oxidation activity of GO was
roughly 3 times higher than that of GOOX-Y300A (Table 5.2). The different substrate
preferences of GOOX-Y300A and GO is likely attributed to the open active site of
GOOX (Huang et al. 2005; Foumani et al. 2011) compared with the deep binding pocket
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of GO which is specifically suited for glucose (Hecht et al. 1993). The activity of GO
reported herein is consistent with an earlier report by Kulys et al. (2001a), which show
weak activity of GO on xylose, maltose, and cellobiose compared to MnCO. Moreover,
due to the reduced substrate inhibition by GOOX-Y300A (Vuong et al. 2013), the current
study suggests that GOOX-Y300A may be particularly advantageous for oxidation of
maltose, as well as xylo- and cello-oligosaccharides generated by cellulases and
xylanases, which are also commonly used enzyme additives in baking applications.
Table 5.2 Specific activities of GOOX-Y300A and GO on selected mono- and oligosaccharides.
a errors represent standard deviations (n=3). b specific activities were calculated in mole basis considering the MW of a monomeric GO (80 KD)
5.3.3.2 H2O2 inactivation
Inactivation of GO by H2O2 has been a bottleneck for industrial applications of GO and
so GO is typically used in combination with catalase, which continuously degrades H2O2.
The requirement for a second enzyme has implications to enzyme cost as well as
environmental benefits of the enzyme approach (Pourbarfani et al. 2004). Thus, an
alternative enzyme with lower H2O2 inactivation is desirable.
Substrates Specific activity (U/µmol)
GOOX-Y300A GO
Glucose 200 ± 20 a 558 ± 7 b
Xylose 270 ± 10 5 ± 2
Maltose 640 ± 30 3.2 ± 0.4
Cellobiose 1000 ± 60 28 ± 3
Cellotetraose 1100 ± 50 0.5 ± 0.2
Mixed Xylooligos 420 ± 10 1.3 ± 0.1
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Studying H2O2 inactivation of oxidases is challenging since most activity assays for this
class of enzymes detect the production of H2O2 as a by-product of the reaction. Using an
oxygen probe to detect oxygen as the co-substrate of GO, Bao et al. (2003) measured
oxygen consumption as a function of GO activity, and in this way, evaluated GO activity
in the presence of increasing H2O2 concentrations.
With a similar approach, an oxygraph was used to measure rates of glucose oxidation by
GO and GOOX-Y300A in the presence of H2O2. As reported in Appendix 3, the analysis
of GO in the presence of H2O2 was consistent with that reported by Bao et al (2003).
Interestingly however, using the oxygraph to measure GOOX-Y300A activity showed an
increase in oxygen concentrations when 2 mM H2O2 was added to the reaction mixture.
This result is presently difficult to explain, but will be further evaluated in future studies.
For the current investigation, high-performance anion-exchange chromatography
(HPAEC) was subsequently used to further compare the impact of H2O2 of GO and
GOOX-Y300A activity.
Liquid chromatography enables the quantitative analysis of oxidized reaction products.
Accordingly, HPAEC was used to measure the concentration of gluconic acid in
reactions containing 1 mM glucose, 0 to 0.2 M H2O2, and either GO or GOOX. Although
the rate of glucose oxidation by GOOX-Y300A is lower than that of GO, the current
analysis revealed that GOOX-Y300A might be slightly more stable than GO in the
presence of high concentration of H2O2 i.e. 0.2 M (Figure 5.9; Table 5.3).
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Figure 5.9 H2O2 tolerance of GOOX-Y300A and GO. Percent activity of the enzymes in the presence of various amount of H2O2.
Table 5.3 Amount of gluconic acid produced by GOOX-Y300A or GO in the presence of various concentrations of H2O2.
a errors represent standard deviations (n=3).
The difference in reduction of the activity of GOOX-Y300A and GO with increased
concentration of H2O2 was statistically significant as evaluated using the SlopesTest
function in Excel (Real Statistics Resource Pack) where the log of the data is taken to
H2O2 (mM) Gluconic acid production (mM) Percent activity
GOOX-Y300A GO GOOX-Y300A GO
0 0.16 ± 0.01 a 0.81 ± 0.1 100 100
25 0.14 ± 0.01 0.63 ± 0.11 83 78
50 0.12 ± 0.02 0.59 ± 0.05 74 73
100 0.1 ± 0.01 0.39 ± 0.11
59 48
150 0.09 ± 0.02 0.31 ± 0.09 52 39
200 0.09 ± 0.02 0.26 ± 0.07 54 32
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generate a linear trend (Figure S5.2) and the corresponding slopes are statistically
compared. This difference is notable when considering the fact that GOOX-Y300A does
not contain catalase while the GO sample used herein is reported to contain
approximately 10 U/mg of catalase activity. Our analysis of GO is consistent with those
report by Kleppe (1996), who show approximately 20% residual GO activity after
exposure to 0.2 M H2O2. Notably, the GO used by Kleppe contained less than 5 U/mg
catalase activity.
It is worth mentioning that for GOOX-Y300A, reduction in activity upon exposure to
H2O2 was not observed in reactions that proceed for only 30 min, or when 0.4 µg of
GOOX-Y300A was used in reactions rather than 0.1 µg. This could be explained by
incomplete oxidation of glucose after 30 min, and perhaps increased stability of GOOX-
Y300A enzyme at higher concentrations. Overall, it is anticipated that the relative
stability of GOOX-Y300A along with its wider substrate specificity will make this
enzyme a suitable substitute to GO in baking applications.
5.3.4 Conclusions
The findings of the current study demonstrate that in dough formulations where
oligosaccharides are in high abundance, GOOX-Y300A may be a better alternative to GO
for the production of aldonic acids and H2O2. Moreover, lower inactivation of GOOX-
Y300A by H2O2 compared to GO is anticipated to reduce requirements for catalase or
protective encapsulation, which could lead to cost savings.
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Chapter 6 : Conclusions
Enzymes are excellent “green” catalysts in that they are site-specific, regio-selective, and
more importantly can be fine-tuned through protein engineering. The main objective of
my PhD study was built upon this latter property of enzymes: tunable catalysts through
molecular engineering. Accordingly, my aim was to use molecular biology techniques to
engineer enzymes from the AA7 family of oligosaccharide oxidase, and in particular the
glucooligosaccharide oxidase from Sarocladium strictum (GOOX), to enhance its activity
on insoluble and high molecular weight polysaccharides.
6.1 Rational design altered the substrate preference of the GOOX
The results presented in chapter 3 validated the hypothesis that site-directed mutagenesis
of residues predicted to determine substrate preference of GOOX could broaden the
specificity of this enzyme. In particular, the characterization of Y300A and Y300N
variants showed over 2-fold increase in catalytic performance of the enzyme on all tested
mono and oligo-saccharides, as well as gain of activity on mannose. Interestingly, the
Y300 mutation was later shown to result in reduced substrate inhibition compared to the
wild-type GOOX (Vuong et al. 2013). The W351F variant also displayed increased
enzyme efficiency on galactose proving the hypothesis that the low activity of GOOX on
substrates with axial OH group is due to the steric hindrance of this hydroxyl with the
tryptophan at 351position (Lee et al. 2005).
Despite the interesting trend in catalytic performance of the GOOX variants, and
improved binding behaviour on tested monosaccharides, the binding capacity of these
104
mutants was reduced on di- and tri- saccharides. Although this is a limitation for
applications of these variants on oligosaccharides, from a scientific point of view this
finding validated the hypothesis that the tyrosine and tryptophan at 300 and 351positions
are indeed involved in a stacking interaction with the second ring of disaccharide and
oligosaccharide substrates, being important residues for substrate binding by the GOOX
enzyme. Thus, the most significant contributions from this part of the study to the
scientific community were:
• To define GOOX residues that affect the substrate binding and preference of this
enzyme, giving clues for engineering of other AA7 enzymes.
• Production of valuable set of industrial enzymes, GOOX-Y300A and GOOX-
Y300N, with increased efficiency on glucose, xylose, galactose and a gain of
activity on mannose, as well as enhanced catalytic performance on
oligosaccharides and reduced substrate inhibition for applications where oxidation
of mono- and/or oligo-saccharides is required and where the oligosaccharide
concentration is not limiting.
6.2 CBM fusion promoted GOOX binding and activity towards polysaccharides
The results presented in chapter 4 demonstrated that fusion of selected CBMs with
affinity towards plant polysaccharides to GOOX could increase the binding and activity
of this enzyme on the corresponding polysaccharides. Specifically, GOOX fusion to
CtCBM3, CtCBM11 and CtCBM44, resulted in a significant enhancement in binding
capacity of GOOX towards Avicel, RAC, β-glucan and xyloglucan depending on the
selectivity of the appended modules. The activity of the fusion enzymes however, was
105
moderately enhanced on certain polysaccharides that contain no branching substitutions
and are linked through β-1->4 glycosidic bonds; namely glucomannan, RAC and Avicel.
This observation suggests that although the appended CBM increases the proximity of the
catalytic module and targeted substrate, favourable interaction with the enzymes active
site remain a bottleneck. Overall, the important breakthroughs and new insights from this
part of the study are as follows:
• Creation of a stable and active cellulose-immobilized CtCBM3-GOOX system
with significant potential in cellulose-biosensor applications and for cost-effective
and easy purification of GOOX enzyme using CtCBM3 as a purification tag.
• The first report on GOOX action towards polysaccharides including glucomannan,
β-glucan, xyloglucan, CMC, RAC, and Avicel broadens our concept of substrate
range among family AA7 oligosaccharide oxidases.
• The effect of linker on catalytic activity and thermostability was a surprise,
underscoring the importance of careful consideration of linker sequences when
constructing CBM fusions to AA7 enzymes as well as other carbohydrate-active
enzymes.
• The correlation between CBM type on binding of GOOX to particular
polysaccharides confirms that the substrate selectivity of the CBM is retained
even when it is linked to a catalytic module with broad substrate selectivity.
6.3 Engineered and wild-type GOOX have potential application in biofuel and food
industries
106
The results demonstrated in chapter 5 showed that the engineered enzymes as well as the
oxidized products presented in this study can be applied in following applications:
1. The oxidized cellooligosaccharides can be used as sugar standards for the
characterization of LPMO enzymes.
2. The fusion enzyme with comparatively high oxidation activity on
oligosaccharides can enhance the nutraceutical value of those compounds.
3. Mutant GOOX with enhanced activity on mono- and oligosaccharides and
reduced substrate inhibition can potentially replace GO in baking applications.
In particular, the application of GOOX products as sugar standards was demonstrated by
NMR and mass spectrometry analysis of reaction products, which confirmed nearly
complete, regio-selective oxidation of hydroxyls at the anomeric carbon of
oligosaccharides regardless of sugar type or chain length of these compounds.
The GOOX oxidation however, did not enhance the prebiotic activity of compounds as
the GOOX-CBM44 oxidized XOS were shown to promote Bifidobacterium longum
proliferation to the same extents of XOS. However, this result in itself was interesting
when considered in light of the negative impacts of aldouronic acid from xylan with
respect to B. longum utilization. It was concluded that unlike aldouronic acids which
obtain the acidic functionality through substitution of gluconic acid to the XOS backbone,
the GOOX-CBM44 oxidized XOS with an aldonic acid group at the anomeric carbon can
still support the growth of B. longum while potentially promoting intestinal mineral
absorption as a result of its acidic functionality.
107
The GOOX variants was shown to be indeed a very good replacing candidate for GO in
baking applications as GOOX-Y300A showed broader substrate specificity as well as
exhibiting lower level of H2O2 inactivation than the commercial GO.
Accordingly the key findings and significant contributions are as follows:
• GOOX-oxidized oligosaccharides as sugar standards would benefit the
bioconversion research community by enabling detailed biochemical analysis of
LPMO enzymes, which enhance cellulose hydrolysis for biofuel and biochemical
production.
• GOOX treatment of xylooligosaccharides was shown to be an important means to
fine-tune the performance of prebiotics and introduce additional nutraceutical
value.
• The GOOX-Y300A variant with wide substrate range is a better alternative to GO
in baking applications, particularly for dough with high fibre content.
Taking one step back from the specific context of this dissertation, the findings from this
study are aligned with the ultimate goal of our research group to discover, characterize
and engineer carbohydrate active enzymes, which could be then applied as industrial
biocatalysts in the production of high-value bioproducts. In the present study the GOOX
enzyme from a fungal origin was recombinantly expressed in P. pastoris and was
successfully engineered with enhanced activity, binding and stability performance.
Moreover, applications has been sought and studied which led to potential industrial
collaborations. Encouragingly, the outcomes from the present study have shown
108
opportunities for continuation or being branched to new projects. Accordingly the next
section will discuss the future recommendations.
109
Chapter 7 : Future directions
The following paragraphs propose future experiments, which could enable the transfer of
GOOX research to industrial practice.
7.1 Incorportaion of GOOX-oxidized oligosaccharides in an LPMO standard assay.
Studies presented in section 5.1 suggest that GOOX oxidized cellooligosaccharides can
be applied as sugar standards for LPMO assays. In particular, the spectrometry results
confirmed that the structure of oxidized oligosaccharides generated by GOOX is the same
as the most abundant product of LPMO action on cellulose. In order to implement this
concept, the GOOX oxidized cellooligosaccharides need to be purified from the oxidation
reaction mixture, and incorporated in a currently used LPMO assay. The LMPO activity
should be then measured using the GOOX products as standards as well as chemically
synthesized oxidized cellooligosaccharides. A direct comparison will advise the practical
feasibility of this application.
7.2 Effect of debranching enzyme on prebiotic activity of polysaccharides
In section 5.2 it was shown that introducing aldonic acid to the reducing end of XOS
using CBM44-GOOX supports proliferation of B. longum, whereas substiution of
gluconic acid on XOS, as presented in aldouronic acids, prevents digestion by B.
longum. It was suggested that the gluconic acid side-chains in aldouronic acid are
responsible for inability of B. longum to utilize this compound. Accordingly, it would be
interesting to remove the side-chains of aldouronic acid and its polysaccharide,
glucuronoxylan, with the aid of debranching enzymes, specifically α-glucoronidases, and
110
to test the effect of these enzyme treatments on the growth of Bifidobacterium cultures.
With that in mind, an α-glucoronidase have been already expressed and purified by a PhD
candidate in our lab, Ruoyu Yan, and following prebiotic experiments is to be conducted
as part of her PhD project. In addition, a study on the effect of oxidized XOS as well as
other GOOX-oxidized oligosaccharides on mineral absoption ought to be performed to
confirm the nutritional benefit of these products.
7.3 Additional value of a CBM fusion GOOX-Y300A for baking application
The results presented in section 5.3 suggest that GOOX-Y300A is suitable to replace GO
in baking applications due to its wider substrate specificity and lower H2O2 inactivation.
However, in this study the free enzyme has been studied whereas industrial applications
typically use immobilized enzyme for easier recycling and higher stability under
operational conditions. Lin et al. (1996) show that immoblization of the wild-type
GOOX-T1 led to significant enhancement in thermal stability and overall performance of
the enzyme. Accordingly, it is suggested to investigate the immobliziation of GOOX-
Y300A which is fused to a CBM specific to a diatery fibre. This way, the fibre -
immobilized enzyme would be added as a baking ingredient.
So far, the fusion enzyme CtCBM22_GOOX-Y300 containing the xylan specific
CtCBM22 has been constructed, expressed and purified by the post doctoral fellow in our
lab, Dr. Vuong; Ben MacCormick is studying the immobilization of this enzyme to oat
spelt xylan, and is assessing the coresponding effects on thermal stability and H2O2
tolerance of this enzyme. In addition, he is investigating the unique behavior of GOOX in
the presence of H2O2 (Figure S5.2) using HPAEC method as descriebed in section 5.3.2.3
with both free and immobilied enzymes to confirm this activation behavior. Ben
111
MacCormick is interested to also test the possibility of purifiying the enzyme with oat
spelt xylan due to the strong affinity of CtCBM22 towards this polysaccharide (Vuong
and Master 2014). This will be an additonal value for intorducing the GOOX variants in
baking applications.
112
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Appendix 1: Supplemental information for chapter 3
Figure S3.1 Multiple sequence alignment of GOOX homologues. The alignment between MnCO (CAI94231-2) from Microdochium nivale, ChitO (XP_391174) from Fusarium graminearum, and GOOX (noted as GOOX-VN) was generated by Megalign (DNASTAR-Lasergene). Amino acids, which were mutated, are highlighted with asterisks.
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Figure S3.2 Stability of wild-type and mutant GOOX at 37°C. Residual activity of GOOX (circle), W351F (square), Y300A (cross) and Y300N (triangle) enzymes on 10 mM maltose after incubation at 37°C
in triplicate for up to 1 h.
Figure S3.3 SDS-PAGE of deglycosylated GOOX and mutant enzymes. SDS-PAGE was performed using a 12 % polyacrylamide gel and proteins were stained with Coomasie Blue. Deglycosylated samples were indicated with asterisks. Supernatant from P. pastoris containing pPICZαA (V) was also included. PageRulerTM Plus prestained protein ladder (Fermentas) was used.
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Appendix 2: Supplemental information for chapter 4
Figure S4.1 Purified wild-type and fusion GOOX proteins on 10% SDS-PAGE. Lane1: PageRuler protein ladder; 2: CtCBM3_GOOX; 3: CtCBM11_GOOX; 4: CtCBM44_GOOX; 5: wild-type GOOX, 6: GOOX_CtCBM3; 7: GOOX_CtCBM11; 8: GOOX_CtCBM44.
Figure S4.2 Binding of wild-type GOOX and CBM fusions to insoluble cellulose as analyzed by SDS-PAGE. Purified proteins were incubated with crystalline cellulose (Avicel) or regenerated amorphous cellulose (RAC) for 2 h on ice with continuous shaking. Unbound (U) and bound (B) protein fractions are shown.
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Figure S4.3 Specific activity of wild-type GOOX and CBM fusions on konjac glucomannan A. 0.1 %, B. 0.3%. All reactions contained 0.5 µg of enzyme. Error bars represents standard deviations; n=3. The doted line represents the specific activity of wild-type GOOX.
Figure S4.4 Adsorbed mass of wild-type GOOX and CtCBM3_GOOX on cellulose-coated sensors. Changes in adsorbed mass during 1.5 µg/mL enzyme addition (1), 50 mM Tris-HCl pH 8 buffer washing (2) and 0.5 mM cellobiose addition (3) in the experiments with CtCBM3_GOOX (green, solid line) and wild-type GOOX (red, dashed line). Mass values were obtained using the Voigt model.
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Figure S4.5 Cellobiose oxidation of enzyme-bound sensors. QCM-D sensors that were previously bound with CtCBM3_GOOX were repeatedly washed and incubated with 0.5 mM cellobiose, and the regeneration of oxidized products was measured by the chromogenic assay.
Figure S4.6 Thermostability of proteins at 45°C. A: wild-type GOOX, B: CtCBM3_GOOX, C: CtCBM11_GOOX, D: CtCBM44_GOOX, E: GOOX_CtCBM3, F: GOOX_CtCBM11, G: GOOX_CtCBM44, H: logarithmic presentation of the graphs used to determine the half-lives.
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Appendix 3: Supplemental information for chapter 5
Observations in experiments using Oxygraph
As mentioned in section 5.3.3.2 the H2O2 inactivation of GO and GOOX-Y300A was
first attempted to be measured by monitoring the oxygen concentration using Oxygraph.
Depending on the susceptibility of the enzyme to H2O2, the rate of oxygen depletion
would decrease with increased addition of H2O2. Notably, the oxygen concentration was
increased when GOOX-Y300A was added in the presence of 0.2 mM H2O2 (Figure S5.1)
resulting in oxygen production instead of oxygen consumption. This phenomenon was
not observed when GO, or solely the buffer was added to the reaction containing H2O2.
To the same reaction when catalase was added, a spontaneous increase of oxygen level
was observed as a result of decomposition of H2O2. Although the increase level of
oxygen with GOOX-Y300A was not as sharp as that with catalase, the similar trend
observed is interesting. These results suggest that GO and GOOX-Y300A will show
different behavior at the presence of H2O2. Further experiments are being designed to
propose a mechanism for explaining this behavior, which is described in the future
direction section.
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Figure S5.1 Behavior of GOOX-Y300A in the presence (A) or absence (B) of H2O2. Reactions contain 10mM glucose, 16 pmole enzyme in Tris-HCl pH 8. The arrows show the time point where enzyme is added.
Figure S5.2 - Log of GluA concentration produced by GOOX-Y300A or GO versus H2O2 concentration.