Biomass, Biofuels, Biochemicals: Advances in Enzyme Catalysis and Technologies [1 ed.] 0128198206, 9780128198209

Table of contents :
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Biomass, Biofuels, Biochemicals: Advances in Enzyme Catalysis and
Technologies
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Contents
List of contributors
Preface_2020_Biomass--Biofuels--Biochemicals
Preface
Section 1: Fundamental aspects of enzymes
1 An introduction to enzyme structure dynamics and enzyme catalysis
1.1 Introduction
1.2 Why do some proteins work as enzyme?
1.2.1 Role of the active site in enzyme functioning
1.2.2 Cofactor, a necessity of enzyme
1.3 What is enzyme catalysis?
1.3.1 Basics of enzyme catalysis
1.3.2 Historical overview of enzyme catalysis theory
1.3.3 Mechanistic view of enzyme catalysis
1.4 Structural dynamics of enzymes
1.5 Ribozymes as a nonprotein catalyst
1.6 Conclusions and perspective
References
2 Classification of enzymes and catalytic properties
2.1 Introduction
2.1.1 Enzyme nomenclature and classification
2.1.2 Enzyme databases
2.1.3 Catalytic properties
2.2 Enzymes classes and properties
2.2.1 Oxidoreductases
2.2.2 Transferases
2.2.3 Hydrolases
2.2.3.1 Amylases
2.2.3.2 Cellulases
2.2.3.3 Xylanases
2.2.3.4 Lipases
2.2.3.5 Proteases
2.2.4 Lyases
2.2.5 Isomerases
2.2.6 Ligases
2.2.7 Translocases
2.3 Conclusions and perspectives
References
Further reading
3 Enzymes and their production strategies
3.1 Introduction
3.2 Enzymes and their classifications
3.3 Enzyme production
3.3.1 Microbial enzyme production
3.3.1.1 Fermentation
3.3.1.2 Recovery
3.3.1.3 Purification
3.3.1.4 Formulation
3.3.2 Enzyme production from plants and animals
3.4 Applications
3.5 Conclusions and perspectives
Acknowledgments
References
4 Robust enzymes designing for efficient biocatalysis
4.1 Introduction
4.2 Biocatalysis engineering—a driving force
4.3 Recent advances in protein engineering
4.3.1 Enzyme immobilization—a drive toward optimum performance
4.3.2 Substrate engineering—a toolkit to harness the enzymatic promiscuity
4.3.3 Structure-assisted protein engineering
4.3.4 Advanced computational modeling
4.3.5 Protein engineering via directed evolution and rational design
4.4 Conclusions and perspectives
4.5 Acknowledgments
4.6 Conflict of interest
References
Section 2: Enzyme engineering for efficient biocatalytic reactions
5 Enzyme engineering strategies to confer thermostability
5.1 Introduction
5.2 Enzyme engineering strategies for thermostabilization
5.2.1 Directed evolution
5.2.2 Rational/semi-rational design
5.2.2.1 Sequence-based engineering
5.2.2.1.1 Comparing sequences with higher thermostability
5.2.2.1.2 Consensus analysis
5.2.2.1.3 Ancestral sequence reconstruction
5.2.2.2 Structure-based engineering
5.2.2.2.1 Beta-factor analysis
5.2.2.2.2 Molecular dynamic simulations
5.2.2.2.3 FoldX and Rosetta_ddg
5.2.2.2.4 Structure-guided sequence-based engineering
5.2.3 De novo design
5.2.4 Comprehensive computational approaches
5.2.4.1 Computational disulfide engineering
5.2.4.2 FRESCO framework
5.2.4.3 FireProt framework
5.3 Conclusions and perspectives
Abbreviations
References
6 Enzyme engineering strategies for catalytic activity in wide pH range
6.1 Introduction
6.2 Bioprospecting of extreme ecological niche for pH stable enzymes
6.2.1 Molecular mechanism of enzyme stability in the wide pH range
6.2.2 Directed evolution of enzyme
6.2.3 Screening strategies for evolved enzymes
6.3 A computational approach for enzyme improvement
6.3.1 Rational design method
6.3.2 De novo design method
6.4 Conclusions and perspective
6.5 Acknowledgments
References
7 Protein engineering approaches for enhanced catalytic efficiency
7.1 Introduction
7.2 Approaches in protein engineering
7.2.1 Rational design
7.2.1.1 Multiple sequence alignment
7.2.1.2 Structure-guided design
7.2.1.3 Computational molecular simulation
7.2.2 Directed evolution
7.2.2.1 Random mutagenesis
7.2.2.2 DNA shuffling
7.2.2.3 In vivo continuous directed evolution
7.2.3 Semi-rational design
7.2.4 De novo protein design
7.3 Advanced technologies assisting protein engineering
7.3.1 Spatial organization
7.3.2 Surface displaying system
7.3.3 Cell-free translation systems
7.3.4 High-throughput screening methods
7.3.5 Cryogenic electron microscopy
7.4 Conclusions and perspectives
References
8 Chimeric enzyme designing for the synthesis of multifunctional biocatalysts
8.1 Introduction
8.1.1 Natural chimeric proteins
8.1.2 Artificial or synthesized or recombinant chimeric proteins
8.2 Methods for generating chimeric/fusion proteins
8.2.1 End-to-end fusion
8.2.1.1 Linker
8.2.1.1.1 Naturally occurring linkers
8.2.1.1.2 Artificial or synthesized linker
Flexible linkers
Rigid linker
Cleavable linkers
In vivo cleavable linkers
Protease-sensitive cleavable linkers
Photo cleavable linker
8.2.1.1.3 Linker designing tools and databases
8.2.1.2 Overlap extension polymerase chain reaction or end-to-end cloning
8.2.2 Domain insertion
8.2.2.1 Construction of domain insertion libraries
8.2.3 Posttranslational conjugation
8.3 Applications of chimeric or fusion proteins
8.3.1 Use of multifunctional enzymes for biomass degradation
8.3.2 Fusion protein-based biopesticides (antimicrobial peptides)
8.3.3 Fusion protein as biopharmaceutical
8.3.4 Fusion protein-based biosensor (protein switching)
8.3.5 Other applications
8.4 Conclusions and perspectives
References
9 Enzyme engineering for enantioselective biotransformations
9.1 Introduction
9.1.1 Enantioselective biotransformations
9.1.2 Enzymes and chirality: the relationship between structure and function
9.1.3 Tools for modifying the enantioselectivity of enzymes
9.2 Directed evolution: a Darwinian approach for tailoring the enantioselectivity of enzymes
9.2.1 Mutant library construction
9.2.1.1 Random approaches
9.2.1.1.1 Error-prone polymerase chain reaction
9.2.1.1.2 Gene shuffling
9.2.1.2 Semi-rational design
9.2.1.2.1 Iterative saturation mutagenesis
9.2.1.2.2 Combinatorial active-site saturation test
9.2.2 High-throughput screening of enantioselective enzymes
9.2.2.1 Chromogenic/fluorogenic substrate surrogate-based assays
9.2.2.2 pH indicator assays
9.2.2.3 Coupled enzyme assays
9.2.2.4 Agar plate–based assays
9.2.2.5 Fluorescence-activated cell/droplet sorting
9.2.2.6 Other assays
9.3 Enzyme engineering: examples in enantioselective biocatalysis
9.3.1 Enhancing enantioselectivity of P450pyr monooxygenase for the hydroxylation of nonactivated carbon atoms
9.3.2 Inverting the enantioselectivity of P450pyr monooxygenase
9.3.3 Engineering amine dehydrogenase for asymmetric reductive amination
9.4 Conclusions and perspectives
References
Section 3: Enzyme immobilization and bioprocess engineering
10 Nanobiocatalyst designing strategies and their applications in food industry
10.1 Introduction
10.2 Design of nanobiocatalyst through nano-immobilization
10.2.1 Adsorption binding of enzymes
10.2.2 Covalent binding of enzymes
10.2.3 Entrapment or encapsulation of enzymes
10.2.4 Cross-linking of enzymes
10.3 Nanobiocatalyst role in food processing with reference to omega-3 oil processing
10.4 Application of nanobiocatalyst in other food processing applications
10.5 Conclusions and perspectives
References
Further Reading
11 Enzyme entrapment approaches and their applications
11.1 Introduction
11.2 Immobilization of enzymes
11.3 History of enzyme immobilization
11.4 Advantages of enzyme immobilization
11.5 Support systems for enzyme conjugation
11.5.1 Organic support materials
11.5.2 Biopolymers
11.5.2.1 Alginate
11.5.2.2 Chitin and chitosan
11.5.2.3 Collagen
11.5.2.4 Carrageenan
11.5.2.5 Gelatin
11.5.2.6 Cellulose
11.5.2.7 Starch
11.5.2.8 Pectin
11.5.3 Hydrogels
11.5.4 Inorganic support materials
11.5.4.1 Zeolites
11.5.4.2 Ceramics
11.5.4.3 Celite
11.5.4.4 Silica
11.5.4.5 Glass
11.5.4.6 Activated carbon
11.5.4.7 Charcoal
11.5.5 Protein-coated microcrystals technique
11.5.6 Smart polymers
11.5.7 Conducting polymers
11.6 Types of immobilization techniques
11.6.1 Support matrix system
11.6.2 Entrapment system
11.6.3 Cross-linking system
11.7 Methods of enzyme immobilization
11.7.1 Physical methods
11.7.1.1 Physical adsorption
11.7.1.2 Ionic binding
11.7.1.3 Physical entrapment
11.7.2 Chemical methods
11.7.2.1 Cross-linking
11.7.2.2 Covalent binding
11.8 Applications of immobilized enzymes
11.8.1 Food industry
11.8.2 Pharmaceutical industry
11.8.3 Brewing industry
11.8.4 Detergent industry
11.8.5 Pulp and paper industry
11.8.6 Textile industry
11.8.7 Biomedical applications
11.9 Conclusions and perspectives
Acknowledgments
References
12 Enzyme immobilization strategies and bioprocessing applications
12.1 Introduction
12.2 Enzymes as industrial biocatalysts
12.2.1 Requirements, advantages, and limitations for the industrial enzymes
12.2.1.1 Stabilization of the enzymes
12.2.1.1.1 About the pH value
12.2.1.1.2 The contribution of temperature
12.2.1.1.3 Grouping methods for stabilization of enzymes
12.2.1.1.4 Use of protein engineering
12.2.1.2 Stabilization and functionality of biocatalysts in nonconventional solvents
12.3 Enzyme immobilization
12.3.1 Changes induced by enzyme immobilization: positive and negative
12.3.1.1 Features due to enzyme immobilization
12.3.1.2 Immobilization improves the activity of an enzyme
12.3.1.3 Improvement of specificity and selectivity of the immobilized enzyme
12.3.1.4 Enzyme stabilization through partitioning
12.3.1.5 Restrictions due to diffusional effects
12.3.2 Strategies, steps, and factors affecting the enzyme immobilization
12.3.2.1 Preimmobilization factors and actions
12.3.2.2 Beneficial steps of enzyme immobilization
12.3.3 Selected methods of enzyme immobilization
12.3.3.1 Adsorbing methods
12.3.3.2 Covalent bonding methods
12.3.3.3 Entrapment methods
12.3.3.4 Cross-linking methods
12.3.3.5 Materials used in manufacturing immobilization matrices
12.4 Applications of immobilized enzymes in various bioprocesses
12.4.1 Food, dairies, juices, cosmetics, pharmaceuticals, and others
12.4.2 Medical applications of immobilized enzymes and biosensors
12.4.3 Bioremediation methods and wastewater treatment
12.4.4 Valorization of food processing wastes—objectives and prospects
12.4.5 Immobilized enzymes in textile industry
12.4.6 Immobilized enzymes in the production and process of cosmetics
12.4.7 Bioprocessing through nanobiocatalysis
12.5 Conclusions and perspectives
References
Section 4: Enzymatic technologies, case studies, and future perspectives
13 Promising enzymes for biomass processing
13.1 Introduction
13.1.1 Biomass
13.1.2 Pretreatment
13.1.3 Biomass-processing enzymes
13.1.3.1 Cellulase
13.1.4 Cellulases for bioconversion
13.2 Strain improvement via mutation
13.2.1 Random mutagenesis
13.2.2 Site-directed mutagenesis
13.2.3 Genetic engineering
13.3 Cellulase accessory and auxiliary enzymes
13.3.1 β-Glucosidase
13.3.2 Xylanase
13.3.2.1 Hemicellulose-degrading enzymes
13.3.2.2 Genetic engineering of xylanases
13.3.3 Laccase
13.3.4 Auxiliary enzymes
13.3.4.1 Lytic polysaccharide monooxygenase
13.3.5 Market scenario
13.4 Conclusion and perspectives
References
14 Enzyme systems for high-value biomolecule production
14.1 Introduction
14.2 Enzymes involved in production of prebiotics
14.2.1 β-Galactosidase
14.2.2 β-Glucosidase
14.2.3 β-Glycosidase
14.2.4 Fructosyltransferase
14.2.5 β-d-Fructofuranoside
14.2.6 Inulinases
14.2.7 Glycoside hydrolases
14.3 Production of prebiotics
14.3.1 Galactooligiosaccharides
14.3.2 Lactulose
14.3.3 Tagatose
14.3.4 Lactosucrose
14.3.5 Fructooligosaccharides
14.3.6 Isomaltooligosaccharides
14.4 Downstream processing of prebiotics
14.5 Global status of prebiotics
14.6 Conclusions and perspectives
References
Further reading
15 Role of enzymatic bioprocesses for the production of functional food and nutraceuticals
15.1 Introduction
15.2 Role of enzymes in production of functional foods
15.2.1 Proteases
15.2.2 Carbohydrate-modifying enzymes
15.2.2.1 β-Galactosidases
15.2.2.2 Enzymes catalyzing fructooligosaccharides production
15.2.2.3 Enzymes catalyzing xylooligosaccharides production
15.2.2.4 Enzymes catalyzing resistant starch production
15.2.2.5 β-Glucosidases
15.2.3 l-Asparaginase
15.2.4 Lipases
15.2.5 Tannase
15.2.6 Phytase
15.3 Production of functional foods using specific enzyme-producing starters
15.4 Genetically modified enzymes in development of functional foods
15.5 Conclusions and perspectives
Acknowledgments
References
16 Biotransformations with crude enzymes and whole cells
16.1 Introduction
16.2 Biotransformations in the pharmaceutical industry
16.2.1 Chiral carboxylic acids
16.2.2 Chiral alcohols
16.2.3 Chiral amines and amino acids
16.2.4 Biosynthesis of natural products
16.3 Biotransformations in food processing
16.3.1 Hydrolysis of naturally occurring large molecules into smaller molecules
16.3.1.1 Trypsin
16.3.1.2 Amylases
16.3.2 Biotransformation of chemicals
16.3.2.1 Glucose isomerase
16.3.2.2 Prebiotics
16.3.2.3 Production of food chemicals by synthetic biology
16.4 Biotransformations in the biofuels industry
16.4.1 Classification and biotransformation of biofuels
16.4.1.1 First-generation biofuels
16.4.1.2 Second-generation biofuels
16.4.1.3 Third-generation biofuels
16.4.1.4 Fourth-generation biofuels
16.4.2 Important examples of biofuels
16.4.2.1 Bioethanol
16.4.2.2 Biodiesel
16.4.2.3 Biohydrogen
16.5 Conclusions and perspectives
Acknowledgments
References
17 Enzymes in the third generation biorefinery for macroalgae biomass
17.1 Introduction
17.2 Biorefinery and biorefining
17.2.1 First and second generation biorefineries
17.2.2 Third generation biorefinery
17.3 Macroalgae as a source of high-value products
17.4 Enzyme production
17.4.1 Solid-state fermentation and semi-solid-state fermentation for enzyme production
17.4.2 Green algae as source for enzyme production
17.4.2.1 Brown algae as source for enzyme production
17.4.2.2 Red algae as source for enzyme production
17.5 Enzymes for biofuels
17.5.1 Enzymes in the bioethanol production
17.5.1.1 Genetic engineering for bioethanol and biofuel production
17.5.2 Enzymatic saccharification of green macroalgae
17.5.2.1 Enzymatic saccharification of brown macroalgae
17.5.2.2 Enzymatic saccharification of red macroalgae
17.5.3 Saccharification by enzymatic complexes, cocktails, and simultaneous fermentation
17.5.4 Biobutanol and acetone, butanol, and ethanol fermentation
17.5.4.1 Enzymatic engineering for acetone, butanol, and ethanol fermentation
17.5.5 Enzymes in biogas production
17.6 Enzymatic extraction of high valued products
17.6.1 Extraction of hydrocolloids
17.6.1.1 Extraction of phenolic compounds
17.6.2 Extraction of pigments
17.7 Conclusions and perspectives
Acknowledgment
References
18 Enzymatic systems for the development of juice clarification strategies
18.1 Introduction
18.2 Why juice clarification?
18.3 Juice clarification strategies
18.3.1 Mechanical process
18.3.1.1 Straining or screening
18.3.1.2 Centrifugation and finishing
18.3.1.3 Clarification by freezing and heating
18.3.1.4 Physical finings
18.3.2 Chemical process
18.3.3 Enzymatic process
18.4 Enzymes used for juice clarification
18.4.1 Pectic enzymes
18.4.1.1 Pectinesterase
18.4.1.2 Polygalacturonase
18.4.1.3 Pectinlyase
18.4.2 Cellulase
18.4.3 Laccase
18.4.4 Amylase
18.4.5 Others
18.5 Conclusions and perspectives
Acknowledgments
References
19 Biocatalyst systems for xylooligosaccharides production from lignocellulosic biomass and their uses
19.1 Introduction
19.2 Xylan: sources and structures
19.3 Xylooligosaccharides
19.4 Production of xylooligosaccharides
19.4.1 Xylanolytic enzymes
19.4.2 Purification of xylooligosaccharides
19.5 Biological properties of xylooligosaccharides
19.6 Xylooligosaccharides applications
19.7 Market and safety aspects of xylooligosaccharides
19.8 Conclusion and perspectives
References
20 Biocatalysis for cascade reactions to produce high-value chemicals
20.1 Introduction
20.1.1 What is a cascade reaction?
20.1.2 Advantages of cascade biocatalysis
20.1.3 Production of high-value chemicals via cascade biocatalysis
20.2 Designs of biocatalytic cascade reactions
20.2.1 Topology of cascade
20.2.1.1 Linear cascade
20.2.1.2 Cyclic cascade
20.2.1.3 Orthogonal cascade
20.2.1.4 Coupled cascade
20.2.1.5 Convergent and divergent cascade
20.2.1.6 Modularized cascade
20.2.2 Pathway design and reaction engineering
20.3 Biocatalysts
20.3.1 Enzymes
20.3.2 Whole-cell
20.3.3 Fused enzymes
20.3.4 Immobilized enzymes
20.4 Examples of cascade biocatalysis for the production of valuable chemicals
20.4.1 Cascade biocatalysis from styrenes
20.4.2 Cascade biocatalysis from l-phenylalanine
20.4.3 Cascade biocatalysis for the synthesis of amines
20.4.3.1 Cascade through alcohol oxidation and reductive amination
20.4.3.2 Cascade through alcohol activation and nucleophilic substitution
20.4.3.3 Cascade through imine formation and asymmetric C–N double bond reduction
20.4.4 Cascade biocatalysis for the synthesis of lactones
20.4.4.1 Cyclic alcohols
20.5 Conclusions and perspectives
References
Further reading
Index
Back Cover

Citation preview

Biomass, Biofuels, Biochemicals Advances in Enzyme Catalysis and Technologies

Series Editor Ashok Pandey Centre for Innovation and Translational Research, CSIR-Indian Institute of Toxicology Research, Lucknow, India

Biomass, Biofuels, Biochemicals Advances in Enzyme Catalysis and Technologies Edited by

Sudhir P. Singh Center of Innovative and Applied Bioprocessing, Mohali, India

Ashok Pandey Centre for Innovation and Translational Research, CSIR-Indian Institute of Toxicology Research, Lucknow, India

Reeta Rani Singhania Center for Energy and Environmental Sustainability-India, Lucknow, India

Christian Larroche Chemical and Biochemical Engineering Laboratory, Institute Pascal, University Clermont Auvergne, Clermont Ferrand, France

Zhi Li Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore

Elsevier Radarweg 29, PO Box 211, 1000 AE Amsterdam, Netherlands The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States Copyright © 2020 Elsevier B.V. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress ISBN: 978-0-12-819820-9 For Information on all Elsevier publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Susan Dennis Acquisitions Editor: Ki Kostas Marinakis Editorial Project Manager: Andrea Dulberger Production Project Manager: R. Vijay Bharath Cover Designer: Greg Harris Typeset by MPS Limited, Chennai, India

Contents List of contributors .............................................................................................................................. xv Preface ................................................................................................................................................ xxi

SECTION 1 FUNDAMENTAL ASPECTS OF ENZYMES CHAPTER 1 An introduction to enzyme structure dynamics and enzyme catalysis ................................................................................................ 3 Jitesh Kumar, Ashok Pandey and Sudhir P. Singh 1.1 Introduction ................................................................................................................ 3 1.2 Why do some proteins work as enzyme? .................................................................. 3 1.2.1 Role of the active site in enzyme functioning ................................................ 4 1.2.2 Cofactor, a necessity of enzyme...................................................................... 4 1.3 What is enzyme catalysis? ......................................................................................... 4 1.3.1 Basics of enzyme catalysis .............................................................................. 4 1.3.2 Historical overview of enzyme catalysis theory ............................................. 5 1.3.3 Mechanistic view of enzyme catalysis ............................................................ 5 1.4 Structural dynamics of enzymes ................................................................................ 6 1.5 Ribozymes as a nonprotein catalyst........................................................................... 7 1.6 Conclusions and perspective ...................................................................................... 8 References................................................................................................................... 8

CHAPTER 2 Classification of enzymes and catalytic properties .......................... 11 Luciana Porto de Souza Vandenberghe, Susan Grace Karp, Maria Giovana Binder Pagnoncelli, Matheus von Linsingen Tavares, Nelson Libardi Junior, ´ Kim Valladares Diestra, Jessica Aparecida Viesser and Carlos Ricardo Soccol 2.1 Introduction .............................................................................................................. 11 2.1.1 Enzyme nomenclature and classification ...................................................... 11 2.1.2 Enzyme databases .......................................................................................... 12 2.1.3 Catalytic properties ........................................................................................ 12 2.2 Enzymes classes and properties ............................................................................... 14 2.2.1 Oxidoreductases ............................................................................................. 15 2.2.2 Transferases.................................................................................................... 16 2.2.3 Hydrolases ...................................................................................................... 17 2.2.4 Lyases............................................................................................................. 21 2.2.5 Isomerases ...................................................................................................... 22 2.2.6 Ligases............................................................................................................ 23 2.2.7 Translocases ................................................................................................... 24

v

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2.3 Conclusions and perspectives .................................................................................. 25 References................................................................................................................. 25 Further reading ......................................................................................................... 30

CHAPTER 3 Enzymes and their production strategies .......................................... 31 3.1 3.2 3.3

3.4 3.5

Himanshu Sharma and Santosh Kumar Upadhyay Introduction .............................................................................................................. 31 Enzymes and their classifications ............................................................................ 32 Enzyme production .................................................................................................. 33 3.3.1 Microbial enzyme production........................................................................ 34 3.3.2 Enzyme production from plants and animals................................................ 40 Applications.............................................................................................................. 41 Conclusions and perspectives .................................................................................. 42 Acknowledgments .................................................................................................... 43 References................................................................................................................. 43

CHAPTER 4 Robust enzymes designing for efficient biocatalysis ....................... 49 4.1 4.2 4.3

4.4 4.5 4.6

Roberto Parra-Saldivar, Ricardo A. Ramirez-Mendoza, Ashutosh Sharma, Goldie Oza, Ricardo Zavala-Yoe and Hafiz M.N. Iqbal Introduction .............................................................................................................. 49 Biocatalysis engineering—a driving force .............................................................. 50 Recent advances in protein engineering .................................................................. 51 4.3.1 Enzyme immobilization—a drive toward optimum performance ................ 51 4.3.2 Substrate engineering—a toolkit to harness the enzymatic promiscuity..................................................................................................... 55 4.3.3 Structure-assisted protein engineering........................................................... 56 4.3.4 Advanced computational modeling ............................................................... 58 4.3.5 Protein engineering via directed evolution and rational design ................... 58 Conclusions and perspectives .................................................................................. 59 Acknowledgments .................................................................................................... 60 Conflict of interest ................................................................................................... 60 References................................................................................................................. 61

SECTION 2 ENZYME ENGINEERING FOR EFFICIENT BIOCATALYTIC REACTIONS CHAPTER 5 Enzyme engineering strategies to confer thermostability ................ 67 Zhe Xu, Ya-Ping Xue, Shu-Ping Zou and Yu-Guo Zheng 5.1 Introduction .............................................................................................................. 67 5.2 Enzyme engineering strategies for thermostabilization .......................................... 68 5.2.1 Directed evolution.......................................................................................... 69

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5.2.2 Rational/semi-rational design ........................................................................ 72 5.2.3 De novo design............................................................................................... 78 5.2.4 Comprehensive computational approaches ................................................... 79 5.3 Conclusions and perspectives .................................................................................. 82 Abbreviations............................................................................................................ 83 References................................................................................................................. 83

CHAPTER 6 Enzyme engineering strategies for catalytic activity in wide pH range................................................................................. 91 Jitesh Kumar, Girija Kaushal and Sudhir P. Singh 6.1 Introduction .............................................................................................................. 91 6.2 Bioprospecting of extreme ecological niche for pH stable enzymes...................... 92 6.2.1 Molecular mechanism of enzyme stability in the wide pH range ................................................................................................ 92 6.2.2 Directed evolution of enzyme ....................................................................... 93 6.2.3 Screening strategies for evolved enzymes .................................................... 95 6.3 A computational approach for enzyme improvement ............................................. 96 6.3.1 Rational design method ................................................................................. 96 6.3.2 De novo design method ................................................................................. 98 6.4 Conclusions and perspective .................................................................................... 98 6.5 Acknowledgments .................................................................................................... 99 References................................................................................................................. 99

CHAPTER 7 Protein engineering approaches for enhanced catalytic efficiency ........................................................................... 103 Guoqiang Zhang, Yukun Chen and Guocheng Du 7.1 Introduction ............................................................................................................ 103 7.2 Approaches in protein engineering ........................................................................ 104 7.2.1 Rational design............................................................................................. 104 7.2.2 Directed evolution........................................................................................ 106 7.2.3 Semi-rational design .................................................................................... 108 7.2.4 De novo protein design ................................................................................ 109 7.3 Advanced technologies assisting protein engineering........................................... 109 7.3.1 Spatial organization ..................................................................................... 109 7.3.2 Surface displaying system ........................................................................... 111 7.3.3 Cell-free translation systems........................................................................ 111 7.3.4 High-throughput screening methods............................................................ 111 7.3.5 Cryogenic electron microscopy ................................................................... 112 7.4 Conclusions and perspectives ................................................................................ 112 References............................................................................................................... 113

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CHAPTER 8 Chimeric enzyme designing for the synthesis of multifunctional biocatalysts ....................................................................................... 119 8.1

8.2

8.3

8.4

Jyoti Singh Jadaun, Lokesh Kumar Narnoliya, Archana Srivastava and Sudhir P. Singh Introduction ............................................................................................................ 119 8.1.1 Natural chimeric proteins ............................................................................ 119 8.1.2 Artificial or synthesized or recombinant chimeric proteins........................ 120 Methods for generating chimeric/fusion proteins.................................................. 122 8.2.1 End-to-end fusion......................................................................................... 122 8.2.2 Domain insertion.......................................................................................... 129 8.2.3 Posttranslational conjugation ....................................................................... 132 Applications of chimeric or fusion proteins .......................................................... 133 8.3.1 Use of multifunctional enzymes for biomass degradation.......................... 133 8.3.2 Fusion protein-based biopesticides (antimicrobial peptides) ...................... 134 8.3.3 Fusion protein as biopharmaceutical ........................................................... 135 8.3.4 Fusion protein-based biosensor (protein switching) ................................... 136 8.3.5 Other applications ........................................................................................ 137 Conclusions and perspectives ................................................................................ 137 References............................................................................................................... 137

CHAPTER 9 Enzyme engineering for enantioselective biotransformations ............................................................................ 145 9.1

9.2

9.3

9.4

Kaiyuan Tian, Balaji Sundara Sekar, Joel Ping Syong Choo and Zhi Li Introduction ............................................................................................................ 145 9.1.1 Enantioselective biotransformations............................................................ 145 9.1.2 Enzymes and chirality: the relationship between structure and function.................................................................................................. 147 9.1.3 Tools for modifying the enantioselectivity of enzymes.............................. 148 Directed evolution: a Darwinian approach for tailoring the enantioselectivity of enzymes ................................................................................ 148 9.2.1 Mutant library construction ......................................................................... 149 9.2.2 High-throughput screening of enantioselective enzymes............................ 152 Enzyme engineering: examples in enantioselective biocatalysis.......................... 155 9.3.1 Enhancing enantioselectivity of P450pyr monooxygenase for the hydroxylation of nonactivated carbon atoms .................................. 156 9.3.2 Inverting the enantioselectivity of P450pyr monooxygenase ..................... 158 9.3.3 Engineering amine dehydrogenase for asymmetric reductive amination...................................................................................................... 159 Conclusions and perspectives ................................................................................ 161 References............................................................................................................... 161

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SECTION 3 ENZYME IMMOBILIZATION AND BIOPROCESS ENGINEERING CHAPTER 10 Nanobiocatalyst designing strategies and their applications in food industry............................................................ 171 Madan L. Verma, Reinu E. Abraham and Munish Puri 10.1 Introduction ............................................................................................................ 171 10.2 Design of nanobiocatalyst through nano-immobilization ..................................... 172 10.2.1 Adsorption binding of enzymes................................................................. 175 10.2.2 Covalent binding of enzymes .................................................................... 176 10.2.3 Entrapment or encapsulation of enzymes.................................................. 176 10.2.4 Cross-linking of enzymes .......................................................................... 177 10.3 Nanobiocatalyst role in food processing with reference to omega-3 oil processing ...................................................................................... 178 10.4 Application of nanobiocatalyst in other food processing applications ................. 179 10.5 Conclusions and perspectives ................................................................................ 180 References............................................................................................................... 181 Further Reading ...................................................................................................... 189

CHAPTER 11 Enzyme entrapment approaches and their applications ................. 191 11.1 11.2 11.3 11.4 11.5

11.6

11.7

11.8

Manisha Sharma and Sudhir P. Singh Introduction ............................................................................................................ 191 Immobilization of enzymes.................................................................................... 192 History of enzyme immobilization ........................................................................ 192 Advantages of enzyme immobilization ................................................................. 192 Support systems for enzyme conjugation .............................................................. 193 11.5.1 Organic support materials.......................................................................... 193 11.5.2 Biopolymers ............................................................................................... 194 11.5.3 Hydrogels ................................................................................................... 196 11.5.4 Inorganic support materials ....................................................................... 196 11.5.5 Protein-coated microcrystals technique..................................................... 197 11.5.6 Smart polymers .......................................................................................... 198 11.5.7 Conducting polymers ................................................................................. 198 Types of immobilization techniques...................................................................... 198 11.6.1 Support matrix system ............................................................................... 198 11.6.2 Entrapment system..................................................................................... 199 11.6.3 Cross-linking system.................................................................................. 199 Methods of enzyme immobilization ...................................................................... 199 11.7.1 Physical methods........................................................................................ 200 11.7.2 Chemical methods...................................................................................... 202 Applications of immobilized enzymes .................................................................. 203

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11.8.1 Food industry ............................................................................................. 203 11.8.2 Pharmaceutical industry............................................................................. 203 11.8.3 Brewing industry........................................................................................ 204 11.8.4 Detergent industry...................................................................................... 205 11.8.5 Pulp and paper industry ............................................................................. 205 11.8.6 Textile industry .......................................................................................... 206 11.8.7 Biomedical applications............................................................................. 206 11.9 Conclusions and perspectives ................................................................................ 206 Acknowledgments .................................................................................................. 207 References............................................................................................................... 207

CHAPTER 12 Enzyme immobilization strategies and bioprocessing applications....................................................................................... 217 12.1 12.2

12.3

12.4

12.5

Emmanuel M. Papamichael and Panagiota-Yiolanda Stergiou Introduction ............................................................................................................ 217 Enzymes as industrial biocatalysts ........................................................................ 219 12.2.1 Requirements, advantages, and limitations for the industrial enzymes ...................................................................................................... 219 Enzyme immobilization ......................................................................................... 223 12.3.1 Changes induced by enzyme immobilization: positive and negative....... 224 12.3.2 Strategies, steps, and factors affecting the enzyme immobilization......... 225 12.3.3 Selected methods of enzyme immobilization ........................................... 226 Applications of immobilized enzymes in various bioprocesses ........................... 230 12.4.1 Food, dairies, juices, cosmetics, pharmaceuticals, and others.................. 230 12.4.2 Medical applications of immobilized enzymes and biosensors................ 232 12.4.3 Bioremediation methods and wastewater treatment ................................. 233 12.4.4 Valorization of food processing wastes—objectives and prospects ......... 234 12.4.5 Immobilized enzymes in textile industry .................................................. 234 12.4.6 Immobilized enzymes in the production and process of cosmetics ......... 235 12.4.7 Bioprocessing through nanobiocatalysis ................................................... 235 Conclusions and perspectives ................................................................................ 235 References............................................................................................................... 236

SECTION 4 ENZYMATIC TECHNOLOGIES, CASE STUDIES, AND FUTURE PERSPECTIVES CHAPTER 13 Promising enzymes for biomass processing ................................... 245 Anil Kumar Patel, Pooja Dixit, Ashok Pandey and Reeta Rani Singhania 13.1 Introduction ............................................................................................................ 245 13.1.1 Biomass ...................................................................................................... 246

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13.1.2 Pretreatment ............................................................................................... 247 13.1.3 Biomass-processing enzymes .................................................................... 247 13.1.4 Cellulases for bioconversion...................................................................... 249 13.2 Strain improvement via mutation .......................................................................... 250 13.2.1 Random mutagenesis ................................................................................. 251 13.2.2 Site-directed mutagenesis .......................................................................... 252 13.2.3 Genetic engineering ................................................................................... 252 13.3 Cellulase accessory and auxiliary enzymes........................................................... 256 13.3.1 β-Glucosidase ............................................................................................. 256 13.3.2 Xylanase ..................................................................................................... 257 13.3.3 Laccase ....................................................................................................... 260 13.3.4 Auxiliary enzymes ..................................................................................... 261 13.3.5 Market scenario.......................................................................................... 262 13.4 Conclusion and perspectives.................................................................................. 263 References............................................................................................................... 264

CHAPTER 14 Enzyme systems for high-value biomolecule production ......................................................................................... 273 Rupinder Kaur and Parmjit S. Panesar 14.1 Introduction ............................................................................................................ 273 14.2 Enzymes involved in production of prebiotics...................................................... 274 14.2.1 β-Galactosidase .......................................................................................... 274 14.2.2 β-Glucosidase ............................................................................................. 275 14.2.3 β-Glycosidase ............................................................................................. 275 14.2.4 Fructosyltransferase ................................................................................... 275 14.2.5 β-D-Fructofuranoside.................................................................................. 275 14.2.6 Inulinases.................................................................................................... 276 14.2.7 Glycoside hydrolases ................................................................................. 276 14.3 Production of prebiotics ......................................................................................... 276 14.3.1 Galactooligiosaccharides............................................................................ 276 14.3.2 Lactulose .................................................................................................... 286 14.3.3 Tagatose ..................................................................................................... 287 14.3.4 Lactosucrose............................................................................................... 288 14.3.5 Fructooligosaccharides............................................................................... 289 14.3.6 Isomaltooligosaccharides ........................................................................... 290 14.4 Downstream processing of prebiotics.................................................................... 292 14.5 Global status of prebiotics ..................................................................................... 294 14.6 Conclusions and perspectives ................................................................................ 295 References............................................................................................................... 296 Further reading ....................................................................................................... 306

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CHAPTER 15 Role of enzymatic bioprocesses for the production of functional food and nutraceuticals.................................................................... 309 15.1 15.2

15.3 15.4 15.5

Rounak Chourasia, Loreni C. Phukon, Sudhir P. Singh, Amit Kumar Rai and Dinabandhu Sahoo Introduction ............................................................................................................ 309 Role of enzymes in production of functional foods.............................................. 310 15.2.1 Proteases..................................................................................................... 310 15.2.2 Carbohydrate-modifying enzymes............................................................. 312 15.2.3 L-Asparaginase ........................................................................................... 317 15.2.4 Lipases........................................................................................................ 318 15.2.5 Tannase....................................................................................................... 320 15.2.6 Phytase ....................................................................................................... 320 Production of functional foods using specific enzyme-producing starters........... 321 Genetically modified enzymes in development of functional foods .................... 322 Conclusions and perspectives ................................................................................ 324 Acknowledgments .................................................................................................. 324 References............................................................................................................... 324

CHAPTER 16 Biotransformations with crude enzymes and whole cells .............. 335 16.1 16.2

16.3

16.4

16.5

Haiquan Yang, Fengyu Qin, Zilong Wang, Xianzhong Chen and Guocheng Du Introduction ............................................................................................................ 335 Biotransformations in the pharmaceutical industry............................................... 336 16.2.1 Chiral carboxylic acids .............................................................................. 336 16.2.2 Chiral alcohols ........................................................................................... 337 16.2.3 Chiral amines and amino acids.................................................................. 340 16.2.4 Biosynthesis of natural products................................................................ 343 Biotransformations in food processing .................................................................. 343 16.3.1 Hydrolysis of naturally occurring large molecules into smaller molecules ............................................................................... 345 16.3.2 Biotransformation of chemicals................................................................. 346 Biotransformations in the biofuels industry .......................................................... 348 16.4.1 Classification and biotransformation of biofuels ...................................... 349 16.4.2 Important examples of biofuels ................................................................. 351 Conclusions and perspectives ................................................................................ 353 Acknowledgments .................................................................................................. 353 References............................................................................................................... 353

CHAPTER 17 Enzymes in the third generation biorefinery for macroalgae biomass......................................................................... 363 Abraham Lara, Rosa M. Rodrı´guez-Jasso, Araceli Loredo-Trevin˜o, ´ Cristo´bal N. Aguilar, Anne S. Meyer and Hector A. Ruiz 17.1 Introduction ............................................................................................................ 363

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17.2 Biorefinery and biorefining.................................................................................... 363 17.2.1 First and second generation biorefineries.................................................. 364 17.2.2 Third generation biorefinery...................................................................... 365 17.3 Macroalgae as a source of high-value products .................................................... 365 17.4 Enzyme production ................................................................................................ 366 17.4.1 Solid-state fermentation and semi-solid-state fermentation for enzyme production ............................................................................... 367 17.4.2 Green algae as source for enzyme production .......................................... 369 17.5 Enzymes for biofuels ............................................................................................. 372 17.5.1 Enzymes in the bioethanol production ...................................................... 372 17.5.2 Enzymatic saccharification of green macroalgae...................................... 373 17.5.3 Saccharification by enzymatic complexes, cocktails, and simultaneous fermentation ......................................................................... 382 17.5.4 Biobutanol and acetone, butanol, and ethanol fermentation ............................................................................................... 383 17.5.5 Enzymes in biogas production................................................................... 384 17.6 Enzymatic extraction of high valued products ...................................................... 385 17.6.1 Extraction of hydrocolloids ....................................................................... 385 17.6.2 Extraction of pigments............................................................................... 386 17.7 Conclusions and perspectives ................................................................................ 387 Acknowledgment .................................................................................................... 387 References............................................................................................................... 388

CHAPTER 18 Enzymatic systems for the development of juice clarification strategies........................................................................................... 397 18.1 18.2 18.3

18.4

18.5

Lokesh Kumar Narnoliya, Jyoti Singh Jadaun, Manisha Chownk and Sudhir P. Singh Introduction ............................................................................................................ 397 Why juice clarification? ......................................................................................... 399 Juice clarification strategies................................................................................... 400 18.3.1 Mechanical process .................................................................................... 400 18.3.2 Chemical process ....................................................................................... 401 18.3.3 Enzymatic process ..................................................................................... 401 Enzymes used for juice clarification ..................................................................... 401 18.4.1 Pectic enzymes........................................................................................... 402 18.4.2 Cellulase ..................................................................................................... 403 18.4.3 Laccase ....................................................................................................... 403 18.4.4 Amylase...................................................................................................... 405 18.4.5 Others ......................................................................................................... 407 Conclusions and perspectives ................................................................................ 407 Acknowledgments .................................................................................................. 408 References............................................................................................................... 408

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CHAPTER 19 Biocatalyst systems for xylooligosaccharides production from lignocellulosic biomass and their uses........................................... 413 19.1 19.2 19.3 19.4

19.5 19.6 19.7 19.8

Michele Michelin and Jose´ A. Teixeira Introduction ............................................................................................................ 413 Xylan: sources and structures ................................................................................ 413 Xylooligosaccharides ............................................................................................. 414 Production of xylooligosaccharides ....................................................................... 414 19.4.1 Xylanolytic enzymes.................................................................................. 417 19.4.2 Purification of xylooligosaccharides ......................................................... 417 Biological properties of xylooligosaccharides....................................................... 418 Xylooligosaccharides applications......................................................................... 419 Market and safety aspects of xylooligosaccharides .............................................. 420 Conclusion and perspectives.................................................................................. 420 References............................................................................................................... 421

CHAPTER 20 Biocatalysis for cascade reactions to produce high-value chemicals ....................................................................... 427 Ji Liu, Kaiyuan Tian and Zhi Li 20.1 Introduction ............................................................................................................ 427 20.1.1 What is a cascade reaction?....................................................................... 427 20.1.2 Advantages of cascade biocatalysis........................................................... 427 20.1.3 Production of high-value chemicals via cascade biocatalysis .................. 428 20.2 Designs of biocatalytic cascade reactions ............................................................. 428 20.2.1 Topology of cascade .................................................................................. 428 20.2.2 Pathway design and reaction engineering ................................................. 430 20.3 Biocatalysts............................................................................................................. 431 20.3.1 Enzymes ..................................................................................................... 431 20.3.2 Whole-cell .................................................................................................. 431 20.3.3 Fused enzymes ........................................................................................... 432 20.3.4 Immobilized enzymes ................................................................................ 432 20.4 Examples of cascade biocatalysis for the production of valuable chemicals ................................................................................................................ 432 20.4.1 Cascade biocatalysis from styrenes ........................................................... 433 20.4.2 Cascade biocatalysis from L-phenylalanine............................................... 435 20.4.3 Cascade biocatalysis for the synthesis of amines ..................................... 436 20.4.4 Cascade biocatalysis for the synthesis of lactones.................................... 440 20.5 Conclusions and perspectives ................................................................................ 442 References............................................................................................................... 443 Further reading ....................................................................................................... 447 Index .................................................................................................................................................. 449

List of contributors Reinu E. Abraham Centre for Marine Bioproducts Development, College of Medicine and Public Health, Flinders University, Adelaide, SA, Australia Cristo´bal N. Aguilar Biorefinery Group, Food Research Department, Faculty of Chemistry Sciences, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico Xianzhong Chen Key Laboratory of Carbohydrate Chemistry & Biotechnology, Ministry of Education, School of Biotechnology, Jiangnan University, Wuxi, P.R. China Yukun Chen National Engineering Laboratory for Cereal Fermentation Technology, Jiangnan University, Wuxi, P.R. China; School of Biotechnology, Jiangnan University, Wuxi, P.R. China Joel Ping Syong Choo Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore Rounak Chourasia Institute of Bioresources and Sustainable Development, Sikkim Centre, Tadong, India Manisha Chownk Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India Luciana Porto de Souza Vandenberghe Department of Bioprocess Engineering and Biotechnology, Federal University of Parana´ (UFPR), Curitiba, Brazil Kim Valladares Diestra Department of Bioprocess Engineering and Biotechnology, Federal University of Parana´ (UFPR), Curitiba, Brazil Pooja Dixit DBT-IOC Centre for Advanced Bioenergy Research, Indian Oil Corporation R&D Centre, Faridabad, India Guocheng Du School of Biotechnology, Jiangnan University, The Key Laboratory of Carbohydrate Chemistry and Biotechnology, Wuxi, P.R. China Hafiz M.N. Iqbal Tecnologico de Monterrey, School of Engineering and Sciences, Campus Monterrey, Monterrey, Mexico Jyoti Singh Jadaun Botany Department, Dayanand Girls Postgraduate College, Kanpur, Uttar Pradesh, India

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Susan Grace Karp Department of Bioprocess Engineering and Biotechnology, Federal University of Parana´ (UFPR), Curitiba, Brazil Rupinder Kaur Food Biotechnology Research Laboratory, Department of Food Engineering & Technology, Sant Longowal Institute of Engineering and Technology, Longowal, India Girija Kaushal Center of Innovative and Applied Bioprocessing, Department of Biotechnology, Mohali, India Jitesh Kumar Center of Innovative and Applied Bioprocessing, Department of Biotechnology, Mohali, India Abraham Lara Biorefinery Group, Food Research Department, Faculty of Chemistry Sciences, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico Zhi Li Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore Nelson Libardi, Junior Department of Environmental Engineering, Federal University of Santa Catarina (UFSC), Floriano´polis, Brazil Ji Liu Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore Araceli Loredo-Trevin˜o Biorefinery Group, Food Research Department, Faculty of Chemistry Sciences, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico Anne S. Meyer Protein Chemistry and Enzyme Technology, DTU Bioengineering, Department of Biotechnology and Biomedicine, Technical University of Denmark, Lyngby, Denmark Michele Michelin CEB—Centre of Biological Engineering, University of Minho, Campus Gualtar, Braga, Portugal Lokesh Kumar Narnoliya Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India Goldie Oza Laboratorio Nacional de Micro y Nanofluidica (LABMyN), CIDETEQ, Queretaro, Mexico Maria Giovana Binder Pagnoncelli Department of Biotechnology and Bioprocess Engineering, Federal Technological University of Parana´ (UTFPR), Curitiba, Brazil Ashok Pandey Centre for Innovation and Translational Research, CSIR-Indian Institute of Toxicology Research, Lucknow, India

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Parmjit S. Panesar Food Biotechnology Research Laboratory, Department of Food Engineering & Technology, Sant Longowal Institute of Engineering and Technology, Longowal, India Emmanuel M. Papamichael Department of Chemistry, University of Ioannina, Ioannina, Greece Roberto Parra-Saldivar Tecnologico de Monterrey, School of Engineering and Sciences, Campus Monterrey, Monterrey, Mexico Anil Kumar Patel Department of Chemical and Biological Engineering, Korea University, Seoul, Republic of Korea Loreni C. Phukon Institute of Bioresources and Sustainable Development, Sikkim Centre, Tadong, India Munish Puri Centre for Marine Bioproducts Development, College of Medicine and Public Health, Flinders University, Adelaide, SA, Australia Fengyu Qin Department of Chemical and Biomolecular Engineering, National University of Singapore, Singapore, Singapore Amit Kumar Rai Institute of Bioresources and Sustainable Development, Sikkim Centre, Tadong, India; Institute of Bioresources and Sustainable Development, Imphal, India Ricardo A. Ramirez-Mendoza Tecnologico de Monterrey, School of Engineering and Sciences, Campus Monterrey, Monterrey, Mexico Rosa M. Rodrı´guez-Jasso Biorefinery Group, Food Research Department, Faculty of Chemistry Sciences, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico ´ Hector A. Ruiz Biorefinery Group, Food Research Department, Faculty of Chemistry Sciences, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico Dinabandhu Sahoo Institute of Bioresources and Sustainable Development, Sikkim Centre, Tadong, India; Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India Balaji Sundara Sekar Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore Ashutosh Sharma Tecnologico de Monterrey, School of Engineering and Sciences, Campus Queretaro, Queretaro, Mexico

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Himanshu Sharma Department of Botany, Panjab University, Chandigarh, India; I.K. Gujral Punjab Technical University, Jalandhar, India Manisha Sharma Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India Sudhir P. Singh Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India Reeta Rani Singhania Center for Energy and Environmental Sustainability-India, Lucknow, India Carlos Ricardo Soccol Department of Bioprocess Engineering and Biotechnology, Federal University of Parana´ (UFPR), Curitiba, Brazil Archana Srivastava Botany Department, Dayanand Girls Postgraduate College, Kanpur, Uttar Pradesh, India Panagiota-Yiolanda Stergiou Department of Chemistry, University of Ioannina, Ioannina, Greece Jose´ A. Teixeira CEB—Centre of Biological Engineering, University of Minho, Campus Gualtar, Braga, Portugal Kaiyuan Tian Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore Santosh Kumar Upadhyay Department of Botany, Panjab University, Chandigarh, India Madan L. Verma Department of Biotechnology, School of Basic Sciences, Indian Institute of Information Technology Una, Una, Himachal Pradesh, India ´ Jessica Aparecida Viesser Department of Bioprocess Engineering and Biotechnology, Federal University of Parana´ (UFPR), Curitiba, Brazil Matheus von Linsingen Tavares Department of Bioprocess Engineering and Biotechnology, Federal University of Parana´ (UFPR), Curitiba, Brazil Zilong Wang Department of Chemical and Biomolecular Engineering, National University of Singapore, Singapore, Singapore Zhe Xu Key Laboratory of Bioorganic Synthesis of Zhejiang Province, College of Biotechnology and Bioengineering, Zhejiang University of Technology, Hangzhou, P.R. China

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Ya-Ping Xue Key Laboratory of Bioorganic Synthesis of Zhejiang Province, College of Biotechnology and Bioengineering, Zhejiang University of Technology, Hangzhou, P.R. China Haiquan Yang Key Laboratory of Carbohydrate Chemistry & Biotechnology, Ministry of Education, School of Biotechnology, Jiangnan University, Wuxi, P.R. China Ricardo Zavala-Yoe Tecnologico de Monterrey, Campus Mexico City, Mexico City, Mexico Guoqiang Zhang National Engineering Laboratory for Cereal Fermentation Technology, Jiangnan University, Wuxi, P.R. China; School of Biotechnology, Jiangnan University, Wuxi, P.R. China Yu-Guo Zheng Key Laboratory of Bioorganic Synthesis of Zhejiang Province, College of Biotechnology and Bioengineering, Zhejiang University of Technology, Hangzhou, P.R. China Shu-Ping Zou Key Laboratory of Bioorganic Synthesis of Zhejiang Province, College of Biotechnology and Bioengineering, Zhejiang University of Technology, Hangzhou, P.R. China

Preface The book titled Advances in Enzyme Catalysis and Technologies is a part of the comprehensive series on Biomass, Biofuels, Biochemicals (Editor-in-Chief: Ashok Pandey). Enzymes are protein macromolecules that catalyze biochemical reactions. Genetic engineering, following the approaches of DNA mutation, directed evolution, and rational designing, modifies the structural features of the native enzyme, conferring it desirable catalytic dispositions. Native and engineered biocatalysts are of paramount importance in developing processing technologies in food, feed, and biopharmaceutical sectors. Advancements in omics, mutation, and bioinformatics research have paved the way for many novel approaches for the discovery and rational designing of enzymes with improved and desirable catalytic dispositions of industrial significance. This book intends to provide the basic structural and functional description and classification of enzymes. The scientific information related to the recombinant enzyme modifications, the discovery of novel enzymes, and the development of synthetic enzymes is presented. The translational aspects of enzyme catalysis and bioprocess technologies have been illustrated by emphasizing the current requirements and future perspectives of industrial biotechnology. The book gives comprehensive information about the protein macromolecules that catalyze biochemical reactions, enzyme kinetics, enzyme production, enzyme engineering, and enzyme-based processing technologies. The basic structural and functional description of the enzymes has been outlined. The fundamental scientific information related to the recombinant enzyme modifications, discovery of novel enzymes, and development of synthetic enzymes has been described. The strategic approaches for the discovery and engineering of novel enzymes have been discussed, along with future perspectives. In brief, this book has covered comprehensive outlook of enzymology and the contemporary translational aspects of the biological catalysts, executing the biological reactions for in vivo or in vitro biosynthesis of high-value molecules, and enzymatic bioprocessing of the plant biomass. The content of the book is organized into four sections, covering different aspects of advancement in enzyme catalysis and technologies. The scientific information is contributed by the global experts of the research field in the form of chapters in the book. The first section deals with basic aspects of enzymes. The chapters of this section narrate the basic information on enzyme-structure dynamics and enzyme catalysis, the nomenclature and classification of the enzymes on the basis of the catalytic behavior. Further, the currently available enzyme production technologies and the state-of-the-art of biocatalysis engineering have been emphasized. Section 2 presents compiled information around the development of efficient enzymes with desirable characteristics in five chapters. These chapters describe the enzyme engineering strategies for the traits of industrial significance such as improvement in the thermal and pH tolerance and enhancement in the catalytic efficiency. These chapters also deal with developing multifunctional biocatalysts and enantioselective biotransformations of enzymes. Section 3 contains the chapters on the approaches for enzyme immobilization and entrapment technologies. Section four cites specific enzyme applications and case studies. The chapters depict the recent advancements and perspectives in enzymatic processing of biomass for achieving value-added bioproducts, functional food development, and different high-value biomolecules of health significance. We highly appreciate the hard work of authors in compiling and presenting the comprehensive scientific and relevant state-of-the-art information in different aspects of enzymology and industrial

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biotechnology. We firmly believe that the enriched scientific materials presented in the book will be of much use to different sections of academic and scientific communities. We gratefully acknowledge the reviewers’ efforts in the critical review of the chapters, which led to the scientific enrichment of this volume. We thank Dr. Kostas Marinakis, Senior Book Acquisition Editor, Andrea Dulberger, Editorial Project Manager, and the entire Elsevier production team for their consistent hard work in the publication of this book. Editors Sudhir P. Singh, Ashok Pandey, Reeta Rani Singhania, Christian Larroche and Zhi Li

CHAPTER

AN INTRODUCTION TO ENZYME STRUCTURE DYNAMICS AND ENZYME CATALYSIS

1

Jitesh Kumar1, Ashok Pandey2 and Sudhir P. Singh3 1

Center of Innovative and Applied Bioprocessing, Department of Biotechnology, Mohali, India Centre for Innovation and Translational Research, CSIR-Indian Institute of Toxicology Research, Lucknow, India 3 Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India

2

1.1 INTRODUCTION Enzymes are the most critical biocatalyst for the catalysis of substrate into the product for all the biological systems to perform metabolism. Not only this, the application of enzymes has been extended to various industries such as textile, feed, and food [1]. The building block of all the enzymes is amino acid (except ribozymes), but not all the proteins have evolved to perform catalysis. The most significant difference between enzyme and protein is the presence of a active sites. The active sites are present in the cleft of the tertiary structure of the enzyme into which substrate enters, and the product diffuses out. The catalytic mechanism of the enzyme has been supported by theories such as lock and key, induced-fit, and transition-state [2
4]. Unlike any chemical catalyst, enzymes are very efficient biocatalyst that speeds up the reaction by lowering the energy of activation. They are also required in low concentration and perform biological reactions without being consumed and undergoing significant changes [5]. Different enzymes have a different mechanism of catalysis, but some of the common catalytic mechanisms include acid
base, electrostatic, and covalent catalysis. Enzymes are dynamic entities that undergo internal changes from local bond vibration to conformational motions. These properties are linked to various biological functions [6]. The role of the structural dynamic of enzymes in their catalytic activity is derived from the molecular dynamics (MD) simulations, nuclear magnetic resonance (NMR) spectroscopy, and mass spectrometry
based methods [7]. Most enzymes are proteins, but ribozymes are the unique example of nonprotein enzymes, which are capable of cleaving phosphodiester link either of a complementary exogenous RNA fragment or of itself. According to RNA world hypothesis, ribozymes would have played an important role in primitive life [8]. This chapter introduces the structure dynamics and catalysis of the enzyme.

1.2 WHY DO SOME PROTEINS WORK AS ENZYME? Proteins are versatile biological macromolecule that performs functions as a structural component, enzymes, hormones, and antibodies in almost all the living beings. However, enzymes are unique Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00001-6 © 2020 Elsevier B.V. All rights reserved.

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CHAPTER 1 AN INTRODUCTION TO ENZYME STRUCTURE DYNAMICS

in their function to catalyze the reactions containing a specific molecule (referred to as a substrate) into the product without being consumed or massive alteration in the structure. Both proteins and enzymes are constituted from the chain of amino acid. Thus, except ribozymes, all enzymes are proteins, whereas all the proteins are not enzymes. Enzymes differ from protein in having active sites and may also require non-protein components (cofactors) for the catalysis [9].

1.2.1 ROLE OF THE ACTIVE SITE IN ENZYME FUNCTIONING Active site is an essential part of the enzyme that is located within the deep pocket of its surface. They play a significant role in the interaction with the substrate and contain key amino acid residues for the catalysis. The catalytic residue acts by characteristic mode of actions such as a nucleophilic attack, acid
base catalysis, and activation of the water molecule [10]. Largely, the majority of the catalytic residues are charged (H, R, K, E, and D) and polar (Q, T, S, N, C, Y, and W), whereas a few are of hydrophobic nature [10]. The charged and/or polar residues participate in charge stabilization and exchange of electrons and protons. The catalytic residues are conserved and have structural rigidity over other residues in the enzyme. Such features make an enzyme, specific toward the substrate.

1.2.2 COFACTOR, A NECESSITY OF ENZYME Another vital aspect of the enzyme, which distinguishes it from protein, is the requirement of a non-protein substance called as cofactors, the essential component for execution of catalytic reaction. Cofactors can further be categorized into prosthetic groups and coenzymes, depending on their type of association with the enzymes [5]. Prosthetic groups are a small molecule, which remains bound to the enzyme; for example, heme is bound to myoglobin and hemoglobin protein as an essential component for oxygen binding. In some cases, metal ions (such as zinc or iron) remain bound to the enzymes, playing a critical role in the catalysis. Coenzymes carry chemical groups; for example, nicotinamide adenine dinucleotide (NAD1) functions as an electron carrier in oxidation
reduction reactions [9]. Interestingly, the presence of cofactor can induce conformational changes in the enzyme that may increase the fitting and interaction of the substrate with the active site [11].

1.3 WHAT IS ENZYME CATALYSIS? 1.3.1 BASICS OF ENZYME CATALYSIS Enzyme catalysis plays a vital role in the metabolism of all the living organisms. Enzymatic reactions are like chemical reactions, which result in the product, and the free energy change (ΔG) during this process is negative. However, the rate of reaction depends on another factor, known as activation energy (EA). If the ΔG is favorable, but still the need of high EA leads to slower progression of the reaction. The biological reaction is required to be catalyzed in the fractions of a second [12]. The slower biochemical changes are made faster (1010
1020) by using the enzymatic actions, without which a reaction may take millions of years to complete [13]. The rate of free

1.3 WHAT IS ENZYME CATALYSIS?

5

energy change, during the progression of enzymatic catalysis, is represented by a bell-shaped curve. The peak in the graph represents transition state (TS), which is an intermediate of the substrate (S) and the product (P). In a reaction, the difference between the free energy of the substrate and the transition state determines the rate of reaction and is referred to as EA. All the catalysts, chemical or biological, are supposed to minimize the EA of a reaction. However, enzymatic reactions are relatively complex as they require substrate binding and forms more than one stable intermediate.

1.3.2 HISTORICAL OVERVIEW OF ENZYME CATALYSIS THEORY The first theory of enzyme catalysis was the “lock and key” model proposed by Emil Fischer, which states that the substrate fits in the enzymes like a key fits into the lock [2]. This was later modified by Koshland’s “induced-fit” theory, proposing that substrate does not fit to the enzyme perfectly, but performs a subtle conformational change in the enzyme’s active site. Later on, Koshland’s induced-fit theory was extended by Eyring’s in 1935 as “transition-state” theory [3], and further by Pauling’s “transition-state stabilization” theory in 1946. Transition-state theory narrates that an enzyme binds preferentially to the transition-state intermediate rather than a substrate or product. This reduces the free energy of the transition state and thereby reduces the energy of activation, achieving an accelerated reaction. Adding to this, the motion of the catalytic residues has been noted to be lesser than the binding residues. However, at the same time, side chains display similar motion [14].

1.3.3 MECHANISTIC VIEW OF ENZYME CATALYSIS The theories explained above are a generalized overview of the mechanism of the enzymes. The actual chemistry of the catalysis depends on the type of enzyme, and inexplicable change occurred during the evolution of enzymes. This section will briefly explain some of the selected mechanisms explored in the enzyme catalysis. Acid
base catalysis is the most homogenous and commonest type of catalytic mechanism. In acid
base catalysis, the charge on the instable transition state is stabilized by donating or accepting a proton from the substrate to the chemical group on the enzyme molecule [15]. The acid
base reaction includes hydrolysis, alcohol lysis, esterification, and condensation. The typical example of enzymes that act through acid
base catalysis is ribonuclease [15]. The hydrolysis of RNA is a two-step reaction in which a cyclic intermediate is formed. The unfavorable positive and negative charges are accumulated on 20 OH and the phosphate oxygen in the first and second steps, which is neutralized during the enzyme catalysis by two histidine residues, capable of donating and accepting protons. The acid
base catalysis depends upon the strength of a catalytic group and its ionization state. So, an enzyme with a strong basic or acidic group is a promising catalyst, because it has a tendency to donate or accept a proton from other groups firmly. In electrostatic catalysis, enzymes such as the ribonuclease in the above example neutralize the charges developed during transition state. A few other enzymes use the opposite charge on the group to balance the charged transition states. These complementary charges, which enhances the catalysis, are available from different groups such as amide group, arginine, and Zn21 in serine protease, Staphylococcal nuclease, and carboxypeptidase [16
18].

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CHAPTER 1 AN INTRODUCTION TO ENZYME STRUCTURE DYNAMICS

Covalent catalysis is one of the strategies adopted by some enzymes (serine proteases, cysteine proteases, protein kinases and phosphatases, and pyridoxal phosphate-using enzymes) that involve the formation of a covalent bond between the substrate and critical residue of the active site or with the cofactor. The process is activated by the chemical modifications of the substrate to speed up the reaction. An example of covalent catalysis is the formation of a Schiff base (electrophilic group) through the condensation of an amine with a carbonyl group. The Schiff base prepares the carbonyl carbon for the nucleophilic attack by another group [19,20]. Covalent catalysis is also carried out by nucleophile that forms a covalently attached intermediate such as the tetrahedral intermediate in case of serine proteases. This helps in easier catalysis of the substrate by the production of reaction intermediates and their subsequent breakdown.

1.4 STRUCTURAL DYNAMICS OF ENZYMES Enzyme catalysis is a dynamic process in which the atomic position of the enzyme molecules changes during enzymatic catalysis in a time-dependent manner. Specifically, protein dynamics have a wide timescale range from picoseconds and nanoseconds, for local flexibility (methyl rotations, side-chain rotations, and loop motions), to microseconds and milliseconds, for collective motions (larger domain motions) [21]. The enzymes undergo conformational changes during the catalytic process for the selective substrate binding, catalysis, and interaction with regulatory molecules. This is essential for enzyme function and activity [22,23]. A variety of techniques such as X-ray crystallography and small-angle scattering, NMR studies, hydrogen
deuterium exchange, neutron scattering, biochemical, and mutational analysis have provided deep insights into the protein dynamics at individual timescale [24]. However, to understand the protein dynamics, information is required at a broad timescale. Additively, the coupled knowledge of theory and computational modeling plays a central role in the understanding of dynamics and catalysis at multiple timescales. In recent years, a myriad of research has proved the interconnection between protein dynamics and enzyme catalysis. Due to this perception, an enzyme Cyclophilin A (CypA) that catalyzes the reversible cis
trans isomerization of prolyl peptide bonds has been studied in detail, biophysically and theoretically [25]. NMR dispersion studies of CypA confirm a relationship between global conformational changes and isomerization of the substrate. Additionally, a characteristic collective motion was detected during the catalysis with a frequency similar to turnover numbers (kcat) [26,27]. The isomerization reaction is also linked to fluctuations in certain regions of the enzyme backbone and the network of protein vibrations. This variation was identified using a combination of MD and umbrella sampling [28,29]. Interestingly, single-molecule studies on the lipase of Candida antarctica have revealed the multiple active conformations that convert slowly at the same timescale of activity [30]. Solvent matters a lot when one talks about the protein dynamics. Protein motion is considered to be Brownian; therefore, the diffusive motion depends upon the friction resulting from the surrounding solvent environment. Ideally, a mobile solvent should support faster motion and flexibility. In contrast to this, a cage of more static/viscous solvent should promote stability in the protein by restricting its motion [21]. This is supported by groundbreaking studies performed in various aqueous solutions of glycerol, ethanol, and ethylene glycol [31]. A relationship between protein

1.5 RIBOZYMES AS A NONPROTEIN CATALYST

7

and solvent dynamics has been inferred in the case of gas binding to heme protein in aqueous solution, which is inversely proportional to viscosity [30]. The studies based on a combination of terahertz spectroscopy, X-ray absorption, and MD illustrate the existence of fast-to-slow waterenzyme motion, correlated with the rearrangements of the active site of a zinc-dependent human metalloproteinase [32]. The importance of small-amplitude rapid vibrations or large-amplitude slow conformational motions is decided by the nature of enzymatic reactions. Substantial conformational changes (microsecond to second timescales) are typically linked with substrate/cofactor binding and product release or the achievement of a specific chemical reaction [25]. Many studies pertaining to protein dynamics have been explained above, but the role of internal motion in accelerating the catalysis rate of the enzyme is challenging and yet to be determined appropriately.

1.5 RIBOZYMES AS A NONPROTEIN CATALYST We have mentioned in the above section that ribozyme is an exception of the non-protein enzyme, that cleaves the ribonucleic acid. Although the enzyme dominates in the living world, the discovery of catalytic RNAs (ribozymes), endorsed the hypothesis that nucleic acid is a vital origin of the biocatalyst, as RNA is a central molecule in the step of protein synthesis in a cellular system. According to the RNA world hypothesis, RNA would have played a dual role of genetic material and biocatalyst in the primitive living cells [33]. As the metabolic functions evolved, RNA-based catalyst might have emerged with a protein-based catalyst. Ribozymes and enzymes of the protein origin have a very different composition. Still, the final goal is catalysis. Like the protein-based enzymes, ribozymes have adopted the primary mechanism of catalysis, involving stabilization of the transition state relative to the ground state at the cost of entropy loss [34,35]. The naturally available ribozymes are of five classes that perform cleavage of phosphodiester bonds in the substrate, RNA. These five classes can be grouped on the basis of their functions— cleavage, splicing, and peptide bond formation. It is interesting that self-cleaving ribozymes have a broad catalytic potential like proteinaceous enzymes [33]. The hammerhead hepatitis δ virus (HDV) and hairpin ribozymes are of B40
120 nucleotide RNAs. These ribozymes perform sitespecific cleavage to yield 20 ,30 -cyclic phosphates and 50 -hydroxyl termini. They are very specific for their target RNAs and use base pairing with the target RNA to bring closer to the active site in comparison to their analogous protein nucleases. The natural hammerhead ribozymes contain additional non-conserved sequences that help in the stabilization at optimum magnesium ion concentration [36,37]. Other ribozymes such as HDV and the hairpin ribozyme catalyze the same reaction by forming an enclosed cleft in which cleavage takes place [38,39]. HDV follows acid
base catalysis, in which ribonucleotides directly contribute to catalysis by donating or accepting proton as discussed above for protein enzymes. Another class of ribozymes performs a two-step reaction, that is, splicing and ligation. The splicing removes the sequence between two exons, that is, introns, and at the same time executes ligation of two exons to produce a mature mRNA transcript. The spliceosome is a dynamic RNA
protein complex that cleaves intervening intronic sequences. Two different classes (group I and II) of self-cleavable autocatalytic spliceosomes are present in protozoan nuclei, fungal mitochondria, algal chloroplasts, bacteria, and bacteriophages. Group I spliceosomes comprise nine

8

CHAPTER 1 AN INTRODUCTION TO ENZYME STRUCTURE DYNAMICS

base-paired elements and perform splicing in two steps of trans-esterification reactions, activated by external guanosine nucleoside or nucleotide [33]. The 30 end of the attached guanosine substrate attacks the 50 splice site phosphate and attaches to the 50 end of the intron. In a further step, 30 OH of the 50 exon targets the phosphate at the 30 -splice junction to join the exons and remove the intron. Group II spliceosomes have a large secondary structure and a different splicing mechanism. The 20 OH group of internal adenosine within the intron serves as the nucleophile, which cleaves the 50 -splice site phosphodiester bond and forms a 20 -50 linkage with the ends of the intron. Further, the 30 OH of the 50 exon attacks the 30 -splice junction phosphate, ligating the exons and discarding the intron.

1.6 CONCLUSIONS AND PERSPECTIVE The difference between protein and enzymes can be very clearly distinguishable, mainly at the functional level. The specificity of the substrate and mechanism used by enzyme makes them able to process multistep reactions without the formation of any by-product that generally occurs in a chemical reaction. As the catalytic residues are found to be conserved in enzymes, they form the basis of novel enzyme identification. Acid
base catalyst is the standard mechanism of the enzyme, which is because every enzymatic reaction involves transfer/acceptance of proton. The role of enzyme dynamics play an essential role in the catalysis of the reaction and is needed in order to tune up the catalysis processes in a different solvent. Ribozymes are the unique class of nonprotein enzyme that play a dual role of genetic material and the catalyst of specific biochemical reactions in the cell. Their role in regulating gene expression in bacteria and eukaryotes highlights their importance in modern biology.

REFERENCES [1] O. Kirk, T.V. Borchert, C.C. Fuglsang, Industrial enzyme applications, Curr. Opin. Biotechnol. 13 (4) (2002) 345
351. [2] S.J. Benkovic, S. Hammes-Schiffer, A perspective on enzyme catalysis, Science 301 (5637) (2003) 1196
1202. [3] K.J. Laidler, M.C. King, Development of transition-state theory, J. Phys. Chem. 87 (15) (1983) 2657
2664. [4] D.E. Koshland Jr., The key
lock theory and the induced fit theory, Angew. Chem. Int. Ed. Engl. 33 (23
24) (1995) 2375
2378. [5] P.K. Robinson, Enzymes: principles and biotechnological applications, Essays Biochem. 59 (2015) 1
41. [6] D. Petrovi´c, et al., Conformational dynamics and enzyme evolution, J. R. Soc. Interface 15 (144) (2018) 20180330. [7] U. Uzuner, et al., Enzyme structure dynamics of xylanase I from Trichoderma longibrachiatum, BMC Bioinform. 11 (2010) S12. [8] T.J. Wilson, Y. Liu, D.M. Lilley, Ribozymes and the mechanisms that underlie RNA catalysis, Front. Chem. Sci. Eng. 10 (2) (2016) 178
185. [9] G.M. Cooper, The Central Role of Enzymes as Biological Catalysts, Sinauer Associates, 2000.

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[10] G.J. Bartlett, et al., Analysis of catalytic residues in enzyme active sites, J. Mol. Biol. 324 (1) (2002) 105
121. [11] D. Biswas, et al., Co-factor binding confers substrate specificity to xylose reductase from Debaryomyces hansenii, PLoS One 7 (9) (2012) e45525. [12] A. Radzicka, R. Wolfenden, A proficient enzyme, Science 267 (5194) (1995) 90
93. [13] R.L. Stein, A process theory of enzyme catalytic power—the interplay of science and metaphysics, Found. Chem. 8 (1) (2006) 3
29. [14] A. Gutteridge, J. Thornton, Conformational changes observed in enzyme crystal structures upon substrate binding, J. Mol. Biol. 346 (1) (2005) 21
28. [15] D.G. Herries, A. Mathias, B. Rabin, The active site and mechanism of action of bovine pancreatic ribonuclease. 3. The pH-dependence of the kinetic parameters for the hydrolysis of cytidine 20 , 30 -phosphate, Biochem. J. 85 (1) (1962) 127. [16] J.K. Judice, et al., Probing the mechanism of staphylococcal nuclease with unnatural amino acids: kinetic and structural studies, Science 261 (5128) (1993) 1578
1581. [17] R.C. Wilmouth, et al., X-ray snapshots of serine protease catalysis reveal a tetrahedral intermediate, Nat. Struct. Mol. Biol. 8 (8) (2001) 689. [18] N.M. Hooper, Families of zinc metalloproteases, FEBS Lett. 354 (1) (1994) 1
6. [19] A.C. Eliot, J.F. Kirsch, Pyridoxal phosphate enzymes: mechanistic, structural, and evolutionary considerations, Annu. Rev. Biochem. 73 (1) (2004) 383
415. [20] M. Pohl, G.A. Sprenger, M. Mu¨ller, A new perspective on thiamine catalysis, Curr. Opin. Biotechnol. 15 (4) (2004) 335
342. [21] J.N. Dahanayake, K.R. Mitchell-Koch, How does solvation layer mobility affect protein structural dynamics? Front. Mol. Biosci. 5 (2018) 65. [22] K. Teilum, J.G. Olsen, B.B. Kragelund, Functional aspects of protein flexibility, Cell. Mol. Life Sci. 66 (14) (2009) 2231. [23] K. Henzler-Wildman, D. Kern, Dynamic personalities of proteins, Nature 450 (7172) (2007) 964. [24] P.K. Agarwal, Enzymes: an integrated view of structure, dynamics and function, Microb. Cell Fact. 5 (1) (2006) 2. [25] S.G. Estacio, In silico strategies toward enzyme function and dynamics, Adv. Protein Chem. Struct. Biol. 87 (2012) 249
292. [26] E.Z. Eisenmesser, et al., Enzyme dynamics during catalysis, Science 295 (5559) (2002) 1520
1523. [27] E.Z. Eisenmesser, et al., Intrinsic dynamics of an enzyme underlies catalysis, Nature 438 (7064) (2005) 117. [28] G.M. Torrie, J.P. Valleau, Nonphysical sampling distributions in Monte Carlo free-energy estimation: umbrella sampling, J. Comput. Phys. 23 (2) (1977) 187
199. [29] J. Ka¨stner, Umbrella sampling, Wiley Interdiscip. Rev. Comput. Mol. Sci. 1 (6) (2011) 932
942. [30] O. Flomenbom, et al., Stretched exponential decay and correlations in the catalytic activity of fluctuating single lipase molecules, Proc. Natl. Acad. Sci. USA 102 (7) (2005) 2368
2372. [31] D. Beece, et al., Solvent viscosity and protein dynamics, Biochemistry 19 (23) (1980) 5147
5157. [32] M. Grossman, et al., Correlated structural kinetics and retarded solvent dynamics at the metalloprotease active site, Nat. Struct. Mol. Biol. 18 (10) (2011) 1102. [33] J.A. Doudna, J.R. Lorsch, Ribozyme catalysis: not different, just worse, Nat. Struct. Mol. Biol. 12 (5) (2005) 395. [34] G.J. Narlikar, D. Herschlag, Mechanistic aspects of enzymatic catalysis: lessons from comparison of RNA and protein enzymes, Annu. Rev. Biochem. 66 (1) (1997) 19
59. [35] A.A. Armstrong, L.M. Amzel, Role of entropy in increased rates of intramolecular reactions, J. Am. Chem. Soc. 125 (47) (2003) 14596
14602.

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[36] A. Khvorova, et al., Sequence elements outside the hammerhead ribozyme catalytic core enable intracellular activity, Nat. Struct. Mol. Biol. 10 (9) (2003) 708. [37] J.C. Penedo, et al., Folding of the natural hammerhead ribozyme is enhanced by interaction of auxiliary elements, RNA 10 (5) (2004) 880
888. [38] A.R. Ferr´e-D’Amar´e, K. Zhou, J.A. Doudna, Crystal structure of a hepatitis delta virus ribozyme, Nature 395 (6702) (1998) 567. [39] P.B. Rupert, A.R. Ferre-D’Amare, Crystal structure of a hairpin ribozyme
inhibitor complex with implications for catalysis, Nature 410 (6830) (2001) 780.

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CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

2

Luciana Porto de Souza Vandenberghe1, Susan Grace Karp1, Maria Giovana Binder Pagnoncelli2, Matheus von Linsingen Tavares1, Nelson Libardi Junior3, Kim Valladares Diestra1, ´ Jessica Aparecida Viesser1 and Carlos Ricardo Soccol1 1

Department of Bioprocess Engineering and Biotechnology, Federal University of Parana´ (UFPR), Curitiba, Brazil Department of Biotechnology and Bioprocess Engineering, Federal Technological University of Parana´ (UTFPR), Curitiba, Brazil 3Department of Environmental Engineering, Federal University of Santa Catarina (UFSC), Floriano´polis, Brazil

2

2.1 INTRODUCTION Enzymes represent the largest group of proteins and play an essential role in different processes including metabolism, gene expression, cell division, and important reactions of the immune system. They are employed in the biotechnology industry and in medicine for diagnostics. A 30%
40% of existent genes encode enzymes [1]. They accelerate chemical reactions in elevated magnitude, promoting coordinated metabolic pathways within cells, and are essential in the fight against pathogenic organisms and in other processes [2]. Enzymes are very specific and their activity and functions are dependent on many characteristics, including their sequence, threedimensional (3D) structure, stability, and interactions with other molecules [3]. The diversity of actions and applications of enzymes are due to different substrate and reaction specificities.

2.1.1 ENZYME NOMENCLATURE AND CLASSIFICATION The first enzyme nomenclature scheme was presented at the International Congress of Biochemistry, Brussels, where the Enzyme Commission (EC) was defined. Then, in 1961, the first version was published. In 1992, the International Union of Biochemistry and Molecular Biology (IUBMB) published the sixth edition including 3196 different enzymes, with supplements published electronically. The classification of enzymes follows the EC number. Enzymes are classified according to the catalyzed reactions [4,5] where the name corresponds to a single enzyme protein and may also be linked to a group of proteins with the same catalytic property. Completely different protein folds are known for catalyzing an identical reaction and thus receiving an identical EC number. These enzymes are called nonhomologous isofunctional enzymes (NISE) [6]. Six enzyme classes or six EC levels were defined, including the oxidoreductases, transferases, hydrolases, lyases, isomerases, and ligases. In August 2018, the EC included a seventh enzyme Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00002-8 © 2020 Elsevier B.V. All rights reserved.

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CHAPTER 2 CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

category (EC 7), which grouped the translocases [7]. The classification of enzymes’ sequences and structures was supported by evolutionary and biophysical models, which allowed the knowledge of sequence and structural similarity [1]. The EC number [2] is composed of four digits: • • • •

First number—shows the class of the enzyme; Second number—gives the subclass; Third number—indicates the sub-subclass; and Fourth number—is the serial number of the enzyme in its sub-subclass.

2.1.2 ENZYME DATABASES Enzyme information is grouped in database sources that include the classification, nomenclature, and description of each characterized enzyme with its corresponding EC number or catalyzed reactions. The BRaunschweig ENzyme DAtabase (BRENDA) is the most used biological database worldwide [2]. In 1987, the collection of enzyme-related data from the scientific literature was started by the German National Research Centre for Biotechnology in Braunschweig. From 1996 to 2007, it was continued at the University of Cologne. Then, it returned to the Department of Bioinformatics and Systems Biology of the Technische Universita¨t Braunschweig. It is now established at the Braunschweig Integrated Centre of Systems Biology. The initial data consisted of a compilation of textual and numeric forms with one dataset for each enzyme class, which were published from 1990 to 2009 in a series of 49 books covering B4900 enzyme classes [2]. Concomitantly, an internet presentation of the database system was created [8,9]. Since then, more than 370,000 enzymes from 6300 enzyme classes were reported. For that, enzyme-related information combined functional data with genomic sequences, enzyme structures, and computed data [1,10]. Another important database is the Protein Data Bank (PDB), which is a repository of 3D macromolecular enzyme structural data determined experimentally by X-ray crystallography and nuclear magnetic resonance (NMR). It consists of information about the 3D shapes of proteins, nucleic acids, and complex assemblies for students and researchers [11]. Some of these databases are presented in Table 2.1.

2.1.3 CATALYTIC PROPERTIES Most enzymes are composed of proteins in a single chain or different chains. Other nonprotein components are also present, such as metal ions or some organic molecules: “the cofactors.” Cofactors consist of vitamins and other molecules. The enzymatic reaction occurs under certain energy conditions, corresponding to a short high-energy transition state, where the substrate is changed to the product. The enzyme catalytic reaction is described below. First, there is the formation of an enzyme complex, with its further dissociation to a product. E 1 S.ES.E 1 P

Enzymes are biocatalyzers that significantly increase the rate of a reaction. The crucial factor for a reaction to happen is the reduction of energy barrier separating the reactants from the

2.1 INTRODUCTION

13

Table 2.1 Enzyme databases for researchers and students’ information. Database

Information

Development/origin

Access

BRENDA

Collection of enzyme functional data

CAZy database

Describes the families of structurally related catalytic and carbohydratebinding modules of enzymes that degrade, modify, or create glycosidic bonds Contains the known enzyme structures that have been deposited in the Protein Data Bank (PDB)

Institute of Biochemistry and Bioinformatics, Technical University, Braunschweig, Germany Architecture et Fonction des ´ Macromolecules Biologiques (AFMB), CNRS, Aix-Marseille Universit´e, France

https://www. brenda-enzymes. org http://www.cazy. org/

Wellcome Trust, United Kingdom

https://www.ebi. ac.uk/thorntonsrv/databases/ enzymes/ https://enzyme. expasy.org/ http://www. enzymedatabase.org/

EC2PDB

EXPAZYENZYME EXPLOREENZ

Repository of information relative to the nomenclature of enzymes Access to the data of the IUBMB Enzyme Nomenclature List

Fun Tree

Provides a range of data resources to detect the evolution of enzyme function within distant structurally related clusters within domain superfamilies Database of enzyme mechanisms

Mechanism and catalytic site atlas PDB

Repository of 3D macromolecular structural data primarily determined experimentally by X-ray crystallography and NMR

Swiss Institute of Bioinformatics Nomenclature Committee of the International Union of Biochemistry and Molecular Biology (NCIUBMB), by Andrew McDonald, School of Biochemistry & Immunology, Trinity College, Dublin 2, Ireland European Molecular Biology Laboratory (EMBL), Cambridge, United Kingdom

Thorton Group, European Bioinformatics Institute, and Mitchell Group, University of St Andrews, United Kingdom Natural Science Foundation, National Institutes of Health

https://www.ebi. ac.uk/thorntonsrv/databases/ FunTree https://www.ebi. ac.uk/thorntonsrv/m-csa/ http://www.rcsb. org/pdb/

products. The activation energy (Ea) is reduced, which favors reactant molecules to overcome this barrier and then the product is formed. Exactly as other catalysts, the enzyme is not consumed or changed by the reaction, which gives the possibility, for example, to recycle or reuse this biocatalyzer for many cycles of catalysis. Enzymatic catalysis occurs through different mechanisms depending on the enzyme. Four catalytic models have been described for enzymes: lock-and-key model, induced-fit model, selected-fit model, and finally, keyhole-lock-key model. In 1984, the first model of enzyme catalysis was presented by Emil Fischer, which was called lock-and-key model [12], and was then expanded to conformational selection model including the induced-fit and the selected-fit models. These models

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CHAPTER 2 CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

consider the flexibility of the ligand and the enzyme. The induced-fit model considers conformational changes in the enzyme, which are induced by ligand binding. In the selected-fit model, there is an equilibrium between multiple conformational states and the enzyme where the ligand can bind to some of them and stabilize the reactive conformation. These induced models describe the enzyme catalysis where the active sites are located on the protein surface. However, more than 60% of enzymes have their active sites placed in deep internal cavities connected to the bulk solvent by access pathways [13,14]. In this way, the keyhole-lock-key model incorporates the passage of the ligands through the tunnels (keyholes) to the catalytic site of the enzyme. The catalysis occurs with: (1) the passage of the ligand through the tunnel, (2) the reorganization of water, and (3) the binding to the catalytic residues [15].

2.2 ENZYMES CLASSES AND PROPERTIES The seven classes of enzymes are described, including the reactions they catalyze, the main group of enzymes that are included in each class, and some of their characteristics (Fig. 2.1).

FIGURE 2.1 Enzyme classification with catalyzed reactions and the main group of enzymes with some of their characteristics.

2.2 ENZYMES CLASSES AND PROPERTIES

15

2.2.1 OXIDOREDUCTASES Oxidoreductases or oxireductases catalyze the oxidation
reduction reaction in the form of A2 1 B-A 1 B2, comprising one-third of all enzymatic activities that are described in BRENDA [16]. Another key characteristic of oxidoreductases is the presence of cofactors such as flavin, heme, and other metal ions. Fungal oxidoreductases are comprised of the heme-containing peroxidases, which belong to the classical lignin-modifying peroxidases (LMPs), dye-decolorizing peroxidases (DyPs), hemecontaining peroxygenases, flavin-containing oxidases and dehydrogenases, copper-containing oxidoreductases, and lytic polysaccharide monooxygenases (LPMOs). Peroxidases (EC 1.11.1.x) catalyze the decomposition of hydrogen peroxide (H2O2) with concomitant oxidation of phenolic and nonphenolic substrates. They are ubiquitous in nature, being found in bacteria, fungi, algae, plants, and animals. They are related to various plant growth processes including cell wall metabolism, lignification, suberization, reactive oxygen species metabolism, auxin metabolism, fruit growth and ripening, and defense against pathogens. They can be successfully applied in various industrial sectors and medicine, with immunological and biotechnological applications [17]. LMPs (EC 1.11.1) include the lignin-peroxidases (LIPs, EC 1.11.1.14), manganese peroxidases (MNPs, EC 1.11.1.13), and versatile peroxidases (VPs, EC 1.11.1.16). These enzymes catalyze the two-electron oxidation of Fe31 by H2O2, forming an oxo-ferryl (Fe41) porphyrin cation radical complex that oxidizes two substrate molecules via one-electron abstraction. White-rot fungi such as Phanerochaete chrysosporium, Trametes versicolor, Pleurotus eryngii, and others typically secrete LIP and MNP. All LMP genes are present in white-rot basidiomycetes but are absent in brown-rot (cellulolytic) basidiomycetes’ and ascomycetes’ (soft-rot) genomes [18]. LIPs are related to the oxidation of recalcitrant polymers and nonphenolic compounds leading to detoxification and delignification processes, by the oxidation of veratryl alcohol and lignin. MNPs are involved in the oxidation of Mn21 to Mn31, mediating the oxidation of phenols, dyes, lignin, synthetic lignin, organopollutants, and bleach-colored compounds. VPs oxidize phenols, amines, dyes, and aromatic alcohols. The LMPs belong to the superfamily of plant peroxidases, which consists of three subclasses comprising (1) class I bacterial and eukaryotic organelle-localized heme peroxidases and catalase-peroxidases, (2) class II secreted fungal heme peroxidases, and (3) class III secreted plant heme peroxidases [17]. Unspecific peroxygenase (UPO, EC 1.11.2.1) and ascomycete chloroperoxidase (EC 1.11.1.10) were recently described and their biotechnological potential has been explored because they have oxygenation and oxidation properties [18]. UPO’s oxidation reactions combine the highly versatile oxygenation and oxidation of P450 monooxygenases with the simplicity of being an enzyme that reacts independently of a cofactor [19]. The DyPs (EC 1.11.1.19) were reported [20] to have the ability to oxidize lignin derivatives, dye substrates, nitrate mononitrophenols and Mn12 to Mn13, sharing some mechanisms of MnP and VP. Flavin-containing oxidases and dehydrogenases include aryl-alcohol oxidase (EC 1.1.3.7), methanol oxidase (EC 1.1.3.13), pyranose 2-oxidase (EC 1.1.3.10), glucose oxidase (EC 1.1.3.7), vanillyl-alcohol oxidase (EC 1.1.3.38), eugenol oxidase (EC, 1.17.99.1) cellobiose dehydrogenase (EC 1.1.99.18), and glucose dehydrogenase (EC 1.1.99.35). As H2O2-generating peroxidases, the

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CHAPTER 2 CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

flavin oxidases and dehydrogenases abstract two electrons from alcohol substrates and the reduced flavin is reoxidized by O2 forming H2O2 [18]. This class of enzymes has been considered for biorefinery applications such as biobleaching, flavor synthesis, oxidation of furfurals, and deracemization of chiral alcohols. Polyphenol oxidases (PPOs) belong to a class of copper (Cu)-containing enzymes performing the oxidation of phenolic compounds to o-quinones, which promote secondary reactions to form melanins and cross-linked polymers. PPOs are divided into(1) tyrosinases, (2) catechol oxidases, and (3) laccases. These classifications are based on the substrate specificity and their mechanism of action. While tyrosinases have both cresolase and catecholase activities, catechol oxidases, catalyze the oxidation of o-diphenols to o-quinones [21]. Laccases (Benzenodiol: oxygen oxidoreductases, EC 1.10.3.2) were initially identified in the Japanese tree, Toxicodendron verniciflua lacquer. Later, their presence in other plants, many insects, and several fungi, especially in ligninolytic basidiomycetes such as Pleurotus [22] was also identified. From the physiological point of view, fungal laccases are involved in developmental processes, morphogenesis, lignin degradation, pathogenesis, detoxification, and pigment formation [23]. In the degradation of lignin, laccases act in synergy with other enzymes or in the removal of toxic compounds formed during the process of degradation mediated by other enzymes [24]. Laccases catalyze the oxidation of phenolic compounds, carrying electrons that are used in the reduction of molecular oxygen to water [24
26]. In addition, they catalyze various reactions such as cleavage of the alkyl phenol bond, Cα
Cβ bonds and phenolic lignin dimers, demethoxylation, demethylation, polymerization, and depolymerization. Laccases belong to a class of oxidases, which do not require the addition or synthesis of cofactors of low molar mass, as in the case of H2O2 for peroxidase. The enzyme uses the molecular oxygen as cosubstrate, already available in the environment, reducing production costs [27]. Although laccases do not require the presence of a cofactor, some substrates cannot be oxidized directly by these enzymes because they are too large to penetrate its active site or because they have a very high redox potential. Some “chemical mediators” act by intermediating the laccase reactions with the substrate [21]. LPMOs are found in saprophytic fungi such as Thermoascus aurantiacus, Gloeophyllum trabeum, Lentinus similis, Pichia pastoris, and Neurospora crassa, as well as in bacteria such as Bacillus amyloliquefaciens and Enterococcus faecalis. The catalysis of LPMO substrates occurs with the binding of active oxygen molecules to the monomeric Cu21, resulting in their interaction with the active site. LPMOs have been used in cellulolytic cocktails because they act on biomass decomposition process through oxidative attack of the C
H bonds (i.e., C-1 and C-4) of cellulose chain using molecular oxygen as external electron donor [18,21].

2.2.2 TRANSFERASES Transferases catalyze the transfer of a specific group from one substance to another. The related groups include methyl, acyl, amino, glycosyl, or phosphate. Glutathione transferases (GSTs, EC 2.5.1.18) include enzymes that promote the detoxification of endogenous and xenobiotic electrophile compounds [28]. Rubber transferase is the common name of the enzyme cis-prenyltransferase (EC 2.5.1.20), which catalyzes the rubber molecule elongation [29], making a sequential condensation of isopentenyl pyrophosphate with prenyl groups [30].

2.2 ENZYMES CLASSES AND PROPERTIES

17

Plant GSTs have been extensively studied due to their herbicide detoxification action (e.g., triazines, thiocarbamates, chloroacetanilides, diphenyl ethers, and aryloxy-phenoxypropionates). Several GSTs are known to be responsible for herbicide tolerance in crops and resistance in weeds [28]. GSTs have been explored in the biosensors technology for herbicides detection because they are able to catalyze conjugation reactions. For example, atrazine was already determined using an immobilized isoenzyme GST I from maize. Cytosolic GSTs can be useful in the diagnosis and monitoring of cancer because GSTs are involved in several chemotherapeutics detoxification and therefore are considered potentially crucial in regulating the susceptibility to cancer [31].

2.2.3 HYDROLASES Hydrolases are enzymes that are able to hydrolyze various bonds and are designated by the EC number 3. According to their substrate specificity, different designations are employed: hydrolases acting on ester bonds (EC 3.1), such as the thioester hydrolases (EC 3.1); glycosylases (EC 3.2); hydrolases that act on ether bonds (EC 3.3); hydrolases that act on peptide bonds (EC 3.4), and others. A very important subclass is the glycoside hydrolases (GHs), which act over the glycosidic bonds between two or more carbohydrates [32]. This group, also called as glycosidases, is described by the Carbohydrate-Active Enzymes (CAZy) database [33]. Some examples of important hydrolases include amylases, xylanases, cellulases, lipases, and proteases. They are presented in detail in the next sections.

2.2.3.1 Amylases Amylases hydrolyze starch molecules to yield various products such as dextrins and successively polymers with fewer glucose units such as maltotriose, maltose, and glucose [34
36]. These enzymes acquire a spatial structure that permits them to wrap around starch and cleave it into smaller units, according to the catalyzed reaction [37,38]. Amylases can be classified into two categories according to how they cleave starch: endoamylases and exoamylases [34
36]. Endoamylases hydrolyze randomly α-1,4-glycosidic bonds in the inner part of starch, producing linear and branched oligosaccharides of diverse chain lengths [34,35,37]. Alpha-amylases (endo-1,4,α-D-glucan glucohydrolase, EC 3.2.1.1) are endoamylases widely applied in the industry [34,38] that randomly hydrolyze the α-1,4-glycosidic bonds in the interior of starch [34,36,38]. Exoamylases, instead, hydrolyze starch from its nonreducing end, progressively generating short products [34,35,37]. Beta-amylases (α-1,4-glucan maltohydrolase, EC 3.2.1.2) and glucoamylases (exo-1,4-α-D-glucan glucanohydrolase, EC 3.2.1.3) are examples of exoamylases [34,37,38]. Beta-amylases cleave nonreducing chain ends of the starch molecule, while glucoamylases hydrolyze single glucose units from nonreducing ends of amylose and amylopectin [37,38]. The most acknowledged catalytic mechanism of α-amylases is the α-retaining double displacement, which comprises two catalytic residues in the enzyme active site: an aspartate and a glutamate [37]. This mechanism contains five steps: (1) bond of the substrate and proton donation from the glutamate, in its acid form, to the glycosidic bond oxygen and aspartate nucleophilic attack to the C1 of a glucose; (2) assembling of an oxocarbonium ion-like transition state followed by the development of a covalent intermediate; (3) release of the protonated glucose from the active site

18

CHAPTER 2 CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

and insertion of a new water or glucose molecule in the active site, which attacks the covalent bond between the aspartate and the glucose molecule; (4) formation of other oxocarbonium ion-like transition state; and (5) glutamate, in its base form, obtains a hydrogen from the new water or glucose molecule, while the oxygen of the new molecule supplants the oxocarbonium bond between a glucose and the aspartate [37]. The other aspartate in the active site, however not participating directly in the catalysis, forms hydrogen bonds with the OH-2 and OH-3 groups of the substrate, strongly contributing to the substrate distortion [37]. The main characteristic of extracellular amylases is to reflect the temperature and pH of the growth media [38]. Most amylases are stable from a pH of 4.0 to 8.0 and the highest activity for many of them is achieved at 50 C [39]. In case of harsh conditions or industrial applications, amylases from extremophilic microorganisms are indicated. Amylase thermostability is of great interest for industrial applications, and, for most amylases, this characteristic is influenced by the presence of calcium ion in their structure [34,38
40]. These hydrolases can be synthesized by plants, animals, and microorganisms [34,38,41]. However, amylases from microbial sources are the most utilized in industry, majorly due to their economical bulk production capacity, ease of genetic manipulation, and enhanced enzyme stability [34,38,41]. Among bacterial sources, most of the commercial amylases are from the genus Bacillus, while fungal amylases are mainly from the genus Aspergillus. Thermostable amylases are obtained from thermophilic microorganisms and detain many commercial applications due to their stability in high temperatures [39].

2.2.3.2 Cellulases Cellulases comprise a complex of cellulolytic enzymes that act synergistically on the conversion of cellulosic substrates to glucose. According to the EC, the cellulases are classified as endoglucanases (EGLs, EC 3.2.1.4), reducing exoglucanases (EC 3.2.1.176), nonreducing exoglucanases (EC 3.2.1.91), and β-glucosidases (BGLs, EC 3.2.1.21) [42]. These multienzyme complexes are capable of hydrolyzing the β-1,4-glucosidic linkages of cellulose, producing mainly glucose, cellobiose, and cello-oligosaccharides. The enzymatic complex acts synergistically with the action of the EGLs, exoglucanases, also named as cellobiohydrolases (CBHs), and BGLs. EGLs produce nicks in the cellulose polymer exposing reducing and nonreducing ends, and then CBHs act upon them to liberate cello-oligosaccharides and cellobiose units. BGLs hydrolyze cellobiose to produce glucose and then complete the hydrolysis [43,44]. Cellulases are produced by a large spectrum of microorganisms such as bacteria, fungi, protozoa, and some animal species including termites and crayfish [44,45]. Fungi are among the most important and explored cellulase producers, especially those of the genus Trichoderma. The same microorganism can eventually secrete more than one type of cellulase. Fungi, mainly Trichoderma reesei and Aspergillus niger, can secrete 12 and 8 hemicellulases, respectively, in high concentrations [45]. T. reesei produces two CBHs (CBHI and CBHII), two EGLs (EGLI and EGLII), as well as seven types of BGLs (BGLI to BGLVII). The CBHI and CBHII together with EGLI and EGLII contribute to 90% of the secreted enzymes, in a ratio of 60:20:10:10, while all the seven BGLs constitute up to 1% of the secreted protein [42]. T. reesei possesses low levels of intracellular BGL as a way to minimize cellobiose feedback inhibition during cellulose hydrolysis [46]. Other fungal species produce different cellulase compositions. A. niger, for example, is a strong BGL producer

2.2 ENZYMES CLASSES AND PROPERTIES

19

comparing to T. reesei. Humicola insolens cellulase system lacks a cellulose-binding domain, and Humicola grisea produces a thermostable cellulase. According to Singhania et al. [44], cellulases are the third largest group of industrial enzymes and their demand increased since 1995 in several industrial applications, such as detergent, textile, animal feed, food, paper, and biofuel industries. Commercial hydrolases are mainly produced by submerged fermentation (SmF) by native or heterologous microbial hosts. Even so, many research projects and publications report the comparison of SmF with solid-state fermentation processes, as an attempt for enzymatic productivity improvement, reduction of costs, and improvement of the viability for commercial applications. The production of cellulases is traditionally carried out in batch SmF processes, mainly using the fungi of the genera Trichoderma and Aspergillus. The relatively recent developments of the cellulosic biofuel industry certainly boosted the interest on cellulases. The actual worldwide cellulosic ethanol production capacity is 480.5 million liters per year among which around 80% are produced using T. reesei cellulases [47]. According to the authors, more than 100 research articles are published per year related to Trichoderma cellulases. All this scientific data reflect the significance of this enzyme for the industrial sector and for a biobased economy.

2.2.3.3 Xylanases Xylanases are hydrolases of the subclass o-glycosylases (EC 3.2.1.8) that hydrolyze the xylan molecule, which is linked by β-1,4 chains of xyloses. Xylan is the second most abundant natural polysaccharide and the major constituent of hemicelluloses [48]. Hemicelluloses consist of complexes of polymeric carbohydrates that include xylan, glucomannan, xyloglucan, galactoglucomannan, and arabinogalactan [49]. Due to the wide variety of xylans, the xylanolytic enzyme system comprises different hydrolytic enzymes such as α-arabinofuranosidase (EC 3.2.1.55), acetylxylan esterase (EC 3.1.1.72), α-glucuronidase (EC 3.2.1.139), β-xylosidase (EC 3.2.1.37), and endoxylanase (EC 3.2.1.8) [50]. These enzymes act synergistically for the conversion of xylan to sugars [51
53]. Among the xylanases, the endoxylanases are directly involved in the hydrolysis of glycosidic bonds and liberation of small stretches of xylooligosaccharides (XOS) [54]. According to the analyses of genomic sequences, these enzymes present a domain that is involved in the degradation of polysaccharides called GH domain [55]. This catalytic domain that comprises between 250 and 450 amino acids is highly conserved, while the affinity differences between its subsites affect xylanases action, substrate preference and product preference [56]. According to the CAZy database (http://www.cazy.org), xylanases are divided into 14 different GH families, but sequence analyses have shown that only 6 out of 14 families contain real specific catalytic domains for xylanase activity [57]. Endo-1,4-β-xylanases, considering true xylanases, are subdivided into two large groups: GH 10 and GH 11 according to their amino-acid sequence, hydrophobic catalytic dominium groups, and the divergent evolution of their 3D structures [58]. The xylanases of the GH 10 family are part of the GH-A clan, generally have a more than 30 kDa and low isoelectric point (pI) values, and their tridimensional structure presents an eightfold (α/β) structure that forms a bowl shape [48,58]. The xylanases of the GH 11 family are part of the GH-C clan and have low molecular mass (,30 kDa) and high pI values [48]. Their tridimensional structure consists mainly of β-sheets that form a β-barrel structure (structure of the “β-jelly roll”). This family hydrolyzes preferably internal bonds of xylan [59]. Differences in the analysis of the GH 11 family crystal structure and kinetic

20

CHAPTER 2 CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

analyses of its activity on XOS showed that it has a high affinity for the long chains of XOS. This fact is probably due to larger substrate-binding clefts with at least seven subsites. Contrarily, GH 10 family has a greater affinity with short chains of XOS with approximately four to five substratebinding clefts [58,60]. The production of xylanases is generally carried out by bacteria, fungi, and protozoa. Some higher organisms such as marine algae, insects, gastropods, arthropods, and plants [57] were also reported. However, microorganisms such as fungi and bacteria are the most used for their production, especially filamentous fungi.

2.2.3.4 Lipases Lipases (triacylglycerol acylhydrolases, EC 3.1.1.3) are enzymes that catalyze the hydrolysis of acylglycerides to fatty acids, glycerol, monoacylglycerols, and diacylglycerols over an oil
water interface [61
63]. They are substrate-specific enzymes and commonly possess elevated stereo-, enantio-, and regioselectivity [61,63
65]. Most lipases hold optimal activity and stability in the pH range of 6.0
8.0 and in the temperature range from 30 C to 40 C, although these values may vary fairly depending on the lipase source [66]. Lipases active site is commonly portrayed as a triad of serine, histidine, and an acid residue, which can be aspartate or glutamate [66]. Lipases catalytic mechanism involves four steps: (1) serine deprotonation mediated by histidine and aspartate; (2) attack of the serine hydroxyl group to the carbonyl carbon from the substrate, resulting in an intermediate; (3) transfer of the acyl group to the lipase and discharge by the water nucleophilic attack; and (4) regeneration of the catalytic site [66]. Lipases are able to catalyze a large number of heterogeneous reactions both in water-soluble media and organic media [64,66,67]. Besides their hydrolytic activity, lipases can also perform synthesis, as esterification and transesterification, when in low-water content medium [61,62,64,66].

2.2.3.5 Proteases Proteases (EC 3.4), also referred to as peptidases or proteinases, consist of a group of hydrolytic enzymes that cleave peptide bonds in protein molecules. According to the E.C., proteases are classified in the group 3 and more specifically in its subgroup 4, which is classified according to substrate specificity, active site, charge, molecular size, and catalytic mechanisms [67,68]. The proteases classification is mainly based on the catalyzed reaction and their site of action. Considering the position of attack, exopeptidases (EC 3.4.11
3.4.19) are those that cleave at the end of the polypeptide chain and endopeptidases (EC 3.4.21
3.4.25, 3.4.99) are those that act in the middle of the chain [69,70]. According to the terminal end of the peptide chain, the exopeptidases can be named as aminopeptidases or carboxypeptidases. The aminopeptidases (EC 3.4.11) hydrolyze peptide bonds at the free N-terminal ends of the protein releasing only single amino acids. If the enzyme releases a fragment with two (dipeptides) or three (tripeptides) amino-acid residues from the N-terminal end, the enzyme is named dipeptidyl peptidase or tripeptidyl peptidase, respectively, being classified as EC 3.4.14. The dipeptidase (EC 3.4.13) can release a single amino acid from a dipeptide. The carboxypeptidases (EC 3.4.16
EC 3.4.18) liberate single amino acids from the C-terminal ends of a polypeptide chain, while peptidyl dipeptidases (EC 3.4.15) release peptides from the C-terminal ends of the protein [68,70,71].

2.2 ENZYMES CLASSES AND PROPERTIES

21

Depending on the type of functional group at the active site and the catalytic mechanism, the peptidases are named as serine endopeptidases (EC 3.4.21), cysteine endopeptidases (EC 3.4.22), aspartic endopeptidases (EC 3.4.23), metalloendopeptidases (EC 3.4.24), threonine endopeptidases (EC 3.4.25), and endopeptidases of unknown catalytic mechanism (EC 3.4.99). Serine proteases (EC 3.4.21) are described by having a serine group in their active site. They are abundant and common in both eukaryotes and prokaryotes. Every single serine protease contains a serine, a histidine, and an aspartate in the active site (catalytic triad). Serine proteases contribute to one-third of all known proteases. Most of the industrial serine proteases are produced by Bacillus sp. [72,73]. Cysteine proteases (EC 3.4.22) occur in both prokaryotes and eukaryotes also. The activity of them was determined by the catalytic triad at the active site (cysteine, histidine, and asparagine). Normally, the cysteine proteases have neutral pH optima. They are not so broadly distributed as was seen with serine and aspartic proteinases. The best-known cysteine protease is papain [74]. Aspartic proteases (EC 3.4.23) or aspartyl proteinases are an acid protease that has two aspartic acid residues in their active site. This class of peptidase performs better on peptide bonds between hydrophobic amino acids (Leu-Tyr, Phe-Phe, Phe-Tyr, etc.). The commercial aspartic enzyme is produced from filamentous fungi [75]. Due to the high structural diversity and specificity of the proteinases, their catalytic properties are varied, with different catalytic mechanisms [68,76]. Acid proteases are specific to aromatic or bulky amino-acid residues’ side chains at both sides of the cleaving bond. Aspartic proteases (EC 3.4.23) are acidic endopeptidases [77]. Neutral proteases have low thermal tolerance and have high affinity for nonpolar amino acids. Normally, cysteine endopeptidases (EC 3.4.22) have neutral pH optima [78,79]. The serine proteases (EC 3.4.21) are generally classified as alkaline, and several alkaline proteases are thermostable [68,77]. Proteases have numerous applications in medical, pharmaceutical, chemical, and food industries. The microbial production of proteases is the most employed, for presenting many advantages if compared with the extraction from plants and animals. Microbial proteases are extracellular, have lower production costs, and present good stability and specificity [68,77,79]. Most commercial proteases, especially neutral and alkaline proteases, are produced by bacteria of the genus Bacillus. Commercially important acid proteases are exclusively produced by filamentous fungi [75,78
80].

2.2.4 LYASES Lyases catalyze the addition or removal of chemical groups through nonhydrolytic bond-breaking reactions. These enzymes cleave C
C, C
O, C
N, C
S, and other bonds, leading to the formation of a double bond or a new ring or adding groups to double bonds [81,82]. The EC number assigns the lyases in class 4 (EC 4). There are eight subclasses of lyases, represented by the second number, which exemplify the bond type involved in the reactions: EC 4.1 carbon
carbon lyases, EC 4.2 carbon
oxygen lyases, EC 4.3 carbon
nitrogen lyases, EC 4.4 carbon
sulfur lyases, EC 4.5 carbon
halide lyases, EC 4.6 phosphorus
oxygen lyases, EC 4.7 carbon
phosphorus lyases, and EC 4.99 other lyases. These subclasses present subcategories, equivalent to the third number, according to the chemical group eliminated [83
85]. This enzyme group includes aldolases, decarboxylases, hydratases, and some pectinases, which catalyze the elimination of aldehyde, carbon dioxide, water, and pectin molecules, respectively [81,86]. They are present in metabolic and anabolic pathways, signal transduction, and in the

22

CHAPTER 2 CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

mechanism of DNA repair [87]. The keyhole-lock-key model is the catalytic property applied to the lyases, which have structures called tunnels and gates that regulate the transport of substances [88,89]. The formation of C
C bonds by the addition of a nucleophilic donor compound, which is usually a ketone, to an electrophilic aldehyde acceptor is catalyzed by aldolases [90]. These enzymes convert their substrates into the aldol products with high yield and specificity, which makes aldolase catalyzed routes for the synthesis of compounds such as carbohydrates, amino acids, and their analogs, from achiral and simple starting materials [91,92]. Aldolases are classified, according to their mechanism, into Type I or II. Type I aldolases, mainly found in plants and animals, are metal cofactor-independent and activate the donor by forming a Schiff base intermediate. On the other hand, the Type II aldolases occur mostly in bacteria and fungi, being Zn21-dependent in the active site to facilitate enolate formation on the donor [87,93]. The decarboxylases, or carboxy-lyases, remove the carboxyl group from a molecule, with the release of CO2, in a process called decarboxylation. They act on the decarboxylation of amino acids, α-keto acids, and β-keto acids [82,94
96]. Amino-acid decarboxylases are involved in the decarboxylation of certain amino acids with the consumption of protons and release of CO2. These enzymes catalyze the synthesis of amines such as cadaverine, putrescine, and β-alanine, catalyzed by lysine decarboxylase, ornithine decarboxylase, and aspartate 1-decarboxylase, respectively, which can be used in different products of the pharmaceutical, food, and chemical industries [95,97]. Hydratases are classified as hydro-lyases by performing the addition and elimination of water at nonactivated C
C double bonds [82,98]. These enzymes can be divided into two groups: cofactordependent hydratases, which add to activated α, β-unsaturated carbonyl compounds, and cofactorindependent hydratases [99]. There has been an increasing interest in exploiting the potential of hydratases from microorganisms, such as fatty acid hydratase, linalool hydratase isomerase, and carotenoid hydratase, for industrial applications [98]. Polygalacturonate lyase and polymethyl galacturonate lyase are pectin lyases involved in pectin degradation through the cleavage of α-1,4-glycosidic linkages in pectic acid and pectin. This occurs with the trans-elimination reaction, forming unsaturated galacturonates and methyl galacturonates [100]. Most pectin lyases reported in scientific studies have been obtained from microorganisms, especially those from the genera Aspergillus, Penicillium, and Fusarium; however, it is possible to observe their presence in plants and animals [101].

2.2.5 ISOMERASES Isomerases are a class of enzymes that constitute the fifth group of the EC Classification (EC 5). They catalyze intramolecular rearrangements or isomerization reactions and are divided into seven subclasses as a function of the type of catalyzed reaction: racemases and epimerases (EC 5.1) that catalyze either the racemization or epimerization of a center of chirality; cis
trans isomerases (EC 5.2) that catalyze the rearrangement of the geometry at double bonds; intramolecular oxidoreductases (EC 5.3) that catalyze the oxidation of a portion of a molecule with concomitant reduction of another portion; intramolecular transferases (EC 5.4) that transfer a group from one position to another within a molecule; intramolecular lyases (EC 5.5) that catalyze reactions in which a group is removed from one portion of a molecule, leaving a double bond, but remains covalently attached

2.2 ENZYMES CLASSES AND PROPERTIES

23

to the molecule (e.g., the breaking of a ring structure); isomerases altering macromolecular conformation (EC 5.6); and other isomerases (EC 5.99) [10]. The catalytic properties of several isomerases have been studied, especially of those with major industrial applications, such as xylose isomerase (also known as glucose isomerase) and sucrose isomerase. Xylose isomerase is a metalloenzyme with two divalent cations binding sites [102], so the presence of divalent metal ions is essential for catalytic activity. Also, thermal stability is desirable for some industrial applications. The most relevant application of isomerases is in the food industry, in the modification of sugars. Xylose isomerase (D-xylose cetol isomerase, EC 5.3.1.5) is often applied in the food industry to produce fructose syrup, through the conversion of D-glucose to D-fructose. It can also be used in the conversion of C-5 sugars derived from biomass to many biochemicals or biofuels (U.S. patents 8,114,974 and 8,093,037). For example, the conversion of xylose to its isomer xylulose makes it assimilable by Saccharomyces cerevisiae, allowing its conversion to ethanol [103]. Other enzymes related to the glycolytic pathway, such as phosphofructokinase and triosephosphate isomerase, have been overexpressed in Corynebacterium glutamicum to enhance the production of D-lactate from a glucose and xylose mixture [104]. Isomerases are essential enzymes in the metabolism of animals, plants, and microorganisms. Despite being present in many different sources in nature, isomerases are most efficiently synthesized by microbial species, either bacteria or fungi. Commercial glucose isomerase, for example, is obtained from Streptomyces murinus, Streptomyces rubiginosis, or Bacillus coagulans [105]. Sucrose isomerase has been identified in many bacterial strains including Enterobacter sp., Erwinia rhapontici, Klebsiella sp., Pantoea dispersa, Pantoea stewartia, Pectobacterium sp., Pseudomonas mesoacidophila, Protaminobacter rubrum, Raoultella planticola, and Serratia plymuthica. All these strains are plant-associated soil microorganisms [106]. D-Arabinose isomerase (EC 5.3.1.3) is mainly obtained from Klebsiella pneumoniae and Escherichia coli [107].

2.2.6 LIGASES The sixth class of enzymes is represented by the ligases (EC 6). Most of them are involved in biologically essential reactions in the central metabolism at cellular level. They catalyze the attachment of two molecules or of parts of them. In some cases, the terms synthase or carboxylase can be used to designate the ligases [e.g., NAD 1 synthase (EC 6.3.1.5), and pyruvate carboxylase (EC 6.4.1.1)] [100]. Ligases catalyze condensation reactions, that is, the formation of carbon
carbon, carbon
sulfide, carbon
nitrogen, and carbon
oxygen, or phosphoric
ester and nitrogen
metal bonds. This class has been divided into six subclasses as defined by the catalyzed reactions, according to Table 2.2. These reactions require energy, in this sense, a phosphate bond is simultaneously hydrolyzed from a nucleotide to release energy and allow the reaction to occur spontaneously. The nucleotides carry chemical energy in the form of adenosine triphosphate (ATP), guanosine triphosphate, cytidine triphosphate, and uridine triphosphate, all named as nucleoside triphosphates [108]. The DNA ligases (EC 6.5.1.1) or polydeoxyribonucleotide synthases are one of the most important tools in the construction of recombinant DNA molecules. They catalyze the formation of a phosphodiester bond between the 50 -phosphate group of one DNA chain with the adjacent

24

CHAPTER 2 CLASSIFICATION OF ENZYMES AND CATALYTIC PROPERTIES

Table 2.2 Reactions catalyzed by ligases and characteristics of the condensation reactions. E.C. subclass

Forming bonds

Characteristics

6.1

Carbon
oxygen

6.2

Carbon
sulfur

6.3

Carbon
nitrogen

6.4

Carbon
carbon

6.5

Phosphoric
ester

6.6

Nitrogen
metal

The reaction on this subclass is characterized by the acylation of a tRNA with the corresponding amino acid (amino-acid-tRNA ligases; EC 6.1.1), (e.g., tyrosine-tRNA ligases; EC 6.1.1.1) For this reaction, the enzymes catalyze the attachment of an acyl group to the sulfur atom of 40 -phosphopantetheine groups in coenzyme A and acyl-binding proteins, or of two cysteine residues forming a cystine The sub-subclasses are Acid-ammonia (or amine) ligases (amide synthases; EC 6.3.1); Acid-amino-acid ligases (peptide synthases; EC 6.3.2); Enzymes forming heterocyclic rings (cyclo-ligases; EC 6.3.3); Other carbon
nitrogen ligases (EC 6.3.4); Enzymes using glutamine as amido-N-donor (EC 6.3.5) These are the carboxylating enzymes, which are mostly biotinyl proteins in a single sub-subclass (EC 6.4.1) This subclass contains a single sub-subclass (EC 6.5.1). This subclass contains enzymes that restore broken phosphodiester bonds in nucleic acids (often called repair enzymes) This subclass contains a single sub-subclass for enzymes that form coordination complexes, that is, form nitrogen
metal bonds (EC 6.6.1)

30 -hydroxyl terminal of another chain. Commercially, the DNA ligases are used in molecular biology, nucleic acid research, and next-generation sequencing applications [109].

2.2.7 TRANSLOCASES Translocases are a new EC class (EC 7) that catalyze the translocation of ions or molecules across cell or their separation within membranes, frequently involving the hydrolysis of ATP. For this reason, some of these enzymes were previously classified as ATPases in the class of hydrolases (EC 3.6.3), although hydrolysis was not their primary function. According to the type of ion or molecule translocated, they are divided into six subclasses: enzymes that catalyze the translocation of hydrons, hydron being the general name for H1 in its natural abundance (EC 7.1); enzymes that catalyze the translocation of inorganic cations and their chelates (EC 7.2); enzymes that catalyze the translocation of inorganic anions (EC 7.3); enzymes that catalyze the translocation of amino acids and peptides (EC 7.4); enzymes catalyzing the translocation of carbohydrates and their derivatives (EC 7.5); and finally, the enzymes catalyzing the translocation of other compounds (EC 7.6) [110].

REFERENCES

25

2.3 CONCLUSIONS AND PERSPECTIVES Enzymes are powerful biocatalyzers that are intensively and continuously studied due to their high and diversified applications in the biotechnology industry and human and animal health. The seven classes of enzymes were presented. They are mainly classified according to the catalyzed reactions: oxireductases, transferases, hydrolases, lyases, isomerases, ligases, and the new recently defined class, the translocases. They are all extensively studied and the information about their classification, identification, and sequence is available in different databases developed in universities and research institutes (BRENDA, PDB, CAZy, EXPLORE-ENZ, and others), which are highly specialized in the subject. With this organized and significant information about the classification and catalytic properties of enzymes, numerous advances and the search for new enzymes will certainly be easily handled by researchers from all over the world.

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FURTHER READING F. Bibi, M. Irshad, Z. Anwar, K.H. Bhatti, Improved catalytic functionalities of purified pristine and chitosanimmobilized polygalacturonase and pectinlyase, Chem. Eng. Res. Des. 128 (2017) 146
154. M.L. Polizeli, A.C. Rizzatti, R. Monti, H.F. Terenzi, J.A. Jorge, D.S. Amorim, Xylanases from fungi: properties and industrial applications, Appl. Microbiol. Biotechnol. 67 (2005) 577
591. S. Riva, Laccases: blue enzymes for green chemistry, Trends Biotechnol. 24 (2006) 219
226. W.W. Windish, N.S. Mhatre, Microbial amylases, Adv. Appl. Microbiol. 7 (1965) 273
304. S. Yan, G. Wu, Secretory pathway of cellulase: a mini-review, Biotechnol. Biofuels 6 (2013) 1
12.

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ENZYMES AND THEIR PRODUCTION STRATEGIES

3

Himanshu Sharma1,2 and Santosh Kumar Upadhyay1 1

Department of Botany, Panjab University, Chandigarh, India 2I.K. Gujral Punjab Technical University, Jalandhar, India

3.1 INTRODUCTION With an exponential increase in knowledge about biochemical processes, it will be true to say that it is impossible to imagine any biological process without enzymes. Enzymes are biocatalysts, which increase the reaction rate to many folds. Enzymes are highly specific and remain unutilized throughout the process. Except for catalytic RNA molecules, all enzymes are proteinaceous in nature and are required for all living organisms [1]. All living organisms produce enzymes, and therefore there can be three different sources of enzymes: microorganisms, plants, and animals. The cost of the production increases with the increase in complexity of the system. Microbes, being the simplest system, are commonly used for enzyme production. Further, factors including low energy, low cost, nontoxic and eco-friendly nature make them popular for many industrial processes [2
4]. Moreover, milder conditions of temperature and pressure are required for the functioning of enzymes, which makes them a favorable alternative to the hazardous chemical catalysts [4
6]. Conventionally, enzymes have been used for making wine, beer, bread, cheese, vinegar, and in the manufacturing of leather and linen. However, the purified form of enzymes has found extensive applications in the industry just a few decades ago [7]. In 1960, plants and animals were the most common source for the production of enzymes [8]. However, approximately after two decades, most people started enzyme production from microbes [9]. The animal-derived enzymes including catalase, lipase, chymotrypsin, trypsin, and rennin are still relevant and represent around 10% of the enzyme market [8]. These enzymes are widely used in food and leather industries. As per Illanes [8], the enzymes produced from plants represent around 5% of the world enzyme market. For instance, papain (derived from Papaya) is an enzyme commonly used in meat tenderization, stain removal, yeast extract production, beer clarification, cosmetics, and medicines [10
13]. Microbial enzyme production covers the remaining market of enzymes. In the year 2017, the global enzyme market size was around USD 6.3 billion and it is estimated to grow at around 6.8% compound annual growth rate (CAGR) up to 2024 [14]. Furthermore, the US enzyme market is expected to hold the dominance; however, Asia-Pacific enzyme market share is expected to get gains of over 7.3% up to 2024. China, India, Japan, and South Korea might be sharing a significant share of enzyme industries [14].

Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00003-X © 2020 Elsevier B.V. All rights reserved.

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CHAPTER 3 ENZYMES AND THEIR PRODUCTION STRATEGIES

Enzymes produced from the microbial system are widely used in various sectors including agriculture, food, chemicals, medicine, and energy. [6]. Moreover, manipulation of microbial strains through DNA recombinant technology and protein engineering techniques has made it possible to meet the increasing demand for enzymes [15]. Furthermore, the microbes are metabolically dynamic, easy to grow on a large scale by fermentation, require simple nutrients, and are not affected by seasonal conditions [8,16]. In addition to the advantages, enzymes and their production methods have some limitations in the field of healthcare and other industries as they may denature under nonoptimal conditions such as alteration in temperature and pH [6]. It may ultimately lead to the discouragement of enzyme use in such conditions; however, they have ample applications under optimal conditions. Additionally, large-scale production of enzymes is a costly process and it directly affects the cost of the final product and indirectly affects the industrial use of such enzymes. Equipment, raw material, and installation are the reasons behind the capital-intensive process of enzyme production [17]. For instance, raw material solely contributes to 28% of operating cost in lignocellulosic enzyme production processes [18].

3.2 ENZYMES AND THEIR CLASSIFICATIONS Enzymes are made up of multiple amino acid molecules, where each amino acid residue is joined with a covalent bond to its neighbor. This amide linkage is known as a peptide bond. They are large macromolecules ranging between kilodaltons to megadaltons in terms of molecular mass [6]. Enzymes are generally made of 20 different amino acids. Asparagine was the first amino acid to be discovered in 1806 [1]. The amino acid residues are exclusively L-stereoisomers in proteins. Merely, small peptides such as a few bacterial cell wall peptides and some peptide antibiotics have D-amino acids isomers. The catalytic activity of enzymes usually depends upon the integrity of protein conformation and is lost if an enzyme is denatured [1]. Enzymes have a specific site called an active site, which is the region of binding of the enzyme to its substrate to form the product. An active site is usually a groove or cleft on the surface of an enzyme that forms a non-polar environment, which helps in efficient binding of a substrate [19]. The enzyme-substrate complex is formed with the help of many weak bonds. Catalytically active residues present in the active sites act on the substrate to convert it into the first transition state and then into the product. The enzyme remains unused and can be used in another cycle of reaction. Substrate binding and catalytic activity are also enhanced from some small non-protein molecules participating along with the enzyme. These molecules are termed as prosthetic groups, cofactors, and coenzymes and extend the catalytic capabilities of some enzymes [20]. Conventionally, there are several names of enzymes, which increase the ambiguity in the names of enzymes. For instance, trypsin name does not denote its name from its substrate and does not tell much about the enzyme. To avoid such ambiguity, it was a necessity to regulate the nomenclature of enzymes internationally. The International Union for Pure and Applied Chemistry along with the International Union of Biochemistry and Molecular Biology established an Enzyme Commission (EC) to issue the guidelines for nomenclature and enzyme classification [21]. This system of classification and nomenclature was complex but unambiguous. In the new system, a unique

3.3 ENZYME PRODUCTION

33

name and code are given to each enzyme that reveals the nature of reaction catalyzed and the substrate involved. For example, adenosine triphosphate (ATP): D-hexose-6-phosphotransferase (EC 2.7.1.1.), commonly known as hexokinase. The code refers to the enzyme of class 2 (transferases), subclass 7 (phosphoryl group transfer), and sub-subclass 1 (alcohol as a phosphoryl acceptor) [19]. Enzymes are broadly classified into six classes by International Union of Biochemists. 1. Oxidoreductases: This group includes the enzymes that catalyze oxidation and reduction reactions. Some examples include dehydrogenase, oxidase, oxygenase, and peroxidases. 2. Transferases: Groups of atoms are transferred from the first molecule to second using these enzymes. Fructosyltransferases, acyltransferases, transketolases, and transaminases are some of the enzymes of transferases. 3. Hydrolases: They help in the hydrolytic cleavage of bonds, such as C
C, C
O, C
N, P
O, and some other bonds such as acid anhydride bonds. Some of the hydrolases are proteases, lipases, cutinases, phosphatases, amylases, and acylases. 4. Lyases: These enzymes catalyze nonhydrolytic cleavage of bonds by elimination, leaving double bonds, and the addition of groups to double bonds. Some examples include pectate lyase, hydratase, dehydratase, decarboxylase, fumarase, and argininosuccinase. 5. Isomerases: This group includes the enzyme catalyzing the structural or geometric change in a single molecule. It includes the transfer of a group from one position to another in a single molecule. Isomerases, racemases, and epimerases are some examples of this class. 6. Ligases: They catalyze the covalent binding of two molecules together with the hydrolysis of a pyrophosphoryl group in ATP or similar nucleoside triphosphate. Ligases and synthases are some examples.

3.3 ENZYME PRODUCTION Since enzymes have a number of applications, there is a hustle in the high production of economically important enzymes. The production ranges from small- to large-scale depending upon the mode of use. Generally, a very refined enzyme is required in the field of healthcare and research [22]. Small-scale production of enzymes is suitable for the production of such specialty enzymes. However, there are other enzymes that are produced in bulk and in tons as per the demand of the industry. The level of production depends on the application and the kind of process used for production [8]. Apart from the glycozymes, which are only produced in plants [23] and animal cell culture [24], all other enzymes are preferably produced in a microbial host. The recombinant DNA technology has become the most advanced helping aid to the improvement of microbial strains. Specific gene is being introduced into the microbial hosts such as Escherichia coli, along with the promoter sequence for an inducer. As soon as the inducer is added to the culture, the gene starts expressing itself and produces the specific enzyme. Moreover, the final yield of the enzyme is an important factor for industries. Recombinant technologies are used to achieve such targets. For instance, Aspergillus oryzae alkaline protease genes have been expressed with native signal peptide and α-factor secretion signal peptide in Pichia pastoris GS115, the yield of recombinant protein has been found to be about 1.5-fold increased, which is approximately 512 mg/L [25]. In another example, R-selective transaminase (R-ATA) from Arthrobacter sp. was produced by the companies

34

CHAPTER 3 ENZYMES AND THEIR PRODUCTION STRATEGIES

Codexis and Merck, which has shown the capacity of converting 200 g/L of prositagliptin ketone to sitagliptin in dimethyl sulfoxide with a purity of 99.95% enantiomeric excess. Furthermore, a 10% increase in yield and a 53% increase in productivity have been achieved as compared to the conventional methods [26].

3.3.1 MICROBIAL ENZYME PRODUCTION As discussed earlier, the microbes share around 90% of the enzyme market. This extensive use of microbes for the production of enzymes is due to their characteristics such as rapid multiplication, easy to manipulate, easy to handle, availability, and easier downstream processing. Microbial enzymes are produced by the fermentation processes. The purification process of enzymes differs according to the mode of their secretion (intracellular or extracellular). For extracellular enzymes, the spent medium is used for purification, whereas intracellular enzymes need the cell rupture processes [27]. Furthermore, strain improvement is the most important part of the microbial enzyme production. Strain improvement is engineering of microbial strains, which includes the addition of a gene into a host for the production of a specific enzyme. The production strain must fall in the generally recognized as safe (GRAS), status given by the Food and Drug Administration, United States. In the United States, food industry enzymes are mandatory to be produced by the organism of GRAS status [8]. It is difficult to achieve a GRAS status for a new organism; therefore, it is preferred to clone an enzyme structural gene into a host of GRAS category [28]. In the case of mycelial microorganism, the morphology and rheological properties are other factors to be considered prior to the enzyme production. Specific activity of an enzyme is the units of enzyme activity per unit mass in the microbes and is one of the important factors for high yield and cost reduction [8]. The high specific activity can be achieved by controlling the environmental factors and genetic manipulation. The significant increase in specific activity of an enzyme will lead to the reduction of cost of fermentation, as well as the cost of downstream processing. A general overview of the enzyme production process has been shown in Fig. 3.1. The upstream processes for the enzyme production include the selection of strain or inoculum and media sterilization. Frozen culture of the strain is inoculated into the medium for primary culture. The media is sterilized prior to the inoculations for primary as well as secondary culture. After inoculation, the primary culture is allowed for the primary fermentation. The primary culture is used as an inoculum for the secondary fermentation process. The downstream processing of the fermented product depends on the nature of the secretion of an enzyme (intracellular or extracellular). If the enzyme is intracellular, the cells are first collected with the help of centrifugation techniques and other filtration techniques. Disruption of cells using different techniques such as sonication and French press is required to get the intracellular content. After the cell disruption, the cell debris is excluded with filtration techniques. In the case of extracellular proteins, the enzymes are secreted in the medium, which is further purified. The product of both intracellular and extracellular fermentation is further subjected to the concentrators for the concentration of the enzyme. The concentrated enzyme is finally purified using various chromatographic techniques. The purification of an enzyme leads to the process of formulation. The formulation may include techniques such as drying, vacuum drying, spray drying, and freezedrying. This process decides how an enzymatic product will be launched in the market. It can be a mixture of two or more enzymes. The formulation is solely based upon the research and

3.3 ENZYME PRODUCTION

35

FIGURE 3.1 General overview of microbial enzyme production.

development of an industry or a lab. All the upstream and downstream processes lead to the formation of the final product.

3.3.1.1 Fermentation Microbial enzymes are produced in a controlled environment with the help of fermentation. Nowadays, microbial strains can also express foreign genes to produce recombinant proteins. There have been experiments with the types of fermentation processes to increase the yield of an enzyme cost-effectively. The submerged fermentation under strictly controlled conditions has been widely used. This method was exponentially developed during the Second World War to fulfill the requirement of antibiotics [29]. It has been the most important area of bioprocess engineering since its inception [8]. The technology of submerged fermentation is highly developed and automated [30] and is the choice for production of many enzymes [31]. Microorganisms are cultivated in a liquid medium with suspended nutrients. As the microbes breakdown the nutrients, the desired enzyme is produced. Parameters including pH, temperature, oxygen consumption, and carbon dioxide are required to be optimized and controlled during the fermentation process. Batch fermentation is the simplest and traditional form of submerged fermentation. The fermenter is filled with the sterilized medium. The enzyme producing strain is inoculated in the medium and incubated under strictly

36

CHAPTER 3 ENZYMES AND THEIR PRODUCTION STRATEGIES

controlled conditions until the enzyme produced reaches to the maximum level. After the whole process, the enzyme is recovered and subjected to downstream processing. The advanced version of the batch reactor is a fed-batch reactor, in which the nutrients are fed after batch cultivation under controlled and sterilized conditions. When the growth of cells reaches the final volume, the enzyme is recovered and purified [8]. As per some publications, the fed-batch reactor is an appealing procedure as it allows the monitoring of metabolic responses of cells along with the simplicity in its operation [32,33]. For example, recombinant β-galactosidase cultivation in fed-batch culture with dissolved oxygen (DO)-stat feeding strategy has produced a low biomass of 18 g/L with a high enzymatic activity of 42,367 U/L [34]. Moreover, it leads to the specific activity of recombinant β-galactosidase to be 40 U/mg [34]. Continuous culture is another form of submerged fermentation, which has been used to study enzyme regulation mainly due to better operational control [35,36]. It involves the addition of fresh media and removal of bioreactor fluid, continuously. Furthermore, the cells can also be immobilized in continuous culture to enhance the productivity of the reactors [37]. The productivity of enzymes in continuous culture is higher than the other two methods; however, there is always a risk of contamination during such processes. The contamination can deplete the whole batch of culture, thus the industry remains reluctant to adopt this method [8]. Many enzymes are also being produced by the solid-state fermentation including cellulolytic enzymes [38], lignocellulosic enzymes [16], and some hydrolases such as amylase [39,40], phytase [40,41], and protease [42]. Solid-state fermentation is also being used for cellulolytic enzyme production from agriculture waste for biofuel purposes [43]. This mode of fermentation represents a good option for low-cost enzyme production [8,44,45]. Localization of the enzyme is an important factor to be considered for enzyme production. Most of the enzymes are intracellular; however, the extracellular enzymes are a significant portion of commodities enzymes [8]. Extracellular enzymes are easier to downstream and thus give a high yield at a low cost. Genetic engineering techniques can be used to make an intracellular enzyme as an extracellular [46]. Furthermore, there are some enzymes that are intracellular in one and extracellular in the other organism [8]. For instance, invertase is generally intracellular in Saccharomyces [47]; however, it is found to be excreted in Streptomyces [48,49] and Candida [49]. Strain improvement experiments are the key to the high yield of enzymes. In addition, the medium used for the fermentation also plays a significant role in the yield. Thus, the medium design is another demand for the enzyme production industry. Furthermore, specific inducers and repressors are used for the control of the synthesis of enzymes. These are used at the level of transcription of the gene. The choice of inducers and repressors has a direct effect on enzyme production. During the precisely designed fermentation process, the inducer or repressor is added to control the enzyme production. Usually, cyclic adenosine monophosphate (AMP) is a positive modulator in gramnegative bacteria. It avoids the blockage of the structural genes encoding the enzymes by a repressor protein [50]. In the case of gram-positive bacteria, yeasts and molds other signal molecules such as polyphosphorylated nucleotides, and cyclic guanosine monophosphate (GMP) are involved in controlling the mechanism of enzyme production [51,52]. The fermentation medium should have an ample amount of inducer in it for efficient production. Structural analogs are preferably used than that of the substrate itself as an inducer. Apart from these, the factors including the growth rate of the microbe, its morphology, and other rheological characters are considered during the processes.

3.3 ENZYME PRODUCTION

37

Another enzyme production technique is a cell-free system. It is an in-vitro approach for the production of enzymes. This system mimics the natural enzyme production process of a whole cell. The cell extract is prepared by mechanical or chemical denaturation of cell wall. Further, substrates, salts, and a DNA template are added for the activation of mRNA and protein synthesis. The technique has been widely accepted for fast protein synthesis [53]. Cell-free system has been used for the synthesis of fusion proteins that may potentially serve as B-lymphoma vaccine. Large-scale production of enzymes with cell-free system is quite difficult and expensive; however, some industries are striving toward the scaling up of cell-free system of enzyme production [54].

3.3.1.2 Recovery The fermentation leads to the chain of downstream processes. Solid
liquid separation is the initial requirement for the recovery of enzymes. Now again, if the enzyme is intracellular, then the cells are recovered. If the enzyme is extracellular, then the medium is used for further processes. Separation can be performed with the help of centrifugation or filtration. Unicellular organisms including bacteria and yeasts are preferentially separated by centrifugation. However, prior flocculation of these organisms is required for the efficient recovery of cells [8]. For organisms, which have filamentous morphology, such as actinomycetes and molds, filtration is the preferred method for solid
liquid separation. Because a compressible cake will be formed, a filter aid is required to be used. The frequently used filters are plate and frame filters and rotary filters [8]. Tubular centrifuges or disk-type centrifuges are used for the process of centrifugation. The microbial cells are smaller and their density is comparable to the spent medium; thus the efficiency of this separation is quite low. Cross-flow microfiltration is an alternative to these methods and is widely accepted [55]. Improvements in various filter membranes and design of microfilters are a need to present purification techniques. Furthermore, the cells are collected in the case of intracellular enzymes. Some enzymes are periplasmic and are easier to extract. They need mild procedures such as osmotic stress. On the other hand, some enzymes are bound to membranes or are in internal cell structures and are difficult to be extracted. In such cases, the cells require cell disruption as well as the use of several detergents that may lead to enzyme denaturation [8]. The methods for the cell disruption of such enzymes include mechanical cell rupture, cell permeability by membrane damage, and thermolysis. Cell rupture methods include pressure, homogenization, sonication, decompression, milling, freezing-thawing, and thermolysis [8]. Cell permeabilization methods that are common in use include alkali treatment, enzymatic lysis, and autolysis [8]. Mechanical disruption methods are generally used for large-scale production of enzymes. Homogenizers and beads mills are the most commonly used methods for cell disruption in industries. Homogenizers force the cells to pass through a very narrow passage under high pressure, which ultimately burst the cells. In bead mills, the cell suspension is mixed with the beads, then subjected to intense stirring, where compression and shear forces lead to cell rupture. Cell permeabilization using enzyme digestion is also very common. For instance, helicase from Helix pomatia has been used for cell disruption of yeasts and molds [8,56]. To reduce the cost of enzyme production on such a large scale, further modifications in these methods are required. The extracellular proteins are recovered from the spent medium of the filtration. The main concern in the production of extracellular proteins is the low concentration of excreted protein in the medium. According to Liu and group, extracellular protein concentration does not increase

38

CHAPTER 3 ENZYMES AND THEIR PRODUCTION STRATEGIES

significantly even under dense culture conditions [57]. Thus the concentration of such enzyme is required. Vacuum evaporation is the most conventional method used for the concentration of such enzymes [58,59]. High-throughput crystallizers have also been designed, which do the concentration of enzyme by water freezing [60]. Ultrafiltration is considered as the best option for the enzyme concentration [8]. The extracted protein has a number of undesired cells and other debris. The debris is removed from the enzyme prior to further purification steps. The debris can be removed by solid
liquid separation techniques including centrifugation, dead-end filtration, and microfiltration. Another method is the aqueous two-phase extraction, which is reported to be a promising method for cell debris removal as well as protein fractionation [61,62]. This is a liquid
liquid extraction where two phases are polymers or one phase is a polymer and the other is a salt [63]. A variety of polymers including poly(oxyethylene glycol), mono ethers, polyoxyethylene alkyl ethers, etc. can be used for this method and has become a conventional operation in pharma and chemical industries. Polyethylene glycol-dextran (PEG/dextran) and polyethylene glycol-salt (PEG/salt) are amongst the most common systems [8].

3.3.1.3 Purification The recovered extracellular enzymes contain the impurity of only some extracellular enzymes and require very less processing [55]. However, the intracellular enzymes contain impurities such as other enzymes, nucleic acids, and cell-constituents, and thus require complex purification. Purification also increases the specific activity of an enzyme during enzyme immobilization [8]. The extracellular enzymes are usually sold as commodities and do not require further purification, for example, hydrolases. Small molecular ions or solutes are removed from the recovered intracellular enzymes by size-exclusion chromatography or diafiltration. The removal of nucleic acid from the intracellular enzyme is customary. Ammonium sulfate is widely used for the removal of nucleic acid and it also serves the purpose for the precipitation of proteins. Unlike other salts, ammonium sulfate is harmless to the enzymes. In cases where more specificity is required, other expensive reagents such as polyethyleneimine, streptomycin sulfate, cetyltrimethyl ammonium bromide (NH4Br), and protamine sulfate are used. However, they are expensive and should only be used where nucleic acid contamination is unacceptable [8]. Bovine pancreatic nucleases treatment has been found to be the most effective treatment in removing the nucleic acids from the enzymes [8]. A series of protein fractionation is required to remove the protein contamination from the recovered enzyme. Salting out is a basic yet very fast technique for protein precipitation. Salting out precipitation is rapid and equilibrium can be achieved easily. However, a good separation can only be achieved by high centrifugal forces, which is not possible in many cases [8]. Conventionally, water
immiscible organic solvent precipitation technique has been used for the fractionation of many enzymes. However, it is required to be done at very low temperature [64]. A three-phase partition system has been developed, which is a combination of salt and solvent precipitation. High purification is achieved by concentrating the enzyme at the water
solvent interface [65,66]. Furthermore, liquid chromatography is an efficient technique for the purification of enzymes and has been widely accepted. In this, solute (enzymes) is dissolved in a mobile phase and passed through a stationary phase (generally a solid matrix). The enzyme gets separated on the basis of varying strengths of interaction with the stationary phase [19]. Column chromatography is broadly used for purification of proteins and enzymes at industrial as well as laboratory scale. In the column chromatography,

3.3 ENZYME PRODUCTION

39

a stationary phase is packed into a column and the mobile phase containing enzyme is allowed to pass through it [1]. These chromatographic techniques are difficult to be scaled up to an industrial level. Size-exclusion chromatography, gel filtration, is another method common in the purification of proteins and used in the final step of enzyme purification [67]. It separates proteins according to their molecular size [19]. Larger molecules leave the column earlier, whereas the small molecules enter the porous structure and are mostly delayed, thus separating proteins. Dextrans, polyacrylamide, polymethacrylate, and agarose are some of the matrices used for size-exclusion chromatography. Scaling up a column is cumbersome and ultimately leads to loss of resolution; thus stacked columns are used for scaling up the process [8]. Ion-exchange chromatography is another technique widely used in laboratories as well as industries. Commercially significant enzymes are purified by ion-exchange chromatography [68]. Ionexchange resins are used as chromatographic support in this method. The interactions between the charged amino acid residues of enzymes with the chromatographic matrix lead to the separation of enzymes [19]. As the protein passes the column, they bind according to their surface charge and pH. The amphoteric nature of proteins makes it helpful to choose a cationic or anionic exchanger. If pH of a protein is more than its isoelectric point, they show a negative charge and bind to anion exchangers, whereas if pH is less than its isoelectric point, they possess a positive charge and tend to bind to cation exchangers. Efficient elution of proteins can be achieved by differing ionic strengths. Some examples of ion exchangers include diethylaminoethyl, triethylaminoethyl, quaternaryaminoethyl, and carboxymethyl. [8]. Affinity chromatography is known to be the most powerful technique of protein purification. It is widely accepted for recombinant protein production because of its high resolution. It is based on the functional properties of an enzyme [69]. The enzymes are specifically purified on the basis of their catalytic properties and immunogenic properties [19]. The yield of this process is high; however, it is costly and difficult to scale-up. The technique is suitable for laboratory-scale production of enzymes and for the enzymes of very high value. Research is going on to reduce the operational and material costs for such a powerful chromatography. A specific ligand interacting with the desired enzyme is attached to the matrix. Ground matrix is usually a polymer, which does not interact with the enzyme. Agarose gel is extensively used as a ground matrix in affinity chromatography techniques. The protein-ligand binding in the process becomes the basis of the separation [19]. Ligands used in such experiments can be a molecule that binds specifically to the enzyme. Substrates, coenzymes, and analogs are the choice of specific ligands for an enzyme. The ligand selection is crucial for the step for the whole process of enzyme purification by affinity chromatography. Hydrophobic chromatography is another form of chromatography used in the process of enzyme fractionation [66,70]. The separation is on the basis of binding of the hydrophobic portion of the enzyme with a hydrophobic gel matrix. The hydrophobic aryl or alkyl groups are coated onto the gel matrix prior to use. Ionic strength should be high for efficient fractionation. For elution, ionic strength or pH of eluent is modified [8]. The high degree of enzyme purification is a demand for some purposes, such as therapeutic use, clinical use, or research. High-end techniques are required for such fine purification, which ultimately increases the cost of the process. The most commonly used method in the industry is the high-pressure liquid chromatography (HPLC). The reverse-phase HPLC is suitable for enzyme fractionation, where enzymes are differentially retarded according to the hydrophobic interaction with

40

CHAPTER 3 ENZYMES AND THEIR PRODUCTION STRATEGIES

the stationary phase [1,19]. The desired enzymes are then eluted by increasing the concentration of organic solvent in the mobile phase. Protein unfolding is a major problem during HPLC purification [8]. Nowadays, a combination of chromatographic techniques has also been used for the purification of proteins.

3.3.1.4 Formulation Formulation of the enzyme includes finishing of the product. The industrial enzymes have to be gone under strict regulations prior to their commercialization [27]. The formulation includes stabilization, standardization, and final polishing of an enzyme. In the case of commodity enzymes, they require final polishing including removal of remaining salts and pH adjustments of the enzyme [8]. For enzymes that are produced for some special purpose such as medicines, they have to undergo stringent processes. These processes include the removal of contaminations such as pyrogens, endotoxins, nucleic acids, and viruses [8]. Furthermore, the shelf life of industrial enzymes is another issue to be resolved. Enzymes in the solid phase are easier to handle and transport; however, the production process has a high risk of allergy among workers due to the dust. Although the shelf life of liquid enzymes is low, they are easy to use and containment is simple. For solid preparations, vacuum drying, spray drying, and freeze-drying are the frequently used methods [9,71,72]. Liquid enzymes require simple finishing steps including desalting and concentration. Diafiltration or size-exclusion chromatography can be used for such desalting operations [73,74]. Industries are also required to obtain safety certificates for the use of enzymes from concerning authorities. For instance, the enzymes used in the detergent industry will require to obtain the certificate that it will be harmless to the user. Some enzymes can be allergic or harmful, and they are needed to get approved from concerning authorities of their country [8]. Guidelines for regulatory and safety aspects of enzyme production are different in all countries. For instance, some countries allow the production of the recombinant enzyme and many others prohibit [27]. A product sheet and certificate of safety should be attached to the enzyme product. Information regarding specific activity, color, solubility, swell factor (in the case of solids), and water content should be provided on the label and good manufacturing process should be followed [8].

3.3.2 ENZYME PRODUCTION FROM PLANTS AND ANIMALS Although majority of the enzymes are produced from microbes, many enzymes derived from plants and animals are still in use. For instance, Catalase (EC 1.11.1.6) is an enzyme used in the food industry, which is produced from liver [75]. Enzymes including tissue plasminogen activator and urokinase are being produced by animal cell culture techniques [24]. Other enzymes such as invertase and acid phosphatase are being produced from plant cells [76,77]. Papain is another plant enzyme, which is widely used in beer clarification, meat tenderization, and medical applications. [10,11,13]. Factors such as high cost, low yield, and risk of contamination affect the use of plants and animals for enzyme production. Plants and animals are favorable sources, only for the enzymes of very high value [8]. For the recovery process, the enzyme is simply extracted from the tissues or corresponding fluids. The extracted crude enzyme is subjected to the downstream processing as discussed in Section 3.1. Fed-batch culture is a good technique for maintaining the animal cell culture and can be made more efficient by modifying the culture medium [24]. Batch fermentation, fedbatch fermentation, perfusion fermentation, and continuous fermentation are various modes to

3.4 APPLICATIONS

41

maintain cell cultures. Plant-derived avidin and β-glucuronidase are commercially produced by big pharma companies at a lower cost. Multimeric proteins such as subunit vaccines, serum, and antibodies have been already produced from plants, which suggest the importance of plant enzyme production [23].

3.4 APPLICATIONS There are numerous applications of various enzymes in diverse industries such as pharmaceutical and analytical industry, food and beverage industry, polymer industry, leather industry, paper and pulp industry, textile industry, detergents, cosmetics, waste treatment, and organic synthesis industry (Table 3.1) [6]. For example, nattokinase (EC 3.4.21.62) is a fibrinolytic enzyme used for thrombosis therapy [78,79]. Moreover, various enzymes are important for diagnostic procedures such as ELISA [80]. Cholesterol oxidase (EC 1.1.3.6) is another enzyme used in the detection and conversion of cholesterol [81
83]. Many enzymes are conventionally popular in the food industry, such as amylase, lipase, xylanase, glucose oxidase, and lipoxygenase, in the baking industry [6]. Further, coagulation of milk into solid curds for cheese production has been done using rennet and around 33% of global cheese is produced by microbial rennet [6]. Furthermore, enzymes have important roles in the beverage industry. Amylase has been used for clarification of juices [6]. Another enzyme naringinase (EC 3.2.1.40) and limoninase hydrolyze the bitter components of citrus juices and improve their quality [2]. Enzymes are also being used as a dietary part of livestock. For example, phytases, proteases, glucanases, xylanases, polygalacturonases, α-amylases, and Table 3.1 Some examples of enzymes produced from microbes, plants, and animals. Enzyme

Source

Use

Lactobacillus sp. Aspergillus niger Aspergillus niger Candida antarctica Trametes hirsuta Aspergillus niger

Ripening of cheese Oxygen removal from beer Debittering of beverages Pitch control in paper industry Polymerization of bisphenol A Color clarification

Rice, turnip Tobacco Soybean Tobacco, potato

Pharmaceuticals Pharmaceuticals Recombinant antibody Subunit vaccines

Liver cells Pancreas

Antioxidant Digestive treatments

Some enzymes produced from microbes [6] Aminopeptidase Glucose oxidase Naringinase Lipase Laccase Cellulase Some enzymes produced from plants [23] α-Interferon Erythropoietin IgG Herpes simplex virus (HSV) Diabetes autoantigen Some enzymes produced from animals [8] Catalase Trypsin

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CHAPTER 3 ENZYMES AND THEIR PRODUCTION STRATEGIES

α-galactosidases are used as part of poultry feed [6,7,84,85]. Many animals are unable to digest the plant-based feed owing to the presence of cellulose and hemicellulose. Addition of xylanase and β-glucanase helps to degrade and digest starch [86]. The use of plastic and polymers has raised concerns for the environment. Few enzymes are used in the production of bioplastic and biopolymers. Biopolymers (polyphosphates, polycarbonates, polymers, etc.) have also been used in various biomedical applications [6]. These applications include tissue engineering, adhesive barriers, control drug-discover, and orthopedic devices [87,88]. Laccase (1.10.3.2) and peroxidase are also involved in biopolymer production [89]. Collectively, these enzymes lead to the production of eco-friendly plastic products. Enzymes such as amylase and lipase are used for deinking and quality enhancement in the paper industry [90,91]. Moreover, laccase is an alternative to chemicals used in the processing of pulp, which leads to the reduction of environmentally hazardous wastes [92]. Furthermore, the leather industry has witnessed the advantage of using eco-friendly enzymes over the use of chemicals for processes including dehairing, bating, soaking, curing, liming, picking, and tanning. [6,93]. Enzymes used in the leather industry include alkaline protease, amylase, lipases, and neutral proteases [6]. Moreover, hydrolase and oxidoreductase enzymes are used in the textile industry. These enzymes are useful in cotton pretreatment and finishing processes [6,93,94]. There are many enzymes used in the cosmetic industry, including the superoxide dismutase (SOD) and peroxidase in a combination for sunscreen creams [95], peroxidase in the formulation of skin creams [96], use of laccase, polyphenol oxidase, oxidase, and peroxidase in hair dying [97]. A number of enzymes have also been used for bioremediation and waste treatment. For instance, oxidoreductase detoxifies toxic compounds through oxidative coupling [98]. Laccase has an important role in bioremediation and removal of phenolic components from industrial effluents [83].

3.5 CONCLUSIONS AND PERSPECTIVES At present, enzymes have become an important part of various industries. However, the production of enzymes has been always a challenge. Enzymes are produced from plants, animals, and microbes. Microbial enzyme production is widely accepted and occupies approximately 90% of the global enzyme market. In microbial enzyme production, the localization of enzyme is a major aspect to be considered. If an enzyme is extracellular, the cost of downstream processing is reduced. However, in the case of intracellular enzymes, it becomes a costly process to purify such enzymes. The purification level also differs with the difference in usage of enzymes. There are some enzymes such as hydrolase, which do not require a high level of purification and are sold as a commodity enzyme. On the other hand, specialty enzymes such as streptokinase are required to be purified finely to be used. The purity of enzyme increases with each increasing step of the fractionation; however, the final yield of the enzyme decreases. Low yield and low specific activity are two areas of the enzyme industry, where the work is still to be done. Technical advancements are required to lower the waste produced during enzyme production and to increase the productivity. Storage and transportation of enzymes are another challenge for the industries. Expensive storage and transportation processes further make the enzymes more costly. Strain improvement in combination with enzyme production can give high yield and specific activity. The enzymes, which are

REFERENCES

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not generally possible to be produced through microorganisms are derived from plants and animals. The processing of plants and animals derived enzyme is costly as compared to microbial enzyme production. Further advancements in bioreactors, extraction and purification techniques, and formulation will lead to greater efficiency of enzyme production. In addition, media designing is required to be assessed to get a high yield of enzymes. The enzymes are continuously replacing the hazardous chemicals used in industries. Apart from industrial products, they can play a role in saving the environment as well as human health.

ACKNOWLEDGMENTS Authors are thankful to Panjab University, Chandigarh, India for providing the research facility and infrastructure. H.S. is thankful to CSIR for a senior research fellowship. H.S. is also thankful to IKGPTU, Jalandhar for Ph.D. registration.

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CHAPTER

ROBUST ENZYMES DESIGNING FOR EFFICIENT BIOCATALYSIS

4

Roberto Parra-Saldivar1, Ricardo A. Ramirez-Mendoza1, Ashutosh Sharma2, Goldie Oza3, Ricardo Zavala-Yoe4 and Hafiz M.N. Iqbal1 1

Tecnologico de Monterrey, School of Engineering and Sciences, Campus Monterrey, Monterrey, Tecnologico de Monterrey, School of Engineering and Sciences, Campus Queretaro, Queretaro, 3 Laboratorio Nacional de Micro y Nanofluidica (LABMyN), CIDETEQ, Queretaro, 4 Tecnologico de Monterrey, Campus Mexico City, Mexico City,

2

Mexico Mexico Mexico Mexico

4.1 INTRODUCTION Biocatalysis, as an emerging approach, has been exploited for several industrial and biotechnological processes. Biocatalysis refers to the use of natural catalysts, such as enzymes, as biological systems or their parts to catalyze the chemical reactions to biotransform the product of interests. Based on the requisite demand, biotransformation processes are performed either using enzymes in their pristine, isolated forms, or collectively as whole cell [1]. A diverse spectrum of multifunctional enzymes is being produced from various naturally occurring resources such as microorganisms (bacteria and fungi) and plants. Further to this, enzyme-driven biocatalysis follows the green agenda to fulfill the ever-increasing demand for effective, sustainable, and economical industrial bioprocesses. Comparative to the chemical-based catalytic agents, so-called chemo-catalysts, enzyme-based biocatalytic cues (simply called “biocatalysts”) hold three-dimensional (3D) structural conformation with numerous contact/access points to target the substrate molecules and thus allow immaculate selectivity [1 3]. Despite the features mentioned above, enzyme biocatalysis also offers an economical and environmentally friendly environment with no protection/deprotection steps involvement as compared to the chemo-catalytic techniques. In contrast to chemocatalytic techniques, enzyme-based catalysis displays three major types of selectivities: (1) regioselectivity and diastereoselectivity, (2) enantioselectivity, and (3) chemoselectivity. Under regioselectivity and diastereoselectivity terms, owing to the unique complex 3D structural features, enzymes are capable of distinguishing numerous functional groups situated in different regions of the substrate molecule and facilitate the reaction efficiently. Almost all enzymes contain L-amino acid residue in their structure and are considered as chiral catalysts under enantioselectivity terms. This further explains that enzymes as chiral catalysts can easily recognize any type of chirality present in the substrate molecule, subject to the formation of the enzyme substrate complex. In chemoselectivity, enzyme-assisted catalyzed reactions tend to be green in nature “cleaner” and most of the impurities generated through side reactions can largely be omitted. As a result, no protection and deprotection steps are essentially required in enzyme-assisted catalyzed reactions. Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00004-1 © 2020 Elsevier B.V. All rights reserved.

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Such anticipated characteristics fulfill the highly requisite demands of sustainable development of robust biocatalysts following green chemistry agenda [4]. Notwithstanding the striking industrial potentialities of native enzymes with unnatural substrates, biocatalysis engineering is highly requisite to fulfill the large-scale implementation demand. In this context, the whole enzyme-based catalysis comprehends numerous parameters including reaction microenvironment, substrate concentration, substrate enzyme ratio, reaction pH, operating temperature, and so on. Herein, this chapter summarizes recent advances related to the protein engineering approaches to design robust biocatalytic cues to catalyze requisite industrial process efficiently. A particular focus has been given to many new and advanced engineering approaches including enzyme immobilization using nanomaterials, microenvironment engineering via substrate engineering, structure-assisted protein tailoring, and advanced computational modeling are discussed with suitable examples to engineer multipurpose biocatalyst. The covered biocatalysis engineering approaches, herein, might offer economically viable and environmentally acceptable biotransformation of industrially relevant products of high interests, efficiently.

4.2 BIOCATALYSIS ENGINEERING—A DRIVING FORCE Notwithstanding several obvious advantages of biocatalyst-assisted reactions over chemo-assisted reactions, there have been fewer applications at large scale, whereas the use of biocatalysis is significantly increased in the past 20 years [5,6]. The rationale behind this negligence was the noncapability to distinguish, accesses, attain, analyze, and optimize biocatalyst with highly requisite attributes in the early days of the industrial revolution. With key scientific advances, protein engineering has now been recognized as the most significant factor that can facilitate the deployment of enzyme-based biocatalysis at large-scale industrial biotransformations [7]. Three distinct waves/ phases of biocatalysis have been witnessed in the last century, and now biocatalysis engineering is entering the fourth wave as a driving force for numerous industrial applications in different sectors. All these biocatalysis phases have been distinguished with a tendency to manipulate or tailor enzymes, regardless of their source and types [2,8,9]. Before the 1980s, the first biocatalysis phase was mostly based on the use of naturally occurring biocatalysts to attain the requisite chemical transformation. Later, in between the 1980s and 1990s, the second phase of biocatalysis wave appeared with more structural information and preliminary directed protein engineering. The second biocatalysis wave was quite effective in broadening the substrate preferences/selectivity of engineered biocatalysts toward the nonnatural and/or unusual products. Following the success of the second phase of biocatalysis, advanced directed evolution techniques appeared in the third wave, which then fast tracked the optimization of biocatalyst [10,11]. Since then, numerous enzymes have been subjected to the fast-growing molecular techniques and genetic engineering approaches accompanied by effective screening methods. The methods mentioned above have significantly improved or induced the industrially relevant catalytic features of enzymes at an accelerating rate. Nevertheless, the adequate timeframe to optimally design a highly efficient biocatalyst is the supreme obstacle to applying biocatalysis in industrial processes. In summary, the field of biocatalysis engineering has been revolutionized with new and/or improved approaches, and each of them is now enabled to offer numerous benefits that were huge challenges in early years of enzyme-based biocatalytic transformations.

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4.3 RECENT ADVANCES IN PROTEIN ENGINEERING Despite the huge potential of numerous naturally occurring biocatalysis for the extraordinary number of industrial and biotechnological applications, several potent enzymes in their native form do not display satisfactory catalytic performance related to the activity, stability, and selectivity. However, some of the abovementioned limitations can be resolved by exploiting new and advanced engineering approaches including enzyme immobilization using nanomaterials, microenvironment engineering via substrate engineering, structure-guided protein engineering, and advanced computational modeling [1,12 16].

4.3.1 ENZYME IMMOBILIZATION—A DRIVE TOWARD OPTIMUM PERFORMANCE Owing to the water solubility, native enzymes are costly and not recoverable from the aqueous medium. Thus, free enzyme forms can be employed for single use and on a throwing-away basis. This shortcoming of native enzymes majorly hinders its industrial application. However, with key scientific advances, the limitation above of free enzyme forms has been tackled by immobilization. This interesting approach have made it possible to engineer enzymes in the form of heterogeneous biocatalyst with added features such as facile recovery, high turnover, high catalytic yield, overall stability, multiple reusability, prolonged shelf-life, and overall cost-effective ratio [1,17]. Moreover, it does not only simplify the reaction procedure but also facilitate the bioproduct recovery in high titers with/without minimum environmental issues [18 27]. Additionally, enzyme immobilization following robust protein engineering approaches as mentioned earlier allows the engineered biocatalyst to be used at a broader spectrum for wide-ranging solvents [26,28]. From the enzyme immobilization viewpoint, numerous materials, regardless of source and type, have been trialed/deployed in diverse geometries by using various strategies and reviewed elsewhere [17,22 25,29 32]. More specifically, various enzyme immobilization approaches, including basic methods, are shown in Fig. 4.1 [31]. Nevertheless, each of them has its own merits and demerits, which has been exemplified comparatively in Table 4.1 [31]. To date, many functionalized materials-based cues at nanolevel have been engineered and utilized as novel support matrices to develop highly efficient biocatalysts. For instance, among different materials, various geometries of nanofibers, nanoparticles (both magnetic and nonmagnetic), hybrid nanoflowers, metal organic frameworks (MOFs), carbon-based nanotubes, mesoporous and nanoporous carriers, thin films, nanocomposites, nanosheets, nanocapsules, and so on have been the focus of current research in nanobiocatalysis. Additionally, regardless of the geometry of the developed nanocarriers with diverse structural and functional attributes, the enzyme immobilization or encapsulation using those robust nanocarriers can also potentially augment or secure the enzymes catalytic potentialities for numerous industrial applications. The functional attributes of engineered material-based nanomatrices, such as available functional units, for example, hydroxyl ( OH), carboxylic ( COOH), amine-containing ( NH2), thiol ( SH), and epoxy groups all facilitate the linking interactions between the enzymes and material-based nanomatrices. Additionally, effective immobilization also improves or induce several characteristics features such as hyperactivity and stability, insolubility during reaction environment, surface area, ease in recovery, reusability, high surface area-to-volume ratio, substrate affinity, maximal enzyme loading, high turnover, inducing

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FIGURE 4.1 Basic methods and submethods of enzyme immobilization. Reprinted from M. Bilal, M. Asgher, R. Parra-Saldivar, H. Hu, W. Wang, X. Zhang, et al., Immobilized ligninolytic enzymes: an innovative and environmental responsive technology to tackle dye-based industrial pollutants—a review, Sci. Total Environ. 576 (2017) 646 659, with permission from Elsevier, Copyright (2016) Elsevier B.V.

product yield, and other catalytic attributes [17,22,33]. Despite the advantages, as mentioned earlier, of nanolevel materials, the left-behind shortcomings and limits should also be considered with care before implying for immobilization purposes. In this context, various technical limitations related to the nanomaterials along with potential strategies to functionalize to overcome those limitations of nanomaterials and impart new functional entities with specific examples are schematically shown in Fig. 4.2 [1]. Very recently, Bilal and coworkers [3] comprehensively compiled various immobilization strategies covering (1) physical adsorption onto a supporting material, (2) entrapment of enzyme in a polymer network, such as alginate, collagen, agarose, gelatin, and polyacrylamide, (3) covalent attachment via multipoint enzyme immobilization, and (4) cross-linked enzyme aggregates (CLEAs) and cross-linked enzyme crystals (CLECs) covering combined CLEAs (combi-CLEAs), multipurpose CLEAs (multi-CLEAs), magnetic CLEAs (mCLEAs), macromolecular CLEAs (M-CLEAs), imprinted CLEAs (iCLEAs), porous CLEAs (p-CLEAs), protein-coated

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Table 4.1 Comparative evaluation of merits and demerits of various immobilization types. Immobilization type Adsorption

Merits

Demerits

ü ü ü ü ü

3 Lower efficacy level 3 Desorption of enzymes from the carrier

ü Covalent bonding

ü ü ü ü ü ü

Entrapment

ü ü ü ü ü ü ü ü ü

Cross-linking

Encapsulation

Easy to carry out No reagents are required No pore diffusion limitation Minimum activation steps involved Comparatively cheap method of immobilization Less disruptive to enzyme than chemical methods Wider applicability Comparatively simple method No leakage or desorption problem A variety of support/carrier available Strong linkage of enzyme to the support Multifunctional groups availability from the support/carrier Mild conditions are required Easy to practice at small scale Fast method of immobilization Can be used for sensing application Cheap (low-cost matrices available) Less chance of conformational changes No matrix or support involved Comparatively simple method Widely used in industrial applications

ü Cost-effective method ü Enzymes are stable for long time ü No extraction/purification steps are required ü “One-pot” immobilization of multiple enzymes ü Native conformation of enzyme is best maintained ü Cell organelles, e.g., mitochondria can be immobilized

3 Competitive inhibition issues 3 Chemical modification of enzyme 3 Loss of functional conformation of enzyme

3 3 3 3

Leakage of enzyme Pore diffusion limitation Chance of microbial contamination Lower level of industrial implementation

3 Polyfunctional reagents are required, e.g., glutaraldehyde 3 Denaturation or structural modification by cross-linker 3 Less concentration of enzymes 3 Generation of unwanted products 3 Modification of end products by other enzymes

Reprinted from M. Bilal, M. Asgher, R. Parra-Saldivar, H. Hu, W. Wang, X. Zhang, et al., Immobilized ligninolytic enzymes: an innovative and environmental responsive technology to tackle dye-based industrial pollutants—a review, Sci. Total Environ. 576 (2017) 646 659, with permission from Elsevier, Copyright (2016) Elsevier B.V.

microcrystals (PCMCs), combi-protein-coated microcrystals (Combi-PCMCs), and cross-linked protein-coated microcrystals (CL-PCMC), and thus are not the focus of this work. Owing to the unique structural and functional diversity, MOFs are highly tunable materials and offer characteristics such as multifunctional and adjustable topography, controlled affinity (both hydrophobicity and hydrophilicity), high surface area, tunable crystallinity, porosity, electronic

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FIGURE 4.2 Technical limitations of nanomaterials and potential strategies to overcome with specific examples. Reprinted from M. Bilal, H.M. Iqbal, Chemical, physical, and biological coordination: an interplay between materials and enzymes as potential platforms for immobilization, Coord. Chem. Rev. 388 (2019) 1 23, with permission from Elsevier, Copyright (2019) Elsevier B.V.

transduction, and ordered structure [16,34 37]. Based on all these features that are highly requisite for industrial exploitation, MOFs and MOFs-assisted support matrices/carriers have gained increasing research interest for enzyme immobilization purposes. Various coupling reactions can be used to immobilize enzymes on the MOFs surface, as support matrices, either by physical adsorption or covalent attachment. To further strengthen the enzyme-support linkage, multipoint coupling (covalent) is postulated as a robust strategy to biomolecules/enzymes linked conjugate. The development of multiple linkages between the enzyme molecules and carrier matrices offer notable features, such as moderates conformational flexibility, leaching issues, and fix the undesirable enzyme denaturation and/or unfolding [38], that limits the industrial exploitation. The abundant functionally active groups on the MOF surfaces, such as amino, carboxyl, and hydroxyl groups can be coupled with available reactive groups of the enzyme. Using this functional group phenomenon, Deng et al. [39] displayed that MOFs can incorporate many different functionalities on linking groups in a way that mixes the linker, rather than forming separate domains. Around 18 multivariate MOF-type structures that contain up to eight distinct functionalities in one phase were designed using 1,4-benzenedicarboxylate and its related derivatives, that is, NH2, Br, (Cl)2, NO2, (CH3)2, C4H4, (OC3H5)2, and (OC7H7)2. Likewise, Cao et al. [40] developed nano-/microscale UiO66-NH2 MOF materials with a uniform size of about 350 400 nm. As developed UiO-66-NH2 MOF was then used to immobilize soybean epoxide hydrolase on the outer surface of UiO-66-NH2

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MOF. The resulting immobilized enzyme-MOF conjugate was designated as SEH@UiO-66-NH2 and used for efficient biosynthesis of enantiopure (R)-1,2-octanediol in deep eutectic solvents, with a yield of around 41.4% and a product i.e., value of 81.2%. Moreover, the characterization profile of SEH@UiO-66-NH2 showed that nano-/microscale UiO-66-NH2 MOF materials are promising for immobilization was high enzyme loading, induced substrate affinity, and stability up to 4 weeks at 4 C. Very recently, Rodriguez-Abetxuko et al. [41] presented a protocol for the synthesis of catalytically responsive hybrid biomaterials and designated as “metal organic enzyme aggregates (MOEAs).” Among other tunable features, the size and morphology of MOEAs can be tailored from small individual particles to macroscopic aggregates. As developed MOEAs were found stable in water and disassemble in the presence of a complexing agent. Also, the catalytic profile of MOEAs showed high transformation rates, significant protein loads, and great thermostability, which revealed MOEAs as an excellent carrierless immobilization system.

4.3.2 SUBSTRATE ENGINEERING—A TOOLKIT TO HARNESS THE ENZYMATIC PROMISCUITY Substrate engineering, which is also termed as substrate promiscuity, is a potential alternative and/or a novel toolkit to expand the substrate specificity to harness the enzymatic promiscuity for efficient catalysis. Fig. 4.3 schematically illustrates three different categories of substrate engineering, that is, switching, expanding, and narrowing the substrate specificity of the enzyme [2]. The attachment of detachable functional entities to an alternate glycosyltransferase substrate encourages a productive binding mode, and thus, facilitate rational control of substrate specificity and regioselectivity of wildtype enzymes [8,42]. Effective promiscuity offers tailoring substrate structure that results in both the optimization and improvement of existing biotransformations using engineered enzymes. Earlier studies also report that during substrate engineering, modifications and/or alterations are required in both, that is, enzyme and substrate to broaden the substrate specificity of enzymes [42,43]. The substrate engineering approach has been extensively deployed to induce the catalytic efficacy of numerous industrial reactions to biotransform the products of interests. This additionally can also lead to the invention of entirely differential reaction types with industrially requisite features. Following substrate engineering, the enzyme features including catalytic turnover, reaction yield, enantioselectivity, conversion efficiencies, and overall process efficiency and steadiness were markedly enhanced using vinyl esters of chiral carboxylic acids than routinely used ethyl esters [44,45]. Furthermore, substrate engineering approach has been found very useful in the biohydroxylation of nonactivated carbon atoms. In addition, it also considered as simple means to influence substrate acceptance and the regioselectivity and stereoselectivity of this transformation. Therefore, this versatile substrate engineering approach has been employed for the hydroxylation of a variety of compounds including cycloalkane carboxylic acids, ketones, amines, amides, and alcohols [46]. Recently, Li et al. [47] discovered a novel ketoreductase, named KmCR2, with a broad substrate spectrum on bioreduction of sterically bulky diaryl- and aryl(heteroaryl)methanones. Two enantiomers, namely diaryl- and aryl(heteroaryl)methanols were used to prepare their stereoselective forms using strategically engineered substrates with a traceless directing group, that is, bromo group. Moreover, they also reported that the combined use of substrate engineering and protein engineering could be a useful strategy to improve the stereoselectivity or switching the stereo-preference of

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FIGURE 4.3 Substrate engineering approaches and manipulation of substrate specificity of the enzyme with suitable schematic examples. E, enzyme; S, substrate; ES, enzyme substrate complex; E , engineered enzyme; P, product. Reprinted from M. Bilal, H.M. Iqbal, Tailoring multipurpose biocatalysts via protein engineering approaches: a review, Catal. Lett. 149 (8) (2019) 2204 2217, with permission from Springer Nature, Copyright (2019) Springer Science Business Media, LLC, part of Springer Nature.

enzymatic processes [47]. In an earlier study, Keina¨nen et al. [48] reported the controlling of the regioselectivity and stereospecificity of polyamine oxidases with the use of amine-attached guide molecules as conformational modulators. Therein, an inverse approach was presented, which focuses on the manipulation of the enzyme substrate rather than the enzyme. Aharoni et al. [49] described that the evolved enzymes exhibited greater promiscuity because the evolution of higher activity toward poor substrates did not impair the parental catalytic activity.

4.3.3 STRUCTURE-ASSISTED PROTEIN ENGINEERING Enzyme-based biocatalysts can also be engineered via modeling by means of structure-assisted protein tailoring or protein-based libraries. The enzymes screening based on a directed evolution can

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potentially be substituted with structure-guided combinatorial designs. Such structure-guided tools further envisioned and enriched the functional attributes by minimizing the required number of optimal variants that need to be screened for requisite features [2]. Based on literature evidence, such structure-guided techniques are not restricted to single enzyme-based catalyzed reactions but also paved the way for engineering multienzyme cascade reaction pathways [9]. Numerous structural and functional identification-based techniques have been or being used to improve the existing enzyme functionalities or impart new attributes to meet the large-scale production and market value (Fig. 4.4) [9]. The structure-guided aid was used to engineer oligonucleotides to impart or introduce modifications at different functional sites that can significantly impact the substrate recognition or catalytic activity. Also, the oligonucleotides are generally engineered in a way that the target regions are usually encoded all or at least a subset of potential amino acids [50]. Earlier, Bell et al. [51] improved the affinity and activity of CYP101D2, which is a cytochrome P450 monooxygenase

FIGURE 4.4 Schematic outline of state-of-the-art strategies to improve enzyme functionalities and/or reengineering enzyme of interest to meet large-scale production and market value. Reprinted from M. Bilal, J. Cui, H.M. Iqbal, Tailoring enzyme microenvironment: state-of-the-art strategy to fulfill the quest for efficient bio-catalysis, Int. J. Biol. Macromol. 130 (2019) 186 196, with permission from Elsevier, Copyright (2019) Elsevier B.V.

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(P450cam) for hydrophobic substrates. They also fine-tuned the site selectivity of cytochrome P450cam by redesigning the crystal structure of the enzyme.

4.3.4 ADVANCED COMPUTATIONAL MODELING The implementation of advanced computational modeling along with machine learning strategies offer innovative substitutes to engineer biocatalysts with requisite catalysis functionalities [52]. However, in the field of advanced computational modeling for protein design, a precise geometry development, fine-tuned structural features, and dynamics of functional proteins are major unsolved challenges. This is because the structural and functional sites of protein exhibit featured folds and structured loops. Unlike idealized protein folds, the structured loops have been found difficult to design via computational-based modeling. Despite these challenges, few examples of successful loop design have been reported. The computational engineering of new protein structures is advancing rapidly which additionally assist to improve the requisite catalytic attributes of biocatalysts, for example, activity of enzymes, selectivity of enzymes, stability of enzymes, and substrate specificity or preference for broader industrial applications. Prior to the experimental implementation and detailed characterization of test variants, molecular dynamics-based simulations are made to correctly identify the target regions for in silico estimation of resultant trait features. In an earlier study by Gordon et al. [53], the use of chemical principles to identify a naturally occurring acid-active peptidase and succeeding use of computational tools have been reported to reengineer the specificity of acid-active peptidase toward immunogenic elements found in gluten. In this study, Gordon et al. [53] used the Rosetta Software Suite and the applied computational modeling results in 261 variants with one to seven amino acid replacements. The kinetic constants result profiles revealed that the engineered enzyme exhibits 116-fold superior proteolytic activity for a model gluten tetrapeptide than the native template enzyme. Moreover, as engineered enzyme also showed over 800-fold switch in the substrate specificity toward immunogenic portions of gluten peptides. In another study, Khoury et al. [54] reported the computational design of Candida boidinii (C. boidinii) xylose reductase for altered cofactor specificity. A computationally driven enzyme redesign workflow was introduced to alter the cofactor specificity from nicotinamide adenine dinucleotide phosphate (NADPH) to nicotinamide adenine dinucleotide (NAD) 1 hydrogen (H) (NADH). The implicit solvation models were integrated in the iterative protein redesign and optimization framework to drive the redesign of C. boidinii xylose reductase (CbXR). The obtained results revealed that out of total 8000 combinations of mutations, only 10 variants were optimally improved with computationally evaluated binding affinity for NADH by presenting mutations in the CbXR binding pocket, whereas the two variants, namely CbXR-RTT and CBXR-EQR, exhibited dual cofactor specificity for both nicotinamide cofactors.

4.3.5 PROTEIN ENGINEERING VIA DIRECTED EVOLUTION AND RATIONAL DESIGN Among numerous protein engineering approaches, the directed evolution strategy at large and rational design-based strategy such as site-directed mutagenesis (SDM) and random mutagenesis (RM), in particular, are innovative genetic engineering approaches for effective protein engineering. Moreover, the SDM approach implicated the construction of point mutations and required meticulous information related to the 3D structure and mechanism of the target enzyme. Whereas comparative to SDM, RM requires no such structural information to construct the mutants

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59

[15]. Likewise, SDM, the rational design for protein engineering also depends on the structural information and design concepts that significantly improve the specific attributes of the enzymes, such as substrate preference/specificity and stability of the enzymes. During the entire engineering process, diverse genes mutants comprising a single mutated parental gene, or any combination thereof, are often screened for requisite attributes. Following that, the resultant mutants with optimal combinations of useful mutations are identified and used for further analysis. For instance, Komeda et al. [55] enhanced the thermostability and catalytic activity of D-stereospecific amino acid amidase from Ochrobactrum anthropi SV3 by directed evolution using error-prone polymerase chain reaction (EP-PCR) together with a filter-based screening method. The simultaneous improvement of the oxidative and thermostability of N-carbamyl-D-amino acid amidohydrolase from Agrobacterium tumefaciens NRRL B11291 via directed evolution using DNA shuffling is reported [56]. Following successive rounds of DNA shuffling, Oh et al. [56] recorded an optimal mutant that had both improved thermal stability and greater resistance to hydrogen-peroxide-mediated inactivation compared with other mutants. In another study, Williams et al. [57] used directed evolution by DNA shuffling to modify the stereochemistry of carbon carbon bond synthesis catalyzed by tagatose-1,6-bisphosphate aldolase. S´anchez-Moreno et al. [58] tuned the phosphoryl donor specificity of dihydroxyacetone kinase from adenosine triphosphate (ATP) to inorganic polyphosphate using a directed evolution program (Fig. 4.5) [58]. In the first evolution cycle, the pristine enzyme was exposed to one round of EP-PCR followed directly (without selection) by a round of DNA shuffling. The theoretical studies based on molecular dynamics simulations and hybrid Quantum Mechanics/Molecular Mechanics optimizations suggest that this mutation affects the binding of the poly-P favoring an adequate position in the active center for the reaction to take place. In summary, with key scientific advances and new advanced technologies, it is now possible to engineer the biocatalysts for efficient biocatalysis by deploying directed evolution approaches.

4.4 CONCLUSIONS AND PERSPECTIVES In conclusion, with key scientific advances in the biotechnology sector at large and biocatalysis or protein engineering, in particular, the development of highly robust and multifunctional biocatalysts is continuously expanding with applied perspectives. The implementation of microenvironment engineering, along with the involvement of molecular engineering approaches and directed evolution strategies all make it possible to engineer robust biocatalysts with efficient catalysis potentialities by lifting-up or tackling many hindering limitations. With such high interests and evolving enzyme-based biocatalysts, in recent years, biocatalysis engineering is anticipated to further grow in numerous biotechnological sectors under green agenda perspectives. There is a demanding need to standardize the preparatory procedures for a proper implementation in numerous industrial bioprocesses. The above-discussed biocatalysis engineering approaches can play a futuristic role in this development following green chemistry principles. Although there is dire demand for such specialized tools to engineer robust biocatalysts because they are generally produced from soluble enzymes that are not reusable in the industrial bioprocess. Therefore, the integration of various inpractice strategies and biocatalysis/protein engineering could possibly be the most efficient approach for creating powerful, recyclable, and catalytically robust biocatalysts that fit best for the real-world biotechnological processes.

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FIGURE 4.5 Directed evolution approach applied to modify the phosphoryl donor specificity of the dihydroxyacetone kinase (DHAK) from Citrobacter freundii. ´ Reprinted from I. Sanchez-Moreno, I. Bordes, R. Castillo, J.J. Ruiz-Pernı´a, V. Moliner, E. Garcı´a-Junceda, Tuning the phosphoryl donor specificity of dihydroxyacetone kinase from ATP to inorganic polyphosphate. An insight from computational studies, Int. J. Mol. Sci. 16 (11) (2015) 27835 27849, an open-access article distributed under the Creative Commons Attribution License.

4.5 ACKNOWLEDGMENTS The authors are grateful to their representative institutional units for providing the literature access.

4.6 CONFLICT OF INTEREST The authors do not have any conflicting, competing, or financial interests.

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[21] K. Antecka, J. Zdarta, K. Siwi´nska-Stefa´nska, G. Sztuk, E. Jankowska, P. Oleskowicz-Popiel, et al., Synergistic degradation of dye wastewaters using binary or ternary oxide systems with immobilized laccase, Catalysts 8 (9) (2018) 402. [22] M. Bilal, Y. Zhao, T. Rasheed, H.M. Iqbal, Magnetic nanoparticles as versatile carriers for enzymes immobilization: a review, Int. J. Biol. Macromol. 120 (2018) 2530 2544. [23] M. Bilal, T. Rasheed, H.M. Iqbal, H. Hu, W. Wang, X. Zhang, Horseradish peroxidase immobilization by copolymerization into cross-linked polyacrylamide gel and its dye degradation and detoxification potential, Int. J. Biol. Macromol. 113 (2018) 983 990. [24] J. Zdarta, A.S. Meyer, T. Jesionowski, M. Pinelo, Developments in support materials for immobilization of oxidoreductases: a comprehensive review, Adv. Colloid Interface Sci. 258 (2018) 1 20. [25] J. Zdarta, A. Meyer, T. Jesionowski, M. Pinelo, A general overview of support materials for enzyme immobilization: characteristics, properties, practical utility, Catalysts 8 (2) (2018) 92. [26] G. Bayramoglu, B. Salih, A. Akbulut, M.Y. Arica, Biodegradation of Cibacron Blue 3GA by insolubilized laccase and identification of enzymatic byproduct using MALDI-ToF-MS: toxicity assessment studies by Daphnia magna and Chlorella vulgaris, Ecotoxicol. Environ. Saf. 170 (2019) 453 460. [27] N. Khan, Q. Husain, Continuous degradation of Direct Red 23 by calcium pectate-bound Ziziphus mauritiana peroxidase: identification of metabolites and degradation routes, Environ. Sci. Pollut. Res. 26 (4) (2019) 3517 3529. [28] M.D. Truppo, H. Strotman, G. Hughes, Development of an immobilized transaminase capable of operating in organic solvent, ChemCatChem 4 (8) (2012) 1071 1074. [29] M. Asgher, M. Shahid, S. Kamal, H.M.N. Iqbal, Recent trends and valorization of immobilization strategies and ligninolytic enzymes by industrial biotechnology, J. Mol. Catal. B: Enzyme 101 (2014) 56 66. [30] T. Jesionowski, J. Zdarta, B. Krajewska, Enzyme immobilization by adsorption: a review, Adsorption 20 (5 6) (2014) 801 821. [31] M. Bilal, M. Asgher, R. Parra-Saldivar, H. Hu, W. Wang, X. Zhang, et al., Immobilized ligninolytic enzymes: an innovative and environmental responsive technology to tackle dye-based industrial pollutants—a review, Sci. Total Environ. 576 (2017) 646 659. [32] M. Adeel, M. Bilal, T. Rasheed, A. Sharma, H.M. Iqbal, Graphene and graphene oxide: functionalization and nano-bio-catalytic system for enzyme immobilization and biotechnological perspective, Int. J. Biol. Macromol. 120 (2018) 1430 1440. [33] E.T. Hwang, M.B. Gu, Enzyme stabilization by nano/microsized hybrid materials, Eng. Life Sci. 13 (1) (2013) 49 61. [34] J. Wang, Electrochemical glucose biosensors, Chem. Rev. 108 (2) (2008) 814 825. [35] H. Furukawa, K.E. Cordova, M. O’Keeffe, O.M. Yaghi, The chemistry and applications of metalorganic frameworks, Science 341 (6149) (2013) 1230444. [36] K.Y.A. Lin, H.A. Chang, R.C. Chen, MOF-derived magnetic carbonaceous nanocomposite as a heterogeneous catalyst to activate oxone for decolorization of rhodamine B in water, Chemosphere 130 (2015) 66 72. [37] Q. Wang, X. Lian, Y. Fang, H.C. Zhou, Applications of immobilized bio-catalyst in metal-organic frameworks, Catalysts 8 (4) (2018) 166. [38] C. Mateo, J.M. Palomo, G. Fernandez-Lorente, J.M. Guisan, R. Fernandez-Lafuente, Improvement of enzyme activity, stability and selectivity via immobilization techniques, Enzyme Microb. Technol. 40 (6) (2007) 1451 1463. [39] H. Deng, C.J. Doonan, H. Furukawa, R.B. Ferreira, J. Towne, C.B. Knobler, et al., Multiple functional groups of varying ratios in metal-organic frameworks, Science 327 (5967) (2010) 846 850. [40] S.L. Cao, D.M. Yue, X.H. Li, T.J. Smith, N. Li, M.H. Zong, et al., Novel nano-/micro-biocatalyst: soybean epoxide hydrolase immobilized on UiO-66-NH2 MOF for efficient biosynthesis of enantiopure (R)1, 2-octanediol in deep eutectic solvents, ACS Sustain. Chem. Eng. 4 (6) (2016) 3586 3595.

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CHAPTER

ENZYME ENGINEERING STRATEGIES TO CONFER THERMOSTABILITY

5

Zhe Xu, Ya-Ping Xue, Shu-Ping Zou and Yu-Guo Zheng Key Laboratory of Bioorganic Synthesis of Zhejiang Province, College of Biotechnology and Bioengineering, Zhejiang University of Technology, Hangzhou, P.R. China

5.1 INTRODUCTION Enzymes, as important macromolecular biological catalysts, are attractive and competitive alternatives to chemical catalysts [1]. Thermostable enzymes are highly competitive and desirable for application in the production of fine chemicals and biofuels, synthetic biology, and protein crystallization [2,3]. However, poor thermostability of natural enzymes becomes a bottleneck that limits the development of biocatalysis. The thermostability poses a fundamental challenge for the exploitation of enzymes for practical-scale syntheses and chemical manufacturing, which often require harsh reaction conditions such as elevated temperature. The thermostable enzyme has good kinetic stability and processes stability under harsh conditions, which benefits the reusability of the biocatalyst and thus reduces the cost of the enzyme [2,4,5]. Enzymes with higher stability also have greater evolutionary potential because they can accept a wider range of beneficial mutations while still retaining native structure [6,7]. Extremophilic microorganisms are a source of high thermostable enzymes with a great variety of industrial applications. The enzymes separated from microorganisms which thrive in thermal vents or hot springs usually have the ability to retain the structure and function in high temperature [8,9]. With the advances in genome sequencing and the metagenomic field, numerous new extremozymes have been identified [10,11]. Unfortunately, compared with mesophilic enzymes, the narrower substrate spectrum and lower activity hindered the application of the extremozymes [4,5]. Due to these limitations, protein stabilization of existing mesophilic enzymes against thermal denaturation has represented a long-standing goal in enzyme design and engineering [12]. The thermostability of enzymes is often structurally predetermined. Plenty of research in crystal structures of the mesophilic and thermophilic proteins showed the important relationship between the protein structure and thermostability. The additional intramolecular/intersubunit interactions (hydrogen bonds, salt bridges, hydrophobic interaction, aromatic interactions, and disulfide bonds) and good conformational structure (well-packed hydrophobic core, more rigid structure, lower entropy of unfolding) are important stabilizing structural features for enzymes.

Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00005-3 © 2020 Elsevier B.V. All rights reserved.

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The analysis of nucleotide sequence and high-resolution, three-dimensional (3D) structures of extremostable proteins widen and deepen the understanding of the thermostability of proteins [13]. The advances in high-throughput screening method and design of “smart” mutant library facilitate the thermostability engineering of the mesophilic enzymes. In this chapter, we will introduce the protein engineering approaches used for the enzyme thermostabilization and discuss the applications of protein thermostability engineering in enzyme thermostabilization.

5.2 ENZYME ENGINEERING STRATEGIES FOR THERMOSTABILIZATION Protein engineering including directed evolution, rational/semi-rational, and de novo design is a powerful tool for modifying the enzyme structure (Fig. 5.1). Traditionally, enzyme engineering by directed evolution which does not require much knowledge about structure/sequence function relationship of protein is the strategy for protein thermostabilization. With the in-depth research of information on structure/sequence function relationship and computational predictive algorithms, more researchers advocate new strategies for designing smaller, higher quality libraries. Rational/semi-rational design and de novo design have become new pets of protein thermostability engineering.

FIGURE 5.1 Schematic representation of protein thermostability engineering.

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5.2.1 DIRECTED EVOLUTION Directed evolution, which simulates the selection process in nature, can generate variants with diversity and evolve enzymes toward certain goals. Utilizing random mutagenesis to construct a molecularly diverse library, followed by high-throughput screening strategy, directed evolution can accumulate beneficial mutations with improved characteristics in iterations of mutagenesis and screening (Fig. 5.2). Directed evolution is now well established as highly effective for protein engineering and optimization. To date, many rules for protein engineering thermostability by rational design are likely to be protein-specific, not universal, and rational design which needs to be guided by detailed structural information may still not apply to all the enzymes. Thus, directed evolution which could be applied to the proteins which are lack of research on protein structure remains the popular approach in protein thermostabilization. According to reported stabilization of α/β hydrolases

FIGURE 5.2 General directed evolution platform.

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summarized by Jones, directed evolution is the main approach to yield a dramatic increase in stability of α/β-hydrolases [14,15]. Polymerase chain reaction (epPCR), as a traditional directed evolution strategy, is one of the most efficient methods to obtain thermostable enzymes. A highly effective and efficient directed evolution strategy is to gradually accumulate single beneficial mutations. Low error-rate random mutagenesis (1 2 amino acid substitutions per gene) by epPCR can avoid the missing beneficial mutations, which may be masked by the much more frequent deleterious ones. Combining other directed evolution strategies such as iterative saturation mutagenesis (ISM), this low error-rate random mutagenesis can generate additional mutations in targeted residues generated by epPCR and identify mutations that only work in concert [16,17]. Rao’s group employed epPCR and sitedirected mutagenesis to enhance thermostability of Bacillus subtilis lipase, providing an excellent example of the application of low error-rate random mutagenesis. A three-tier screening protocol, namely Petri Plate assay based on halos screening, 96-well plate assay with p-nitrophenyl butyrate (pNPB) after exposing to heat, and the final screening tube assay for determining thermal inactivation profiles, was applied to screen thermostable variant in the mutant library. Multiple rounds of epPCR and recombination generated a thermostable variant with 12 accumulated mutations, which showed substantial enhancement in melting temperature (ΔTm 5 122.2 C) (Fig. 5.3) [18 20]. Although traditional epPCR-based random mutagenesis is one of the soundest and easiest to implement directed evolution method, there are also some limitations in constructing highly diverse mutant libraries, such as commonly low mutagenic frequency, the redundancy of the genetic code, and mutagenic “hot spots” (mostly conservative) caused by propensity of polymerases [21,22]. The sequence saturation mutagenesis (SeSaM) method is a state-of-the-art directed evolution approach which could overcome these shortcomings of epPCR [23]. A four-step chemoenzymatic process, including the incorporation of phosphorothioate nucleotides, chemical cleavage of phosphorothioate bonds, and introduction of universal bases and their replacement by standard nucleotides inserting point mutations, was applied to generate diverse mutations. Shivange et al. took advantage of SeSaM to evolve Yersinia mollaretii phytase (Ymphytase). A 96-well plate assay was implemented to compare activities of phytase variants before and after incubation at 60 C based on fluorescence. After two rounds of SeSaM protocol, nine mutations were identified in the thermostable Ymphytase variants. Iteratively combining the site-saturation mutagenesis and site-directed mutagenesis, the key beneficial mutations were identified, and the resultant variants M6 (T77K/G187S/Q154H/K289Q) and M3 (T77K/G187S/Q154H) displayed 3.0 C and 2.5 C enhancement in Tm. The so-called iterative key-residue interrogation of the wild type with substitutions identified in directed evolution (KeySIDE) approach was proved to be a potential protein thermostabilization technology [24]. Different from the nonrecombinant directed evolution approaches such as epPCR and SeSaM, DNA shuffling which refers to the recombinant approach is an efficient method to generate hybrids or chimeric forms with unique properties. Additionally, in the DNA shuffling, new point mutagenesis will also be introduced at a relatively high rate (0.7%) [25]. The traditional DNA shuffling has been widely applied to improve the protein thermostabilization. Akbulut et al. constructed a lipase chimera L3-3 utilized DNA shuffling, whose t1/2 at 50 C was 9.2- and 8.8-fold improvement compared with the parent enzymes from Bacillus pumilus strain L5 and L21. The additional point mutagenesis G14S, A15G, and V109S (not stem from the parent lipases) in chimera L3-3 played significant roles in the rigidification of loop region [26].

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FIGURE 5.3 Thermostabilization of Bacillus subtilis lipase by multiple rounds of low error-rate epPCR and iterative saturation mutagenesis.

The ability of directed evolution to generate a diverse mutant library is its greatest strength and central weakness. Picking up desired variants in the large and diverse libraries of mutants requires much time and labor. Therefore, a robust high-throughput screening method is necessary for directed evolution. The main strategy for screening for enzyme thermostability relies on measuring residual enzyme activity after incubating at high temperature [27]. Asial et al. described a generic activityindependent biophysical screen inside the cell for thermostability screens, named dHot-CoFi, which is expected to be applied to a wide range of proteins [28]. A high-throughput thermal scanning method based on differential scanning fluorimetry was also designed by Seabrook et al. which had adequate reliability [29,30]. In addition, some indirect ultrahigh-throughput screening method, such as the absorbance-activated droplet sorting [31], fluorescence-activated droplet sorting [32], Fo¨rster resonance energy transfer, fluorescence resonance energy transfer (FRET) probe-based high-throughput

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screening for in vivo enzyme activity [33] and utilization of robotic systems [34] or microfluidics [35] are helpful to the protein thermostability engineering.

5.2.2 RATIONAL/SEMI-RATIONAL DESIGN Different from directed evolution, rational and semi-rational design focus on several amino acids based on well understanding of structure/sequence function relationship of proteins, which can dramatically reduce the size of mutant library and labor intensive for screening. Sequence-based engineering and structure-based engineering are two important tools for rational/semi-rational design to localize flexible regions in proteins and predict thermostability change brought by the introduction of mutations. Then, site-directed mutagenesis, site-saturation mutagenesis, and focus epPCR are applied to bring stabilizing mutations into these potential “hot spots” (Fig. 5.4). In recent years, a number of excellent computational tools ranging from bioinformatic analysis of primary sequences to simulations of tertiary structures have been developed and established a manipulate platform that aimed at monitoring the flexible region and predicting thermostable mutation, which promotes the application of rational/semi-rational design (Table 5.1).

5.2.2.1 Sequence-based engineering The protein primary structure to a large extent determines the 3D structure, thus affecting the protein thermostability. According to many sequence-based statistical studies, a large amount of charged residues, hydrophobic residues, and prolines are usually present in the thermophilic

FIGURE 5.4 Overview of concepts used to create thermostable enzymes by rational/semi-rational design.

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Table 5.1 Summary of several important software for monitor flexibility or predicting mutation for protein stabilization. Protein engineering approaches Sequence-based engineering Structure-based engineering

Function

Software

Website

Reference

Consensus analysis

Consensus finder B-FITTER

http://kazlab.umn.edu/

[14]

http://www.kofo.mpg.de/en/ research/biocatalysis Gromacs, Amber, NAMD, YASARA or Charmm http://flexweb.asu.edu/ http://kinari.cs.umass.edu/Site/ index.html http://cpclab.uni-duesseldorf. de/cna/

[36]

Prediction of flexible sites

MD simulations FIRST KINARI

Prediction of stabilizing mutations

De novo design

Design of disulfide bonds

Constraint network analysis I-mutant 3.0

BridgeD RosettaVIP

http://gpcr2.biocomp.unibo.it/ cgi/predictors/I-Mutant3.0/IMutant3.0.cgi foldx.crg.es www.rosettacommons.org http://dezyme.com http://cptweb.cpt.wayne.edu/ DbD2/ http://caps.ncbs.res.in/iws/ modip.html http://hazeslab.med.ualberta.ca/ forms/ssbond.html http://biodev.cea.fr/bridged/ www.rosettacommons.org

FireProt framework

https://loschmidt.chemi.muni. cz/fireprot/

FoldX Rosetta_ddg PoPMuSiC Disulfide by design MODIP SSBOND

Comprehensive computational approaches

Redesign of hydrophobic core Prediction of stabilizing mutations based on multiple algorithms

[37] [38] [39] [40]

[41]

[42] [43] [44] [45] [46] [47] [48] [49] [50]

FIRST, floppy inclusion and rigid substructure topography; MD simulations, molecular dynamic simulations; MODIP, modeling of disulfide bonds in proteins.

proteins [51]. Analyzing the sequence information of homologous proteins, discovering the correlation between the protein sequence and their thermostability, identifying potential “hot spots” that affect thermostability has been one of the most commonly used strategies in protein thermostability engineering. This sequence-based protein engineering strategy has the following obvious characteristic and superiority: (1) sequence-based engineering does not require the construction of a large library of clones and (2) compared to structure-based engineering, the sequence-based engineering

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strategy requires less knowledge about protein structure. The sequence information of proteins (especially thermostable proteins) available in public databases is a treasure trove, which offers abundant valuable propositional information and may guild the protein engineering. The sequencebased engineering based on the analysis of homologous sequences has been proved to be an attractive protein engineering approach for proteins that have no available structure [52 54].

5.2.2.1.1 Comparing sequences with higher thermostability Multiple sequence alignment (MSA) is a common-used and easy-to-use sequence-based engineering method for rational design. At first, find homologous sequences for the target sequence in the database using search tools such as basic local alignment search tool (BLAST). The selected protein sequences should consist mainly of characterized thermophilic and hyperthermophilic sequences with established thermostability. More thermophilic and hyperthermophilic sequences with high identity used in MSA are more favorable to identify the key residues. Then, sequence alignment is implemented to find key changes functional to enhance the thermostability. During the analysis of sequence differences, the hydrophobic (or hydrophilic) residues in target sequence for which their counterparts in hyperthermophiles are hydrophilic (or hydrophobic), and the position occupied by flexible residues such as glycine should receive more attention [55,56]. This method has been applied successfully into the thermostabilization of proteins including lipase [56], L-asparaginase [57], amylase [58], lyase, xylanase [59], and β-1,3-1,4-glucanase [60]. For example, MSA was used to improve the thermostabilization of Candida rugosa Lipase1 (Lip1). The flexible glycine at residue 414 in Lip1 is distinguished from other thermostable lipases. Four mutants (G414A, G414M, G414H, and G414W) with obvious enhancement in thermostability were obtained by site-saturation mutagenesis. Among them, G414W displayed the most remarkable enhancement in thermostability, which showed a 6.5-fold longer t1/2 at 60 C, and a 14 C higher T5015 [56].

5.2.2.1.2 Consensus analysis Key residues analysis using sequence alignment with thermophilic homologs suffers from the number of characterized thermophilic/hyperthermophilic protein sequences. In long-term nature selection, the destabilizing random mutations in proteins have a high probability to be purged by nature. Thus, the residues appearing at a specific position most frequently among homologous structures tend to be more conducive to the stability than other residues at the same position. Based on this assumption, consensus analysis (CA) was developed for designing thermostable and functional enzyme variant according to the comparison of a set of homologous sequences in a family, in which thermostable/hyperthermostable homologs are not necessary [61 63]. CA involves the following four steps: (1) acquisition of homologous sequences, (2) MSA, (3) calculation of conservation and generation of consensus sequence which reflects the most frequent residues at each position, and (4) substitution of nonconsensus residue in target protein with consensus one. Nowadays, the CA approach has been widely successful in improving the stabilities of various proteins [64 67]. The leucine-rich repeat sequence in mouse leucine-rich repeat transmembrane neuronal 2 (LRRTM2) was redesigned using CA. The resultant consensus protein represented a very respectable increase of Tm of 32 C [68]. However, in consensus sequence, there still exists a large proportion of neutral or destabilizing residues. About 10% of conserved residues are stability neutral and about 40% are destabilizing, which lead to challenges during implementation.

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5.2.2.1.3 Ancestral sequence reconstruction Ancestral sequence reconstruction (ASR) is a powerful tool to infer the sequence of ancient proteins based on extant proteins and understand the ways in which proteins evolve in response to changing environments. With a history of around 50 years, ASR has grown to be one of the most popular protein engineering technologies [69,70]. Many lines of evidences showed that due to the cooling trend of the earth, the thermal stabilities of the proteins declined during evolutionary [71 73]. For example, the Tm value of thioredoxin declined at a rate of 6 C per billion years during the course of evolution [74]. Therefore, high thermostable variants can be achieved by replacing modern sequences with ancestral residues. In recent years, due to the explosion in genome sequence data and advances in bioinformatics tools, ASR has become ever more feasible. The process of ASR from extant homologs mainly involves the following stages: (1) collection of homologous sequences using NCBI, UniProt, and so on; (2) MSA; (3) construction of phylogenetic tree; and (4) reconstruction of ancestral sequence. The computer software packages such as phylogenetic analysis using parsimony (PAUP), phylogenetic analysis by maximum likelihood (PAML), MrBayes, and FastML [69,75] are available for the phylogenetic analysis and ancestral sequence inference. The obtained ancestral sequence can be implemented in protein thermostability engineering (ancestral mutation method). The simplest approach is the introduction of ancestral residues into a modern protein sequence. To enhance the thermostability of Bacillus circulans β-amylase, an ancestral sequence was reconstructed based on 22 amylase sequences from higher plant and bacterial. After introduction of ancestral residues, the variant contained seven ancestral mutations, which resulted in a 3.2 C increase in Tm [76]. Analyzing phenotypic and sequence differences between two subfamilies in a phylogenetic tree to identify the potential key residues responding to the change in thermostability, which is called the reconstructing evolutionary adaptive path (REAP) approach, is also an efficient way to find mutations for protein thermostabilization [77,78]. In addition, combining ASR and random mutagenesis to generate ancestral libraries containing random ancestral mutation is beneficial to the construction of “smart libraries” for directed evolution and discovery of more improved mutants [79].

5.2.2.2 Structure-based engineering As with sequence-based engineering, structure-based engineering is another important method to propose mutations for protein thermostabilization. Structure-based engineering takes advantage of protein structures and molecular modeling data to analyze molecular interactions and predict specific substitutions. Recently, rigidify flexible site (RFS) approach becomes the research hotspot. Mechanically weak regions of proteins are the most likely unfolding initiators at elevated temperatures. In this strategy, the flexible residues or region which may be the mechanically weak point in protein structure will be pinpointed based on the structural data. Then, site-directed mutagenesis, site-saturation mutagenesis, or gene fusion is employed to introduce appropriate mutations to strengthen the flexible structure. Computational protein design has been an efficient tool to find the flexible region in protein, which automates and improves the efficiency of protein structure analysis that facilitates the identification of flexible region and design of mutations. In this section, we will introduce some important approaches employed in structure-based engineering for improving protein thermostabilization.

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5.2.2.2.1 Beta-factor analysis The protein beta-factor (B-factor) or temperature factor (Debye Waller factor), which reflects the diffusion of atomic electron density, was used to evaluate the flexibility of the protein structure. In the comparison of various enzymes, the B-factor value decreased when the temperature optimum and thermostability increased, which implied that thermostability is increased with the rigidification of protein structures [80,81]. Different algorithms, such as normal mode analysis [82], the elastic network model [83], the Gaussian network model [84], the flexibility and rigidity index methods [85], and multiscale weighted colored graph model, have been developed for the accurate prediction of protein B-factor. Nowadays, the protein B-factor can be obtained by the B-factor DataBank (BDB) database (http://www.cmbi.ru.nl/bdb/) [86] or using computational tools such as B-FITTER, YASARA. B-factor profiles have been proven effective in analyzing the flexible sites of proteins. In 2007 Reetz et al. showed a standard protocol of B-factor profiles for protein thermostability [36]. The X-ray structure of target protein B. subtilis lipase A was analyzed by B-FITTER and 10 amino acids showing the highest average B-factor values were chosen as sites for randomization using standard saturation mutagenesis. After the ISM, the best variants X and XI were found with Tm increased from 48 C to 89 C and 93 C. In addition, the B-factor concept can easily cooperate with other rational design tools, which better guides the protein thermostabilization. Land et al. present a rational design strategy based on proline substitutions in flexible regions of the Chromobacterium violaceum amine transaminase identified by B-factors using YASAR. The best mutant displayed a 2.7-fold enhancement in t1/2 values at 60 C [87]. Incidentally, there are several restrictions or limitations with the B-factor concept. The accurate prediction of protein B-factor requires higher protein structure quality. The B-factor values may vary significantly between proteins depending on the structural refinement and the crystal quality. Although some computational methods such as support vector regression [88] and PROFbval [89] could predict B-factor value from the amino acid sequence, a high-resolution X-ray structure of monomeric protein without nonresolved atoms and external molecule bounds is a favor for the prediction of B-factor value [36].

5.2.2.2.2 Molecular dynamic simulations Molecular dynamic (MD) simulation is a powerful method to investigate the thermostability of proteins by characterizing the dynamic properties of protein structures at atomic-level resolution during the protein unfolding process. Modern computers allow simulation of a long-time scale of unfolding processes (several nanoseconds) at highly elevated temperature, which make MD simulations an attractive means to identify the weak points in protein structure. The root mean square fluctuation (RMSF) calculated by MD simulations at different temperature (300K 600 K) can represent the overall flexibility of the proteins. The higher RMSF values indicate higher mobility of amino acids, which may be the “short-board” regions that can serve as a target for thermostability enhancement. The B-factor value can also be calculated using RMSF (RMSF2 5 3B/8π2, where B is the B-factor value). Compared with the experimental B-factor predicted by B-FITTER and crystallographic data, the RMSF or B-factor from the simulations sometimes reflect the flexibility of some regions of proteins more accurately due to the crystal lattice packing effects [90]. Additionally, MD simulations can also be utilized to analyze RMSF, root mean square deviation (RMSD), secondary structure, hydrogen bonds, salt bridges, radius of gyration, and solvent accessible surface area at

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different temperatures, which contribute to the understanding of thermostabilizing factors and mechanisms [91]. The typical protocols for MD simulations for the thermostability analysis usually consist of three steps: (1) preparation of 3D structure of the target protein from database or homology modeling, (2) addition of explicit water molecules and minimization of energy, and (3) equilibration at room temperature and simulation at elevated temperatures. For the prediction of protein flexibility, a length of simulations ranging from 2 to 100 ns is long enough and the time step is set on the picosecond or femtosecond time scale [92]. Various MD simulation packages, such as Amber, Gromacs, and nanoscale molecular dynamics (NAMD), have been widely used in identifying flexible regions and studying the effect of mutations on thermostability [93 95]. Modarres et al. used NAMD simulation package to analyze the structure of LigTh1519 DNA ligase from the thermophilic archaeon Thermococcus sp. 1519 at different temperatures (280K 500 K). The thermosensitive region L300-E350 was detected based on the RMSF analysis. Combining the salt bridge analysis by visual molecular dynamics (VMD) and sequence analysis by MSA, four mutations (A287K, G304D, S364I, and A387K) were introduced to extend salt bridge in the detected thermosensitive region, which yielded an increase in t1/2 at 94 C of 25 and 15 min, a twofold to threefold improvement [55].

5.2.2.2.3 FoldX and Rosetta_ddg FoldX is a frequently used force field algorithm and applied in the studies about protein stabilization experiments. FoldX can calculate the Gibbs free energy (ΔΔGFold) change brought by single mutations, thereby predicting beneficial single and multiple-point mutation sites for protein thermostabilization. The algorithm of FoldX was first described by Guerois et al. and has become a popular calculation tool after years of development [42]. FoldX showed higher accuracy than other algorithms in many studies [96]. The crystal structure quality has an important influence on the calculation result. In addition, compared with other popular protein engineering tools for stabilization, FoldX with a graphical interface is user friendly to the researchers who are not familiar with programming languages such as Python, Java, and R-script. In many cases, FoldX can perform a complete calculation of every single mutation (including self-mutated structures), overall protein, or a specific region in the protein. The candidate mutant lowering ΔΔGFold below a certain value will be selected for experimental verification. Huang et al. used FoldX to scan the beneficial mutants in the higher B-factor region G129-D134 in amine transaminase from Aspergillus terreus, and six potential mutations (ΔΔGFold , 0) were selected. The resultant variant T130M/E133F showed a 3.3-fold improvement in t1/2 at 40 C [97]. Buß et al. calculated ΔΔGFold for all possible single mutations in Variovorax paradoxus transaminase by FoldX. 11 most stabilizing mutations with ΔΔGFold ranging from 230.1 to 254.3 kcal/mol were selected. The Tm of the best performing mutant was increased to 59.3 C, an increase of 4.0 C in comparison with the wild-type enzyme [98]. Rosetta_ddg is a module in RosettaDesign, which is also a usual tool to predict protein stability changes due to ΔΔGFold of mutations [43]. Similar to FoldX, Rosetta_ddg can predict potential stabilizing mutation by calculating the ΔΔGFold values of all possible single substitutions. Yu et al. investigated the ΔΔGFold and stability changes for all mutations in the flexible loops of Escherichia coli transketolase and the qualitative prediction accuracy of the Rosetta_ddg reached 65.3% [99].

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In addition, FoldX and Rosetta_ddg are more commonly used as a filter to exam preselection mutant sites generated by other protein thermostability engineering approaches. The combination of FoldX, Rosetta_ddg, and other algorithms usually get more stabilizing mutation predictions. These make them important parts in some comprehensive computational approaches such as FireProt [100] and FRESCO [101].

5.2.2.2.4 Structure-guided sequence-based engineering Although traditional sequence-based engineering allows for protein thermostabilization without requiring detail structural information, these approaches have pitfalls in the prediction of accuracy and library size. Using sequence-based approaches to generate mutations at positions identified by structural approaches can make up these defects. For example, combining the consensus approach and in silico structural analysis, the structure-guided consensus approach results in low screening requirements of just about 20 40 site-directed mutations per protein [102]. V´azquez-Figueroa et al. used the structure-guided consensus approach to enhance the stability of glucose dehydrogenase. When focused on the subunit subunit interactions, it suggested a success rate of thermostability engineering from 46% to 86% [103]. Structure-guided consensus approach has been proved to be a considerable potential method for improving protein thermostability. The structure-based MSA and ASR are also feasible approaches which can predict stabilizing mutations in association with the protein thermostability more efficiently. Utilizing sequence alignment and structure comparison, 16 candidate amino acids, which correlates to the loss of 17 hydrogen bonds in the Bacillus lipase when compared with its thermostable homolog Bacillus stearothermophilus lipase P1, were identified. The mutation P247S exhibited a 60-folds increase in thermostability at 60 C [104]. Nguyen et al. focused on adenylate kinase (Adk) and reconstructed a phylogenetic tree for the Adks from Firmicutes. After analyzing the crystal structures of several important nodes in the tree, it was found that several unique salt bridges sequentially disappeared during evolution toward colder environments and then reappear in species that subsequently adapt to hot niches. According to the REAP approach, the residues, which were responsible for the salt bridge in an ancestral Adk ANC1, were mutated into the corresponding positions found in Adk from B. subtilis, resulting in a dramatic effect on stability with a c.20 C increase in Tm [105].

5.2.3 DE NOVO DESIGN De novo design is a promising tool for designing proteins with desired properties based on the identification of protein sequences [106]. The folding strategy adopted by proteins is to keep the conformation which lies at the free energy minimum. De novo protein design can seek sequences that will have a free energy minimum in the fold of proteins. Compared with rational design approaches, de novo design needs more sufficient knowledge of the structure function relationship. It is because of the stringent requirements for structure function relationship, only a limited number of de novo design tools are available for protein thermostabilization. The hydrophobic residues in the core of proteins play important roles in protein folding and preserving the thermostability. Many computational approaches for predicting flexible sites like BFITTER and MD simulations only identify hot spots which are usually located on the surface of a protein. The de novo design tools RosettaVIP can redesign a well-packed hydrophobic core of protein, which can identify mutations to improve the quality of core packing, resulting in a more

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thermostable protein [107]. Borgo and Havranek first used the RosettaVIP protocol to automatically design stabilizing mutations for methionine aminopeptidase [49]. The five mutations (V24I, C45L, V207I, A152I, and F156L) led to drastic reduction in buried void volume, which indicated the improved packing of the hydrophobic core. The fivefold mutant exhibits an increase in Tm of 17.6 C. De novo design also has been explored to introduce intramolecular or intersubunit interactions for improving protein thermostability. Disulfide bond is an important covalent bond, which can ensure the proper folding of proteins, increase protein stability, and reduce self-aggregation [108,109]. Disulfide engineering is a powerful strategy for protein thermostabilization. The rapid growth of protein structures in the Protein Data Bank (PDB) spurred the development of computational methods to the de novo design of disulfide bonds. For example, with the assistance of software Disulfide by Design (DdD), two de novo disulfide bonds were designed and engineered in formate dehydrogenase from Candida boidinii, which eliminated the easily oxidized free cysteine, resulting in a 6.7-fold increase in t1/2 at 60 C and a remarkable improvement in resistance to Cu21 [110]. The combination of de novo design of disulfide bonds and in silico fast screening gives greater precision to the introduction of disulfide bonds. Due to the high cooperativity and sensitivity, hydrogen bond networks were difficult to be designed by many existing computational methods. Recently, a new Monte Carlo-based sampling protocol in program Rosetta (MC HBNet) was developed to search for mutations which can form self-contained hydrogen bond networks [111]. It is believed that the improvement of algorithm prediction technology would help the precise de novo design of interaction in the near further.

5.2.4 COMPREHENSIVE COMPUTATIONAL APPROACHES In the last decades, there have been significant advances toward more rational design approaches. The development of computational tools has greatly reduced the experimental efforts to generate highly thermostable proteins. But all algorithms for predicting stabilization mutations still suffer from low accuracy. When using an algorithm alone such as FoldX, Rosetta_ddg, I-Mutants3.0, and PoPMuSiC, the accuracy only ranged from 0.38 to 0.85 [96]. The low accuracy of these algorithms decides that all algorithms are not able to design or predict single mutation events towards trustworthy one mutation protein designs. Thus, combining multiple algorithms and various identification approaches for the flexible region can further reduce the mutation library and improved the accuracy of prediction.

5.2.4.1 Computational disulfide engineering Based on the geometric model of native disulfide bonds, various algorithms have been developed to assess all the possible residue pairs in the protein structure to determine if they meet the geometric criteria for disulfide bond formation. Nowadays, disulfide engineering tools such as DbD [45], modeling of disulfide bonds in proteins (MODIP) [46], SSBOND [47], and BridgeD [48] have been widely used, which make disulfide bonds easily de novo designed [112,113]. Similar to other algorithms, these computational tools predict a large number of disulfide bonds but lack identification of valid candidates, which may lead to the generation of false negatives. Thus, a mature protocol of de novo disulfide engineering has been developed and widely used: (1) identification of the flexible region in a target protein (optional); (2) designing disulfide bonds by computational methods such

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as DbD, MODIP, and BridgeD; (3) filtering out unreasonable disulfide bonds based on the geometric structure (distance, dihedral angles, and steric hindrance), local flexibility (destabilizing energy, B-factor, and RMSD), and location (away from the functional region), the unreasonable disulfide bonds will be eliminated before the experimental screening; (4) verifying the formation of the introduced disulfide bonds by Ellman’s assay; and (5) thermostability analysis of the mutants. This disulfide engineering approach used for the introduction of disulfide bond is believed to be a practical technology for improving protein thermostabilization [114,115]. This disulfide engineering approach was used effectively by Li et al. to enhance the thermostability of Rhizomucor miehei Lipase. At first, DbD2, SSBOND, MODIP, and BridgeD were applied to predict potential disulfide bonds and a total of 155 residue pairs were predicted to form disulfide bonds. After eliminating the residues ˚ of the catalytic triad and other imporhaving high conservation score and the residues within 5 A tant regions, 56 potential disulfide bonds were prepared for the stability assessment by FoldX. Eight residue pairs having negative ΔΔGFold were obtained and further observed in the optimized model viewed by YASARA to determine whether a disulfide bond was formed. Finally, two disulfide bonds S56C/N63C and V189C/D238C were confirmed and contributed to 1.2 C and 4.2 C higher Tm, respectively [116].

5.2.4.2 FRESCO framework Framework for rapid enzyme stabilization by computational libraries (FRESCO) is a computational workflow for enzyme stabilization based on the calculation of folding energies with multiple algorithms, disulfide-bond designs, and in silico fast screening. FRESCO framework was proposed as early as 2014 by the group of Janssen [101]. The procedure of FRESCO framework consists of three steps (Fig. 5.5): (1) mutation prediction based on the folding energies calculated by FoldX and Rosetta, the single mutations with a ΔΔGFold higher than a certain value will be discarded; (2) an optional computational approach for the de novo design of disulfide bonds or a backbonebased amino acid usage survey [117] for calculation of the statistical energy function and van der Waals energy; and (3) assessment of stabilizing mutations. The unreasonable mutations affecting the important residues responding for activity, increasing flexibility, distorting protein structure, introducing internal cavities or unsatisfied hydrogen bond, exposing hydrophobic residues at the surface or destroying existed interactions will be eliminated based on the visual inspection and MD simulations and (4) experimental verification of single mutation and combination of stabilizing mutations. The combination of multiple algorithms and evaluation of stabilizing mutations strongly reduces the amount of experimental screening required to stabilize an enzyme, which makes FRESCO framework a commonly used computational prediction tool [118 122]. In the thermostabilization of limonene epoxide hydrolase from Rhodococcus erythropolis DCL14, 248 variants were obtained at Step 1, and 28 potential disulfide bonds were predicted by dynamic disulfide discovery in MD simulations at Step 2. After the filtration, 130 variants were discarded due to their chemical unreasonableness and 54 variants were eliminated due to the increased local flexibility. Finally, 21 of 64 variants were proved to increase thermostability and three combined variants exhibited a dramatically increased Tm (ΔTm 5 120 C 2 35.5 C) [101].

5.2.4.3 FireProt framework FireProt is also a computational strategy integrating energy-based and evolution-based approaches for predicting highly stable multiple-point mutants. FireProt framework was first developed by

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FIGURE 5.5 General protocol of framework for rapid enzyme stabilization by computational libraries flamework.

Bednar et al. and verified by increasing the thermal robustness of model proteins haloalkane dehalogenase (HLD) DhaA and γ-hexachlorocyclohexane dehydrochlorinase LinA [100]. In 2017 FireProt framework was refined and became popular as webtool by Musil et al. [50]. Currently, FireProt framework which integrates 16 computational tools provides researchers an alternative for designing stable multiple-point mutants. In general, FireProt framework is constructed by three distinct strategies (Fig. 5.6): (1) energy-based approach. FoldX and Rosetta_ddg were applied to calculate the ΔΔGFold of single mutations on nonconservative residues. (2) Evolution-based approach. CA was used to identify potentially stabilizing substitutions. (3) Combination of energy-based approach and evolution-based approach. During these three approaches, FoldX and Rosetta will be used for crosschecking the accuracy of the calculations. Antagonistic effect of mutations pairs could be minimized by using Rosetta to assist the combination of putative beneficial mutations to gain further improvement of thermostability. In 2015 FireProt was verified by increasing the thermal robustness of model proteins HLD DhaA and γ-hexachlorocyclohexane dehydrochlorinase LinA. Combining the evolution-based approach and energy-based approach resulted in the best variant DhaA115 and LinA01 showing about 24 C and 21 C increase in the Tm value indicated by differential scanning calorimetry. Compared with other protein thermostability engineering methods,

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FIGURE 5.6 Workflow of the FireProt framework.

FireProt framework could identify stabilizing mutations with a higher success rate of 80% [100]. Due to robust identification of stabilizing mutations, FireProt framework has been implemented in the thermostabilization of various proteins [123,124].

5.3 CONCLUSIONS AND PERSPECTIVES Enzymes have become a potential alternative for chemical catalysts duo to their unique selectivity, and mild reaction conditions. The global enzyme market has achieved great success and is expected to reach $17.50 billion by 2024. The poor thermostability of natural enzymes remains one of the largest obstacles in the path to their application in industry. Thus, enzyme engineering has received more and more attention as a powerful tool for protein thermostabilization. From traditional directed evolution to de novo protein design, the protein thermostability engineering has become simpler and more efficient. However, it still lacks a single unique tool that will allow us to create thermostable protein catalysts accurately. With the increase in the number of biological protein structures in public databases and sustained advances in the computation for analysis of structure information, it is expected that there will be explosive growth in the research about understanding

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in elucidation of structure function relationship of proteins. The construction of the crucial association between protein structure and thermostability will push rational modifications and de novo design to a new stage.

ABBREVIATIONS RMSF RMSD epPCR RFS B-factor ISM SeSaM MSA CA ASR REAP MD simulations FRESCO

root mean square fluctuation root mean square deviation error-prone PCR rigidify flexible sites approach the protein beta-factor iterative saturation mutagenesis sequence saturation mutagenesis multiple sequence alignment consensus analysis ancestral sequence reconstruction reconstructing evolutionary adaptive paths molecular dynamic simulations framework for rapid enzyme stabilization by computational libraries

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6

Jitesh Kumar1, Girija Kaushal1 and Sudhir P. Singh2 1

Center of Innovative and Applied Bioprocessing, Department of Biotechnology, Mohali, India 2 Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India

6.1 INTRODUCTION Enzymes are the macromolecular biological catalyst that speeds up the rate of reaction by lowering the activation energy inside the living cell or in the cell-free form. The majority of enzymes used currently are of microbial origin because they are relatively more stable and have diverse properties than plant and animal-derived enzymes [1]. The activity of microbial enzymes is dependent on the different physical and biochemical conditions in which they are designed to exhibit their optimum activity. Among them, pH is one of the critical parameters that affect the stability and function of enzymes. Various categories of microbial enzymes operational under a wide pH range have been characterized, making them special to survive the harsh biotechnological processes [2]. Enzymes found in the extreme environment are evolved to maintain its threedimensional (3D) native structure during the course of evolution. But, extreme environment is known to contain less diverse microbial population, and the smaller size of the extremophile genome limits the number of interesting biocatalysts [2]. Therefore, methods to improve the stability of the mesophilic enzymes based on the sequence and structural information are useful for the property not found in natural enzymes. But, they again require rigorous studies and a huge computational tool to perform the mutations. Also, the interactions such as hydrophobic, covalent, Van Der Walls, hydrogen bond, and electrostatic among amino acid residues conferring stability of enzyme to extreme conditions are challenging to redesign. At present, modern protein engineering approaches such as random evolution [error-prone polymerase chain reaction (PCR) or DNA shuffling] are being used to improve the desired properties of naturally available enzymes [3 5]. The screening of mutant protein libraries produced from random evolution is usually extensive, but various high-throughput assays based on the colorimetric substrate are available to handle such difficulties [6]. The current chapter discusses the availability of natural enzymes in alkaline and acidic pH. It also expands the understanding of the mechanism of their stability. The tools for rational and modern protein engineering approaches have also been presented.

Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00006-5 © 2020 Elsevier B.V. All rights reserved.

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6.2 BIOPROSPECTING OF EXTREME ECOLOGICAL NICHE FOR PH STABLE ENZYMES Extreme environments such as acidic and alkaline soil and geothermal hot springs have been on the top priority list of the biologist for the discovery of stable enzymes or enzymes with improved properties. Studies suggest that microorganisms thriving in such type of extreme conditions are equipped with special molecular mechanisms [7] and their evolutionary rates of proteins are faster [8]. Similarly, their proteins are customized accordingly [9]. Therefore, genes can be isolated from the microorganism by culturing (culture dependent) or extracting metagenomic DNA (culture independent), followed by next-generation sequencing and bioinformatics analysis [10]. In the metagenomic study from the alkaline hot spring (76 C, pH 5 8.2) of Galicia, Spain, the gene encoding for lipase was identified, and upon characterization, it exhibited maximal activity at 40 C and pH 7.5. Interestingly, the enzyme possessed 80% of the maximum activity in the range of pH from 6.5 to 8 [11]. On the contrary, microorganisms living in acidic environments such as sulfidic mine areas, marine volcanic vents, and acidic river are acidophilic, which can grow at pH below 3 [7]. Enzymes isolated from such sources may also show stability toward acidic pH. Maltase (α-glucosidase) was isolated from a bacterium, Sulfolobus solfataricus found in an acidic hot spring. This enzyme has an optimum pH of 4.5 at 85 C and retains its 50% activity remaining in the pH range of 3.5 6.5 [12]. Enzymes from acidic and alkaline niche vary in their amino acid composition. Amino acid comparison of acid-tolerant enzymes revealed to contain more Trp, Tyr, Thr, and Ser, and less Glu, Lys, Met, and Arg. The alkaline enzyme has slightly more Trp, Ala, and Cys and less Lys, Arg, and Glu [13]. Such studies reflected the importance of alkaline and acidic environment for mining the novel enzymes with unique sequence composition to sustain the surrounding pH. In addition to this, the composition of their amino acid of the enzymes from the extreme environment is helpful in improving the stability or catalytic efficiencies of the existing enzymes.

6.2.1 MOLECULAR MECHANISM OF ENZYME STABILITY IN THE WIDE PH RANGE The mechanism of enzyme stability defines the way for the development of novel properties and is critical in sustaining the introduced mutation for the desired phenotype. Prior knowledge of factors, which determine the stability, is very much crucial for the structure function relationship of enzymes in the wide pH range [14]. Enzymes operational in the extreme environment should be able to retain its stability for the maintenance of biologically active conformations. The protein in its native state usually forms a compact-specific 3D structure, which is stabilized by the different physical forces such as covalent bonds, noncovalent hydrophobic, electrostatic, Van der Waals interactions, and hydrogen bonds [15]. The role of hydrogen bonding has been found to be the force behind stabilizing the interaction between two mutated amino acid residues of penicillin acylases (PA) at neutral and alkaline pH [14]. The mutant showed a ninefold improvement in the stability at pH 10. Two residues in the wild-type PA, Gluβ482 and Aspβ484, involved in the interacting network of two antiparallel β-sheets, become negatively charged at alkaline pH, leading to repulsion among their side chains and ultimately leading to disruption of the interactions. Studies have suggested that to alkaline

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environment is observed by a less number of charged amino acid and lysine residues and more of arginine and neutral hydrophilic residues (histidine, asparagine, and glutamine) [16,17]. At alkaline pH, the structure of the protein is also destabilized by the deamidation of two residues, that is, asparagine and glutamine, and mutation in these residues can improve the stability [18 20]. Electrostatic interactions among the charged amino acid residues are known to play a crucial role in enzyme adaptation to extreme pH. Subsequently, low pH stability can be further enhanced by the substitution of the acidic and basic residue and vice versa to improve the balance of net charge [21,22]. The pH-dependent stability of the enzyme is known to be regulated by the different ionization states of the side chain of its titrable amino acid [23]. Therefore, changing the pK(a) value of the key residue is a critical approach to improve the pH stability profile of the enzyme. Such type of strategy has been used to increase the stability of endo-beta-1,4-xylanase and protein G at acidic and alkaline pH, respectively [20,23]. Research studies have witnessed the various mechanisms, as described above, for the stability of enzyme structure; however, it is still not clear that whether charged residue on the surface or core of the protein could confer acidic tolerance. Some studies have suggested that electrostatic interactions on the outer surface area of least importance to protein stability, whereas the buried protein core plays a more critical role in maintaining stability [14,24]. As ionizable residues are prone to deprotonation/protonation, they determine an important role in stability. Therefore, strategies can be focussed on analyzing interactions of such residues whose side chains are buried inside the protein core.

6.2.2 DIRECTED EVOLUTION OF ENZYME Directed evolution is one of the prominent approaches used currently by protein biologists to obtain a mutant protein exhibiting the evolved function of their interest. This approach mimics the natural evolutionary process of introducing mutations in the protein in a very short timescale without understanding the function of the protein. Directed evolution is a well-established approach for obtaining an enzyme with altered stability, catalytic activity, or substrate specificity from the existing wild-type versions [25]. In the laboratory, the protein of interest is subjected to iterative cycles of error-prone polymerase chain reaction (epPCR) using their nucleotide sequence as the template (Fig. 6.1). The rate of mutation in each step of epPCR is regulated by controlling the frequency of wrong nucleotide base incorporation through the DNA polymerase. epPCR-enabled libraries are efficient in producing more than 103 variants in a few hours [5]. Gene shuffling is another approach that is used sometimes in combination with epPCR or independently to generate a library of evolved enzymes. The technique relies on the assembly of randomly digested gene pools to create a recombined and evolved enzyme [26]. A plethora of microbial enzymes has been discovered from different ecological niches and tailored for achieving improved activity. However, a limited number of carbohydrate-active enzymes have been improved to enhance their activity in a wide pH range. During the industrial processing of substrates, enzymes undergo various acidic and alkaline processes that affect their stability, leading to a loss in the activity. A thermostable alpha-amylase from Bacillus licheniformis (BLA) was engineered for acid tolerance by directed evolution in order to sustain acidic conditions during industrial starch hydrolysis [3]. Random mutation using epPCR followed by the screening of 5500 clones identified a double mutation (Thr353Ile/His400Arg) in BLA, which showed remarkable

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Second round of mutation

• Generation of mutant gene libraries using error-prone PCR • Cloning and expresssion of mutant libraries for, e.g., in E. coli • Transformation into appropriate host

• Validation of the selected mutant • Selection of best mutant for the second round of mutation

Validation

Diversity creation

Library screening

• Whole cell analysis of mutant gene libraries • Cell lysis and protein activity determination • High-throughput screening using fluorescent or colorimetric substrate

Discard negative mutants

FIGURE 6.1 Methods of enzyme improvement using directed evolution approach.

enhancement in the amylase activity of 11.3-fold higher than wild type. Further, single mutant (Thr353Ile and His400Arg) showed higher amylase activity (3.5- to 6.0-fold) at pH 4.5 than wild type. Acid stability of Thr353Ile and His400Arg mutant was attributed to the hydrogen bonding, hydrophobicity, helix propensity, and electrostatic field interaction. Further, the directed evolution approach was employed to improve both alkaline tolerance and thermal stability of enzymes simultaneously. One such example is xylanase A from Bacillus subtilis (XynA), which has wide applications in bio-bleaching, food, feed and biorefineries, and degradation of lignocellulosic biomass [27,28]. The natural source of XynA derived from Bacillus sp. inhabiting the alkaline soil shows optimum activity at neutral pH. Therefore, the development of thermoalkaline XynA is needed to process Kraft pulps at high pH $ 10 and temperature # 80 C. The successful XynA mutant exhibiting thermoalkaline property was obtained after combining two rounds of epPCR and DNA shuffling on thermostable mutants and native XynA [27]. This step identified a mutant having mutations (Q7H/G13R/S22P/S31Y/T44A/I51V/I107L/S179C) with a temperature optimum of 80 C, a threefold increase in the specific activity as compared to the wildtype enzyme at pH 8.0. The above examples demonstrated the potential application of directed evolution in improving the functions of native carbohydrate-active enzymes for catalytic activity at a wide pH range. However, directed evolution is often faced by challenges such as mutant library preparation, robust screening,

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and biasness in the codon introduction. But, it offers improved functions to the enzyme, which would otherwise take years through natural evolution. The most critical aspects of directed evolution are genetic diversity and high-throughput screening (HTS) of protein mutants, which are crucial for the successful introduction of desired properties in the protein of interest. The genetic diversity of the mutant library of the gene should contain rare beneficial mutations, functional and properly folded protein, nonredundant gene sequences, biased amino acid substitution based on the desired properties, and cost-effective [29]. There are many methods available for creating genetic diversity in the sequences using random mutagenesis, but epPCR is used widely. A good random mutagenesis should favor equal nucleotide substitution such that three-nucleotide representing respective codon for amino acid should change [30]. But, the experimental findings suggest that the ideal mutagenesis should bring out all the possible transitions (Ts) and transversions (Tv) with a 16.67% substitution for each nucleotide pair and a Ts/Tv ratio of 0.5 [30,31]. Apart from this, no deletion and insertion are desirable in the mutant library.

6.2.3 SCREENING STRATEGIES FOR EVOLVED ENZYMES The protein library created by random mutagenesis is usually large (B104 6), which increases the difficulty of finding a variant with desired enzymatic trait [32]. To get rid of tedious screening processes, HTS methods have been devised to screen the protein library of mutants against specific functions based on their true or analogous substrate. Currently, HTS methods can be categorized based on the nature of throughput method, enzyme property (activity, stability, and specificity), and the type of enzymes. During HTS, a direct indicator is preferred to screen the activity of the improved enzyme properties at a wide range of pH. For example, an alkaline stable mutant of xylanase from directed evolution was screened using a substrate dye conjugate, remazol brilliant blue-xylan [33]. Further, flask culture is used to confirm the properties gained by mutants. Such type of indicator eases the process of tedious screening of mutants obtained from directed evolution. There are many other useful screening strategies such as cell- and phage-display that subjects the accessibility of enzymes to substrate and makes the detection process faster [34]. At present, unlike traditional microtiter plate assay, ultrahigh-throughput microfluidic screening system is designed to screen B107 protein variants per day [35]. This technique allows a precise screening of enzymatic activity within a confined microdroplet containing single or multiple substrates. The fluorescently activated droplet, showing positive fluorescent activity due to enzymatic activity, can be sorted at a rate of 400 2000 droplets per second. The microdroplets are created to form an emulsion generated in such a way that reactions are dispersed in oil to form water-in-oil droplets. Such screening platform has been used to screen and enrich microbial cells expressing cellulase from a pool of positive and negative library [36] and even from heterogeneous enzyme libraries [6,37]. Such techniques could be helpful in improving the limiting properties of several carbohydrate-active enzymes, which will help them to survive wide pH environments.

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6.3 A COMPUTATIONAL APPROACH FOR ENZYME IMPROVEMENT Enzymes are known to have wide applications in different industrial processes. A major drawback faced by enzymatic processes at industrial scale is the lack of suitable enzymes, which could be stable at different pH conditions [38]. All enzymes have a defined range of pH stability due to which they either show the least activity or become inactive if used out of this range. Therefore, there is a great need for enzymes that could act in a wide range of pH. There are mainly two approaches to find such enzymes, that is, either by screening environmental samples having acidic/ alkaline pH or enzyme engineering using a computational approach to incorporate desired changes in an enzyme at the sequence and structural levels. In the former approach, it takes a longer time to screen enzymes with the desired activity, as this method involves screening and validation via molecular biology techniques for the prediction of a stable novel enzyme. Also, in the case of random mutagenesis, a wide range of mutant screening is required [39]. This strategy demands more resources and is cost-effective. On the other side, the development of several computational tools provides great assistance in predicting enzyme modifications for molding them as per the desired activity. The bioinformatics analysis of these enzymes using various algorithms resulted in a more precise engineering approach. Unlike the experimental approach, computational enzyme designing is less laborious and more time-effective [40]. Therefore, the computational enzyme designing/redesigning has gained a great interest over the past years. There are several computational methods available for tailoring the enzyme for a specific activity. These can suggest mutations at the sequence as well as structural levels, based on the algorithm incorporated in a particular tool [41]. These tools can deal with a large amount of data available in various sequence and structural databases to suggest best possible modifications for the improved activity. These improvements can be made at specific sequence site, generating site-specific mutants or can be designed by de novo enzyme designing. Therefore, computational enzyme engineering can be categorized into two dominant classes, that is, rational enzyme designing and de novo approach (Fig. 6.2 and Table 6.1) [40].

6.3.1 RATIONAL DESIGN METHOD The rational design method is based on the sequence and structural information of protein [42]. In order to introduce an accurate mutation, this strategy demands precise information about amino acid arrangement and behavior of protein under different environmental conditions such as temperature and pH [43]. Site-directed mutation plays a major role to improve the catalytic properties of proteins. Rational enzyme designing can be used to introduce mutation at a specific position based on prior knowledge about role of amino acid to be modified [42]. It is considered to be the most significant method in predicting site-directed mutations for protein engineering. In order to propose a reliable mutation, mutants are screened based on their structural properties [44]. The mutants generated by rational engineering approach are modeled and evaluated by various bioinformatics tools such as Procheck, Verify-3D, and What-check. These tools evaluate the modeled mutant protein structure base on their stereochemical properties [42]. This approach increases the probability of beneficial mutations with reduced library size, which further reduces the time and effort compared to traditional mutational methods. This method has been widely used for improving the

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FIGURE 6.2 Overview of computational design of enzyme.

Table 6.1 Tools used for de novo enzyme and rational enzyme designing. Tool name

Link

De novo enzyme designing ROSSETTA FoldX AMBER

https://www.rosettacommons.org/software/ http://foldxsuite.crg.eu/ http://ambermd.org/

Rational enzyme designing ZEBRA HotWizard PopMusic

http://biokinet.belozersky.msu.ru/zebra https://loschmidt.chemi.muni.cz/hotspotwizard http://babylone.ulb.ac.be/popmusic

thermostability of various industrially important enzymes in order to improve their activity over a wide range of temperature. For example, thermostability and catalytical efficiency of proteases have been improved using rational enzyme designing approach [45]. To enhance pH stability, complete information of amino acid sequence and their properties at the structural level play an important role [46]. Therefore, concerning the available structural information, planning mutations at a specific site can serve better results. In order to design an enzyme with acidic/alkaline pH stability, homologous variants of the gene of interest (GOI), which are functional at desired pH should be searched using BLASTx tool. All the homologous sequences should be aligned using multiple sequence alignment tools such as ClustalW, T-Coffee, and Muscle. This will check sequence-level divergence from GOI, and mutations can be planned at such sites. These mutants should be modeled using a homologous structural template via computational tools such as Phyre and Modeller.

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This structural folding signifies the probability for the functionality of a mutated enzyme. The major drawback of this approach is gathering the complete structural and biochemical information about the enzyme of interest, which requires expertise in structural bioinformatics [44]. This approach cannot be applied for enzymes for which there is unavailability of sufficient structural data.

6.3.2 DE NOVO DESIGN METHOD De novo approach of protein engineering is another promising method for generating mutants with desired activities. It is based on the study of physical principles that guide protein folding. The understanding of biochemical and biophysical parameters, which is the basis of enzyme functioning, can help in designing a customized enzyme. This will help to address the various challenges related to the generation of beneficial mutants. Computation biology has introduced several algorithms and tools to design protein structures with accuracy at the atomic level from scratch [47]. In this approach, a defined backbone structure and the complete sequence of the protein are unknown. It is very difficult to accommodate a sequence with precise core packaging and accurate hydrogen bonds. Therefore, thousands of conformational designs are generated, starting with either algebraic equations to calculate geometry parameters [48] or assembling small peptide fragments [49]. Combinatorial sequence-optimization calculations are performed for each of the backbone conformations to check for the sequence of lowest energy for a particular conformation. To strengthen these in silico predictions, lowest energy confirmation for a modeled structure is calculated via ab initio structure-prediction calculations. This approach led to an understanding of the principles behind guiding protein structure and folding, which can help in achieving great success for designing a stable protein with atomic-level accuracy [50]. The proteins designed via this method is predicted to be computationally stable but does not guarantee its expression in the surrogate host [47]. Therefore, this turns out to be a significant drawback of this approach. A very small number of designed proteins can adopt a stable folded structure. The possible reasons for such failures are insolubility and polydispersity, which could be the result of intermolecular hydrophobic interactions [47]. Therefore, this approach requires more improved criteria for energy calculations and screening of the best-fit protein designs.

6.4 CONCLUSIONS AND PERSPECTIVE Enzyme performing in extreme pH is a very attractive property that is required during industrial processes. Therefore, enzyme mining from highly acidic or alkaline ecological niches has proven that these sources could be the reservoir of many unexplored stable enzymes. The role of various interactions such as hydrophobic, covalent, Van Der Walls, hydrogen bond, and electrostatic among amino acid residues could be the force behind the stability of pH-tolerant enzymes. Understanding of these stabilization forces in enzymes from extremophiles could be the way to mutate the homologous enzyme from mesophilic source. However, directed evolution is a promising approach over rational approaches that mimic the process of natural selection, directing the enzyme toward

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developing pH-tolerant mechanism. This has been used a lot for enzymes discussed above but less for biomass hydrolyzing biocatalyst. Making an enzyme tolerant to wide pH conditions could bring the solution for the difficulties of enzyme inactivation or low catalytic efficiencies during the process of abundant biomass. The computational designing of the enzyme could predict the amino acid substitution to be done in the enzyme based on the known information of key residues and algorithm that can mimic the reaction conditions. The available in silico and wet lab validation methods provide an enormous opportunity for the improvement in pH tolerance of the enzymes of industrial importance.

6.5 ACKNOWLEDGMENTS The authors acknowledge the Center of Innovative and Applied Bioprocessing (CIAB), Department of Biotechnology (DBT), Government of India for providing the facility to compile this work. JK acknowledges the CIAB for Research Associateship and GK acknowledges the Department of Science and Technology (DST), Govt. of India for Doctoral Inspire Fellowship.

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7

Guoqiang Zhang1,2, Yukun Chen1,2 and Guocheng Du3 1

National Engineering Laboratory for Cereal Fermentation Technology, Jiangnan University, Wuxi, P.R. China School of Biotechnology, Jiangnan University, Wuxi, P.R. China 3School of Biotechnology, Jiangnan University, The Key Laboratory of Carbohydrate Chemistry and Biotechnology, Wuxi, P.R. China

2

7.1 INTRODUCTION Biocatalysis has been extensively used as a greener sustainable process in industry for production of bulk/fine chemicals and pharmaceuticals [1 3]. In this process, enzymes are commonly used as highly versatile and proficient biocatalysts, and they can greatly accelerate green chemical reactions with exquisite specificities such as enantioselectivity and stereoselectivity [4]. Furthermore, over the past decade, more and more novel and potentially useful biocatalysts increased exponentially due to the development of metagenomic mining [5]. These performance features and progress make protein enzyme a competitive candidate for chemical synthesis in the food, chemical, and pharmaceutical industries [6,7]. However, the industrial processes are always harsh, and there is often a significant gap between natural enzyme properties and specific requirements for applications. To make the biocatalyst become competitive and attractive in rigid industrial processes, it is necessary to design and modify enzymes for preferable catalytic activity, stability, and specificity in further. [8]. Protein engineering is a powerful strategy to modify DNA or protein sequence and structure to generate desired, more useful industrial catalysts, which was also used to insight their biochemistry and catalytic mechanism [9]. It involves designing diversity mutant library, screening and identifying mutants with improved properties such as improved activity, increased stability/tolerance, or altered substrate specificity. Complex molecular structures with completely new functions could be evolved and gained by modifying the sequence of individual proteins [10]. Currently, many protein engineering strategies were developed to improve their performance and the most commonly applied methods for protein engineering, include rational design, directed evolution, semi-rational design, and de novo enzyme (Fig. 7.1). With the rapid development of structural biology and bioinformatics, a tremendous amount of progress was made in the protein modeling to achieve the efficient design of mutant libraries for analysis of structure function mechanism [11]. This review focuses on current dominant approaches and the roles of protein engineering played in enhancing enzyme performance in biocatalysis process. Especially, we also highlight and discuss

Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00007-7 © 2020 Elsevier B.V. All rights reserved.

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FIGURE 7.1 Current dominant approaches for protein engineering.

recent advanced strategies for assisting design, screening, and analysis in protein engineering for the effective biocatalysis and cell factories research.

7.2 APPROACHES IN PROTEIN ENGINEERING 7.2.1 RATIONAL DESIGN Based on the rapid development in structural biology, molecular biology technology, and bioinformatics, a variety of strategies in protein engineering have emerged over the past decades. As the most classical method in protein engineering, rational design approach was achieved by sitedirected mutagenesis followed by rational analysis of proteins [12]. Based on this approach, scientists can build a smart library and gain desired enzyme performance from detailed insights of sequence structure function of homologous protein [13]. This approach was inexpensive and technically easy because it is not necessary to screen a large capacity mutant library. In the following part, the popular methods of rational design would be described and discussed, including multiple sequence alignment (MSA), structure-based design, and computational molecular simulation.

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7.2.1.1 Multiple sequence alignment Protein structure and function depend on DNA sequence and the core of protein engineering is modifying the DNA sequence. Therefore, it is a popular and effective strategy to analyze and identify function-related regions based on sequence alignment. For example, MSA can help identify regions of conformational flexibility based on structural similarity, which is a crucial step to predict protein secondary structure and navigate functional residues in different bioinformatics methods [14]. As standard tools for the alignment of evolutionarily related sequences, MSA methods have been used as an algorithmic solution to explore amino acid functionality and evolution relationships from homologous protein sequences [7,15]. It is helpful to establish evolutionary and/or functional cladogram based on known structures and enable successful design of new functional sequences [16]. As a classic MSA method, Multiple Sequence Comparison by Log-Expectation (MUSCLE) was first developed by Robert C. Edgar for MSA of protein and DNA sequences [17,18], which could achieve the high accuracy and processing speed for large numbers of sequences. MUSCLE provides improved speed and alignment accuracy compared with other frequently used programs such as TCoffee (Tree-based Consistency Objective Function for Alignment Evaluation) and MAFFT (multiple alignment using fast Fourier transform, https://mafft.cbrc.jp/alignment/software/). Pervez et al. focused on developing a simulated protein sequences database (SAliBASE) based on different parameters such as insertion rate, deletion rate, sequence length, and so on [14]. Besides, Rozewicki et al. described a Database of Aligned Structural Homologs (DASH) in web version, that integrates MAFFT MSA tool with structural alignments. It provides the alignments for different proteins at domain and sequence levels in the Protein Data Bank [19]. More recently, genome mining, highthroughput sequencing, and coevolution analysis have been integrated and significantly enhanced the accuracy of protein structure prediction, which is also making scalable and accurate sequence alignment possible [20,21].

7.2.1.2 Structure-guided design Structural biology techniques have been used for the determination of protein structures to atomic resolutions and identifying mutagenesis hotspots by molecular modeling [22]. The currently used techniques were protein crystallography or nuclear magnetic resonance (NMR) spectrometry. Based on the information of structures and electrostatic characteristics, enzyme chemists attempt to redesign existing enzymes to enhance their catalytic efficiency [23]. For example, enzyme structure stability can be improved by efficient fixation of N-terminal owning to N-terminal always takes part in maintaining structure [24]. As a classic method of protein stability and ability to resistant denaturation, disulfide bonds could form between two cysteine residues, which is helpful to adapt nonoptimal reaction conditions and eventually enhance their catalysis efficiency [25,26]. Bashirova and coworkers [27] designed disulfide bonds based structure modeling to improve the activity of endoglucanases (EGLII) under industrial conditions. Two positive enzyme variants, S127C/A165C and Y171C/L201C, showed improvement of enzyme activity by 15% 21% with carboxymethylcellulose and glucan as substrate. The structure simulations result also revealed that the introduced disulfide bonds in the two variants stabilized their global structures and enhanced activity. Due to the charged transition states of catalytic reactions, the stabilization of charges is another important tool to improve catalytic efficiency such as introduction of hydrogen bonds. Russell and his colleague

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[28] designed serine protease as a model based on surface charges. They found that pH profile of enzyme can be engineered by altering its surface charges, which had been named as Russell rule for directing the protein design. Bai et al. [29] compared different xylanases under alkaline, neutral, and acidic conditions, which also showed a similar principle to above Russell rule. Based on these structure information, they engineered F11 xylanase and achieved the pH catalytic adaptation. In general, electrostatic characteristic-based structure engineering has shown enough efficiency to redesign the enzymes of which the reaction proceeds via charged translation state.

7.2.1.3 Computational molecular simulation Although the detailed information of catalysis could be obtained from the structural information of the intermediates of enzymes with transition states through X-ray and NMR methods [13], the lacking of detailed structural knowledge for most proteins is still a major drawback [30]. To accomplish the efficient rational design, computational modeling and design increasingly contribute to efforts to improve catalytic characteristics of enzymes [31]. For example, researchers could analyze potential hotspot and predicate mutant properties with help of energy calculations and molecular dynamics (MD) simulations, which provided new efficient ways for solving enzyme rational design problems [32,33]. The dynamic interaction between enzymes and substrates could be simulated by MD simulations and the most optimal variants which consider the accurate positioning of the enzymes by analyzing transition states. These methods included analysis of amino acid sequences, computer simulations of protein folding structures, and screening of desired variants, which provided a guidance to design optimal biocatalysts experimentally. Khoury et al. [34] used computer simulation to analyze the protein binding capacity with cofactor. In this process, they designed the cofactor preference of xylose reductase via cofactor binding energy calculations and effectively changed the native cofactor NADPH dependent to the NADH dependent. After modification, several variants showed significantly higher NADH preference as up to B100-fold change, and others presented cofactor specificity for both NADH and NADPH. Arnold and coworkers [35] also successfully applied computational modeling to design cytochrome P450 BM-3 with altered substrate specificity. It is essential and basic to develop an efficient and appropriate design tool from diversiform available algorithms for protein engineering. For instance, the FoldX was developed incipiently from force field algorithm and has become a powerful and popular web tool [36]. More recently, Buß et al. successfully applied FoldX to analyze folding motifs of proteins and create a more stable ω-transaminase [37]. They have also compared different algorithms with FoldX for the prediction of beneficial mutation sites and showed that the performance of the design tool also depended on protein properties [38]. Compared to random mutation methods, these rational design libraries were efficient to enrich more functional variants, because the computational design algorithms could generate smaller but higher quality libraries based on immensely useful structure function modeling and analysis.

7.2.2 DIRECTED EVOLUTION The rational design requires knowledge of protein structure and active-site function. However, very few enzymes have been studied this intensively. By simulating the Darwinian natural evolution, directed evolution in the laboratory has been developed as a powerful technique to modify the enzyme

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properties, especially for the protein without available information. So far, it has been successfully used to enhance catalytic activity, change substrate specificity and other features [39]. Generally, directed evolution comprises a group of molecular biology techniques, including the random mutagenesis of different starting genes, DNA shuffling for rearrangement and screening enzyme variants with expected properties. The process can be iterated to improve the required properties step by step. The most common strategies are random mutagenesis and DNA shuffling. With the development of biological technologies, some fast, low-cost, and flexible synthetic methods were also developed for building combinatorial variant libraries [40].

7.2.2.1 Random mutagenesis Error-prone PCR is a most frequently used random mutagenesis technique to build a mutant library and it could introduce random mutants into any position of DNA [41]. In this technique, a proofreading ability-deficient Taq DNA polymerase was used for low-fidelity polymerase chain reaction (PCR) and further lowered by alternating the PCR temperature and changing the concentration or proportion of the PCR system [26,42]. Based on previous studies, it is the optimal condition for protein modification if the substitution frequency of DNA could be controlled at 1 3 bp/kb. Liu et al. [43] built a mutant library using error-prone PCR method and isolated a α-amylase BLA (Bacillus licheniformis α-amylase) mutant with T353I and H400R, which showed an B11-fold higher kcat/KM than that of the starting enzyme. Yao et al. [44] used the similar way to get a α-amylase AmyS mutant with K82E and S405R, showing a Bsevenfold improvement in α-amylase activity. Xu et al. [45] improved the enzyme activity of Arabidopsis thaliana LACS9 (AtLACS9) by introducing random mutations into its cDNA using error-prone PCR. Two AtLACS9 variants containing multiple amino acid residue substitutions were identified with enhanced enzyme activity. To improve the efficiency of directed evolution for cytochrome P450 carbene transferase, Arnold et al. built an efficient strategy by combining random and targeted mutagenesis, which showed a power of directed evolution to improve selective functionalization of cyclic compounds [46]. Further evolutionary analysis revealed the beneficial amino acid site, which provides valuable information for lipid biosynthesis.

7.2.2.2 DNA shuffling DNA shuffling is also an efficient way for protein directed evolution in the laboratory, which includes both mutations and recombination [47]. In this process, the gene of interest is randomly cut into segments of different length, then the segments are reassembled by overlap PCR or appropriate DNA polymerase with overlap segments. A library of genes with segments from different genes is then constructed [42]. Yao et al. [48] performed a DNA shuffling of the b3s1 gene for glycine oxidase. A mutantB4S7 was isolated and showed a 3.9-fold increase in the specificity constant. The further analysis of the mutant showed that mutation position P247S is nearby the active site, which plays a crucial role in regulating the substrate preference. Researchers developed a fast and flexible strategy-“mix-and-match” including asymmetric PCR to produce mutagenic fragments and shuffling of variant mutagenic fragments. It could efficiently generate diverse and combinatorial mutation libraries [40]. Recently, computer simulations-assisted directed evolution of protein have been an emerging powerful strategy to generate recombination of blocks of point mutagenesis sequence. For example, a new developed software Shuffle Optimizer is applied to increase the nucleotide homology between DNA of interest and shuffled together for desired performance [49].

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7.2.2.3 In vivo continuous directed evolution Compared to traditional directed evolution in vitro, in vivo continuous directed evolution is becoming more and more attractive [50,51]. The autonomous cycles of orthogonal self-replication and selection have been used to accumulate positive mutations, resulting in the rapid directed evolution of relevant proteins, pathways, or organisms for industrial applications. Liu et al. [52] created an efficient, controllable, powerful mutagenesis system in Escherichia coli, which increase mutation rate B3 3 105-fold over background levels, which is much higher than other widely used mutagenesis methods. They demonstrated that this system can be used to speed up continuous evolution of T7 RNA polymerase in short time. More recently, Ravikumar et al. [53,54] developed a potent, inducible, and scalable orthogonal DNA replication (OrthoRep) system in yeast, which efficiently generates B1 3 105-fold mutation higher than the background genome. Using OrthoRep system, they continuously and rapidly gained malarial dihydrofolate reductases with improved drug resistant through serial passaging. Now in vivo targeted mutagenesis systems have emerged as a new and powerful strategy for rapid directed evolution of protein. Additionally, the high-throughput scale of directed evolution would be achieved with the development of automation and screening platform. Compared to other protein engineering methods, directed evolution is feasible for desired performance caused by random and unexpected mutations. Indeed, a recent study indicated that beneficial mutations can also occur in many regions other than active/binding sites of a protein [55]. These examples support the notion that diverse regions within a protein have the potential to influence function. Although the directed evolution has provided an efficient strategy for protein modification, there are drawbacks existed and need to be resolved, such as a large number of variants often requires high-throughput screening and not all desired performance can be linked to an efficient screening method.

7.2.3 SEMI-RATIONAL DESIGN Although directed evolution has provided an effective approach for improving its catalytic properties [56,57], it mainly relies on efficient screening method for huge mutant library. To further improve the efficiency of protein engineering, the combination of rational design and random method, referred to as semi-rational design, has become an effective strategy in protein engineering [58]. Briefly, this approach preselected potential target amino acid position and limited diversity through collecting information about protein structure and computational predictive algorithms [59 61]. For example, Blomberg et al. applied computational and semi-rational design to modify Kemp eliminase. The targeted enzyme was designed and screened, resulting in accelerated the reaction 6 3 108-fold through combinable rounds of error-prone PCR, DNA shuffling, and rational designed mutagenesis [62]. Aslan et al. [63] used semi-rational design as to target two positions simultaneously for modifying L-lactate dehydrogenase. Eight variants with enhanced activity toward the selected α-keto acids were isolated after screening in a small library. Dulcey et al. performed a comprehensive semi-rational mutagenesis of alkanoate synthase RhlA by comparative modeling and chimeric-based methods [64]. It is helpful to improve efficiency of directed evolution with help of rational design, and it can build smaller but higher quality libraries. Because these smart libraries can be designed by

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modifying a limited hotspot sites, which was probable to affect the performance. Such regions can be preselected using tools for analyzing catalytic mechanism [65] or detecting binding pockets. Bendl et al. [66] developed a web server HotSpot Wizard 2.0 (http://loschmidt.chemi.muni.cz/hotspotwizard) for the rapid detection of hotspots for enhanced performance. It provides a comprehensive understanding of protein structure function relationship and protein ligand reaction, which is helpful for semi-rational design in protein engineering.

7.2.4 DE NOVO PROTEIN DESIGN With the deeper understanding of protein folding and function mechanism, de novo protein synthesis has become mainstream by exploring the protein structure and function relationship, guided by the biophysical and chemical principles [67,68]. The structure and computational simulation-based design and machine-learning strategies have also provided efficient methods to analyze the impact of different amino acids on protein folding and performance [69]. In earlier studies, scientists attempted to design different protein scaffold or structure as expected. Based on the principle of appropriate arrangement of polar and nonpolar residues directing amphipathic structures, Xu et al. [70] employed a “Binary Patterning” strategy for protein design, resulting in self-assembled monolayers from de novo beta-sheet proteins. To design a functional protein as de novo enzyme, it needs to understand the reaction catalytic mechanism, select proper active sites, and design the scaffolds to keep structure stability. Finally, the designs are optimized from their protein folding, thermodynamics, and protein ligand interaction after considering binding pocket and the protein dynamic structure [69]. Besides, modification of existing proteins to generate different properties is another potential method to explore toward novel enzymatic activities through semi-synthesis [71]. It is critical for biological function that how does protein adjust conformation to adapt to the changing environment. Recently, Boyken et al. [72] designed a pH-responsive protein, and they successfully organized histidine with conformation change in buried hydrogen bond. This study demonstrated that de novo protein design can also achieve environmentally triggered conformational changes. It is critical for biological function that how does protein adjust conformation to adapt to the changing environment. Recently, Boyken et al. [72] designed a pH-responsive protein, and they successfully organized histidine with conformation change in buried hydrogen bond. This study demonstrated that de novo protein design can also achieve environmentally triggered conformational changes. Although the development of de novo enzymes is at the outset and the performance is not good enough, the combination of protein evolution and efficient screening methods might be used to optimize those de novo enzymes with poor performance in further.

7.3 ADVANCED TECHNOLOGIES ASSISTING PROTEIN ENGINEERING 7.3.1 SPATIAL ORGANIZATION Spatial organization is a common theme in many naturally occurring multi-enzyme cascades to improves reaction efficiency. Reengineering enzyme catalysts by spatial organization, including

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compartment or scaffold, represent a simple but effective route. These are summarized here in ascending order of complexity: direct fusion of enzymes; colocalization of enzymes on a solid scaffolding of enzymes [73]; and encapsulation of enzymes within containers [74] for biocatalysis (Fig. 7.2A). With advanced synthetic biology development, it has achieved the rational organization of enzymes aided by nucleic acid scaffold and protein scaffold in vivo or in vitro [75,76]. Zhang et al. studied the kinetics of enzyme cascade with glucose oxidase and horseradish peroxidase in a DNA scaffold system. They found the activity of enzymes can be enhanced by suitable pH environment after binding DNA scaffolds [77,78]. Similarly, Zhang et al. designed the self-assembling protein scaffold and used them for alcohol dehydrogenase and amine dehydrogenase, which also showed enhanced activity and stability in enzyme cascade reaction [76]. Further analysis confirmed the effect mechanism of immobilizations and microenvironment would be helpful for enhanced performance [79]. Several successful cases have presented that spatially organized enzymes could display better performance than native enzymes in both single- and multiple-enzyme systems. However, it was problematic that the choice of spatial organization system often affects enzyme activity and stability. It would be highly beneficial to develop a computational method to predicate the effect of spatial organization on reaction kinetics prior to time-consuming testing and optimization.

FIGURE 7.2 Advanced technologies for protein engineering. (A) Spatial organization of enzyme can be designed to modify reaction microenviroment, increase flux or enhance enzyme stability. (B) Cell-free translation systems are based on the cellular ribosomal protein synthesis system, and it can be used to produce target protein in short time. (C) Cell surface display techniques allow for the expression of a target peptide or protein on the surface of a cell through linkage with a genetically fused anchor protein. (D) High throughput screening method can be used to rapidly screen larger enzyme libraries with acceptable coverage.

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7.3.2 SURFACE DISPLAYING SYSTEM Surface display techniques allow for the expression of a target protein on the surface of a cell or biomaterial through linkage with a genetically fused anchor protein, which have been adopted as an important technology for protein engineering (Fig. 7.2C) [80,81]. There are several advantages for the surface display system. Firstly, it is not necessary to cross a cell membrane barrier for substrate and product of the enzymatic reaction. Secondly, the fixed enzymes could avoid proteolytic degradation and keep stable activity. Finally, it can also be easily separated from the reaction mixture achieving reutilization in industrial process [82,83]. For example, surface displaying has been used in yeast to enhance the catalytic activity, stability, and other features of enzymes, including luciferase, lipase90, and lipoic acid ligase [84]. Han et al. [85] developed yeast surface displayingassisted directed evolution system to modify a split peroxidase. There showed noticeably improved activity after rounds of fluorescence-activated cell sorting (FACS)-based screening and isolation. In addition, cell-free display system has also been developed for protein engineering, including ribosome display, DNA scaffold display, and lipid compartment [86]. Compared with cell-surface display systems, cell-free display is a faster, easier and more controllable method for protein display, especially for enzymes involving in cytotoxic reaction.

7.3.3 CELL-FREE TRANSLATION SYSTEMS Cell-free translation systems have developed rapidly over the last decades, and facilitate rapid target proteins production in laboratory. They are based on the cellular ribosomal protein synthesis system and represent an alternative to in vivo expression, which is gaining attention for its simplicity and high degree of controllability [86,87]. Generally, the cell-free translation system is composed of a cell extract from cells such as Escherichia coli. These extracts are supernatants from centrifugation and contain essential components including ribosomes, translation factors, aminoacyl-tRNA synthetases, and tRNAs. Besides, additional RNA polymerase, as well as competent for energy regeneration is also required for efficient protein production (Fig. 7.2B). Recently, these systems have been applied in protein engineering, such as introducing unnatural amino acids into proteins expression. Moreover, these systems do not need to perform in living cells, which would a perfect chassis for synthesis of unstable or cytotoxic proteins in vitro. In the postgenome sequencing era, cell-free protein translation represents a major advantage over other approaches because this approach substantially increases the production rate of expressed proteins. Accordingly, cell-free translation systems are particularly important as high-throughput methods to synthesize proteins of the whole genome, which have contributed to the elucidation of many previously unknown protein functions in massive genomic database [88].

7.3.4 HIGH-THROUGHPUT SCREENING METHODS Protein engineering always involves screening large and diverse protein libraries to isolate mutants with desired properties. To lift the screening efficiency, various high- or ultrahigh-throughput screening methods have been developed to identify variants with improved properties such as increased activity, altered substrate specificity, and increased stability [89]. Previously enzymatic high-throughput screening is commonly performed in 96-well microtiter plates, allowing researchers

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to deal with collection composed only of few thousands of cells. Currently, flow cytometer [90] or microfluidic devices [91] might be the more efficient throughput platform (Fig. 7.2D). In these systems, enzyme performance could be linked to a biochemical or visible signal, such as fluorescence, biosensor-based on enzyme activity [92,93]. Gielen et al. [94] built an ultrahigh rate microfluidic absorbance-activated droplet sorter (AADS), which can screen target enzymes about 300 droplets per second. They integrated trace detection module for screening of phenylalanine dehydrogenases. Fourteen hits showed increased activity after two rounds of directed evolution toward its native substrate. The AADS module paves the way for the implementation of droplet microfluidics in protein engineering. The various high-throughput tools including microtiter plates, flow cytometer and microfluidic technologies are still developing and appear as promising technologies for discover novel enzyme and/or optimize the enzymatic conditions. In further, the combination of bioinformatics analysis to guide library design would help to optimize target enzyme much faster and accurately, and reduce time and labor in the process.

7.3.5 CRYOGENIC ELECTRON MICROSCOPY The recent emerging cryogenic electron microscopy (CryoEM) is a modified and updated electron microscopy technique that has been widely used to determine the protein structure of molecular samples without crystallization [95]. With the development of detector technology and software algorithms, it has presented a significant advantage on analysis of structure of protein complexes, and membrane proteins, which was difficult to determine by other traditional methods. CryoEM has become an important supplementary to crystallography and NMR spectroscopy, especially for analysis of macromolecular structure. For example, researchers have enhanced structural understanding of the respiratory chain with help of advanced CryoEM method. They have studied the structural and functional mechanisms of ATP synthase and found the proton-translocation domain was organized in an unexpected pattern [96,97]. Beyond that, the CryoEM is also used to determine other significant protein complexes such as cytochrome bc1 [98] and respirasome [99]. This progress offers deep understanding of the respiratory-chain complexes in the mitochondria. Fitzpatrick et al. [100] also integrated time factor into the technique and developed 4D CryoEM. It could be used to detect movements of protein molecules at a nanosecond time scale. In recent studies, Muench et al. also integrated X-ray free-electron lasers, crystallographic beamlines, and CryoEM, and it would present a powerful strategy for protein complex determination in the future [99].

7.4 CONCLUSIONS AND PERSPECTIVES Protein engineering is essential to modify and optimize natural enzyme and proteins. It makes enzymes more stable, higher activity, or different substrates/cofactor specificity. Its applications cover a broad range, including biocatalysis for food and medical industry. The development of protein engineering is accompanied by understanding the structure function relationship and catalytic mechanisms. However, it is not yet clear enough to understand the detailed relationship between protein structure and function and it is necessary to develop more and more innovative tools to optimize the protein engineering. Recently, precise statistical models and machine

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[82] Z. Xu, D.W. Kulp, Protein engineering and particulate display of B-cell epitopes to facilitate development of novel vaccines, Curr. Opin. Immunol. 59 (2019) 49 56. [83] I. Chen, B.M. Dorr, D.R. Liu, A general strategy for the evolution of bond-forming enzymes using yeast display, Proc. Natl. Acad. Sci. U.S.A. 108 (28) (2011) 11399 11404. [84] G.M. Cherf, J.R. Cochran, Applications of yeast surface display for protein engineering, Meth. Mol. Biol. 1319 (2015) 155 175. [85] Y. Han, T.C. Branon, J.D. Martell, D. Boassa, D. Shechner, M.H. Ellisman, et al., Directed evolution of split APEX2 peroxidase, ACS Chem. Biol. 14 (4) (2019) 619 635. [86] Y. Shimizu, Y. Kuruma, B.W. Ying, S. Umekage, T. Ueda, Cell-free translation systems for protein engineering, FEBS J. 273 (18) (2006) 4133 4140. [87] H. Ohashi, E. Miyamoto-Sato, Cell-free technologies for proteomics and protein engineering, Protein Pept. Lett. 23 (9) (2016) 819 827. [88] D.M. Leippe, K.Q. Zhao, K. Hsiao, M.R. Slater, Cell-free expression of protein kinase a for rapid activity assays, Anal. Chem. Insights 5 (2010) 25 36. [89] C.K. Longwell, L. Labanieh, J.R. Cochran, High-throughput screening technologies for enzyme engineering, Curr. Opin. Biotechnol. 48 (2017) 196 202. [90] J.C. Baret, O.J. Miller, V. Taly, M. Ryckelynck, A. El-Harrak, L. Frenz, et al., Fluorescence-activated droplet sorting (FADS): efficient microfluidic cell sorting based on enzymatic activity, Lab Chip 9 (13) (2009) 1850 1858. [91] P.A. Romero, T.M. Tran, A.R. Abate, Dissecting enzyme function with microfluidic-based deep mutational scanning, Proc. Natl. Acad. Sci. U.S.A. 112 (23) (2015) 7159 7164. [92] J.D. Bloom, M.M. Meyer, P. Meinhold, C.R. Otey, D. MacMillan, F.H. Arnold, Evolving strategies for enzyme engineering, Curr. Opin. Struct. Biol. 15 (4) (2005) 447 452. [93] P.S. Daugherty, B.L. Iverson, G. Georgiou, Flow cytometric screening of cell-based libraries, J. Immunol. Meth. 243 (1-2) (2000) 211 227. [94] F. Gielen, R. Hours, S. Emond, M. Fischlechner, U. Schell, F. Hollfelder, Ultrahigh-throughputdirected enzyme evolution by absorbance-activated droplet sorting (AADS), Proc. Natl. Acad. Sci. U.S. A. 113 (47) (2016) E7383 E7389. [95] R.A. McLeod, J. Kowal, P. Ringler, H. Stahlberg, Robust image alignment for cryogenic transmission electron microscopy, J. Struct. Biol. 197 (3) (2017) 279 293. [96] A. Zhou, A. Rohou, D.G. Schep, J.V. Bason, M.G. Montgomery, J.E. Walker, et al., Structure and conformational states of the bovine mitochondrial ATP synthase by cryo-EM, Elife 4 (2015) e10180. [97] W. Ku¨hlbrandt, K.M. Davies, Rotary ATPases: a new twist to an ancient machine, Trends Biochem. Sci. 41 (1) (2016) 106 116. [98] K. Amporndanai, R.M. Johnson, P.M. O’Neill, C.W.G. Fishwick, A.H. Jamson, S. Rawson, et al., Xray and cryo-EM structures of inhibitor-bound cytochrome bc1 complexes for structure-based drug discovery, IUCrJ. 5 (Pt2) (2018) 200 210. [99] S.P. Muench, S.V. Antonyuk, S.S. Hasnain, The expanding toolkit for structural biology: synchrotrons, X-ray lasers and cryoEM, IUCrJ. 6 (Pt 2) (2019) 167 177. [100] A.W. Fitzpatrick, U.J. Lorenz, G.M. Vanacore, A.H. Zewail, 4D cryo-electron microscopy of proteins, J. Am. Chem. Soc. 135 (51) (2013) 19123 19126.

CHAPTER

CHIMERIC ENZYME DESIGNING FOR THE SYNTHESIS OF MULTIFUNCTIONAL BIOCATALYSTS

8

Jyoti Singh Jadaun1, Lokesh Kumar Narnoliya2, Archana Srivastava1 and Sudhir P. Singh2 1

Botany Department, Dayanand Girls Postgraduate College, Kanpur, Uttar Pradesh, India 2Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India

8.1 INTRODUCTION The chimeric or fusion protein is termed for the protein that is produced as a translational result of fusion or chimeric gene, which is constituted by combining the divergent segments from two or more separate genes. Protein engineers also termed “hybrid” for such chimeric proteins as these are derived from partial or full consorting of two or more proteins, which may differ in their source, functions, or intracellular locations. Chimeric proteins are of two types: (1) natural and (2) artificial or synthesized.

8.1.1 NATURAL CHIMERIC PROTEINS Some chimeric proteins exist naturally as they play a crucial role in the speciation process through evolution in protein families [1]. Predominantly, chimeric proteins exhibit additional function(s) without losing the original functionality of the partner proteins [2]. Evolution of protein families is executed as a result of gene reorganization exercises such as gene duplication, horizontal gene transfer, chromosome translocation, exon shuffling, alternative splicing, conversion of a noncoding region into coding element, and repositioning of transposable elements [3,4]. Genome reorganization leads to the production of a protein with modified or entirely new function, or sometimes a novel phenotype is generated after repetitive gene reorganization [5]. BCR-ABL1 is one of the most studied examples of naturally occurring chimeric proteins. It is considered as oncogene associated with chronic myeloid leukemia, and it is responsible for the occurrence of Philadelphia (Ph) chromosome due to the activity of BCR-ABL1 fusion protein [6]. In Caribbean Sea whip coral, Plexaura homomalla, a fusion protein of heme peroxidase and lipoxygenase is reported, which cumulatively generates allene oxide, a precursor for prostaglandin-like molecules [7]. Another suitable example of a chimeric gene is jingwei (jgw) gene family of Drosophila, which plays a vital role in hormonal metabolism [8,9]. Drosophila nuclear transport factor-2-related (Dntf-2r) gene is evolved through the retroposition process of parental Dntf-2 gene, and Dntf-2r is male-specific [10]. There are many other examples of naturally occurring fusion proteins such as PvuII type Restriction-modification (R-M) system of Niabella soli strain DSM 19437. The fusion of restriction Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00008-9 © 2020 Elsevier B.V. All rights reserved.

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endonuclease (REase) and its transcriptional regulator C protein lead to protective methylation of DNA of the newly formed host cell [11]. Another example is pyrroline-5-carboxylate synthase, a bifunctional enzyme, which plays a vital role in proline biosynthesis [12].

8.1.2 ARTIFICIAL OR SYNTHESIZED OR RECOMBINANT CHIMERIC PROTEINS Protein engineering is one of the most exciting topics of research, and it is so much vast and deep that still, we are on periphery in the knowledge of this subject. Since 1980, various methods and techniques have been invented for deciphering the nature, structure including active site determination, and catalytic activity of the targeted protein. Each individual protein has its characteristic functional domains for executing its own specific catalytic activity. A protein can catalyze biochemical reaction efficiently but may not ensure good thermostability or high optimum temperature. Sometimes, this restriction is due to pH or metallic inducer or any other factor of the reaction mixture or reaction conditions; therefore one functional enzyme is unable to fulfill the demand. Thus to maximize the capabilities of an enzyme, concept of the chimeric enzyme is raised. This concept is also useful for determining the functional regions in the polypeptide or the critical amino acids of the active site, which participate in the catalytic reaction. Rational design and directed evolution are the most commonly used approaches for designing of chimeric enzymes [13]. Construction of fusion or chimeric proteins is the result of consecutive research leading to understand the interaction of DNA and protein. Earlier, protein engineering was centered on the identification and characterization of a single enzyme, but now advanced technologies can figure out the role of multiple enzymes in a single shot. Thus fusion proteins can be synthesized in a more robotic manner, and these proteins can minimize the compatibility issue of two proteins when they are used simultaneously in the reaction as a mixture. Currently, designer enzymes are synthesized according to their requirement in the process. With the aid of recombinant DNA technology and bioinformatic tools, chimeric protein can be easily synthesized by following a planned strategy. In a simple procedure, corresponding genes of targeting proteins are fused with or without linker and cloned in the suitable expression vector, and the vector is directed into a worthy host. Mostly bacterial cells (e.g., Escherichia coli), yeast cells, and mammalian cell lines HEK293 (human embryonic kidney-293) and CHO (Chinese hamster ovary) are regularly used as a host for chimeric protein production. Any plant cell or rarely insect cells can be used for this purpose. In vivo study of chimeric protein predicted the phenological or physiological role while in vitro study reveals their modified catalytic characteristics [14]. After screening of combinations of genes, a useful version of the fusion protein is constructed. Bi, tri, or multifunctional protein can be produced by the fusion of two or more proteins or by insertion of functional and catalytic domains of different proteins into single peptide [15] (Fig. 8.1). There are several enzymes, which naturally have multifunctional properties, so by mimicking the natural pathways, some attempts have been performed, and many of them are working successfully [16]. Generally, fusion protein has the characteristics of each partner protein, although, in some instances, reduction or loss in functionality of parent proteins is also reported [17]. Research interest in the creation of fusion enzymes is expanding tremendously since the last decade due to their applicability in agriculture, industries, and therapeutic sectors [18
20].

8.1 INTRODUCTION

121

FIGURE 8.1 A schematic diagram representing the different methods used for fusion of domains or proteins: (A) end-to-end gene fusion, (B) domain insertion, and (C) posttranslational conjugation.

Multifunctional proteins are synthesized, having the desirable qualities of different proteins into single peptide [18]. Recently, fusion protein having xylanase and cellulase activity is used for degradation of lignocellulosic biomass, and it performed more efficiently than the individual enzymatic treatment [21]. Efficient hydrolysis of complicated polymers (xylan and cellulose) leads to the generation of simple sugars (xylose and glucose), which can be further channelized for production of other valuable products such as biofuels [22,23]. Another advantage of such reaction is the management of the agro-industrial waste in less effort [24]. For the deinking of waste paper, synthesized chimeric enzyme lipase-cutinase (Lip-Cut) performed more efficiently than individual or mixture of parental proteins [25]. The major role of fusion proteins is reported in the development of chimeric therapeutic agents. Many chimeric drugs are approved by the FDA, and these are synthesized and marketed by several drug companies [26,27]. Multifunctional chimeric antibodies have great potential as Fc fusion-based proteins are proving very useful for the treatment of cancer [28]. By seeing the importance of fusion protein in each sector, it becomes necessary to explore more about this area because such protein engineering technologies are very advantageous for minimizing various issues such as fuels shortage, therapy of diseases and genetic disorders, issues in drug delivery, waste management, and issues of proteomics research. In this chapter, we will describe different methods of production of the fusion protein with their diverse applications.

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8.2 METHODS FOR GENERATING CHIMERIC/FUSION PROTEINS Designing of the chimeric enzyme is needed to develop enzymes with improved or novel functionality (Fig. 8.1). Hence, by following the nature’s footsteps, experiments have been carried out at laboratory scale, and success has been achieved up to a significant level [29
31]. Three models are commonly used for designing the fusion proteins: (1) end-to-end fusion, (2) domain insertion, and (3) posttranslational conjugation, and these are described in detail below.

8.2.1 END-TO-END FUSION This is the classical approach for designing and construction of fusion proteins. This secures wide applications in tagging of protein, display action of targeted protein, and clinical therapeutics [32
34]. Two proteins are joined into end-to-end manner directly by ligation of C-terminal end of one protein with the N-terminal end of another protein (Fig. 8.1). The one gene should be free of the stop codon, and another gene should lack a ribosome binding site. Three fates can be adopted by the fusion product such as loss in activity of one protein or both protein and loss in activity of one with induction in another activity or induction in the activity of one or both protein. To generate such fusion, one prerequisite is the complete study about the component proteins including their structural pattern, kinetic parameters (optimum temperature, optimum pH, substrate affinity, thermostability), substrate binding sites, and their inducer or inhibitors. Another requirement is to check the compatibility of component protein with others. After confirmation of partner proteins, another major factor is the order of partner proteins, sequential appearance in chimeric protein, because the order of protein decides the functional activity of the chimeric protein. Sometimes, there is no effect of the arrangement of domains, but sometimes it is critical for proper folding to show catalytic activity. Hence, different combinations of acting domains as well as their sequential arrangement should be tried. Tandem fusion is a simple method and just joining of the corresponding gene of partner protein resulted in a single polypeptide having the coding region of both the genes. If C- and N-terminal ends are flexible, they act as a bridge and provide space for conventional folding of amino acid residues. Occasionally, the rigidity of terminal ends creates steric hindrance, which renders appropriate folding of the polypeptide [35]. However, direct ligation of coding sequences showed some undesired consequences such as poor solubility of the fusion protein, formation of inclusion bodies, and misfolding, which hampers the catalytic activity [36]. Use of linker overcomes the limitations of direct fusion such as solubility, yield, catalytic activity, biological activity, and protein folding [34] (Fig. 8.1).

8.2.1.1 Linker Linker is a short chain of amino acid residues or oligopeptide, which introduces between the domains of proteins. Linker plays a crucial role in the synthesis of the fusion protein as it affects the solubility and flexibility of amino acid residues for attaining maximum activity. Choice of the linker is dependent on the nature of target protein or the features which one desire to introduce in targeted fusion product. Prime features of linker are (1) the length of linker, (2) composition of amino acid residues, (3) the sequence of amino acids, (4) flexibility, (5) folding, (6) stability, (7) hydrophobicity, and (8) secondary structure confirmations such as helical, coil/bend, β-strand,

8.2 METHODS FOR GENERATING CHIMERIC/FUSION PROTEINS

123

and turns [34,37,38]. The main intention to add linker between domains is to extend the conformation for allowing maximal flexibility of domains [36]. Length of the linker can be increased to create space between the domains for excluding the steric hindrance [39]. The linker may occur naturally in some proteins, or it may be synthesized as per need.

8.2.1.1.1 Naturally occurring linkers Naturally, linker molecules are present between the domains of the multidomain protein, and several linkers are reported on the margins of functional domains of the protein [40]. Various roles of natural linkers are suggested by authors such as structural support to domain, cooperative interdomain interaction, stability to partner protein, catalysis, transport or signal transmission, biological activities, protein dynamics, and allostery [41
45]. Study reveals that average length of linker varies from 4.5 to 21.0 amino acid residues and length of linker affects the hydrophobicity in reverse order, so linker of long length has high solvent accessibility [36]. In linker region, frequently occurring amino acid residues are serine, glycine, proline, arginine, phenylalanine, threonine, glutamic acid, glutamine, aspartic acid, lysine, asparagine, and alanine [36,46]. Due to smaller size, it is favorable to include glycine-rich linker for flexibility; therefore many chimeric proteins have glycine-rich linker (R33GIGRGG39), which modulates the functionality by bridging helices α1 and α2 [47]. Proline is also prevalent in occurrence, but it provides stiffness and rigidity to structure due to its cyclic nature [48]. Natural linker acts as a spacer between domains and functions independently without interacting with adjacent domains. Linker undergoes such conformational transitions, which is suitable for exposing the active site toward the substrate. Thus interpeptide linkers in multidomain proteins play a key role in the interaction of domains [49]. A detailed study of natural linkers favors creating artificial linkers for fusion protein [34,46,50] (Table 8.1).

8.2.1.1.2 Artificial or synthesized linker By employing the tools of molecular biology and synthetic biology, artificial linkers can be synthesized by using the backbone of natural linkers. Various linkers have been synthesized for the fusion of proteins (Table 8.1). Linkers can be categorized into three types on the basis of the structure module and amino acid composition. Flexible linkers. Flexible linkers are used to access freedom for the advantageous movement of the domains. Repeated glycine (nonpolar), serine, or threonine (polar) sequence is routinely used for synthesis of flexible linkers because glycine aids for mobility and serine
threonine forms hydrogen bond with water molecules, which prevent unacceptable interactions of the linker with the domain [46]. Some examples of flexible linkers are KESGSVS, SEQLAQFRSLD, EGKSSGSGSESKST, AMGPSSGAPGGGGS, and GGGGS [34,58,59]. The flexibility of linker also affects the solubility of the protein; in a study, it is reported that linker flexibility affects the formation of active inclusion bodies of green fluorescence protein (GFP)-PhoC (acid phosphatase) fusion protein [60]. Other flexible linkers, (GSSGSS)n, (GSSSSS)n, or (S6)n, where n is the number of repeats, are also used for rational designing of multidomain protein [61]. The effect of linker on catalytic activity is represented by fusing the artificial linker (GSGSGSGSG) of variable flexibility on variants of BsCel5A, cellulase 5A from Bacillus subtilis [49]. Rigid linker. It is not a compulsion that the use of flexible linkers will always produce desirable results. In some demonstrations, it is reported that flexibility limits the proper functioning of domains. It may be due to destructive interdomain interactions or unfavorable close proximity of

Table 8.1 Application of multifunctional enzymes for biomass degradation. Fusion enzyme

Donor organism

Host

Fusion plasmid

Method

Activity

Reference

1. Cel (cellulase)
CBM (carbohydratebinding module) 2. Cel
CBM
CE (carbohydrate esterase) 3. Xyn (xylanase)
CBM
Cel Cellulosome (xylanase)
laccase

Camel rumen metagenome data

Escherichia coli BL21 (DE3)

pET26b-cel-cbm pET26b-cel-cbm-ce pET26b-xyn-cbm-cel

Tandem fusion

[21]

xylanase from Geobacillus stearothermophilus and laccase from the aerobic bacterium Thermobifida fusca

E. coli BL21 (DE3)

pETduet-Xyn-c-Lac

A tetravalent scaffolding

Expansin (EXLX1)
endoglucanase (CelD)

Expansin from Bacillus subtilis and endoglucanase from Clostridium thermocellum

BL21 (DE3)

Using linkers 1. GS3 linker— (GGGGS)3 2. GS6 linker— (GGGGS)6

β-1,4-endoglucanase (Endo5A) and β-1,4endoxylanase (Xyl11D)

Paenibacillus strain ICGEB2008

E. coli DH5 strain

1. pET-22b-(EXLX1CelD) 2. pET-22b-EXLX1GS3-CelD 3. pET-22b-EXLX1GS6-CelD pQE30 (End5A-GSXyl11D)

Fusion protein XynCBM-Cel exhibited synergistic xylanase/ cellulase activities and more efficiently hydrolyzed rice and barley straws than the native (XylC and CelC) proteins The fusion protein Xync-Lac showed twofold enhancement in release of reducing sugars from wheat straw compared to protein lacking the laccase Fusion protein EXLX1GS3-CelD showed highest cellulose degradation activity

[52]

lipase-cutinase (Lip-Cut)

Cutinase (Cut) from Thielavia terrestris NRRL 8126 and lipase (Lip) from Thermomy ceslanuginosus

Pichia pastoris

pPICZαA-Lip-Cut

End-to-end fusion by using linker (GPTTTSSAPN PTSSGCPNATK)

Cellulase/xylanase (Cel48A exoglucanase-Cel5A endoglucanase

Thermobifida fusca

E. coli BL21

Scaf  BTFA

Using tetravalent scaffolding

Fusion protein (End5AGS-Xyl11D) has 1.6- and 2.3-fold higher specific activity than Endo5A and Xyl11D Ink removal efficiencies of fusion enzyme were 25.8% and 16.2% higher than that of the controltreated laser-printed paper and newspaper, respectively Tetravalent (Cel48ACel5A-Xyn10B-Xyn 11A) cellulosome system showed approximately twofold more enzymatic

Glycine-serine linker (GGGGSGGGGS)

[23]

[51]

[25]

[53]

endoxylanasesXyn10B and Xyn11A) 1. CutinaseCBMCel6A 2. CutinaseCBMCenA

Xylanasearabinofuranosidase (Xyln
Ara)

xylosidase
arabinosidase (Xar)xylanase (XynA)

Xylnase-arabinofura nosidase Xylnase
xylosidase

CBMs Cel6A and CenA of cellulase from T. fusca and Cellulomonas fimi, respectively Carboxyl terminus of cutinase from T. fusca Catalytic region of xylanase gene (xynZ) from C. thermocellum and the full-length arabinofuranosidase gene (abfA) form Geobacillusstearo thermophilus xylosidase
arabinosidase (Xar) from Thermoanaerobacter ethanolicus and xylanase (XynA) from Thermomyces lanuginosus

b-D-xylanase from Clostridium thermocellum, arabinofuranosidase from Bacillus stearothermophilus and a xylosidase from Thermoaerobacterium sp.

activity than mixture of divalent enzymes In presence of pectinase, both chimeric proteins showed threefold higher catalytic efficiency than native cutinase

E. coli BL21 (DE3)

1. pET20b/cutinaseCBMCel6A 2. pET20b/cutinaseCBMCenA

Overlap PCR

Tobacco (Nicotiana tabacum L. cv. Samsun-NN)

mp2300Xyln-Ara

24-amino acid linker (GGGGGADQ LAIG PMYNQ VVYQYPN)

B19 mg active enzymes per gram of total soluble protein of leaf of T1 transgenic plants

[18]

E. coli JM109 (DE3)

1. pET-20b-XarXynA 2. pET-20bXar
L1
XynA 3. pET-20bXar
L2
XynA 4. pET-20bXar
L3
XynA 5. pET-20bXar
L4
XynA 6. pET-20bXar
L5
XynA pET-29b-Xyln
ara and pET-29bXyln
xylo

With the help of linkers: L1—SAGSSAA GSGSG L2—(SAGSSAA GSGSG)2 L3—(GGGGS)2 L4—SGGSSAA GSGSG L5—SAGSSAA ASASG

As compared to other construct, Xar
L1
XynA and Xar
L4
XynA more efficiently release the reducing sugar

[54]

24-amino acid linker (GGGGGA DQLAIGPMYN QVVYQYPN)

Xyln
Ara fusion releases B30% more xylose from wheat arabinoxylan than the mixture of parental enzymes (xylanase/ arabinofura nosidase)

[55]

E. coli strain BL21 (DE3)

[33]

(Continued)

Table 8.1 Application of multifunctional enzymes for biomass degradation. Continued Fusion enzyme

Donor organism

Host

Fusion plasmid

Method

Activity

Reference

Cellulase-bglucosidase

Thermotoga maritima MSB8

E. coli strain BL21(DE3)

pET-28a(1)/ Cel5C-BglB

Overlap PCR

[56]

Glucanase-xylanase

Glucanase from Bacillus amyloliquefaciens and xylanase from Bacillus subtilis

E. coli

pET-30a(1)Glu-Xyl

Overlap extension PCR

Carbohydrate-binding module (CBM family 1)-feruloyl esterase A (FAEA)-endoxylanase B (XYNB)

CBM and feruloyl esterase A (FAEA) and endoxylanase B (XYNB) from Aspergillus niger

Aspergillus niger strain D15#26

1. pAN52.3-FAEAXYNB 2. pAN52.3-CBMFAEA-XYNB

Overlap extension

Both cellulase (Cel5C) and β-glucosidase (BglB) activities only in construct having BglBfused downstream of cel5C Bifunctional (Glu-Xyl), with greatly enhanced glucanase activity associated with a decrease in xylanase activity Both constructs are more efficient than free enzyme Release of ferulic acid from corn bran was increased after addition of CBM

PCR and linker (GSTYSSGSSS GSGSSSSS) was added between genes

[36]

[57]

8.2 METHODS FOR GENERATING CHIMERIC/FUSION PROTEINS

127

domains and linker or improper space for orientation of individual domains [62]. To avoid these limitations, there is an advancement in the synthesis of rigid linkers, which creates proper space between domains to evade the interaction of domains. These linkers have Pro, Arg, Phe, Thr, Glu, and Gln residues as these residues provide stiffness and rigidity to the linker [36]. To overcome the limit of flexible linkers, an empirical rigid linker A(EAAAK)nA (n 5 2
5) was developed by a group of researchers [63]. This linker generates α-helices such as secondary structure, which support rigidity to regulate the distance and undesirable interaction between the domains. Another type of rigid linker includes proline-rich sequences such as (XP)n, where X may be any amino acid, and n denotes number. Naturally, proline-rich sequences are also found and presence of proline support stiffness which allows peculiar distance between domains [48,64]. In a study, various types of linkers were screened (flexible, rigid, and mix-type linker) for a fusion protein of nitrilase of Thermotoga maritima MSB8 and CotG (outer coat protein) of B. subtilis to obtain optimum catalytic features [65]. Flexible linker includes GGGGS, (GGGGS)2, and (GGGGS)3 and rigid linkers are (EAAAK), (EAAAK)2, and (EAAAK)3; another assortmenttype linkers are GGGGSEAAAKGGGGS and GGGGSEAAAK. Among these linkers, linker (GGGGSEAAAKGGGGS) displayed desirable improvement in thermostability and pH, which was about 2.67- and 1.9-fold in contrast to protein without linker [65]. So mixing of flexible and rigid type of linker may produce more effective consequences, so further trials are needed to find out the most advantageous sequence of the linker. Cleavable linkers. Earlier described linkers have a stable connection with the domains by covalent bonding, so they cannot be separated from domains during the reaction. In certain cases, the attachment of linker to protein may negatively affect its bioactivity, stability, or steric hindrance; therefore cleavable linkers are designed. In vivo cleavable linkers Cleavable linkers can be removed from the fusion protein through chemical agents or in vivo cleavage. An in vivo cleavable linker was designed by using the advantage of reversibility of disulfide linkage. A most fascinating example is dithiocyclopeptide linker, which contains a thrombin-sensitive sequence and an intramolecular disulfide bond. The fusion protein of transferrin and granulocyte colony-stimulating factor is constructed by using this linker. For removal of the linker, the fusion protein was treated with thrombin, and consequently, thrombinsensitive sequence is released, although protein domains are connected by a disulfide bond. For separation of proteins, this disulfide bond is reduced so that each partner protein can show their realistic functions without any hindrance [66]. Protease-sensitive cleavable linkers In vivo cleavable linkers can be designed on the basis of protease sensitivity of linkers. Specific protease is released by a peculiar cell in a pathogenic state and specificity can extend up to cells, or tissue or cellular compartment. The linker may be designed in such manner, which should contain protease cleaving sequence. In a study, protease cleavable linker (VSQTSKLTRAETVFPDV) was used for fusion of recombinant coagulation factor IX (rFIX) and albumin protein. This fusion product has 10- to 30-fold higher activity than the proteins devoid of the cleavable linker, and simultaneously half-life of protein was also enhanced [67,68]. In another study, efficiency of antibody
drug conjugates (ADCs) is increased as an anticancer drug. Here F16 immunoglobulin was fused with monomethyl auristatin E, a potent cytotoxic drug, using dipeptide linker with four variants such as Val-Ala (valine
alanine), Val-cit (valinecitrulline), Val-Lys (valine
lysin) and Val-Arg (valine
arginine). Among them, Val-Ala displayed better performance [69].

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CHAPTER 8 CHIMERIC ENZYME DESIGNING FOR THE SYNTHESIS

Photo cleavable linker Kakiyama and his coworkers developed a linker design [5-(and-6)carboxytetramethylrhodamine (TMR)-GKLAKLAKKLAKLAKKLAKLAKGC (TMR-KLA-C) peptide], which is cleavable under UV light irradiation, so it may enhance the authenticity of cell toxicity
related studies [70].

8.2.1.1.3 Linker designing tools and databases Due to exceeding interest in the production of fusion proteins, linkers become unavoidable tools in protein engineering. It is necessary to set a depository of linker molecules, whether artificial or natural. Steps have been carried out in this direction, and some databases and tools are programmed for linker designing. There are various types of linker molecules available in database; moreover, linker molecules can be synthesized according to the need [71]. LINKER is the most frequently used database, and it provides a set of linkers according to the user’s specifications such as length, choice of amino acid, and secondary structures such as a helix. This database is the earliest step in the linker designing research area. This database contains four categories to make easy searching of target linker: (1) linker: they only connect two uninterrupted domains, (2) helical: more than 30% residue of this linker engaged in helix formation, (3) nonhelical: less than 30% amino acids are engaged in helix formation, and (4) all linkers: it contains all type linkers in a single platform. This database is based on a loop library, which contains 14,734 loop sequences, which is derived from Brookhaven Protein Data Bank (PDB). This library contains a large fraction of short length (4
9) sequences, 12% of medium loops (10
20 residues), and a small fraction of longer loops. The main input required for this tool is the length of the linker, and many additional options are also available, such as restriction cleavage site or specific protease cleavage site. Moreover, the user can give a choice for amino acid residues. Although this was a good practice, it is not in use currently due to the small data set. Therefore another data set is developed for interdomain linker designing, this database has higher sequences, and it is divided into two classes: helical and nonhelical. It accepts the query in any format such as length of linker, PDB code, and helical content. In this database, after entering a query, a set of linkers are displayed in a hyperlink. By clicking this hyperlink, a new web page is open, which showed the 3D atomic structure of linker [36]. Unfortunately, this database became outdated and is not so much accessed by users due to lack of amendments according to the updated number of protein sequences in PDB. Synlinker, a web-based program, is developed for the generation of the fusion protein with the help of linkers. There is a collection of about 2150 natural linkers and 110 artificial/empirical linkers. Other than linker length, it contains additional criteria for query searching such as solvent accessibility, terminal amino acids of the linker, compositional bias, proteolytic site, natural or artificial linker type, and flexibility, and the most favorable feature is that it shows the conformational structure of linker in JSmol viewer [72]. It is publically accessible at http://synlinker.syncti.org/.

8.2.1.2 Overlap extension polymerase chain reaction or end-to-end cloning Overlap extension polymerase chain reaction (PCR) is one of the most suitable techniques to join two or multiple genes together [73]. If we wish to combine two genes together, primers of each gene should be synthesized separately but with some modifications. The forward primer of the first gene should have a restriction site compatible with the targeted expression vector and reverse primer has an overlapping sequence of 5’ end of the second gene. The forward primer of the second

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gene has an overlapping sequence of the first gene, while reverse primer has a restriction site compatible with the expression vector. Both genes are amplified separately and then ligated together. Ligation product can be checked by performing PCR with the forward primer of the first gene and reverse primer of second gene. If the size of the product is the sum of base pairs of both genes, then proceed to clone in the expression vector. In expression vector, the order of gene can be further confirmed by restriction digestion analysis of the product. By following similar strategy, two, three, or multiple enzymes can be fused for the production of the multifunctional protein.

8.2.2 DOMAIN INSERTION Domain is the functional unit of protein, and a number of the domain may vary from one to many as in multidomain proteins. More than two-third and 30% proteins of prokaryotic and eukaryotes, respectively, are devoted to multidomain protein families. Natural occurrence of these proteins shows their great impact in evolutionary processes [30]. Generally, in multidomain proteins, domains are arranged continuously, but in some cases, discontinuous domains are also reported [74]. In discontinuous domains, continuity of domain breaks down due to insertion of another domain. Hence the occurrence of discontinuous multidomain proteins became the backbone for protein engineering through directed evolution approach [75,76]. This domain insertion may be single (single-time insertion of domain) or multiple (more than one domain insertion), and further multiple insertions can be of three types: (1) nested insertion (insertion of new domain into intermediate domain), (2) two-domain insertions, (3) three-domain insertions (Fig. 8.2). Although domain insertion is more laborious than tandem fusion, still it is gaining attention for construction of fusion protein. Single domain proteins are favorable for domain insertion, although more than 50% of proteins are with a single domain, so this fact is not so much attentive [77]. In this strategy, two key points are needed to consider: there should be close proximity between N- and C-terminal of a parent polypeptide, and another is the selection of insertion site in parent protein. Insertion of the domain without creating any disturbance in the integrity of the original polypeptide chain is very critical; therefore selection site plays a decisive role in getting success. (A)

N

(B)

N

(C)

N

(D)

N

C

Domain A

N-terminal domain A

Domain A

Domain A

C-terminal domain A

Domain B

Domain B

Domain B

Domain C

C

C

C

FIGURE 8.2 An overview of different types of domain insertions in proteins: (A) single insertion, (B) nested insertion, (C) two-domain insertion, and (D) three-domain insertion. N and C represent the N- and C-terminal end of parent domain or protein, respectively.

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Surface loops and turns are the most preferred locations for using as insertion site due to their negligible participation in the functionality of protein [30]. Ehrmann and his coworkers followed the sandwich fusion of alkaline phosphatase with MalF (multispanning cytoplasmic membrane protein) to develop a tool for studying the topology of several cytoplasmic membrane proteins [75,78]. They produced several chimeric constructs by varying the cutting site in MalF, and they obtained alteration in the activity of both genes in different chimeras. Another most acclaimed example of domain insertion approach is the fusion of dihydrofolate reductase and β-lactamase (BLA) into phosphoglycerate kinase [79]. The resulted protein showed all types of activity of corresponding partners, so this showed that parent protein and host cell could sustain the genetic load if domains are positioned in right conformations. Domain insertion technique is used to regulate, or switching of enzymes such as allosteric enzymes, and several attempts have been performed for the production of biosensors or reporter based on this strategy. Switch protein is the protein that can undergo conformational changes according to the strength of the signal, which may be chemical or physical. The fusion of TEM BLA into the MalE (maltose-binding protein) produced chimeric enzyme showing penicillinase and maltose-binding activities, simultaneously. By controlling the concentration of maltose, the conformation of MalE protein can be regulated, which indirectly determines the activity of lactamase [80]. Further improvements have been made to explore more about the switch functioning of BLAMalE fusion protein [81]. Another most potent reporter gene is GFP, which may be used as insertion domain, or it can be used as parent protein for insertion of other protein. For the construction of the cAMP sensor, random insertions of GFP in cAMP were directed, and the further library was screened out to find the best insertion site [82]. In another study for creating an optical biosensor, the BLA gene was inserted into the loop of GFP protein [83].

8.2.2.1 Construction of domain insertion libraries In domain insertion technology it seems difficult to get the predicted insertion of the domain into protein so random insertion libraries are opted out to make it easier (Fig. 8.3). Library construction can be done by using any of the following methods: 1. DNase I nuclease: For the creation of domain insertion library, the prerequisite is to create a single break in double-stranded (ds) DNA, which is the acceptor gene cloned in the expression vector. DNase I is an endonuclease, so it can break down the single or ds DNA. It cleaves the phosphodiester bonds of ds DNA and leaves phosphate group at 50 end and OH at 30 end. By optimizing the quantity of DNase I, the number of breaks in DNA can be controlled, so it is used to create random insertion libraries or random circular permutation libraries [82,84]. The activity of DNase I depends on its concentration, quantity of targeted DNA, and reaction time, although it is very sensitive to temperature. Moreover, the occurrence of large deletions in the acceptor DNA molecule may lead to a frameshift. It is tough to get DNA with only a single break even after meticulous optimization. So limitations of this strategy are: needed a very high quantity of DNA, optimization of reaction with each preparation of plasmid, a high number of “junk” type DNA fragments, difficulty to screen, and maintain a large library. Other methods have been developed to reduce such complicity. 2. S1 nuclease: S1 nuclease is also endonuclease, but preferentially it digests the ss DNA and RNA, so commonly it is used for blunting of DNA and for RNA mapping [85]. Although it can

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FIGURE 8.3 A schematic representation of preparation of domain insertion library. It includes following steps: generation of single break in acceptor gene (orange color in plasmid) and then insertion of target DNA (red color insert) in the acceptor gene having single cut. Ligation is followed by recircularization of plasmid, which is then transformed into host for screening of the plasmid having target gene.

liberalize the supercoiled plasmid DNA under restricted conditions, so it can be used as an alternative of DNase I in construction of insertion library. Study reveals that it breaks mostly in inverted repeat-rich regions, which occur mostly in the origin of replication. Any disturbance in origin of replication may be dangerous for replication of plasmid, so modification in the location of S1 digestion is required. It is found that the addition of cofactor Zn11 instead of Mg11 resulted in a diverse pattern of digestion. Tullman and his colleagues applied this feature to create domain insertion library by using S1 endonuclease, and they successfully created a library, which contains TEM-1 BLA gene (circularly permuted change in the order of amino acids) inserted into ribose-binding protein of E. coli [86]. Generation of a large number of junk library members is a major limitation of this method. 3. Multiplex inverse PCR: It is a variant of PCR in which two diverse primers are used for amplification of complete plasmid sequence [87]. This is a more focused method because here insertion site can be chosen according to the permissive region of acceptor gene. Primers are designed to have some overlapping regions, which may be three or six or more than six base pairs. In the final construct, one and two additional amino acids are added on

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overhangs of the acceptor gene by addition of three and six base pairs, respectively. This method can be used for deletion or tandem duplications in the acceptor gene by excluding or including some codon, respectively. The advantages of this method are: information about the location of gene information, insertion is only in acceptor gene, no disturbance in the backbone of plasmid, low rate of a combinatorial library, rare chance of frameshifting, and less effort to maintain the library. The limitation is only that a separate set of primers is needed for each insertion site, cost of reaction setup, and time for optimization of PCR conditions. 4. Use of engineered transposon: Transposons are known as “jumping genes” as they can move from one location to another in the genome. Engineered transposon can be used as an alternative of DNase I to create a break in the acceptor gene [88]. For carrying forward this concept, an engineered Mu transposon called as MuDel is applied for creation of domain insertion library [89]. Earlier, this MuDel or mini Mu is used for trinucleotide deletion or insertion [90]. MuDel is DNA construct, which can be inserted into genome efficiently and accurately, but the location is not predicted. Transposition is carried out by using MuA transposase. This MuDel is engineered to carry the recognition sequence for MlyI, a type II restriction enzyme that cuts outside the recognition sequence, along with transposon insertion mechanism. Integration of MuDel in target DNA is followed by digestion with MlyI, which creates a loss of three nucleotides (Fig. 8.4). This deletion opens the site for deletion or exchange of nucleotides. A DNA cassette is designed, which is termed SubSeq and it reverses back the lost nucleotide triplet by inserting the donating sequences [89]. The fusion protein of cytochrome b 562 (cyt b) and BLA TEM-1 is created by applying MuDel transposon/MlyI methodology [76]. In another study, MuST transposon is used for insertion of xylanase from Bacillus circulans with interdomain linker into lacZa gene as an acceptor in plasmid PUC19 [91].

8.2.3 POSTTRANSLATIONAL CONJUGATION The term protein conjugation is used for joining together of two different proteins by covalent bonds. Cross-linking or conjugation between proteins may be induced by physicochemical processes such as heat, mechanical pressure, or photooxidative treatment or by shifting to alkaline pH conditions [92]. Cross-linking can be performed by using cross-linking chemical agents that have bifunctional groups having reactivity with functional groups of proteins such as glutaraldehyde and carbodiimides [93]. Enzymatic or chemoenzymatic methods are also used for cross-linking of proteins, and this method is supported by the mechanism of posttranslational modification of a protein by a specific enzymatic set [94]. Some examples of natural intermolecular protein cross-linking are ubiquitination, collagen, and elastin cross-linking, fibrin cross-linking, and these reactions are catalyzed by specified enzymes [94,95]. The enzymes of these processes are used for in vitro crosslinking between peptides. Cross-linking may be performed in two ways: (1) direct covalent bond formation as performed by transferases and (2) indirect covalent bond formation as through reactive species generated by an intermediate enzyme. After activation, proteins themselves form the covalent bond between functional groups. Enzymes used for cross-linking of protein are listed as transferases and hydrolases, transglutaminases, lysyl oxidases, oxidoreductases, peptidases, laccases, tyrosinases, and peroxidases [94].

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FIGURE 8.4 An outline of transposon (MuDel)-based domain insertion method. MuDel transposon having MlyI restriction site is inserted into DNA and then followed by restriction digestion with MlyI. MuDel is removed out from DNA but it also carries extra three nucleotides of DNA. So this gap of three nucleotides becomes the site for deletion or substitution or domain insertion.

8.3 APPLICATIONS OF CHIMERIC OR FUSION PROTEINS 8.3.1 USE OF MULTIFUNCTIONAL ENZYMES FOR BIOMASS DEGRADATION Lignocellulose is the major part of plant biomass, and it is constituted by lignin, cellulose, and hemicellulose. Cellulose is the most abundant polymer in nature, and it is made up of glucose subunits linked via β-1,4-glycosidic bonds [96]. Hemicellulose is another predominant organic polymer, and it is constituted by xylan. Xylan is characterized by the presence of a β-1,4-linked D-xylopyranosyl as linear chain along with diverse side chains of monosaccharide or oligosaccharide molecules [97]. Lignin is a natural heteropolymer of phenyl propene subunits such as coniferyl alcohol, sinapyl alcohol, and p-coumaryl alcohol. These subunits are joined together by cross-linking by C
C or ester or ether bonds. It constitutes 10%
25% of plant biomass [98].

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Due to complex and heterogeneous nature of plant cell wall, it is hard to break down these polymeric substances. It requires a series of enzymes such as exoglucanase, endoglucanase, xylanse, β-xylosidase, and laccase [98,99]. Nature evolves some cellulosomes as structural entities, which occur in some anaerobic microbes, and they effectively degrade the lignocelluloses [100]. Cellulosome is multifunctional, and it is based on cohesin
dockerin interaction for assembly and surface layer homology domain for attachment to the cell surface. In cellulosome, a carbohydrate-binding module is also present on noncatalytic scaffolding protein, which expedites the degradation of cellulose [101]. Several trials have been executed on the basis of this natural cellulosome model, and fusion proteins are synthesized by joining cellulosome or its few domains with other enzymes like laccases [23]. Another method to degrade such biopolymers is to build up the fusion proteins having domains of different proteins such as cellulase and xylanase into a single polypeptide. Several examples are available in the literature, which present that chimeric enzyme had better catalytic efficiency than individual native proteins or their mixture (Table 8.1). For optimizing the suitable condition of xylan hydrolysis, several chimeric xylanases are also designed by end-to-end or domain insertion technology [20]. For making a bifunctional enzyme having xylanase and mannanase activity together, four constructs are designed: (1) xyn-S4-man (xS4m), (2) man-S4-xyn (mS4x), (3) xyn-E4-man (xE4m), and (4) man-E4-xyn (mE4x). For joining these proteins, A[EAAAK] 3 linker is used, and these constructs were expressed in Pichia pastoris strain X33. Results revealed that chimera XE3M greatly enhanced the hydrolysis of luffa cylindrical fiber [102]. So it is a cost-efficient process to hydrolyze these complicated polymers into soluble sugars, which are further utilized to manufacture some valuable products such as biofuel [98].

8.3.2 FUSION PROTEIN-BASED BIOPESTICIDES (ANTIMICROBIAL PEPTIDES) In animals and crops, pathogenic attack causes severe physical or economic loss, although many solutions are available but none is worthy to carry for longer time. The uninterrupted practice of chemical pesticides presented a horrible scenario in front of us as they harshly affected the environment and human health. To overcome the limitations of chemical pesticides, the use of biopesticides is emerging as an eco-friendly solution to this problem as they are natural, cheaper, and biodegradable. These are frequently used for preventing fungal and bacterial attacks. Several types of biopesticides are developed and in continuation of this, antimicrobial peptides (AMPs) have emerged as a potential tool for avoiding the harmful effects of pathogens. The AMPs are oligopeptides having five to more than 100 amino acid residues, and these AMPs may be natural or synthetic [103]. Due to proteinaceous nature, it is quite easy to synthesize these peptides in in vitro conditions and their production can be scaled up after optimizing the conditions. Pathogens have also diversified their nature and behavior according to the defense mechanism of the host. After a certain time period, they develop resistance; therefore a more advanced defensive technology is needed, which should be equipped with more powerful tools to throwback the parasitic attacks. Recently, chimeric biopesticides are found more influential than other conventional pest management approaches. Chimeric biopesticides are fusion peptides that are derived by joining two or more peptides, having antimicrobial effects against different pathogens. Their synergistic effect helps them to kill a range of pathogens [104]. A chimeric AMP “super-Blad” is

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developed, which functions against filamentous fungi, yeast, human, and plant pathogenic bacteria. It is the combination of Blad protein and SP10-5/Sub5 protein [105]. Blad is a polypeptide (20 kDa) having fungicidal properties, while SP10-5 (12 amino acids) or Sub5 (12 amino acids) peptides possess antibacterial properties against gram-positive and gram-negative bacteria and certain fungi such as Candida albicans [106,107]. Applications of fusion protein technology may provide a new dimension to synthesize novel AMP with more versatility. Moreover, bifunctional AMPs are used for surface coating on titanium as substrate, while free ends have antimicrobial activity [108]. So this method can be adopted for the synthesis of engineered biomaterial having the antimicrobial property for inhibiting the bacterial colonization.

8.3.3 FUSION PROTEIN AS BIOPHARMACEUTICAL Recombinant proteins are used as therapeutic agents for the last two decades, and recombinant insulin represents the most widely used biotherapeutic agent. Currently, the use of recombinant fusion proteins is boosted up in the treatment of several challenging diseases due to huge amelioration in protein engineering techniques [109]. Synthesis of therapeutic fusion protein is executed to meet the following outputs: (1) fusion of polypeptide to pharmacokinetically active peptide to extend its half-life and improve the pharmacokinetic properties to increase the duration of action, (2) fusion of tissue or cell targeting domain to active molecule (peptide or toxin) leads to cellspecific action, (3) fusion of two differently active protein domains to construct multifunctional peptide [28]. Although there are several reasons for bending toward the fusion protein therapeutic technology, the most emphasizing factor is the extension of the half-life of pharmacologically active molecules [110]. The fusion protein may be natural or synthetic, and generally, one partner has a molecular recognition facility, and another partner provides features such as stability, target specificity, or delivery route leader. To increase the serum half-life of active protein, fusion partner should have low immunogenicity, proteolysis resistant, less or negligible disturbance in the activity of protein, and stability or low rate of renal clearance. Examples of fusion partners include IgG Fc fragment, human serum albumin, transferrin, elastin-like peptide, artificial gelatin-like protein, homo-amino acid polymer (repeat of same amino acid), polypeptide Pro-Ala-Ser, and PEGylation (chemical cross-linking to polyethylene glycol) [28,110]. Among them, Fc fusion proteins have received more attention because Fc domain prevents the degradation of fused protein due to binding with neonatal Fc receptor (FcRn), which aid in recycling of protein. The first chimeric drug was developed by fusing Fc fragment of IgG1 with tumor necrosis factor alpha receptor-2 P75 [111]. To date, up to 74 modified antibodies or chimeric proteins are approved drugs for selling in the market, and 11 out of them are Fc fusion drugs. Some examples of fusion protein approved for marketing are Rituxan (rituximab), Zenapax (daclizumab), Remicade (Infliximab), Simulect (Basiliximab), Unituxin (dinutuximab), and Anthim (obiltoxaximab) [27]. Innovative antibodies are also used as therapeutic agents, and these modifications of antibodies can be performed by various methods: (1) Fc fusion, (2) ADCs, (3) immunocytokines, (4) checkpoint modulators, (5) bispecific antibodies, (6) antibody mixtures (7) T-cell redirection, (8) Chimeric Antigen Receptor (CAR) -T cells and T-cell receptor (TCR) -T cells [27].

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Successful clinical testing of chimeric or fusion drug established a direction to explore new findings in this area.

8.3.4 FUSION PROTEIN-BASED BIOSENSOR (PROTEIN SWITCHING) Protein switch refers to the protein that can exist in “on” (active) or “off” (inactive) state as it recognizes the input signal and responds it through an output signal. This input signal may be any stimuli such as concentration of ligand, variation in pH or temperature, change in ionic strength, fluctuation in light intensity and wavelength, or binding with other molecules by covalent or noncovalent bonds (hydrogen bonds or hydrophobic effect or van der Waals forces) while output signal may be an enzymatic reaction or binding affinity [81]. This protein switch can convert its conformational state according to the strength of the signal. Therefore this versatility and sensitivity of protein switch favor the development of biosensors. In the last decade, a large number of biosensors have been developed, and few of them are commercially available. In biosensor, the output signal is measured by coupling it with “read out” devices, which may read the optical (fluorescence emission) or biochemical (catalytic or binding) or electrochemical (electron transfer) activities. The ability of modifying protein switches has great potential to produce new protein switches or modification in the existing ones. The activity of partner protein can be regulated by joining the two different domains together like allosteric regulation [83]. Several fluorescent sensors are developed by using the optical signal transduction activity of green fluorescent protein (GFP) by domain insertion method. Insertion of calcium-binding calmodulin protein in GFP protein has been achieved as calmodulin is sensitive to Ca11, so it shows sevenfold increments upon saturation with Ca11 ions [112]. Fo¨rster resonance energy transfer (FRET) is becoming a potential tool to study the protein
protein interactions, and many biosensors are designed by using FRET technique [113]. The transfer of excited-state energy from one fluorophore (donor) to the other fluorophore (acceptor) through dipole
dipole resonance coupling, and this is a nonradiative process and dependent on the distance between the fluorophores. In FRET, many variants of GFP protein are produced to act as fluorophore partners, for example, a cyan fluorescent protein (CFP) and yellow fluorescent protein (YFP) or GFP/Cherry [114]. These fluorescent proteins are attached to either N- or C-terminals of the target protein and FRET measurement is based on the increment in fluorescence of acceptor protein and loss of fluorescence of donor protein [115]. FRET has applications such as measuring the calcium ion in living cells, heavy metal detection, protein phosphorylation, and in vivo detection of Zn11 ions, and it is also used for measurement of intramolecular distances within proteins [113,115]. Cameleon, a fluorescent indicator of Ca11, is a chimeric protein and it consists of blue fluorescent protein or CFP (FRET donor), calmodulin (CaM), CaM-binding peptide of myosin light chain kinase (MLCKp), and YFP (FRET acceptor) [116]. Many variants of cameleon are available, which are generated by altering the properties of GFP proteins and calcium-binding modules [113]. By using the fluorescence property of GFP, bimolecular fluorescence complementation (BiFC) technique is also developed for examining the protein
protein binding interaction. In this technique, YFP or CFP protein is split into two domains which unable to fluorophore, although rejoining resulted in a bright fluorescence. Proteins under examination are ligated separately to the splitted N- or C-terminal fragment of YFP or CFP protein. If proteins are interacting, then fluorescence can be seen either in in vitro or in vivo condition [117].

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8.3.5 OTHER APPLICATIONS Fusion proteins have several applications in diverse areas of biological sciences. Tagging of the small peptide to target protein fastens its purification from the mixture of proteins. The most frequently used protein tag is hexahistidine tag, and by adding this tag, protein can be purified in simple steps by using affinity chromatography. His- tag has an affinity for metal (nickel, zinc, or cobalt ions), so immobilized metal ion binds specifically to the protein having His- tag [118,119]. Other tags such as glutathione S-transferase protein, FLAG peptide, and S-tag are also used for this purpose [120,121]. Sometimes, fusion of peptide or protein domain improves the catalytic features of protein [122]. Another advantage of fusion protein is to locate cellular localization of target gene [123]. Fusion proteins can be used as reporter gene or used to synthesize the display libraries of targeted gene [124]. Recently, fusion proteins are used for tissue engineering [125].

8.4 CONCLUSIONS AND PERSPECTIVES Designing of fusion protein became a thrust area for protein engineers to decipher recent challenges. Chimeric protein engineering is a very dynamic and versatile process and this field is too vast to explore even after two decades of focused research; still, we have scarce knowledge about the structure and functional relationship of domains in different types of proteins. The major constraint is that a similar technique is not applicable for all proteins due to the diverse composition of amino acids and folding patterns of the protein. In the last decade, there is significant progress in fusion protein engineering, which increases the scope of their applicability in medical, agriculture, and enzyme-dependent industries, development of biosensors, etc. By applying fusion protein and innovative antibodies technique, we can develop drugs with a broad spectrum with enhanced stability, which can be constructive input for the welfare of society. Protein engineering technique, thus, has a remarkable socioeconomic impact. The major drawback of fusion drugs is immunogenicity, so further investigation should focus to curtail this bottleneck to the satisfactory level. It is required to combine the synthetic biology and bioinformatics to develop more efficient fusion proteins. It is easier to get the genomic and transcriptomic data of any organism. By applying these data inputs, researchers can get a new roadmap for proteomics. Study of protein structure, function, and protein
protein interactions with the computer-assisted programs can accelerate protein engineering research. On the basis of in silico outputs, studies can be optimized, which will save time and provide the precise outcomes.

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CHAPTER

ENZYME ENGINEERING FOR ENANTIOSELECTIVE BIOTRANSFORMATIONS

9

Kaiyuan Tian, Balaji Sundara Sekar, Joel Ping Syong Choo and Zhi Li Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore

9.1 INTRODUCTION Biocatalysis is being increasingly recognized as a useful tool for chemical synthesis in many industries. This is largely due to the unique advantages enzymes enjoy over chemical catalysts, such as being able to function efficiently at mild temperatures and pressures without the use of harsh organic solvents, their nontoxic and biodegradable nature, their ability to work in tandem with other enzymes to create one-pot cascade reactions, and their exceptional regio-, chemo-, and enantioselectivity [1
6]. The latter is perhaps the most important benefit of biocatalysis—the naturally high enantioselectivity of many enzymes enables a wide variety of synthetic approaches for yielding enantiopure products and intermediates [1
3,6
10]. Such enantioselective biotransformations have been proven to be extremely useful in the synthesis of various chiral chemicals.

9.1.1 ENANTIOSELECTIVE BIOTRANSFORMATIONS The earliest example of enantioselective biotransformation was a kinetic resolution reported by Louis Pasteur, in which he reacted racemic ammonium tartrate with a Penicillium glaucum mold and isolated the remaining levorotatory tartaric acid [11]. Since then, biocatalytic kinetic resolution has been widely studied and reported, as well as used for a range of industrial processes (Fig. 9.1A). Classical examples are the use of acylation enzymes (amidases, proteases, esterases, and lipases) to prepare a variety of chiral amino acids, amides, alcohols, and esters from the corresponding racemates [12
17]. Degussa has commercialized an industrial process using acylases for synthesizing enantiopure amino acids [18]. Other excellent examples such as the application of epoxide hydrolases for the resolution of racemic epoxides have also been reported [19
23]. A more elegant application of enantioselective enzymes can be found in dynamic kinetic resolution, which involves the synthesis of a chiral product from one enantiomer of a racemic substrate, while the other enantiomer is continuously being racemized during the reaction (Fig. 9.1B). Many enzymatic dynamic kinetic resolution methods have been reported, using a metal- or enzyme-catalyzed racemization step in tandem with an enzymatic asymmetric transformation step [24
26]. For instance, lipases are capable of producing enantiopure esters or amides from racemic Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00009-0 © 2020 Elsevier B.V. All rights reserved.

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(A) Kinetic resolution (R)-selective alcohol dehydrogenase

OH R1

R2

OH R2

R1

Racemic alcohol

O +

K1

R2

Ketone

(S)-alcohol

(B) Dynamic kinetic resolution O R1

racemization OH

(S)-acid

(R)-selective lipase/esterase

O R1

OH

(R)-acid

R2OH

(C) Asymmetric synthesis (S)-selective amine O transaminase K1

ketone

Amine donor

Coproduct

OR2

(R)-ester

NH2 R1

R2

O R1

R2

(S)-amine

FIGURE 9.1 Applications of enantioselective biotransformations. (A) Kinetic solution of a racemic alcohol with an (R)selective alcohol dehydrogenase to obtain the enantiopure (S)-alcohol. (B) Dynamic kinetic resolution of a racemic acid with an (R)-selective lipase/esterase to obtain the enantiopure (R)-ester. (C) Asymmetric synthesis of an enantiopure (S)-amine from the prochiral ketone with an (S)-selective amine transaminase.

secondary alcohols or amines in combination with a ruthenium- or palladium-based catalyst for racemization [27
30]. Apart from (dynamic) kinetic resolution, the asymmetric synthesis of chiral products directly from prochiral substrates is also possible with enantioselective biocatalysts (Fig. 9.1C). Asymmetric preparation methods are highly desired because they turn nonchiral compounds into chiral ones. A well-studied example is the use of alcohol dehydrogenases (ADH) to obtain enantiopure secondary alcohols from the corresponding prochiral ketones via asymmetric reduction [31
34]. Other instances of asymmetric synthesis include the use of lyases for aldol formation [35
37], transaminases for amine and amino acid production [34,38
40], and multiple types of C 5 C bond functionalization catalyzed by enzymes such as monooxygenases [41,42], dioxygenases [43,44], haloperoxidases [45,46], hydratases [47,48], and ammonia lyases [49,50], among others [51]. Despite all the examples of enantioselective biocatalysis available in the literature, there is still a lack of enzymes with sufficient or correct enantioselectivity, resulting in a need for the development of new enzymes with desired enantioselectivities. To develop these new enzymes, we must first ask ourselves: what makes an enzyme enantioselective?

9.1 INTRODUCTION

147

9.1.2 ENZYMES AND CHIRALITY: THE RELATIONSHIP BETWEEN STRUCTURE AND FUNCTION Enzymes are biomacromolecules made up of a sequence of amino acids. The sequence and physical properties of these amino acids define the secondary, tertiary, and quaternary structures of the protein, which in turn determine its functional nature. Because of Emil Fischer’s “lock-and-key” and Daniel Koshland Jr.’s “induced-fit” theories, we have a rudimentary understanding of how a protein’s structure contributes to its unique and precise catalytic functions [52]. For an enantiospecific enzyme, the 3D structure of the protein may favor the binding of one enantiomer of the substrate (or product) over the other enantiomer at the active site for the chemical reaction to occur, thus giving rise to enantioselectivity (Fig. 9.2). Unfortunately, a fundamental understanding of the exact relationship between sequence, structure, and function still remains to be elucidated, making the rational or de novo design of enzymes from the ground up extremely challenging. To date, there are only a few reports of de novo enzyme design, using the Rosetta computational tool developed by the Baker Lab, and they include an esterase [53], an organophosphate hydrolase [54], a retroaldolase [55], and a Kemp eliminase [56]. However, these computationally designed enzymes exhibited poor catalytic activities in the lab; therefore, the current knowledge of sequence
structure
function relationships is still insufficient for true rational design of proteins. Thus other methods are typically favored for tailoring enzymes for the needs of enantioselective biotransformations.

FIGURE 9.2 The “lock-and-key” theory explains how the structure of an enzyme can discriminate between different enantiomers of a substrate.

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9.1.3 TOOLS FOR MODIFYING THE ENANTIOSELECTIVITY OF ENZYMES In 2018 Frances H. Arnold was awarded the Nobel Prize in Chemistry for her pioneering work on modifying enzymes by directed evolution. In her seminal work, she used directed evolution to improve the solvent tolerance of subtilisin E, allowing it to function in high concentrations of dimethylformamide [57]. Subsequently, directed evolution has been widely used as a powerful tool for engineering many enzymes to suit a range of different catalytic requirements [58
60]. This general approach has been demonstrated a number of times to improve or even invert the enantioselectivity of enzymes toward nonnatural substrates [61,62]. One early example is the improvement of enantioselectivity of a lipase for the hydrolysis of (S)-2-methyldecanoic acid p-nitrophenyl ester [63]. In another powerful example showcasing the versatility of directed evolution, an epoxide hydrolase with poor enantioselectivity [2% (S,S)] toward cyclohexene oxide was evolved to produce two separate mutants: one with enhanced enantioselectivity [94% ee (S,S)] and another with inverted enantioselectivity [80% (R,R)] [64]. In contrast to rational design, directed evolution was originally conceived as a random approach in combination with screening or selection of desired enzyme properties. Despite the obvious disparity between these approaches, both methods have recently been converging toward a semirational design methodology for the creation of small libraries with high-quality mutants, decreasing the screening effort required while increasing the probability of obtaining a desired enzyme. In this work, we will discuss various enzyme engineering strategies for the modification of enzyme enantioselectivity, including random and semi-rational approaches, along with highthroughput screening assays for measuring enzyme enantioselectivity. Recent examples of enzymes engineered for enantioselective biotransformations will be reviewed as well.

9.2 DIRECTED EVOLUTION: A DARWINIAN APPROACH FOR TAILORING THE ENANTIOSELECTIVITY OF ENZYMES Directed evolution is a selection process performed in the laboratory to engineer enzymes with improved properties by modifying the DNA sequence, or gene, encoding the protein. The general strategy behind directed evolution is inspired by Darwin’s theory of evolution and natural selection: random and small mutations in the genotype during the reproduction process can give rise to a generation of multiple individuals with phenotypes differing from the parents and/or each other. A selection bias is then applied, allowing the fittest individuals to go on to produce the next generation, with mutations of their own [65]. With the advent of routine polymerase chain reaction (PCR) techniques in the laboratory, it soon became possible to generate millions of copies of a single gene while adding small yet significant mutations in the DNA. These genes are then cloned in suitable hosts and the corresponding enzymes expressed, resulting in a library of mutant enzymes. Selection is then carried out in the form of a high-throughput screening method, which has to be designed for rapid, sensitive, and accurate measurement of the desired trait to be obtained. This cycle may then be carried out multiple times to achieve the target enzyme. Therefore, the successful engineering of an enzyme heavily depends on efficient library construction and robust screening methods.

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9.2.1 MUTANT LIBRARY CONSTRUCTION As previously mentioned, mutant library construction may be random or semi-rational. Random approaches do not require a priori knowledge of an enzyme other than the encoding gene information, and are thus favored for applications where little is known about an enzyme’s structure. The drawback of such random mutagenesis methods is that a statistically high proportion of the mutants generated are unlikely to exhibit any improvement over the parent enzyme, thus the screening effort required is laborious. On the other hand, semi-rational approaches make additional use of the structural information of the enzyme in combination with the enzyme engineer’s insight to predict the locations of “hotspots”—amino acid residues which harbor a high probability of modifying the enzyme’s traits or functions when mutated. Semi-rational methods typically result in small yet “smart” libraries which require less screening effort, and therefore time, to obtain positive mutants. In this section, we will cover both random and semi-rational methods for constructing mutant libraries.

9.2.1.1 Random approaches Random mutagenesis is a very useful tool for creating a huge library of mutants. Combined with an effective high-throughput screening assay, this technique can successfully create proteins with desired aspects even in the absence of crucial information such as crystal structure of the protein, active site of the enzyme, and so on [66
69]. Here, we discuss two of the most widely used approaches reported for random mutagenesis [66,70
73].

9.2.1.1.1 Error-prone polymerase chain reaction PCR is used to amplify desired DNA targets by providing DNA polymerase, template DNA, DNA primers and deoxyribonucleotide triphosphates (dNTPs) [74]. The fidelity of the DNA polymerase can be reduced by modifying the composition of the reaction mixture or by using mutated DNA polymerases [75
77]. This reduced fidelity introduces mutations randomly in the amplified DNA and the process is called error-prone PCR. Error-prone PCR is the simplest method of random mutagenesis and therefore is a popular approach for creating huge libraries of random mutations [78]. The typical methodology for performing error-prone PCR is to provide unbalanced nucleotide concentrations of dNTPs for the reaction [79]. By doing so, misincorporation of nucleotides is forced to occur during elongation. High concentrations of DNA polymerase is also beneficial while performing error-prone PCR as it will guarantee replication even with base-pair mismatches [80]. Another approach is by changing the cofactor concentration of Taq polymerase. Magnesium ions present in the PCR reaction buffer act as cofactors for Taq polymerase, hence increasing the magnesium ion concentration or replacing the magnesium with manganese can decrease the fidelity of the polymerase and increase the mutation efficiency [81,82]. Mutant DNA polymerases with reduced proofreading capability have also been reported [83,84].

9.2.1.1.2 Gene shuffling To create entirely new functions in an enzyme, drastic changes are required in the coding sequence, which limits the application of error-prone PCR because it can only generate several site mutations at a time. To overcome this limitation, gene shuffling incorporates a huge number of beneficial

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mutations from multiple genes within a single step of the library creation process [85]. It involves the in vitro recombination of two or more genes to generate an array of shuffled genes that may code for enzymes with desired properties. DNA shuffling performed with similar protein-coding sequences ( . 70% identity) utilizes the naturally occurring genetic diversity of the same class of enzyme and could result in hybrid genes with beneficial mutations [86]. One such hybrid gene could code for a highly efficient enzyme with multiple desired properties such as high activity, thermostability, and enantioselectivity [66]. The workflow of gene shuffling is explained in Fig. 9.3. In this method, the parent genes were digested into several fragments using enzymes such as DNase I or restriction endonucleases to generate fragments of different length followed by unification of random fragments using PCR or blunt-end ligation [87]. These chimeric hybrids can then be cloned into an expression vector and screened for desired function using high-throughput assays.

FIGURE 9.3 Workflow of DNA shuffling for achieving enzyme with desired characteristics.

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9.2.1.2 Semi-rational design Though random mutagenesis has proven successful in many cases, the size of the library is large compared to the number of improved mutants obtained [88
90]. In fact, even with high-throughput screening methods, the screening process is tedious and time consuming for the libraries created by random mutagenesis. Recent advances in bioinformatics and protein crystallography have led to the gradual accumulation of knowledge regarding protein structures and the active sites of enzymes, thus facilitating the rational design of enzyme engineering approaches. This combination of random mutagenesis with rational design has resulted in interesting semi-rational methods for directed evolution, which can bypass some limitations experienced in random and rational approaches [91]. The semi-rational design of directed evolution targets specific residues in multiple positions based on the structural and functional knowledge. The smart libraries obtained by semi-rational design are more likely to yield enzymes with the desired functions while requiring relatively small library sizes compared to random mutagenesis. In this section, we will discuss two of the widely used methods in semi-rational approaches.

9.2.1.2.1 Iterative saturation mutagenesis Iterative saturation mutagenesis (ISM) has arisen as a valuable mutagenesis approach in protein engineering [89]. The first step is to analyze the protein and identify the hotspots that are crucial for achieving the desired characteristics (i.e., improved activity, broad substrate scope, and improved enantioselectivity) [92
94]. The mutation sites are chosen on the basis of structural data, after which each particular site undergoes saturation mutagenesis and is replaced by any of the 19 other possible amino acid residues [89]. The model workflow of ISM is shown in Fig. 9.4 [89]. Saturation mutagenesis is performed on the chosen hotspots (A, B, C. . .) using mutagenic primers during PCR amplification of the target DNA. Cloning and expression of these mutated DNA fragments lead to one or more libraries of enzyme mutants. The libraries are then screened by high-throughput assays and the best hits from each library are identified and sequenced. In the second round of mutation, the best-performing mutant(s) is used as a template and the other unmutated hotspots will be subjected to saturation mutagenesis. This process is repeated until the enzyme with the most desired characteristic is achieved.

9.2.1.2.2 Combinatorial active-site saturation test Combinatorial active-site saturation test (CAST) is a semi-rational mutagenesis method performed with the residues lining the active site of the enzyme [95,96]. In the CAST approach, the key amino acid residues involved in activity and stereoselectivity can be identified. Saturation mutagenesis of these target residues is usually performed in groups of two or three spatially close residues, resulting in relatively small libraries of mutant enzymes that can be easily screened for activity [95,97]. CAST is a very useful technique for enhancing activity, modifying stereoselectivity, and widening the substrate scope [98]. Iterative cycles of CASTing can be performed, in a similar manner as ISM, for enhancing the activity of the obtained mutants [97
99].

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FIGURE 9.4 Workflow of iterative saturation mutagenesis for evolving an enzyme with desired characteristics.

9.2.2 HIGH-THROUGHPUT SCREENING OF ENANTIOSELECTIVE ENZYMES Due to the (semi-)random nature of mutant library construction, large quantities of enzyme mutants are generated, yet only a handful of these may exhibit an improvement in the desired catalytic property. Therefore, it is necessary to use a high-throughput screening assay—a fast, sensitive, and accurate means of screening the enzyme library to obtain the target enzyme. Most high-throughput screening assays rely on the detection of a spectroscopic signal upon reaction, which allows the assay to be carried out rapidly in 96-well microplates using inexpensive spectroscopy equipment [62,100,101]. It is important to note that there is no singular general-purpose high-throughput assay—they often need to be designed specifically for a single target. Some particularly successful assays for screening the enantioselectivity of enzymes are reviewed here.

9.2.2.1 Chromogenic/fluorogenic substrate surrogate-based assays The first assay for the screening of enantioselective biocatalysts used the hydrolysis of para-nitrophenyl (PNP) esters (Fig. 9.5). The product released upon hydrolysis, para-nitrophenol, exhibits UV
Vis absorption at 405 nm in solutions above pH 7. Thus, lipase activity for the hydrolysis of achiral PNP esters can be easily tracked by measuring absorption against time [102]. For chiral PNP esters, however, the kinetic resolution of the racemate only measures the overall reaction rate and not the enantioselectivity; therefore, Reetz et al. used enantiopure (R)- and (S)-esters separately, enabling the determination of the enantiomeric ratio (E) of each lipase mutant. This strategy allowed the screening of enantioselective lipases, and a mutant with vastly improved enantioselectivity was engineered in four rounds of random mutagenesis and screening cycles [103]. Similar

9.2 DIRECTED EVOLUTION: A DARWINIAN APPROACH

R2 O

R1 O

PNP. umbelliferone, or resofurin moeity

R2

Lipase/esterase R1

O

R1 O

PNP. umbelliferone, or resofurin moeity

+

HO

PNP. umbelliferone, or resorufin moeity

O Spectroscopic signal

(S)-acid

(S)-ester R2

OH

153

R2

Lipase/esterase R1

OH

+

HO

O (R)-acid

(R)-ester

PNP. umbelliferone, or resorufin moeity Spectroscopic signal

N HO

NO2 HO PNP λ = 405 nm

O

O

HO

O

Umbelliferone

Resorufin

λex = 330 nm

λex = 325 nm

λem = 460 nm

λem = 425 nm

O

FIGURE 9.5 Chromogenic/fluorogenic enantioselectivity assay via the hydrolysis of enantiopure para-nitrophenyl esters, umbelliferone esters, and resorufin esters.

screening assays involving the use of enantiopure umbelliferone esters or resorufin esters have also been reported, where the quantity of hydrolyzed product umbelliferone or resorufin is measured by fluorescence spectroscopy [104,105]. The problem with such assays is that these esters are usually not of industrial interest, hence an assay screening for high activity on such substrate surrogates may not necessarily yield an industrially useful lipase.

9.2.2.2 pH indicator assays The lipase-catalyzed hydrolysis of an ester produces an acid, which causes the pH of the reaction mixture to decrease. Similar to the approach above, concurrent assays can be conducted using (S)and (R)-esters separately with appropriate colorimetric pH indicators. The color change can be tracked easily using UV
Vis spectroscopy and the E of the enzyme calculated. It is important to note that the pK values of the buffer and the indicator should be within the same range. Some reported pH indicators used in enantioselective hydrolysis screening assays are para-nitrophenol [106] and bromothymol blue [107].

9.2.2.3 Coupled enzyme assays The direct detection of the target enantiomer molecules is a more desirable screening method than those discussed so far. An elegant approach is to use a coupled enzyme assay, where one of the product enantiomers is selectively converted into a secondary product by another enzyme, producing a spectroscopic signal (Fig. 9.6). In one example, an assay was developed for measuring the ee of a secondary alcohol (a potential product of enzymes such as monooxygenases and lipases,

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(A)

Vis absorption λ = 400 nm 2 ABTS cation

(B)

NAD(P)+ OH R1

NAD(P)H + H+

R2

(S)-alcohol

UV absorption λ = 340 nm

(R OH R1

R2

(R)-alcohol

NAD(P)+

R2

O R1

DH )- A

O2 R1

(S)-alcohol

(S ) -A DH

HR P

OH

(S

)-A Ox

H2O2

O

R2

R1

Ketone NAD(P)H + H+ UV absorption λ = 340 nm

(R)

x - AO

OH R1

R2

2 ABTS

R2

Ketone H2O2 2 ABTS

O2

(R)-alcohol

P HR

2 ABTS cation Vis absorption λ = 400 nm

FIGURE 9.6 Coupled enzyme UV
Vis spectroscopic assays for detecting the ee of racemic alcohol solutions with (A) (S)- and (R)-specific alcohol dehydrogenases (ADH) and (B) (S)- and (R)-specific alcohol oxidases (AOx) and horseradish peroxidase (HRP).

among others) by using an (R)-specific alcohol dehydrogenase and an (S)-specific alcohol dehydrogenase separately, to oxidize the respective alcohol enantiomer to a ketone. The individual reaction rates were measured by the formation of NAD(P)H over time using UV
Vis absorption at 340 nm, and the ee of the alcohol could be calculated from the kinetic constants of each enzyme, with only 6 9% uncertainty [108]. Another coupled enzyme assay also measures the ee of secondary alcohols by means of a horseradish peroxidase (HRP) with an (R)- or (S)-specific alcohol oxidase, respectively. The enantiospecific alcohol oxidase converts the corresponding alcohol enantiomer into a ketone and releases H2O2 in the process, which the HRP uses to oxidize a chromogenic precursor such as 2,20 -azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) into a colored product easily measured by UV
Vis spectrometry. In this way, the ee of various secondary alcohol mixtures produced by ketone reductases could be rapidly determined at a rate of 1.7/min [109].

9.2.2.4 Agar plate
based assays Most assays require the random picking of mutant colonies from an agar plate—either using a colony picker or manually—followed by growth and enzyme expression in 96-well plates, before being subjected to high-throughput screening. Agar plate 2 based assays can significantly reduce the number of colonies picked and screened, by associating the desired enzyme characteristic with

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155

an easy-to-recognize phenotype displayed directly on the plate, such as a colorimetric or fluorescent reaction [110]. The tributyrin agar plate assay is a classic method used to measure the hydrolytic activity of esterases and lipases [111]. When tributyrin is added to an agar plate, it results in a turbid emulsion. Therefore, microorganisms producing enzymes with lipolytic activity would be able to hydrolyze the tributyrin, forming clear halos around the corresponding colonies. One example of an enantioselective agar plate-based assay is the oxidation of enantiopure (R)- or (S)-amine by a monoamine oxidase on an agar plate containing the chromogenic precursor 3,30 -diaminobenzidine [112]. The H2O2 produced by the oxidase then reacts with 3,30 -diaminobenzidine to produce a dark brown color. By duplicating the colonies on agar plates containing only the (R)- or (S)-amine, respectively, colonies with high activity toward only one enantiomer could be selected.

9.2.2.5 Fluorescence-activated cell/droplet sorting Fluorescence-activated cell sorting (FACS) has recently been shown to be capable of rapidly screening extremely large enzyme libraries (108
1010). This approach requires the enzyme to be displayed on the surface of the host cell—enzymatic conversion of a chiral chromogenic screening substrate results in the formation of a fluorescent product that binds to the cell membrane. The cells are then sorted to isolate those displaying high fluorescence signal, and the genetic information of the positive enzyme variants is recovered. FACS has been applied for the evolution of enantioselective HRP [113] and lipase [114]. In a similar approach, fluorescence-activated droplet sorting (FADS) was also demonstrated for evolving enantioselective esterase [115]. FADS differs from FACS by achieving compartmentalization and sorting with water-in-oil droplets instead of cells.

9.2.2.6 Other assays Other types of “medium-throughput” assays may be less favored due to their relatively low throughput (102
103) compared to the methods described above; however, these assays may be necessary in niche applications and so should not be ignored. Many detection methods have been reported for enantioselective screening of enzyme libraries, including chiral GC [116], chiral HPLC [117], mass spectrometry with isotope labeling [118,119], nuclear magnetic resonance spectroscopy [120], and surface-enhanced resonance Raman scattering [121]. The interested reader may refer to published literature on the topic [101,122].

9.3 ENZYME ENGINEERING: EXAMPLES IN ENANTIOSELECTIVE BIOCATALYSIS In this section, we showcase a series of successful attempts at using directed evolution to tune the enantioselectivity of a natural enzyme, as a demonstration of the directed evolution techniques discussed in earlier subsections. Several other examples of successful manipulations of enantioselectivity have also been reported on other enzymes, such as with Baeyer
Villiger monooxygenases, lipases, and ADH. In any case, before embarking on any directed evolution attempt, it is important to scan prior literature for any structural and/or mechanistic data to aid in the selection of evolution methodology; perhaps one may even uncover mutations that have already been examined on the enzyme of interest on other substrates. While not essential to a successful directed evolution effort, factors such

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as structural and mechanistic knowledge, moderate base selectivity, and an efficient bacterial expression system would all increase the likelihood for the attempt to achieve desired results.

9.3.1 ENHANCING ENANTIOSELECTIVITY OF P450PYR MONOOXYGENASE FOR THE HYDROXYLATION OF NONACTIVATED CARBON ATOMS Cytochrome P450 monooxygenases are among the most well-studied enzymes by directed evolution and belong to the largest subfamily of monooxygenases reported for asymmetric hydroxylation (RsH - RsOH). Its mechanism is well known: an activated heme iron-peroxo species likely forms a CsO bond after radical H abstraction from the CsH bond. This subsection discusses the directed evolution efforts on a P450pyr monooxygenase from Sphingomonas sp. HXN-200 to enhance its mediocre enantioselectivity on various substrates of interest. One study on the P450pyr enzyme sought to improve the natural enantioselectivity of the enzyme to more industrially acceptable levels, such that an essentially enantiopure product may be achieved using N-benzyl-pyrrolidine as a substrate [123]. The wild-type (WT) enzyme could produce (S)-Nbenzyl 3-hydroxypyrrolidine in 43% ee, which was unsatisfactory, thus the authors embarked on an attempt to modify the enantioselectivity of P450pyr by directed evolution (Fig. 9.7A). In this study, OH (A)

P450pyrTM N Bn

N Bn N-benzyl pyrrolidine

(S)-N-benzyl 3hydroxypyrrolidine 98% ee

(B)

P450pyrSM1 n-Octane

OH (S)-2-octanol >98% ee

(C)

P450pyrSM2 OH Propylbenzene

(S)-1-phenyl-2-propanol 95% ee

FIGURE 9.7 Engineering for P450pyr for enhanced enantioselectivity. (A) Engineered P450pyr for highly S-enantioselective hydroxylation of N-benzyl-pyrrolidine. (B) Engineered P450pyr for highly regio- and enantioselective hydroxylation of n-octane. (C) Engineered P450pyr for highly regio- and enantioselective hydroxylation of propylbenzene.

9.3 ENZYME ENGINEERING

157

amino acid residues were identified based on the X-ray crystallography structure of the protein and 20 residues were identified as hotspots for mutation using the ISM method. In the first round of evolution, a new mutant I83H, which improved the (S)-product ee to 78%—was engineered. In the second round of ISM, the highly promising I83H variant was subjected to mutagenesis on the other 20 residues, and a double mutant (I83H/M305Q) was found to have improved enantioselectivity, giving the product in 94% (S) ee. A third and final round was conducted and a triple mutant, P450pyrTM (I83H/M305Q/A77S), was found to give a remarkable 98% product ee for the (S)-hydroxylated product, while maintaining a similar level of activity as the WT enzyme. Molecular docking of the substrate in the structures of WT and triple-mutant enzymes revealed that the main factor of enhanced enantioselectivity was due to conformational changes that resulted in a comparatively closer distance between the heme oxygen and the pro-S hydrogen in the mutant ˚ ; mutant: 2.4 A ˚ ). One conclusion that can be drawn is that a small number of amino (WT: 2.8 A acid mutations may sometimes result in a significant improvement in the enantioselectivity of an enzyme, if rational choices are made in library design. A similar directed evolution attempt was also carried out for P450pyr on the asymmetric hydroxylation of another nonactivated carbon—the subterminal carbon of an alkane [124]. The natural selectivity of the WT enzyme favored the hydroxylation of the terminal carbon of an alkane, resulting in an achiral terminal alcohol. Hence, the evolution target was not confined to enantioselectivity, but instead included regioselectivity, and the enzyme was ambitiously engineered to perform the subterminal hydroxylation of alkanes in a stereoselective manner (Fig. 9.7B). The first three rounds of ISM were carried out using a model substrate, 4-nitrophenetole, which decomposes into a chromophore upon subterminal hydroxylation. This aided the facile highthroughput screening of mutant libraries generated for the 22 hotspot residues located close to or leading to the substrate-binding site. The resulting best mutant (N100S/T186I/F403I) had moderate regioselectivity (40%) on the subterminal carbon of octane, which proved to be a vast improvement from the WT enzyme. Despite this, the enantioselectivity of the mutant was still poor, as there was no selection bias in the screening assay for the enantiopurity of the product. To better screen for enantioselectivity, a novel coupled enzyme high-throughput screening assay was developed with stereo- and regio-complementary oxidative enzymes, which allowed the use of a sensitive colorimetric detection method for end-point product characterization. For this screening effort, the target substrate (n-octane) could be used instead of a substrate surrogate, significantly improving the assay viability. The usefulness of this assay was apparent immediately as an additional L302V mutation was uncovered in the fourth round that greatly improved regioselectivity, stereoselectivity, and activity. The sequence space was explored a further two rounds with the newly developed high-throughput screening assay, finally yielding a sextuple mutant P450pyrSM1 (A77Q/I83F/N100S/T186I/L302V/F403I). The mutant P450 exhibited complete subterminal selectivity ( . 99%) and excellent stereoselectivity (98% ee) in the hydroxylation of n-octane to (S)-2octanol, while still retaining 90% of its activity compared to the WT enzyme. Further kinetic studies showed that the catalytic efficiency (kcat/Km ratio) of the sextuple mutant was nearly identical to the WT enzyme. The observant reader might notice some commonly mutated residues between these two examples, namely A77, I83, N100, T186, and F403. It may be the case that these residues are positions that affect the shape of the binding pocket in such a way that enantioselectivity may be alerted significantly, no matter the substrate. Thus, it could be interesting for the protein engineer to test

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multiple substrates with the most promising mutants obtained in previous directed evolution efforts, with the hope that these binding pocket changes would be universally beneficial in obtaining a higher stereoselectivity across a broader substrate range. One can observe this idea of testing previous successful mutants in several instances. In the same study that produced the sextuple P450pyr mutant, the authors repeated the ISM procedure on a different substrate, propylbenzene, using a more focused library with the best mutants found in the previous evolution. The result was another sextuple mutant P450pyrSM2, with identical mutations except for L251V instead of A77Q. P450pyrSM2 could perform the subterminal hydroxylation of propylbenzene to give (S)-1-phenyl-2-propanol in 95% ee and 98% subterminal selectivity (Fig. 9.7C). We can also observe this universality of enantioselectivity in the triple mutant which catalyzed the (S)-hydroxylation of N-benzyl-pyrrolidine, introduced at the beginning of this subsection. P450pyrTM exhibited high regio- and enantioselectivity on the hydroxylation of other substrates, such as N-benzyl-pyrrolidinone and N-benzylpiperidinone [125]. Surprisingly, P450pyrTM also proved to be useful for carrying out a different enantiospecific reaction—the asymmetric epoxidation of para-arylalkenes. The triple mutant generated for the (S)-hydroxylation of N-benzylpyrrolidine showed apparent reversed enantioselectivity for para-arylalkenes, with enhanced (R)epoxidation of para-substituted styrene compounds observed compared to the WT enzyme [126]. Therefore, mutants generated from one study evolving enantioselective enzymes suited for one reaction may be transferrable to a different reaction entirely, highlighting the importance of prescreening any “hotspot” mutations discovered in prior studies to avoid duplicate discovery. After obtaining an enantioselective mutant, adjustments to the reaction system can be applied to optimize for better enantioselectivity. Further use of the mutant P450pyr enzymes was demonstrated in a sulfoxidation system, where the previously evolved triple mutant was deconvoluted, with the single mutant I83H resulting in the highest product ee of 85% of the (R)-sulfoxide when coupled with a glucose
dehydrogenase-based cofactor recycling system [127]. When the system was then modified to a biphasic aqueous/ionic liquid (IL) system, enantioselectivity improved to greater than 95% ee of the (R)-sulfoxide, with much better substrate tolerance. It was later proposed that the improvement of ee from the biphasic system was the result of having a low concentration of substrate present within the aqueous phase, which could be correlated with higher product ee. In some cases, when enzyme engineering does not produce a satisfactory result, it could be worthwhile to explore methods for enhancing enzyme enantioselectivity with reaction engineering to improve product ee.

9.3.2 INVERTING THE ENANTIOSELECTIVITY OF P450PYR MONOOXYGENASE Another study on engineering enantioselectivity of P450pyr details an ambitious attempt to completely invert the enantioselectivity of the enzyme for the asymmetric hydroxylation of Nbenzyl-pyrrolidine to (R)-N-benzyl 3-hydroxypyrrolidine (Fig. 9.8) [128]. A principal focal point of the research effort was the development of an efficient high-throughput screening method for the detection of product ee, thus reducing the screening effort, a step which could potentially be a major bottleneck. The use of two stereo-complementary ADH coupled with a colorimetric NBTformazan assay provided a quick and reliable detection method for ee, without the need for chiral chromatographic separation.

9.3 ENZYME ENGINEERING

P450pyr mutant with inverted stereoselectivity N Bn N-benzyl pyrrolidine

159

OH N Bn (R)-N-benzyl 3hydroxypyrrolidine 83% ee

FIGURE 9.8 Inversion of stereoselectivity of P450pyr by directed evolution for the R-hydroxylation of N-benzyl pyrrolidine.

˚ of the docked substrate For this directed evolution study, 17 residues were identified within 5 A to serve as hotspots for the first round of ISM. In this first round of evolution, two important mutants were uncovered: one with improved (S)-selectivity (F403L, 65% ee), and one exhibiting (R)-selectivity (N100S, 42% ee). While the improved (S)-selective mutant did not yield any further noteworthy double mutants in the second round, the (R)-selective template produced multiple improved double mutants, of which the best mutant retained its regioselectivity while being highly (R)-selective (N100S/T186I, 83% ee). Analysis of the homology model of the mutant suggested that conformational changes at the entrance of a cavity allowing substrate access to the binding pocket (caused by mutations at the 100 and 186 positions) resulted in the inversion of enantioselectivity. It is noteworthy that a single point mutation in the P450pyr enzyme could invert enantioselectivity and serve as a basis for more successful mutants to be evolved, thereby demonstrating the potentiality of directed evolution to affect a gradual inversion of enantioselectivity.

9.3.3 ENGINEERING AMINE DEHYDROGENASE FOR ASYMMETRIC REDUCTIVE AMINATION One approach to engineering an enantioselective enzyme is to repurpose a highly enantiospecific enzyme by expanding or redesigning its substrate scope to include nonnatural substrates. The importance of chiral amines in the pharmaceutical industry has propelled the development of enzymes for the asymmetric synthesis of amines. Asymmetric amine functionalization from a prochiral carbonyl-based compound is particularly attractive, and this reaction can be achieved by a variety of different classes of enzymes. Amine dehydrogenases (AmDHs) are a newly developed class of enzymes that have been engineered from amino acid dehydrogenases by directed evolution (Fig. 9.9). These AmDHs offer distinct advantages over their transaminase counterparts, such as the ability to use free ammonia as the amine donor and the availability of an efficient cofactor recycling system to drive the reaction to completion. This subsection discusses some of the early work on engineering enantioselective AmDHs by expanding the substrate scope of natural enzymes. The first example of evolving an AmDH involved the re-orienting of substrate scope of Bacillus stearothermophilus leucine dehydrogenase (LeuDH) to accept ketones without the carboxylic acid

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Natural stereoselective enzyme for L-amino acid production Amino acid dehydrogenase

O R

OH

O R

O

OH NH2

Keto acid

L-amino acid

Engineered enzyme with new substrate scope for (R)-amine production Amine dehydrogenase

R O Ketone

R NH2 (R)-amine

FIGURE 9.9 Engineering of an amino acid dehydrogenase into an amine dehydrogenase for chiral amine production.

group [129]. A modified CASTing procedure was adopted, loosely following ISM principles. Based on the homology model of the enzyme, the researchers identified residues that were close to the active site, with additional attention given to residues that were close to the carboxyl moiety. Rounds of mutant libraries were created with degenerate codons, some of which were multisites that included previously mutated residues, to discover any possible synergistic effects. The final enzyme was a quadruple mutant K68S/E114V/N261L/V291C, which exhibited measurable reductive amination activities on several ketone compounds. The choice of high-throughput screening assay has also posed some key challenges for AmDH evolution. The amination reaction involves the oxidation of NADH to NAD1; however, it is preferable to measure the formation of NADH rather than its consumption for technical reasons. As a result, AmDH evolution efforts often use the reverse reaction—the deamination of a ketone—as a screening method so as to produce NADH, which can then be easily tracked. Following this success, a follow-up study was conducted which aimed to transfer the LeuDH directed evolution data to the engineering of Bacillus badius phenylalanine dehydrogenase (PheDH) [130]. Sequence alignment on the two dehydrogenases identified the analogous residues responsible for substrate specificity. Through screening of various mutants, it was found, rather logically, that the two hotspots responsible for the substrate specificity were the lysine (K77) and asparagine (N276) residues that closely interacted with the carboxyl group of the substrate. Since there was evidence of the synergistic effect of mutating both sites simultaneously, a two-site DDK library was generated on K77 and N276, with the best mutants selected for detailed characterization. The resulting double mutant K77S/N276L proved to be highly enantioselective [ . 99.8% ee (R)] and could be driven to high conversion when conducting the asymmetric amination of parafluorophenylacetone. It is apparent from the evolution of this AmDH that any successful mutations from a prior evolution attempt may be successfully applied to engineer novel enantioselective enzymes from homologous templates, allowing a good combination of directed evolution and rational design techniques to be applied.

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Indeed, it was also found through another study of the evolution of Rhodococcus PheDH that despite only sharing 32% identity with B. badius PheDH, the analogous lysine and asparagine residues were important for the carboxyl selectivity [131]. In the first round of two-site NNK mutagenesis, several positive clones were identified, with best-performing double mutants being K66Q/N262L and K66Q/N262C. When the individual mutations were deconvoluted, it was found that K66Q was the key enabling mutation, while the additional mutation on N262 provided positive epistatic effects. It was later confirmed through molecular docking that the mutation of K66Q allowed the carbonyl group access to the catalytic sites. A final round of NNK single-site mutagenesis then resulted in a more active triple mutant K66Q/S149G/N262C, which could catalyze the reductive amination of phenylacetone and 4-phenyl-2-butanone to afford the respective chiral (R)-amines in .98% ee. These examples demonstrate the expansion of substrate scope of naturally stereoselective enzymes by increasing their substrate scope, allowing the enzyme to tolerate the absence of a moiety that would otherwise be necessary for the enzyme to carry out its function. The engineering of the AmDH class of enzymes shows the effectiveness of combining rational techniques with the power of directed evolution to access a larger reaction space in enantioselective biocatalysis.

9.4 CONCLUSIONS AND PERSPECTIVES This chapter gave an overview of the emerging importance of biocatalysis for chemical synthesis, and how the directed evolution of enzymes can play an essential role in developing new biocatalysts for enantioselective biotransformations. Various methods for mutant library generation including random and semi-rational approaches were described and a number of recently developed enantioselective high-throughput screening assays were reviewed. The considerations for choosing the most appropriate methodologies were discussed and should provide a useful basis for aspiring enzyme engineers. Several accounts of recent and excellent examples of enzymes engineered for enantioselective biotransformations were highlighted, such as P450 monooxygenases for asymmetric hydroxylation and amine dehydrogenases for asymmetric reductive amination. The incomplete understanding of the relationship between enzyme structure and function has so far excluded us from accessing the vast realm of unexplored enzyme space. Going forward, advanced computational tools such as artificial intelligence will play an increasingly important role in the prediction of enzyme structures and their corresponding traits and functionalities, allowing for the more accurate and rational design of artificial enzymes. Until then, directed evolution continues to be the tool of choice for modifying enzymes; thus the search for ever more effective high-throughput screening assays will continue. In particular, the use of microfluidic platforms for ultrahigh-throughput cell and droplet sorting shows great potential for the expanding field of directed evolution.

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3124. [125] S.Q. Pham, P. Gao, Z. Li, Engineering of recombinant E. coli cells co-expressing P450pyrTM monooxygenase and glucose dehydrogenase for highly regio- and stereoselective hydroxylation of alicycles with cofactor recycling, Biotechnol. Bioeng. 110 (2) (2013) 363
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11574. [127] P. Gao, et al., Enhancing enantioselectivity and productivity of P450-catalyzed asymmetric sulfoxidation with an aqueous/ionic liquid biphasic system, ACS Catal. 4 (10) (2014) 3763
3771. [128] W.L. Tang, Z. Li, H. Zhao, Inverting the enantioselectivity of P450pyr monooxygenase by directed evolution, Chem. Commun. 46 (30) (2010) 5461
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3972. [130] M.J. Abrahamson, J.W. Wong, A.S. Bommarius, The evolution of an amine dehydrogenase biocatalyst for the asymmetric production of chiral amines, Adv. Synth. Catal. 355 (9) (2013) 1780
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1122.

CHAPTER

NANOBIOCATALYST DESIGNING STRATEGIES AND THEIR APPLICATIONS IN FOOD INDUSTRY

10

Madan L. Verma1, , Reinu E. Abraham2, and Munish Puri2 1

Department of Biotechnology, School of Basic Sciences, Indian Institute of Information Technology Una, Una, Himachal Pradesh, India 2Centre for Marine Bioproducts Development, College of Medicine and Public Health, Flinders University, Adelaide, SA, Australia

10.1 INTRODUCTION Enzymes, also termed as “biocatalyst,” have been playing a considerable role in shaping the research and development in food industry [1 4]. Enzymes are considered as “Generally Recognized As Safe (GRAS)” biological entities that are used either as ingredients or processing aids in various food processing applications. Based on a recent report released from the Global Market Insights Inc., food enzymes market size will surpass $3.6 billion by 2024 [5]. These encouraging trends warrant research effort that may improve upon stability and cost of the enzyme, which is a major concern for industrial application. The downstream processing during enzyme purification requires multiple steps that can enhance the cost of the purified enzymes. Moreover, purified enzymes are prone to denaturation as compared to in vivo form of enzymes due to harsh physicochemical environment at the industrial setting. The inherent property of the biocatalyst(s) is to work in mild conditions that hamper their wider applications. Thus, the use of enzyme-based bioprocesses is generally not favorable due to the higher cost and ease of instability of the biocatalyst [6 8]. However, nanotechnology is playing an important role in providing stability and costeffectiveness in the enzyme-mediated food processes [9]. Recently, nanotechnology has significantly contributed to the biotechnological processes that have applications in diverse fields ranging from food to bioenergy sectors [10 13]. Nanotechnology refers to the synthesis of the nanomaterials in the range of 1 100 nm and its potential applications. Synthesis of the nanomaterial can be achieved by the top-down and bottom-up approaches [14 18]. Nanomaterial synthesis can be achieved either by physicochemical or biological methods, whereas biological method is finding more preference. Green chemistry mediated biological process leads to negligible by-product without using any harsh chemicals.



Both authors have contributed equally in the write-up.

Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00010-7 © 2020 Elsevier B.V. All rights reserved.

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Nanomaterials occupy a prominent position in the biological system due to inherent multitasking superior functions [19 26]. Nanomaterial forms such as nanoparticle, nanofiber, nanotube, and nanocomposite are used as nanocarrier for biocatalysts. Nanocarrier-bound biocatalyst, referred to as nanobiocatalyst, can be developed using tailor-made nano-immobilization methods [27,28]. Nanomaterials can load a higher amount of enzymes due to higher surface area-to-volume ratio. Nanomaterial-loaded enzyme possesses minimum mass-transfer resistance that makes majority of biocatalytic processes more efficient. The surface of nanomaterial can be quite easily tailored with a plethora of functionalized agents. Thus, one type of nanomaterials can be used for immobilisation of a variety of enzymes based on tailoring the surface chemistry for the specific nanomaterial. Ease of purification of the nanomaterial-bound enzyme facilitates its cost-effective downstream processing [29 33]. Magnetic nanomaterials or their composites can be quickly separated from the product under the influence of external magnetic field. The present chapter discusses the types of nano-immobilization methods and various strategies employed for the development of nanobiocatalyst. Application of nanobiocatalysts in food processing is also discussed. Role of nanobiocatalyst in food processing industry with reference to omega-3 fatty acid processing is elaborated.

10.2 DESIGN OF NANOBIOCATALYST THROUGH NANO-IMMOBILIZATION Enzyme immobilization is a well-established technology that is employed for various applications in various industries [12 18]. The term “immobilized enzyme” was first accepted in the Enzyme Engineering Conference in 1971. The meaning of immobilization as a word is confinement/localization of biocatalyst in a defined region of space or movement limiting as binding of free enzymes to supports limits their mobility [34 41]. Enzymes have a plethora of applications in improving the food quality ranging from bioactive extraction to fortified nutraceutical delivery [42 44]. Enzymes can be isolated from diverse sources such as animals, plants, and microorganisms. However, enzymes sourced from microbes are the preferred choice due to ease of production and scale up at industrial setting. Some of the applications of free enzymes in food industry are listed in Table 10.1. Enzymes mostly work in a mesophilic range that can be prone to denaturation [45,46]. Such fragile biocatalyst moiety can be protected using advanced functional nanomaterials [47]. Thus, enzyme protection from external harsh environment conditions is achieved using enzyme immobilization methods (Fig. 10.1). Enzyme immobilization provides operational stability and reusability that make any bioprocess cost-effective. Enzyme immobilization techniques are classified further based on the nature of interaction between enzyme and the carrier (support) [48 51]. Enzyme could be immobilized by physical adsorption, covalent binding, cross-linking, and entrapment methods (Fig. 10.2). However, each and every technique has its own advantages and disadvantages. Physical adsorption is the most simplest and cost-effective method where enzyme and support interact through weak forces. Enzyme leaking is the problem associated with this method. The covalent immobilization methods involved strong bonding between enzyme and support that can facilitate several times recycles of enzymes without loss of biocatalytic function. Cross-linking is a form of covalent

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Table 10.1 List of commonly used enzymes in food industry and their potential applications. Enzyme

Polysaccharide

Industrial application

References

α-Amylase

Starch, glycogen

[5]

α-Acetolactate decarboxylase α-Galactosidase, β-galactosidase β-Fructofuranosidase/ invertase Cellulase

Pyruvate

Cellulose

Catalase

Hydrogen peroxide

Esterase

Esters

Glucose oxidase

Glucose

Glucoamylases

Cleaves 1,4α-glycosidic bonds from glycosidic chains

Glucanase

β-Glucans

Hemicellulase

Hemicellulose

Isomerases Naringinase

Glucose isomerization to fructose Hydrolysis of naringin

Pectinases

Pectin

Pullulanase

Pullulan

Xylanase

Xylan/hemicellulose

Baking, starch liquefaction such as in paper industry (to improve the property of starch—to make low viscosity, high molecular weight), brewing alcohol, clarification of fruit juice Dairy industry (flavor and aroma in dairy product) Prebiotic food ingredients, producing hydrolyzed milk products Production of fructooligosaccharides, production of sugar syrup Extraction of olive oils in combination with pectinase, xylanase, improve the quality and food product’s shelf life, beverage industry such as fruit juice for breakdown of cell wall Food processing of dairy products such as cheese, sourdough, brewing industry: beer, wine, and vinegar Producing flavoring and fragrance in food processing, improve flavor in sausages Used in foods and beverages to remove residual glucose and oxygen such as in egg powder, used in baking industry to improve crumbing property of bread and croissants Saccharification of starch or dextrin into glucose: beer production Improving quality of bread Textile and pharmaceutical industries Sugar syrup production such as high glucose and high fructose syrups Prebiotic sausage, gluten-free bread Baked, rice starch bread formulated using yeast glucan Baking industry: break down the hemicellulose in wheat flour Production of high-fructose corn syrup, sweeteners Debittering enzyme commercially used for citrus juice Preparation of functional foods and dietary fibers Starch-based industries for saccharification of starch, production of high-maltose, and fructose-rich corn syrup Dough conditioning

β-Galactosides Sucrose

[6] [7] [8] [9]

[10]

[11] [12]

[13]

[14]

[15] [16] [17] [18] [19]

[20]

FIGURE 10.1 Types of nano-immobilization methods employed for the development of nanobiocatalysts.

FIGURE 10.2 Schematic representation of developing a nanobiocatalyst, enzyme-nanomaterial bioconjugate.

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175

binding where an enzyme is linked to another enzyme by covalent bonding without using supports. This is often referred to as cross-linked enzyme aggregates (CLEA). Entrapment involves trapping the enzyme inside a support; however, this technique is limited by enzyme leakage and substrate diffusion. Based on the pros and cons of enzyme immobilization methods, enzyme immobilization is followed using two approaches, namely physical adsorption and covalent binding [28 32]. Like the conventional method of enzyme immobilization technology, nano-immobilization is also categorized into four types based on the interaction between the nanocarrier and a biocatalyst [52 57]. Such methods are primarily adsorption method, covalent method, entrapment method, cross-linked method, and a combination of two methods such as adsorption followed by crosslinked method.

10.2.1 ADSORPTION BINDING OF ENZYMES Adsorption method employs a simple and reversible process that binds the enzyme to the support material through weak forces such as hydrophobic and van der Waals interactions [58 63]. The enzymes can be temporarily bound to the nanocarrier using the physical stimuli such as ionic strength, temperature, and pH. It is a quick process that is achieved simply by mixing aqueous solution of the enzyme with the nanocarrier. At optimized reaction conditions, the excess enzyme from the immobilized enzyme on the nanomaterial is removed through extensive washing with distilled water [64]. Performance of the adsorption method is considerably improved by tailoring the key factors of the enzyme-dissolved buffer, such as the pH and the ionic strength and surface properties of nanocarrier such as pore size, shape, and nature of hydrophilicity and hydrophobicity. This method is the first priority to employ for studying the enzyme and nanocarrier interaction due to ease of its simplicity, and free of chemical additives [65]. The other salient feature of adsorption method is the quick recovery of the costly nanocarrier due to ease of reversibility. For example, costly amyloid nanocarrier can be easily used and recovered by using the reversible enzyme immobilization method. However, the major concern with this method is the long-term stability and recyclability of the nanocarrier-bound enzyme due to leaching of the weakly bound enzymes from the nanocarrier under harsh reaction condition. This limitation of excessive enzyme leaching from the support can be controlled by using a combination of adsorption followed by covalent method or cross-linker method. Nanobiocatalyst is developed using adsorption method for achieving the desired product. For example, lipase sourced from Candida rugosa immobilized on surfacemodified magnetite nanoparticles for quick bioseparation and reuse [66]. Lipase from Burkholderia sp. was also immobilized on surface-modified ferric silica nanocomposite through physical binding. As a result, only 29 mg /g lipase was bound to the nanocarrier [67]. The nanobiocatalyst improved the thermal stability and the product formation reduced to approximately half the yield after eight reuses [68]. Researchers found that the adsorption entrapment technique was found to be more effective than surface adsorption [69]. Lipase sourced from C. rugosa, CRL, was also immobilized using peptide nanotubes, through hydrogen bonding between amide groups of the nanotube and complementary functional groups on the surface of enzymes. The nanobiocatalyst exhibited higher biocatalytic potential than free lipases [70]. In general, this physical binding method is known to achieve weak interactions between the enzyme and the nanocarrier, even though the ionic binding method is generally stronger than other interactions [71].

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10.2.2 COVALENT BINDING OF ENZYMES This method is mostly employed to achieve cost-effective application of immobilized enzyme at industrial setting [72 74]. Covalent binding is a typical and effective immobilization method that involved covalent binding between the nanocarrier and enzyme of interest [75 80]. It involves formation of covalent bonds between a functionalized nanocarrier and the active amino acid residues of the surface of the enzyme [81 84]. Several types of nanocarrier have been used, such as nanopores, nanoparticles, or nanofibers to immobilize the enzymes through covalent binding [85 88]. The main advantage of the covalent binding is the interactions between lipase and nanocarrier that prevent enzyme leaching even after repeated use [89 92]. Recently, researchers reported a technique for immobilization of CRL on magnetic chitosan microspheres for transesterification of soybean oil [93]. As a result, the nanobiocatalyst was determined as an effective biocatalyst for the conversion of soybean oil. It was shown to retain high original activity during multiple repetitive transesterification cycles [93]. Two different types of lipase immobilization were employed to effectively synthesize the biodiesel production [94]. Enzyme namely β-glucosidase was immobilized on polymer nanofibers by glutaraldehyde treatment onto covalently attached to achieve high stability and enzyme activity [95]. Lipases sourced from Rhizopus oryzae and C. rugosa were covalently bound to silica, which was used for the biodiesel conversion from crude canola oil. The substrate conversion rate of degummed crude canola oil to fatty acid methyl esters was achieved higher as compared to the free enzyme [96]. The rare limitation of covalently bound enzymes is a lower biocatalytic activity than the native enzymes due to blockage/altered original active site of the enzyme during immobilization reaction. Researchers employed silica nanoparticles as support material for laccase immobilization and reported an easy recovery of product [97].

10.2.3 ENTRAPMENT OR ENCAPSULATION OF ENZYMES Entrapment method involves enclosing a biocatalyst into polymeric or inorganic network [98,99]. It retains the biocatalyst with ease of substrate permeation inside the gel and consequently moves the product out is achieved simultaneously [100]. This method employs the mixing of enzyme aqueous solution and polymer solution together followed by cross-linking the polymer to form the lattice structure for entrapping the enzyme [101 108]. The entrapment method is widely used for industrial applications due to the protective environment for biocatalyst inside the gel under the mild reaction conditions [109,110]. This method has a wider application and is further categorized into three categories: gel entrapping, fiber entrapping, and microencapsulation. For example, a wide variety of enzymes has been used for enzyme immobilization in sol gel matrices [109,110]. The sol gel-entrapped biocatalyst was found to have improved stability than the lipase immobilized by adsorption on the same material [111]. It is probably due to enzyme leakage from the adsorbed preparation under the provided reaction conditions. Lipase sourced from Candida antarctica was entrapped inside the thermosensitive polymer at higher temperature, causing shrinkage of the polymer. The shrinkage of the polymer was reversible, so enzyme leaked out of the particles. A significant degree of enzyme inactivation was reported as compared to the enzyme entrapment in the low temperature [112]. The limitation of this entrapment method is confined to mass transfer and low enzyme loading due

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to the act of the support as barrier. The nanoencapsulation or microencapsulation creates an insoluble polymeric matrix for enzyme entrapment [108]. Alginate microbeads or microspheres have been used in the enzymatic reactor process. As a result, mechanical stability limits their use to few applications [108]. Nanoencapsulation is holding a great promise in food industry [104,106,107]. It delivers the fortified value-added products through this method. Nanocarrier such as nanopore gold and single/ multiwalled nanotube is frequently employed to retain the biocatalyst through these immobilization techniques.

10.2.4 CROSS-LINKING OF ENZYMES Nanotechnology has leveraged the conventional carrier-free immobilization method, that is, crosslinking method. Recently, application of nanocarrier in particular to magnetic nanoparticles plays a vital role in the development of magnetic cross-linked enzyme aggregate for quick bioseparation and efficient bioprocessing [113 120]. The cross-linking method involves covalent interaction of one enzyme molecule to other enzyme molecules through a suitable cross-linker [117,121]. The advantage of cross-linking enzyme is obtaining a stable nanobiocatalyst with better reusability due to the strong covalent bonding between the nanocarrier and the biocatalyst [118,119,122,123]. The most commonly employed cross-linker, glutaraldehyde, is employed for CLEA. Magnetic nanoparticle functionalized with amino group was employed to develop a novel magnetic CLEA [120,124]. Magnetic CLEA retained more activity and stability than CLEA. Magnetic CLEA of glucoamylase was developed using a novel cross-linker of dialdehydic pectin [121,125]. The use of the most commonly employed cross-linker glutaraldehyde was compared with that of the dialdehydic pectin cross-linker. However, magnetic CLEA exhibited 22-fold higher biocatalytic activity than CLEA. Lipase sourced from C. rugosa was also immobilized on glutaraldehyde-activated poly(vinyl alcohol-co-ethylene) nanofibers through covalent bonds with the aldehyde groups. This method yielded higher enzyme activity due to higher loading of enzymes [122,126]. Biocatalyst immobilization on nanomaterials using glutaraldehyde as a cross-linker is a common method of covalent binding [123,127]. It provides higher enzyme activity due to higher enzyme loading capacity. It is reported in most of the studies that the increment of cross-linker initial concentration induced the higher enzyme loading that led to decrement in the biocatalytic activity of the immobilized preparation [122,126]. In addition, glutaraldehyde and its derivatives cause environmental concerns [124,125,128,129]. Other immobilization chemistries have been developed. In one approach, the nitrile groups of the polyacrylonitrile nanofibers were activated by an amidination reaction followed by reaction with lipase in phosphate buffer solution [126,130]. Activated nanofibers showed higher enzyme loading efficiency (43 mg lipase/g fibers) and retained 80% original activity of lipase as compared with free enzyme, which is much higher than is the typical case for glutaraldehyde. Lipase can also be covalently attached to multiwalled carbon nanotubes through carbodiimide activation. It resulted in improved stability at high temperatures [127,131]. A study was conducted on the preparation of Pseudomonas cepacia lipase CLEA [132]. Authors reported that the crosslinked lipases have a greater stability than the free enzymes in denaturing conditions. The enzyme was found to catalyze the conversion of fatty acid methyl esters from butter tree (Madhuca indica)

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oil, where transesterification is difficult by chemical routes due to its high free fatty acid content. The conversion obtained after 2.5 h was 92%. Researchers investigated a modification procedure for preparation of cross-linked lipase of Geotrichum sp. [129,133]. The obtained cross-linked lipase exhibited improved pH and thermal stability as compared to free lipase. The relative biodiesel yield obtained was 85% through transesterification of waste cooking oil with methanol [129,133]. However, enzyme immobilization method may cause alteration in enzyme structure, in particular to active site of enzyme that leads to poor interaction of enzyme substrate molecules, rendering poor biocatalytic activity.

10.3 NANOBIOCATALYST ROLE IN FOOD PROCESSING WITH REFERENCE TO OMEGA-3 OIL PROCESSING Recently, few studies have been documented using nanobiocatalyst for enrichment of omega-3 fatty acids (Table 10.2). The fish oil hydrolysis for selective fatty acid enrichment is discussed in this section. Researchers demonstrated the fish oil hydrolysis for omega-3 fatty acids using magnetic nanoparticle-bound lipase [130,134]. Nanobiocatalysts exhibited higher selectivity for docosahexaenoic acid (DHA). However, nanobiocatalyst showed the same degree of selectivity for eicosapentaenoic acid (EPA) as by the free enzyme. Recently, lipase sourced from recombinant Bacillus subtilis was employed for nanoparticle immobilization using covalent method [134]. Nanobiocatalyst exhibited good biocatalytic activity for fish oil hydrolysis. The optimized covalent binding method provided higher enzyme loading efficiency of up to 95%. The nanoparticle enzyme interaction after immobilization studies showed minimum alteration in the biocatalyst structure as revealed through electron microscopy and spectroscopy studies. The operational stability of nanobiocatalyst improved twofold and reusability up to 20 cycles with loss of 50% original biocatalytic potential immobilized Table 10.2 Nanobiocatalyst application in fish oil hydrolysis for omega-3 fatty acids enrichment. Nanomaterial type

Enzyme source

Nano-immobilization method

Application

References

Magnetic nanoparticle

Lipase from Thermomyces lanuginosus Lipase from Bacillus subtilis

Covalent method

[132]

Lipase from Thermomyces lanuginosus Lipase from Yarrowia lipolytica

Physical method

4.4-fold DHA content higher compared to free enzyme 1.5-fold DHA higher as compared to free lipase 1.6-fold DHA higher as compared to free lipase 90% higher DHA content as compared to free enzyme

Magnetic nanoparticle

Carbon nanotube

Magnetic nanoparticle

Covalent method

Covalent method

[130]

[131]

[91]

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179

on magnetic nanoparticles by a facile covalent method for fish oil hydrolysis. The biocatalytic potential of immobilized lipase was compared with free enzyme and biochemically characterized with respect to different parameters such as pH, temperature, substrate concentrations, and substrate specificity. Fish oil hydrolysis for omega-3 fatty acids enrichment by immobilized thermostable lipase was further improved. Nanobiocatalyst has selective hydrolytic action to DHA as compared to EPA. It exhibited 1.5 times higher DHA selectivity than free enzyme. However, nanobiocatalyst and free enzyme showed similar hydrolytic selectivity toward EPA. Robust nanobiocatalyst by immobilizing lipase sourced from Thermomyces lanuginosus on the surface of magnetic nanoparticles was developed [131,135]. Nanobiocatalyst showed a higher protein loading of 4.4 mg of enzyme per gram magnetic nanomaterial support. The role of water content improved the biocatalytic potential. At 50% water content, nanobiocatalyst improved the selective enrichment of DHA fish oil up to 4.4-fold the initial content of DHA present. Nanobiocatalyst retained the 82% original catalytic potential even after six recycles of fish oil hydrolysis. A facile methodology for the preparation of robust nanobiocatalyst for fish oil hydrolysis was developed using carbon nanotube carrier [132,136]. Authors employed carbon nanotube support for physical adsorption of lipase sourced from T. lanuginosus. A higher enzyme loading of 6 mg per support was achieved. Carbon nanotube-bound lipase showed 1.6-fold selective DHA enrichment of initial fish oil at optimized 50% water content. Nanobiocatalyst retained 80% operational stability even after the sixth reusability. Lipase sourced from Yarrowia lipolytica was immobilized on the surface of magnetic nanoparticle in a reverse micelles system [95]. Surface-functionalized nanoparticle was achieved by silanization agent, an aminopropyl triethoxysilane. Nanobiocatalyst was further characterized by circular dichroism spectroscopy that revealed an increase in alpha helix and a decrease in beta-sheet content after enzyme immobilization. The resulted nanobiocatalyst was quite active in the reverse micelles system as compared to aqueous system. The nanobiocatalyst exhibited 90% DHA enrichment of the fish oil and was quite robust after reuse of 20 cycles for real substrate hydrolysis.

10.4 APPLICATION OF NANOBIOCATALYST IN OTHER FOOD PROCESSING APPLICATIONS Applications of enzymes in industry setting are well documented. Enzymes namely lipase, trypsin, amylase, glucose isomerase, papain, pectinase, and naringinase have been employed in the food industries for many decades [133 135]. Various products ranging from glucose, high fructose corn syrup, prebiotics, juice debittering, and clarification have been achieved through industrial enzymes immobilized using bulk materials [136 138] but not on nanomaterials. Food safety is a major concern in the food industry. Notably, the GRAS microorganisms and their products have been used in the food industry [3,139]. Additionally, food grade materials have been used as carrier in the food industries. The most used nanomaterial has been documented as a safe material in the in vivo and in vitro studies [140]. The magnetic nanoparticles were employed for stabilization of two enzymes namely cellulase and pectinase and further applications in pigment extraction from the orange peel,

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waste material of Nagpur mandarin orange [141]. The size of nanoparticle in the range of 40 90 nm was surface-functionalized using aminopropyl triethoxysilane. The enzymes were covalently immobilized on the surface with the help of a glutaraldehyde cross-linker. At the optimal reaction condition of pH 5.0 and temperature 50 C, nano-immobilized enzymes achieved ninefold higher carotenoid pigments extraction than free enzyme. Robust nanobiocatalysts retained the 85% original activity even after the three recycles of real substrate hydrolysis. Graphene nanosheet was used as a nanocarrier for α-galactosidase sourced from chickpea [142]. Authors employed response surface methodology for optimization of the enzyme immobilization. The nanocarrier-bound enzyme showed better affinity than the native enzyme. It had one-third lowered Km value as compared to soluble enzyme. Nanobiocatalyst was employed to hydrolyze the raffinose family oligosaccharide, the main cause of flatulence in soybean-derived foods. Nanobiocatalyst was stable even after six repeated usage with real substrate hydrolysis and authors demonstrated a potential application of this study in food processing industries. β-Galactosidase sourced from chickpea was covalently immobilized using functionalized graphene nanosheet [143]. Higher enzyme immobilization efficiency was achieved using optimizing the reaction condition through the statistical approaches of response surface methodology. Nanobiocatalyst was characterized by electron microscopy and infrared spectroscopy techniques to investigate the correlation at the structure function level. Nanobiocatalyst exhibited the same activation energy and temperature optimal value as compared to free enzyme. Authors reported enhanced thermal stability as well as excellent operational stability. Nanobiocatalyst showed more than 90% retention of original activity even after ten reusable cycles. β-Galactosidase sourced from pea plant was covalently immobilized onto the surface of gold nanoparticle [144]. The developed nanobiocatalyst was employing for sensing the lactose content in food products. The spacer arm namely cysteamine-glutaraldehyde was inserted on the surface of nanoparticle. Enzyme was covalently bound through this spacer arm. Authors reported an enhancement of catalytic activity and operational stability up to 6 months. Nanotechnology has developed techniques that can enhance the activity of enzyme to work at extreme temperatures, chemical condition while maintaining the necessary stability, and activity. The use of nano-immobilized enzyme in industrial setting is evolving; authors could not find any reference in the literature; thus this is a huge opportunity.

10.5 CONCLUSIONS AND PERSPECTIVES Applications of nanomaterials as biocatalysts carrier hold a great promise in food industries in the near future as this can address challenges such as stability of enzyme, biocatalytic processing and engineering, and recovery in downstream processing. Advancement in enzyme engineering modalities for discovery of the high turn-over biocatalyst of food industrial relevance is also the main research and development activity all over the world to meet the ever-growing population demand for quality food requirements. Design and application of a suitable nanobiocatalyst to meet commercial feasibility that employs less use of water and energy are the need of the hour in the food industry. It is clearly demonstrated from the above-discussed studies that nanobiocatalyst is the key player in diverse applications of food industry ranging from pigment extraction from

REFERENCES

181

citrus peel to enrichment of omega-3 fatty acids. Nevertheless, this field is still evolving to improve nanocomposites in a multitasking role to produce more value-added products. This may be feasible with the amalgamation of other suitable nanomaterials such as polymeric nanomaterials, graphene, and sheet to develop a suitable and robust nanobiocatalyst. It is anticipated that food quality and safety will be improved significantly in the near future in food industry with the intervention of nanotechnology. Thus, it is strongly advocated that food grade material, while using nanomaterial for enzyme immobilization, adhering to safety standards (GRAS) should be practiced in the food processing industry.

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[125] Y. Yurekli, S.A. Altinkaya, Catalytic performances of chemically immobilized urease under static and dynamic conditions: a comparative study, J. Mol. Catal. B Enzym. 71 (1) (2011) 36 44. [126] S.F. Li, Y.H. Fan, R.F. Hu, W.T. Wu, Pseudomonas cepacia lipase immobilized onto the electrospun PAN nanofibrous membranes for biodiesel production from soybean oil, J. Mol. Catal. B Enzym. 72 (2011) 40 45. [127] P. Ji, H. Tan, X. Xu, W. Feng, Lipase covalently attached to multiwalled carbon nanotubes as an efficient catalyst in organic solvent, AIChE J. 56 (2010) 3005 3011. [128] V. Kumari, S. Shah, M.N. Gupta, Preparation of biodiesel by lipase-catalyzed transesterification of high free fatty acid containing oil from Madhuca indica, Energy Fuels 21 (1) (2007) 368 372. [129] J. Yan, Y. Yan, S. Liu, J. Hu, G. Wang, Preparation of cross-linked lipase-coated micro-crystals for biodiesel production from waste cooking oil, Bioresour. Technol. 102 (7) (2011) 4755 4758. [130] M.L. Verma, N.M. Rao, T. Tsuzuki, C.J. Barrow, C. Puri, Suitability of recombinant lipase immobilised on functionalised magnetic nanoparticles for fish oil hydrolysis, Catalysts 9 (2019) 420. [131] N. Matuoog, K. Li, Y. Yan, Immobilization of Thermomyces lanuginosus lipase on multi-walled carbon nanotubes and its application in the hydrolysis of fish oil, Mater. Res. Express 4 (2017) 25402. [132] N. Matuoog, K. Li, Y. Yan, Thermomyces lanuginosus lipase immobilized on magnetic nanoparticles and its application in the hydrolysis of fish oil, J. Food Biochem. 42 (5) (2018) e12549. [133] S. Kapoor, A. Rafiq, S. Sharma, Protein engineering and its applications in food industry, Crit. Rev. Food Sci. Nutr. 57 (2017) 2321 2329. [134] K. Elavarasan, B.A. Shamasundar, Effect of oven drying and freeze drying on the antioxidant and functional properties of protein hydrosylates derived from freshwater fish (Cirrhinus mrigala) using papain enzyme, J. Food Sci. Technol. 53 (2016) 1303 1311. [135] M.-C. Liu, S.-J. Yang, D. Hong, J.-P. Yang, M. Liu, Y. Lin, et al., A simple and convenient method for the preparation of antioxidant peptides from walnut (Juglans regia L.) protein hydrosylates, Chem. Cent. J. 10 (2016) 39. [136] K. Vosmann, B. Wiege, P. Weitkamp, N. Weber, Preparation of lipophilic alkyl (hydroxy)benzoates by solvent-free lipase-catalyzed esterification and transesterification, Appl. Microbiol. Biotechnol. 80 (2008) 29 36. [137] A.G.A. SA, A.C.D. Meneses, P.H.H.D. Araujo, D.D. Oliveira, A review on enzymatic synthesis of aromatic esters used as flavor ingredients for food, cosmetics and pharmaceutical industries, Trends Food Sci. Technol. 69 (2017) 95 105. [138] K.C. Badgujar, P.A. Pai, B.M. Bhanage, Enhanced biocatalytic activity of immobilized Pseudomonas cepacia lipase under sonicated condition, Bioprocess Biosyst. Eng. 39 (2016) 211 221. [139] R. Porta, A. Pandey, C.M. Rosell, Enzymes as additives or processing aids in food biotechnology, Enzyme Res. 2010 (2010) 436859. [140] D.J. McClements, H. Xiao, Is nano safe in foods? Establishing the factors impacting the gastrointestinal fate and toxicity of organic and inorganic food-grade nanoparticles, npj Sci. Food 1 (2017) 6. [141] S. Kumar, P. Sharma, P. Ratrey, B. Datta, Reusable nanobiocatalysts for the efficient extraction of pigments from orange peel, J. Food Sci. Technol. 53 (7) (2016) 3013 3019. [142] N. Singh, G. Srivatava, M. Talat, H. Raghubanshi, O.N. Srivastava, A.M. Kayastha, Cicer α-galactosidase immobilization onto functionalized grapheme nanosheets using response surface methods and its applications, Food Chem. 142 (2014) 430 438. [143] D. Kishore, M. Talat, O.N. Srivastava, A.M. Kayastha, Immobilization of β-galactosidase onto functionalized graphene nano-sheets using response surface methodology and its analytical applications, PLoS One 7 (7) (2012) e40708. [144] A. Dwevedi, A.K. Singh, D.P. Singh, O.N. Srivastava, A.M. Kayastha, Lactose nano-probe optimized using response surface methodology, Biosens. Bioelectron. 25 (4) (2009) 784 790.

FURTHER READING

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FURTHER READING S.B. Bankar, M.V. Bule, R.S. Singhal, L. Ananthanarayan, Glucose oxidase—an overview, Biotechnol. Adv. 27 (4) (2009) 489 501. S.H. Bhosale, M.B. Rao, V.V. Deshpande, Molecular and industrial aspects of glucose isomerase, Microbiol. Rev. 60 (2) (1996) 280 300. M. Curic, B. Stuer-Lauridsen, P. Renault, D. Nilsson, A general method for selection of α-acetolactate decarboxylase-deficient Lactococcus lactis mutants to improve diacetyl formation, Appl. Environ. Microbiol. 65 (3) (1999) 1202 1206. P.M. de Souza, P. de Oliveira Magalha˜es, Application of microbial α-amylase in industry—a review, Braz. J. Microbiol. 41 (2010) 850 861. P. Fernandes, Enzymes in food processing: a condensed overview on strategies for better biocatalysts, Enzyme Res. 2010 (2010). 862537-862537. A.D. Harris, C. Ramalingam, Xylanases and its application in food industry: a review, J. Exp. Sci. 1 (7) (2010) 1 11. S.L. Hii, J.S. Tan, T.C. Ling, A.B. Ariff, Pullulanase: role in starch hydrolysis and potential industrial applications, Enzyme Res. 2012 (2012) 921362. J. Kaushal, S. Mehandia, G. Singh, A. Raina, S.K. Arya, Catalase enzyme: application in bioremediation and food industry, Biocatal. Agric. Biotechnol. 16 (2018) 192 199. M. Khan, E. Nakkeeran, S. Umesh-Kumar, Potential application of pectinase in developing functional foods, Annu Rev. Food Sci. Technol. 4 (1) (2013) 21 34. O. Kirk, T.V. Borchert, C.C. Fuglsang, Industrial enzyme applications, Curr. Opin. Biotechnol. 13 (4) (2002) 345 351. P. Kumar, T. Satyanarayana, Microbial glucoamylases: characteristics and applications, Crit. Rev. Biotechnol. 29 (3) (2009) 225 255. A.V. Kumar, R.S.C. Kurup, C. Snishamol, G. Prabhu, Role of cellulases in food, feed, and beverage industries, in: B. Parameswaran, et al. (Eds.), Green Bio-processes: Enzymes in Industrial Food Processing, Springer, Singapore, 2019, pp. 323 343. T. Panda, B.S. Gowrishankar, Production and applications of esterases, Appl. Microbiol. Biotechnol. 67 (2) (2005) 160 169. M. Puri, U.C. Banerjee, Production, purification, and characterization of the debittering enzyme naringinase, Biotechnol. Adv. 18 (3) (2000) 207 217. S. Saqib, A. Akram, S.A. Halim, R. Tassaduq, Sources of β-galactosidase and its applications in food industry, 3 Biotech 7 (1) (2017). 79-79. F. Zhu, B. Du, B. Xu, A critical review on production and industrial applications of beta-glucans, Food Hydrocoll. 52 (2016) 275 288.

CHAPTER

ENZYME ENTRAPMENT APPROACHES AND THEIR APPLICATIONS

11

Manisha Sharma and Sudhir P. Singh Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India

11.1 INTRODUCTION Enzymes as a biological catalyst have changed the scenario by bringing a drastic change, and thus considered a remarkable discovery in the world of industries. This has been further proven by its utilization in diverse sectors, including food, pharmaceutical and textile industries, etc. The enhanced usage of enzymes over chemical approaches is due to their substrate specificity, green chemistry, and biological mode of catalyst production. Even under mild conditions, enzymes could initiate the reaction, catalyzing the biosynthesis of specific products, unlike the chemical approach of the product synthesis. Since past ancient times, enzymes play an integral role in our lives, for instance, in the processing of various products such as linen, leather, and indigo, etc. The term enzyme has been originated from the Greek word “ενζυμoν”, coined by Wilhelm Friedrich Kuhne, from the University of Heidelberg in 1877 [1]. However, the application of these kinds of biomolecules as biocatalysts was described by Gottlieb Sigismund Kirchhoff in 1812 during his investigation of the process of conversion of starch into glucose [2]. The first enzyme discovered was diastase in 1833 by a French chemist, Anselme Payen [3], whereas, in 1835, Swedish scientist Jons Jacob Berzelius carried out the hydrolysis of starch by diastase. In 1846, Dubonfout described the activity of invertase, and later in 1862, Louis Pasteur and team studied the fermentation of sugar to alcohol [4]. The first pure enzyme urease was isolated by James B. Sumner from Cornell University in 1926. Subsequently, various enzymes have been discovered, produced, and purified worldwide. Thus, from ancient times to the beginning of the 20th century, a vast expansion of the technological use of the biological catalysts has been experienced from household applications to industrial applications. Various carbohydrate- and protein- active enzymes such as proteases, amylases, cellulases, dextransucrases, and dextranases have been categorized under the group of the most actively used enzymes in the starch, food, baking, and bioprocessing industries [5]. In spite of having so many applications, the extraction of enzyme is quite expensive. The recovery of the enzyme contaminant from the treated sample is difficult. From the commercial point of view, the recovery and reusability factor of enzyme molecules is very demanding to make the process economical. Keeping this in mind, enzyme immobilization methods have been developed, enhancing the enzyme’s stability and reusability for multiple reaction cycles. Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00011-9 © 2020 Elsevier B.V. All rights reserved.

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FIGURE 11.1 Flowchart representation of comparison between free (A) and immobilized (B) enzymatic processing.

11.2 IMMOBILIZATION OF ENZYMES The immobilized enzymes are defined as the state of the enzyme where it is physically confined to a matrix without any change in their catalytic properties, but it could be used repeatedly. Immobilized biocatalysts are helpful in cost reduction of the enzymatically formed product, due to multiple duty cycles of the catalytic reactions (Fig. 11.1).

11.3 HISTORY OF ENZYME IMMOBILIZATION Several enzymatic processing-based technologies have been developed in food, pharma, and textile industries. The number of papers and patents on the immobilized enzymes has been found to be more than 5000 to date. The adsorption of invertase on charcoal and alumina was the first published report of enzyme immobilization by Nelson and Griffin in the year 1916. Several inorganic and organic carriers have been utilized for attachment of enzymes via adsorption and covalentfixation approaches. A few enzymes, in their immobilized forms, have been utilized in various industrial processes. This has revealed the commercial importance of some of the immobilized enzymes due to their higher stability, unaltered activity, and reusability [6].

11.4 ADVANTAGES OF ENZYME IMMOBILIZATION Multidimensional researches have been performed to target the issues associated with enzymatic bioprocesses, such as high cost of enzyme production, enzyme purification, enzyme instability, and

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Table 11.1 The advantages and disadvantages of immobilization of enzymes. Immobilization of enzymes Advantages

Enzyme repetitive usability Ease in separation of product from enzyme Stability of enzyme from harsh environment Easy process with minimum contamination

Disadvantages

Reduction or loss of activity Limitation in diffusion of substrate and product High cost of carriers Support systems require costly and highly designed engineering approach

tedious enzyme recovery after completion of catalytic reaction. Immobilization of enzymes confers several advantages on the aforementioned aspects, for example, stabilization of the enzyme structure, increased tolerance to harsh conditions such as pH, temperature, organic solvents, easier recovery of enzyme from the reaction mixture, reusability of the enzyme, and development of the continuous process of the catalytic reaction. These improvements in the biocatalyst properties help in the economics of the enzymatic bioprocess applications in food, textile, and pharmaceutical industries [7]. Table 11.1 depicts the summary of the advantages and disadvantages of immobilized enzymes.

11.5 SUPPORT SYSTEMS FOR ENZYME CONJUGATION In order to immobilize, the properties of both the enzymes as well as support matrix are taken into consideration. Both organic and inorganic support systems have been employed for the immobilization of the enzyme (Fig. 11.2).

11.5.1 ORGANIC SUPPORT MATERIALS Eupergit C is one of the organic supportive materials used for the immobilization of enzymes. It is a highly stable polymer, both chemically and mechanically. The porous structure of Eupergit C polymer has
N,N-methylene-bis(methacrylamide), glycidyl methacrylate, allyl glycidyl ether, and methacrylamide, as components for the covalent interaction with the amino group of the enzyme. The more the density of oxirane groups, the more binding of enzyme occurs on the surface of Eupergit C. This property has made Eupergit C a desirable matrix to develop industrial bioprocesses [8
10].

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FIGURE 11.2 Different categories of carrier or support systems employed in enzyme immobilization.

11.5.2 BIOPOLYMERS The biopolymers, alginate, cellulose, starch, chitosan, and pectin are the biopolymers that have been widely demonstrated as suitable matrix for enzyme immobilization. Some protein-based materials such as albumin and gelatin have also been utilized as an attachment matrix [11].

11.5.2.1 Alginate Alginate is one of the most commonly used biopolymers for the immobilization of enzymes. It is derived from brown algae and comprised of calcium, magnesium, and sodium salts of alginic acid. Various enzymes have been immobilized using alginate, exhibiting improved operational stability and reusability for many cycles, such as α-amylase [12], pectinases, [13,14], glucoamylase [15], dextransucrases [16], and dextranases [17]. Research findings in several reports endorsed enhancement in the stability of enzyme after its attachment to the alginate material, cross-linked with divalent ions (such as Ca21) and glutaraldehyde [18,19].

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195

11.5.2.2 Chitin and chitosan Chitin and chitosan have been utilized in immobilization of several enzymes [20
22]. It has also been used in combination with alginate [23]. The alginate
chitosan-immobilized enzyme showed reduced leaching as compared to alginate-immobilized enzyme. The strong association between the enzyme and support matrix could be developed due to the abundance of hydroxyl and amino groups, high porosity, and hydrophilicity of the chitosan compound.

11.5.2.3 Collagen The natural polymer, collagen, has been mostly used in combination with glutaraldehyde crosslinker, for example, for tannase immobilization [24], and Fe31 supporting matrix, for example, for catalase immobilization [25].

11.5.2.4 Carrageenan Carrageenan is a linear, sulfated polysaccharide that has been explored for the immobilization of lipase [26] and galactosidase enzymes [27,28]. Biodiesel production has also been carried out by co-extrusion method with 42.6% immobilization efficiency [29].

11.5.2.5 Gelatin The amino acid-rich hydrocolloid, gelatin, has an indefinite shelf life, which makes it an attractive matrix for biocatalyst fixation. Gelatin has been used in complex with polyacrylamide cross-linked with chromium (III) acetate or chromium (III) sulfate or potassium chromium (III) sulfate [30]. Further, efficient enzyme immobilization including laccase [31], β-glucosidase [32], α-amylase [33], and glucoamylase [34] have been demonstrated by using gelatin along with calcium alginate, polyester films, and titanium (IV) species [35,36].

11.5.2.6 Cellulose Cellulose, the most abundant polymer, has been explored for enzyme immobilization experiments, such as with fungal laccase, penicillin G acylase (PGA), glucoamylase, α-amylase, tyrosinase, lipase, and β-galactosidase [37
43]. Cellulose has also been used with magnetite nanoparticles and dialdehyde-coated magnetite nanoparticles for attaching starch-degrading biocatalysts [40]. The ionic liquid
cellulose film, after activation with glutaraldehyde, could provide a better formability and flexibility after immobilization [43].

11.5.2.7 Starch Starch is a homopolysaccharide of monomeric glucose units linked together by α-1,4 linkage. It has linear amylose and branched amylopectin chains. Immobilization of bitter gourd peroxidase was carried out using calcium alginate
starch hybrid supports. This not only caused enhancement in the enzyme’s activity but also provided protection to the enzyme from some denaturants such as urea [44].

11.5.2.8 Pectin Pectin, a heteropolysaccharide with a backbone of galacturonic acid, in combination with alginate, has been found a suitable matrix for many enzymes, for example, papain, dextransucrase, and

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dextranase [45,46]. The attachment enhanced thermal tolerance with significantly reduced leakage of the enzyme. This might be because of the stable interaction in the form of polyelectrolyte complexes between the enzyme and the pectin-coated support matrix [47,48].

11.5.3 HYDROGELS The hydrogel or cryogel, natural or synthetic, facilitate immobilization of enzyme in nonaqueous media. The whole-cell secreting enzyme has also been attempted to get immobilized using polyvinyl alcohol (PVA) cryogels [49]. PVA can be synthesized either through freeze-drying [50] or through partial drying at room temperature. The PVA formed through incomplete drying has been found to be more stable. PVA has also been explored for the immobilization of lipase enzyme from Candida rugosa [51], alcohol dehydrogenase [52], and glucose oxidase [53]. Jekel reported the lens-shaped hydrogels, known as Lentikats, formed at room temperature [54]. It has been exploited for the immobilization of whole cells of Rhodococcus equi A4 [55], dextransucrase [56], and β-glucosidase [57]. Generally, the small-sized proteins leach out from the matrix, but in the case of PVA gels, the use of organic media, as well as cross-linkers, prevent the leaching of the enzyme from the matrix [58].

11.5.4 INORGANIC SUPPORT MATERIALS 11.5.4.1 Zeolites Zeolites are the microporous crystalline solids with characteristic structure and shape. It has multiple adsorption sites involved in the interaction with enzyme molecules. The hydroxyl groups in microporous zeolites interact with the enzyme through strong hydrogen bonds, as compared with microporous dealuminized ones [59]. It has been used for the immobilization of α-chymotrypsin [60]. Chang and Chu reported a higher activity of lysozyme after immobilization with NaY zeolite [61].

11.5.4.2 Ceramics This support has been employed for the immobilization of lipase from Candida antarctica [62]. The different pore sizes of the ceramic membrane could reduce the diffusion rate and increase the specific surface area [63].

11.5.4.3 Celite It is a highly porous and diatomaceous bioaffinity material with low polarity and large adhesion area. It is an economical material that has been utilized for the immobilization of lipase, polyphenol oxidases, and β-galactosidase [64
66]. It provides protection to the enzyme from denaturation caused by high pH, temperature, and denaturants [64]. Transaminases have been immobilized to Celite as sol
gel matrix. The inertness and interconnected pore structure of Celite make it a preferable material for enzyme fixation [67].

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11.5.4.4 Silica Silica has been explored for the immobilization of lignin peroxidase and horseradish peroxidase (HRP). It removes chlorolignins from eucalyptus kraft effluent [68]. The cleaning efficiency of detergents is enhanced after immobilizing α-amylase on silica nanoparticles. The desirable characteristics of silica are the ordered arrangement, characteristic structural arrangement, surface area, and tolerance to chemical and mechanical forces [69]. The incorporation of methyl or PVA groups, hydroxyl, and reactive siloxane groups have been found to strengthen the efficiency of silica for enzyme binding [70
72].

11.5.4.5 Glass Glass is a viscous liquid material that could be used for immobilization of enzymes such as α-amylase. Glass beads functionalized with amino group-containing phthaloyl chloride is robust and renewable material for enzyme attachment [73]. The immobilization of nitrite reductase on controlled pore glass beads has been employed as a biosensor for continuous monitoring [74]. Similarly, glass pH electrodes used for immobilization of urease have been demonstrated to sense urea in the blood sample [75].

11.5.4.6 Activated carbon The activated carbon, both natural and modified with hydrochloric acid, is used as the matrix for the absorption of the enzyme [76]. This has been demonstrated as a promising material for enzyme’s reusability. Even after immobilization of acid protease and acidic lipases onto the mesoporous-activated carbon particles, the enzymes maintained the catalytic efficiency after 21 cycles of reuse [77,78]. The suitable characteristics of activated carbon for enzyme’s immobilization are large surface area and presence of pore volume in the range of 600
1000 m2/g, and ˚ , respectively [79]. 300
1000 A

11.5.4.7 Charcoal Charcoal has been considered as a superior material for enzyme’s adsorption [80]. Amyloglucosidase, immobilized onto charcoal without using any cross-linking agent, has been established as a promising biocatalyst system for starch hydrolysis [81]. Papain enzyme molecules have been grafted onto charcoal particles, and this immobilized matrix system has been demonstrated useful industrial wastewater treatment [82].

11.5.5 PROTEIN-COATED MICROCRYSTALS TECHNIQUE The inorganic support protein-coated microcrystals (PCMCs) has been explored for the immobilization of freeze-dried enzymes using lyoprotectants and other inorganic salts. The PCMC is mainly formed by mixing enzyme with salts such as potassium sulfate, amino acid, or any sugar in aqueous form and adding dropwise into the isopropanol. The micro-sized crystals so formed contain immobilized enzymes. Many enzymes have been immobilized in such ways, for example, lipases, oxidoreductases, catalase, soybean peroxidase, and HRP [83].

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11.5.6 SMART POLYMERS Smart polymers are highly responsive to the change in environmental parameters such as pH, temperature, and ionic conditions [84
88]. Poly-N-isopropylacrylamide (polyNIPAM) polymer is one such responsive polymer that has been conjugated with enzyme either through the introduction of vinyl groups into the enzymes or through reaction with NH3 group [89]. Ivanov described the immobilization of penicillin G amidase (PA) using NIPAM, which is rich in highly reactive functional groups, through condensation reaction [90]. NIPAM is readily dissolved in the water below its critical solution temperature (LCST), that is, 32 C, and gets precipitated above its LCST. It could be employed for reducing the diffusional limitations as well as loss in enzyme’s activity. The enzyme could be easily recovered by precipitating the immobilized enzyme by raising the temperature above the LCST.

11.5.7 CONDUCTING POLYMERS Conducting polymers are the polymers exhibiting electrical conductivity. Due to its potential in various biotechnological applications, synthesis and characterization of conducting polymers are in great demand [91]. The charge transfer complexes/radical ion salts, organometallic species, and conjugated organic polymers are desirable for transporting organic compounds. The intrinsically conducting polymers or electro-active conjugated polymers are a newly emerging class of polymers exhibiting the electrical and optical properties. Such electronically conducting polymers were applied for the application of enzyme immobilization and biosensor production [92,93]. In various studies, enzymes were immobilized using conducting polymers. Alcohol oxidase was conjugated with carboxylic acid functionalized multiwall carbon nanotubes (f-MWCNTs) as conducting polymer to evaluate the amount of alcohol in the beverages [94]. Glutamate oxidase, immobilized with the functionalized conducting polymer of 5,20 :50 ,2v-terthiophene-30 -carboxylic acid on a platinum microelectrode, was exploited in studying in vivo release of extracellular glutamate due to cocaine stimulation [95]. Invertase was attached to polypyrrole and polyamide/polypyrrole electrodes, which retained its activity for several months [96].

11.6 TYPES OF IMMOBILIZATION TECHNIQUES The broad categories of immobilization techniques have been shown in Fig. 11.3.

11.6.1 SUPPORT MATRIX SYSTEM In this system, the enzyme is attached to a support through physical, ionic, or covalent bonding. Among these, physical interactions, including hydrophobic and van der Waals interactions, are not able to withstand and keep the enzyme fixed for a long period of time, especially under industrial conditions of high temperature and pH. However, ionic and covalent bonding is relatively stronger, and therefore, preferred over other physical interaction methods. It also offers reduced enzyme leakage [97].

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FIGURE 11.3 Flowchart showing different types of immobilization techniques.

11.6.2 ENTRAPMENT SYSTEM In the case of entrapment system, sol
gel or polymer network is involved in the immobilization of enzyme. The entrapment should be made capable of preventing the leakage of the enzyme, efficiently. Therefore, additional covalent interaction is recommended to enhance the enzyme binding with the support system [97].

11.6.3 CROSS-LINKING SYSTEM Cross-linking-type approach is a useful strategy for the immobilization of the enzyme. Crosslinking of enzyme is achieved without the utilization of any carrier, either as crystals or aggregates, known as cross-linking enzyme crystals, and cross-linking enzyme aggregates (CLEAs), respectively [98]. This approach has many advantages, such as an increase in the activity and stability of enzyme and reduction in the cost of production and processing [97].

11.7 METHODS OF ENZYME IMMOBILIZATION The method for the immobilization of enzymes is mainly based on physical or chemical interactions. Physical method is a type of noncovalent interaction, whereas in chemical interaction covalent bond formation occurs between the enzyme and support. These immobilization methods do not bring any modification in the structure of the enzyme, therefore, show very less adverse effect on the activity of the enzyme. Physical methods of enzyme immobilization have been classified into the following subcategories: physical adsorption, ionic bonding, and physical entrapment. In the same way, chemical methods involve cross-linking and covalent bonding (Fig. 11.4).

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FIGURE 11.4 Flowchart representation of different methods of immobilization.

11.7.1 PHYSICAL METHODS 11.7.1.1 Physical adsorption The dipole
dipole and hydrophobic interactions, van der Waals forces, or hydrogen bonding are involved in the physical adsorption based on the characteristic properties of the substrate and the adsorbate [99]. Being a simple and economical technique, with the capacity to retain high catalytic activity, physical adsorption has been used for enzyme immobilization in the industry [100]. The technique has been proven economical for industrial applications, as it allows the reusability of supportive materials. This technique, on the other hand, is less stable and sometimes leads to leakage of molecules during reuse processes [101]. The adsorption also protects the enzyme from making aggregates, getting lysed, and interacting with hydrophobic interfaces [102]. In 1916, Nelson and Griffin carried out the immobilization of invertase onto aluminum hydroxide through physical adsorption. Some of the eco-friendly supports including coconut fibers, microcrystalline cellulose, kaolin, and micro/mesoporous materials have been utilized by researchers. They were found to have good water-holding and cation exchange properties, making the enzyme’s performance better in reduction and oxidation reactions [42,103
107]. The polypropylene-based hydrophobic granules/Accurel EP-100, used for the immobilization of lipase, were found to enhance the reaction rate [108,109]. The octyl-agarose and octadecyl-sepabeads were explored for the immobilization of Yarrowia lipolytica lipase by physical adsorption. This controlled process with economic production led to higher yields with 10-fold more stability as compared to free lipase. This might be because of the high affinity between the enzyme and support system due to the hydrophobicity of octadecyl-sepabeads [110]. Another biodegradable material utilized for the immobilization of lipase obtained from C. rugosa via adsorption showed the reusability of the enzyme up to 12 cycles with 94% activity retention after 4 h at 50 C [111]. Due to crystallinity and less toughness, these aforementioned supports are more preferred than polyhydroxybutyrate. Mishra et al 2011 reported the

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utilization of 1,4-butenediol diglycidyl ether-activated byssus threads for the immobilization of urease with enhanced pH stability and retention in 50% of enzyme activity [112]. The supports of biological origin are eco-friendly, cost-effective, as well as free of ethical issues. One such example included the use of biocompatible mesoporous silica nanoparticles (MSNs) as a support system for enzymes [113].

11.7.1.2 Ionic binding The electrostatic interaction between two oppositely charged groups is called ionic bonding. This ionic bonding between oppositely charged groups of the carrier and the enzyme molecules constitute another type of immobilization technique. The ion exchangers (either anion or cation) prepared from organic polymers such as dextran and cellulose are also considered good carriers for enzyme immobilization.

11.7.1.3 Physical entrapment In order to protect the enzymes from getting affected by external agents such as proteases, a noncovalently linked cage-like protection has been provided to the enzymes known as entrapment-based immobilization [114]. This has been further classified into two subgroups—one is gel entrapment, and the other is microencapsulation. The development of microencapsulation was initiated for the encapsulation of enzymes, which were later modified with various upgrading techniques, namely carrier-bound immobilization encapsulation, cross-linking encapsulation, and entrapment encapsulation. The sol
gel is one of the well-established techniques for the entrapment by retaining the active functional state of the enzyme [115
117]. Some of the practices of the encapsulation methods have been reported in the literature including the formation of liposomes, where an aqueous solution of the enzyme was encapsulated in the lipid layer forming amphiphilic molecules. Such vesicles containing enzymes have been applied in the medical or biomedical field for the enzymereplacement therapy as well as in the process of cheese ripening and controlled flavor development process [118]. Another type of encapsulation, as an alternative to liposomes, has been practiced with biodegradable polymers in nano-size so as to allow them in various drug delivery processes, for example, poly (lactic acid) (PLA) and poly (lactic-co-glycolic acid) (PLGA). They have been found to be extensively biocompatible with the potential capability of releasing therapeutic proteins over a long period of time in a controlled manner. Alginate has been reported to be a preffered matrix for the immobilization of the enzymes. The utilization of alginate for the entrapment of the enzyme has been proven to be cheap, safe, nontoxic and foodgrade. Many enzymes have been immobilized using alginate, for example, dextransucrase entrapped with alginate showed immobilization yield ranging from 57% to 98% [119
123], β-glucosidases immobilized with calcium alginate and cross-linker showed higher thermal and pH stability with increased affinity with the substrate, etc [124]. The major drawback found in alginate-immobilized enzyme is the leakage of the enzyme into the medium after repeated cycles of use. This problem has been tried to overcome with the utilization of certain polymers, polysaccharides in combination with alginate, which was found to reduce the leakage of the enzyme, with improved catalytic properties. Thus, the immobilization of β-galactosidase with alginate
gelatin
calcium hybrid carriers [35] and immobilization of dextransucrase and dextranase using alginate
pectin beads [46] have been demonstrated to reduce the leakage and loss in activity of enzymes after repeated cycles. Similarly, chitosan was also utilized for entrapment of lipase from C. rugosa with significant entrapment efficiency and enzyme

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activity. Chitosan based support has also been considered to be nontoxic and biocompatible with good affinity with proteins [125]. The mesoporous silica has been employed for the entrapment of the enzymes due to significant characteristics related to its surface area, pore distribution, and adsorption capacity [126]. The utilization of carrageenan was carried out for the entrapment of lipase with improved thermostability and tolerance toward organic solvent [26,29].

11.7.2 CHEMICAL METHODS 11.7.2.1 Cross-linking Cross-linking is one of the chemical methods of enzyme immobilization in which enzyme is attached to each other through covalent bond via bi- or multifunctional reagents. Glutaraldehyde is one such common solvent to be utilized as a linker reagent. They make the enzyme insoluble in water by forming three-dimensional cross-linked aggregates [127]. The immobilized enzymes are generally of controlled size ranging from 1
100 μm with high productivity, operational stability, and easy recycling of the catalyst. The incorporation of salts, water-miscible organic solvents, or nonionic polymers also led to the precipitation of enzymes from the aqueous solution of the reaction mixture by forming CLEAs [128]. These physical aggregates cross-linked by noncovalent bonding are found to be permanently insoluble with maintained structure and catalytic activity. Such type of immobilized enzymes is thus seen to be quite effective in terms of production cost, catalytic activity, and reusability [129]. This immobilization technique has been applied to attach cofactor-dependent oxidoreductases [128] and penicillin acylase with remarkable improvement in their activities [130].

11.7.2.2 Covalent binding Covalent immobilization has brought a breakthrough and new development in the field of enzyme immobilization because the covalent attachment of the enzyme with support matrix does not require the functional groups of the catalytic site of the enzyme. Mainly the targeted functional groups of the enzyme for the covalent immobilization involve hydroxyl and amino groups and to some extent, sulfhydryl groups [127]. The drawback associated with covalent immobilization is the inactivation or loss in activity of enzyme due to unwanted conformational structural changes. Thus, covalent immobilization also proceeded with some modifications [127]. The intense contact between the support matrix and enzyme does not allow the leakage of the enzyme into the reaction sample. A study on the immobilization through linkage of protein with peptide-modified surfaces led to higher stability and activity of protein with maintained protein orientation [131]. Similarly, covalent immobilization of enzymes using cyanogen bromide (CNBr)-agarose and CNBr-activated Sepharose were found to enhance the thermal stability of enzymes [22,110]. The modification of silica gel carriers after eliminating unreacted aldehyde groups, and SBA-15 supports (having cage-like pores with Si
F moieties lining) have been exploited for the covalent immobilization of enzymes [132,133]. The utilization of mesoporous silica, chitosan, etc., as a support system for covalent immobilization, enhanced the thermal stability and half-life of the enzymes [22,126]. The thiol group of cysteine residues of the enzyme was covalently conjugated with unsaturated carbonyls, which led to the formation of stable thioether bonds [134
136].

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11.8 APPLICATIONS OF IMMOBILIZED ENZYMES 11.8.1 FOOD INDUSTRY The processing and analysis of food ingredients are the main functional aspects of food industries. These have been more pronounced with the utilization of immobilized enzymes [137], as the fixed biocatalysts minimize the cost of processing [138]. The processes where enzymes are used in the food industries include processing of starch and cheese, preservation of food, hydrolysis of lipid, etc. α-amylase is one of the most common enzymes used in the food processing industry. Due to the reduced operational stability and reusability, a focus has been given to the immobilization of α-amylase. Different attempts have been made for the immobilization of α-amylase using ionexchange resin beads [139] and CLEAs [140] with better stability and improved processing of starch. Pectinases are another group of enzymes utilized in the food industries for clarification of juices. Various reports have been mentioned in the literature showing immobilization of pectinase using calcium alginate [141], PVA sponge [142], silica-coated magnetite nanoparticles [143], and magnetic cornstarch microspheres [144]. Immobilization has been found to improve the optimum pH, temperature, and thermal stability as well as the reusability of the pectinases even after several cycles of its use. The most important enzyme of dairy or food industries, β-galactosidase, catalyzed the breakdown of lactose in milk or whey. It has also been immobilized using various support materials in order to enhance its utilization in continuous batch processes. The immobilization of β-galactosidase with calcium alginate and in complex with concanavalin A led to the stability of the enzyme against denaturants with the efficient hydrolysis of whey or milk hydrolysis [145]. The increased thermal stability of the recombinant β-galactosidase from Bacillus stearothermophilus after immobilization using tris(hydroxymethyl)phosphine and glutaraldehyde was also studied [146]. For the deproteinization, proteases are widely applied in protein containingstuff in food industries. The chitosan beads have been employed for the immobilization of alkaline protease to perform the deproteinization of natural rubber, which led to the reduction of the 96% of the nitrogen level with the reusability of the enzyme up to five cycles [147]. A group carried out the covalent immobilization of a novel cysteine protease, procerain B, on glutaraldehyde-activated chitosan matrix with improved reusability for use in food industries [148].

11.8.2 PHARMACEUTICAL INDUSTRY The enzyme immobilization has gained an important place in the pharmaceutical industry. The production of various biotechnological products, have been carried out using immobilized enzymes [149]. Cysteine proteinases, asparaginase, streptokinase, urokinase, deoxyribonuclease I, hyaluronidase, pegademase, and glucocerebrosidase are some of the enzymes used in the pharmaceutical industry. The major application of immobilization of enzyme employed in pharmaceutical industries is the production of penicillin using immobilized penicillin amidase [150,151]. In the world, about 19% of the worldwide antibiotic market share has gone into the pocket of penicillin [150]. Therefore, for the production of semisynthetic penicillin, the PGA enzyme plays an important role in catalyzing penicillin G to produce 6-aminopenicillanic acid (6-APA). The immobilization of PGA enzyme on metal affinity membrane (IMAM) showed more activity, stability, and reusability up to 16 cycles [152]. The macro-mesoporous silica spheres were also employed for the

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immobilization of PGA to produce penicillin G [153]. The κ-carrageenan was employed for the immobilization of bacterial cells. This led to the continuous production of APA from penicillin G [154]. Apart from 6-APA, 7-aminodeacetoxycephalosporanic acid (7-ADCA), 7aminocephalosporanic acid (7-ACA), and deacetyl-7-aminocephalosporanic acid (D-7-ACA) are also utilized for the production of semisynthetic antibiotics of penicillins or cephalosporins. The production of fine chemicals in the pharmaceutical industries has been carried by employing an important enzyme, D-amino acid oxidase. To protect from the exposure of gas
liquid interface, D-amino acid oxidase from Trigonopsis variabilis (TvDAO) was immobilized with functionalized silica carrier [155]. After the entrapment with calcium alginate matrix, D-amino acid oxidase showed good operational stability and provided an economic process for the industries [156]. Another essential enzyme from pharma point of view is L-asparaginase, generally used for the treatment of acute lymphoblastic leukemia and lymphosarcoma. Various support materials have been explored for the immobilization of L-asparaginase and the host microorganisms such as polyaniline nanofibers matrix [157], silica gel carrier [158], and chitosan tripolyphosphate nanoparticles [159]. It has been found that immobilization enhanced the affinity of the enzyme for the substrate as well as improved the reusability of the enzyme. Streptokinase is one of the most widely used enzymes in the pharmaceutical industries for the treatment of vein thrombosis, pulmonary embolism, and acute myocardial infarction. In order to enhance the shelf life of the enzyme, various immobilization approaches have been employed, involving chitosan nanoparticles [160], and PVC-g-PEGMA [161].

11.8.3 BREWING INDUSTRY From ancient times, brewing has been considered and practiced as a fermentation process. Advancement in fermentation technologies has led to the development of various brewing industries [162]. The immobilized yeast cells have become popular in brewing processes [163]. But the process optimization, cost of production, and altered beer flavor due to physiological changes are reasons behind the failure of the continuous production of beer using yeast cells in a bioreactor [164]. These problems have been overcome through various approaches, but the immobilization has been considered a better option for the production of beer with improved characteristics of color, flavor, etc. Researchers have studied the production of beer using free or alginate-immobilized yeast cells in which not only the rate of fermentation remains unaltered but also sensory qualities of the beer remain the same using immobilized cells [165]. The immobilization of a cryotolerant Saccharomyces cerevisiae was studied using porous cellulosic material [166]. The utilization of low-cost support material such as corn cob has also been employed for the immobilization of the S. cerevisiae for ethanol production. This could offer an alternative method for winemaking [167]. The support systems such as calcium alginate
magnetic nanoparticles, chitosan
magnetite microparticles, and cellulose-coated magnetic nanoparticles have been explored for the immobilization of yeast S. cerevisiae for ethanol fermentation. But the immobilization with magnetic particles showed more stability during fermentation. It has been observed that immobilization has improved the quality and production of ethanol in a bioreactor [168].

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11.8.4 DETERGENT INDUSTRY The enzymes have gained an important place in the production of detergents. The most common enzymes utilized in the detergent industries include amylases [169], lipases [170], and cellulases [171]. The immobilization of α-amylase has been carried out on the silica nanoparticles to employ as a detergent powder for cleaning and removal of starch. The reason for the demand of immobilization of enzymes for detergent industries is to protect the stability and activity of the enzymes from the negative effect of the compounds present in the detergents. In a comparative analysis, α-amylase immobilized on silica nanoparticles was able to remove starch from cotton fabrics with better efficiency than the free form of the enzyme. Immobilization has been shown to improve thermal stability and humidity tolerance of enzyme [172]. The immobilization of Bacillus megaterium MTCC2444 with calcium alginate via entrapment was found to enhance the production of enzyme, alkaline protease, for its use in the detergent industry. The enzyme produced from immobilized cells gave a higher yield and better stability against oxidants, surfactants, and commercial detergents [173]. The covalent co-immobilization of commercial Aspergillus niger α-amylase, Trichoderma viride cellulase, protease, and lipase from porcine pancreas onto a plastic bucket and brush was demonstrated to be useful in cloth washing. The immobilized enzymes also exhibited improvement in pH and temperature tolerance. The conjugation yield was observed to be higher on the brush than on the beaker wall, which could be because of larger surface area in case of the brush. The matrix
enzyme conjugates were used along with detergent in the washing of starch, grass, egg albumin, and oil-stained cotton cloth pieces. The results confirmed the better performance of detergents in combination with immobilized enzymes than that of sole detergent [174].

11.8.5 PULP AND PAPER INDUSTRY A huge amount of waste is generated by the pulp and paper industry. This has become a serious environmental pollutant that needs to be eliminated. Biological means are preferred to address these industrial wastes [175]. Immobilized enzymes could be a promising biological mean for the removal of the effluents. For example, a bench-top bioreactor containing the immobilized form of two basidiomycetous and a deuteromycetous fungi, extracted from pulp and paper mill effluents, was demonstrated to reduce the color and degrade lignin in the effluent [176]. One of the major pollutants from the paper and pulp industry is chlorophenols. The covalent immobilization of peroxidases from horseradish and soybean onto aldehyde glass through their amine groups evaluated the effect of immobilized peroxidases in the removal of chlorophenols in comparison with the free enzyme [177]. It has been reported that immobilized form was able to remove the contaminate in a shorter time period. Laccase from Streptomyces psammoticus, encapsulated in alginate beads, removed significantly reduced the color and phenol content [178]. The immobilized laccase retained 50% of its activity even after eight reaction cycles, and thus it is useful for the development of an economical process. The critical issues related to the presence of pitch particles originated from whitewater during papermaking has been attempted to get solved by employing chitosan-immobilized lipase. This has shown a significant decrease in the pitch particles with improved operational stability [179a,b]. The bleaching process carried out in pulp and paper industry releases a lot of oxalate as a by-product. This has been handled using oxalate decarboxylase immobilized to Eupergit C [180].

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11.8.6 TEXTILE INDUSTRY The enzymes have gained an important place in the field of the textile industry [181]. Many enzymes used in the textile processing include cellulase for the finishing and softening of denim and cotton, respectively, amylase for alteration of sizes, pectate lyase for scouring, catalase for terminating bleaching, laccase for carrying bleaching process, and peroxidase for the removal of dye [182]. The immobilization of these enzymes has shown a better performance in the textile industries; for example, immobilized commercial acid cellulases effectively adsorbed abrade indigo-dyed denim fabrics [183]. Immobilized lipase onto zirconia-coated alkylamine glass beads by glutaraldehyde coupling was employed in washing cotton clothes [184]. Calcium alginate-entrapped peroxidase showed a better efficiency to treat effluents [185]. Immobilized laccase on poly (4-VP) grafted and/or Cu (II) ions chelated magnetic beads could be utilized for large-scale applications [186].

11.8.7 BIOMEDICAL APPLICATIONS It is well known that enzymes have been playing an important role in various medical applications since ancient times. This has been implemented nowadays by isolating and purifying various clinically important enzymes. In 2004, Reis and Rom´an elaborated the importance of immobilized enzymes in biomedical applications [187]. Mainly due to the stability, life-span, and safety issues related to enzymes, immobilized enzymes are preferred [188]. In biomedical industries, polyphosphazenes as hybrid inorganic
organic polymers with remarkable physicochemical and biocompatible properties play critical roles in drug delivery, biosensing, enzyme immobilization, etc. Another material for the immobilization is core/sheath nanofiber membrane, formed by using poly [bis(pmethylphenoxy)] phosphazene (PMPPh). The lipase immobilized using nanofiber and polyphosphazenes system was found to show enhanced activity and better results for biomedical applications [189]. Similarly, enzymes were immobilized on different support systems for biomedical industries, for example, egg-white lysozyme was utilized as antibacterial agent after immobilization on electrospun chitosan nanofiber [190]. Acetylcholinesterase and choline oxidase were exploited for biosensing after co-immobilization on poly-2-hydroxyethyl methacrylate membrane.

11.9 CONCLUSIONS AND PERSPECTIVES The immobilization technology has been found to enhance the activity and stability of the enzyme in different reaction conditions. This has been clearly reflected in the evolutionary and remarkable phase change occurred in the enzyme technology in terms of its production and utilization from household activities to industrial scale. Different support materials have been tested for many enzymes of industrial importance. The support matrix has the capability to alter enzymatic activity and stability. The choice of support material should be made according to the compatibility of the enzyme for conjugation, which could improve the operational stability of the biocatalytic processes. Immobilization technology can provide several acceptable enzymatic bioprocesses in the field of food, pharma, and textile industries.

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The technology of immobilization of enzymes has marked a peculiar place at industrial-level production of products. But, most of the enzymes exploited in the industries for the production of food ingredients, antibiotics, and detergents are still difficult to be utilized in the immobilized form, mainly for continuous processes. The process of immobilization of enzymes needs to be understood in a better way so that an enzyme can be immobilized without affecting the structural arrangements of the protein molecule. Intensive researches should be carried out to curtail any chance of enzyme leakage from the attachment matrix. This will increase the use of recombinant enzymes in the food processing industry. The development of new approaches is required for the immobilization, which utilizes low-cost and nontoxic support that could make the whole process of immobilization as well as product formation in an economical and sustainable manner.

ACKNOWLEDGMENTS The authors acknowledge the Department of Biotechnology (DBT), Government of India, and Center of Innovative and Applied Bioprocessing, Mohali. MS acknowledges the Council of Scientific and Industrial Research (CSIR), for fellowship.

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771. [184] S.V. Malik, V. Kalia, C.S. Pundir, Immobilization of porcine pancreas lipase on zirconia coated alkylamine glass using glutaraldehyde, Indian J. Chem. Technol. 7 (2) (2000) 64
67. [185] M. Matto, Q. Husain, Calcium alginate
starch hybrid support for both surface immobilization and entrapment of bitter gourd Momordica charantia peroxidase, J. Mol. Catal. B: Enzym. 57 (1) (2009) 164
170. [186] G. Bayramo˘glu, M. Yilmaz, Y.M. Arica, Reversible immobilization of laccase to poly-4-vinylpyridine grafted and Cu (II) chelated magnetic beads: biodegradation of reactive dyes, Bioresour. Technol. 101 (17) (2010) 6615
6621. [187] R.L. Reis, J.S. Rom´an, Biodegradable Systems in Tissue Engineering and Regenerative Medicine, CRC Press, Boca Raton, FL, 2004, pp. 359
367. [188] V.P. Torchilin, Immobilised enzymes as drugs, Adv. Drug Deliv. Rev. 1 (1) (1987) 41
86. [189] S.G. Wang, X. Jiang, P.C. Chen, A.G. Yu, X.J. Huang, Preparation of coaxial-electrospun poly [bis(pmethylphenoxy)] phosphazene nanofiber membrane for enzyme immobilization, Int. J. Mol. Sci. 13 (11) (2012) 14136
14148. [190] J.M. Park, M. Kim, H.S. Park, A. Jang, J. Min, Y.H. Kim, Immobilization of lysozyme-CLEA onto electrospun chitosan nanofiber for effective antibacterial applications, Int. J. Biol. Macromol. 54 (2013) 37
43.

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ENZYME IMMOBILIZATION STRATEGIES AND BIOPROCESSING APPLICATIONS

12

Emmanuel M. Papamichael and Panagiota-Yiolanda Stergiou Department of Chemistry, University of Ioannina, Ioannina, Greece

12.1 INTRODUCTION These magnificent biocatalysts, the enzymes, are macromolecules sensitive in denaturation and in most cases lost their activity due to a series of reaction conditions, including pH, temperature, ionic strength, presence of organic solvents, and high speed of agitation. When the rather costly enzymes are used to catalyze high-scale bioprocesses, their aforementioned sensitivities along with one single use of the biocatalyst constitute disadvantages with serious synthetic and economic consequences, which should be precluded [1]. These drawbacks are widely avoided through immobilization of the enzymes onto inert matrices. The immobilization of enzymes emulates their native mode of action in vivo, where in most cases they act being attached on various biomembranes [2]. The immobilized enzymes are reusable and make available the use of continuously fed flow reactors. In these cases, enzymes act as separated from both their substrates and products, which in turn can be transferred between the phases in biphasic reaction media [2]. Inert and insoluble matrices of various types, forms, and structures have been used for enzyme immobilization purposes. These tools of immobilization were found effectively stable versus the pH value and temperature of the reaction media; they enhance the catalytic efficiency of the immobilized enzymes and eliminate substrate and product inhibition [3]; nevertheless, this is not always the case. Therefore a variety of enzyme immobilization techniques have been reported and applied by depending on the immobilization matrix, the enzyme, and the positive or negative effect of immobilization on enzyme’s catalytic efficiency and stability [4]. Hence, we consider that it may be essential and/or prerequisite to search for valid and specific strategies of enzyme immobilization to facilitate and improve particular bioprocessing applications. Various enzyme immobilization strategies have been reported where novel noncustomary methods were applied, which may increase the catalytic efficiency of enzymes by improving their stability, reuse, and resistance versus inhibitors, etc. It follows that crucial structural and kinetic attributes of the biocatalyst under consideration are taken into account (e.g., catalytic effectiveness, faster operation, and multitasking function) [5]. An already applied immobilization strategy was based on the use of laboratory available appropriate solids, where the enzymes were immobilized, reversibly or irreversibly, through either physical or chemical methods; in that latter situation,

Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00012-0 © 2020 Elsevier B.V. All rights reserved.

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an additional supporting arm was commonly used [1,6]. Further strategies of immobilization comprised enzyme trapping, cross-linking, formation of enzyme aggregates [7
11], and other advanced strategies based on suitable design [12,13]. Usually, by enzyme engineering they are prepared enzymes which are more compatible with the immobilization solid supports, and thus are achieved higher yields in the course of a biocatalyzed reaction [14]. It should be pointed out that immobilization matrices must be chosen so that to fulfill crucial requirements such as stability during the reaction time, resistance to deterioration by water and/or organic solvents, flexibility in the mechanical stirring, and the type of the utilized reactor [15]. On the other hand, novel and/or usual immobilization strategies require suitable evaluation with respect to the followed reaction process [16
18]. Methodologically, the emerged strategies of immobilization are mainly distinguished in vitro multistep and in situ one-step schemes. In the former case, the enzyme is chemically cross-linked or adsorbed noncovalently onto a solid support and/or is encapsulated and surrounded by some polymer gel, etc. [19]. Recently, attention has been given to design necessary structural modifications of selected matrices in order to immobilize specific enzymes. Consequently, when the enzyme structure and mechanism of the reaction under consideration are well known, the application of immobilized enzyme would be economically and productively more successful [20]. In biochemical terminology, bioprocessing could be designated, generally, as the knowledge and practice in investigating, developing, and composing valuable products through biocatalytic treatment of industrial and other wastes. Foods and food additives, pharmaceuticals and cosmetics, biofuels, etc. are a few examples of the potential materials for rebuilding. Food making by means of bioprocessing is rather an ancient approach of creative transformation of specific wastes into valuable products [21,22]. The released wastes, largely originating from food and other related industries, as well as their by-products, contain compounds as monomeric and polymeric sugars, proteins, lipids, organic and inorganic salts, nonvolatile organic solvents, organic and inorganic acids, etc. A few of these wastes could not be characterized as environmental pollutants; on the other hand, the environment cannot assimilate the huge quantities of these compounds, harmful or not, which are streaming in the soil, rivers, and underground waters. Conversely, it is a pity to lose the previously mentioned key substances in the junk, and not to transform them through enzymatic catalysis into high-valued products, whereas they retain high economic value [23]. Among the prerequisites for the application of bioprocessing procedures is the thorough control over the entire followed enzymatically catalyzed synthetic process, which could also provide a continually scientific advance of the applied methodologies and their kinetics. A deeper knowledge of the kinetics will promote the research for new enzymatically catalyzed bioprocesses and their applications, in both conventional and nonconventional reaction media, in biphasic systems, ionic media, and so forth, for novel reactions’ management and increase the safety in food making [21,22,24]. Therefore selected enzymes could attain bioprocessing applications by both decomposing the available wastes and synthesize new useful compounds without the formation of by-products. In this work, we review in detail various strategies and methodological tools of enzyme immobilization in order to be utilized in bioprocessing applications.

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12.2 ENZYMES AS INDUSTRIAL BIOCATALYSTS Over the past thousands of years, human beings used enzymes empirically in making bread, cheese, wine, as well as in other everyday activities (e.g., legume seeds treatment, such as peas, beans, and lentils). Only early in the 20th century, scientists isolated, and started to exploit the secreted enzymes by microorganisms, as well as to understand their protein nature, whereas several decades later, enzymes are used in industrial manufacturing of beneficial products [25]. Globally, enzymes are well established as the all-purpose advantageous catalysts, due to their high specificity, exceptional reactivity, and effectiveness under unfavorable conditions. Enzymes, in contrast to the conventional inorganic and/or organic catalysts, are environment-friendly and fully biodegradable, although less stable and costly.

12.2.1 REQUIREMENTS, ADVANTAGES, AND LIMITATIONS FOR THE INDUSTRIAL ENZYMES 12.2.1.1 Stabilization of the enzymes The stabilization of enzymes and especially those used for industrial mass production is a matter of specific knowledge, research, and applications. Under certain conditions, and in both aqueous and nonaqueous (organic) reaction environments, the structure of enzymes may denature. Extremely acidic and/or alkaline media, reactions that take place at very high or low temperatures, solutions of high ionic strength, elevated concentrations of surfactants, presence or absence of metallic salts, etc. may influence strongly the structure and consequently the activity of enzymes. Numerous approaches have been elaborated, as devoted to the stabilization of enzyme structure, activity, and catalytic efficiency (e.g., enzyme engineering, modifications by means of chemical reactions, and different methods of enzymes’ immobilization in various inert matrices). In all of the aforementioned stabilization methods of the enzymatic catalytic power, potential unfavorable interactions may occur among the employed biocatalyst and the various used reactants. The common case is to stabilize enzyme molecules and their structures (secondary, tertiary, etc.); this task could be more effective by considering suitable inactivation models [26]. The simplest model is referred to the denaturation of enzymes’ native structure (ns) toward their inactivation (i), by passing through a reversible unfolded state (us), that is, ns "K us
!k i: The reversible reaction concerns the thermodynamic stabilization, while the irreversible reaction is referred to the kinetic stabilization of the enzyme molecule. Practical reasons suggest to focus on the irreversible step, which commonly proceeds through a first-order exponential decay according to equation At 5 A0e2kt, where At, A0, k, and t represent the activities of the enzyme under consideration at time t and time zero, the first-order rate constant, and the time, respectively. The estimation of the half-life t1/2 is a measure of the enzyme stability. Researchers usualy face destabilization of enzymes, which undergo denaturation, through complicated pathways. A double exponential decay model equation At 5 A0[Γe2αt 1 (1 2 Γ)e2βt] comprising three parameters and linear combination of the individual rate constants of enzyme destabilization, may describe these cases. In this equation, Γ, α, and β represent either complex functions or expressions of individual rate constants. If Γ 5 0, then the previous equation degenerates to a first-order exponential decay, whereas when β 5 0, it means that the enzyme molecule

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preserves a small part of its activity during the inactivation state [27]. The aforementioned theory may not provide an accurate description of the destabilization of a biocatalyst. Measurements that are more accurate have been achieved by means of specific bioreactors where enzyme and substrate react at a low temperature and after a period of time at some higher temperature; in both the previous situations, the activities of the enzyme are measured and compared, as well as the rate of reduction of substrate [28]. A similar methodology can be applied by examining various destabilization factors of enzymes’ structure. As we have already commented, the molecular occurrences of stabilization/destabilization
denaturation of enzymes depend on two dissimilar states, that is, the thermodynamic (to keep the active conformation of enzyme) and the kinetic (to avoid irreversible destabilization of the active conformation of enzyme within the time course of the reaction). Nevertheless, we should point out that this knowledge is crucial in keeping biocatalysts stable, which is a key task as far as it concerns the cost, competition, and further development of production in industrial and biotechnological scale [29]. Furthermore, one question: “what about the industrial and biotechnological requirements for their potential biocatalysts?,” is emerging. The enzymes, and because of their protein nature, will be advantageous biocatalysts in industry and biotechnology if and only if they may be kept on stable as concerning their native structure. Factors, which in general have been referred in the beginning of this part, such as the pH value, the concentrations of metallic salts, the temperature, the organic solvents, the surfactants, potential inhibitory concentrations of substrates, and/or known or unknown inhibitors, which are contained in the reaction media, could most likely destabilize enzymes’ structure, function, and activity. More specifically, in organic and/or nonaqueous media, the enzymes catalyze a huge variety of reactions based on the potential higher solubility of their substrates, as well as on advantageous thermodynamics. Efforts to stabilize the enzymes in these media have been focused mainly on proper modification of their protein molecules by well-studied methods. Furthermore, the accurate meaning of one particular factor, out of these mentioned just above, is customarily simplified, but not on purpose. Although the pH-value is valid for nonaqueous and/or nonpolar organic solvents, its definition and effect is substantially different in comparison to ordinary aqueous solutions.

12.2.1.1.1 About the pH value Generally, variations in the pH value of reaction media where enzyme catalysis is performed affect stability and activity of the biocatalyst. At extreme pH values (both acidic and/or alkaline), the enzyme structure is vastly affected, most likely irreversibly, leading to complete inactivation and destabilization of the enzyme molecule [30]. At the optimum pH value, the structure, and consequently the function and activity of the enzymes are the highest ones. These phenomena are explained due to ionization of side chains (pKa values) of the residual amino acids of the backbone of enzyme molecule. Therefore protonations and/or deprotonations of the catalytic residues of the enzyme molecule mainly occur, as well as ionization of the residues that occupy the enzyme’s binding pockets, may also take place. The pH profiles of important equilibrium and/or rate constants, which are calculated during an enzymatic reaction, usually are bell-shaped curves and are described by variations of the Henderson
Hasselbalch general Eq. (12.1). We have explained the parameters and the symbolism of Eq. (12.1) in detail earlier [31]. Next, the estimated pKas are informative for the identification of key residues of the enzyme molecule which affect its stability,

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function, and activity. However, the aforementioned are valid in aqueous reaction media, whereas in nonaqueous and/or organic media, the concept of pH differs essentially, as it has been explained earlier [31]. kobs 5

n X i51

kilim 11

n P j51

!

(12.1)

Bij

12.2.1.1.2 The contribution of temperature It should be emphasized from the beginning that the viewpoint, which generally suggests the use of higher temperatures to increase reaction rates, is scientifically false. One has first to detect whether or not the reaction under consideration is homogeneous or heterogeneous encompassing diffusional limitations; in that latter case, high temperatures may be fatal for the followed reaction. More specifically, by dealing with enzyme-catalyzed reactions, temperature variation needs additional consideration and handling [31]. Usually enzymes are sensitive at extreme temperatures, namely very low and very high ones, due to their protein nature. By increasing the temperature of the medium of an enzymatic reaction, a reversible gradual increase of its reactivity is observed, up to some upper limit; after that, the enzymatic reactivity is gradually decreased up to some lower limit. Further increase in temperature of the enzymatic reaction medium may cause irreversible destabilization of the enzyme molecule due to the destruction of its tertiary structure. These temperature variations are perceptible by drawing absolute temperature profiles of the Michaelis
Menten parameters (e.g., kcat, kcat/Km, and/or Km), and may be equally observed by decreasing the temperature of the enzymatic reaction medium. Therefore the concept to increase the reaction rate of an enzyme-catalyzed reaction, by increasing the temperature of the reaction medium, and vice versa, is true only under the abovementioned conditions and concerns, that is, by taking into serious account the stabilization of the employed biocatalyst.

12.2.1.1.3 Grouping methods for stabilization of enzymes Commonly, more than one method is required to stabilize an enzyme structure, function, and catalytic activity; besides, in cases where the biocatalyst is under denaturant conditions, a combination of various methods may be used. Various immobilization techniques coupled with protein engineering procedures have been successfully used comprising chemical modification followed by immobilization based on different types of solid supports [32
34].

12.2.1.1.4 Use of protein engineering The stabilization of enzymes by protein engineering is rather a complex task. Hence, various experimentally different approaches have been proposed aiming towards the improved functioning of the enzyme molecule at particular reaction circumstances. In this context, approaches are described, which were considered as relatively more efficient in stabilizing enzyme molecules for future biotechnological and/or industrial use [35,36].

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The impressive results from the application of the directed evolution technique in protein engineering, which is an iterative method, should be emphasized. This experimental approach requires only a rough knowledge or even no knowledge on the molecular structure of enzyme under investigation. The directed evolution is achieved by means of a random generation of a diversity of genes, their collection, and the investigation of the attained mutant
enzyme molecules. Subsequently, the best qualified enzyme mutant constitutes a basis for the next iteration until the achievement of the optimum one, by considering certain attributes [37]. When the researcher has access to information concerning the combination of structure and function of an enzyme, then the method of rational design can be applied. Suitable assumptions are put forward for consideration to reduce a diversity focused to decrease the number of the expected mutants, as well as their analyses. This method is valuable in cases where the protein engineering research is focused on the modification of either the surface and/or the catalytic site of the enzyme under investigation, and in order to enhance its reactivity [38,39]. Both the aforementioned directed evolution and rational design, as well as a certain combination of them, may be applied to promote the biocatalytic performance of industrial enzymes (e.g., stability versus pH, temperature, salinity, and nonaqueous reaction media) [40,41]. The approach of polypeptide chain extension is based on the addition of relatively small peptide chains to either the amino- and/or the carboxyl-edge of enzyme macromolecule. These extensions may help in improving the thermostability of the enzyme under consideration, as the extended polypeptide chains through random mutagenesis provide a thermostable enzyme structure, due to that it occupies a wider area. When a polypeptide chain extension results via random mutagenesis jointly with random point mutations, the enzyme which is formed through directed evolution possesses new attributes. Unfortunately, the polypeptide chain extension is not applicable to all enzymes [42]. The important technique of site-specific mutagenesis although provided so far many toward the enhancement and preservation of enzyme stability is however useful in relatively limited situations where the structure of the employed enzyme is known. This is the case where the investigation of many mutants is unfeasible. Furthermore, and regardless that in the literature are recently described applications of novel genetic methods, there are stuations where the enzyme molecule underwent chemical modification in order to be stabilized [43]. The development of new methods of enzyme mutagenesis leads to potential increase of the diversity records of the utilized natural amino acid residues. The number of these residues should be decreased by the use of suitable programming and/or laboratory probes of predicting the outcome of any amino acid substitution, and according to the expected result [37,44]. Furthermore, by the direct evolution, and through iterative algorithms, the properties of enzymes can be modified so that these biocatalysts to be more helpful in biotechnology and other applications. Conversely, the increased industrial demands require faster and cheaper protein engineering strategies such as that termed as KnowVolution (i.e., knowledge gaining directed evolution). The described protein engineering strategy aims to the collection and analysis of as much as possible information on the primary and secondary structures of the enzyme molecules. Furthermore, all enzyme structures (i.e., primary up to tertiary) are strictly related to their function and activity. Subsequently, the target of these pioneer protein engineering methodologies is the increased probabilities for production of properly improved enzyme mutants during every iterative cycle [44].

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12.2.1.2 Stabilization and functionality of biocatalysts in nonconventional solvents Nowadays, many biotechnological products are manufactured by means of enzyme-catalyzed reactions in nonaqueous media and/or organic solvents. Therefore the stabilization and the catalytic function of enzymes in these environments along with their specific and/or increased reactivity is the requisite. Hence, currently, efforts have been made in the development of specialized technologies on the improvement of both the stability and the catalytic performance of industrial enzymes to function in nonconventional media [45]. There is a good reason for that. In nonaqueous media and/or organic solvents, in both polar and nonpolar, numerous substrates are fully soluble, as well as the synthetic action of enzymes is favored thermodynamically versus the hydrolysis [2]. Synthetic processes of this kind are advantageous as the biocatalyst exhibits higher selectivity, reactivity, and reaction rates; additionally, they are considered as green and thus are preferred. Nevertheless, the key of these methods’ success is to pay attention in keeping the structural and functional stability of the used biocatalyst [46]. In any case, the reader should recall in memory that all free enzymes, after their isolation and purification procedures are found in aqueous media and are enveloped by a hydration (solvation) shell. A hydration shell still also exists around lyophilized enzyme molecules [47], which undergo a mutual dehydration and partial solvation when they found in organic solvents. This later process strongly affects the structure and stability of the enzyme molecule, and it is more profound in hydrophobic organic solvents than the hydrophilic ones [48]. Therefore and after all this information it sounds rather necessary for the researcher to think more techniques, which will help in a decisive way in using stabilized enzymes in nonaqueous and/or organic media, as it is commented below.

12.3 ENZYME IMMOBILIZATION As it has been mentioned in Section 12.1, enzymes are highly expensive chemicals, whose cost potentially slows down the industrial and technological development [1]. Furthermore, the enzymes, which are sensitive macromolecules, should be operated carefully; this is not an easy task. For these reasons are required quick and efficient solutions to this drawback. Enzyme immobilization is the concept, which so far offered advantages as it is the recycling and/or the long-time continuous application of these extraordinary biocatalysts for manufacturing purposes [4,13]. Nevertheless, the enzyme immobilization processes are costly, and thus the employed immobilization techniques should fulfill at least several fundamental characteristics, that are: (1) simplicity; (2) low cost of the immobilization matrix, and the process; (3) stability, longstanding, recycling, and easy recovering of the immobilized enzyme; and (4) maintenance of activity and selectivity of the immobilized enzyme. Recently, the enlarged industrial requirements demand immobilized enzymes possessing additional attributes such as low toxicity, extended use in multiple reaction systems, as well as for various industrial applications [26,49]. Moreover, and in all cases, immobilized enzymes should act quite similar versus their free form, although enzymes in free form are much more stable than immobilized ones (e.g., the rate of self-hydrolysis of immobilized proteases approaches zero) [13].

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12.3.1 CHANGES INDUCED BY ENZYME IMMOBILIZATION: POSITIVE AND NEGATIVE 12.3.1.1 Features due to enzyme immobilization The enhancement of stability, function, and stereoselectivity of immobilized enzymes versus temperature, pH value, solvent, metallic cations, surfactants, time of storage, etc., which follow their immobilization, are considered as significant. Usually, in extremely high and/or low temperature or pH value of the reaction media, free enzyme molecules are subject to irreversible inactivation, mainly through unfolding; similar denaturation of the free enzyme molecule occurs during their maintenance in specific organic solvents or in aqueous media, containing high metallic salts concentrations. For industrial and biotechnological production plans, it gets of great importance to prevent various kinds of irreversible enzyme inactivation due to the use of these biocatalysts in immobilized form; in this way, different immobilization strategies have been developed. The employed and ever-renewed strategies and methods of enzyme immobilization contribute positively to the stabilization, improved function, reuse, and other beneficial properties of these biocatalysts. On the other hand, immobilization should not be faced as the panacea to solve all the raising negative aspects of biocatalysis in laboratory and industrial scale. The basis of immobilization, that is, the solid immobilization matrix, might become source of unfavorable results affecting the structure and thus the stability, reactivity, and selectivity of the immobilized enzyme. Although the methods of immobilization will be discussed below, however, they play a crucial role for successful enzyme immobilization. It may be the material of the immobilization matrix and its interaction with the enzyme molecule, the mode of enzyme binding onto the matrix, the bonding network between enzyme and matrix, the use of spacer, and the degrees of mobility of the immobilized enzyme molecule [50].

12.3.1.2 Immobilization improves the activity of an enzyme Despite the fact that native enzymes may catalyze only a limited range of reactions, the question is whether it is likely that these biocatalysts may be improved toward their usefulness in industry and biotechnology. Moreover, these immobilized enzymes exhibit reduced activity as compared to their native form most likely due to the restricted accessibility by their substrates. Recently, encouraging results have been reported, which ascertain that there is no rule without exceptions and that immobilized enzymes displayed higher catalytic activities than in their free form [51]. These results were explained on the basis of properly chosen matrices versus the immobilized enzymes, their substrates, and the reaction media. More encouraging results concerning higher catalytic performances of immobilized enzymes were attributed to favored orientation of the immobilized biocatalyst so that to be in closer contact versus its substrate [50]. The aforementioned results were obtained subsequent to extensive inspection and designed optimization of a range of factors affecting enzyme immobilization and function (i.e., temperature and pH value of the immobilization medium, the material of the used matrices and the immobilization method, the bonding network between matrix and immobilized enzyme, etc.) [52].

12.3.1.3 Improvement of specificity and selectivity of the immobilized enzyme The selectivity of industrial biocatalysts is considered lately among their important features as far as it concerns the enzyme-catalyzed enantioselective synthesis of chiral chemical compounds,

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and the formation of different percentages of diastereoisomers. The nowadays chemistry faces elevated demands of production of diastereoisomers especially in the field of medical therapy; the pharmaceutical diastereoisomers possess different biological reactivity [50,53]. Immobilization of enzymes by adsorption, entrapment, and by covalent bonds on proper matrices may adjust their selectivity and specificity. Additional factors, which affect positively the catalytic efficiency along with the enantioselectivity of the immobilized enzymes, are the nature of the immobilization matrix, the temperature, pH value and water activity (in biphasic media) of the biocatalyzed reaction; these factors create various microenvironments, in the vicinity of the immobilized enzyme, which alter the nature of the enzyme
substrate interactions [54].

12.3.1.4 Enzyme stabilization through partitioning It is well known that when a solid matrix, and independently of carrying an immobilized biocatalyst or not, when it is immersed in a solution a Helmholtz
Stern double layer is formed creating a partition of hydrophobic and hydrophilic microenvironments. This property has been used as a stabilization probe against molecules, which may inactivate the immobilized biocatalyst. When the solid matrix, which is supported by the immobilized enzyme, is immersed in binary reaction media containing both water and organic solvents, then a decrease of the enzyme activity could be observed due to a potential modification of the enzyme structure [55].

12.3.1.5 Restrictions due to diffusional effects Diffusional effects are observed in systems where solid matrices are immersed in water solutions, and thus, Helmholtz
Stern double layers are formed. These phenomena should be studied by taking into account the reaction mechanism, which was followed in the absence of diffusional effects, as well as by means of the well-known Fick’s laws of diffusion, and the assistance of earlier reasonable assumptions [31]. Furthermore, the researcher may consider that under the above circumstances’ basic entities, such as the pH value, the substrate and product concentrations hold on unusual meanings due to their gradient transport through the double layer. The values of these later entities along with the enzyme’s activity differ substantially in the bulk solvent versus those on the surface of immobilized enzyme; however, the thermodynamics of the system remains unaffected. The method of enzyme immobilization (e.g., adsorption, entrapment in a porous matrix, and covalently bonded on the matrix surface through a spacer) strongly contribute on either to simplify or to complicate the occurrence of the diffusional effects.

12.3.2 STRATEGIES, STEPS, AND FACTORS AFFECTING THE ENZYME IMMOBILIZATION 12.3.2.1 Preimmobilization factors and actions The protein nature of the enzymes warns on their sensitivity, as well as on the physicochemical modifications, which will occur and affect these molecules after immobilization. The established microenvironment in the vicinity of the immobilized enzyme, the resulting changes which were discussed above, the enzyme
substrate and enzyme
product interactions, all they affect the stability, and potentially the kinetic behavior of the immobilized biocatalyst. Additionally, the

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immobilization matrix may positively or negatively contribute in sustaining the secondary and tertiary structures of the immobilized enzyme through hydrogen and/or covalent bonding among them. Therefore the followed enzyme immobilization method and procedure should focus on maintaining the potentially higher percentage of activity of the biocatalyst. The above may be achieved by the proper choice of the immobilization matrix in order to fulfill particular requirements concerning the immobilized enzyme, the suitable method of immobilization and the optimized design of the reaction conditions [56].

12.3.2.2 Beneficial steps of enzyme immobilization In general, the following steps and actions are ensued toward the immobilization of an enzyme: 1. The proper matrix should be chosen according to the chemical nature of its material and surface, as well as its porosity. 2. The running of an optimized design is necessary concerning the requirements of the biocatalyzed reaction (i.e., method of immobilization, reaction medium, activity and stability of the immobilized enzyme, details of the experimental conditions, product yield, reaction time, and temperature), which will be monitored, according to the used enzyme and its substrate [57]. Nevertheless, additional attention should be given on the abovementioned steps and actions as far as it concerns the influence and enhancement of leading concepts such as the structural stability, reactivity, and the reuse of the immobilized biocatalyst [57].

12.3.3 SELECTED METHODS OF ENZYME IMMOBILIZATION By taking into account the experimental and theoretical origins of the reported standard strategies, which are utilized for enzyme immobilization, one can objectively distinguish their advantages and disadvantages. Criteria of choosing the most appropriate strategy, as well as the experimental methodology for enzyme immobilization, have been described above; specific biochemical and physicochemical factors should be fulfilled, which concern the interactions between the immobilization matrix and the immobilized enzyme [58]. These standard strategies might be characterized according to their chemical and physicochemical basis into four groups as depicted in Fig. 12.1. 1. Adsorbing methods based on weak-bonding interactions: Although are simple and inexpensive methods leaving the enzyme unaffected, however, desorption of enzyme is likely. 2. Methods based on covalent bonds between reactive groups of the immobilization matrix and the immobilized enzyme: Enzyme destabilizing methods. 3. Entrapment methods, where the enzyme is naturally restricted within a polymeric porous network of the immobilization matrix, allowing the substrate and products to cross, though retain the enzyme intact. Highly applied methods exhibit diffusional effects. 4. Methods based on the formation of cross-linked aggregates of the enzyme molecules with a proper reactant (functional and/or dysfunctional): Expensive methods, which stabilize the immobilized enzyme exhibiting diffusional effects.

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Methods of enzyme imobilization

Irreversible methods

Encapsulation and microoencapsulation

E E

E

E E

E

Reversible methods

Entrapment Covalent Cross-linking methods methods bonding methods E E E E E E E E E E E E E E E E E E

Adsorbing methods E E E E E E

FIGURE 12.1 Schematic representation of enzyme immobilization methods, where E stands for a molecule of free enzyme, as modified from Sirisha et al. [20].

12.3.3.1 Adsorbing methods The immobilization is achieved by immersing the chosen matrix in an aqueous solution of enzyme for a predetermined time through appropriate experimental design. The aqueous solution contains the necessary reagents, according to the references, in order to maintain the enzyme activity intact. Subsequently, the matrix is removed from the aqueous solution, is washed with a suitable buffer to remove the free enzyme molecules, and in most cases is dried by lyophilization and stored under anhydrous conditions. The immobilized enzyme by adsorption is unstable in solutions of extreme pH values and/or ionic strength due to partial enzyme desorption. Moreover, in the adsorption methods, it is likely a partial reduction of the catalytic efficiency of the immobilized enzyme due to a potential blockage of its catalytic site. Although, a huge variety of natural materials have been used as adsorption matrices, which are friendly to the environment, however, efforts were reported on the chemical modification of natural matrices to improve their adsorbing properties [59].

12.3.3.2 Covalent bonding methods Enzyme immobilization onto suitable matrices by means of covalent bonds usually follows activation of the matrix. Frequently, this activation comprises the covalent binding of a linker or spacer onto the matrix surface. The linker or spacer may be glutaraldehyde, epichlorohydrin, different oligomeric compounds of glyoxal, which form Schiff bases with available side-chain amino group of the enzyme, side chains of amino acid residues such as of lysine, cysteine, aspartic and glutamic acid, and histidine. Subsequently, the enzyme appropriately reacts covalently and irreversibly with the activated linker [44,59]. A variety of linkers have been used, which are appropriate for matrices of different materials, as well as different immobilization methodologies were employed in order to match the used enzyme. Although in the chemical affinity methods only few quantities of enzymes are

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immobilized, however the stability of the immobilized biocatalyst is increased whereas its activity may be decreased [59]. Obviously, elevated activities of the immobilized enzyme are observed as much as its catalytic residues remain unaffected by the binding with the linker. Finally, increased stabilization of the immobilized enzyme may be achieved through multipoint covalent bonding [60].

12.3.3.3 Entrapment methods Immobilization of enzymes by means of the entrapment methods is considered as irreversible; they have been recognized as the easiest ones, without charging the enzyme with structural modifications. Besides, entrapment methods deliver poor biocatalysts, which exhibit diffusional limitations [57]. To some extent, encapsulation and microencapsulation are similar methods, where the biocatalyst is locked in the inner space of semiporous spheres, of micrometer scale [50]. The entrapment methods of enzyme immobilization may be carried out easily as follows: in an aqueous solution of the enzyme is added a solution of the chosen monomer, which will be polymerized chemically through specific catalysts and/or by adjusting the experimental status. It should be taken into account that both, the chosen monomer and its polymeric form, to be inert versus the enzyme. These methods are advantageous as far as it concerns the stability and the well-reduced leaching and denaturation of the immobilized enzyme. An additional advantage of the entrapment methods is the capability of optimizing the entrapment conditions so that to manage the microenvironment of immobilized enzyme, as well as to have limited control on its optimal pH value and other parameters [59].

12.3.3.4 Cross-linking methods Cross-linking techniques comprise another group of irreversible enzyme immobilization methods. They are based on the development of three-dimension intermolecular cross-linked structures, which connect covalently the molecules of the enzyme to be immobilized [61]. These methods are designated by the absence of immobilization solid supports, which allow the development of stable, more active and less costly immobilized biocatalysts [12,62]. In the case of cross-linking methods, linkers are required similar to those described in the covalent bonding methods, excluding glutaraldehyde, which is not recommended due to that it might introduce a reduction of activity of the immobilized enzyme [12]. The reported cross-linking immobilization procedures may be distinguished into two types; they are performed either via crosslinked enzyme aggregates (CLEAs) or via cross-linked enzyme crystals (CLECs). In the former case (CLEAs), the immobilized biocatalysts function also in aqueous media, and they are stable against elevated concentrations of metallic salts, organic polar, and nonpolar solvents. The immobilized biocatalysts of the latter case (CLECs) have been considered as very stable. CLEAs are prepared by dissolving the selected enzyme in appropriate aqueous buffer and at low temperature; then the aggregation is performed by the stepwise addition of an aqueous solution of either (NH4)2SO4 or polyethylene glycol (PEG). The oversaturated solution is maintained at the low temperature for about 24 h, whereas the aggregates may be separated by centrifugation. Subsequently, a sufficient quantity of the enzyme aggregates is added in the solution of a chosen linker, in n-propanol, to undergo cross-linking under mild agitation on a suitable platform at room temperature. The excess of the linker is usually removed by washing the preparation of the crosslinked enzyme by the same buffer, where the CLEAs were formed. In the case of using PEG

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instead of (NH4)2SO4, it is necessary to add a small percentage of a surfactant in the solution of the preparation of the CLEAs [63].

12.3.3.5 Materials used in manufacturing immobilization matrices The chemical and structural features of an immobilization matrix are key factors, which affect the operative value of the immobilized enzyme as biocatalyst. The reader understands that a wrong choice of the immobilization matrix could be destructive for the enzyme as well as costly; the opposite is beneficial and it contributes to the development of at least a biochemically and kinetically adequate immobilized biocatalyst [62]. Moreover, a matrix which might be designated as the most appropriate among many should be inert versus the enzyme, compatible to be used in sensitive industrial branches such as food, pharmaceutical, and cosmetic and to be available at low cost [64]. Therefore the perfect choice of the matrix material sounds difficult, and it leans rather on the properties and the kinetics of the enzyme under immobilization [65]. According to the cited experience, the immobilization matrices should meet certain features as the following [20]: 1. Inexpensive, easily available, environmentally friendly, thoroughly inert, stable, and improving the specificity and reactivity of the immobilized enzyme. 2. Be tolerant versus temperature, pH value, mechanical stress, and organic solvents, thus yielding stable immobilized biocatalyst, under various situations. 3. Be regenerative after the period of life of the immobilized biocatalyst. 4. To load relatively elevated amounts of the enzyme. 5. To deliver disinfectant properties. The immobilization solid supports are grouped as inorganic or organic, whereas both of them may be distinguished in natural and/or synthetic (polymeric). Furthermore, and beyond the chemical nature of the immobilization matrices, their physical attributes (e.g., pore and particle size, and diameter, resistance in mechanical stress and rigorous agitation) are significant, and they may facilitate or disturb, accordingly, the functioning of the immobilized biocatalyst. The physical attributes of the immobilization matrices are decisive, and they will affect the kind of an employed reactor, whenever it is necessary. For example, porous matrices usually exhibit many diffusional limitations, though the enzyme is located in an environmentally protected area [66]. Dealing with materials used in manufacturing immobilization matrices, it should be pointed out a specific technique in this field; it is the immobilization of enzymes by means of nanoparticles. This later technique is beneficial by the reasons that: (1) it is based on an easy and simple synthesis of nanoparticles, which contain the immobilized enzyme, though without surfactants and other toxic compounds; (2) potentially, the size of nanoparticles may be fitted within the experimental boundaries;, and (3) the achievement of equally sized nanoparticles is quite possible [67]. In Table 12.1, various natural and artificial materials of inorganic and/or organic origin are summarized, which have been used for the development of immobilization matrices. In summary, therefore, three essential issues should be taken into account for the development of immobilized biocatalysts, that is, the proper selection of: (1) matrix, (2) method, and (3) conditions of immobilization. These later issues contribute effectively in the formation of immobilized enzymes with particular biochemical and physicochemical properties, which determine their applicability to specific industrial and biotechnological processes.

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Table 12.1 A collection of materials, which have been used for the manufacture of immobilization matrices. Inorganic

Organic

Natural: Bentonite, silica (sol
gels, hydrogels, celite), hydroxyapatite, and natural ceramics

Natural polymers: Polysaccharides (cellulose derivatives, dextrans, starch, agar, agarose, chitin, alginate), proteins (collagen, albumin), and carbon (active charcoal) Synthetic polymers: Polystyrene, modified sepharose, polyacrylates, polymethacrylates, polyacrylamide, vinyland allyl- polymers, active membranes, smart polymers, protein-coated polymers Processed polymers: Cellulose and carbon nanoparticles, nanofibers, nanotubes, and nanocomposites

Processed: Artificial ceramics, glass (porous, nonporous, coated), controlled porous metal oxides, and magnetic nanoparticles

12.4 APPLICATIONS OF IMMOBILIZED ENZYMES IN VARIOUS BIOPROCESSES Nowadays, bioprocessing deals with the modification and exploitation of useless and/or pollutant derivatives, which are by-products of various biochemical and technological activities, and they contain handy compounds. These bioprocesses are performed through the use of either living cells or their secreted biocatalysts (enzymes), in order to convert the aforesaid by-products into functional and effective, materials, feedstocks, new enzymes, etc., for further industrial manufacturing. Our specific interest, which will be commented below, focus on the requirements and application of bioprocesses based on the utilization of immobilized enzymes, their recycling competence and appropriate use in various areas such as: (1) industrial and biotechnological production (e.g., pharmaceuticals, foods, winemaking, and textiles) and (2) analytical and synthetic chemistry and biochemistry, in laboratory scale [20,68].

12.4.1 FOOD, DAIRIES, JUICES, COSMETICS, PHARMACEUTICALS, AND OTHERS The use of immobilized biocatalysts in food processing is beneficial in saving time and cost; besides, the processed food is easily recovered from the reactor and it is free of contaminants. Examples of food bioprocessing by using immobilized enzymes are as follows: 1. The hydrolysis of starch by means of α-amylases (EC 3.2.1.1) immobilized on calcium alginate matrix prevents the formation of gelatin subproducts [69]. 2. Many polysaccharides are selectively hydrolyzed by immobilized glucoamylases (EC 3.2.1.3) at consecutive scissile glycosidic α-1,4 and α-1,6 bonds, and at the nonreducing ends of the polymer chains, as it is represented in (Fig. 12.2). Glucoamylase and its immobilized forms have found a broad use as safe food hydrolytic biocatalysts. Potential results of the biocatalytic action of immobilized glucoamylase are different syrups of glucose or fructose and/or other

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FIGURE 12.2 Representation of a part of the molecular structure of a polysaccharide; immobilized glucoamylases hydrolyze selectively glycosidic α-1,4 and α-1,6 bonds.

hydrolysates, according to the hydrolyzed polysaccharide [70,71]. Significant contributions of immobilized amylases, glucoamylases, and other glucosidases have been reported and found useful, in the industrial isomerization of manufactured glucose to the sweeter fructose [72]. 3. Specific hydrolytic enzymes, the pectinases, were engaged in the processing of juices from fruits and other vegetables (e.g., tomatoes and pomegranates), according to their specificity in hydrolyzing α-1,4 bonds or removing acetyl and methoxyl protective groups of pectin substrates. Immobilized pectinases have been used in industrial scale reactors, and in low-cost processes of refinement, mainly, of fruit juices, by decreasing the juice thickness [71]. 4. A few more years ago, the milk whey, which contains a high percentage of lactose was considered as a pollutant by-product of dairy industries, triggering serious ecological concerns; the presence of lactose in the environment is related with elevated biological oxygen demand and chemical oxygen demand [73]. Nowadays, the biocatalytic action of immobilized β-galactosidases on the substrate milk whey, in continuous bioreactors, it outputs lactose hydrolysates (i.e., galactose and glucose), which contribute in the improvement of sweetness in many manufactured food additives [74]. Moreover, and through standardized use of the biocatalytic action of immobilized β-galactosidases, novel processed, lactose-free dairy products, including lactose-free milk, are now available in the market, as the intolerance of lactose among humans is common. Whole milk passes through a column consisting of immobilized β-galactosidase; the eluted milk contains only glucose and galactose. People who inherently lack the proper enzymatic system in order to transform galactose to glucose may not digest galactose; the aforementioned system comprises the enzyme galactose mutarotase, galactokinase, galactose-1-phosphate uridyl transferase, Uridine

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diphosphate-galactose-4-epimerase. If galactose is not transformed into glucose in the organism, then it reduces to galactitol, a poisonous hexanol, the excess of which leads to the development of cataract in the lens of eyes [71,75]. 5. Immobilized proteolytic enzymes, which are considered as safe, have been also employed in the fermentation and process of foods, and in the manufacture of cosmetics and pharmaceuticals. A classic example is papain, a plant protease, which is found unique as tenderizer of beef meat, through the hydrolysis of the protein fibers. Proteases, and especially in their immobilized form, are recognized as key enzymes in the manufacture of wine, brewing, bread, milk coagulation, etc. Essentially, proteases add value to foods and food additives by enhancing their flavors, aromas, and texture, as well as may improve their dietary significance [72,76]. The immobilized proteases have been applied successfully in the industrial processing of leather [71].

12.4.2 MEDICAL APPLICATIONS OF IMMOBILIZED ENZYMES AND BIOSENSORS There are many advantages to employ immobilized biocatalysts in medical applications, rather due to they are widely suitable, efficient, less costly, stable, and reliable. For example [70], immobilized enzymes may be useful as: 1. Probes in clinical analyses of various samples (i.e., blood serum, sputum, urine, gastric fluids, etc.). 2. Specific biosensors for the quantification of crucial measurable diagnostic metabolites (e.g. certain previously applied chemical methodologies) have been already replaced. These probes of immobilized biocatalysts are specific also in qualitative analysis of pesticides, heavy metals, organophosphorus chlorine, etc., where free enzymes may be denaturized, whereas the immobilized ones are tolerant [77]. Biosensors are specific and sensitive in detecting and measuring near to zero concentrations of the chemical constituent under consideration, as well as they distinguish the targeted one among many; biosensors are also less costly compared to other probes, and ecologically accepted [78]. An example is the glucose oxidase—β-D-glucose: oxygen-1-oxidoreductase (EC 1.1.3.4)—immobilized on polycarbonate membrane via glutaraldehyde-activated poly-(L-lysine) [79]. 3. Utilities for the treatment of certain diseases and development of relative pharmaceuticals, where the research in this field tries to take advantage of the biocatalytic power of immobilized enzymes [80]. Therefore and by targeting to particular achievements, they have been developed different immobilized biocatalysts, dedicated to the treatment of the following: 1. Thrombolysis, thrombophlebitis, phlebitis, and venous thrombosis, using streptokinase (EC 3.4.99.0) immobilized in gelatin gel; the complexes of streptokinase with the human plasminogen produce hydrolytically plasmin [81]. 2. Liver illnesses, using yeast alcohol dehydrogenase (EC 1.1.1.1) immobilized on magnetic nanoparticles of Fe3O4 activated by carbodiimide. Nevertheless, this treatment scheme was applied to mice and it was characterized as promising, only after a continuous research [82].

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12.4.3 BIOREMEDIATION METHODS AND WASTEWATER TREATMENT Billions of tons of various urban, industrial, and agricultural pollutants, that is, household garbage, organic and other chemicals, accumulated heavy metals, azo-dyes, pesticides, different agricultural wastes, hydrocarbons, halogenated compounds, wastewaters, etc., are produced and released untreated in streams, rivers, and sea, and they are transformed to toxic. The biocatalytic degradation of the aforementioned pollutants by means of immobilized oxidative enzymes has been applied as an effective alternative versus the use of living microorganisms [70]. These procedures, which generally may be termed as bioremediation, are referred to the treatment of polluted soil, subterranean zones and waters, by means of the systematic degradation of the implicated pollutants. Examples of bioremediation approaches are the following: 1. Immobilized plant and fungi laccases (EC 1.10.3.2), neutralize phenolic compounds and other similar pollutants, through the cleavage of their aromatic rings, and according to the general reaction R1OOR2 1 2e2 1 2H13R1OH 1 R2OH. A representative Laccase-catalyzed bioremediation reaction is depicted in Scheme 12.1 [83,84]. 2. Immobilized peroxidases (E.C 1.11.1.7) are active on organic hydroperoxides (R-O-OH) substrates according to the example reaction in scheme 12.2 [85]. 3. Immobilized azoreductase (EC 1.7.1.6) which were employed in the degeneration of various azo-dyes, nitro-aromatic compounds, etc.; the initial stage is the reduction of azo bond leading to the total collapse of azo dye. Azoreductases are oxidoreductases using as coenzymes NAD1 or NADP1; in the case of immobilized azoreductases, the coenzymes act as dissolved in the liquid phase. Accordingly, the reaction in Scheme 12.3 is a representative example of the oxidative activity of azoreductases [86,87].

OH R

O2

H2O

R

OH

COOH C(CH3)3

Laccase C(CH3)3

O OH

SCHEME 12.1 Laccases degrade phenolic compounds, through the cleavage of their aromatic rings.

R O OH +

NH2 Peroxidase NH2

R OH+H2O

+

NH NH

SCHEME 12.2 Representative reaction of the oxidative activity of peroxidases on hyperoxides; in this reaction, the coenzyme is the o-phenylenediamine.

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R2 N R1

N

+ 2 NAD(P)H

Azoreductase

R2 NH2 + H2N

+ 2 NAD(P)+

R1

SCHEME 12.3 Representative reaction of the oxidation of an azo-dye by azoreductase; in this reaction, the coenzyme is either the NADH or NADPH.

12.4.4 VALORIZATION OF FOOD PROCESSING WASTES—OBJECTIVES AND PROSPECTS Food industries and agroindustrial units, as all the manufacturing activities, produce wastes and by-products, which contain enough quantities of compounds covering the whole spectrum of the basic foodstuff (carbohydrates, lipids, and proteins). The previous should not be considered as just garbage, and apart from the fact that they affect the environment negatively, they include a huge economical value, which might be recovered through their appropriate transformation [23]. These wastes and by-products have complex structure and are mostly polymeric chemical compounds, which highly charge the environment; thus immobilized enzymes are matched as the best solution for their valorization [88]. The accomplishment of the aforementioned valorization may be performed through the relative basic methodologies of Section 12.4.2, though some additional enzymatic processes could be acknowledged. Esterification of organic fatty acids and alcohols, as well as esterification of carbohydrates and specifically starch, has been reported earlier by means of immobilized enzymes in biphasic, nonaqueous organic media, and in solvent-free systems; transesterification processes of lipids and fats have been reported as well. The results of these valorization activities comprise the formation of valuable products useful in everyday needs as biodiesel, biofuel, surfactants, soaps, biodegradable plastics, and adhesives [2,89
92]. The nowadays demands promote the research of novel modification methods of the industrial and agroindustrial by-products and wastes, in order to enhance their economic sustainability, exploitation, as well as the environmental protection along with their valorization processes. Hence, attempts were focused at first to reduce the cost of construction of immobilized enzymes whose about 50% corresponds to the cost of immobilization matrix; then, cheaper matrix material was chosen (e.g., cellulosic matrices), as well as matrix-free immobilization methods were established (e.g., CLEAs), and were described in Section 12.3.3.4. Nevertheless, these are not adequate. Additional research showed that a further stabilization of the structure of immobilized enzyme, along with a deeper knowledge of its kinetic mechanism, and the reaction conditions, are necessary towards a low-cost and effective biocatalytic process [93].

12.4.5 IMMOBILIZED ENZYMES IN TEXTILE INDUSTRY The requirements of using immobilized oxidoreductases (laccases) in the textile industry originate, first, upon the much higher cost of the previously employed chemical treatments; in our time, industries and public opinion take into account the environmental, as well as the cost of processing of the harmful chemical wastes. Additionally, the use of immobilized enzymes contributes

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235

significantly to the advantageous enhancement of the industrial yield of the processed textiles versus the past nonenzymatic treatments [94]. It seems likely that both the previous reasons prompted the research, as well as and the full application of the biocatalyzed processes of textiles through immobilized enzymes. Similarly, immobilized oxidoreductases have been applied to other industrial branches, that is, of paper and pulp [95].

12.4.6 IMMOBILIZED ENZYMES IN THE PRODUCTION AND PROCESS OF COSMETICS As cosmetics are considered various products, which appear in the market through different forms (e.g., creams, foams, solutions, gels, powders, and shampoos) and are designed for specific reasons, that is, for the care of face, lips, skin, hair, etc.; cosmetics are sensitive in long-time preservation and thus should be produced continuously and in relatively small quantities. In the cosmetics industry, the lipases are largely involved. Lipases are able to hydrolyze and synthesize esters, depending on the employed solvent system and the reaction conditions [2,31]. Additionally, lipases play a double role in the manufacturing of cosmetics, as they are both the proper biocatalysts and working components as well. The enzyme lipases are suitable for the care of sensitive tissues (e.g., the face), as well as are used as retarding agents dedicated for the regulated delivery of specific forms of cosmetics in situ. By using immobilized lipases in the production and process of cosmetics, the formation and release of by-products into the environment are decreased, and along with their nontoxic and biodegradable nature have evolved the cosmetics industry as a green one [96].

12.4.7 BIOPROCESSING THROUGH NANOBIOCATALYSIS The use of nanoscale matrices for enzyme immobilization is currently steadily grown and introduces new enhanced biocatalysts. This progress is succeeded by the immobilization of enzymes on the nanomatrices, and it secures their structure, as well as their selectivity and kinetics versus substrates and products. Furthermore, this approach makes available the immobilization of much more enzyme molecules on the same quantity of the matrix material due to its elevated surface area, put forward many advantages concerning various bioprocesses [97]. Nanotubes, nanoparticles, and nanosheets are several of these new immobilization matrices [98
101], which offered suitable biocatalysts used in various biotechnological applications mostly at laboratory scale [76].

12.5 CONCLUSIONS AND PERSPECTIVES In this work, various enzyme immobilization strategies and methods were reviewed and described in detail, which originated as a result of the application of different chemical and/or biochemical backgrounds. Moreover, it has been elucidated that enzyme immobilization is more or less costly, complex, and not at all an easy issue, where numerous factors and demands are involved which should be fulfilled at the accomplishment of this process. These latter requirements are related firmly to the sustainability of structural and functional stability, selectivity, activity, thermal and

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pH value tolerance, as well as to the environmental acceptability of the immobilized biocatalyst. In contrast, the kinetic behavior may be different from those exhibited by the free enzyme form, due to dissimilar reaction conditions of using free and immobilized enzyme. An emphasis has been given to both the beneficial use and the drawbacks due to the use of each of the described immobilization methods, as well as depending on the material of immobilization matrix. Furthermore, herein were described in details selected cases of applications of immobilized enzymes in various bioprocesses concerning foods, dairies, pharmaceuticals, medical applications, biosensors, bioremediation methods (in treatment and valorization of food processing wastes), textile industry and cosmetics. Special emphasis has been given to the bioprocessing through nanobiocatalysis. The vast biotechnological progress, which was emerged due to the use of immobilized enzymes, will find potentially a further evolution toward an expanding field of applications in foods, chemicals, agriculture, fuel, more efficient control of environmental protection, etc. This evolution will be much more rapid and targeted, as far as new theories and practices will be elaborated in the detailed designed immobilization of enzymes.

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2986.

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13

Anil Kumar Patel1, Pooja Dixit2, Ashok Pandey3 and Reeta Rani Singhania4 1

Department of Chemical and Biological Engineering, Korea University, Seoul, Republic of Korea 2DBT-IOC Centre for Advanced Bioenergy Research, Indian Oil Corporation R&D Centre, Faridabad, India 3Centre for Innovation and Translational Research, CSIR-Indian Institute of Toxicology Research, Lucknow, India 4Center for Energy and Environmental Sustainability-India, Lucknow, India

13.1 INTRODUCTION The world is approaching on the edge of irreversible damage to the global environment caused by climate change. The continuous rise in emissions of greenhouse gas (GHG) due to an everexpanding population’s demand for energy and other utility is a major cause for concern. If this growth remains uncontrolled, the consequences could be catastrophic for the global ecosystem. Globally, there is an alarming situation that has led researchers to look for possible solutions to reduce GHG emissions. In this context, renewable resources could play a tremendously positive role to move humankind toward sustainable solutions. Lignocellulosic biomass is foreseen as the most abundant renewable resource, which could be utilized for the benefit of mankind. This group of biomass includes surplus biomass that does not have food or feed applications. This biomass is comparatively rigid in structure and consists of several components such as cellulose, hemicelluloses, lignin, and pectin. It can be hydrolyzed chemically as well as enzymatically. As we envisage having an eco-friendly environment and moving toward sustainable solutions for the future; enzymes would be the first choice being green. These are the natural catalysts, which are highly specific. Biomass can be hydrolyzed into simple sugars by using enzymes, which can further be employed for conversion into useful products for various purposes, for example, ethanol. Ethanol from biomass has revolutionized the present era, specifically due to its application as liquid fuel. Our dependency on fossil fuels has led the way to reach the present deleterious situation. These are nonrenewable resources that are proposed to get exhausted in the near future. Gasoline used in the transport sector is the second largest contributor to GHG emissions after industries [1]. Bioethanol produced from Lignocellulosic biomass is not carbon neutral but reduces GHG emission significantly comparative to gasoline [2]. If we need to save our planet, the use of renewable sources of energy is inevitable. As public awareness of this problem is growing, there has been an increased investment into alternative sources of energy which will lower global dependence on fossil fuels as well as GHG emissions. The concept of biorefinery holds a broad range of technologies capable of separating biomass resources such as wood, grass, corn, etc., into their constituents that is, carbohydrates, proteins, Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00013-2 © 2020 Elsevier B.V. All rights reserved.

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triglycerides, etc., which can then be converted into important products, which are biofuels and chemicals [3]. Innovative technologies to access compounds from a single feedstock are among the new movement of research around the world. Biorefineries are eco-friendly as well as very similar to oil refineries [4]. The aim of this chapter is to establish the biorefinery context of best utilization of feedstocks by the synergistic action of several hydrolytic enzymes with their appropriate concentrations. Thus it is proposed that the surplus feedstock’s available worldwide which do not have any other alternative applications can be used for bioconversion via enzymes into useful commodities for mankind.

13.1.1 BIOMASS Lignocellulosic biomass could be utilized for the benefits of mankind for converting them into value-added products and is a valuable source to modern society. Recently, there has been a renaissance in the use of lignocellulosic biomass for two decades for biofuel, which was otherwise lost in oblivion. Cellulose is the most common organic polymer, representing about 1.5 3 125 tons of annual production of biomass via photosynthesis, particularly in the tropic regions. It is the only inexhaustible, renewable, and dominating agricultural waste material available in abundant to mankind. It has the great advantage that it is not utilized as food such as starchy biomass is used. Various types of biomass are abundantly available to be utilized for bioconversion, which are not utilized for any other purposes. Lignocellulosic biomass is mainly composed of three biopolymers: cellulose, hemicelluloses, and lignin, and the amount varies from 30%
50%, 20%
40%, and 20%
30%, respectively. All these components of biomass are an interlinked naturally unique arrangement to give the best rigidity to biomass and offer great resistance to get digested by enzymes. Here cellulose is the main structural component and also the most common organic biopolymer, which is a nearly interminable source of raw feedstock that can be used for raising different products. Cellulose shows crystalline nature, which is an unusual feature among biopolymers. Approximately 500
15,000 anhydrous glucose units are interlinked via β-1,4-glycosidic bonds and forming a linear homopolysaccharide. Intermolecular and intramolecular hydrogen bonding is carried out from β-1,4 orientation of the glucosidic bonds, resulting in natural cellulose into extremely crystalline, insoluble, and highly resistant to the enzyme attack. In each cellulose crystal, cellulose chains are usually hardened by H-bonds at inter and intrachain and neighboring polymeric layers which superimpose to one another are interlinked by weak Van der Waals forces. The extremely crystalline region of polymer is distinct from the other less amorphous region [5]. Cellulose is usually embedded with other structural biopolymers such as hemicelluloses and lignin to be protected naturally by any challenges; hence, it is rare that cellulose is present in nature without copolymers in the pure state. It possesses a unique characteristic of having permeability for larger molecules of enzymes as well as for smaller water molecules. Hemicellulose consists of a short, highly branched polymer of pentoses (e.g., D-xylose and Larabinose) and hexoses (e.g., D-mannose, D-galactose, and D-glucose) with 50
200 units. The acetate group of hemicelluloses is arbitrarily connected with ester linkages to the hydroxyl groups of the sugar rings. Hemicelluloses provide a connection between lignin and cellulose [6]. Lignin is a heterogeneous, amorphous, and cross-linked aromatic polymer, which is composed of trans-coniferyl, trans-sinapyl, and trans-p-coumaryl alcohols. Lignin encompasses the cellulose

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microfibril forming a complex matrix, which is covalently attached to side groups on different hemicelluloses. The percentage of lignin varies from 2%
40%. The carbon
carbon (C
C) and ether (C
O
C) bond in the lignin provides plant cell wall stability and protects it from microbial attack [7]. Enzymes-mediated bioconversion of lignocellulosic biomass includes three key steps: (1) pretreatment of biomass which reduces the rigidity of the biomass by loosening the fibers and, or by separating lignin from biomass; (2) enzymatic hydrolysis of pretreated biomass into sugars mainly glucose; and (3) conversion of glucose into value-added products. Each step has its own challenges. The ratio of components in biomass varies for different biomass and is the main reason that none of the set processes of pretreatment and enzymes could work universally.

13.1.2 PRETREATMENT The recalcitrance of lignocellulosic biomass is a major challenge for the enzyme’s action on the biomass. In pretreatment, the structure of cellulose and lignin attached by hemicellulose breaks down, resulting in decreased crystallinity of cellulose and an increased fraction of amorphous cellulose, the most acceptable matrix for enzymatic hydrolysis. Pretreatment of biomass enables cellulose to be accessible to the enzymes either by removing part of the lignin or hemicellulose. Lignin is the most recalcitrant component among different components of biomass to digestion and is responsible for the protection of biomass from microbial attacks. It restricts cellulase to access cellulose. Hence, it has been rightly considered as one of the biggest barriers to economic bioconversion of biomass [8]. It is assumed that after pretreatment, the hydrolysis rate increases up to 90% from 20% theoretically [6,9]. Various pretreatment methods have been proposed in the last few years. These pretreatment methods fall into different classes: physical pretreatment (e.g., milling, grinding, and irradiation), chemical pretreatment (e.g., alkali, dilute acid, oxidizing agents, and organic solvents), physicochemical pretreatment (e.g., steam pretreatment/autohydrolysis, hydrothermolysis, ammonia fiber explosion, and wet oxidation), and biological pretreatment or combination of these [10]. The abovementioned pretreatment methods can be selected based on the nature of biomass as well as its composition. Biorefining process of biomass usually starts with a pretreatment, followed by the conversion of cellulose to glucose, and finally downstream products [11,12]. However, the process is often hampered by the unsatisfying performance of biomass-processing enzymes, leading to the consumption of large amounts of enzymes and thus high production costs.

13.1.3 BIOMASS-PROCESSING ENZYMES Nature has given biomass rigidity due to its role in forming a structural base to plants and trees. It is this rigidity that enables them to stand straight and withstand microbial attacks. When it is a boon, it also throws challenge having enzymes to break it into simpler molecules. As biomass is not a single component rather a complex structure made up of cellulose, hemicelluloses, lignin pectin, etc., a single enzyme cannot work on it; rather, a cocktail of enzymes is required to effectively breakdown the complex biomass. Thus, a set of enzymes consisting of cellulase, xylanase, pectinase, auxiliary enzymes, etc. in different proportions are required for breaking up of complex biomass.

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The understanding of lignocellulose matrix structure and conversion mechanism of hydrolytic and auxiliary enzymes can provide significant insights on the performance of cellulase cocktails for enhancing saccharification efficiency [13]. Therefore, screening of novel microorganisms and bioprospecting in search of superior enzymes is an ambitious project that needs a targeted approach. Resulted superior enzymes may act synergistically with existing commercial cellulases for increasing its hydrolytic efficiency. There is a commercial need for a complex and coherent enzyme cocktail, which may act synergistically to unlock the complexity of lignocellulose polymer and convert it into fermentable sugars to a greater extent possible. These enzymes are only the major expensive inputs in the overall bioprocess and bringing down the budget of enzymatic saccharification is a current major challenge.

13.1.3.1 Cellulase Microbial degradation of the lignocellulosic biomass is attained by rigorous action of several enzymes among them the cellulase is the most prominent one. Though bacteria and fungi, both are capable of producing cellulases, fungi are considered as the better candidate for production of cellulase due to their capacity to secrete a great amount of cellulases extracellularly. Cellulase is a complex enzyme comprising of three major enzymatic components such as endoglucanase (EG, EC-3.2.1.4), exoglucanase (cellobiohydrolase, CBH, EC-3.2.1.91), and β-glucosidase (BGL, EC-3.2.1.21). Along with the above components, auxiliary enzymes such as lytic polysaccharide monooxygenases (LPMOs) also came into picture which perform a substantial role in the cellulose hydrolysis and are considered among cellulase components. EGs cleave cellulose polymer from in between of two units for uncovering both reducing as well as nonreducing ends, later CBH performs upon both of these ends to release cellobiose and cello-oligosaccharides, and finally BGL chops cellobiose to release glucose and thus resulting an appropriate hydrolysis. LPMOs are nonhydrolytic proteins, which help cellulase components to access cellulose fibers by producing nicks in between the chain and accelerate the action of EG. Thus, all of them act together in perfect synergism, as shown in Fig. 13.1. These enzymes act in perfect synergism and tight regulation under natural conditions and convert the cellulose polymer into its monomers [14]. Usually, CBH and EG are produced in sufficient amount, whereas BGL is produced meagerly by microorganisms. Commonly, cellulases exhibit a typical two-domain configuration [15,16], which contains a cellulose/carbohydrate-binding domain (CBD) or cellulose-binding module (CBM) and a catalytic domain (CD) linked via a linker peptide. Catalytic or core domain comprises a catalytic site while CBDs assist for the enzyme attachment to cellulose. Hydrolysis of natural cellulose needs various levels of collaboration among three key components of cellulases. A unique synergism occurs among exoglucanases and EGs (endo
exo synergism) and also between exoglucanases. BGL has been regarded as the rate-limiting enzyme as it completes the hydrolysis by hydrolyzing cellobiose, which otherwise if remains in the medium inhibit CBHs production [17]. Mostly, the common BGL undergoes feedback inhibition when the glucose gets accumulated in the system; thus the recent emphasis is on the necessity of having glucose-tolerant BGLs to drive the reaction forward and avoid feedback inhibition even in the presence of higher concentration of glucose [18]. Trichoderma reesei is a potential producer of cellulase among well producers, which has been extensively reviewed and exploited for commercial cellulase production. Cellulases obtained from T. reesei are basically inducible enzyme and its production is well regulated by activation [19] and repression mechanisms of genes are coordinately regulated [20]. T. reesei cellulase is rich in CBH

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FIGURE 13.1 Synergistic action of cellulase components to hydrolyze completely.

and has sufficient EG also but is deficient in BGL. Several studies have been on improving the BGL content of T. reesei to produce an excellent repertoire of cellulase for efficient hydrolysis of biomass. In several cases, heterologous BGL has been overexpressed in T. reesei and was successful too in improving cellulase content [21]. Fungal cellulases are thus the most promising candidates and are exploited for improved production [19].

13.1.4 CELLULASES FOR BIOCONVERSION Microbial cellulases find applications in a variety of industries where cellulases of varying degrees of purity are desired. Though cellulases were initially investigated several decades back for the bioconversion of biomass, this later became unattractive and the other industrial applications of the enzyme as in animal feed, food, textiles, and detergents and in the paper industry were predominantly pursued. However, with the shortage of fossil fuels and the arising need to find alternative sources for renewable energy and fuels, there is a renewal of interest in the bioconversion of lignocellulosic biomass using cellulases and other enzymes. The performance of cellulase mixtures in biomass conversion processes depends on several of its properties including stability, product inhibition, synergism among the different enzymes, productive binding to the cellulose, physical state as well as the composition of cellulosic biomass. Cellulases are available in the market under different names or trademark for different applications, which could also be tried for biomass hydrolysis. Nieves et al. [22] and Kabel et al. [23] analyzed and evaluated the potential of several commercial cellulases for biomass conversion. They performed the standard assays for different enzymes such as filter paper activity (FPU), CMCase, BGL, and xylanase. It would not be feasible to predict the efficiency of cellulases for bioconversion based on standard assays, as there are no clear relationships between cellulase activities on soluble substrates and those on insoluble

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substrates. So the soluble substrates should not be used to predict the efficiency of cellulases for processing relevant solid substrates, such as plant cell walls. The choice of the enzyme preparation for particular biomass would be more dependent on biomass characteristics rather than on standard enzyme activities measured. Preparations having higher FPUs are desirable for bioconversion since filter paper is a highly crystalline cellulose, the degradation of which depends on the combination of activities of EG and CBH, where the EG creates new chain ends for the CBH to split off cellobiose which further gets attacked by BGL to give glucose. Preparations of cellulase from a single organism may not be very efficient for hydrolysis of a particular feedstock. Though the filamentous fungi are the major source of cellulases and hemicellulases and the mutant strains of Trichoderma including T. reesei, Trichoderma viride, and Trichoderma longibrachium are the best-known producers of the enzyme, it is also well known that these species of Trichoderma have a low level of BGL activity. Cellulases for biomass conversion could be a blend or enzyme cocktail containing endo- and exo-cellulase, xylanase, BGL, pectinase, etc., which could vary for different biomass based on their composition. The hydrolytic efficiency of a multienzyme complex for lignocellulose saccharification depends both on properties of individual enzymes and their ratio in the multienzyme cocktail. The ideal cellulase complex must be highly active on the intended biomass feedstock, able to completely hydrolyze the biomass, operate well at mildly acidic pH, withstand process stress, and be cost-effective. The success of any lignocellulosic ethanol project will depend on the ability to develop such cellulase systems. The key to developing cellulases that are effective toward a particular biomass feedstock is to artificially construct them either by enzyme assembly to form cocktails or to engineer the cellulase producers to express the desired combination of cellulase enzymes. Both these approaches have been tried with success. Enzyme cocktails have been developed by mixing T. reesei cellulase with other enzymes including xylanases, pectinases, and BGLs, and these cocktails were tried for hydrolysis of various feedstock. One of the recent examples of cocktails developed include the multienzyme complex developed based on highly active Chrysosporium lucknowense cellulases. Characterization of cellulases has been achieved to a considerable extent by site-directed mutagenesis. These studies as well as X-ray crystallography of the enzymes have led to the identification and characterization of putative catalytic and binding residues and the trapping of enzyme
substrate complexes. Nevertheless, understanding the mechanism of degradation of the natural substrate
crystalline cellulose remains a great challenge. Physical and nutritional parameters optimization have led to enhanced cellulase production, however, to have further enhancement, strain improvement is inevitable. Strain can be improved by genetic modification to increase the cellulase production as well as its efficiency.

13.2 STRAIN IMPROVEMENT VIA MUTATION Strain improvement is inevitable if cellulase production has to be reached to an industrially feasible level. To increase the properties in order to meet the robust commercial applications, engineering of the cellulases was adopted as common practice. The majority of filamentous fungi naturally produce numerous cellulases during growth up on lignocelluloses biomass. Random mutagenesis and site-specific mutagenesis found to be a standard method for improving the secretion of cellulase in

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fungi. Moreover, both mutagenesis in their combinations were explored to develop tailor-made enzymes for commercial applications [24]. For the last 20 years, Penicillium oxalicum mutant (JUA10-T) produces cellulase enzyme preparations at industrial scale in China [25].

13.2.1 RANDOM MUTAGENESIS T. reesei RUT-C30 is the result of 30 cycles of random mutation [26]. This is of the most exploited fungal strain for commercial cellulase production. The reason for being most exploited is that it has been researched exclusively and other cellulase sources were neglected during earlier days. At present, Penicillium sp., Aspergillus sp., Myeciliophthora, and Humicola sp. are giving tough competition to T. reesei [27]. Among other methods of strain improvement, classical mutation has been the most accepted to date. Several articles have been published on fungal mutations and thereby increased cellulase production. Most of the fungal strains employed for commercial cellulase production are the mutant strains though few are genetically modified too. Table 13.1 summarizes

Table 13.1 Cellulase production improvement via mutagenesis among microorganisms. Microorganism

Type of mutation

Recombinant properties

References

Penicillium oxalicum

Functional mutation

[28]

Aspergillus species

Sequential random mutation Random mutation

Enhanced cellulase production (160 FPU/ L/h) Enhanced cellulase production

[30]

[37] [38] [39]

Cellulolytic fungi Penicillium janthinellum NCIM1171 Trichoderma atroviride Humicola insolens Aspergillus sp. Su14 Penicillium decumbens 114

Random mutation Random mutation

Trichoderma viride Aspergillus terreus AUMC 10138 Fusarium oxisporum Cellulomonas Penicillium Funiculosum

Random mutation Random mutation

Enhanced β-glucosidase activity and FPase activity Stable and enhanced cellulase production Enhanced cellulase production and hydrolysis Enhanced cellulase production Enhanced cellulase production Enhanced cellulase production Enhanced cellulase and hemicellulase production Enhanced cellulase production Enhanced alkaline cellulase production

Random mutation Random mutation Site-directed mutagenesis Site-directed mutagenesis Site-directed mutagenesis

Enhanced cellulase production Enhanced xylanolytic activity Increased enzyme activity against crystalline cellulose Increased thermostability and activity toward Avicel Modified endoglucanase for hydrolysis of β-glucans

Acremonium cellulolyticus

Thermotoga maritima Macrophomina phaseolina

Random Random Random Random

mutation mutation mutation mutation

[29]

[31] [32] [33] [27] [34] [35] [33] [36]

[40] [41]

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several strains that have been improved for cellulase production by mutation. Most of the industrially exploited strains for cellulase production are robust mutants.

13.2.2 SITE-DIRECTED MUTAGENESIS When mutagenesis is done to bring the desired changes in the DNA sequences of a gene, it is called site-directed mutagenesis. It is time-consuming as one should know the sequence of the genome and the region to target. Then with the help of specific primers, amino acid sequence can be changed and then by homologous recombination, the changes get incorporated into the genome to bring the desired changes in the property of the protein. It is usually done to improve the property of a protein by changing the amino acid sequence. For improvement of the thermostable property of xylanase obtained from Aspergillus niger BCC14405, Thr and Ser residues on the xylanase surface were substituted by four-five Arg residues. The modified enzyme showed maximal activity as compared to the wild-type strain which showed only a 15% increase in activity, half-life of the mutant enzyme was also increased respectively, that is, 257 6 16 and 285 6 10 min for both four- and five-Arg mutants, whereas wild enzyme only showed 14 6 1 min increase, therefore from above it was observed that arginine substitution increased the consistency through 18
20 fold. A kinetic study was carried out, and fiveArg substitution enzyme showed lower substrate affinity and catalytic rate [42]. An EG from Thermotoga maritima cel5A underwent engineering via site-directed mutagenesis and CBM to obtain a hyper-thermostability [40]. Commercial cellulase formulations obtained by fungal mutants find great application in modern biotechnology for production of 2G biofuels from lignocellulosic biomass [43]. Tremendous attention has been given for obtaining cellulases with better stability, greater specific activity, reduced inhibition susceptibility and increasing characteristics of explored enzymes through advanced techniques of genetic engineering [44
48]. With the advancement of heterologous expression, the recombinant enzyme production systems have been proved as the best platform for effective production of commercial cellulase [49].

13.2.3 GENETIC ENGINEERING Genetic engineering is one of the most popular tools, which enables to combine multiple desirable traits into a single organism. It can be used to engineer microbes for high metabolite production which could vary from simple proteins to highly specific therapeutic proteins. However, all the cases may not be as simple as it sounds because of the inherent complexity of the organisms itself or the metabolites needed to be expressed. The most important information which is required is the genetic makeup of the organisms. Knowledge of whole genome sequence and function makes it easier to decide the target sites for genetic alterations. For improved production of cellulases in fungi, metabolic engineering as well as targeted strain engineering demands an effective insertion method for directed genetic alterations. Singh et al. [50] gave an account of improvement in cellulase production among microorganisms by employing genetic engineering as a tool. For a long time, the low-efficiency of directed gene insertion has been a key challenge to get sufficient transformants through the homologous insertion/deletion of expression cassette. This is mostly due to the complexities associated with the presence of introns

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in their genes and due to complexities associated with glycosylation. Filamentous fungi are known to produce a broad range of metabolites in significant quantities. Their ability to grow on the cheaper substrate has made them a potential source of metabolites for industrial applications. Knowledge of fungal genetics has accumulated since years of research and industrial applications. Along with the ability to secrete a protein, they can perform posttranslational modification such as glycosylation and disulfidation [51]. Most of the filamentous fungi are transformed with a plasmid, which integrates into their genome and thereby provides stability to the fungal transformants. Thus filamentous fungi have tremendous potential to be employed as hosts for recombinant DNA. But there exist challenges on the nonhomologous recombination in fungi. This difficulty was resolved by elements disabling of DNA repair in the nonhomologous end-joining pathway [52,53]. Both Aspergillus and Trichoderma were used as expression hosts for numerous fungal and nonfungal genes. Table 13.2 gives an account of the heterologous gene expression for enhancing the cellulase synthesis as well as the ratio of different components to increase its bioconversion efficiency. T. reesei genome [64] revealed that despite being the best-known producer of cellulases, the genome of the fungus contains fewer cellulases and hemicellulases than any other sequenced fungi. Table 13.3 shows few cellulase genes available in the host. Thus, the mechanism and reason for high secretion level are still to be revealed. It has been observed that cellulase and hemicellulase gene of T. reesei is regulated under cre1 regulatory gene through catabolic repression. The gene cre1 was fully replaced/removed through a truncated cre1-1 mutant variant, which was formerly found in mutant strain RUT-C30 with improved enzyme synthesis. This resulted in a morphological change in the colony of mutants as compared to wild strain, colonies of mutant become smaller in size with few aerial hyphae and spores. When cultured in glucose medium, these transformants showed a decrease in the production of cellulase and hemicellulase. However, they showed increased hydrolytic enzyme quantities when cultured in fermentation media through the induction of hydrolase gene. This study suggested that cre1 performs as an expressing modulator for the cellulase and hemicellulase genes under inducing and noninducing environments. When different experiments on the mutant strain with cre1 and cre1-1 were performed, no phenotypic difference was reported, which showed that gene cre1-1 was basically a null allele. From the above, it was determined that cre1 is a promising gene for the strain engineering to increase the enzyme yield [62]. In cellulolytic filamentous fungi, synchronous regulation of both induction, and catabolic repression is important for cellulolytic enzyme expression and its precise reactions with carbon sources. Several proteins, especially regulatory transcription factors which are responsible for above processes, have been recognized and engineered in several cellulolytic fungi including P. oxalicum, but after these excessive struggles such as transforming a solitary target for the cellulase production is scarce. For improving lignocellulolytic enzyme production, a strategic methodology was set to genetically transform P. oxalicum. Moreover, Clr-2 is a fungal specific transcription factor domaincontaining protein, which is also known as cellulose degradation regulator. In Neurospora crassa, out of 212 genes responding to Avicel, 135 genes were regulated through Clr-1/Clr-2 [72]. The clrB is an ortholog in P. oxalicum and well conserved in all ascomycete fungi, and it is also vital for induction of cellulase expression. Therefore first target level of clrB was increased constitutively by overexpressing the gene with the promoter gpdA from Aspergillus nidulans. Major intracellular BGL plays a negative role in the induction of cellulases and xylanases. In addition to this,

Table 13.2 Heterologous expression for enhanced cellulase production. Recombinant properties

Source

Host

Strategy used

Promoter gpdA from Aspergillus nidulans

Penicillium oxalicum

Enhancement of cellulase production

[17]

Penicillium decumbens

Trichoderma reesei (RUTC30) T. reesei

Amplifying induction along with depression (knockout strategy) Agrobacterium-mediated transformation

Enhanced BGL activity

[54]

Transformation via plasmidcarrying selectable marker

Enhanced cellulase production with higher BGL content

[55]

Transformation via plasmid constructed by prochymosin cDNA fused with promoter, signal sequence, and terminator of the cbhl gene CBH gene inactivation

[56]

Cloned and expressed eg7

Production of eukaryotic heterologous protein for enhanced cellulase production Enhanced cellulase production Thermostable cellulase

Cloned and expressed bgl

glucose-tolerant BGL

[59]

Overexpressed BGL under constitutive and inducible promoter Heterologous expression of bgl and LPMO under the control of gla1 promoter Gene cre1 knocked out and replaced by a cre1-1 Cloned and expressed in protein-secreting vector pEXPYR using ligation-free cloning

65-fold higher yield comparative to wild type Production of heterologous enzyme simultaneously Enhanced cellulase production Enhanced LPMO level in cellulase produced

[60]

Escherichia coli BGL and acetamidase gene (amdS) or the argB gene of A. nidulans Chymosin (rennin) isolated from stomach of young calves

Expression vector PAMH10 Myceliophthora thermophila Penicillium funiculosum NCL1 P. oxalicum

bgl from Aspergillus niger/TrLPMO from T. Reesei RUT-C30 AN3046LPMO from A. nidulans

T. reesei

T. reesei Pichia pastoris P. pastoris P. oxalicum

Penicillium verruculosum T. reesei A. nidulans

References

[57] [58]

[61]

[62] [63]

BGL, β-Glucosidase; CBH, cellobiohydrolase; LPMO, lytic polysaccharide monooxygenase.

Table 13.3 Cellulase or cellulase encoding genes present in microorganisms. Microorganisms

Cellulase/genes

References

Trichoderma reesei

Two CBHs, eight EGs, seven BGLs, 16 hemicellulases, and nine LPMO genes Two CBHs, five EGs, lacking CBM Three CBHs, four EGs, and one BGLs Three CBHs, 11 EGs, and 11 BGLs Eight EGs, seven CBHs, nine BGLs, and 25 LPMO genes

[65,66]

Two CBHs, two EGs, five BGLs, and seven LPMO genes 12 endo- and exo-cellulases

[69,70] [71]

Humicola insolens Neurospora crassa Penicillium decumbens Myceliophthora thermophila Aspergillus niger Clostridium thermocellum

[67] [68] [35] [58]

BGL, β-Glucosidase; CBH, cellobiohydrolase; CBM, cellulose-binding module; EG, endoglucanase; LPMO, lytic polysaccharide monooxygenase.

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the expression of cellulases and hemicellulases was repressed when the preferred carbon sources were available, which is mediated by transcription factor CreA. To overcome repression, gene bgl2 was replaced by marker gene hph along with clrB overexpression and gene CreA was deleted by using gene bar as the marker, obtaining a trigenic recombinant strain “RE-10.” Several screenings were carried out for comparative analysis of “RE-10,” wild type, and JU-A10-T [17]. Creating fungal strain overexpressing positive regulatory genes and knocking out negative regulatory genes would improve cellulase expression. For the last 20 years, P. oxalicum mutant (JUA10-T) produces cellulase enzyme preparations at industrial scale in China [25]. Recently, its genome has been sequenced, providing valuable information to mine novel components that play key roles in biomass saccharification [35]. Moreover, the mechanism of the synthesis of P. oxalicum cellulase gene via describing the inducer transportation, signal transduction, and transcription regulation has been partly revealed. This advancement has been led by numerous methods such as gene knockout techniques, genome sequencing, whole transcriptome shotgun sequencing, and proteomics. Recognition of the three key transporters of cellodextrin, for example, CdtC, CdtD, and CdtG, which are known to play the vital roles in induction of cellulase production, has been enabled through gene disruption analysis [28]. The downregulation of the cellulolytic gene’s expression has been documented to be occurred by signaling G protein-cAMP pathway [73]. Moreover, library of single-gene deletion was established for approximately 470 transcription factors, four key, and 15 novel, transcriptional regulators. The precise roles of these factors in the regulatory network of cellulase expression were also recognized and described [74]. The Reconstruction of Expression Regulatory Network (REXRN) technology has been developed as a new strategy to engineer fungi that enhance cellulase and protein production. One mutant in particular (RE-10 from REXRN) displayed a drastic increase in cellulase and hemicellulase production and produced even a higher value compared to the industrial strain JU-A10-T [17]. However, the BGL of RE-10 has not yet been improved to the same level as other inducible cellulases. This makes further improvement of its BGL activity necessary. Yao et al. [60] have presented a systematic overexpression analysis of nine BGL encoding genes in the wild-type strain 114-2 of P. oxalicum. They overexpressed BGL1, BGL4, or BGL5 using both constitutive as well as inducible promoters. High-yielding BGL producers were obtained and the yields were elevated from twofold to 65-fold. With the help of genetic engineering, cellulolytic enzyme production could be improved significantly. These improvements could also be achieved by gene inactivation and changing the promoter. The promoter of the cellulase encoding gene, which is reported to be strong, and highly inducible promoter reported was cellobiohydrolase I (CBH I). CBH I was also being used in the eukaryotic heterologous production of protein in T. reesei [56]. CBHs are usually glycosylated such as other fungal enzymes. As a rule, the N-glycosylation is a characteristic of CDs, while peptide linkers, rich in Ser and Thr residues, are heavily decorated with O-linked glycans [75
77]. N-linked glycosylation, is the attachment of an oligosaccharide molecule, known as glycan to a nitrogen atom (amide nitrogen of Asn residue of a protein), in a process called N-glycosylation. This type of linkage is important for structure and function of some eukaryotic proteins. The roles of N- and O-glycosylation in the structure and function of cellulases were not well understood until recent years. It has been thought that both types of glycans may participate in the correct folding of a protein and maintaining its stability, while the O-linked glycans in a linker may protect it against protease action [76,77]. One study has reported that an over-N-

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glycosylated recombinant CBH I of T. reesei (rTrCel7A) expressed in A. niger displays a reduced activity on cellulose relative to the native enzyme [78]. Then the same team of researchers showed that eliminating an existing N-glycosylation site in rTrCel7A or introducing a new N-glycosylation site in recombinant Cel7A of Penicillium funiculosum (rPfCel7A) by site-directed mutagenesis leads to an increased enzyme activity against crystalline cellulose [39]. Gusakov et al. [79] applied this approach for boosting the specific activity of recombinant CBH I from Penicillium verruculosum (rPvCel7A expressed in Penicillium canescens) and showed that the N-linked glycans are involved in complex interactions with a polymeric substrate in catalysis [80]. Gusakov et al. [79] got a deep insight through site-directed mutagenesis to discover in recombinant P. verruculosum about the N-linked glycans in the CD “CBH II (rPvCel6A)” denote a vital segment of the enzyme-mediated catalytic machinery. From complete data analysis, it was discovered about extremely active Myceliophthora thermophila CBH IIb (MtCel6B), the N-glycan mechanism affects both GH7 and GH6 processive CBHs [81].

13.3 CELLULASE ACCESSORY AND AUXILIARY ENZYMES 13.3.1 β-GLUCOSIDASE BGL is the rate-limiting component because it is easily inhibited by its own product glucose. The inhibitory effect leads to the accumulation of its substrate cellobiose, a strong inhibitor to EGs and CBHs, and thus slows down the whole cellulose degradation process through cascade feedback suppression [18,21]. Most of the potent cellulase-producing fungi lack BGL required for efficient breakdown of cellulose fibers. It is required that the deficient enzyme must be supplemented to the base enzyme such that all the components should be in optimum concentration. Supplementation of BGL blended cellulase is an effective strategy for maximizing saccharification efficiency. Over the past decade, several efforts were made for obtaining the BGLs with excellent glucose tolerance by protein engineering as well as by isolating it from diverse environmental resources [82]. There is a possibility that the deficient component could be overexpressed in base enzymepreparing organism to get the desired cocktail with all the components of cellulases. The selection of enzyme component to be expressed/overexpressed is usually based on the absence of that enzyme component in base preparation. Most of the commercial cellulase production is being done by recombinant fungal strains. Novozymes Inc. used T. reesei as a parent or base strain and isolated effective cellulolytic as well as noncellulolytic genes from other organisms and transformed them into this strain for effective preparation of enzyme mixture in a single organism [83]. Similarly, Dyadic’s recombinant strain, which also produces few accessory enzymes. Cofermentation of P. verruculosum (basic strain) with another P. verruculosum with the heterologously expressed Aspergillus sp. β-glucosidase produced up to 19% of β-glucosidase as compared to 4% in basic strain alone. This cofermented enzyme preparation gave 113% increased hydrolysis of Avicel than basic strain enzyme preparation [84]. In another report, the formulation of enzyme obtained from recombinant P. verruculosum, secreting the heterologous AnBGL (BGL from A. niger) or TrLPMO (LPMO from T. reesei) under gene promoter gla1 to demonstrate effective lignocellulosic biomass hydrolysis than that of control formulations having no heterologous enzymes. The enzyme preparation comprising TrLPMO and AnBGL equally exhibited the best saccharification of lignocellulosic

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biomass and claiming to develop a potential fungal strain accomplished to provide both the heterologous enzymes concurrently [61]. The bgl genes from a large number of bacterial, mold, yeast, plant, and animal systems have been cloned and expressed in both Escherichia coli and eukaryotic hosts such as Saccharomyces cerevisiae, Pichia stipitis, and filamentous fungi. Filamentous fungi are known to be a good BGL producer and several BGLs from glycosyl hydrolase family 3 and 1 have been purified and characterized from various fungi. In spite of being good producers of the enzyme, reports on cloning of BGL from fungi are relatively low due to complexities. Even Pichia and Saccharomyces have also been employed as host of expressing eukaryotic bgl genes. But several thermotolerant BGLs from bacteria have been cloned and expressed in E. coli. Table 13.2 provides an account of the heterologous enzyme expressed in various hosts with improved cellulase production and efficiency as well. A coding sequence “BGL I” from Penicillium decumbens was inserted with CBH I promoter of T. reesei which is widely used in enzyme industry to enhance BGL activity in T. reesei to degrade cellulose efficiently. The ligated sequences of BGL I coding sequence and CBH I promoter sequence were added in the T. reesei RUT-C30 genome through Agrobacterium-based transformation. Two transformed strain were selected based on their filter paper and BGL activity, which were 6-8 and 30-folds, respectively, higher compared to wild strain. Furthermore, pBGL1 (heterologously synthesized) was purified and supplemented into the T. reesei RUT-C30 secreted enzyme complex and was observed that during hydrolysis of cornstalk, the yield of glucose increased up to 80%. This result showed that the heterologous secretion of a BGL in T. reesei could yield a balanced enzyme blend [54]. There has always been a debate on using cocktail supplemented with deficient component of cellulase into the base enzyme or expressing the deficient component in a suitable host having the other entire component expressed in optimum amount. For the effective or complete degradation of cellulose, biopolymer requires to work all three key cellulases in the synergistic manner. Moreover, the usage of a complete culture to deliberates numerous advantages, for example, reduction in the budget of manufacturing enzymes, probability for synthesizing whole range of enzymes and comparatively easy process scale for production of bulk and fine chemicals [85]. Thus, whole-cell engineering of fungal strain is the best choice for improved cellulase expression.

13.3.2 XYLANASE There are some other components also produced, which could increase cellulase efficiency significantly such as xylanase. About 20%
40% of biomass is hemicellulose, which acts as a physical barrier and limits the accessibility of the cellulases to cellulose [86]. Endo-1,4-β-xylanase, β-xylosidase, α-glucuronidase, α-L-arabinofuranosidase, and acetylxylan esterase for xylan degradation reacts on different heteropolymers whereas for digestion of glucomannan, β-mannose, and β-mannosidase breaks down the polymer backbone [6,87]. Blending of hemicellulases especially xylanases in the enzymatic cocktails shows synergistic effect in cellulase activities and increases overall saccharification efficiency [13].

13.3.2.1 Hemicellulose-degrading enzymes Xylan is the second most abundant polysaccharide present in plants after cellulose. This polymer comprised of 1,4-linked several units of β-D-xylopyranose attached with several substituents such

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as O-acetyl, α-L-arabinofuranosyl, 4-O-methylglucuronic, or α-1,2-linked glucuronic acids attached as side groups [88]. The presence of acidic substituents is not always ubiquitous in the xylan structure and the unsubstituted linear xylan has also been isolated from several other sources [89]. The degree of polymerization in case of hardwood xylan is 2X or more as compared to softwood xylan, with the former existing as O-acetyl-4-O-methylglucuronoxylans, and arabino-4-O-methylglucuronoxylans. The interaction of xylan with cellulose and lignin occurs with the help of various covalent and noncovalent linkages involving these side groups, which also help in determining the functional properties of xylan, including solubility. The different types of xylan such as linear homoxylan, arabinoxylan, glucuronoxylan, and glucuronoarabinoxylan have been categorized based on the common side groups attached to its backbone. The differences between the different types of xylan can be further attributed to its branching properties. The different O-acetyl groups attached to xylosyl residues help in the protection of acetylxylan from degradation by xylanases, possibly due to steric hindrance. The complete hydrolysis of acetylxylan thus depends on the synergistic action by the xylanases and acetylxylan esterases. In a nutshell, the side chains play a vital role in solubility, physical conformation, and degradability of the xylan molecule [90,91]. Multiple enzymes can synergistically act together to completely degrade xylo-oligomers into xylose monomer residues. Two main xylanases enzymes are β-1,4-endoxylanase and β-1,4 xylosidase. Fungal β-1,4-endoxylanases are the enzymes that degrade linear chain of 1,4-linked D-xylose residues present in xylan which belong to GH10 and GH11 family. GH10 endoxylanase has broader substrate specificity than members of GH11 family. GH10 family enzymes are capable of degrading xylan backbones with higher degree of substitution and smaller xylo-oligosaccharides; hence, these enzymes are capable of degrading substituted xylan completely. The released xylooligosaccharides are subsequently degraded by β-xylosidase, and they belong to GH3 family. The xylan substituent groups are degraded by accessory xylanases which include, but are not limited to, L-arabinofuranosidases, p-coumaric esterases, acetylxylan esterases, ferulic acid esterases, etc. Xylanases from bacterial, archaeal, and fungal origin have been explored widely; however, the filamentous fungi hold a special place as excellent enzyme producer. Many filamentous fungi are known for their xylanase production such as Malbranchea cinnamomea, A. niger, Humicola insolens, and P. oxalicum which have been used for laboratory and industrial research. Owing to their potential industrial applications, xylanases have also been widely characterized for novel applications and produced many industrial patents as well (Table 13.4).

13.3.2.2 Genetic engineering of xylanases To increase their industrial suitability, xylanases have been improved by genetic engineering approaches. Genetic modulation of xylanases has been done to improve their industrial suitability such as an increase in activity, increase in substrate specificity, and stability increase in extreme industrial conditions. The different genes encoding xylanases were cloned in both the homologous and heterologous hosts for overproducing the enzymes and their utilization in commercial applications. Several fungal xylanase genes were cloned as well as expressed in the number of hosts, for example, bacteria, yeast, and fungus. However, for the expression of fungal xylanases, yeast, and fungus expression, hosts provide advantages of a proper protein folding and protein glycosylation of expressed

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Table 13.4 Expression hosts and summarized information of patented industrial xylanases. Source of gene

Gene/enzyme

Expression host

Reference

Humicola insolens Aspergillus niger Trichoderma reesei Neocallimastix patriciarum Penicillium griseofulvum A160 Aspergillus tubingensis DS16813 Orpinomyce ssp. strain 2 T. reesei

Xyn11B XAn11 Xyn2 Endoxylanase Endoxylanase Endoxylanase B

Pichia pastoris P. pastoris Escherichia coli Bacillus napus Aspergillus oryzae A. niger N593

[92] [93] [94] [95] [96] [97]

Endoxylanase Endoxylanase

[98] [99]

T. reesei T. reesei A. niger A. niger

Endoxylanase Endoxylanase β-Xylosidase β-Xylosidase

E. coli Trichoderma spp., Humicola spp., Neurospora spp., Aspergillus spp., and Fusarium spp. H. insolens Hansenula polymorpha A. niger H. polymorpha

[100] [101] [102] [103]

recombinant proteins. Use of native signal sequence for expression of recombinant enzymes is a common approach to obtain high-enzyme yielding recombinant clones. Some of the representative examples of industrially applicable recombinant fungal xylanase are compiled in Table 13.4. Homologous expression of T. reesei M2C38 endoxylanases was improved by creating mutant containing a new N-glycosylation sites for the expressed enzymes. The addition was done at the conserved amino acid site of GH11 xylanases by replacing Asn at 131 position along with Ser/Thr at 133 position. The new variant displayed an enhanced expression by 40% as compared with wild type [104]. Another example of T. reesei endoxylanase II modification was observed in an earlier report of Sung and Tolan [105]. The genetic modification of this enzyme was done to improve its activity at the higher temperature, extreme alkaline conditions and thermostability to make it suitable for paper and pulp industries applications. Thermophilicity of this enzyme was improved by replacing its native amino acids at positions 10, 27, and 89, respectively, with His, Met, and Leu. The aminoterminal sequence of this enzyme was also replaced by N-terminal sequence of Thermomonospora fusca. Another modification for the addition of ten additional amino acids from N-terminus of Clostridium acetobutylicum xyn B which was added to N-terminus of T. reesei endoxylanase. All these modifications improved the activity of mutant xylanases from 55 C to 75 C and active pH range of the enzyme was increased from 7.5 to 9.0. The same endoxylanase underwent another series of modification for the improvement of this enzyme at the higher temperature of 62.5 C and pH of 5.5. Replacement of Val108, Ser110, Asp154, and Ala158 was done with Cys, and Glu162 was replaced with His. The mutant produced after this modification enhanced activity and the thermostability stability of this enzyme was improved from 55 C to 62.5 C [105].

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13.3.3 LACCASE Lignin denotes 30%
35% of lignocellulosic biomass and the second most abundant carbon source on earth, after cellulose. This is one of the most recalcitrant parts of lignocellulosic biomass which acts a “glue” to bind cellulosic and hemicellulosic portions of biomass together [106
108]. In order to obtain the fermentable sugars from lignocellulosic biomass, delignification of lignocelluloses is advantageous for increasing cellulose accessibility. Naturally, lignin degradation is carried out by lignin-degrading enzymes, which include laccases, peroxidases, and manganese peroxidases [109]. As laccases do not require any additional enzymes/mediators to oxidize lignin
phenolic substrates, these enzymes are self-sufficient for lignin degradation. Laccases can be produced by fungi, bacteria, and plants as well as insects [110]. As compared to the laccase of plant or bacterial origin, fungal laccases are holding the highest reduction potential for oxidizing polyphenols, which makes them important industrial biocatalysts. Laccases are blue multicopper oxidases that have the ability to oxidize polyphenolic substrates that are present in lignin [111]. The active site of these enzymes is characterized by a catalytic center of four copper ions, which are involved in oxidative reactions [112]. The reduction of oxygen to water is coupled with oxidation of polyphenols to carry out depolymerization of lignin. Oxidized phenolic substrates are very unstable reaction intermediates, which subsequently induce next oxidation reactions or other nonenzymatic reactions to cleave bonds present in lignin, such as Cα
oxidation, Cα
Cβ, and aryl
alkyl cleavage. Nonphenolic lignin component such as α-hydroxyl group present in β-O-4-ether bonds represents a major part of lignin polymer. However, due to lower reduction potential of laccases, nonphenolic components cannot be oxidized by these enzymes. However, small molecules known as mediators, which usually have higher reduction potential, are able to oxidize and degrade β-O-4ether bonds in lignin [113]. These mediators can be natural fungal metabolites such as 3-hydroxyanthranilic acid, or they can be derived from lignin such as acetosyringone and syringaldehyde, or they can also be the synthetic molecules such as 2,20 -azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) [114]. After laccase-mediated oxidation of these small mediators, these oxidized radicals can diffuse in lignin polymer and effectively degrade the polymeric form of lignin. This system is known as laccase-mediated system for lignin degradation and has been employed in vitro for lignin pretreatment in biorefineries. This mechanism has shown to be quite effective in lignin degradation and has also supported enhanced biomass saccharification [115,116]. Fungal laccases are produced by a variety of fungi; however, white-rot fungi are known for efficient lignin degradation. White-rot fungi selectively depolymerize lignin, leaving cellulosic portions out of the biomass, which gives a white coloration to the wood; hence these fungi are termed as white-rot fungi. Some of the popularly known white-rot fungi are Phanerochaete chrysosporium, Trametes versicolor, Pleurotus ostreatus, etc., which contain high lignocellulolytic activities and can be exploited as biological pretreatment agents in biorefineries. However, longer growth periods of white-rot fungi and lower enzyme production yield limit their large-scale application in biorefineries. These problems might be improved by selecting hypersecreting laccase strains or by molecular intervention to modify organisms that could either produce laccases in higher titer or could reduce the longer incubation days of fungi, hence can improve the economic viability of laccase applications.

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13.3.4 AUXILIARY ENZYMES Auxiliary enzymes play an important role in cellulose degradation and are regarded as a part of cellulase system.

13.3.4.1 Lytic polysaccharide monooxygenase In recent years, LPMOs’ discovery has been regarded as the biggest revolution in cellulose degradation. In recent years, slight investigations were devoted toward finding out the role of LPMOs, which are auxiliary nonhydrolytic enzymes that help in hydrolysis of cellulose. Apart from endo
exo synergism, these auxiliary enzymes are also supportive for the dissolution of crystalline property of cellulose via oxidative breakdown of glycosidic linkages. LPMOs improve the saccharification by offering fresh attachment sites for the action of BGL, and CBHs. These LPMOs facilitate a novel oxidative process for effective degradation of the polysaccharide. Moreover, it was presumed that these auxiliary enzymes act on the fibrillary surface of crystalline cellulose and rendering its surface to become more prone to cellulases action [117
119]. Interestingly, LPMOs can extract the desirable and required electrons from lignin through far-off electron transfer [120]. In the carbohydrate-active enzyme (CAZy) database, all the LPMOs are classified as the members of GH61, and CBM33 families. But now again these are classified as “auxiliary activities” (AA) contrary to former classification as GH61, and now they are placed in 9
11, and 13 AA families [121]. LPMOs are copper-dependent metalloenzymes that initiate the biomass deconstruction process and subsequently synergize with cellulases, hemicellulases, and other accessory enzymes to enhance deconstruction of lignocellulosic biomass [122]. Their discovery has changed our view toward cellulose degradation mechanism. Initially, it was thought that cellulase degradation machinery consists of three fundamental enzymes, namely EGs/endo-β-(1,4)-glucanases, exoglucanases/exo-β-(1,4)-D-glucanases, and BGLs. However, the discovery of LPMOs has established their active role in cellulose degradation and these enzymes have become an integral part of canonical cellulase machinery. LPMOs are one of the powerful candidates of fungal enzymatic machinery that are mainly responsible for increasing cellulose accessibility to hydrolases. Initially, these enzymes were classified as GH61 enzyme due to their observed weak EG activity [123,124]. However, the first indication for these enzymes as nonhydrolases came through the first crystal structure of GH61 from Hypocrea jecorina, TrCel61B [125]. From the characterized structure, it was revealed that these enzymes contain a unique conserved metal-binding flat active site unlike the cleft or tunnel such as an active site of cellulases. This enzyme was also found to lack the conserved carboxylate residue in its active site, which is a hallmark of cellulases catalytic site. Currently, LPMO spans in five different classes, AA9 LPMOs consist mostly of cellulosedegrading LPMOs that are derived from fungi, AA10 LPMOs are chitin, and cellulose-acting enzymes and are derived from bacteria. The oxidative LPMO from fungus mainly acts on cellulose as the main substrate, AA11 LPMOs contain mainly fungal LPMOs that act on chitin, AA13 LPMO enzymes oxidize to starch, and lastly AA14 class of LPMOs which target on xylan degradation. These ubiquitous enzymes are present in nearly all cellulolytic fungi spanning Basidiomycota as well as Ascomycota group ascomycetes and some of the genes have been characterized from brown-rot fungi as well. Table 13.5 summarizes some of the known fungal strains with AA9

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CHAPTER 13 PROMISING ENZYMES FOR BIOMASS PROCESSING

Table 13.5 Some representative examples of AA9 LPMO proteins acting as cellulase-boosting enzymes. Source of LPMOs

Substrate

Expression host

Increase (%) or (folds increase)

References

Thermoascus aurantiacus

Pretreated corn stover Pretreated corn stover Pretreated corn stover Avicel

Trichoderma sp.

18

[117]

Trichoderma sp.

13

[117]

Trichoderma sp.

15

[126]

Pichia pastoris

25

[127]

Oak residue Filter paper Sorghum stover Wheat straw

P. pastoris P. pastoris A. nidulans P. pastoris KM71H P. pastoris GS115

26 5 37 54

[128] [129] [63] [130]

1.93-folds

[131]

Thelavia terrestris T. aurantiacus Phanerochaete chrysosporium Gleophyllum trabeum Chaetomium globosum Aspergillus nidulans G. trabeum Aspergillus niger

Avicel

LPMOs activities for improved cellulase performances. Apart from fungi, LPMOs are widespread in bacteria, and some yeasts are also reported to bear LPMOs. These enzymes have gained the attention of researchers worldwide when they were first reported to be capable of reducing cellulosic ethanol production costs. Addition of these enigmatic GH61/LPMO enzymes in cellulases helped in reducing cellulase protein by at least twofolds [117]. Later, researchers also observed a correlation between enhanced biomass saccharification activity of newer Novozymes cellulase preparations, Cellic Ctec2, and their oxidative LPMO activities [126,132]. Though LPMOs do not display any significant hydrolytic abilities on their own, they have the capability to increase biomass hydrolysis potential when they are present in a blend of cellulase cocktails. There are plenty of reports that have characterized these enzymes and have also demonstrated their synergistic abilities with canonical cellulases. Therefore, utilization of synergistically performing proteins is a great strategy to enhance the biomass hydrolysis via enzymes, which can remove the economic constrain of cellulase usage as well as the hydrolysis of lignocellulose. Hence, the discovery of these wonderful enzymes holds great avenues to deliver an innovative solution for making cellulosic ethanol production process a commercial reality.

13.3.5 MARKET SCENARIO Enzyme applications in industries are profitable as it is a biocatalyst and barely requires any toxic metal ions for its action on the substrate. Therefore, enzymes play a very important role in its action in an eco-friendly way [133]. In the current scenario, industries are focusing their interest toward biorefinery, not only for the use of biofuel (bioethanol) but also for the variable range of value-added products and at this point, enzymes play a very essential part in the bioconversion of

13.4 CONCLUSION AND PERSPECTIVES

263

LC biomass [133]. In the present scenario, cellulase enzymes market is considered to be the second largest industry in the world as they have a vast application in terms of processing of cotton, paper recycling, in juice extraction, as detergent enzymes, and animal feed additives. Nevertheless, cellulase may become a huge industrial sector if it is used in the production of ethanol (biofuel) and bioethanol be used as the main transportation fuel [14]. Cellulase demand is continuously increasing because of its wide application in industries; therefore several companies are involved in the production of cellulase. There are two known cellulase-producing companies and they are “Novozymes,” and “Genencor.” These two companies are doing a significant amount of work on their part in lowering the cost of cellulase. In the past, Genencor (part of DuPont now) produced accelerase 1500, which was a cellulase complex, aimed for industries working on LC biomass processing [14], whereas Novozymes’ recent products are cellicCtec 2, cellicCtec 3, and HTec 3 [134]. It was estimated that in 2014, the value of industrial enzymes was 4.2 billion, and it was assumed that at a compound annual growth rate there would be approximately 7% increase during the year ranging from 2015 to 2020 to achieve the value of 6.2 billion [135]. According to Littlewood et al. [136], enzyme costs impart 18%
43% of the minimum ethanol selling price, which relies on the loading of enzymes during enzymatic saccharification. Khajeeram and Unrean [137] reported that the cost of enzymes forms 44% of the raw material price. It was suggested that on-site or near-site enzyme production is a favorable approach to the noteworthy depletion of enzyme cost up to 30%
70% expecting to its simplified purification and logistics [137]. The formulation of advanced enzymes from both the companies comprising BGL as well as LPMOs supplements, which is an indicator to understand the significance of these auxiliary enzymes for hydrolysis of lignocellulosic biomass.

13.4 CONCLUSION AND PERSPECTIVES There are many challenges that have yet to be overcome, for example, the recalcitrance of lignocellulosic biomass, which necessitates the pretreatment step to open up the fibers and decrease the crystallinity of cellulose, which again adds to the cost of lignocellulosic-bioprocess technology. Pretreatment methods also need to vary from biomass to biomass based on their compositional characteristics. Second, the overall cost of the technology of biomass processing is far from being economically feasible. The most important component that is biomass-processing enzyme still needs improvement in terms of productivity as well as improved properties for enhanced efficiencies. The majority of the available cellulases in commercial applications are produced from both strains of A. niger, and T. reesei in which cellulases from T. reesei are usually deficient of adequate BGL quantity and other auxiliary enzymes to facilitate efficient biomass hydrolysis. Thus, the cellobiose accumulated due to an incomplete conversion caused by the limiting amounts of BGL inhibits exo- and endoglucanases. BGLs are also subject to product inhibition by the glucose beyond certain levels that vary between the different preparations and sources of the enzyme. This issue has been addressed by supplementing BGL having glucose tolerance into the hydrolysis mixture comprising other auxiliary components of cellulase and the resulting cocktail mixture for hydrolysis of lignocellulosic biomass to improve the hydrolysis efficiency.

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CHAPTER 13 PROMISING ENZYMES FOR BIOMASS PROCESSING

Improvement in cellulase efficiency could be done by supplementing deficient component of cellulase into base enzyme or by genetic modification of the host. The latter enables to culture source of cellulase for its production in one vessel for enzyme production, taking care of economic stress. LPMOs gave a new direction to cellulose hydrolysis by enabling cellulase to hydrolyze cellulose easily and are being considered an integral part of cellulase. By genetic modifications, several hosts have been modified to overexpress all the components required for optimum hydrolysis of biomass. Xylanases have been reported to improve hydrolysis significantly when present in high amount in cellulase cocktail. All these enzyme components have been overexpressed as well as their properties such as thermostability have been improved, which allows operating hydrolysis of biomass at elevated temperature and thus less usage of an enzyme. Thus, these improvements have led to realize the dream of lignocelluloses bioprocessing.

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203.

CHAPTER

ENZYME SYSTEMS FOR HIGH-VALUE BIOMOLECULE PRODUCTION

14

Rupinder Kaur and Parmjit S. Panesar Food Biotechnology Research Laboratory, Department of Food Engineering & Technology, Sant Longowal Institute of Engineering and Technology, Longowal, India

14.1 INTRODUCTION The change in the lifestyle of the world population has a strong impact on the eating habits, such as increase in the consumption of processed as well as fast-foods that have an adverse effect on the health and lead to the onset of various diseases, such as diabetes, cancer, allergies, obesity, and others [1]. However, in the current scenario, consumers have become aware of the relationship between diet and health, which has formed the basis for the development of functional foods. Functional foods have become a recent area of research and can be implied to those foods or ingredients that have health benefits beyond their nutritional value [2,3]. Since its introduction, the market size of the functional foods has increased significantly. The global market size for these foods was valued to USD 150 billion, which is expected to rise to USD 250 billion in the coming years Fig. 14.1. The major leaders in this market are the United States, Europe, and Japan. Nevertheless, China, India, and Latin America, as well as other Asian countries are emerging as the fastestgrowing functional food market [5]. Amongst all, prebiotics are considered as one of the outstanding functional foods ingredients having varied health benefits [6]. Prebiotics can be defined as the “selectively fermented food ingredient that allows specific changes both in the composition and/or activity in the gastrointestinal microbiota, thus conferring various health benefits upon the host” [7,8]. For any food to be considered as prebiotic, there are certain criterion such as (1) the food must not be hydrolyzed or digested in the upper intestinal tract by the gastrointestinal enzymes and pass into the colon in the intact form, (2) must be fermented in the colon by the colonic microflora, and (3) must stimulate the activity and/or growth of the beneficial microflora, especially Lactobacilli and Bifidobacteria [9,10]. Among all the foodstuffs, nondigestible oligosaccharides (NDOs) meet all the requirements and therefore are considered as potential prebiotics owing to their structural backbone, which enables them to resist the digestion by the gastrointestinal enzymes [11]. The fermentation of prebiotics in the colon by the colonic microflora results in the production of short-chain fatty acids such as acetate, butyrate, propionate along with other metabolites that impart health benefits on the host [12,13]. The several health beneficial activities of prebiotics include an increase in the bioavailability of minerals, modulation of the immune system, prevention Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00014-4 © 2020 Elsevier B.V. All rights reserved.

273

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CHAPTER 14 ENZYME SYSTEMS FOR HIGH-VALUE BIOMOLECULE

FIGURE 14.1 Revenue of functional food market worldwide. From ,https://www.grandviewresearch.com/industry-analysis/functional-food-market/ . [4].

of cancer, regulation of metabolic disorders such as obesity, prevention of irritable bowel syndrome and other inflammatory conditions, and reduction in different gastrointestinal infections [14 16]. Prebiotics are consumed on a daily basis based on the fact that it is the natural ingredient found in fruits, vegetables, and whole grains. The natural sources include Jerusalem artichoke, chicory, rye, milk, honey, asparagus, onion, leeks, banana, garlic, wheat bran, flour and barley in the range of 0.3% 6% of fresh weight. However, owing to the less concentration, these sources may not be sufficient to impart various functionalities in the human body. Therefore, in order to meet the increasing demand of the consumers, there is a necessity for the production of prebiotics and its incorporation into various foods [17]. Generally, most of the prebiotics are extracted from plants, synthesized either by the enzymatic hydrolysis of plant polysaccharides or by enzymatic transgalactosylation of carbohydrates [18,19], enzymatic transgalactosylation being preferred over the others for its various techno-economic benefits. Keeping the above in view, the chapter provides comprehensive information about the various enzymes involved in biomolecule production, enzymatic transgalactosylation strategies for the production of prebiotics, their extraction and purification approaches and the worldwide status of prebiotics.

14.2 ENZYMES INVOLVED IN PRODUCTION OF PREBIOTICS Enzymatic transgalactosylation reaction occurs by the transfer of the galactosyl moiety or any other sugar molecule to the other carbohydrate molecule [20]. The various enzymes involved in the transgalactosylation reaction for the production of prebiotics have been discussed in the following sections.

14.2.1 β-GALACTOSIDASE β-Galactosidase (E.C. 3.2.1.23), commonly known as lactase is one of the versatile enzymes that have significant potential in food and pharmaceutical sectors owing to its ability to catalyze the

14.2 ENZYMES INVOLVED IN PRODUCTION OF PREBIOTICS

275

hydrolysis reaction as well as transgalactosylation reaction for the production of glucose, galactose, and prebiotics, respectively [21,22]. Generally, lactase belongs to the family of GH 2, however, enzymes of thermophiles, psychrophiles, halophiles also belong to GH 42. In addition, lactase present in multigene plants and fungi are the members of the GH 35 family and that of plants and archaea are of GH 1 [23,24]. It is a tetramer having four identical polypeptide chains, each chain comprising of up to 1023 amino acids that bond together to form five structural domains. Among these five domains, one is a jelly roll barrel, whereas others are fibronectin, β-sandwich and central domain with a TIM-type barrel. This TIM-type barrel is made up of tetramer subunits and acts as the active site for the enzyme. [25]. From the sequence analysis of amino acids, it was revealed that the terminal amino acids consist of α-peptide that is responsible for the subunit interface. The dissociation of tetramer to the dimer form results in the inactivation of the active site [26].

14.2.2 β-GLUCOSIDASE β-Glucosidases (E.C. 3.2.1.21) are small, monomeric enzymes having the functional catalytic properties on a single polypeptide chain. These enzymes catalyze the hydrolysis of O-glycosidic bonds of the carbohydrate molecule to release glycosyl residues and oligosaccharide [27]. These enzymes are present in bacteria and archaea, as well as in eukaryotes and have potential applications in food sector [28].

14.2.3 β-GLYCOSIDASE β-Glycosidases, one of the enzymes of the glycoside hydrolase family, are ubiquitous in nature and can be isolated and characterized from different microorganisms such as archaea, eubacteria, and eukaryotes [29]. These are small monomers having a molecular weight of 49 kDa and are responsible for catalyzing the hydrolysis of (1,4 ) - glycosidic bond for the production of oligosaccharides [30]. Although the different enzymes from the glycoside hydrolase family have been explored for the production of prebiotics, however much of the work has been focused on the application of β-galactosidase in the synthesis of prebiotics.

14.2.4 FRUCTOSYLTRANSFERASE Fructosyltransferase (E.C. 2.4.1.9) or FTase is used for catalyzing the production of fructooligosaccharides (FOS) and possess high transfructosylating activity. It catalyzes the transfer of fructosyl moiety from a sucrose molecule to another molecule acting as an acceptor group [31]. It has been reported that about 91 sequences of FTase belong the glycoside hydrolase family of GH 32 as well as GH 68. These are further grouped into seven clades, which are divided into five for plants, one for bacteria, and one for fungi [32].

14.2.5 β-D-FRUCTOFURANOSIDE β-D-Fructofuranoside (FFase) or invertase (E.C. 3.2.1.26) is another enzyme that catalyzes the production of FOS constituting of one ketose, nystose, and 1-fructofuranosyl nystose [33]. Besides

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CHAPTER 14 ENZYME SYSTEMS FOR HIGH-VALUE BIOMOLECULE

transfructosylating activity, this enzyme is also responsible for the hydrolytic reaction. Similar to fructosyltransferase, it also catalyzes the transfer of fructosyl group from the sucrose molecule to another sucrose molecule by cleavage of β (1,2) linkages. It can be extracted from plants, bacteria, and fungi [34].

14.2.6 INULINASES Another food grade enzyme having the capability of catalyzing the production of FOS is inulinases. On the basis of the cleavage of β (2,1) linkage pattern of inulin, inulinase can be classified as exoinulinases and endoinulinases [35]. Exoinulinases (3.2.1.80) cleave the β- 2,1 bond at the nonreducing end of inulin; thereby releasing fructose and glucose molecule. On the other hand, endoinulinases (3.2.1.7) cleave randomly the internal bonds of inulin and yield FOS [36]. However, both these enzymes exhibit specificity toward sucrose.

14.2.7 GLYCOSIDE HYDROLASES A combination of enzymes belonging to α-glucosidases family and having amylolytic, branching or transferase activity has been used for the production of isomaltooligosaccharides (IMO). α-Glucosidases of the GH 31 family (E.C. 3.2.1.20) utilize maltose as a substrate during the production of IMO [37]. The enzymes belonging to GH 13 family are further subdivided into 40 or more subfamilies, among which subfamily 20 is responsible for the isomaltooligosaccharides (IM) synthesis, which includes maltogenic amylases (E.C. 3.2.1.133), neopullulanases (E.C. 3.2.1.135), and cyclomaltodextranases (E.C. 3.2.1.54) [38,39]. The enzymes of GH 70 family possess transferase activity, whereas those belonging to GH 57 family have branching activity [38,40]. The enzymes of the GH 66 the α-1,6 glycosidic linkages of dextran and result in the production of IMO of varying length [41].

14.3 PRODUCTION OF PREBIOTICS Production of different prebiotics has been carried out by the catalytic activity of the enzymes using varied substrates. Different strategies such as whole cells, crude enzymes, immobilized enzymes/cells, and recombinant enzymes have been employed in order to enhance the productivity and/or improve the bioprocess. The biotechnological routes for the enzymatic synthesis of prebiotics have been discussed in the following sections. The detailed description of the production of prebiotics using lactose and other sugars as a substrate has been depicted in Table 14.1 as well as in Table 14.2.

14.3.1 GALACTOOLIGIOSACCHARIDES Galactooligosaccharides (GOS), one of the promising NDOs, have emerged as the potential prebiotics owing to its various physiochemical and physiological properties. These constitute 2 8 or more monosaccharide units, with glucose being the terminal unit and the other remaining units of galactose linked by glycosidic bonds as shown in Fig. 14.2 [114].

Table 14.1 Production of prebiotics using lactose as a substrate. Process conditions Prebiotics

Microbial source

Equipment

Volume

Enzyme (concentrated)

ILC

pH

Temperature ( C)

2.75 U

30%

7

47.5

15 U/mL

400 g/L

7.5

50

Time

Yield

References

Whole cells GOS

Lactulose

Pediococcus acidilactici Pantoea antophila Bac 55.2 P. antophila Bac 69.1 Sporobolomyces singularis and yeast isolate (NUTIDY007) Kluyveromyces lactis

50 mL

[42] 24 h

136 g/L

[43]

145 g/L Erlenmeyer flasks

100 mL

7%

25%

30

60 h

65%

[44]

20 mL

400 g/L

6.8

40

6h

177 g/L

[45]

40%

7.0

60

3h

20 g/L

[46]

30 h

6.3 g/L

[47]

40%

[48]

K. lactis

10 mL

1.2 U/mL (5.5 mg cell mass/mL) 10.4 g/L

Aspergillus lacticoffeatus Lactobacillus delbruckii Lactobacillus reuteri P. acidilactici Aspergillus oryzae and K. lactis A. oryzae 1 glucose oxidase 1 catalase

5 mL

5 mL

300 g/L

4.5

37

100 mL

50 U/mL

40%

7

40

Crude enzyme GOS

Propionibacterium acidipropionici K. lactis (lactozym 3000 L HP G) K. lactis (Maxilact LGX 5000) A. oryzae

35% 2.75 U 50 U ONPG/g lactose Biostat B fermentation system Microtubes

0.5 L

30% 200 g/L

7 6.5

200 g/L

4.5

47.5 45

1 mL

1.3 U/mL

300 g/L

6.5

45

20 mL

1.5 U/mL

400 g/L

7.0

40

[42] [49]

3 h 20 min

33.1%

4h

54.1 6 1.9 g/L

[50]

24 h

26.8%

[51]

160 g/L

[45]

154 g/L Erlenmeyer flasks

100 mL

0.05%

40%

4.5

50

4h

23%

[52]

(Continued)

Table 14.1 Production of prebiotics using lactose as a substrate. Continued Process conditions Prebiotics

Microbial source

Lactulose

A. lacticoffeatus A. oryzae A. oryzae

K. lactis

A. oryzae A. oryzae

Lactosucrose

Arthrobacter sp. K. lactis Bacillus circulans

Equipment

Volume 5 mL

Fed batch reactor UFX membrane reactor (MWCO 10 kDa) Syringe (50 mL): nonconventional media Aqueous media Erlenmeyer flasks Stirred glass reactor Enzyme reactor Eppendorf tubes

Enzyme (concentrated)

ILC

pH

Temperature ( C)

Time

Yield

References

5 mL 100 IUH/g

300 g/L 50%

4.5 4.5

37 55

30 h 5h

2.5 g/L 40.4 g/L

[47] [53]

10 U/mL

500 g/L

4.6

40

9h

8.67 g/L

[54]

50 μL

10%

6.7

40

150 min

10.4 g/L

[55]

90 min 200 IU/g

50%

4.5

40

5

37

0.90 mL 1 U/mL

0.1 mol/ L 40% 250 g/L 60%

8 g/L 0.282 g/g lactose 30 mmol/L

6 7.5 6.0

37 40 40

250 g/L

6.5

40% 300 g/L

30 mL

1 mL

6

[56] [57]

4h

8 g/L 146 g/L

[58] [59] [60]

59

24 h

20.2%

[61]

4.5 5

40 35

45 min

[62] [63]

100 g/L

5.5

45

48 h

23.3% 24.4 kg/kg lactose 24.2 g/L

0.08 mg/mL

300 g/L

4.5

30

30 h

71.7 g/L

[65]

3 spheres/mL (1 mg/mL) 10 U/g lactose

300 g/L

3

30

8h

30.79 g/L

[66]

43%

6.5

55

42% 44%

[67]

Immobilized cells/enzymes GOS

Aspergillus aculeatus A. oryzae A. oryzae

Round bottomed flask Bioreactor Batch reactor

B. circulans

Packed bed reactor

A. oryzae and Kluyveromyces marxianus B. circulans Bifidobacterium bifidum NCIMB 41171

100 mL 1 kg 10 mL 1 mL 50 mL

Duran bottles

0.499 g

[64]

Lactulose

A. oryzae A. oryzae A. oryzae

Lactosucrose Tagatose

Packed bed reactor Erlenmeyer flasks Erlenmeyer flasks

45 mL

K. lactis B. circulans Commercial β-galactosidase and Lactobacillus fermentum CGMCC2921 cells

Packed bed reactor

700 mL (for hydrolysis) 150 mL (production)

50%

4.5

50

0.6 g/g

[68]

100 IU/g

50%

4.5

50

0.32 g/g

[69]

100 IU/g

50%

4.5

50

0.263 g/g

[70]

12 U/mL 3 spheres/mL (1 mg/mL) 2 g enzyme and 6 g cells

40% 300 g/L

7.5 3

47 30

60 min 8h

15.80 g/L 42.08 g/L

[71] [66]

200 g/L

6.5

39 (initial) and 65 (final)

3 h (initial)

39.9 g

[72]

59.8% 6 0.3%

[73]

24 h (production)

Recombinant enzymes/cells GOS

A. oryzae

Tagatose

Marinomonas sp. BSi20414 Bifidobacterium longum RD 47 Yarrowia lipolytica CGMCC7326 Paenibacillus barengoltzii Thermotoga naphthophila RKU10 Escherichia coli

Lactulose

E. coli Caldivirga maquilingensis IC167 Bacillus stearothermophilus Lactobacillus plantarum FMNP01 Sulfolobus solfataricus

60%

4.5

40

7h

480 mM

7.0

40

5h

36%

8.5

45

140 min

268.3 g/L

[75]

5 mg/mL

500 g/L

5.5

60

6h

160 g/L

[76]

5 U/mL

350 g/L

7.5

40

8h

47.9%

[77]

3.5 U

Flask

500 mL

[74]

Reaction bottles

5 mL

4 mg/mL

0.2 mM

7.0

75

24 h

207.63 mM

[78]

Flask

100 mL

6.2 g of wet cells 20 g wet cells 15 U/mL

50 g/L

6.5

34

24 h

43%

[79]

500 g/L 70%

7.0 4.5

70 85

16 h 4h

20.2% 108 g/L

[80] [81]

200 g/L

6.0

75

4h

8.8 g/L

[82]

1 U/mL

60%

7.0

50

6h

[83]

3 U/mL

40%

6.0

80

6h

18.38 6 2.17 g/L 50 g/L

100 mL

10 mL Tube

[84]

(Continued)

Table 14.1 Production of prebiotics using lactose as a substrate. Continued Process conditions Prebiotics

Microbial source

Equipment

Volume

Enzyme (concentrated)

ILC

pH

Temperature ( C)

Time

Yield

References

400 g/L

102 g/L 40% 50%

[85] [86] [87]

300 g/L

75%

[88]

46 mmol/L

[57]

Other enzymes GOS

Lactulose

S. solfataricus Thermus sp. Halothermothrix orenii Thermus caldophilus Pyrococcus furiosus (free β-glycosidase) P. furiosus (immobilized β-glycosidase) P. furiosus (immobilized β-glycosidase) P. furiosus (free β-glycosidase)

270 g/L

Stirred glass reactor Continuous packed bed reactor Packed bed reactor

30 mL

Membrane reactor

10 mL

0.1 mol/ L

15 g Amberlite IRA-93 15 mg

34 g/L

5

5

75

75

3.5 h

47 mmol/L

5h

43%

1 day

41%

[89]

Table 14.2 Production of prebiotics using other substrates. Process conditions

Prebiotics

Substrate concentrated

pH

Temperature ( C)

810 g/L

5.5

50

104 spores/mL

20%

5.5

30

48 h

Sucrose

40 g/L

600 g/L

50

72 h

Dextran

100 mL

2%

Burdock roots Inulin 1 Sucrose Agavins

30 U/g

100 g

5.68 U/g 100 U/g substrate

5.95% 1 59.87% 30% 33%

4 U/mL

55%

Enzyme

Enzyme source

Substrate

β-Fructofuranosidase

Schwanniomyces occidentalis and Saccharomyces cerevisiae Penicillium citreonigrum

Sucrose

Sucrose

Cladosporium cladosporioides MUT 5506 Bacillus circulans

Enzyme concentrated

Time

Yield

References

166 g/L

[90]

0.65 6 0.06 g FOS/ g sucrose 344 g/L

[91]

Whole cells FOS

β-Fructofuranosidase

β-Fructofuranosidase

IMO

Cyclodextran glucanotransferase

5.5

40

[92]

[93]

Crude enzyme FOS

Endoinulinase Inulinase Endoinulinase Exoinulinase (25%) and Endoinulinase (75%) Inulinase

IMO

Fungamyl 800L transglucosidase α-Glucosidase α-Amylase 1 transglucosidase

Aspergillus. niger

Kluyveromyces marxianus ATCC 16045

A. niger

Sucrose 1 organic solvent (25%) Banana flour Maltose Rice starch

0.3 mL each

50

24 h

6.5

55

9h

4.5

60

48 h 90 min

5.75

45

5.5

60

12 h

1.6 U/mL

600 g/L

4.0

65

24 h

Amylase (3 U/g) 1 transglucosidase (1 U/g)

35%

6.0

60

12 h

29.2% 6 0.8% 674.82 mg/g substrate 20% 10%

[94]

14%

[97]

76.67 6 2.71 g/L 0.56 g IMO/ g maltose 84%

[98]

[95] [96]

[99] [100]

(Continued)

Table 14.2 Production of prebiotics using other substrates. Continued Process conditions

Prebiotics

Enzyme

Enzyme source

Substrate

Enzyme concentrated

Substrate concentrated

pH

Temperature ( C)

Time

Yield

Leuconostoc mesenteroides MTCC 10508

Sucrose

100 mg/L

100 g/L

6.0

30

5h

34.23% 1.546% 30.56% 2.94% 30.52% 2.51% 27.40% 2.761%

References

Recombinant enzymes/cells FOS

Levansucrase

Table sugar Jaggary

β-Fructosidase β-Fructofuranosidase

Endoinulinase

β-Fructofuranosidase

Endoinulinase IMO

Thermotoga maritima Aureobasidium melangogenum 11-1 Yarrowia lipolytica

Candida apicola and Torulaspora delbrueckii S. cerevisiae

Maltogenic amylase 1 αglucotransferase

Bacillus stearothermophilus, Thermatoga maritima

α-Glucosidase

Pichia pastoris

Sugarcane molasses (diluted) Sweet sorghum juice Sucrose

3 μg/mL

1.75 M

5.5

60

Sucrose

117 U/g sucrose

300 mg/mL

4.5

50

60

[101]

6 6 6

28.054% 6 1.0814%

Inulin

786.5 g/L

Sucrose

6 IU

420 g/L

Chicory inulin Liquefied corn starch

10%

200 g/L

Amylase (200 U/g substrate); glucotransferase (6000 AU/g substrate) 62.5 mg/mL

30%

6.5

50

30%

4.0

55

Maltose

6

5

37.9%

[102]

7h

0.66 g FOS/ g sucrose

[103]

2h

0.912 6 0.012 g FOS/g inulin 14.61 g/L

[104]

0.9 g FOS/g inulin 68%

[106]

45

40 14 h

[105]

[107]

[108]

Immobilized enzymes/cells FOS

β-Fructofuranosidase Inulosucrase

Sucrose Sucrose

5 g beads 10 U/mL

60% 200 g/L

5 5.5

62 40

12 h 24 h

59% 35.4%

[109] [110]

Date byproducts

15.6 U

150 mL/100 g

5.0

50

1.5 h

123 g/L

[111]

Dextransucrase

A. japonicus Lactobacillus reuteri 121 Aspergillus awamori NBRC 4033 L. mesenteroides

Sucrose 1 glucose

60 beads or 5.2 g fiber

5.3

25

24 h

α-Glucosidase

A. niger

Maltose

1500 U

1M sucrose 1 1 M glucose 200 g/L

5.0

55

β-Fructofuranosidase

[112]

50.83%

[113]

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CHAPTER 14 ENZYME SYSTEMS FOR HIGH-VALUE BIOMOLECULE

FIGURE 14.2 Chemical structure of galactooligosaccharides.

The enzymatic reaction for the production of GOS is carried out in two steps. The first step involves the formation of enzyme galactosyl complex with the release of the glucose molecule. The second step is carried out by the transfer of the galactose moiety to a nucleophile, preferably a carbohydrate rather than water [115]. Production of GOS is a kinetically controlled reaction, where the hydrolysis and the transgalactosylation reaction compete with each other. In the second step, if the acceptor is water, the reaction process with the hydrolysis resulting in the production of glucose and galactose. However, if the terminal galactosyl acceptor is a carbohydrate (lactose), the reaction leads to the formation of GOS [85]. The source of the enzyme, initial lactose concentration, temperature, and reaction time have a profound influence on the structure, degree of polymerization (DP) as well as glycosidic linkage of GOS [116]. A high concentration of lactose favors the GOS production, thus the initial lactose concentration should be high in order to facilitate the transgalactosylation reaction and increase the yield. However, as the reaction progresses, hydrolytic reaction is favored, which decreases the productivity of GOS. In order to achieve higher GOS yield, the reaction should be stopped close to the point where the yield is maximum. Apart from this, temperature during the course of the reaction is also one of the crucial factors, since it affects the rate of reaction, lactose solubility, and enzyme stability. Above all, optimum pH, temperature as well as kinetic characterization varies with the organisms; therefore source of the enzyme affects the properties of GOS [18,117]. Despite this, other factors that affect the yield are nonconventional media and the reactor configuration. Various aqueous or organic solvent systems have been tested for the GOS production. Biphasic systems have also been explored to achieve high GOS yield and studies have revealed an increase in GOS yield by 19% using cyclohexane and water in the ratio of 95:5 [118]. Apart from this, the reverse micelle system constituting of dicotyl sodium sulfosuccinate/isooctane enhanced the yield to 55% from 31% [119]. Aqueous two-phase system or biphasic systems have proved to be a promising technique due to the partition of products, inhibitors, and enzymes between two phases. However, these systems have certain limitations in terms of lactose solubility, reaction rate,

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285

enzyme activity, and process economics, which hinder their applications at the industrial scale [120,121]. Generally, production of GOS is carried out in a batch reactor; however, product inhibition is one of the major constraints. To overcome this, continuous reactors can be used, which facilitates the continuous removal of glucose and galactose, thereby improving the yield. Although these reactors are economically feasible, residence time, feed flow rate, and permeate collection rate affect the efficiency of the process in these reactors [122,123]. Therefore, reactor designs are the prime factors that affect the productivity of GOS. Various strategies such as crude enzymes, purified enzymes, recombinant enzymes, immobilized enzymes, and whole cells have been investigated for GOS production. Whole cells from Bifidobacterium bifidum, Lactobacillus plantarum, Kluyveromyces lactis as well as other sources were also utilized for the biotransformation of lactose to GOS. Owing to their ease of availability and production, these are considered over the purified enzymes. Moreover, whole cells overcome the limitation of enzyme purification, thus reducing the cost of production. However, permeability barrier is one of the major hindrances, which results in low reaction rate and can be overcome by permeabilization of the cells [124]. Commercial production of GOS is carried out using β-galactosidase from different sources. β-Galactosidase from Aspergillus oryzae, Aspergillus niger, Bacillus circulans, Bifidobacterium infantis, Escherichia coli, Kluyveromyces marxianus, and K. lactis have been used for the production of GOS. The source of the enzyme generally determines the type of GOS synthesized during the reaction. Lactase from bacteria results in the formation of tetra- and pentasaccharides, whereas those obtained from yeast and fungi produce trisaccharides with a small amount of tetrasaccharides. High yields of GOS (90 and 86 g/L) have been achieved from crude as well as immobilized β-galactosidase obtained from Lactobacillus acidophilus ATCC 4356 using 400 g/L lactose at 50 C [125]. In addition, other strategies, such as genetic engineering and protein engineering are being tested to enhance the productivity of GOS. A thermostable β-galactosidase from Marinomonas sp. Bsi20414, expressed in E. coli resulted in high GOS yield constituting of β-1,3 linkages having a better bifidogenic factor as compared to the β-1,4 and β-1,6 linkages [74]. Site-directed mutagenesis of A. oryzae was carried out to produce mutants (N140C, W806F, and N140C/ W806F) that have better transgalactosylation activity than the wild type [73]. The galactooligosaccharide yields 50.7% (N140C), 49.3% (W806F), and 59.8% (N140C/W806F) indicated their efficiency in catalyzing the transgalactosylation reaction as compared to the wild-type strain (35.7%). Apart from lactose, whey has also been used for the production of GOS. Enzyme immobilized on functionalized glass beads was evaluated for their capability in continuous flow packed bed reactor using whey as a substrate [126]. The biotransformation of lactose to GOS in a packed bed reactor is strongly influenced by the feed flow rate and with the increase in the flow rate, GOS concentration increased due to the higher residence time of the reactants in the bioreactor. Although different microbial sources have been explored for GOS production, however, other potential sources having high transgalactosylation activity can also be explored. Besides, productivity of GOS can also be enhanced through enzyme engineering and by the development of efficient as well as cost-effective purification techniques. The detailed information about the biotechnological strategies used for the GOS production has been provided by Panesar et al. [127].

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14.3.2 LACTULOSE 4-O-β-D-Galactopyranosyl-D-fructofuranose, also known as lactulose is a synthetic disaccharide composed of fructose and galactose linked together with β-1,4 glycosidic linkages (Fig. 14.3). These β linkages enable the lactulose to pass into the colon without degradation or digestion in the upper gastrointestinal tract [128,129]. Production of lactulose is generally carried out by the isomerization of lactose by regrouping the glucose molecule on the fructose. This isomerization technique can be divided into three groups, electro-activation, chemical, and enzymatic methods. Various catalysts such as alkaline agents, complex reagents, and enzymes can be employed for the synthesis of lactulose; however, these catalysts must be nontoxic and economical and facilitate the separation from the reaction mixture [129]. Owing to its complexity, less specificity, requirement of high amount of catalyst, hot alkaline conditions, synthesis of undesirable by-products and expensive downstream processing have limited its application for the large-scale production of lactulose [130]. Thus, another suitable approach that has received significant attention in recent years is the use of enzymes for the biotransformation of lactose. In contrast to the chemical synthesis, enzyme-catalyzed reactions are eco-friendly, cost-effective, and do not have the requirement of pure substrates [131]. Enzymatic catalysis is generally being carried out by the enzyme β-galactosidase and glycosidase, which are also responsible for catalyzing the transgalactosylation reaction of GOS. During enzymatic catalysis, lactose gets hydrolyzed into its respective monosaccharides with the action of β-galactosidase to produce galactosyl-enzyme complex. In the second step, the galactosyl moiety is transferred to the fructose molecule; thereby resulting in the production of lactulose [130]. β-Galactosidase from the genera Kluyveromyces and Aspergillus is widely used for lactulose production. Crude β-galactosidase or in whole-cell and immobilized form have been used for the production of lactulose. Biocatalytic activities of the enzyme from the different microbial sources, for example, A. oryzae, E. coli, Kluyveromyces fragilis, and K. lactis were compared, which revealed the highest bioconversion rate from the enzyme of K. lactis [46]. Yet, the yield obtained from enzyme-catalyzed

FIGURE 14.3 Structure of lactulose.

14.3 PRODUCTION OF PREBIOTICS

287

reaction is comparatively lower than by chemical synthesis, which can be attributed to the variation in the enzyme substrate complex formation as well as the structure of the biocatalyst [132]. Apart from this, in order to achieve high yield, higher concentrations of fructose have to be added that lead to the significant proportions of unreacted fructose in the mixture [70]. However, this can be overcome by using fed-batch system, since this system is compatible with those enzyme-catalyzed reactions with poor soluble substrates or at high substrate concentrations. Thus, it was able to obtain high lactulose yield in a fed-batch system, in contrast to the batch system even at high fructose concentration [53]. Lactulose production is a kinetically controlled reaction, where the hydrolysis overweighs the transgalactosylation reaction; thus time is one of the prime factors while carrying out batch production [133]. Regardless of the method used for the production, in order to accomplish higher yields, the process must facilitate the efficient utilization of the biocatalyst that can be accomplished by the recovery of the enzyme at the end of the reaction [134]. Immobilization of the enzyme has a significant effect on the properties of the enzyme as well as on its stability. Thus, the method, as well as matrices for enzyme immobilization, must be selected cautiously. The efficiency of immobilized biocatalyst was assessed in a repeated-batch mode reactor, operated at different lactose-to-fructose ratio [70]. An increase in lactulose yield per unit mass of biocatalyst as well as cumulative productivity was observed in this bioreactor. Moreover, fructose-to-lactose concentration had a considerable effect, not only on the yield, but also on the product distribution, which further affected enzyme stability. Even though lactulose yield is obtained at high fructose/lactose molar mass ratio, it is convenient to work with low fructose/ lactose molar mass ratio in a repeated-batch process, regardless of the type of the immobilization method used and reduction in the yield as well as productivity of lactulose [69]. Low fructose/ lactose mass ratio indicates less residual fructose in the bioreactor that further affects the cost of purification and overall production economy. Whey has also been utilized as a potential substrate for the production of lactulose. Lactulose synthesis was carried out both in the batch and the continuous reactors using immobilized enzyme. In a batch system, the inhibitory effects of galactose and glucose in the reaction mixture were higher; however immobilized enzyme tends to decrease this effect to some extent. In contrast to the batch system, continuous system increased the lactulose yield because it was successful in diminishing the inhibitory effects of the monosaccharides [71]. Approximately 17.3 g/L of lactulose was synthesized at 50 C and pH 7 using immobilized β-galactosidase [135]. Still, the operational stability was restricted as the enzyme subunits get lost during the washings, thus there is a need to execute strategies to stabilize the multimeric enzymes for transgalactosylation reactions. In spite of various biotechnological strategies for lactulose production, still, extensive research is needed to explore the other cost-effective strategies to not only improve and/or enhance the productivity, but also scale-up the production process.

14.3.3 TAGATOSE Tagatose, an isomer of D-galactose is a hexoketose monosaccharide sugar that is seldom found in nature (Fig. 14.4). It has sweetness similar to that of sucrose and is estimated 92% as sweet as sucrose in 10% solution. Yet it has a calorific value of 1.5 kcal/g in humans. For this reason, tagatose has gained tremendous attention in recent years as low-calorie sugar-substitute sweetener [136].

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CHAPTER 14 ENZYME SYSTEMS FOR HIGH-VALUE BIOMOLECULE

FIGURE 14.4 Structure of tagatose.

Chemical production of tagatose can be carried out from D-galactose using calcium catalyst and a strong acid. In spite of being economical, the use of high temperature, pressure and its subsequent effect on the environment have limited its application at industrial scale [137]. Another alternative strategy that has been studied extensively over recent years is the application of biological catalyst, preferably L-arabinose isomerase (E.C. 5.3.1.4) that catalyzes the isomerization of D-galactose to D-tagatose [138]. Generally, tagatose production has been carried out using galactose or dulcitol (galactitol) and only few studies have been done using lactose as a substrate. Lactose, when used as a substrate, is hydrolyzed by β-galactosidase and the synthesized galactose is transformed by the action of L-arabinose isomerase to D-tagatose. Alginate immobilized Lactobacillus fermentum cells along with commercial β-galactosidase were investigated for successful biotransformation of lactose to D-tagatose in a packed bed reactor [72]. Despite being a low-calorie sweetener, the market of tagatose is limited owing to its high price. Therefore, an efficient process using lactose as a substrate and immobilization of Lactobacilli cells can circumvent these problems. Moreover, having generally recognized as safe (GRAS) status, production of tagatose from L. fermentum cells could open up new possibilities for the application of tagatose in food and pharmaceutical sectors. Further research is being carried out to simplify the reaction process for the production of tagatose. So, there is a need for the development of a strategy that can result in the single-step production of high quantities of tagatose. However, the major drawbacks are that the strains of isomerase and β-galactosidase have different operating conditions, especially pH. Hence, a recombinant E. coli strain was constructed by Zhan et al. [79] to coexpress β-galactosidase and L-arabinose isomerase mutant (ECAI-Q299K), having the optimal conditions similar to that of β-galactosidase. The study concluded the efficiency of this process, as higher yields of tagatose were generated. Alginate immobilized L. plantarum cells and β-galactosidase from E. coli for the simultaneous hydrolysis and biotransformation of whey lactose to tagatose resulted in a maximum conversion rate of 42.3% at 50 C and pH 7 during 48 h of reaction [139]. Simultaneous hydrolysis and isomerization reaction for the production of D-tagatose is one of the challenging techniques especially when enzymes with Km values are used because they have low affinity for substrates and require a large amount of substrate concentration to initiate the reaction. Higher yields can be used by using borate as a coupling agent to D-tagatose molecules, thereby shifting the reaction equilibrium. The addition of borate during the reaction did not affect the lactose hydrolysis but had a significant effect on the yield of the tagatose.

14.3.4 LACTOSUCROSE Lactosucrose (O-β-D-galactopyranosyl-(1,4)-O-α-D-glucopyranosyl-(1,2)-β-D-fructofuranoside), also known as lactosylfrucoside or 4G-β-D-galactosylsucrose, is a rare trisaccharide synthesized from

14.3 PRODUCTION OF PREBIOTICS

289

FIGURE 14.5 Structure of lactosucrose.

lactose and sucrose by the transgalactosylation reaction as depicted in Fig. 14.5 [140]. Unlike other oligosaccharides, it has 30% of sweetness as compared to sucrose and is stable for 1 h at pH 4.5 and 120 C [141]. Lactosucrose can be synthesized by the transgalactosylation reaction catalyzed either by β-fructofuransoide as well as other levansucrases or by β-galactosidase [142]. In general, β-galactosidase from B. circulans has been widely used, which catalyzes the galactosyl moiety from the lactose molecule to sucrose that acts as an acceptor and thus results in the formation of tri- and tetrasaccharides having β-1,4 linkages as demonstrated in Fig. 7 [143]. During the lactosucrose production, substrate concentration, reaction time, temperature, and quantity of enzyme are the important factors that have a profound influence on the yield [60]. Similarly, immobilized enzyme from B. circulans was also found to be efficient in the biotransformation of lactose [66]. On the other hand, it was observed that production of lactosucrose is carried out at low temperatures, in contrast to GOS, where the reaction takes place at high temperatures. Although research is being undertaken to enhance the yield, strategies must be devised for the application of thermostable enzymes as well as those enzymes that have high specificity for lactosucrose production and result in higher conversion rate.

14.3.5 FRUCTOOLIGOSACCHARIDES FOS are fructose oligomers composed of 1-ketose (GF2) as shown in Fig. 14.6. Production of FOS is carried out by the transfructosylation of sucrose, in which the β-2,1 of sucrose is cleaved followed by the transfer of fructose moiety to the another molecule, preferably sucrose or fructooligosaccharide catalyzed by fructosyltransferases [144,145]. FOS is about 0.4 0.6 sweet as sucrose and thus can be used as a functional sweetener, thereby expanding its application both in the pharmaceutical and food sectors [146]. Commercial FOS are synthesized by the action of enzymes having transfructosylating activity extracted from fungal sources (Aspergillus japonicas, Aureobasidium, A. oryzae, and A. niger), bacteria (Arthrobacter, Lactobacillus reuteri, and Bacillus macerans), and yeast (Candida and Kluyveromyces sp.). Industrial production is generally carried out from sucrose or inulin as substrates [147]. Unlike GOS, the synthesis of FOS is a kinetically controlled reaction because several reactions occur simultaneously. Various techniques such as whole cells, crude enzyme, and immobilized cells/enzymes have been explored for the FOS synthesis. Biotransformation of sucrose using dried A. niger CGMCC 6640 mycelia as whole-cell biocatalyst had been carried out, which revealed the maximum yield of 314.60 g/L at pH 7.0 and 33 C for 40 h

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CHAPTER 14 ENZYME SYSTEMS FOR HIGH-VALUE BIOMOLECULE

FIGURE 14.6 Structure of fructooligosaccharides.

[148]. Being a simple, efficient, and economic process, this technique has the potential for the scaleup at the industrial scale. Several attempts are being carried out to enhance the productivity of FOS. One step fermentation using Aureobasidium pullulans cells yielded in 0.63 g FOS/initial g sucrose in a shorter duration [149] in contrast to the other studies [150,151] carried out in a 5 L bioreactor. Different enzymes such as FFase and FTase have been used for the production of FOS. Mixed enzyme system constituting of β-fructofuranosidase and glucose oxidase resulted in high FOS yield (approximately 98%) in a stirred tank reactor using glucose and sucrose [152]. Furthermore, FTase from A. oryzae yielded 53% of FOS using 60% sucrose at pH 5.15 and 55 C [153]. Apart from FTase and FFase, inulinases have also been used for FOS production. The catalytic action of inulinase constitutes two parallel paths; one is its transfructosylating reaction for the production of FOS and the other is the hydrolytic reaction that results in the formation of fructose and inulooligosaccharides [97]. Biocatalytic action of inulinases for FOS production gave a maximum yield of 674.82 mg/ g substrate FOS in a mixed substrate of inulin and sucrose during a reaction period of 9 h [95]. Another potential enzyme is levansucrase from Bacillus subtilis that resulted in the higher production of FOS (41.3 g/L) besides levan from 350 g/L sucrose at 35 C for 36 h in a bioreactor [154]. Simultaneous enzymatic production of FOS and levan from sucrose presents a suitable alternative for industrial production because these are biomolecules that have potential health benefits and can satisfy the growing demands of consumers for healthy foods. Above all, other strategies such as genetic engineering were also tested to enhance the enzyme yield and subsequent FOS production. Polyethylene-glycol-mediated transformed A. niger strain ATCC 20611 demonstrated an increase in FFase activity up to 58% and further enhancement in the FOS production in contrast to its wild strain [155]. Improvement in the strain provides an alternative for cost reduction in any bioprocess. However, an efficient genetic engineering approach for the transfer of the desired traits into the host more accurately is an essential requirement. The detailed description of the biocatalytic strategies for the production of FOS has been reported by Bali et al. [33].

14.3.6 ISOMALTOOLIGOSACCHARIDES IMO has emerged as potential prebiotics and has significant applications as low-calorie sweeteners and cariostatic components [156]. These are the branched or cyclic glucooligosaccharide composed

14.3 PRODUCTION OF PREBIOTICS

291

FIGURE 14.7 Structure of isomaltooligosaccharides.

of varying units of glucose (between 2 and 10) linked by α-1,2-, α-1,3-, or α-1,6- glucosidic linkages in combination with α-1,4- bonds (Fig. 14.7). The different types of IMOs include maltooligosaccharides, isomaltose, panose, isopanose, isomaltotetraose, isomaltotriose, nigerose, centose, and kojibose [157]. The DP, type of linkages, its position and proportion influences both the structure and properties of IMO [158]. Moreover, enzymes also play an important role in the diverse structure of IMO. For example, branching enzyme catalyzes the cleavage of the α-1,4 bonds with the transfer of the glucan unit to the glucose unit, thereby resulting in the formation of the branched structure [159]. Similarly, IMO was synthesized from 30% and 50% cassava starch using two methods; first being the simultaneous catalysis of α-amylase and branching enzyme followed by α-transglucosidase. The second method involves the application of α-amylase and branching enzyme followed by β-amylase and α-transglucosidase. Both these methods produced panose and branched IMO having α-1,6 glucosidic linkages and DP between 4 and 7 [156]. To improve the yield as well as process efficiency, mixture of enzymes (pullulanase from recombinant Bacillus naganoensis in Bacillus licheniformis; α-amylase from Bacillus amyloliquefaciens, β-amylase from barley bran and α-transglucosidase from A. niger) were employed for saccharification and transgalactosylation of starch for the production of IMO. Approximately, 49.09% IMO constituting of a mixture of isomaltose, isomaltotriose, and panose was produced after 13 h of reaction [160]. Amongst all, pullulanase is responsible for the acceleration of the saccharification process, thus reducing the time of transgalactosylation reaction. Besides this, recombinant wild type or Y101 mutated amylomaltase in combination with transglucosidase from A. niger was used for the IMO production from the tapioca starch. The highest IMO yield was obtained using 30% of the substrate 120 units of both types of amylomaltase and 6 units of transglucosidase. However, the incubation time varied from 30 min to 1 h while using recombinant wild type amylomaltase and mutated amylomaltase, respectively [161]. Whole Microbacterium cells having 25 IU α-glucosidase activity yielded 85 g/L IMO comprising a mixture of isomaltotriose, isomaltotetraose, isomaltopentaose, and isomaltohexaose using 40% initial maltose concentration after 24 h. Moreover, the addition of LiCl to the reaction mixture enhanced the yield by 15% [162]. Further, scale-up using 100 mL of immobilized cells in a column increased the productivity by 4.2 with a conversion of 24%.

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14.4 DOWNSTREAM PROCESSING OF PREBIOTICS During the biotransformation reactions, the reaction mixture contains appreciable amounts of byproducts such as galactose, glucose, residual lactose, and other sugars, besides the product. As prebiotics are synthesized for human consumption, a high degree of purity is mandatory, which is therefore one of the important processes during prebiotic manufacture [163]. Product purification not only determines the quality of the final product (functional properties and applications), but also its market value and thus is one of the most critical and expensive process operations [164]. Various techniques such as solvent extraction, activated charcoal treatment, and chromatographic techniques such as ion-exchange resins have been frequently used. However, the choice of the technique depends on its impact on the environment, food safety, ease of large scale processing, and cost of the process [165]. In the case of lactulose, alkalizing hydroxides precipitates during the addition of either acids or salts, or even by the incorporation of ethanol. Acidification of the reaction mixture also results in the precipitation of borate or aluminate into boric acid as well as aluminum hydroxide, respectively, resulting in the separation of lactulose [166]. Additionally, borate can also be removed from the lactulose mixture by three methods, namely (1) crystallization in combination of two-column liquid chromatography, (2) passing through the column packed with different resins, and (3) distillation with methanol [167]. Alternatively, ethanol can be used for the crystallization of lactose and thereby its effective separation from lactulose. Moreover, the difference in the lactose and lactulose in terms of alcohol solubility can be used while carrying out the purification. Lactulose purification had been carried out by pressurized liquid extraction at 1500 psi for 30 min, resulting in the purity of over 90% [168]. On the contrary, purification of GOS is quite complex, because the reaction mixture constitutes ample quantities of residual sugars, glucose, as well as unreacted lactose, which is one of the major challenges during its efficient isolation and purification [169]. Purification of GOS using adsorption on activated charcoal not only depends on the molecular weight but also on the functional groups linked with the adsorbent in terms of its type, position, and orientation [151]. Ever since this technique was first reported by Whistler and Durso in 1950 [170], for the separation of sugars, this technique has been widely used for the purification of GOS [171], FOS [144], or other oligosaccharides and sugars [172] using various combinations of adsorbents and ethanol concentrations. Activated charcoal has a large surface area to pore volume, good adsorption capacity, and ease of regeneration of sorbents. In addition, the process is simple, facilitates the separation of large quantities of sugars/oligosaccharides in a single column and is economical, thus have gained the interest of researchers for the separation of biomolecules from the reaction mixture [173]. Fractionation of oligosaccharides using water as a solvent resulted in the removal of monosaccharides as well as 20% of the unreacted disaccharides; yet, selective removal of these sugars was obtained using water and ethanol mixture [174]. The effect of the chemical structure of the carbohydrate on the adsorption behavior results in an efficient separation of different carbohydrates having the same molecular weight but different chemical structure. Selective fermentation of carbohydrate mixture with microbial sources, preferably yeast (Saccharomyces cerevisiae and K. marxianus) is one of the promising techniques in terms of cost, environmental sustainability, and obtainment of high purity products [175]. S. cerevisiae cells have

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the ability to ferment monosaccharides from the reaction mixture; however, these cells are unable to remove disaccharides such as lactose [18]. On the contrary, K. lactis cells have been reported to effectively ferment both monosaccharides as well as disaccharides; thereby resulting in high purity GOS [132]. In contrast to the conventional technique, this technique has a significant technological advantage in terms of high productivity and purity within less time. Apart from this, purification of lactulose had also been carried out using S. cerevisiae and K. marxianus cells, respectively by Guerrero et al. [176]. In addition to this, low-purity IMO produced from rice crumbs and tapioca flour was purified by selective fermentation with yeasts S. cerevisiae and Saccharomyces carlsbergensis and it proved to be an efficient technique for the removal of the sugars from the reaction mixture [177]. Furthermore, being a rich source of nutrients, fermentation rates were higher for IMO syrup obtained from rice crumbs rather than IMO from tapioca flour. Supercritical fluid technology has also been explored for the carbohydrate purification. In the past, this technique was utilized for the extraction of nonpolar components; however in recent years, novel strategies are being applied to separate the polar components such as carbohydrates by using solvent and CO2 mixture [178]. This technique has been applied for the purification of lactulose and tagatose from their respective sugar mixtures. Lactulose purity of 95% or higher with recovery approximately 50% was obtained at 100 bar, 100 C, and 18% of the solvent: ethanol and water in the ratio of 95:5 [179]. However, for tagatose, the optimal operating conditions were 300 bar, 60 C, and 30% isopropanol; under these conditions, 75% of the tagatose was recovered with maximum purity close to 90% [180]. Three-step supercritical extraction processes were employed for the fractionation of GOS [178]. The first step (100 bar, 80 C, and 21% of ethanol and water in the ratio 95:5) resulted in 90% purity with the removal of monosaccharides, whereas second step (100 bar, 100 C, and 14% of ethanol and water in 95:5 ratio) indicated the recovery of 100% disaccharides with purity of around 80%, and third step (150 bar, 80 C, and 21% of ethanol and water in the ratio of 90:10) provided 100% trisaccharides having purity of around 80%. Above all, recently the focus has been shifted toward the use of membrane technology for purification of oligosaccharides. Membrane nanofiltration is advantageous over the other techniques in terms of time, energy, and ease of scale-up [165]. Thus, the efficiency of this technique has been investigated for the separation of sugars from the reaction mixture. However, oligosaccharides with the same molecular weight, such as lactose and trisaccharides, create problem during separation [163]. Thus, in order to obtain high purity products, two alternative strategies can be used; either the use of multistage nanofiltration system [165] or prehydrolysis of lactose prior to purification [181]. Comparative studies were undertaken using two nanofiltration membranes (Synder NFG and TriSep XN45) for the purification of both raw and hydrolyzed GOS in a stirred dead-end cell [181]. Results indicated that incorporation of lactose hydrolysis facilitated the retention of GOS and removal of monosaccharides as well as lactose, thereby improving the GOS purification. On the other hand, the use of low molecular weight cutoff membrane (TriSep XN45) also had a significant influence on the purification yield. Nanofiltration membranes have also been found to be efficient during the purification of FOS. These not only have high rejection for FOS, but also impart low rejection for sugars [182]. To date, constant volume diafiltration has been successfully employed for the separation of sugars from the FOS mixture; however, this technique utilizes ample amounts of dilution solution, which limits its application [183]. In order to overcome this, some studies had reported the recycling of the permeate as in counter-current diafiltration or two-stage diafiltration to reduce the consumption of liquid.

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Yet, these strategies were not efficient because membranes have the selectivity toward the FOS [184]. Thus, another alternative, variable volume diafiltration was proposed. Several strategies have been proposed and investigated for the purification of oligosaccharides; however, most of these are energy-intensive with low levels of purity, thereby limiting its application for industrial use. Membrane technology is a suitable approach for the fractionation of carbohydrates; however, membrane fouling is a major limitation while using this technique. This can be improved by carrying out the process operations under critical transmembrane pressure and under defined temperature, cross-flow velocity, flux stability, and membrane selectivity. In recent years, supercritical extraction technology could also emerge as a potential green and cost-effective technique that has significant industrial applications.

14.5 GLOBAL STATUS OF PREBIOTICS The global status of prebiotics was estimated to be USD 3.34 billion in 2016 and is anticipated to rise at the compound annual growth rate (CAGR) of 10% [185]. The overall market size of prebiotics is predicted to reach USD 7.91 billion by 2025. The major factor for the growth of prebiotics in recent years is rising concerns about the health among the consumers [186]. The prebiotic market is divided on the basis of application, ingredients, and geography. On the basis of applications, the prebiotic market is segregated into various sectors such as food, beverage, dietary supplements, and animal feed. The food and beverage section includes diary, baked foods, cereals, dry food, and fermented meat products, whereas nutritional supplements, infant formula, as well as other food supplements come under the category of dietary supplements. However, the higher demand being the food and beverage sector, especially in the dairy sector where more than 80% of the global market share was accounted for in 2016 and is expected to gain a rise by 9%. This can be attributed to the ease of availability and accessibility of the dairy products that have a significant impact on the growth of the prebiotics market [187]. In 2015, the demand of prebiotics for animal feed was approximately USD 281.9 million; a rise in growth is estimated at the CAGR of 10.5% from 2016 to 2024. The importance of animal protein and its impact on the animal gut health has boosted the demand and global market of prebiotics in animal feed. In terms of consumption, more than 600 kiloton prebiotics was reported to be consumed in 2016 and the market may view the consumption of more than 1.35 million tons by 2024 [188]. On the basis of ingredients, FOS have high demand because of its low caloric value and capability to enhance the food texture. FOS has significant applications in food and beverages, dietary supplements, animal feed, as well as in pharmaceuticals, the major players being China, Japan, United States, Germany, and Europe. Owing to the large demand, FOS market is expected to witness considerable growth by 2024, which is around USD 3.52 billion [189]. The global FOS market is segmented on the basis of its substrate for its production, inulin or sucrose. Sucrose FOSs are mostly used due to the availability of the raw material and in 2015 has accounted for almost 50% of the global market share, followed by inulin, which is the second-largest substrate for FOS production and adds up to 40% of the total market share. After FOS, GOS holds the second-largest market share. The market share of GOS in 2019 was estimated to be USD 600 million and would

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rise up to USD 860 million by 2024 at the CAGR of 6.2% [190]. The market of GOS is segmented into liquid and powder form; the proportion of powder form in 2015 is 65.72% [191]. In contrast to GOS and FOS, according to the global lactulose market report, 2017, lactulose revenue share was USD 144 million in 2013, which rose to USD 148 million in 2016. Further, according to the study conducted by Global Info Research, the market share is expected to increase by up to USD 159 million by 2021 and USD 180 million by 2024 [192]. In terms of Geographical areas, the market is divided into Europe, North America, Asia Pacific, and Rest of the World. Amongst all, Europe led by UK, Germany, and France dominates the prebiotic market and holds the largest market share (40.3%). Further, the market value of prebiotics is anticipated to expand at the CAGR of 10.2% from 2016 to 2024. On the other hand, prebiotic market in North America is also expanding rapidly and may witness growth at 9.7% from 2016 to 2024. The driving factor for rapid growth in prebiotic ingredient in the American market is due to the prevalence of different diseases such as bovine spongiform encephalopathy, porcine epidemic diarrhea virus, and swine flu that has led to the increase in awareness about the animal feed. The demand for prebiotics in Asia Pacific countries such as Japan, China, and India has also increased tremendously over recent years due to the rise in chronic diseases. The estimated revenue from the Asia Pacific market is expected to grow at the rate of 10.7% from 2016 to 2024 [193]. Several companies are involved in the production of prebiotics worldwide. The key players in the prebiotic industry include Kraft Foods Inc., Cargill Inc., Abbott Laboratories, Friesland Campina Domo, Yakult Honsha Co. Ltd., Clasado Ltd., Solvay Pharmaceuticals, and many more. Different companies are focused on the implementation of the new and novel technological strategies for the introduction of new products in the market with an aim to impart various health benefits, besides meeting the increasing demand of the consumers [185].

14.6 CONCLUSIONS AND PERSPECTIVES With the augment of the scientific evidence describing the correlation among the diet, gut microflora, as well as human health, there has been an increasing trend among the consumers for the consumption of functional foods. Prebiotics are one such acclaimed candidate that influence the activity or growth of the gut microbes and impart a positive effect on the host. In order to cut down the price and enhance its availability, enzymatic processes for the production of prebiotics have been investigated. Several enzymes have been tested for their efficiency in catalyzing the biotransformation reaction from various substrates. Although being ubiquitous in nature, microbial sources are preferred owing to various advantages over the other existing sources. Despite various studies have been carried out on the microbial enzymes, there is a need for the novel microbial sources to increase the yield as well as properties of not only enzymes but also the prebiotics. Besides the synthetic substrates, agro-industrial wastes have also been explored for the production of prebiotics from different enzymes. In spite of being a rich source of nutrients, especially sugars, these are generally considered as wastes and disposed off into the environment, thus posing serious environmental issues. Therefore, valorization of the agro-industrial residues can be carried out for the production of high-valued biomolecules. This strategy not only improves the cost of production but also provides a suitable alternative for its disposal. However, to date, only few residues have been

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explored for enzymatic production of prebiotics. Therefore, much research has to be focused to employ various agro-industrial by-products for the production of several other prebiotics and strategies have to be developed to increase the yield. Prebiotics are known to exhibit various health-promoting effects in the host; thus they can be incorporated into a variety of food products as nutritional supplements for the development of functional foods. Although significant scientific evidence has revealed the relationship between the colonic microflora and human health, there are certain challenges in terms of effective, stable, and economical development of functional food that may help to deliver positive health properties on the host. Moreover, the clinical significance of prebiotics, its efficiency, and underlying mechanism remains to be explored. Therefore, research should be focused on the mechanism of prebiotic action, the relevance of its structure corresponding to the function and further to the health. Metagenomics, metabolomics, glycomics, analytical chemistry, clinical human trials, and various ohmic strategies can be considered while examining the mechanism of prebiotics and its function in the host. In vitro studies are also one of the useful tools through which the potential of prebiotics can be validated by animal models. Owing to the various health claims as well as its varied preventive properties, prebiotic market is likely to grow and would replace antibiotics in recent years.

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[177] Y.C. Pan, W.C. Lee, Production of high-purity isomalto-oligosaccharides syrup by the enzymatic conversion of transglucosidase and fermentation of yeast cells, Biotechnol. Bioeng. 89 (7) (2005) 797 804. [178] F. Montan˜e´ s, T. Fornari, A. Olano, E. Ib´an˜ez, Supercritical fluid purification of complex carbohydrate mixtures produced by enzimatic transglycosilation and isomerized with complexating reagents, J. Supercrit. Fluids 53 (1-3) (2010) 25 33. ´ lvarez, A. Montilla, N. Corzo, A. Olano, et al., Selective frac[179] F. Montan˜e´ s, T. Fornari, P.J. Martı´n-A tionation of disaccharide mixtures by supercritical CO2 with ethanol as co-solvent, J. Supercrit. Fluids 41 (1) (2007) 61 67. ´ lvarez, N. Corzo, A. Olano, E. Ib´an˜ez, Selective recovery of [180] F. Montan˜e´ s, T. Fornari, P.J. Martı´n-A tagatose from mixtures with galactose by direct extraction with supercritical CO2 and different cosolvents, J. Agric. Food Chem. 54 (21) (2006) 8340 8345. [181] L. Santib´an˜ez, L. Fern´andez-Arrojo, C. Guerrero, F.J. Plou, A. Illanes, Removal of lactose in crude galacto-oligosaccharides by β-galactosidase from Kluyveromyces lactis, J. Mol. Catal. B Enzym. 133 (2016) 85 91. [182] W.Y. Li, J. Li, J. Shen, C.X. Chen, Study on purifying fructooligosaccharides by membrane separation, in: Proceedings of the Fourth National Membrane Science and Technology Conference Report, Nanjing, P.R. China, 2002, p. 10. [183] X.L. Wang, C.H. Zhang, P.K. Ouyang, The possibility of separating saccharides from a NaCl solution by using nanofiltration in diafiltration mode, J. Membr. Sci. 204 (2002) 271. [184] F. Lipnizki, J. Boelsmand, R.F. Madsen, Concepts of industrial-scale diafiltration systems, Desalination 144 (2002) 179 184. [185] Available from: https://www.grandviewresearch.com/industry-analysis/prebiotics-market. [186] Available from: https://www.marketwatch.com/press-release/global-prebiotics-market-size-to-reach-usd791-billion-by-2025-hexa-research-2019-02-25?mod 5 mw_quote_news. [187] Available from: https://www.benzinga.com/pressreleases/18/10/r12439511/global-prebiotics-marketsize-worth-usd-7-91-billion-by-2025-hexa-res. [188] Available from: https://www.marketwatch.com/press-release/prebiotics-market-size-global-industryanalysis-statistics-forecasts-to-2024-2019-02-08. [189] Available from: https://www.grandviewresearch.com/press-release/global-fructooligosaccharides-fosmarket. [190] Available from: https://www.erienewsnow.com/story/40135577/galactooligosaccharides-gos-market2019-analysis-and-market-expert-inspection-report. [191] Available from: https://www.marketresearchstore.com/report/global-galactooligosaccharidesgos-market2018-industry-production-trends-sales-298551. [192] Available from: http://www.arcognizance.com/report/global-lactulose-market-2019-by-manufacturersregions-type-and-application-forecast-to-2024. [193] Available from: https://www.hexaresearch.com/research-report/prebiotics-industry.

FURTHER READING S.M. An, J.H. Wu, L.F. Qian, Y.L. Gao, Y. Wu, G.P. Yu, Applications of ultrafiltration nanofiltration membrane continuous combination technology for refining of milk derived oligosaccharides, Adv. Mater. Res. 634-638 (2013) 1429 1434.

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M. Barboza, D.A. Sela, C. Pirim, R.G. LoCascio, S.L. Freeman, J.B. German, et al., Glycoprofiling bifidobacterial consumption of galactooligosaccharides by mass spectrometry reveals strain-specific, preferential consumption of glycans, Appl. Environ. Microbiol. 75 (23) (2009) 7319 7325. L.F. Ch´avez-Flores, H.I. Beltran, D. Arrieta-Baez, D. Reyes-Duarte, Regioselective synthesis of lactulose esters by Candida antarctica and Thermomyces lanuginosus lipases, Catalysts 7 (9) (2017) 1 15. A. Co´rdova, C. Astudillo, L. Santiban˜ez, A. Cassano, R. Ruby-Figueroa, A. Illanes, Purification of galactooligosaccharides (GOS) by three-stage serial nanofiltration units under critical transmembrane pressure conditions, Chem. Eng. Res. Des. 117 (2017) 488 499. J. Cui, Y. Li, Q. Wang, J. Li, Y. Ou, J. Wang, et al., Production, purification and analysis of the isomaltooligosaccharides from Chinese chestnut (Castanea mollissima blume) and the prebiotics effects of them on proliferation of Lactobacillus, Food Bioprod. Process. 106 (2017) 75 81. K. Dendene, L. Guihard, B. Balannec, B. Bariou, Study of the separation of lactose, lactulose and galactose by liquid chromatography using cationic ion-exchange resin columns, Chromatographia 41 (9-10) (1995) 561 567. J. Gao, D. Li, Y. Jiang, L. Ma, L. Zhou, Y. He, Pickering emulsion stabilized by immobilized bienzyme nanoparticles: a novel and robust system for enzymatic purification of isomaltooligosaccharide, Ind. Eng. Chem. Res. 53 (47) (2014) 18163 18169. A.K. Goulas, P.G. Kapasakalidis, H.R. Sinclair, R.A. Rastall, A.S. Grandison, Purification of oligosaccharides by nanofiltration, J. Membr. Sci. 209 (2002) 321 335. M. Grambliˇcka, M. Polakoviˇc, Adsorption equilibria of glucose, fructose, sucrose, and fructooligosaccharides on cation exchange resins, J. Chem. Eng. Data 52 (2) (2007) 345 350. O. Hern´andez, A.I. Ruiz-Matute, A. Olano, F.J. Moreno, M.L. Sanz, Comparison of fractionation techniques to obtain prebiotic galactooligosaccharides, Int. Dairy J. 19 (2009) 531 536. K.B. Hong, J.H. Kim, H.K. Kwon, S.H. Han, Y. Park, H.O. Suh, Evaluation of prebiotic effects of high-purity galactooligosaccharides in vitro and in vivo, Food Technol. Biotechnol. 54 (2) (2016) 156 163. R.C. Kuhn, F.M. Filho, Purification of fructooligosaccharides in an activated charcoal fixed bed column, New Biotechnol. 27 (6) (2010) 862 869. R.C. Kuhn, F. Maugeri Filho, V. Silva, L. Palacio, A. Hern´andez, P. Pr´adanos, Mass transfer and transport during purification of fructooligosaccharides by nanofiltration, J. Membr. Sci. 365 (1-2) (2010) 356 365. R.C. Kuhn, M.A. Mazutti, F.M. Filho, Separation and purification of fructooligosaccharides on a zeolite fixedbed column, J. Sep. Sci. 37 (8) (2014) 927 933. R.C. Kuhn, L. Palacio, P. Pr´adanos, A. Hern´andez, F.M. Filho, Selection of membranes for purification of fructooligosaccharides, Desalin. Water Treat. 27 (2011) 18 24. S.E. Ladirat, H.A. Schols, A. Nauta, M.H.C. Schoterman, F.H.J. Schuren, H. Gruppen, In vitro fermentation of galacto-oligosaccharides and its specific size-fractions using non-treated and amoxicillin-treated human inoculums, Bioact. Carbohydr. Diet. Fibre 3 (2014) 59 70. W. Li, Y. Sun, H. Ye, X. Zeng, Synthesis of oligosaccharides with lactose and N-acetyl glucosamine as substrates by using β-galactosidase from Bacillus circulans, Eur. Food Res. Technol. 231 (2010) 55 63. L. Lu, J. Wu, D. Song, H. Zhao, G. Gu, Y. Guo, et al., Purification of fructooligosaccharides by immobilized yeast cells and identification of ethyl β-D-fructofuranoside as a novel glycoside formed during the process, Bioresour. Technol. 132 (2013) 365 369. M. Michelon, A.P. Manera, A.L. Carvalho, F.M. Filho, Concentration and purification of galactooligosaccharides using nanofiltration membranes, Int. J. Food Sci. Technol. 49 (2014) 1953 1961. A. Nath, C. Bhattacharjee, R. Chowdhury, Synthesis and separation of galacto-oligosaccharides using membrane bioreactor, Desalination 316 (2013) 31 41. Q.D. Nguyen, F. Mattes, A. Hoschke, J. Rezessy-Szabo´, M.K. Bhat, Production, purification and identification of fructooligosaccharides produced by β-fructofuranosidase from Aspergillus niger IMI 303386, Biotechnol. Lett. 21 (1999) 183 186.

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K. Nishizawa, M. Nakajima, H. Nabetani, Kinetic study on transfructosylation by β-fructofuranosidase from Aspergillus niger ATCC 20611 and availability of a membrane reactor for fructooligosaccharide production, Food Sci. Technol. Res. 7 (1) (2001) 39 44. S. Pruksasri, T.H. Nguyen, D. Haltrich, S. Novalin, Fractionation of a galacto-oligosaccharides solution at low and high temperature using nanofiltration, Sep. Purif. Technol. 151 (2015) 124 130. M.C. Rabelo, C.S.M. Pereira, S. Rodrigues, A.E. Rodrigues, D.C.S. Azevedo, Chromatographic separation of isomaltooligosaccharides on ion-exchange resins: effect of the cationic form, Adsorpt. Sci. Technol. 30 (8-9) (2012) 773 784. H. Ren, J. Fei, X. Shi, T. Zhao, H. Cheng, N. Zhao, et al., Continuous ultrafiltration membrane reactor coupled with nanofiltration for the enzymatic synthesis and purification of galactosyl-oligosaccharides, Sep. Purif. Technol. 144 (2015) 70 79. M.C. Rodrigues da Cruz, R.Z. Basseto, T.M.B. Bonfim, D. Brand, N.C. Chiquetto, M.M. de Almeida, Production and partial purification of galacto-oligosaccharides by sequential fermentation, Braz. J. Food Res. 8 (4) (2017) 38 56. B. Rodriguez-Colinas, S. Kolida, M. Baran, A.O. Ballesteros, R.A. Rastall, F.J. Plou, Analysis of fermentation selectivity of purified galacto-oligosaccharides by in vitro human faecal fermentation, Appl. Microbiol. Biotechnol. 97 (13) (2013) 5743 5752. V. Sangwan, S.K. Tomar, B. Ali, R.R.B. Singh, A.K. Singh, S. Mandal, Galactooligosaccharides purification using microbial fermentation and assessment of its prebiotic potential by in vitro method, Int. J. Curr. Microbiol. Appl. Sci. 3 (4) (2014) 573 585. L. Santib´an˜ez, A. Co´rdova, C. Astudillo-Castro, A. Illanes, Effect of the lactose hydrolysis on galactooligosaccharides mixtures subjected to nanofiltration: a detailed fractionation analysis, Sep. Purif. Technol. 222 (2019) 342 351. Available from: https://doi.org/10.1016/j.seppur.2019.04.020. J.I. Sanz-Valero, Production of Galacto-Oligosaccharides From Lactose by Immobilized β-Galactosidase and Posterior Chromatographic Separation (Ph.D. Thesis), Ohio State University, Columbus, OH, 2009, p. 70. D. Sen, A. Gosling, G.W. Stevens, P.K. Bhattacharya, A.R. Barber, S.E. Kentish, et al., Galactosyl oligosaccharide purification by ethanol precipitation, Food Chem. 128 (3) (2011) 773 777. K.M.J. Van Laere, T. Abee, H. Schols, G. Beldman, A.J. Voragen, Characterization of a novel β-galactosidase from Bifidobacterium adolescentis DSM 20083 active towards transgalactooligosaccharides, Appl. Environ. Microbiol. 66 (2000) 1379 1384. K. Vaˇnkov´a, Z. Onderkov´a, M. Antoˇsov´a, M. Polakoviˇc, Design and economics of industrial production of fructooligosaccharides, Chem. Pap. 62 (4) (2008) 375 381. M. Wang, H. Admassu, M.A.A. Gasmalla, X. Hua, R. Yang, Preparation of high-purity lactulose through efficient recycling of catalyst sodium aluminate and nanofiltration: a pilot-scale purification, J. Sci. Food Agric. 98 (14) (2018) 5352 5360. L. Wi´sniewski, M. Antoˇsovaʹ, M. Polakoviˇc, Simulated moving bed chromatography separation of galactooligosaccharides, Acta Chim. Slov. 6 (2) (2013) 206 210. Z. Zhang, R. Yang, S. Zhang, H. Zhao, X. Hua, Purification of lactulose syrup by using nanofiltration in a diafiltration mode, J. Food Eng. 105 (1) (2011) 112 118. B. Zhao, L. Zhou, L. Ma, Y. He, J. Gao, D. Li, et al., Co-immobilization of glucose oxidase and catalase in silica inverse opals for glucose removal from commercial isomaltooligosaccharide, Int. J. Biol. Macromol. 107 (2018) 2034 2043.

CHAPTER

ROLE OF ENZYMATIC BIOPROCESSES FOR THE PRODUCTION OF FUNCTIONAL FOOD AND NUTRACEUTICALS

15

Rounak Chourasia1, Loreni C. Phukon1, Sudhir P. Singh2, Amit Kumar Rai1,3 and Dinabandhu Sahoo1,2 1

Institute of Bioresources and Sustainable Development, Sikkim Centre, Tadong, India 2Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India 3Institute of Bioresources and Sustainable Development, Imphal, India

15.1 INTRODUCTION In recent years, growing consumer awareness on influence of dietary components on health has led to increasing demand of functional foods and nutraceuticals [1]. Functional foods are defined as regular foods, which provide health benefits for the prevention or treatment of any specific diseases in addition to basic nutrition properties [2
4]. Enhancement of bioactive compounds in foods is carried out by fermentation using microorganisms [2,4,5] and using commercial enzymes from different sources [6
8]. Food bioprocesses including fermentation and enzymatic hydrolysis have resulted in production of functional foods with one or more health benefits responsible for reduction of risk of specific disease [2,3,9]. The highly efficient catalytic ability of enzymes makes them a preferred candidate in the food industry and attracts researchers in development of enzymatic bioprocess for functional food development [10]. These enzymes provide solubility, stability, and enhanced bioavailability to bioactive compounds apart from disabling antinutritional properties [11]. Commercial enzymes from different sources have been applied in several bioprocesses for (1) production of bioactive compounds by hydrolysis or transformation [7,12] and (2) reduction of antinutritional factors [13
15]. Apart from enhancement of bioactive compounds in foods, enzymatic bioprocess has also been applied for recovery of high-value nutraceuticals from food processing by-products [16,17]. Food bioprocess employing enzyme has resulted in release of various bioactive compounds responsible for functional properties such as antihypertensive, antithrombotic, antioxidant, anticancer, immunomodulatory, opioid, and antiinflammatory activities [3,8,18
28]. The most important enzymes used in bioprocesses for improving food-derived functionality and reduction of antinutritional factors are carbohydrate-modifying enzymes, proteases, lipases, phytases, tannases, and L-asparaginase. Although microbial enzymes are preferred over other sources due to factors including low cost of production, process controllability, easy optimization, and manipulation Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00015-6 © 2020 Elsevier B.V. All rights reserved.

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of enzymes for target function, enzymes extracted from plants and animals have found great commercial importance in the food industry [6,9,29
34]. This chapter highlights the research based on the diverse enzymes-based bioprocesses applied for the production of functional foods.

15.2 ROLE OF ENZYMES IN PRODUCTION OF FUNCTIONAL FOODS Enzymes are the backbone of several food bioprocesses for production of functional foods and nutraceuticals. These enzymes are responsible for production of functional foods and reduction of antinutritional factors (Fig. 15.1). The role of different enzymes and their role in production of functional foods are discussed in this section.

15.2.1 PROTEASES Hydrolysis of peptide bonds by proteases releases protein hydrolysates, peptides, and amino acids, resulting in enhanced functional and nutritional properties of foods such as bioactive peptide liberation and alleviation of protein allergy [10,20]. Proteases are classified into seven groups: aspartic, serine, threonine, cysteine, glutamic, and metalloproteases and asparagine peptide lyases and have promising potential in functional food production. Food-derived bioactive peptides produced by

FIGURE 15.1 Role of enzymes derived from plant, animal, and microbial sources in development of bioprocesses for production of functional foods and nutraceutical compounds. GOS, Galactooligosaccharides; FOS, fructooligosaccharides; RS, resistant starch; PUFA, polyunsaturated fatty acids; CLA, conjugated linoleic acid.

15.2 ROLE OF ENZYMES IN PRODUCTION OF FUNCTIONAL FOODS

311

protease-mediated hydrolysis during enzymatic or fermentation processes demonstrate various functional properties including antihypertensive, antithrombotic, antioxidant, anticancer, immunomodulatory, opioid, and antiinflammatory activities [4,35]. The activities exerted by bioactive peptides on health depend on peptide size and amino acid sequence; thus specific proteases along with protein substrate and hydrolysis conditions affect biological activity of peptides generated (Table 15.1). Apart from proteases from plants and animals’ origin, proteases produced by microorganisms inhabiting extreme environments have expressed high stability and catalytic properties. Thus a huge plethora of proteases can be exploited for development of foods with diverse and effective functional properties. Angiotensin-I-converting enzyme (ACE) inhibitory peptides have been widely studied for treatment of hypertension and related target organ damage [5]. Preparation of ACE inhibitory peptides from food proteins such as casein and whey proteins by enzymatic hydrolysis offers several advantages such as low cost, retained nutritional value of proteins, specificity of enzymes in the type of

Table 15.1 Proteases associated with bioactive peptide production and peptide bioactivity. Substrate

Enzyme

Source

Bioactivity

References

α-Casein

Proteinase

ACE inhibition

[36]

κ-Casein

Cysteine protease Rennet Pepsin Alkaline protease Corolase Trypsin Papain

Lactobacillus helveticus CP790 Ficusjohannis

Immunomodulation

[32]

Mucor miehei Pig gastric mucosa Bacillus licheniformis Pig pancreas Bovine pancreas Carica papaya

ACE inhibition, metal chelation Antioxidant properties ACE inhibition

[33] [37] [8]

Antioxidant properties Reduced allergenicity ACE inhibition

[38] [39] [29]

Pig pancreas Bacillus thermoproteolyticus Bacillus amyloliquefaciens C. papaya Thunnus alalunga

Anxiolytic properties ACE inhibition

[40] [36]

Antioxidant, scavenging, metal chelation, ACE inhibition Antioxidant activity Antioxidant activity

[6]

Corolase LAP Alcalase

Aspergillus sojae

ACE inhibition

[41]

Bacillus subtilis

ACE inhibition

[41]

Flavourzyme

Aspergillus oryzae

ACE inhibition

[23]

Casein α-Casein Casein β-Lactoglobulin β-Lactoglobulin Bovine serum Albumin Ovolin α-Zein Egg-yolk protein preparation Fish protein Abalistes stellaris muscles Whey protein concentrate Whey protein concentrate Whey protein concentrate

Trypsin Thermolysin Neutrase Papain Trypsin

ACE, Angiotensin-I-converting enzyme; LAP, Leucyl aminopeptidase.

[9] [31]

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CHAPTER 15 ROLE OF ENZYMATIC BIOPROCESSES

peptide production by hydrolysis of protein at specific position, and a control over the hydrolysis process [8,42]. Hydrolysis of skimmed goat milk by alkaline protease showed a high degree of hydrolysis and ACE inhibitory activity, thus indicating the potential use of enzyme hydrolysis of milk in industrial production of functional milk products [8]. Production of hard cow milk cheese using rennet from microbial origin resulted in release of peptides with high antioxidant activity [33]. Peptides EIVPN and DKIHPF in addition to the newly identified peptide VAPFPQ showed high ACE inhibitory activity and revealed high metal chelating activity. Papain, a plant cysteine endopeptidase has been observed to generate bioactive protein hydrolysates from fish with antioxidant, antidiabetic, immunomodulatory, ACE inhibitory, and metal-binding properties [3]. The hydrolytic potential of papain has been studied in the production of cheddar cheese using different types of milk (soy, cow, and goat) [43]. Fish protein hydrolysates produced using papain have demonstrated high antioxidant activity showing its potential as functional food additive for application in functional food industry [9]. Walnut protein hydrolyzed by papain resulted in release of hydrolysates with 81% purity and demonstrated one of the highest antioxidant properties [44]. In addition to bioactive protein hydrolysates, papain reduced allergenicity of soy protein isolates, while enhancing sensory, technical, and functional properties [45]. The ability of papain to recognize Gln-Pro-rich allergenic wheat proteins such as gliadins has allowed the use of papain in production of hypoallergenic flour [9,46]. Trypsin, a serine protease of animal origin has been observed to hydrolyze bovine β-lactoglobulin, resulting in reduced allergenicity in milk-based foods [39]. In another study, trypsin from liver of albacore tuna (Thunnus alalunga) was used to hydrolyze starry triggerfish (Abalistes stellaris) muscle resulting in the production of protein hydrolysates with high antioxidant and metal chelating activity, which can be used as a bioactive ingredient in functional foods and as natural antioxidants in lipid food systems [31]. Low production cost and high yield resulting in large-scale production of microbial proteases have promoted the growth of microbial protease market as compared to plant and animal proteases [47]. Microbial rennet produced by strains of Rhizomucor miehei, Endothia parasitica, Aspergillus oryzae, and Rhizomucor pusillus has provided cheese industry with an effective solution to the high demands of the animal rennet chymosin, such that microbial rennet is being used for 33% cheese production worldwide [48]. Whey protein hydrolysates produced using a mixture of proteases from Lactobacillus helveticus LB13 showed 92.2% ACE inhibition activity [49]. Pancreatin and proteases from Bacillus licheniformis along with proteases from Asp. oryzae and Aspergillus sojae have been used as alternatives of trypsin and protease 2A for denaturation of whey proteins lactalbumin and lactoglobulin with high effectiveness [23,41]. Neutrase, a commercial protease produced by Bacillus amyloliquefaciens has been used to hydrolyze egg-yolk protein preparation (YP), obtained as by-products of lysozyme, lecithin, and cystatin isolation and white protein preparation (WP) obtained during ethanol extraction of lecithin from egg yolk. YP and WP hydrolysates produced using neutrase showed high antioxidant activity, scavenging capacity, and chelating property along with significant ACE inhibitory activity, indicating that YP and WP hydrolysates can be used as enhancers of functionality and natural antioxidants in food [6].

15.2.2 CARBOHYDRATE-MODIFYING ENZYMES Carbohydrases catalyze the breakdown of carbohydrates and have wide applications in the functional food industry. The most important carbohydrate-modifying enzymes belong to the microbial

15.2 ROLE OF ENZYMES IN PRODUCTION OF FUNCTIONAL FOODS

313

origin and include amylases, inulinases, galactosidases, glucosidases, fructosyltransferases, glucosyltransferases, and pectinases. These enzymes catalyze the formation of different functional compounds such as prebiotics, including galactooligosaccharides (GOS), fructooligosaccharides (FOS), and resistant starch (RS), and reduce lactose in milk [26] (Table 15.2).

15.2.2.1 β-Galactosidases β-Galactosidases are exoglycosidases that cleave the β-bond of galactosides to release galactose and another organic moiety. Gene products of lac Z and member of family 1 of the glycoside hydrolases belonging to carbohydrate active enzymes (CAZymes) [57], and some β-galactosidases show the ability to cleave α-L-arabinosides, β-D-fucosides, and β-D-glucosides [20]. β-Galactosidase catalyzes two reactions in milk: hydrolysis of lactose and conversion of lactose to allolactose [58]. Hydrolysis of lactose results in the production of lactose-free milk, which serves functionality to Table 15.2 Carbohydrase-catalyzed production of functional foods and associated bioactivities. Enzyme

Source

Product

Bioactivity

References

β-Galactosidase

Lactobacillus delbrueckii subsp. bulgaricus ATCC 11842 Aspergillus oryzae Asp.s oryzae DIAMF Aureobasidium pullulans Lactobacillus gasseri DSM 20604 Bacillus lentus Penicillium multicolor Bifidobacterium longum KACC 91563 Lactobacillus kimchi

Lactose-free dairy foods

[50]

GOS FOS

Lactose hydrolysis, digestibility of dairy foods by lactoseintolerant individuals Prebiotic Prebiotic

FOS

Prebiotic

[25]

FOS, MFOS

Prebiotic

[28]

RS Gentiooligosaccharides

Prebiotic Prebiotic

[52] [18]

Ginsenoside Rd and spirostane glycosides (bioactive saponins) Resveratrol

[53]

Jeotgalibacillus malaysiensis Thermotoga naphthophila RKU-10

Salicin

Antitumor, antiinflammatory, and antiallergic properties Antiinflammatory, cardioprotective, antioxidant, and anticancer properties Antiinflammatory property Prebiotic

β-Galactosidase Fructosyltransferase (FTase) Fructosyltransferase (FTase) Inulosucrase

Isoamylase β-Glycosidase β-Glucosidase (BlBG3) β-Glucosidase

β-Glucosidase (BglD5) β-Glucosidase

Galactotrisaccharides (GOS3)

GOS, Galactooligosaccharides; FOS, fructooligosaccharides; MFOS, Maltosylfructosides; RS, resistant starch.

[26] [51]

[54]

[55] [56]

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product against lactose intolerance, a condition that affects 70% of the adult population [20]. β-Galactosidase was produced from Lactobacillus delbrueckii subsp. bulgaricus ATCC 11842, and the enzyme was used for the hydrolysis of lactose in milk and dairy products [50]. The conversion of lactose to allolactose occurs through the transgalactosylation mechanism, which results in the increased production of oligosaccharides [22]. β-Galactosidase from Asp. oryzae was used to synthesize GOS from partially dissolved and supersaturated solution of lactose [59]. GOS are a group of nondigestible prebiotic fibers that stimulate the growth of probiotic bacteria in the gut such as lactic acid bacteria (LAB), bifidobacteria, and bacteroides [60]. Probiotic bacteria control certain central nervous system disorders via biochemical signaling in the microbiome brain-gut axis [61]. A new class of probiotics named as psychobiotics impart antidepressant and anxiolytic activities that relieve symptoms of chronic fatigue syndrome and depression [62]. Apart from acting as “food” for probiotic bacteria, GOS upon fermentation in the large intestine produce bioactive compounds such as short-chain fatty acids (SCFAs). SCFAs such as butyric acid, acetic acid, and propionic acid promote colon mucosa cell health and thus help control colon inflammation [63]. GOS are highly valued functional food ingredient and prebiotic with proved health benefits demonstrated by human clinical trials [64] and are used in a wide variety of functional foods [22].

15.2.2.2 Enzymes catalyzing fructooligosaccharides production FOS are small dietary fibers with low caloric value, Generally Recognized as Safe status, and prebiotic properties [65]. Rapid growth in industrial production of FOS has been observed due to various aspects including rise of healthy sugar market relying on FOS’s prebiotic function [66] and conversion of sugar industry waste into nutraceutical products [67]. The substrates involved in FOS production include sucrose and inulin, the conversion of which to FOS is catalyzed by fructosyltransferases and inulinases, respectively. FOS are synthesized from sucrose by the enzymes β-Dfructosyltransferase or β-fructofuranosidase via sucrose hydrolysis, releasing fructose and glucose and transferring the fructosyl moiety to another acceptor molecule of FOS or sucrose [68]. Fungal enzymes are commonly used for production of FOS from sucrose including enzymes from Aspergillus, Penicillium, and Aureobasidium [51,69,70]. One of the most widely followed protocol for FOS production from sucrose is via the transfructosylation of β-fructofuranosidase from Aureobasidium pullullans [25]. Inulosucrase, isolated from Lactobacillus gasseri catalyzes the transfer of fructose moieties of sucrose to the reducing end C-1 and at C-6 of the nonreducing end of maltose, resulting in the release of neoerlose and erlose [28]. Levansucrases are another example of bacterial extracellular enzymes that act on sucrose and produce β-2,6-linked fructans of different chain sizes [71]. Inulin is the second most abundant polysaccharide in nature after starch and its hydrolysis catalyzed by endo-inulinases results in the release of FOS. Although higher content of FOS is released upon catalysis by endo-inulinases that with transfructosylation of sucrose, endo-inulinases have not been considerably used in scale-up production of FOS [72]. Microorganisms that produce considerable amounts of endo-inulinases include Pseudomonas sp., Aspergillus sp., Bacillus sp., Yarrowia sp., Penicillium sp., and Xanthomonas sp. [65]. The production of FOS using endo-inulases from Pseudomonas sp. achieved higher yields (75.6% and 80%) as compared to FOS production using commercial endo-inulinases (65%), highlighting the scope of exploiting microbial endo-inulinases for efficient commercial production of FOS [73,74]. FOS production using purified and commercial enzymes has been studied extensively. A study using a stirred tank reactor under controlled

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conditions employed a mixture of two enzymes, glucose oxidase, and β-fructofuranosidase (Aspergillus niger ATCC 20611) [75]. Various studies focused on FOS production catalyzed by immobilized enzymes [76] used pretreated molasses as the substrate and the reaction was catalyzed by a membrane reactor with immobilized enzymes. In another study, FOS production was studied using free, immobilized, and pretreated immobilized inulinases on an aqueous-organic system [77]. Production of FOS with sucrose as the substrate using aqueous and aqueous-organic system with immobilized inulinase was indicated by Ref. [78].

15.2.2.3 Enzymes catalyzing xylooligosaccharides production Xylooligosaccharides (XOS) have recently emerged as prebiotics with more desirable properties as compared to FOS and inulin due to its stability at low pH and high temperatures, thus making it suitable for being incorporated as prebiotic ingredient in functional foods such as yogurt and other fermented dairy products [79]. Functional properties of XOS include antioxidant effects, lipid metabolism, reduction in cholesterol levels, antidiabetic properties, reduced risk of colon cancer, improved calcium absorption, and promotion of growth of probiotic intestinal microbiota [80
82]. A study reported that consumption of 4 g of XOS daily by elderly individuals above 65 years old led to improvement of intestinal microbial population [83]. Hemicellulose, a heteropolymer comprising mainly of xylans and arabinans is among the three major components of lignocellulosic biomass besides lignin and cellulose [82]. The xylan fraction of hemicellulose ranges from 60% to 85%, which can be converted to XOS upon breakdown by either thermochemical methods or enzymatic processes [79]. Large-scale production of XOS prebiotics from lignocellulosic biomass generated from agricultural and industrial processing could be very beneficial to the food industry [84,85]. However, XOS production by thermochemical processes results in the generation of undesirable by-products such as furfural and hydroxymethylfurfural [79]. Thus the development of effective and cheaper XOS production methods including enzymatic processes has been attaining greater significance lately. Xylanase (endo-β-1,4-xylanase), β-xylosidase, and acetyl esterases catalyze the breakdown of xylan into xylose and XOS. Xylobiose and xylotriose, the most active prebiotic XOS were produced from pineapple peel extract by enzymatic hydrolysis using endo-1,4β-xylanase M1 from Trichoderma viride. At optimum operating parameters, XOS production peaked at 25.6/100 g hemicellulose [86]. In another study, endoxylanase from Thermomyces lanuginosus was immobilized on Fe-based magnetic nanoparticles and used for the production of XOS by hydrolysis of almond shell hemicellulose [87]. A new endoxylanase produced by the marine bacteria Pseudoalteromonas atlantica was active on heteroxylans from various sources [88]. XOS production from wheat straw hemicellulose by a combination of endo-1,4-β-xylanase and β-glucosidase resulted in achievement of 91% conversion rate of substrate to XOS [89]. As compared to thermochemical production of XOS, enzymatic processes provide environmentally friendly method that avoids the use of poisonous chemicals and release of noxious by-products due to higher specificity and efficiency presented by xylanolytic enzymes [90]. Recently, several studies have approached optimization of enzymatic processing methods for reduction of XOS production costs. The most effective and simplified method for XOS production devised includes enzymatic hydrolysis and direct fermentation of lignocellulosic biomass by microbes [80]. Direct enzymatic hydrolysis of premilled rice straw, corn cobs, and rice husk resulted in production of cheap XOS [91,92]. In another study, direct

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fermentation of Brewer’s spent grain to produce arabinoxylooligosaccharides was found to be more advantageous as compared to hydrolysis by commercial enzymes [93,94].

15.2.2.4 Enzymes catalyzing resistant starch production Starch is the main source of carbohydrates in legumes, cereals, and roots [95]. However, a significant amount of starch called Resistant Starch (RS) is not completely digested and undergoes fermentation in the colon [96]. Fermentation of RS in the colon yields SCFAs, organic acid, alcohol, and gases. Due to this reason, the biological effects of RS are comparable to those of dietary fibers [97,98]. Foods rich in RS reduce postprandial glycemia in humans and thus can be used in the control of type II diabetes [96]. The slow digestion of RS reduces glycemic response along with the release of postprandial insulin adding the increase in period of satiety. SCFAs bind to peripheral tissues and function in the signaling of insulinemic responses and glucose homeostasis. In addition, SCFAs obtained from RS have shown to exert alteration in the expression of hypothalamic neuropeptides [98], participate in the integrity of intestinal epithelial barrier resulting in attenuation of local and systemic inflammation, stimulate intestinal blood flow resulting in the prevention of intestinal ischemia, and impart positive effects in the chronic kidney disease (CKD) population [99]. Manufacture of RS by physical methods results in an unsatisfactory yield of 20%
30% and also arouses food safety issues [10,100
103]. Production of RS by enzymatic or a combination of enzymatic and physical methods reduce the amount of substrate required for RS production and also avoid food safety problems related to RSrich food production [52]. Enzymatic processes for RS production utilize debranching enzymes such as pullulanase and isoamylase that reduce molecular weight of the substrate and increase amylose yield, increasing the RS content up to 50% [52]. Thermostable isoamylase has been observed to dismantle starch with great specificity and efficiency, releasing smaller branches composed of 1 and 3 glucose residues differing from the substrate specificity of other known isoamylases. In another study, a novel strategy was developed for RS production combining isoamylase and amylase [52].

15.2.2.5 β-Glucosidases β-Glucosidases hydrolyze glycosidic bonds of soluble cellodextrins and cellobiose into glucose molecules from nonresidues in β-D-glucosides and are important for hydrolytic glucose degradation essentially in lignocellulosic biomass [104
106]. Fungal β-glucosidases are preferred for industrial use because of thermal and pH stability and high activity of the enzymes [107]. Nevertheless, many studies have shown the potential use of β-glucosidases extracted from other sources as well such as yeasts and intestinal bacteria in the production of bioactive compounds that can be considered as functional food or can be used as an ingredient in the development of functional food [12,53,108]. Gentiooligosaccharides including gentiotetraose, gentiopentaose, and gentiohexaose are produced from hydrolysis of substrates such as lichen polysaccharide [109] and gentiobiose [18] by twostage transglycosylation through a combination of β-glucosidase and endo-β-(1,6)-glucanase from Penicillium multicolor. Novel glucose and ethanol-tolerant β-glucosidase produced by Pichia guilliermondii G1.2 was studied for its potential application in winemaking and saccharification of cellulose [21]. Resveratrol is known for various bioactive properties including antioxidant, antiaging, antimicrobial, and antiviral effects and is also used to treat menopausal disorders [110
112].

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Saponins are natural glycosides that exert antitumor, antiinflammatory, and antiallergic effects upon consumption but are dependent on the enzymatic metabolism by intestinal microbiota [113
115]. Intestinal bacteria belonging to genera Eubacterium, Bacteroides, Clostridium, Fusobacterium, Bifidobacterium, and Prevotella are known for participation in saponin biotransformation via secretion of various carbohydrate-hydrolyzing enzymes. A putative β-glucosidase (BlBG3) of the glycoside hydrolase (GH) 3 family was isolated from Bifidobacterium longum KACC 91563 and upon in-depth molecular, structural, and functional characterization was found to hydrolyze saponins [53]. In another study, a stilbene glucosidespecific β-glucosidase was isolated from Lactobacillus kimchi and purified, and functional characterization was performed [54]. The enzyme converted polydatin into resveratrol with a 100% yield and was used effectively in the production of bioactive compounds. Several studies have reported the isolation, purification, and characterization of β-glucosidases from extremophilic bacterial strains. A novel β-glucosidase (BglD5) belonging to the glycoside hydrolase family 1 (GH 1) produced by the halophilic bacteria Jeotgalibacillus malaysiensis was purified and characterized [55]. The enzyme efficiently hydrolyzed salicin, improving antiinflammatory property of the compound, along with other compounds such as cellobiose, cellotriose, cellotetraose, cellopentose, and cellohexanose. In another study, β-glucosidase from the hyperthermophilic anaerobe Thermotoga naphthophila RKU-10 was used for the production of prebiotic galactotrisaccharides (GOS3) with milk processing waste lactose as the substrate [56].

15.2.3 L-ASPARAGINASE Fried and baked foods are liked and consumed worldwide and these foods are derived from plant sources such as potato and various grains [116]. These food products are associated with high generation of the carcinogen acrylamide. Reaction between the amino acid asparagine and reducing sugars via Maillard reaction generates high levels of acrylamide in fried and baked foods [116]. Strategies aiming reduction of acrylamide in fried and baked foods are application of additives; controlling the pH, temperature, and cooking time; and reduction of acrylamide precursors [117,118]. However, transformation of the main precursor of acrylamide, asparagine by the hydrolyzing enzyme L-asparaginase into aspartic acid and ammonia results in decreased acrylamide with minor impact on the technological properties of the product [119]. The effects of L-asparaginase activity on gingerbread, crispbread, wheat-oat bread, french fries, biscuits, fried dough, coffee, potato chips, tortilla chips, and cookies have been studied [10]. L-Asparaginase from Asp. oryzae CCT 3940 decreased acrylamide concentration by 72% in fried potatoes, with additional increase of product quality by maintaining the glutamine levels in the sample [120]. Novel asparaginase extracted from Aquabacterium sp. A7-Y reduced 88.2% acrylamide from potato strips [115]. In another study, the use of a purified asparaginase from Pseudomonas oryzihabitans resulted in 90% decrease of acrylamide in fried potatoes [15]. L-Asparaginases are produced by a wide variety of organisms including microorganisms, plants, and animals and all these enzymes have proven to be effective in the reduction of acrylamide in baked and fried products [14,116]. The production of the first completely “acrylamide-free” commercially available biscuits in Germany was announced in 2008 where the biscuits were treated with preventase (extracted from A. niger) [10]. Another commercially available asparaginase is

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Acrylaway, which is extracted from Asp. oryzae and manufactured by Novozymes [10]. These enzymes are used in more than 30 countries for the industrial production of coffee, biscuits, snacks, and French fries [10].

15.2.4 LIPASES Lipases hydrolyze triacylglycerols (TAG) at the lipid
water interface to free fatty acids and glycerol. The ability of lipases to catalyze the production of fatty-acid derivatives such as fatty acid esters of antioxidants, structured lipids, and polyunsaturated fatty acids (PUFA) places the enzyme among the most important enzymes in the food industry [121] (Fig. 15.2). Mono and diacylglycerides (DAG)-enriched oils have demonstrated antiobesity properties and are soon expected to replace TAG-enriched oils [122]. DAG oils are considered as good fat substituent as they positively affect lipid metabolism, decrease TAG in adipose tissue, reduce body weight, and lower abdominal and liver fat concentration [123]. Lipases have earned importance in the enzyme-catalyzed production

FIGURE 15.2 Lipase-catalyzed production of bioactive molecules including the polyunsaturated fatty acids: diacylglycerides (DAG) and monoacylglycerides (MAG); omega-3 fatty acids: eicosapentaenoic acid (EPA), docosahexaenoic acid (DHA), and conjugated linoleic acid (CLA). These fatty-acid derivatives demonstrate functional properties such as antiobesity, antihypertensive, antiinflammatory, and antioxidative activity.

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of novel structured lipids (modified TAGs) as an advantageous alternative to chemical processes by presenting functional properties to edible oils and fats [10]. Omega-3 fatty acids including stearidonic acid, α-linolenic acid, docosapentaenoic acid, eicosapentaenoic acid (EPA), and docosahexaenoic acid (DHA) are PUFAs that are well known for health beneficial effects and have high potential in functional food industry [20,124]. DHA and EPA demonstrate protective effects against cardiovascular diseases such as arrhythmias, hypertension, and atherothrombosis in addition to being potent regulators of antiinflammatory, antiapoptotic, neuroprotective, and improved attentional and physiological functions, particularly those involving complex cortical processing [125]. Omega-3 fatty acids are concentrated by lipase hydrolysis of oil, which includes partial removal of monounsaturated and saturated fatty acids. For this, the location of the fatty acids in oil and hydrolysis selectivity of lipase is of great importance. Lipases release fatty acids upon hydrolysis at specific positions on the glycerol backbone. Most fish oils have DHA and EPA located at the middle (sn-2) position and their concentration requires lipases that hydrolyze fatty acids on the outer (sn-1 and sn-3) positions of the glycerol backbone. Lipase TL 100 L from T. lanuginosus has been used successfully for concentration of DHA and EPA from anchovy oil [126]. An increase in DHA and EPA concentration from 15% and 30% to 35% and 60%, respectively, was observed upon hydrolysis of anchovy and seal oils by porcine pancreatic lipase [127]. Lipase A (CAL-A) from Candida antarctica combined with T. lanuginosus lipase (TL 100 L) was used to hydrolyze alga oil resulting in generation of approx. 90% pure DHA concentrate [128,129]. Conjugated linoleic acid (CLA) consists of a group of linoleic acid (LA) isomers with trans-10, cis-12, and cis-9, cis-12 being the dominant isomers. CLA is the most active antioxidant in milk fat and is present in various dairy products along with ruminant meat. Functional properties of CLA include body fat mass reduction, antiatherogenic and antidiabetogenic activities, and positive effects on atherosclerosis [130
133]. In addition, CLA shows better inhibition of neoplastic growth than tocopherols and omega-3 fatty acids [134]. A multifunctional α-enolase protein was isolated from Lactobacillus plantarum ATCC 8014 cells, which possesses the ability to produce 9cis-11-trans-CLA from LA upon combination with the membrane-associated protein fraction [24]. Linoleate isomerase (LAI) protein isolated from Lactobacillus acidophilus L1 and Lactobacillus reuteri (ATCC 23272 and ATCC 55739) strains catalyzed conversion of CLA from LA, indicating the potential use of LAB LAI in the production of CLA through enzymatic isomerization [135]. CLA from plant oils has been successfully synthesized using a synergistic biocatalytic system based on immobilized Rhizopus oryzae lipase and Propionibacterium acnes isomerase [30]. The method resulted in a very high production of trans-10, cis-12-CLA and is a promising method for largescale CLA production. Lipids in food spontaneously react with atmospheric oxygen (autooxidation) resulting in degradation of technological and functional property of food [136]. Lipases act as catalysts in production of antioxidant fatty acid esters that reduce oxidative damage, thus maintaining the structural and functional property of food [121]. Lipases catalyze the formation of transference of ester bonds and thus can be used for acylation of antioxidants [137]. L-Ascorbic acid esters are used as autooxidation retarding agents for stabilization of food products rich in oil and are synthesized using lipases as catalysts and vinyl, alkyl esters, and saturated and unsaturated free fatty acids as acyl donors [121]. Novozyme 435, a commercially available lipase acrylic resin from C. antarctica was used with stearic acid methyl ester as substrate to produce 6-O-L-ascorbyl-stearate with 99.8% purity [138]. Similarly, synthesis of ascorbyl palmitate was catalyzed by Novozyme 435 with tert-amyl

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alcohol, proving to be the best solvent for this reaction [139]. Lipase B from C. antarctica proved to be the best catalyst among 15 lipases, esterases, and proteases for the enzymatic acetylation of Vitamin E (tocopherol ester), which is used as an efficient alternative to the chemically produced tocopherol [140]. Natural phenolic compounds protect food-based products from oxidation and increase shelf life of lipid-containing products having nutritional property [141]. Use of natural phenolics is restricted due to low solubility of these compounds in hydrophobic media, resulting in poor absorption and low concentration in circulatory system [142]. Lipophilization of natural phenolics addresses this problem, resulting in the formation of lipophilic antioxidants [143].

15.2.5 TANNASE A wide variety of foods in our diet include phenolic compounds but the functional effects of these compounds are not expressed to the full potential in our health due to the complex formation of phenolic compounds with cellulose matrices [144] present in glycosidic, esteric, or polymeric forms that have low absorption rate [145]. Tannin acyl hydrolase (Tannase) is an extracellular enzyme produced by bacteria, yeast, and fungi in the presence of tannic acid and has been known to hydrolyze the ester bond (galloyl ester of an alcohol component) and the depside bond (galloyl ester of gallic acid) of tannins [146]. The action of tannase on hydrolysis of complex polyphenols has been identified and the ability of tannase to cleave cross-links between cell-wall polymers is expected to release simpler phenolic compounds resulting in increased absorption of polyphenols in the small intestine [10]. Many studies have reported increased polyphenol content in food residues that were subjected to treatment with tannase. Treatment of grape pomace with tannase resulted in increased content of total phenols along with enhanced antioxidant activity, aglycones, and phenolic acids such as gallic acid, and monomeric polyphenols [147,148]. Biotransformation of soy isoflavones daidzein and genistin by tannase has been reported recently along with an increase in antioxidant activity and total phenolic content of soy milk. However, the most important result of enzymatic biotransformation of soy milk by tannase was the production of the nutraceutically superior equol from soy isoflavones, which was observed for the first time [149]. Treatment of green tea with tannase caused reduction in gallocatechin gallate, epicatechin gallate, and epigallocatechin gallate, decreased toxicity along with reduction in adipocyte lipid accumulation and increased antioxidant activity, immunomodulatory activity, and gallic acid content. In addition, tannase treatment resulted in inhibition of the potential carcinogen N-nitrosodimethylamine formation [150
154]. Increase in bioactive phenolic composition of other phenol-rich beverages upon tannase treatment has also been reported. Tannase treatment of orange juice resulted in the generation of a higher functional product with increased antiproliferative and antioxidant activities on human cells due to transformation of glycosides in aglycones (hesperidin and naringin) [121,155]. In another study, oil extraction from oleic matrices was enhanced upon tannase treatment, resulting in an increase in phenolic compound extraction and antioxidant activity [156].

15.2.6 PHYTASE Phytases are generally produced by microorganisms that inhabit plants and have been known for sequential dephosphorylation of phytic acids generating products with high solubility, lower chelating capacity, and a decreased effect of inhibition on mineral absorption [157]. Phytic acids in their

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salt form express antinutritional effects in food products by forming complexes with essential minerals (K, Fe, Ca, Mg, Zn, and Mn) and interacting with proteins in food and during digestion [10]. Phytase has great potential in development of functional foods, importantly for the reduction of mineral deficiency risk in susceptible people [158]. Several studies have reported the use of phytases for phytate dephosphorylation during food processing, thus leading to the development of low-phytate foods with increased nutritional and functional profile beneficial for human health [159]. Reduction of phytate accompanied by reduced oxalate in food due to phytase treatment resulted in reduction in kidney stone development [160]. Application of phytase during production of whole grain bread showed phytate reduction accompanied by increased fiber and mineral bioaccessibility, resulting in enhanced nutritional value of bread without affecting the bread quality [161
164]. Consumption of supplemental dietary fiber has been reported to increase bone mineral density and Zn status in rats fed a low-Zn diet [165]. The bioavailability of Mg, P, and Ca was enhanced upon consumption of phytase-treated lupin [13]. Similarly, bioavailability of Zn and Fe was enhanced upon consumption of phytase-treated rice, sorghum, and wheat seed [166,167]. In vivo studies based on absorption of Fe by rats fed with phytase-treated pea flour protein and faba bean flour confirmed increased Fe absorption and digestive utilization correlating the higher levels of Fe in the sternum of rats [168]. In another study, rats were fed sorghum treated with phytase and tannase, which resulted in increased phosphorous digestibility, lower excretion of phosphorous, and higher biochemical indices for cholesterol and glucose was observed [169]. As no phytase-treated food product has been developed yet for human application, commercially available microbial phytases can be used for controlled and balanced degradation of phytate resulting in development of foods with known content of myo-inositol phosphate esters exerting lower antinutritional effects, increased antioxidant activity, and enhanced health benefits [10].

15.3 PRODUCTION OF FUNCTIONAL FOODS USING SPECIFIC ENZYMEPRODUCING STARTERS In recent years, fermentation using specific enzyme-producing microorganisms has been applied in food bioprocess for specific transformation for bioactive compounds and development of functional foods [2,4,170]. Isomerase and reductase enzymes of LAB have been observed to increase CLA content in dairy products during microbial fermentation [170]. The linoleate isomerase complex of L. plantarum catalyzes the conversion of 10-hydroxy-12-cis-octadecenoic acid (10-HOE), the hydration product of LA to produce monoenoic acids and CLA isomers [171]. Penicillium spp. derived β-glucosidases have been used in the production of gentiooligosaccharides, which are used in the functional food industry for their bifidogenic activity [172]. β-Glucosidase from bacterial strains involved in fermented food production has shown excellent potential in use for enzymatic enhancement of functional properties in food industry [2,173]. β-Glucosidase along with α-amylase and protease was produced using Bacillus spp. during fermentation of kinema, resulting in the generation of soybean bioactive hydrolysate rich in polyphenols [2]. Similarly, β-glucosidase produced by stress-induced L. acidophilus BCRC 10695 increased the aglycone fraction in total isoflavones in fermented soy milk from 21.8% initially to 97.9% in 24 h [173]. Fermentation of citrus

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residues with tannase producing fungi, Paecilomyces variotii increased the content of phenolic aglycones such as naringenin, ellagic acid, and hesperetin, resulting in enhanced antilipogenic effects on adipocytes and antioxidant activity [155,174]. Similar fermentation of castor bean residues resulted in an increase in gallic acid content and a reduction of cytotoxicity in macrophages and reduced levels of the toxic compound ricin [155]. Yeasts play an important role in the production of β-glucosidase and subsequent transformation of bound polyphenol to their aglycone forms, improving the bioactive profile of fermented foods and beverages [1,5]. A quantitative and qualitative screening of β-glucosidase activity from Saccharomyces strains isolated from must was reported recently [175]. Other yeasts, including those belonging to the genus Pichia have been observed to secrete β-glucosidase [176]. β-Glucosidase produced from Dekkera bruxellensis was used for enzymatic cleavage of sugar moiety of piceid (Polygonum cuspidatum) to release resveratrol [12]. Exploring specific enzymeproducing microorganisms can have several applications in functional food industry.

15.4 GENETICALLY MODIFIED ENZYMES IN DEVELOPMENT OF FUNCTIONAL FOODS The food enzyme market consists mainly of natural food enzymes for food processing and functional food production. However, these enzymes do not satisfy the requirements of the sophisticated and precise food processing conditions that require enzymes to function under extreme environmental conditions such as pH, temperatures, salinity, and pressure and deliver higher yields. Genetically modified enzymes are designed to improve the enzymological properties and enhance characteristics such as specificity, catalytic efficiency, purity, multifunctionality, surface property, and yield. These changes are attributed to altered amino acid sequence due to manipulation of genetic sequence of the protein. Application of recombinant enzymes in development of functional foods is met with several challenges and safety concerns, yet the potential benefits of its utilization in the food industry, the environment, and consumers have encouraged several studies dedicated to recombinant enzymes and related processes. Escherichia coli and Pichia pastoris are the most common microbial model systems for the expression of genetically modified enzymes [177]. Carbohydrases are the most widely-used enzymes in the food industry and genetic modifications of these enzymes have demonstrated increased functional food yield and efficacy. Novel α-amylase was obtained from B. licheniformis by replacing His residues in the active site domain with Arg and Asp residues that resulted in increased activity at acidic pH (pH 4.5) as compared to the wildtype enzyme which led to enhanced starch liquefaction, saccharification, and fermentation [177]. Enhancement of substrate specificity of Thermotoga maritima β-glucosidases for quercetin glucosides by site-directed mutagenesis led to increased aglycone production [178]. Cellobiose hydrolytic efficiency of Aspergillus aculeatus β-glucosidase was enhanced by site-saturation mutagenesis resulting in accelerated saccharification capability of the enzyme [179]. A variant of endo-β-1,4xylanase generated by site-directed mutagenesis demonstrated high catalytic activity for cleavage of β-1,4 glycosidic linkages in xylans and this enzyme was used in the production of XOS from wheat straw under alkaline and thermal conditions [180]. Site-saturation mutagenesis of His 286 was used for the production of modified α-amylase from Rhi. oryzae, resulting in an increase in the

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optimum temperature of the enzyme to 60 C and a lower optimum pH of 4.5 as compared to the wild-type enzyme [181]. Substitution of the hydrophobic Lys 48 residue with Ala and Leu in β-1,31,4-glucanase led to enhanced thermostability and halostability of the enzyme along with improved catalytic efficiency for hydrolysis of β-1,3-1,4-glucan of oats, barley, avena, and rice [182]. The ability to breakdown sucrose into glucose and fructose was increased by engineering the production of a novel Saccharomyces cerevisiae invertase with optimal pH and stability ranges via substitution of hydrophilic residues in the peripheral loops or active site region with hydrophobic amino acids [183]. Similar modifications were engineered for the enhancement of transfructosylating activity of another S. cerevisiae invertase, which was used for efficient production of prebiotic FOS [184]. For the enhanced production of prebiotic GOS, the transglycosylating activity of modified thermoresistant T. maritima β-galactosidase was improved by rational designing [185]. Directed evolution was used for the production of a β-galactosidase variant that showed high activity during milk processing in industrial type conditions at 8 C [186]. Similar modifications are performed for the production of commercial β-galactosidase and lactases demonstrating activities at low temperatures for applications in lactose-free milk product processing [177]. Lipases have shown activities over wide temperature ranges and pH with some lipases possessing fatty acid specificity [177]. Lipase from Malassezia globosa was modified for improvement of thermostability, increasing its specific activity on mono- and diacylglycerol and making it a potent biocatalyst for diacylglycerol synthesis in edible oils with enhanced health benefits [187]. Thermostable T1 lipases have been engineered for the production of SCFAs in milk, resulting in increased bioactivity and enhanced flavor in dairy products [188]. Plant and animal-derived food proteases unlike lipases and carbohydrases are not usually genetically modified because these enzymes are adequately compatible with the environmental factors of industrial food processing [177]. Other reasons include the wide availability of microbial proteases for industrial use. However, studies have reported the increased catalytic efficacy of microbial proteases for function food development. Increased efficiency in production of antihypertensive peptides, processing of food oil, and debittering activity was observed by truncated neutral protease from Asp. oryzae with optimum temperature of 55 C and a pH of 8 [189]. A serine protease, subtilisin nattokinase from Bacillus subtilis var. natto was subjected to modification by site-directed mutagenesis, resulting in higher specific activity, oxidative stability, and strong fibrinolytic activity [190]. Similarly, the glycine-releasing activity of an acid protease was improved and produced from a mutant Asp. oryzae strain using potato pulp powder fermentation [191]. A cellobiose 2-epimerases was engineered from Caldicellulosiruptor saccharolyticus that showed heightened specific activity to convert lactose to lactulose directly without the coproduction of epilactose [192]. This enzyme has shown potential use in the commercial production of lactulose that can be used as an additive in medicines and prebiotic food. Site-directed mutagenesis was used to increase the substrate-binding affinity, thermostability, and catalytic efficiency of D-psicose 3epimerase in the isomerization of D-fructose to D-psicose, which is an ultralow-calorie sweetener with potentially desirable physiological properties [193]. Genetic modification of phytases from Yersinia via site-directed mutagenesis resulted in improvement of thermostability of the enzyme along with trypsin and pepsin resistance. This enabled enhanced ability and potential industrial use of the enzyme for efficient phytic acid hydrolysis into inorganic phosphate for improved nutrient uptake from foods [194]. Rational designing was used to engineer a novel di-D-fructose dianhydride I (DFA I)-forming Streptomyces davawensis inulin fructotransferase with enhanced catalytic

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activity and thermostability for the production of prebiotic DFA I using inulin as substrate via industrial production processes [195].

15.5 CONCLUSIONS AND PERSPECTIVES The current chapter explains the importance of enzyme bioprocesses in the production of various bioactive compounds and development of functional foods. The roles of enzymes from different sources with varying catalyzing abilities and environmental parameter requirements have been highlighted. However, there is a need for development of economical and efficient enzymatic bioprocesses for the production of some of the nutraceuticals that can replace the existing chemical and physical processes. Exogenous enzymes are feasible catalysts for the production of functional food ingredients. Identification of novel enzymes with superior catalytic properties is a priority of recent research. The cost of isolation, purification, and concentration of target bioactive compounds produced from enzymatic bioprocessing is higher than chemical production of the compounds. Research attention toward reducing the cost of eco-friendly enzymatic bioprocess is needed. Extensive research needs to be conducted to study the in vivo stability and absorption during human digestion of bioactive compounds produced from enzymatic processes. In addition, studies need to be conducted to validate the safety and in vivo bioactivity of enzyme-catalyzed functional foods. Enzyme-treated functional foods can have enhanced bioavailability in the gastrointestinal tract, which can directly influence the bioefficacy. The ability of extremozymes to catalyze reactions under extreme environmental conditions can be exploited for efficient and cheaper industrial production of functional foods. Development of novel techniques for enzyme immobilization can provide additional advantage of repetitive use, easy separation, and stability of enzymes from substrates in reaction mixtures, thus increasing utility of enzymes in industrial applications. Genetically modified enzymes find better industrial applications than wild-type enzymes. Diverse genetic modifications and engineering of novel highly catalytic enzymes are required in the functional food industry. However, studies based on the safe use of genetically modified enzymes for the production of functional food ingredients need to be conducted.

ACKNOWLEDGMENTS The authors of the manuscript thank the Institute of Bioresources and Sustainable Development (IBSD), a National Institute of Department of Biotechnology, Govt. of India for all the support, encouragement, and providing necessary help to undertake the study.

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16

Haiquan Yang1, Fengyu Qin2, Zilong Wang2, Xianzhong Chen1 and Guocheng Du3 1

Key Laboratory of Carbohydrate Chemistry & Biotechnology, Ministry of Education, School of Biotechnology, Jiangnan University, Wuxi, P.R. China 2Department of Chemical and Biomolecular Engineering, National University of Singapore, Singapore, Singapore 3School of Biotechnology, Jiangnan University, The Key Laboratory of Carbohydrate Chemistry and Biotechnology, Wuxi, P.R. China

16.1 INTRODUCTION Cell-free enzymes and whole cells are the main types of biocatalysts used in the biotransformation of chemicals. Among crude and purified cell-free enzymes, crude enzymes are particularly important for biotransformations because the purification of enzymes raises the cost of biomanufacturing [1]. Additionally, high-purity enzymes are not necessary for cell-free chemical biosynthesis [2]. Cell-free chemical biosynthesis, especially multienzyme cascades, has several advantages over whole cells, including diverse reaction conditions, direct control of the reaction environment, and flexible cascade assembly [2]. Of course, whole cells are also important for chemical synthesis. Due to the complexity of the structures and synthetic pathways of chemicals, cell-free enzyme systems can struggle to synthesize certain types of chemicals, whereas whole cells can often produce such chemicals efficiently following metabolic and fermentation engineering. Pharmaceuticals, food, and biofuel industries account for most biotransformations. The pharmaceutical industry is driven by an increasing demand for novel molecules with higher therapeutic effectiveness and safety. Additionally, reliable manufacturing processes that deliver medicines on a large scale are of equal importance. More and more complex drug molecules are being designed and developed, and many are synthesized in multiple steps [3]. The key challenge for such synthetic pathways is often the synthesis of key intermediates, especially the introduction of stereocenters [3]. Conventional synthetic chemistry has greatly accelerated the development of the pharmaceutical industry through the development of synthetic methods [4]. Biotransformations also play an important role in food processing, especially the hydrolysis of large natural products such as proteins to smaller functional peptides and other molecules, and the biotransformation of one chemical (e.g., sucrose) into another (e.g., functional oligosaccharides). Meanwhile, the demand and production of biofuels are ever-increasing worldwide. The utilization of biofuels is attractive, not only due to the various advantages including renewability, high energy, CO2 mitigation, and eco-friendliness, but also because they can be produced from biomass [5]. Biofuels are produced by renewable biomaterials rich in starch, cellulose, hemicellulose, and Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00016-8 © 2020 Elsevier B.V. All rights reserved.

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lignins, such as corn, cassava, lignocellulosic biomass, coconut, rice bran, linseed, algae, soybean, jatropha, palm, jojoba, castor, and many others. For instance, the majority of ethanol in the United States is produced by biotransforming corn as a feedstock, while Brazil mainly uses sugar to biotransform ethanol [6]. In this chapter, we discuss advanced biotransformations that use crude enzymes and whole cells in the pharmaceutical, food, and biofuel industries.

16.2 BIOTRANSFORMATIONS IN THE PHARMACEUTICAL INDUSTRY Recent advances in biocatalysis and leveraging of biocatalysts in drug development are leading to new breakthroughs in the pharmaceutical industry. Tremendous advances in DNA sequencing, DNA synthesis, metagenomics, bioinformatics, protein engineering, and biosynthesis enable the discovery and utilization of new enzyme activities and provide an ever-expanding toolbox of biocatalysts [7
10]. Many characteristics of biocatalysts meet the demands of pharmaceutical research and development, including exquisite selectivity for the synthesis of chiral medicines, and the ability to catalyze reactions under mild conditions [3,7,10]. A number of elegant reviews have been published on biocatalytic organic synthesis [11
14]. Herein, we provide some selected examples of biocatalytic chiral synthesis and biosynthesis of active pharmaceutical ingredients (APIs).

16.2.1 CHIRAL CARBOXYLIC ACIDS Independent of cofactors, enzymatic hydrolysis is a low-cost biocatalytic technology that can provide chiral APIs [15,16]. In addition, strict stereoselectivity, broad substrate scope, and stability under organic synthesis conditions enable enzymes to be introduced during the initial stages of the biocatalytic preparation of pharmaceutical chemicals [15,16]. (S)-Pregabalin, a lipophilic γ-aminobutyric acid analog, was developed by Pfizer for the treatment of epilepsy, neuropathic pain, fibromyalgia, and generalized anxiety disorder [17]. In the initial development of Pregabalin, several chemical synthesis routes of high yield and low cost were established, but late-stage resolution was needed for enantiopure synthesis of the active molecule [18]. In another chemical route, (S)-3-cyano-5-methyl hexanoate, the key intermediate for S-Pregabalin, was provided via asymmetric hydrogenation. However, the noble metal, Rhodium, was used as catalyst in this process, which increased the cost of purification and applied environmental pressure [19]. To avert these disadvantages, a new synthesis route was developed using lipolase, a lipase from Thermomyces lanuginosus, as biocatalyst in the key step. Inexpensive racemic ester 1 was used as the starting material to provide (S)-1, the intermediate for S-Pregabalin [20]. (S)-1 was hydrolyzed to produce (S)-2 with high enantioselectivity (E value .200), and R-1 was recycled by racemization in the initial stage of the route (Fig. 16.1). Biocatalytic conditions were subsequently optimized by adding calcium acetate in the reaction medium to repress product inhibition, and a high conversion (48%) was accomplished at high substrate concentration (765 g/L) after 24 h [20]. Another famous enzymatic hydrolysis is catalyzed by Candida antarctica lipase B (CALB), a lipase from C. antarctica. CALB was developed to prepare APIs for the production of Ramatroban [21], a thrombin receptor antagonist developed by Bayer, (S,S)-reboxetine [22], an antidepressant marketed by Pfizer, and Odanacatib [23], an inhibitor of cathepsin K developed by Merck & Co. Using protein engineering, researchers tuned the enantioselectivity of CALB to expand its potential

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337

FIGURE 16.1 Preparation of (S)-3-cyano-5-methyl hexanoate by biocatalytic hydrolysis.

FIGURE 16.2 Kinetic resolution of profen esters by engineered CALB.

in the pharmaceutical industry [24
26]. Profens such as ibuprofen, flurbiprofen, and ketoprofen are very important nonsteroidal antiinflammatory drugs that are frequently used as analgesics, antipyretics, and antiinflammatory agents. Using cell-free extracts of engineered CALB as a biocatalyst and racemic profen esters as the starting material, (S)-enantiomers of profens, more effective than (R)-enantiomers, could be produced at up to 99% ee in the case of ketoprofen (Fig. 16.2) [24
26].

16.2.2 CHIRAL ALCOHOLS Ketoreductases (KREDs, carbonyl reductases, alcohol dehydrogenases) could use NAD(P)H as a cofactor to reduce aldehydes and ketones to alcohols [27
30]. To minimize loading of the expensive cofactor, high-efficiency cofactor recycling systems have been constructed, and glucose, formate, and 2-propanol can serve as a terminal reductant. The reduction of prochiral carbonyl groups into chiral alcohols is an example of great atom economy, and chiral alcohols are widely used in the pharmaceutical industry as critical building blocks [31
33]. Biotransformation makes the preparation of such chiral molecules more environmentally friendly [31
33]. (S)-Licarbazepine is a voltage-gated sodium channel inhibitor used in the treatment of epileptic seizures and commercialized as an acetate ester prodrug [34]. A whole-cell-mediated asymmetric

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reduction of oxcarbazepine 6 to S-licarbazepine 7 has been developed. In this biocatalytic system, the expensive cofactor can be recycled using glucose as the cosubstrate (Fig. 16.3) [35]. To improve the application potential of the biocatalytic reduction of oxcarbazepine, a KRED from Lactobacillus kefi has been engineered by Codexis, and the biocatalytic process using this enzyme can tolerate high concentrations of substrate (100 g/L) and achieve a high isolate yield (96%) [36]. KREDs are readily engineered for higher activity and enlarged substrate scope [37]. A ketoreductase from Candida glabrata (CgKR1) has been engineered by tuning two rational residues to generate a combinatorial variant (CgKR1-F92C/F94W) that exhibits higher activity toward 28 structurally diverse substrates than the wild-type enzyme [38], with specific activity over 50 U/mg for 13 substrates. Moreover, five substrates at high loading ( . 100 g/L) were reduced completely at gram-scale quantities in preparative reactions, and chiral building blocks for Imbruvica (commercialized as a lymphoma treatment) could be produced in high space time yield (589 g/L/day) using whole cells harboring the enzyme variant as the biocatalyst and a cell-free extract of glucose dehydrogenase as the coenzyme (Fig. 16.4) [38]. Regarding stereoselectivity, Prelog’s rule should be generally followed when prochiral carbonyl compounds are asymmetrically reduced by most KREDs, and KREDs with antiPrelog stereoselectivity are relatively rare in nature [39,40]. As both Prelog and antiPrelog alcohols are used as important building blocks in the pharmaceutical industry [31], switching the selectivity of KREDs from Prelog to antiPrelog has been exploited via structure-guided rational designs. Additionally, using a short-chain dehydrogenase/reductase from Lactobacillus fermentum (LfSDR1) as the

FIGURE 16.3 Asymmetric reduction of oxcarbazepine.

FIGURE 16.4 Biosynthesis of the Imbruvica intermediate.

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339

template, a switch for tuning the stereoselectivity of SDRs between Prelog and antiprelog using halogen-substituted acetophenones was recently discovered. Knowledge obtained from Lf SDR1 could be further transferred to other SDRs to tune their enantioselectivity between Prelog and antiPrelog (Fig. 16.5) [41]. Using a cell-free extract containing engineered enantio-complementary SDRs as biocatalysts, both enantiomers of some valuable chiral alcohols for the synthesis of pharmaceutical agents became accessible. More and more KREDs are becoming accessible for the biocatalytic reduction of carbonyl groups, facilitating the use of chiral alcohols at large scale in commercial processes. In addition to focusing on the mining of new KREDs, attention is being paid to widening the substrate scope, improving enantioselectivity and activity, and applying KREDs as biocatalysts for generating two contiguous chiral centers via enzymatic dynamic reductive kinetic resolution (DYRKR) [42]. In the process of DYRKR catalyzed by KREDs, one enantiomer of a racemic substrate is reduced rapidly and the other enantiomer undergoes racemization (Fig. 16.6). Consequently, the carbonyl group is reduced asymmetrically, and DKR takes place on the adjacent chiral center at the same time [42]. For multistereocenter medicines, DYRKR provides a very useful synthetic approach. A variety of β-keto esters and other β-keto chemicals have been used as starting reactants to develop high-efficiency drug synthesis routes [42]. A DYRKR approach affording (2S,3R)-methyl-2benzamido-m-ethyl-3-hydroxybutyrate, a key building block for carbapenems, has been exploited

FIGURE 16.5 Asymmetric reduction of acetophenones using enantio-complementary short-chain dehydrogenase/reductase (SDRs).

FIGURE 16.6 An example of the DYRKR process.

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FIGURE 16.7 Biosynthesis of intermediates of carbapenems (A) and Vibegron (B) via the DYRKR process.

using alcohol dehydrogenase from Burkholderia gladioli as the biocatalyst [43]. Furthermore, Merck & Co. recently developed a highly efficient DYRKR route for Vibegron, a potent and selective β3-adrenergic receptor agonist, which has been implemented at the manufacturing scale (Fig. 16.7) [44].

16.2.3 CHIRAL AMINES AND AMINO ACIDS It is estimated that 40% of existing pharmaceuticals contain a chiral amine component in their structure [45]. Thus the development of efficient approaches to synthesize chiral amines is of great

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value in the pharmaceutical industry [46]. As a potential green alternative to metal-catalyzed reactions, biocatalysts have been introduced in this field. Using pyridoxal-50 -phosphate as a cofactor, transaminase transfers an amine group from an amine donor to a ketone or aldehyde [47]. Both (R)- and (S)-selective transaminases have been discovered and engineered to provide chiral amines for APIs or intermediates of APIs [48,49]. The best biocatalytic route to date was developed by Merck and Codexis in 2010, using engineered transaminase ATA117 Rd11 in the direct asymmetric synthesis of the antidiabetic drug, Sitagliptin (Fig. 16.8) [50]. Recently, amine dehydrogenases that catalyze the reductive amination of prochiral ketones using ammonium as the amine donor have been engineered from amino acid dehydrogenases discovered via gene mining [51,52]. Both transaminases and amine dehydrogenases prefer to produce primary amines. To obtain secondary and tertiary amines, approaches using imine reductase (IRED) as the biocatalyst have been developed [53,54]. Furthermore, a reductive aminase from Aspergillus oryzae (AspRedAm), an IRED homolog, has been discovered and used at a preparative scale for the synthesis of (R)-Rasagiline (Fig. 16.9), a drug used to treat symptoms of Parkinson’s disease [53]. In addition, chiral amino acids are extensively applied as significant building blocks for APIs. As summarized in a recent elegant review, enzymatic asymmetric synthesis is a unique method of producing chiral amino acids [55]. A whole-cell catalyst, recombinant Escherichia coli expressing a leucine dehydrogenase from Bacillus cereus (BcLeuDH) and a formate dehydrogenase from Candida boidinii (CbFDH) to regenerate the cofactor NADH, has been

FIGURE 16.8 Biosynthesis of Sitagliptin.

FIGURE 16.9 Biosynthesis of (R)-Rasagiline via reductive amination.

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FIGURE 16.10 Biosynthesis of the intermediate of boceprevir, telaprevir, and atazanavir.

developed to asymmetrically aminate 2-ketocarboxylic acid, affording L-tert-leucine, a valuable precursor for a variety of active pharmaceuticals such as boceprevir, telaprevir, and atazanavir (Fig. 16.10). In this process, ammonium formate acts as both amine donor and terminal reductant [56]. In addition to α-amino acids, biocatalysts have also been exploited to asymmetrically synthesize other nonnatural amino acids. Recently, Δ1-pyrroline-5-carboxylate reductases, ketimine reductases, and N-methylamino acid dehydrogenases have been used to asymmetrically synthesize N-functional amino acids by GSK [57]. Additionally, such reactions were performed to produce N-alkyl-functionalized chemicals (optically pure) at gram scale [57]. Using α,β-unsaturated carboxylic acids, aspartase from Bacillus sp. YM55-1 was redesigned to synthesize β-amino acids at large scale (up to 300 g/L) [58].

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FIGURE 16.11 Metabolic route for the biosynthesis of parthenolide. GAS, (1)-germacrene A synthase; GAO, germacrene A oxidase.

16.2.4 BIOSYNTHESIS OF NATURAL PRODUCTS Natural products and their derivatives are important sources for drug discovery [59
61]. Cost-efficient production has hindered the development of new drugs that depend on natural products and their analogs [62
64]. Isolation from officinal plants applies great environmental pressure [62
64]. The complex structures and multiple stereocentres of natural products make total synthesis very complicated using standard synthetic chemistry approaches [64]. However, under ambient conditions, cells are perfect for chemical transformations since they provide complex chemicals using renewable feedstocks. Through heterologous expression of the biosynthetic pathway from Tanacetum parthenium, the promising anticancer drug, parthenolide, was produced by yeast (Fig. 16.11) [65]. By contrast, 18 steps were required for chemical synthesis, achieving only a 2.3% yield [66]. Biosynthesis has proved to be revolutionary for producing chemicals. Advances in metabolic engineering make it possible to increase material and energy flux toward precursors of highvalue products [62
64]. Using simple sugars as a starting feedstock, yeast and E. coli cells have been engineered as microfactories to produce medicinal natural products and intermediates (Fig. 16.12), exemplified by tetrahydrocannabinol (THC) [67], cannabidiol (CBD) [67], taxadiene [68], artemisinic acid [69], noscapine [70], thebaine [71], and codeinone [71]. In this process, enzymes from different organisms are recruited as biocatalysts to catalyze the necessary reactions in vivo.

16.3 BIOTRANSFORMATIONS IN FOOD PROCESSING For food processing, most biotransformations involve the hydrolysis of naturally occurring large molecules into small functional molecules and biotransformation of cheap chemicals into more important and useful chemicals (Fig. 16.13). For the hydrolysis of naturally occurring large molecules into smaller molecules, trypsin and amylase are the most widely applied enzymes. Meanwhile, for

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FIGURE 16.12 Biosynthesis of plant-derived natural products using simple sugars as feedstock. GPP, Geranyl pyrophosphate; IPP, isopentenyl pyrophosphate; DMAPP, dimethylallyl pyrophosphate; GGPP, geranylgeranyl pyrophosphate; FPP, farnesyl pyrophosphate; Tyr, tyrosine; GBCA, cannabigerolic acid; CBD, cannabidiol; THC, tetrahydrocannabinol.

FIGURE 16.13 Biotransformation in food industry.

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biotransforming one chemical into another, glucose isomerase and fructosyltransferase are most widely utilized. In synthetic biology related to food chemicals, whole cells are most important for biotransformations.

16.3.1 HYDROLYSIS OF NATURALLY OCCURRING LARGE MOLECULES INTO SMALLER MOLECULES 16.3.1.1 Trypsin Trypsin (EC 3.4.21.4) is widely used in food, pharmaceutical, and other industries [72,73]. Compared with proteins, peptides possess many advantages, including high functionality and reduced allergenicity [74,75]. During food processing, trypsin can be used to biotransform proteins into functional foods. Industrial production of trypsin has mainly focused on extraction from vertebrate tissues, as exemplified by bovine pancreatic trypsin [72,76,77]. Microbial production of trypsin is also important, and microbial hosts include E. coli, Streptomyces griseus, Pichia pastoris, Bacillus subtilis, and many others [72,78
80]. For trypsin-mediated biotransformation of proteins into functional peptides, purified and crude enzymes are used, but not as widely as whole cells because trypsin used in commercial production is mainly obtained from vertebrate tissues. Crude trypsin from Thunnus alalunga has been used to convert starry triggerfish muscle into protein hydrolysates with potential for use in functional foods [76]. Although trypsin can hydrolyze proteins and thereby improve nutritional value [75,81], its wide application in the food industry is limited by costly production and purification processes [82,83]. Conventional protein digestion by trypsin normally requires a long time ( . 10 h), and during this process, hydrolyzed samples can be contaminated by trypsin and its autohydrolysis products [84]. The purification of trypsin-digested products can also be very costly and complex. For example, ammonium sulfate precipitation and chromatography steps using diethylaminoethyl sepharose, soybean trypsin inhibitor sepharose 4B, and other resins have been used for purification of trypsin from Litopenaeus vannamei hepatopancreas [85]. The immobilization of enzymes has been applied for several decades and can improve the stability, activity, and durability of enzymes [74]. Due to reuse, high enzyme concentration, short digestion period, production of enzyme-free hydrolysates, and the low degree of auto-digestion and immobilization has been applied for trypsin hydrolysis of proteins [83,84,86,87]. Cell-envelope proteinases from lactic acid bacteria as industrial enzymes are not economically viable due to poor stability and reusability. Immobilization of a cell-envelope proteinase from Lactobacillus delbrueckii and trypsin by nonwoven polyester fabrics can efficiently hydrolyze proteins, as demonstrated by bovine serum albumin and casein [88].

16.3.1.2 Amylases Starch is an important raw material in the food industry because it is inexpensive and renewable [89]. Amylases are the most important enzymes for degrading raw starch, which can be hydrolyzed by amylases into short oligosaccharide chains, maltose units and glucose [90]. In order to increase the value of starch in food processes, amylase is used to hydrolyze starch to produce products of high value, including maltooligosaccharides such as maltose [91]. α-Amylase hydrolyzes starch to

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produce high levels of maltose and is mainly expressed in fungal and bacterial sources such as Penicillium expansum and Streptomyces praecox [91]. Crude or partially purified amylases are used in the starch industry rather than high-purity amylases, and partially purified and pure α-amylases from Bacillus licheniformis exhibit no differences in hydrolysis efficiency or adsorption of raw starch [92]. In one study, crude α-amylase from Brevibacterium sp. was used to hydrolyze Tacca tuber starch to produce maltooligosaccharides in the functional food industry [93]. Additionally, the purified recombinant Corallococcus sp. amylase, CoMA, expressed in E. coli was used to investigate the conversion of maltooligosaccharides and soluble starch into maltose [91]. During baking, amylases are used to improve dough properties and baked product quantities, and amylases are used to increase bread textural properties during breadmaking [94]. Furthermore, crude commercial amylase and amyloglucosidase enzymes have been employed to improve bread properties, including loaf height and shelf life [94]. Whole cells are rarely used during the hydrolysis of starch to produce glucose and sugars such as maltose in food processes. In one study, whole cells of Saccharomyces cerevisiae strains expressing fungal amylases were used to hydrolyze raw starch, and could replace more than 90% of amylases needed for the hydrolysis of raw starch [90]. Meanwhile, Song et al. used cell-surface display technology to produce polyhydroxyalkanoates from starch by anchoring Streptococcus bovis amylase on the external surface of the cell membrane of Corynebacterium glutamicum [95]. However, whole cells expressing amylase are generally used in the production of biofuel (e.g., bioethanol) and biomaterials (e.g., polyhydroxyalkanoates) rather than food processes. In order to improve the stability, activity, durability, and hydrolysis efficiency of α-amylase in the hydrolysis of starch, immobilization has been widely applied. For example, magnetic nanoparticles were employed to immobilize α-amylase, and analyses of the stability and activity of immobilized and crude free α-amylase showed that immobilization increased starch digestion activity, and raised the stability and durability, compared with the free enzyme [96]. Meanwhile, EI-Sayed et al. immobilized amylase from Laceyella sp. DS3 to improve durability and metal tolerance [97], and Konovalova et al. used a cellulose ultrafiltration affinity membranes to immobilize α-amylase and thereby improve both the mass transfer coefficient and starch hydrolysis [98].

16.3.2 BIOTRANSFORMATION OF CHEMICALS 16.3.2.1 Glucose isomerase Glucose isomerase is an important food enzyme that catalyzes D-glucose to produce D-fructose, which could be used to generate high-fructose corn syrup. Corn starch is hydrolyzed by α-amylase and glucoamylase to corn syrup (mainly glucose), and this is acted on by glucose isomerase to produce high-fructose corn syrup (mainly fructose) [99]. High-fructose corn syrup has several important applications in the food industry, including enhancing functionality, stability, enjoyment, and taste [99]. Compared with α-amylase and glucoamylase, glucose isomerase is expensive [99]. In order to improve the utilization efficiency of the enzyme and decrease the cost of biocatalysts, immobilization (e.g., using columns) has been used to produce D-fructose from the biocatalysis of D-glucose using glucose isomerase. Kamal et al. used poly-acrylic acid (PAA) and poly-acrylic acid-co-2-acrylamido-2-methylpropanesulfonic acid (PAA-co-AMPS) polymer networks to immobilize glucose isomerase, resulting in excellent recyclability and stability [100]. Meanwhile, Jia et al.

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used immobilized whole cells expressing glucose isomerase to efficiently produce high-fructose corn syrup at high D-fructose concentrations, which reduced the cost of scale-up [101].

16.3.2.2 Prebiotics Prebiotics are functional nutrients that play an important role in human health and mainly include fructooligosaccharides (FOS), galactooligosaccharides (GOS), xylooligosaccharides (XOS). FOS, prebiotics used in functional and low-calorie food industries [102], possess many useful properties, such as enhancing Ca21 and Mg21 adsorption and inhibiting the growth of intestine pathogenic microorganisms [102]. FOS are mainly produced by biotransforming sucrose with microbial enzymes [103]. Fructosyltransferase (EC 2.4.1.9), used in industrial FOS production, converts sucrose to FOS of different molecular weights [102]. During industrial production, FOS are typically obtained from the transformation of sucrose by free or immobilized fructosyltransferases, mainly from Aspergillus species such as A. niger [103
105]. In one study, crude sucrose:sucrose 1-fructosyltransferase from Schedonorus arundinaceus was expressed in P. pastoris to efficiently produce FOS from sucrose [106], while another group used the immobilization of Rhodotorula sp. fructosyltransferase to optimize FOS synthesis, increasing the composition of nystose by 40% under optimal conditions [107]. However, there are several disadvantages for the production of FOS from transformation by fructosyltransferases, including high concentrations of by-products (e.g., fructose and glucose) and low FOS purity. Whole-cell one-pot transformation processes can also be used to produce FOS, as demonstrated with Aspergillus japonicus, engineered Yarrowia lipolytica, engineered S. cerevisiae, Aureobasidium pullulans, and Penicillium expansum [104,108,109]. FOS production using this approach has several advantages compared with normal enzyme applications, including low cost, minimal by-products, high FOS purity, and saving time and labor [104]. Zhang et al. expressed A. oryzae fructosyltransferase on Y. lipolytica cell surface to transform sucrose into FOS, and productivity was 160 g/L/h [104]. Meanwhile, Marı´n-Navarro et al. expressed engineered invertase in S. cerevisiae to produce FOS in a one-pot process and used a two-stage temperature control process to improve the yield to 200 g/L [109]. GOS, another important typical probiotic, can stimulate the growth and activity of bifidobacterial microbial communities (e.g., gut microbiota) in consumers [110,111]. GOS has been widely applied to stimulate growth of Bifidobacteria and Lactobacilli [112]. Some commercial GOS are Bimuno and Vivinal [113]. However, Liu et al. also found that GOS might inhibit microbes producing butyrate and thereby affect glucose metabolism, although it improves growth of bifidobacterial microbial communities [114]. GOS compounds are mainly produced by β-galactosidase catalyzing lactose at high concentrations in industrial production process, involving a variety of different microorganisms (including Kluyveromyces lactis, Bacillus circulans, Streptococcus thermophiles, and A. oryzae) [113]. Three kinds of β-galactosidases have been used for production of GOS, including free β-galactosidase, immobilized β-galactosidase, and whole cells expressing β-galactosidase. For hydrolysis of lactose in dairy industry, free β-galactosidase has mainly been employed [115]. Yin et al. determined and compared reaction kinetics of free β-galactosidases purified from K. lactis, B. circulans, and A. oryzae on GOS production [113]. In order to increase the stability and cyclic utilization of enzymes, β-galactosidase can be immobilized by different methods (e.g., covalent binding) [115]. Gu¨lec¸ et al. used cellulose acetate membrane modified to immobilize β-galactosidase to produce GOS, and maximum production reached 27% (w/w) [116]. Meanwhile,

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whole cells with β-galactosidase have also been used to produce GOS. Lee et al. immobilized E. coli cells with β-galactosidase inclusion bodies (active) to catalyze lactose to produce GOS, and the yield reached B32% [117]. XOS, a potential prebiotic found in milk, fruits, and honey, was found to be composed of xylose units [118]. Chemical and enzymatic methods are mainly used to produce XOS. During enzymatic XOS production, xylan is first incubated under alkali conditions, and xylanases from Thermoascus aurantiacus, Streptomyces olivaceoviridis, Sporotrichum thermophile, and Thermotoga maritime have been applied [118].

16.3.2.3 Production of food chemicals by synthetic biology Natural products from plants and other sources have been widely used in the food industry, but many cannot be efficiently produced or extracted from plants. The extraction and production of many food chemicals are limited by environmental and weather conditions. Microorganism cells offer an alternative platform for the production of natural compounds from renewable resources [119]. Genetic and metabolic engineering are used to optimize host microorganisms to efficiently produce food chemicals. For example, many isoprenoids serve as food additives in the food industry, such as lycopene and coenzyme Q10. Lycopene is a biological antioxidant that is important for human health because it inhibits the oxidation of cholesterol and lipoproteins [120,121]. Several microorganisms are used to produce lycopene based on genetic and metabolic engineering, including E. coli, Y. lipolytica, C. glutamicum, Rhodospirillum rubrum, Blakeslea trispora, P. pastoris, and Barringtonia racemosa [120,122
127]. Several carbon sources (e.g., glucose and N-acetylglucosamine) can be used for the biotransformation of lycopene by whole cells. For example, Sun et al. constructed an engineered E. coli strain by deleting several key genes, including zeaxanthin glucosyltransferase gene crtX and modulating the expression levels of key genes, such as the gene dxs encoding 1-deoxy-D-xylulose-5-phosphate synthase, resulting in whole cells that can produce 3.52 g lycopene/L during fed-batch fermentation [120]. Y. lipolytica cells were also used to efficiently transform glucose to produce lycopene via codon optimization of key genes (e.g., synthetic construct phytoene synthase gene crtB), overexpression of key genes (e.g., GGS1), and controlling growth-limiting conditions [125]. Matano et al. expressed the EII permease gene, nagE, from Corynebacterium glycinophilum, along with N-acetylglucosamine-6-phosphate deacetylase and glucosamine-6P deaminase genes, nagA and nagB, to improve the production of lycopene by transforming the chitin-derived amino sugar N-acetylglucosamine [124].

16.4 BIOTRANSFORMATIONS IN THE BIOFUELS INDUSTRY Biofuels are produced via biological processes (e.g., biotransformations) rather than geological processes such as the formation of fossil fuels. People are consuming fuels at an increasing rate during the rapid development of economy. However, fossil fuels are likely to become depleted in the coming decades because they are not renewable. Meanwhile, the large-scale exploitation and application of fossil fuels have a detrimental effect on the environment, resulting in global warming, damage to ozone layer, biosphere, geosphere, and widespread ecological destruction [128]. Biofuels are a sustainable and efficient alternative to fossil fuels, and addition of biofuels into fossil

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fuels is helpful for reducing the overall cost of fuels. Bioethanol is popular in the United States and Brazil. Biodiesel is the most commonly used biofuel in Europe. In 2016, the production of global biofuel increased by 2.6%, but the demand for biofuel is increasing at a rate of 6.5% per annum [129]. The United States and Brazil are the largest producers of bioethanol, accounting for B85% of worldwide production. During 2015, B15 billion gallons of bioethanol were produced by the United States alone [130]. Biofuels include solids, liquids, or gases, but the term generally refers only to liquid biofuels for transport. During production of biofuels, crude enzymes (e.g., cellulase and amylase) are used for hydrolyzing biomass raw material, and whole cells are used for production of biofuels via biotransforming biomass raw material hydrolyzed.

16.4.1 CLASSIFICATION AND BIOTRANSFORMATION OF BIOFUELS Well-known biofuels include bioethanol, biodiesel, and biogas. Bioethanol and biodiesel are the most widely used liquid biofuels for transport. Biofuels are classified into four generations including first, second, third, and fourth based on feedstock and production technologies. In this section, we describe several important biofuels and their production by biotransformation of other biomaterials.

16.4.1.1 First-generation biofuels First-generation biofuels are produced by transforming crops containing sugars (such as sugar beet), starch-rich crops (such as corn), plants with oil (such as soybean and canola), and animal fats. Starch, sugar, and oil from plants are processed and eventually transformed into biodiesel or ethanol via whole yeast fermentation or transesterification [131]. The application of such biofuels in food products is increasingly questioned due to concerns about competition for raw materials and land. [132].

16.4.1.2 Second-generation biofuels Lignocellulosic material was used to produce second-generation biofuels, and this is reported to overcome some of the disadvantages associated with first-generation biofuels [132]. Cellulose is abundant in nature. Indeed, cellulosic biomass is believed to be the most abundant biological material on earth [133]. Nonfood crops are cellulose-rich materials and can therefore be used as feedstocks to produce biofuels. Second-generation biofuels are made mostly from agriculture and forestry residues such as waste plant biomass, lignocellulosic nonfood crops, and other waste resources [134]. However, it may be difficult to produce these fuels because cellulosic biomass is more resistant to be broken down than starch, sugars, and oils. For instance, a series of physical, chemical, mechanical or biological pre-treatments may be required to deconstruct lignocellulosic biomass to less complex polymer molecules for the further production of liquid fuels [135]. Pre-treatment is technically difficult, although research on developing efficient and costeffective ways of carrying out the process is ongoing. Cellulose, hemicellulose, and lignin are used to produce lignocellulose. However, the cellulosic content is composed of hemicellulose, lignin, and other materials, making it difficult to degrade due to complex bonding of lignocellulosic plant structures [136]. In order to release sugars from lignocellulosic biomasses efficiently, lignin must be removed. However, delignification is costly and challenging and requires some technical improvements, especially for pilot-scale applications [137]. During the production process of

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bioethanol, saccharification of pretreated biomasses is the key step. Biomass can be treated with either acid or enzymatic methods to release sugars; however, crude enzymes are more preferred for saccharification due to the milder processing conditions and environmental friendliness. The enzymatic hydrolysis of biomass involves complex enzymatic catalysis. The production of soluble sugars hydrolyzed from cellulose is based on the coordinated activities of multiple enzymes including β-glucosidase, β-endoglucanase, and β-exoglucanase. Trichoderma reesei and Humicola insolens have been used for the simultaneous enzyme production and hydrolysis of cellulose [138]. Cellulase enzymes from Humicola and Trichoderma are highly active during alkali pretreatment process. Feruloyl esterases, mannases, xylanases, and other auxiliary enzymes can be used for the enzymatic depolymerization of lignocelluloses [139,140]. Recently, lytic polysaccharide monooxygenases belonging to a new class of nonhydrolytic enzymes have been developed that are reportedly capable of lowering the cellulase dosage and overall cost of the process [141]. Cellulose and hemicellulose are hydrolyzed to corresponding monosaccharide (pentose-rich and hexose-rich sugar syrup) that are the main substrates for production of ethanol. However, ethanol production from crop waste feedstocks requires more complex steps than typical sugars fermentation processing from food feedstocks. In order to use lignocellulosic biomass as feedstock to produce ethanol in an efficient way, several different fermentation strategies have been employed by scientists around the world (Fig. 16.14) [135,142]. The different strategies include consolidated

FIGURE 16.14 Schematic diagrams of different available methods of ethanol production by biotransformation [135,142]. SHF, Separate hydrolysis and fermentation; SSF, simultaneous saccharification and fermentation; SSCF, simultaneous saccharification and cofermentation; CBP, consolidated bioprocessing.

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bioprocessing, simultaneous saccharification and cofermentation, simultaneous saccharification and fermentation, and separate hydrolysis and fermentation strategies [135,142].

16.4.1.3 Third-generation biofuels Third-generation biofuels are related to algal biomass, recently developed as new source for biofuel synthesis in bioenergetics. Compared with traditional oil plants, algae biomass is a better source that is richer in lipid contents [143]. Much attention has been directed toward algae due to impressive carbon dioxide assimilation, high yield, and simple operational procedures. Moreover, due to cultivation in unproductive dry lands, marginal farmlands, seawater, and even wastewater, growing algae avoids competition for arable land and freshwater.

16.4.1.4 Fourth-generation biofuels Fourth-generation biofuels are similar to third-generation biofuels in terms of using nonarable land, but this approach employs genetically modified algae with improved photosynthetic efficiencies, increased light penetration, and reduced photoinhibition to enhance biofuel production [144,145]. Fourth-generation biofuels are mainly divided into two types: photobiological solar biofuels and electrofuels. Some fourth-generation biofuels are carbon-neutral, resulting into net carbon or greenhouse gas emissions. Truncated chlorophyll antennae in chloroplasts have been used to enhance light penetration in dense microalgae cultures [146], along with minimizing light absorption and pigment manipulation [147]. To improve the photosynthetic efficiency of microalgae, the absorbing spectrum range has been expanded [148]. Additionally, the lipid and carbohydrate contents can be significantly improved via metabolic engineering of microalgae [149].

16.4.2 IMPORTANT EXAMPLES OF BIOFUELS 16.4.2.1 Bioethanol Sugars and starches can be used as substrates to prepare fermentative ethanol as a commercial (liquid) biofuel. Ethanol-producing microorganisms such as S. cerevisiae [150] and Zymomonas mobilis [151], are usually used to transform sugars into bioethanol. Nonfood crop wastes are increasingly taking the place of food-based resources in biofuel fermentation [152]. This robust technology can transform glucose into ethanol, similar to conventional fermentation, the final concentration of ethanol reach 8%
16%. Meanwhile, the biotransformation of cellulose to ethanol can be divided into the following steps: (1) disintegrating biomass, (2) saccharification, (3) transformation of sugars into ethanol, and (4) purifying products. Briefly, cellulosic and hemicellulosic components of plant materials must first be broken down into sugars, which are biotransformed to produce ethanol. Nonetheless, it is of great significance to develop and engineer new microorganisms for fermentation that can directly use complex polymeric substrates [153]. Various microorganisms are used for ethanol production from lignocellulosic biomass, including Mucor circinelloides, Kluyveromyces marxianus, T. reesei, S. cerevisiae, and Scheffersomyces stipitis [154]. Although producing bioethanol from biomass has been carried out for many years, the efficient and cheap production of bioethanol by biotransformation still remains challenging. During biotransformation processing, there are major technological bottlenecks such as the availability and

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development of low-cost and substrate-specific enzymes, the design of highly efficient reactors, and efficient biotransformation strains [130].

16.4.2.2 Biodiesel Biodiesel, as an alternative biofuel, can be produced based on transesterification of monohydric alcohols and triglyceride oils. After ethanol, biodiesel should be currently the second most abundant biofuel in the world. Biodiesel is biodegradable, nontoxic, and convenient for various purposes. Biodiesel can be produced from residues of vegetable oils, such as rapeseed, palm, or soya, which reduces dependence on chemical-based fossil fuels [155,156]. Transesterification involves the reaction of an acyl donor (ester) with and acyl receptors such as glycerol or alcohol via glycerolysis or alcoholysis. Lipases (e.g., acyl ester hydrolases) can be used to hydrolyze triacylglycerols to obtain monoacylglycerides, diacylglycerides, and other substances. Under certain conditions, lipases can also catalyze esterification, transesterification, and interesterification [157]. The production of biodiesel by lipase-catalyzed transesterification of triglycerides and alcohols is an environmentally friendly and sustainable route for fuel production due to mild reaction conditions and the use of low-cost feedstocks containing water and free fatty acids, such as nonedible oils and waste oils [158]. Although using lipases as biocatalysts includes several advantages, their isolation and purification can be costly. Additionally, the structure of lipases is often unstable when isolated from their natural environment. In industrial processes, immobilized enzymes or cells may exhibit improved enzymatic properties, including activity and stability, and resistance to inhibition by reaction products. Studies on the production of biodiesel using algae are increasing. Both macro- and microalgae possess superior photosynthetic potential. As a major source of biodiesel, algae is the most abundant feedstock for biodiesel production, achieving .250-fold oil production per acre, compared with soybeans. Algae also produce much more biodiesel than palm oil, and it is widely believed that gasoline may be replaced by biodiesel automotive fuel produced by algae. Various types of microalgae have been used to produce biodiesel, including Botryococcus braunii, Chlorella species, and Scenedesmus species. B. braunii possesses oil levels of 20% 2 50% and favorable productivity, while Chlorella might also be suitable for the production of biodiesel [159].

16.4.2.3 Biohydrogen Biohydrogen is most commonly produced from organic waste materials by eukaryotes (e.g., algae) and prokaryotes (e.g., bacteria and archaea). Biohydrogen produced by algae is a clean fuel that could replace gasoline [160]. Enterobacter aerogenes is the organism most widely explored for hydrogen production. Biohydrogen and other biogases (e.g., methane) have been achieved using the anaerobic fermentation of residues by Chlorella reinhardtii biomasses [161]. The production of biogas by Chlamydomonas reinhardtii fermentation is efficient, with a yield of up to 587 6 8.8 mL biogas/g (volatile solids). Interestingly, the yield of biogas during the hydrogen production cycle can be improved by 1.23-fold, possibly because storage compounds (e.g., starch) have a high fermentative potential for production of biogas. A mixed biorefinery fermentation of Scenedesmus sp. and Chlamydomonas sp. has been reported, and the maximal yield reached 1.91 mL H2/L after 4 days [162].

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16.5 CONCLUSIONS AND PERSPECTIVES Crude enzymes and whole cells are the main forms employed in biotransformations in pharmaceutical, food, and biofuel industries. In the pharmaceutical industry, crude enzymes and whole cells have been used in biotransformations to synthesize important biomedical intermediates and final products. Biocatalysts utilize ambient conditions and can exhibit exquisite selectivity; hence, their appeal enhances sustainability in the pharmaceutical and other industries. In food processing, crude enzymes play a key role in the hydrolysis of large molecules (e.g., proteins) to produce small functional molecules (e.g., peptides) and in the synthesis of important molecules from cheap chemicals (e.g., sucrose). Meanwhile, whole cells are also important for the production of food chemicals (e.g., food additives), especially through metabolic engineering and synthetic biology approaches. Sugarcane and corn are widely used to produce bioethanol and triglycerides are used to produce biodiesel. Production of cellulosic ethanol is a major aim because it has the potential to make the energy sector of each country self-sufficient, while lowering environmental degradation. The production of biofuels has been focused on nonfood crop waste resources but food-based resources. However, in order to decrease the price of biofuels, technical and economic aspects of biofuel production should be optimized. Existing methods for reducing costs mainly include decreasing the number of reaction steps and enzymes and enhancing sustainable production of biofuels. Such approaches require crude enzymes with high catalytic efficiencies and whole cells that perform well in terms of enzyme expression levels and high conversion efficiency. Genomic DNA sequencing and recombinant DNA technologies are enabling the discovery of new biocatalysts, and enzymes can be readily engineered to adapt them to manufacturing processes. Additionally, immobilization can improve stability, activity, and durability, which applies for the biotransformation by enzymes and whole cells.

ACKNOWLEDGMENTS This study was funded by the 111 Project (No.1112-06) and Postgraduate Education Research and Practice Project of Jiangnan University (YJSJG2017004).

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CHAPTER

ENZYMES IN THE THIRD GENERATION BIOREFINERY FOR MACROALGAE BIOMASS

17

Abraham Lara1, Rosa M. Rodrı´guez-Jasso1, Araceli Loredo-Trevin˜o1, Cristo´bal N. Aguilar1, Anne ´ S. Meyer2 and Hector A. Ruiz1 1

Biorefinery Group, Food Research Department, Faculty of Chemistry Sciences, Autonomous University of Coahuila, Saltillo, Coahuila, Mexico 2Protein Chemistry and Enzyme Technology, DTU Bioengineering, Department of Biotechnology and Biomedicine, Technical University of Denmark, Lyngby, Denmark

17.1 INTRODUCTION Third generation biorefineries, based on seaweed as a feedstock, are envisaged to become an important type of processing facilities for sustainable production of chemicals, new materials, and renewable energy, in order to mitigate a dual problematic: the exhaustion of fossil fuels and the increasing levels of greenhouse gases. Both problematics represent an imminent threat to the biodiversity at global scale [1]. With this problematic looming, more governments are looking forward to a bio-based solution, capable to harness renewable energy from a variety of organic feedstocks. Macroalgae biomass has been found suitable to be transformed into biogas, bioethanol, and biobutanol, having the capacity to replace fossil fuels [2]. However there is a bottleneck for this technology to be currently massified, the low rentability because of the downstream processing [3], third generation biorefineries profits can be boosted by the production of high-value products, to ease the future transition of an oil-centered economy to a bio-based economy. Enzymes can be produced direct from macroalgal biomass to cut operative costs, also used in saccharification process to improve sugar yields, improve biofuel conversion, and obtain high-value products in order to achieve a renewable and cost-effective process [4]. This chapter will focus on some technologies developed with the goal to achieve a sustainable biofuel production and high-added value compounds, involving enzymatic interactions for the three generations of macroalgae biomass in terms of biorefinery concept.

17.2 BIOREFINERY AND BIOREFINING The concept of biorefining can be defined according to the Department of Energy in the United States as: the integrated biorefineries uses different biomasses (energy crops, agricultural residues, algae) as feedstock and bioconversion technologies to produce a range of chemicals with highadded value (or other materials), animal feed, heat, power, and biofuels [5]. Also, the definition Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00017-X © 2020 Elsevier B.V. All rights reserved.

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of the IEA Bioenergy “Task 42 Biorefining,” the biorefining in a circular bioeconomy is the sustainable biomass processing (fractionation) for the production of high added products and bioenergy [6]. A biorefinery is thus a facility that employs the biorefining concept for biomass processing to produce fuels, chemicals, and higher value products from biomass [7]. The main differentiation is the renewable nature of the biomass utilized; additionally, the biofuels produced have low emission rates in comparison with fossil fuels. Historically, three iterations of biorefineries are acknowledged in relation to the nature of the feedstocks implemented in the process, and the main differences are summarized in Fig. 17.1. Contrary to the classic fossil refinery, biorefinery is nourished by the concept of sustainability.

17.2.1 FIRST AND SECOND GENERATION BIOREFINERIES First generation biofuels have sugar, starch, vegetal and animal fat as primary source. The main quandary of this iteration is the harvestable soil competition over energy crops and crops for human and animal consumption. Although it is currently available in some developed countries, some concerns have arisen due to the direct relation between food prices and the lack of farmable lands [8]. The second generation of biofuels takes advantage of cellulosic (or lignocellulosic) feedstocks; cellulose is known to be the most abundant organic compound on the planet surface [9], and this

FIGURE 17.1 Biorefinery concept for macroalgal biomass sustainability.

17.3 MACROALGAE AS A SOURCE OF HIGH-VALUE PRODUCTS

365

iteration of biofuel production aims to use lignocellulosic biomass derived from agricultural industry wastes such as cereal straw, forest residues, bagasse, energy crops, and rotation forest [10] to produce biofuels and other commodities. However, this kind of biomass requires a pretreatment in order to hydrolyze the reducible sugars from the highly intricate polymeric matrix to make them available to fermentation. This procedure can be attained with physical, chemical, physicochemical, solvent-derived extraction, and biological treatments that in themselves consume energy and commonly used pretreatment procedures, for example, hydrothermal biomass pretreatment moreover results in the production of carbohydrate degradation products that are cellulase enzyme inhibitors. The complex lignocellulosic matrix and the pretreatment requirement are the main difficulties of this generation [11,12].

17.2.2 THIRD GENERATION BIOREFINERY This iteration is characterized for the employment of aquatic biomass as feedstock for refining, including primarily macroalgae and microalgae. As the feedstock is an aquatic biomass, there is no competition for farmable lands. Biorefinery of algal biomass is widely considered a promising alternative for actual fuels against its generational counterparts [12,13] and are sought to be a direct replacement for, gasoline, diesel, and others kind of fuels used nowadays. Macroalgae present a high content of carbohydrates, and a matrix not as intricated as the lignocellulosic biomass utilized in the previous generational iteration, mean a theorical fivefold production potential in terms of energy potential in comparison to terrestrial biomass [7].

17.3 MACROALGAE AS A SOURCE OF HIGH-VALUE PRODUCTS Even though the technological advances and the multiple advantages in productivity terms macroalgae offer, the sole production of biofuels from macroalgae it does not directly imply economic feasibility, thus the biorefinery concept of product fractionation is a must for the production of high-value products, to overcome the commercial deficit, becoming an analogous concept of the traditional petroleum refineries [14]. Several pathways have been identified to meet a satisfactory solution for this problematic. A range of high-value products are already identified, and genetic tools only expand the scope of possible commodities achievable by biorefineries. There are two perspectives in the biorefinery processes: the direct cascade principle, where higher value bioproducts are reaped first and the residues are then used for biofuels production, and in the case of the market demand biofuels over high-value products, the prioritization is reversed in what is known as an inverse-cascade principle [15]. The polysaccharides of the algae cell wall are a versatile source of active compounds across all the three types of algae. Red algae, or Rhodophyta, cell wall is mainly constituted by agar and carrageenans (CRGs) both constituted by sulfated galactans [16]; Green algae, or Chlorophyta, main cell wall has a complex matrix of mainly three heteropolysaccharides: glucuronoxylorhamnans, glucuronoxylorhamnogalactans, or xyloarabinogalactans [17]; and in brown algae, or Phaeophyceae, the cell wall is essentially composed of alginate [18], a uronide polymer composed of residues of mannuronate and guluronate. Brown macroalgae also contain laminarin as storage compound;

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laminarin is made up of a backbone of β-1,3-linked glucose moieties with β-1,6-linked branches. The glucose base of laminarin means that laminarin has a direct potential as a source for bioethanol production. Some of the macroalgae polysaccharides, that is, CRGs, agar, and alginates are already widely used in pharmaceutical and food applications, thus providing the base for a biorefinery concept where high-value substances can be produced in conjunction with bioenergy. Moreover, in brown algae, some of the residue polysaccharides, notably fucoidans (fucose-rich sulfated polysaccharides), are currently studied for their bioactive properties to fully exploit its bioactive traits: biocompatibility, biodegradability, and therapeutic activity [19], similarly providing for use of brown macroalgae in biorefining for production of a fan of products. Compared to terrestrial plant materials, the knowledge and commercial availability of enzymes for modification and saccharification of algae carbohydrates are much less. Due to their catalytic selectivity, enzymes represent a huge potential in relation to use in algal biorefining, but in order to achieve commercially competitive processes, significant research and development are required with regard to development of enzymes for algal biorefining processes—with regard to enzyme discovery, feasible enzyme production, and attainment of robust and highly active enzyme mixtures.

17.4 ENZYME PRODUCTION Furthermore, studies have indicated the emergence of new functions and the redevelopment of former lost functions, meaning a continuous process of acquiring and losing genetic material. Although it is theorized that the evolutionary pathway is determined by a conjunction of biochemical, biophysical, and regulatory factors [20]. Bacteria and fungi have evolved to break down matter in order to survive, these qualities are exploited in the biotechnology industry as a result of this factor; enzyme production is rather an intrinsic part of the nature of the biorefinery process, from the depolymerization of the base sugars of the algae through an enzymatic pretreatment to the alcohol conversion of fermentable sugars. Enzymes are always present in any kind of fermentation and bacteria and fungi are amply used in biotechnological processes, so the enzyme production does not represent a deviation of the process. Fungi task in nature is to degrade biomass, plant cell wall degrading enzymes of higher fungus can be traced back to a common ancestor gene from an early lineage of aerobic zoosporic fungi [21]. In the biofuel production, the most important enzymes are hydrolytic in nature: amylases, xylanases, and cellulases [22]. Thermoresistant mesophiles and thermophile microorganisms are mainly used in biorefinery; these microorganisms are a good enzymes producer of cellulases, hemicellulases, proteases, amylases, phosphatases, and laccases, to name some, which are produced and amply utilized in several industries such as food, pharmaceutical, and chemical industries. Another perk of utilizing fungi and bacteria resistant to heat is the thermostability of its metabolic products optimal for its industrial use and less susceptible to contamination [23]. Several microbial enzymes are utilized to modify and obtain oligomers from polysaccharide hydrolysis, with the complication of the need for the utilization of a diverse array of enzymes due to the structural complexity and diversity. The highly specific activity of enzymes is often used to depolymerize the polysaccharides to oligomers of interest or elucidate the main structure of the original molecule [19].

17.4 ENZYME PRODUCTION

367

17.4.1 SOLID-STATE FERMENTATION AND SEMI-SOLID-STATE FERMENTATION FOR ENZYME PRODUCTION Fermentation process for production of enzymes can be conducted using solid medium (solid-state fermentation, SSF) or liquid medium (submerged fermentation, SmF). SSF has been largely adopted in past generation of biorefinery as platform to obtain high-value products, and SmF is largely employed for large-scale operations [24,25], as macroalgae feedstock is more easily depolymerized than lignocellulosic feedstock, macroalgae have recently been shown to be workable as a C- and N-source in enzyme production, and both SmF and SSF are proven to be effective platforms for enzyme production [24]. A summary of common advantages and disadvantages is shown in Fig. 17.2. Several reactor designs have been developed for enzyme production, such as SSF using a rotating drum bioreactor [25], tray bioreactor, packed bed reactor, air-pressure pulsation bioreactor, and intermittent or continuously mixed SSF bioreactor. Several advantages of SSF are considered over SmF, lower reactor volume required, lower susceptibility of bacterial contamination, no foam formation, lower cost of effluents, lower cost of fermentation medium, minimal mineral concentration are needed, higher production yield for fungal based products, improved downstream process due high concentrations of the product, use of wastes as substrate is possible [26]. Although enzyme production heavily depends on the optimal conditions of the microorganisms employed and its nutrimental needs. The enzymatic production from macroalgae biomass via different microorganisms and fermentations is summarized in Table 17.1.

FIGURE 17.2 Advantages and disadvantages for fermentations strategies.

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CHAPTER 17 ENZYMES IN THE THIRD GENERATION BIOREFINERY

Table 17.1 Enzymes obtainable from macroalgae and derivate polysaccharides. Microorganism used

Enzymes produced

Technique

References

Ulva fasciata

Cladosporium sphaerospermum

Cellulases

SSF

[27]

Ulva lactuca

Bacillus sp. BT21

Xylanases

SSF

[84]

Algal waste

Cellulomonas uda (NCIM 2353)

Cellulases

SSF

[29]

Sargassum sp.

Bacillus sp.

Laminarases

SmF

[44]

Fucus vesiculosus

Aspergillus niger PSH and Mucor sp. 3P

Fucoidanases

SSF

[25]

Padina tetrastromatica

Bacillus sp. BT21

Xylanases

SSF

[84]

n/a

Escherichia coli DH5 strain for storage and E. coli BL21(DE3) strain for protein expression

Glucanase ZgLamA

Auto-induction by ZYP-5052, Autoinduction Media

[55]

Saccharina japonica

Defluviitalea phaphyphila sp.

Alginate lyases, uronic acid reductase, 2-keto3-deoxy-Dgluconate (KDG) kinases, mannitol 1-phosphate dehydrogenase, laminarase

SmF

[32]

n/a

Bacillus sp. Alg07

Alginate lyase

SmF production using purified sodium alginate.

[29]

Gracilaria verrucosa

E. coli DH5α, E. coli BL21(DE3), Pseudoalteromonas carrageenovora, Zobellia galactanivorans, Clostridium cellulovorans

Agarase, carrageenase, and neoagarobiose hydrolase

SmF with purified agarose and carrageenan

[37]

n/a

Bacillus sp. Lc50-1

Thermostable λcarrageenase, CgaL50

SSF using agar for colony selection and SmF with λ-carrageenan

[31]

Group

Algae

Green algae

Brown algae

Red algae

17.4 ENZYME PRODUCTION

369

Table 17.1 Enzymes obtainable from macroalgae and derivate polysaccharides. Continued Group

Microorganism used

Enzymes produced

Technique

References

Ahnfeltia plicata

Bacillus sp. BT21

Xylanases

SSF

[84]

n/a

Cellulosimicrobium cellulans

κ-Carrageenase

Fermentation over purified k-carrageenan

[56]

Gigartina skottsbergii

Pseudoalteromonas carrageenovora

κ-Carrageenases, ι-Carrageenases

P. carrageenovora was grown in the presence of λ-carrageenan

[32]

Gelidium amansii

E. coli with overexpressed β-agarase gen of Pseudoalteromonas sp. AG52

β-Agarase

Enzyme production induced by culture media and isopropyl-ßthiogalactopyranoside

[33]

n/a

E. coli DH5α with genes (Aga50A, Aga50D, and NABH) from Saccharophagus degradans

β-Agarases

Enzyme production induced by culture media and isopropyl-ßthiogalactopyranoside

[94]

Gelidium amansii

Raoultella ornithinolytica B6-JMP12

3,6-Anhydro-Lgalactose dehydrogenase

SmF

[84]

Algae

NABH, Neoagarobiose hydrolase; SmF, submerged fermentation; SSF, solid-state fermentation.

17.4.2 GREEN ALGAE AS SOURCE FOR ENZYME PRODUCTION Cellulose is the most abundant organic compound in the earth, a β-1,4-linked homopolymer of β-Dglucose, in which the polysaccharide chain presents a reducing end and a nonreducing end with the capacity to form an aldehyde and take the linear form. Enzymes that cleave cellulose are known as cellulases and must overcome a myriad of challenges because the β-1,4-linked polysaccharides present covalent bonding and present cellulose microfibrils in plants (and in some algae), forming lattices with few binding sites for cellulase action, and can be packed into several crystalline forms (polymorphs) that can be presented in the form of Cellulose Iβ and Iα polymorphs in several algae. The difference between Cellulose Iβ and Iα consists in the hydrogen bonding patterns and the interlayer array. Cellulose Iβ is composed of two layers: “the center” and “the origin”, where cellulose Iα is composed of a sole chain [41]. Cellulase production can be attained by SSF means, using cellulose-rich algal biomass as carbon source; green macroalgae Ulva fasciata was used as a raw carbon source for Cladosporium sphaerospermum a cellulase-producing fungus. After the SSFderived cellulase was obtained, its enzymatic activity was tested over, the same macroalgae for

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CHAPTER 17 ENZYMES IN THE THIRD GENERATION BIOREFINERY

saccharification treatment obtaining an optimized yield of 112 6 10 mg/g dry weight; after the alcoholic fermentation (with S. cerevisiae) with ethanol yield of 0.47 g/g reducing sugar and conversion efficiency was 93.81% [27]. The case in point can be taken as an example in the approach to enzyme production, taking advantage of structural polysaccharides of macroalgae as carbon source for the obtention of enzymes of interest by fermentation means. Enzyme production is also applied in lignocellulosic biomass biorefinery, but the enzyme-producing organisms have a more difficult catalytic pathway to obtain reducible sugars because the cellulose (35% 50%) is packed with hemicellulose (25% 30%) and intertwined lignin (25% 30%), forming a complex matrix [42]. Lignin has been found to hinder the enzymatic activity, because lignin binds to the cellulose hydrophobic faces, limiting the hydrolyzation of the cellulose crystalline structure and also binds preferentially to the specific residues on the catalytic site [43,44]. Lignin and hemicellulose are less prevalent in the common structure of green macroalgae; Ulva lactuca is known to be a green seaweed with a relatively high content of lignin and hemicellulose (9% cellulose, 17% hemicellulose, 14% lignin), in comparison with the usual traces contained in macroalgae [45].

17.4.2.1 Brown algae as source for enzyme production Alginate from kelp is the most abundant marine biomass, amply used in food, textile, paper, cosmetics, and pharmaceutical industries due its low cost, safety as a result of its low toxicity, thickening properties when in aqueous solution, and cross-linking in presence of divalent metal ions, Ca21, Sr21, Zn21, Pb21, Cu21, and Cd21, with the added value of being thermostable [46]. Alginate is a polysaccharide composed of C-5 epimers; β-D-mannuronic acid (M) and α-L-guluronic acid (G) linked by a 1-4 bond, arranged in block sequences of continuous M or G residues (M- or G-block), and M and G residues (MG-block), commonly arranged in a block copolymer with long homopolymeric sequences of MM-block(s) residues and GG-block(s) residues are separated by MG-block(s) [47], its properties are primarily given by the chain elongation, G residues content, and position; acetylation content prevents further epimerization [48]. The variations among alginates originate from the C-5 epimerase, which catalyzes the conversion of β-D-mannuronic acid to α-L-guluronic acid; some bacterial strains produce this enzyme, but can also be found in the brown algae Saccharina japonica in its diploid multicellular reproductive stage, known as sporophyte. Bacterial epimerase activity is widely investigated; Pseudomonas spp. and Azotobacter vinelandii are well-known bacteria that produce a biofilm mainly constituted by alginate, although Sac. japonica has a heterologous alginate biosynthetic system compared with the bacteria; brown algae lack the alginate lyase gene in the alginate synthesis encoding cluster, meaning that it is possible that some C-5 epimerases have similar hydrolytic action [49]. The only known functional alginate lyase activity in brown algae was reported recently in the algal tissue of Sac. japonica, preparing a library of complementary DNA of its sporophyte [50]. The stock market price of alginate escalates significantly due the hydrocolloid properties of alginate, and its specific application, pharmaceutical uses such as biocompatibility allow alginate to be used in cell encapsulation of mammalian cells; initially, this procedure was investigated for immunoisolation of analeptic proteins: fibroblasts, kidney cells, myoblasts, fibroblasts, and islet cells. And it was later used in regenerative medicine of tissues: dorsal root ganglia and neural progenitor cells for neural tissue, fibroblasts for dermis, chondrocytes for cartilage, mesenchymal stem cells and osteoblasts for bones, and hepatocytes for liver [51]. Mannunoran C-5 epimerases can be used to achieve a tailored alginate chains in vitro to enhance the base structure of the

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polysaccharide, improving the content of G-block (being the factor that affects directly on the gelation properties), switching M-blocks to MG-blocks, in order to customize the porosity, gel strength, and biocompatibility [47,48]. Enzyme production can be attained by fermentation means, over the algal biomass. Alginate lyases from endogenous microorganisms have been screened for alginate lyase production in some brown algae species Sac. japonica, Sargassum horneri, and Sargassum siliquastrum, from where 196 strains of bacterial producers of alginate lyase enzyme have been identified and Bacillus halosaccharovorans had the best enzymatic production titles [52], and with the correct fermentation approach, it might be an optimal process for alginate lyase production, for the obtention of alginate oligomers. Fucoidanase enzymes can be produced from fucoidan-rich biomass such as Fucus vesiculosus, through SSF using fungal strains with encoding genes for fucoidan hydrolytic enzymes, and Mucor sp. P3 had the best enzyme activity [25]. Laminariase production can also be obtained through SSF, using Sac. japonica as substrate using Talaromyces amestolkiae GT11 [53], also using raw laminarin extracted from Sargassum sp. has been proven an effective way to produce laminarase using Bacillus sp. [54]. Laminariase production has also been achieved using a known encoding gene for β-1,3-glucanase of Zobellia galactanivorans (ZgLamAGH16) and transforming Escherichia coli into E. coli DH5α strain for storage, and in E. coli, BL21(DE3) strain for protein expression, using a plasmid, induces the enzyme production using an auto-inductor medium [55].

17.4.2.2 Red algae as source for enzyme production Agar, alginates, and CRG are the main polysaccharides and the one most exploitable compound comprising the red algae, which are known to have valuable bioactive properties broadly used in industrial processes of food, pharmaceutical, cosmetic, paper, and textile as emulsifier and gelling agents [18]. CRG is one of the main components of some varieties of red algae cell wall. This polysaccharide is a gel-forming hydrocolloid composed of linear chains, comprised of D-galactose and D-anhydrogalactose residues, alternating α-1,3- and β-1,4-linkages with ester sulfates (15% 40% content). There are three main varieties of CRG: κ-CRG has one sulfate group for each disaccharide unit; ι-CRG has two and λ-CRG has three groups. Sulfate ester quantity confers different properties, λ-CRG is the most soluble of them all, in both hot and cold aqueous solutions, whereas κ-CRG is only soluble in hot aqueous solutions. CRGs are amply used in pharmaceutical investigation for drug formulation enhancements, improve drug release times, and formulate delivery systems based on pH and temperature [30]. For each type of CRG, there exists a homologous hydrolytic enzyme, κ-, ι-, and λ-carrageenase, which can be used to obtain carrageenan oligosaccharides (COS) with valuable pharmacological activities. The λ-carrageenase can be used to obtain neo-λ-carrabiose, said enzyme can be produced by a number of organisms, some thermoresistant bacteria such as Bacillus sp. Lc50-1 can be isolated from hot springs, propagated by SSF using agar for colony selection, and induce the thermostable λ-carrageenase production using SmF with λ-CRG as sole carbon source, its thermostable nature grants an advantage exploitable in industrial processes [31]. Microorganisms isolated directly from algal tissue, or decaying tissue, can be proven effective to detect potential and efficient enzyme producers; Cellulosimicrobium cellulans can be isolated from life specimens of Gelidium sesquipedale, Alsidium corallium, and Ceramium rubrum; these endogenous microorganisms use the sugars present in the algal cell wall as carbon source using hydrolytic enzymes; this characteristic can be exploited and optimized for

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κ-carrageenase production, using protocols for cell immobilization by matrix absorption using purifiedκ-CRG as carbon source, obtaining better results than a free cell medium [56]. Agar is another main polysaccharide comprising some families of red algae; Gracilariaceae, Gelidiaceae, Pterocladiaceae, and Gelidiellaceae. Its principal function is like that of hemicellulose in land plants. Agar is a wide term that englobes a mixture of D- and L-galactose polysaccharides. Agarose is a neutral polysaccharide composed of repeated D-galactose and 3,6-anhydro-L-galactose (L-AHG) units linked by β-1,3- and α-1,4-glycosidic bonds, comprising the 70% of the agar structure. The main agar chain has substituents such as sulfate ester, methoxyl group, and pyruvate ketal (e.g., alternating D- and L-galactose) and to a minor extent in agarose; the said components lessen the gelling property of agar. Agars have diverse industrial applications such as phycocolloids in food, pharmaceuticals, cosmetic, medical and biotechnology industries [35,57]. Other kinds of enzymes obtainable of red macroalgae and green algae are the enzymes in charge of the tolerance of metals, antioxidant response, Acanthophora spicifera, Chaetomorpha antennina, and Ulva reticulata. Superoxide dismutase, ascorbate peroxidase, glutathione peroxidase, and catalase are increased when copper and cadmium are present in excess; these enzymes can be used as a bioindicator of marine pollution and metal tolerance and thus can be implicated in bioaccumulation [36,58].

17.5 ENZYMES FOR BIOFUELS The use of enzymes in the biorefineries of third generation is fundamental for the production of fermentable sugars in the production of biofuels and structural and chemical modification of macroalgae polysaccharides and it has been studied different strategies for enzyme effective utilization, enzymatic pretreatments for polysaccharide hydrolysis (saccharification), simultaneous saccharification and fermentation, immobilization, enzymatic cocktails, and genetic engineering [59].

17.5.1 ENZYMES IN THE BIOETHANOL PRODUCTION Considering the high count of carbohydrates present across almost all genera of macroalgae, the little to nonpresence of lignin, bioethanol may well be the immediate choice for biofuel production. Although the algal cell wall is considerably less convoluted as lignocellulosic materials, pretreatment is still widely used, even before enzymatic saccharification, because the reducible sugars are not available for the alcohologenic microorganisms to produce ethanol. Mechanical, physicochemical, alkali, and acid pretreatment are still used to increment the reaction area, break the bonds between structural polysaccharides, otherwise inaccessible for enzymatic hydrolytic action [60]. One of the most widely used pretreatments is acid hydrolysis and/or hydrothermal treatment, due to low cost and relatively low concentrations of acid used but has one detriment, the glucose byproducts formation, namely 5-hydroxymethylfurfural and levulinic acid, both compounds have a direct negative effect in bioethanol production. Hydrothermal pretreatment is a cost-effective and low residue pretreatment; hot water is used as solvent usually from 100 C to 374 C coupled with high pressures, usually from 10 to 60 bar, to induce an autohydrolysis phenomenon, also known as self-ionization of the water, where the H2O deprotonates and forms a hydroxide ion OH2, and the

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hydron ion protonates another water molecule forming H3O1; it is theorized that this phenomenon is catalyzed by the sulfated groups (2SO3H) and the sulfated index of the macroalgae [12,16,59,61]. Production of biofuels such as butanol, bioethanol, and hydrogen from macroalgae using different microorganisms is summarized in Table 17.2.

17.5.1.1 Genetic engineering for bioethanol and biofuel production The genetic manipulation is a useful tool, elucidating the metabolic pathways to find key genes to enhance fermentation, which could be exploited through bioinformatic and genetic engineering, for example, recently the genome of TetV-1, mimiviridae (uncommonly large virus) infecting green algae, was characterized, founding the mannitol metabolism enzyme mannitol 1-phosphate dehydrogenase, the saccharide degradation enzyme α-galactosidase, and the key fermentation genes pyruvate formate-lyase and pyruvate formate-lyase activating enzyme [77]; these genes can be transfected to an expressing vector, to enhance green macroalgae enzymatic saccharification. Or take advantage of the intrinsic metabolic characteristic such as the Sphingomonas sp. A1 superconduct that enables to engulf soluble and/or insoluble alginate macromolecules with a genetic overexpression of Zymomonas mobilis pdc and adhB (promoter and alcohol dehydrogenase) [78], modify genetic restrictors of catabolic pathways [79], confer thermoresistant properties [23], halotolerant properties [80], improve binding capabilities [81], or export metabolic pathways to alcohologenic microorganisms to broad the assimilable carbon sources. Table 17.3 shows the genetic engineering of enzymes and the enzymatic action of macroalgae polysaccharides.

17.5.2 ENZYMATIC SACCHARIFICATION OF GREEN MACROALGAE After the pretreatment takes place, enzymatic treatment has proven to be an effective way of saccharification of the remaining glycosidic bonds of the polysaccharides. Marine bacteria Bacillus sp. strain BT21 are known to produce xylanases. Endoxylanases cleave the β-1,4-glycosidic linkages of the xylan, an organic polymer more commonly found in lignocellulosic biomass. Saccharification in SSF of pretreated biomass of xylan-rich macroalgae; Ahnfeltia plicata, Padina tetrastromatica, and U. lactuca (a red, brown, and green algae, respectively) using Bacillus sp. strain BT21 was conducted in a study to measure the saccharification activity of xylanases after a hot water pretreatment; Ahnfeltia sp. and Padina sp. biomass was more effectively hydrolyzed by the pretreatment; A. plicata biomass after the pretreatment and enzymatic saccharification released about 57% of reducing sugars; P. tetrasomica biomass were fucose-free, meaning the enzyme activity nor the enzymatic treatment released fucose from the cell wall, but achieved 53% of reducing sugars released after the enzymatic treatment; U. lactuca presented 37% of glucose and xylose release, pointing out a possible xyloglucan-like structure in macroalgae [84]. Vibrio parahaemolyticus, a halotolerant bacterium, has outstanding cellulose hydrolytic activity, was used for the enzymatic hydrolysis of Ulva intestinalis and U. lactuca, green algae known to have cellulose-rich composition. V. parahaemolyticus was isolated from its marine habitat and propagated in a carboxymethyl cellulose medium as sole carbon source, to produce halotolerant cellulose; the highest titer was achieved at 24 h and 14% of salt concentration, obtaining 2.11 U/mL and specific activity of 6.05 U/mg. After a dilute acidic pretreatment of Ulva intestinalis and Ulva lactuca and the enzymatic hydrolysis have taken place 135.9 and 107.6 mg/g [28].

Table 17.2 Biofuels from macroalgae. Algae

Microorganism

Enzymes involved

Products yielded

Yield

References

Ulva lactuca

Clostridium acetobutylicum

Acetoacetate decarboxylase (adc), alcohol/aldehyde dehydrogenase (adhE), and butyrateacetoacetate CoAtransferase (ctfA/B)

Acetone, butanol, and ethanol (ABE)

0.35 g ABE/g sugar consumed

[107]

Laminaria hyperborea

Zymobacter palmae

Mannitol dehydrogenase

Bioethanol

0.61 g ethanol (g mannitol)21 after 21.9 h under oxygenlimiting conditions

[40]

Sac. japonica

Bacillus sp. JS-1/Pichia angophorae KCTC 17574

Glucosidases, mannitol dehydrogenases, xylanases

Bioethanol

7.7 g/L (9.8 mL/L)

[63]

Sac. japonica

Escherichia coli engineered with Zymomonas mobilis pyruvate decarboxylase (Pdc) and alcohol dehydrogenase B (AdhB)

Alginate lyase, improved function of mannitol dehydrogenase, and alcohologenic enzymes

Bioethanol

20 g/L or 2.4% v/v

[64]

Gelidium amansii

Brettanomyces custersii KCTC 18154P

β-Agarases and α-neoagarobiose hydrolases

Bioethanol

11.8 g/L of bioethanol in batch reactor, 27.6 g/L of bioethanol in continuous reactor

[65]

n/a

Vibrio sp. EJY3

Endo-β-agarase I (DagA), exo-β-agarase II (Aga50D), and neoagarobiose hydrolase (NABH)

Bioethanol

37.1% theoretical maximum yield of bioethanol

[66]

Laminaria japonica

E. coli KO11 (Zymomonas mobilis ethanol production genes) and Saccharomyces cerevisiae

Mannitol dehydrogenase and alcohologenic enzymes

Bioethanol

Ethanol yield 23 29 g/L

[67]

Sac. japonica

Defluviitalea phaphyphila Alg1

Glucosidases, mannitol dehydrogenases, alginate lyase

Bioethanol

0.47 g/g-mannitol, 0.44 g/ g-glucose, and 0.3 g/galginate; 0.25 g/g-kelp

[68]

U. lactuca

Clostridium beijerinckii, Clostridium saccharoperbutylacetonicum

Acetoacetate decarboxylase (adc), alcohol/aldehyde dehydrogenase (adhE), and butyrateacetoacetate CoAtransferase (ctfA/B)

Butanol

15.2 g/L of reducing sugars

[69]

Ahnfeltia plicata, Padina tetrastromatica, and U. lactuca

Bacillus sp. strain BT21

Xylanase

Depolymerization to obtain fermentable sugars

233 6 5.3, 100 6 6.1 and 73.3 6 4.1 lμg/mg of seaweed biomass

[84]

Laminaria spp.

S. cerevisiae

1 U/kg laminarase

Ethanol

Ethanol yield of 0.45% (v/v) w/o treatment

[70]

Hydrolysates of Ulva intestinalis and U. lactuca

Vibrio parahaemolyticus

Cellulases

Fermentable sugars

135.9 mg/g and 107.6 mg/g of reducing sugar, respectively

[28]

Sac. japonica

Clostridium sp.

Acetoacetate decarboxylase (adc), alcohol/aldehyde dehydrogenase (adhE), and butyrateacetoacetate CoAtransferase (ctfA/B)

Hydrogen and VFAs

Maximum hydrogen of 179 mL/g-VS and VFAs concentration of 9.8 g/L were produced from 35 g/L of S. japonica within 5 days of anaerobic fermentation

[113]

Sac. japonica

Clostridium sp.

Hydrogenase

Hydrogen and volatile fatty acids (VFAs)

Hydrogen production of 179 mL/g-VS and VFAs concentration of 9.8 g/L were produced from 35 g/L

[113]

U. lactuca

Formosa agariphila

Ulvan lyase, an unsaturated glucuronyl hydrolase, one xylosidase, and two rhamnosidases

Oligosaccharide depolymerized to monomeric sugars

n/a

[71]

(Continued)

Table 17.2 Biofuels from macroalgae. Continued Algae

Microorganism

Enzymes involved

Products yielded

Yield

References

Sac. japonica

V. harveyi ATCC 14126 or Vibrio alginolyticus ATCC 17749

alginate lyases

VFAs

The amount of VFA produced from 40 g/L of S. japonica increased from 8.3 g/L (control) to 15.6 g/L when it was biologically pretreated with Vibrio harveyi

[74]

Sac. japonica treated with β-cyclodextrin

Clostridium

Hydrogenase

VFAs and biohydrogen

VFAs (12.5 g/L) at 38.6 C and 7.4 g/L β-CD

[75]

Table 17.3 Genetic engineering of enzymes. Original enzyme producer

Recombinant/ cloning vector

Vibrio splendidus 12B01

Enzyme

Modification

Objective

References

Escherichia coli BL21 (DE3)

Oligoalginate lyases (OalA, OalB, and OalC)

The genes were amplified from the genomic DNA of V. splendidus 12B01 using the oligonucleotide primers. The sequences of the oligonucleotide primers are based on the DNA sequences of the oligoalginate lyases (OalA, OalB, and OalC) from V. splendidus 12B01

Depolymerize constitutive monomers

[82]

Vibrio sp. QY101

E. coli BL-21 LysS.

Alginate lyase (AlyVI)

Site-directed mutagenesis of AlyVI cDNA

Depolymerize oligomeric alginates

[76]

Stenotrophomonas maltophilia k279a

E. coli smlt1473

Polysaccharide lyase (Smlt1473)

Single point mutations to H221F and R312L for increased activity and specificity for poly-Dglucuronic acid (polyGlcUA), and poly-Dmannuronic acid

Enhance binding capabilities of polysaccharide lyase

[81]

Zymomonas mobilis

Sphingomonas sp. A1

Alginate lyase, alcohol dehydrogenase B (adh B), pyruvate dehydrogenase complex (pdc)

Disrupting the lactate dehydrogenase gene, and overexpressing adh B and pdc of Z. mobilis in Sphingomonas sp. A1

Harness the superconduct in Sphingomonas sp. A1 to production of ethanol from alginate as sole carbon source

[78]

Vibrio sp.

E. coli BL21 (DE3)

3,6-Anhydrogalactonate and 2-keto-3deoxygalactonate

The catabolic enzymes for 3,6-anhydro-Lgalactose (AHG0) were inserted in a recombining bacterium

Use AHG as sole carbon source for biofuel production and industrial chemicals

[92]

(Continued)

Table 17.3 Genetic engineering of enzymes. Continued Original enzyme producer

Recombinant/ cloning vector

Saccharophagus degradans 2-40T

Enzyme

Modification

Objective

References

E. coli BL21 (DE3)

Bgl1B, β-glucosidase

The catabolic enzymes for β-glucosidase of S. degradans 2-40T were inserted in a recombining bacterium

Break down the cleaving β-1,3-, β-1,4-, and β-1,6glycosidic linkages of laminarin and produce high-value laminarioligosaccharides

[87]

Vibrio sp. EJY3

E. coli BL21 (DE3)

Agarolytic β 1-4-Dgalactosidase (VejABG)

The catabolic enzymes for agarose hydrolases were introduced into a recombinant vector for expression

D-Galactose

and 3,6anhydro-L-galactose

[91]

Thermoanaerobacter brockii

E. coli and Brevibacillus choshinensis

Thermostable βglucosidase

The cglT gene encoding a thermostable BGL in T. brockii DSM1457 was amplified and inserted into the plasmid pQE30 at BamHI and PstI sites in E. coli JM109 and B. subtilis RIK1285

Thermostable enzymes for a cost effective cellulose saccharification

[103]

Cellulophaga sp. QY3

E. coli strains DH5a and BL21 (DE3)

i-Carrageenase (CgiA_Ce)

The CgiA_Ce gene encoding a thermostable icarrageenase was introduced into a recombinant vector

Cleaved i-carrageenan yielding neo-i-carrabiose and neo-i-carratetraose as the main end-products, and neo-i-carrahexaose was the minimum substrate

[83]

Pseudoalteromonas carrageenovora

E. coli DH5α

λ-Carrageenase

The gene CglA encoding the catabolic enzyme for λ-carrageenase was introduced into a recombinant E. coli

Hydrolyzes the β-(1-4) linkage of λ-carrageenan, neo-λ-carratetraose, and neo-λ-carrahexaose

[32]

Postechiella marina M091, Pseudoalteromonas atlantica T6c, Streptomyces coelicolor A3 (2), Thermoplasma acidophilum, Aspergillus niger

E. coli DH5α, BL21 (DE3) and Rosetta (DE3)

α-Neoagarobiose hydrolase, 3,6-anhydro-Lgalactonate cycloisomerase, 3,6anhydro-L-galactose dehydrogenase, 2,5diketo-3-deoxy-Lgalactonate 5-reductase, 2-keto-3-deoxy-Lgalactonate 5dehydrogenase

Predict the enzyme function of four core genes in the L-AnG gene cluster

Elucidate the metabolic pathway to obtain 3,6anhydro-L-galactose and D-galactose from agarose

[88]

E. coli

Engineered E. coli

Synthetic promoters and 50-UTRs were designed to deregulate the control system of glucose and galactose consumption

Remove carbon catabolite repression and amplify galactose utilization rate

Simultaneously assimilate galactose and glucose and improve galactose assimilation rate.

[79]

S. degradans 2-40, S. coelicolor A3

E. coli BL21 (DE3)

Aga16B, Aga50D and NABH coding genes from S. degradans 2-40 and dagA gene from S. coelicolor A3

Insert the encoding genes from three agarases and α-neoagarobiose hydrolase (NABH) to a recombinant expression vector

Simultaneous saccharification and fermentation from agarose to obtain bioethanol

[89]

Neurospora crassa

Saccharomyces cerevisiae

Cellodextrin transporter (cdt-1) and intracellular β-glucosidase (gh1-1)

Insertion of the genes for cellodextrin transportation and an intracellular glucosidase to enable the simultaneous fermentation of cellobiose and galactose to an expression vector

Simultaneous saccharification and fermentation of cellobiose and galactose

[90]

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17.5.2.1 Enzymatic saccharification of brown macroalgae Regarding alginate saccharification and oligosaccharide production, it was initially complicated because no ethanologenic microorganisms can degrade alginate [78], but there are many microorganisms, especially bacteria such as Sphingomonas and Flavobacterium spp. which are known to produce alginate lyases capable of catalyzing cleavage of alginate for utilizing alginate as carbon source. Alginate lyases also help reduce the viscosity of brown algae during processing. Manns et al. [72] studied the substrate specificity and substrate viscosity effect on different microbial alginate lyases. They concluded that the alginate lyase from Sphingomonas sp. (SALy) had high enzymatic activity on polyguluronic acid, making this enzyme promising candidate to support glucose release from brown macroalgae. Vibrio splendidus 12B01 is also known to have three functional oligoalginate lyases (OalA, OalB, and OalC) with a notable substrate action, di-, tri-, tetra- and pentasaccharides, and extracellular endo-acting and exo-acting alginate lyases (A1yA, AlyB, AlyC, and AlyD). Functionality for the oligoalginate lyases was studied using a recombinant bacteria; OalA seems to have the higher specific activity (28.5 U/mg) for alginate, cleaving various sizes of poly(M), but as di- or trisaccharides were not observed in the alginate degradation products of the recombinant bacteria, probably due exo-processive action [82]. Also, for alginate depolymerization, the bacterium Stenotrophomonas maltophilia k279a is known to be able to cleave multiple anionic polysaccharides via β-elimination mechanism, because its polysaccharide lyase (Smlt1473) have a strong activity against hyaluronan, poly-β-D-glucuronic acid, and poly-β-D-mannuronic acid, which can be effectively improved by single point mutations; the hyaluronan degrading activity was almost eliminated, while the poly-β-D-glucuronic acid and poly-β-D-mannuronic acid activity was improved by 35% [81]. Laminarin saccharification from brown algae can be realized to some extent by commercial cellulase mixtures developed for terrestrial cellulosic biomass—presumably due to a broad specificity and ability of some of the β-glucanases to catalyze cleavage of the β-1,3 bonds in laminarin. Manns et al. [86] studied the effect of different milling pretreatments on enzymatic hydrolysis for glucose production from brown seaweed Laminaria digitata with high content of glucan. They concluded that the small particle size of brown seaweed did not increase the surface area in the enzymatic hydrolysis process, and the fungal cellulases were able to hydrolyze the laminarin (glucan) from brown seaweed to glucose. Also, laminarin saccharification was accomplished by a recently discovered β-glucosidase from Saccharophagus degradans 2-40T; Bgl1B (a gene studied by overexpression in a recombinant bacteria), which is able to hydrolyze, laminaribiose, cellobiose, gentiobiose, lactose, and agarobiose, by cleaving β-1,3-, β-1,4-, and β-1,6-glycosidic linkages, but it shows more affinity for laminaribiose using as a sole carbon source to produce glucose, additionally Bgl1B can produce laminarioligosaccharides with various degrees of polymerization by transglycosylation, meaning a glycosidic bond formation [87].

17.5.2.2 Enzymatic saccharification of red macroalgae Agarose can be effectively broken down to agarooligosaccharides (AOSs) by Vibrio sp. strain EJY3, by its particular β-galactosidase, unlike the lacZ-encoded β-galactosidase of E. coli, does not hydrolyze lactose, cleaving β1-4 linkages at the nonreducing end, to release galactose and neoagarooligosaccharides (NAOSs) from AOSs, improving the percentage of reducing sugars obtained

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from acetic acid-hydrolyzed agarose, β1-3 linkages are commonly hydrolyzed by weak acid pretreatment. It was found that Vibro sp. EJY3 has a unique enzyme, VejABG the first β-galactosidase found to be capable of hydrolyzing AOSs (agarolytic β1-4-D-galactosidase); This novel enzyme acts on the β1-4 linkage only at the nonreducing end of AOSs between galactose and L-AHG [91]. β-Agarase I hydrolyzes agarose to produce neoagarohexaose and neoagarotetraose and later hydrolyzes to neoagarobiose by β-agarase II [85]. Residual NAOSs, mainly neoagarotetraose may not be hydrolyzed. Complete saccharification of AOSs to D-galactose and L-AHG can be achieved using a combination of NAOS hydrolase or a neoagarobiose hydrolase (NABH) from S. degradans 2-40T (SdNABH) [87] and VejABG, cleaving both the β1-3 and β1-4 linkages, respectively, at the nonreducing end of a neoagarotetraose molecule, successfully hydrolyzing agarose to monomers [91]. Nevertheless, the catabolic pathway for L-AHG of Vibrio sp. EJY3 has been identified in the genes VejAHGD and VejACI and studied through amplification in an alcohologenic E. coli KO11 to successfully convert L-AHG or agarose hydrolysates, as sole carbon source, to bioethanol at a 1.2-fold higher than the control strain [92]. The complete catabolic pathway of LAHG is still not fully investigated, but different metabolic intermediates have been elucidated to help fully utilize L-AHG in biorefinery processing [93]. Notwithstanding, L-AHG has been recently studied to serve as anticariogenic sugar, obtaining better inhibitory effects in comparison with xylitol against Streptococcus mutans [39]. Agar from Gracilaria verrucosa, an industrially important red algae can be hydrolyzed completely by an acid pretreatment combined with enzymatic saccharification, using recombinant agarases Aga50D and NABH produced originally in S. degradans 2 40 and later overexpressed in E. coli DH5α, said enzymes improving the total reducing sugars released from G. verrucosa; 34.9% total reducing sugars (TRS) yield (only with acid pretreatment), and a yield of 47.4% TRS with enzymatic treatment was achieved [94]. CRG is a complex polysaccharide with specific binding sites in the backbone for precise enzymes, κ-, ι-, and λ-carrageenases, which are endohydrolases that cleave β1-4 bonds obtaining oligogalactans, neocarrabiose, or neoagarobiose [95]. These oligosaccharides have been shown to exhibit antitumor and antiviral properties [96], and its hydrolytic enzymes have industrial applications outside the bioethanol production; textile as a biological dye thickener, detergent additive, since CRG is a widely used as food additive, carrageenases will ease the removal of stains, since CRG are highly affined to cellulose fibers, and algal protoplast isolation, useful in genetic engineering [95]. Recently, a multifunctional enzyme was found; with amylase activity, with unspecific agarase and carrageenase action, were discovered in a marine bacterium Vibrio alginolyticus 63; the enzyme was coined Amy63 and was expressed in recombinant E. coli improving its starch, agar, and CRG degradation capacities [97]. The said enzyme could be induced by red algae biomass and there are ubiquitous properties in enzymes of marine organisms such as hyperthermoestability, barophilicity, pH and salt tolerance, resilience against extreme cold conditions, and product of the evolutionary adaptation due to the harsh conditions of its environment. Until recently, fungi with agarolytic and carrageenase activity were not extensively documented; carrageenase activity was only documented in Aspergillus ochraceus, Aspergillus terreus, and Phoma sp. [98]; however, a study of microorganisms retrieved from the Antarctic showed that Beauveria bassiana, Cladosporium sp. 2, Doratomyces sp., Penicillium chrysogenum, Penicillium citrinum, Penicillium sp., and Pseudogymnoascus sp. were able to produce agarolytic and/or

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carrageenolytic activities. A thalli sample of Iridaea cordata (an Antarctic red algae) contained the fungi Penicillium sp. UFMGCB 10066 that showed the best carrageenase activity, 17.49 6 3.21 U/ mL (amount of enzyme required to recover 1 μg of galactose per minute). Cold-adapted enzymes have broad industrial potential because of its high specific activities ranging from low to moderate temperatures and moderate temperature increases induce inactivation [25,85].

17.5.3 SACCHARIFICATION BY ENZYMATIC COMPLEXES, COCKTAILS, AND SIMULTANEOUS FERMENTATION Enzyme complexes are hydrolytic enzymes contained in scaffolding protein or scaffoldin (CbpA), one of the most studied enzymatic complex regarding algal biomass saccharification is the cellulosome derived from Clostridium sp.; it has hydrolytic enzyme modules dockerin domain linked to a cohesion domain in the scaffoldin that also presents a carbohydrate-binding molecule, or surface layer homology domain (SLH) useful to ease the purification process or enable an enzymatic immobilization. [93]. It is reported that the complex enhances the enzymatic activity due to the distance restriction, improving its synergic activity in comparison to a single enzyme activity [37]. The effectivity of enzymatic complexes was studied in red macroalgae, G. verrucosa assembling an enzymatic complex using the chimeric AgaB and AhgA genes for the β-agarase, anhydrogalactosidase from Z. galactanivorans and CgkA gene for κ-carrageenase from Pseudoalteromonas carrageenovora were expressed in a recombinant bacteria, purified, and fused in a complex using mCbpA as scaffoldin protein randomly, as it was found that more than one band were present in the western blot analysis, and it was also found an improvement of 3.6-fold higher of sugar release; cAgaB yielded 188.6 against 679 mg/L obtained from the enzymatic complex [37]. Another study [99] of the integration of the bacterial expansin of Bacillus pumilus, into a recombinant scaffoldin miniCbpA from Clostridium cellulovorans containing chimeric β-agarase cAgaB from Z. galactanivorans. Expansis are nonhydrolytic enzyme that loosens hydrogen bonds, and this activity was coupled with agarolytic complexes proven to increase enzymatic activity over purified agar and natural biomass of Gelidium amansii obtaining a 3.7-fold and 3.3-fold higher, respectively, in comparison to a single enzyme. Concluding that expansin can effectively improve hydrolysis in agarolytic enzymatic complexes, and also modify the gelling of the agar. Enzymatic cocktails are a combination of different enzymes with the objective to break down complex biomass. Endemic microorganisms of the macroalgae have been proven an optimal source to produce enzymatic cocktails, Saccharina digitate was degraded up to 90% using enzymes isolated from a marine fungus [34]. Aspergillus niger, a highly utilized fungi in biotechnology was used to produce a cocktail to degrade the polysaccharides of Ulva rigida, the cocktail contained β-glucosidase (109 IU/mL), pectinase(76 IU/mL) and carboxymethyl cellulase (4.6 IU/mL) and it was produced using U. rigida as nitrogen source. This cocktail was applied over raw algae extract as pretreatment and then fermented using anaerobic sludge, in anaerobic fermentation conditions to produce biogas [100]. Simultaneous saccharification is a joint process merging the saccharification process with the fermentation phase using only one reactor, using specialized microorganisms or enzymes to reduce complex biomass to its monomers during simultaneous fermentation of the monomers to, for example, ethanol [101]. A study [102], comprising a new enzymatic cocktail (Haliatase) that contained a

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mixture of β-glucanase (1875 U/g), carrageenase (315 U/g,) and agarase (440 U/g) obtained from the hepatopancreas of abalone snails, and milling procedures of two macroalgae; Enzymatic simultaneous saccharification and fermentation using the cocktail and S. cerevisiae for ethanol production of U. lactuca was not improved by centrifugal milling obtaining 6 g ethanol/100 g TS (total sugars) after 72 h; G. sesquipedale was improved with centrifugal milling from 2 to 4 g of ethanol/ 100 g TS. Although the difficulty is that alcohologenic microorganisms usually do not consume pentoses, metabolically engineered bacteria capable of producing ethanol from formerly not consumable compounds, recombinant bacteria are useful to achieve this kind of process, a study expressing the β-glucosidase gene from Thermoanaerobacter brockii in E. coli and Brevibacillus choshinensis for simultaneous enzyme production and saccharification from cellulosic biomass; 0.5 U/mL of β-glucosidase activity was found in E. coli, and 0.74 U/mL of β-glucosidase activity in B. choshinensis, implying that its metabolic pathways are more suited for recombination [103]. This could be proven useful in the construction of consolidated bioprocess in the future for cellulosic-rich macroalgae.

17.5.4 BIOBUTANOL AND ACETONE, BUTANOL, AND ETHANOL FERMENTATION The concept of butanol as a biofuel is a promising one, being an alcohol with a long alcohol chain, nonpolar, and bears an energy content similar to that of fossil fuels, and thus additional modifications in combustion engines are not needed; it can be used in a pure solution or in a mixture in a gasoline blend [104], and can also be shipped in existing pipelines [101]. As the actual trend in stagnant sales of flex-fuel vehicles (FFVs, which are vehicles that can run in a bioethanol blend up to 85%), and the overall reduction in production of FFV models by some automakers [105] makes biobutanol a possible viable alternative. The acetone, butanol, and ethanol (ABE) fermentation is a strict anaerobic process, involving classically bacteria of the Clostridia genera, transforming carbohydrates into acetone, butanol, and ethanol. Algal biomass is rich in carbohydrates and some studies have been conducted to explore the biobutanol production, acid hydrolysates of U. lactuca with an average reducing sugar content of 15.2 g/L, was fermented in a larger scale using a classic ABE reactor, and in a two-phase ABE reactor, the difference is that the classic fermentation is a batch fermentation using bacteria with both acidogenic and solventogenic phases, such as Clostridium acetobutylicum, with recommended product removal in situ of products, as butanol is known to be a fermentation inhibitor. In the two-phase process, sugars are initially converted to butyric acid by anacidogenic bacteria such as Clostridium tryobutyricum and then fed to a second reactor with a solventogenic bacteria such as Clostridium beijerinckii, the study shown an average maximum yield of 4 g/L of biobutanol after 8 days, showing no relevant difference between both process [106]. Another study using the same algae, U. lactuca, implementing a hydrothermal pretreatment, followed by enzymatic saccharification via commercial cellulases prior to fermentation, using C. beijerinckii; obtained a yield of 0.35 g ABE/g sugar consumed. However, it was also found that C. beijerinckii in a glucose and rhamnose-rich culture produced 1,2-propanediol (a high valued chemical), as sole product [107]. Algal extracts of Saccharina spp. were also studied for ABE fermentation by C. acetobutylicum obtaining butanol and total solvent yields of 0.12 and 0.16 g/g, respectively. It was discussed that if alginate could also be metabolized to produce solvents, the yield would be higher [108].

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17.5.4.1 Enzymatic engineering for acetone, butanol, and ethanol fermentation The yields of the fermentation via biochemical conversion are low compared with chemical synthesis from alcohol, [104] and efforts are being made in improving the conversion rates, In the butanol synthetic pathway, six enzymes are needed to complete the conversion of acetyl-CoA to butanol, including, acetyl-CoA acetyltransferase (THL, or thiolase), β-hydroxybutyryl-CoA dehydrogenase (BHBD), 3-hydroxybutyryl-CoA dehydratase (CRT or crotonase), butyryl-CoA dehydrogenase (BCD), butyraldehyde dehydrogenase (BAD), and butanol dehydrogenase (BDH). Although the bottleneck is the lack of understanding of the clostridia metabolic regulations involving the shift of the acidogenic to the solventogenic phase, and how it is intertwined with the rest of the regulatory networks. There has been made some studies to boost solvent tolerance, in C. acetobutylicum M5, blocking butyrate kinase pathways to reduce butyrate and improve butanol formation obtaining 0.84 g butanol/g ABE, in comparison withB0.6 g butanol/g ABE produced by the wild-type, transferring the clostridial fermentative genes to E. coli and Bacillus subtilis for butanol production [62,109], enlarge the assimilable carbon sources such as cellulose, by introducing a minicellulosome to degrade cellulose more effectively, or implementing a dual fermentation using β-galactosidase Bga2 of Clostridium stercorarium to improve the function of the endoglucanase Cel9D from Clostridium thermocellum to degrade xyloglucans [110,111], can be also implemented in cellulose-rich macroalgae, to enhance butanol production or to obtain oligosaccharides with prebiotic activity.

17.5.5 ENZYMES IN BIOGAS PRODUCTION The anaerobic digestion is a fermentation approach to obtain biofuels as a result of a series of reactions, hydrolysis, acidogenesis, acetogenesis, and methanogenesis. Sugars and amino acids are converted into alcohols and carboxylic acids, known as volatile fatty acids (VFA), which are intermediary products of acidogenic bacteria, then transformed into acetic acid, hydrogen, and CO2. Later VFAs are converted into methane and CO2 by methanogenic bacteria [112]. Biomethane yield production reported for macroalgae ranges from 0.12 to 0.48 m3 CH4/kg VS (volatile solids), but there are several challenges to overcome; high nitrogen content of macroalgae results in high levels of ammonia, and inhibitory to methanogen producers, pretreatments are needed to depolymerize cell walls, and endogenous alkaline metals of macroalgae can inhibit anaerobic fermentation [60]. Biomethane enhancing production via enzymatic engineering is more investigated in microalgae-applied protocols. Biohydrogen and VFAs can be produced through Sac. japonica biomass anaerobic fermentation by Clostridium sp. obtaining 9.8 g/L and 179 mL/g-VS of hydrogen were obtained from 35 g/L of Sac. japonica within 5 days, within the same study it was found that the selection of methanogenic inhibitors such as β-cyclodextrin and iodoform, β-cyclodextrin being the most effective one [113]. Macroalgae biomass or sludge can be used with a cosubstrate such as waste paper or manure, regulates de carbon-nitrogen rate obtaining improved yields of biohydrogen [15,60]. Additionally, zero-valent Fe0 nanoparticles promote favorable acid formation of hydrogenproducing strains Clostridium and Terrisporobacter sp. over macroalgae in dark fermentation procedures using Sac. japonica as substrate obtaining 20.25 mL H2/g VS [114]. Enzyme employment in biohydrogen production is more oriented toward the saccharification of macroalgal biomass. A study comparing the production of hydrogen between galactose and

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enzymatic hydrolysates of agar using the recombinant enzymes agarase (AgaXa) and neoagarobiose hydrolase (NH852) and hydrothermal treatment showed a 3.86-fold higher yield for the agar hydrolysates; 303.2 mL/g of hydrogen was obtained in the cultivation media containing 5.87 g/L of galactose, whereas 5047 6 228 mL/L of hydrogen was obtained from 50 g/L of agar hydrolysate [73]. The efficiency of enzymatic saccharification to produce biogas; using U. rigida as nitrogen source to produce an enzymatic cocktail using Aspergillus niger, said cocktail was rich in β-glucosidase, pectinase, and cellulases, producing 1175 mL/ g COD of biogas, is theorized that enzymatic digestion improves anaerobic fermentation [100].

17.6 ENZYMATIC EXTRACTION OF HIGH VALUED PRODUCTS Macroalgae offer an extensive array of obtainable substances with bioactive properties, contained in the cell walls; the pursuit for cleaner extraction procedures offers eco-friendly alternatives to classic solvent, alkaline, and acid extractions; pressurized extraction, ultrasound-assisted, microwave-assisted and case in point, enzymatic-assisted extraction (EAE) [115]. The main advantage over the other procedures is the catalytic efficiency, specific hydrolytic activity, wide range of reactive conditions [116], low energy usage, and reduced extraction times. It can be implemented industrially but nowadays is restricted by the enzyme cost, and commercially available enzymes [116,117].

17.6.1 EXTRACTION OF HYDROCOLLOIDS Some of the polysaccharides, such as alginate from brown seaweed, agar, and CRGs from red macroalgae, have important commercial niches that can be integrated into the biorefinery process to improve profits [118]; these polysaccharides are intertwined with proteins, polymeric phenols, calcium, or potassium ions [119]. A study exploring agar extraction with alkaline, enzyme-assisted extraction with commercial enzymes using 8 U/mL cellulase and 26.6 U/mL arylsulfatase (Venzyme: Wseaweeds 5 20:1) at 50 C, for 3 h and a combination of acidic, alkaline, and enzymatic protocols, 4 U/mL (20:1) cellulase at 50 C for 1 h, followed by a treatment with NaOH solution at 87 C for 3 h, rinsed, and treated with an acid solution with 0.064% using red alga Gracilaria lemaneiformis biomass; the alkaline extraction showed the optimal performance obtaining the best gel strength, sulfate content, and L-AHG content, the combined protocol obtained similar results but generated lower sewage discharge, and the EAE obtained good agar integrity, and it was concluded that it is a promising alternative to alkaline extraction [115]. Ultrasound-assisted extraction (UAE) coupled with EAE was studied using Sargassum muticum as raw biomass and residual algae from alginate production; commercial enzyme complexes were tested; Celluclast had the best fractionation performance in the raw biomass; Celluclast and amylase were the best formulations for the alginate free residue; it was found that neither the joint extraction process, nor individually, improves dramatically the phenolic extraction yields. Nevertheless, it was found that UAE process was better at longer prolonged times, due to synergism with the enzyme treatment, possibly meaning a facilitated substrate enzyme interaction and disruption of the cell wall to selectively extract phenolic compounds [120]. It is reported the

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importance for the optimization in coupling of extraction protocols such as subcritical water extraction, a combination of high-pressure (50 300 psi) and high-temperature (50 200 C), with EAE to obtain polyphenols and phlorotannins and other bioactive compounds [121,122]. Several algal biomasses have been studied for optimization parameters of polysaccharide extraction including Ecklonia radiata [123], G. verrucosa [124], G. amansii [125], Sac. japonica [125], among others.

17.6.1.1 Extraction of phenolic compounds Phenolic compounds have multiple health benefits; it has antioxidant, antiinflammatory, and antitumor properties; they can be considered as functional food ingredients with antioxidant and antimicrobial properties. Polyphenols can be conjugated glycosides (flavonoids) or insoluble (phenolic acids) [119]. A study involving extracts of Ulva armoricana for the obtention of antivirals and antioxidants through EAE using a multiple-mix of commercial glycosyl hydrolases and an exo-β-1,3(4)-glucanase (Novozymes, Bagsværd, Denmark) obtaining hydrolysates 2000 and 600 kDa (ulvans), and a group of oligosaccharides (3 kDa), the half-maximal effective concentration (EC50) was found to be 320.9 6 33.6 μg/mL of the hydrolysate fraction against herpes simplex virus type 1 (HSV-1). Also, some of the enzymatic combinations resulted in concentrated amounts of polyphenols, from 0.6% to 1.0%. Nevertheless, inhibiting concentration (IC50) was not significant against the Butylated hydroxyanisol and butylated hydroxytoluene standards [126]. Enzymatic hydrolysates of Red seaweed Solieria chordalis was also studied for antiviral activities against HSV-1; three proteases (subtilisin, neutral metalloproteinase, and exopeptidase) and five carbohydrases (amyloglucosidase, α-amylase, endoxylanase, endo-β-glucanase, polygalacturonase, β-glucanase), all commercial enzymes, where the extract using proteases obtained the best antiherpetic activities, EC50 of 86.0 μg/mL, also it was reported the increased yield of sulfated polysaccharides [127]. Enzymatic extracts of Chondrus crispus and Codium fragile obtained with commercial proteases and carbohydrases was also tested against HSV-1, reporting a significant activity, EC50 of 77.6 126.8 μg/mL for C. crispus and 36.5 41.3 μg/mL for C. Fragile [128]. Antioxidants from Palmariapalmata, a red macroalgae, were extracted using protease and carbohydrase treatments; Umamizyme, a commercial protease, was found to have the best performance in extracting phenols; thus the hydrolysates of the Umamizyme extract had the best radical scavenging capacity against 2,2-diphenyl-1-picrylhydrazyl and peroxy radicals, the extracts could be implemented as natural antioxidants and functional food ingredients [129]. It is important to consider that phenolic compounds may be hepatotoxic, nephrotoxic, inflammatory, and even carcinogenic [119].

17.6.2 EXTRACTION OF PIGMENTS It is reported that extraction of algal bioactive compounds using enzymes can potentially improve yields and safety, and also cellulose of some macroalgae hinders the solvent extraction. Some pigments are valuable bioactive compounds, such as fucoxanthin a photosynthetic xanthophyll present in brown macroalgae, a study regarding the optimization of an enzymatic extraction of fucoxanthin from F. vesiculosus using a commercial enzyme (Viscozyme) to eliminate the use of solvents; showed a eco-friendly process with no toxic waste, that converts water-insoluble raw materials into partially soluble materials, obtaining a similar fucoxanthin yield of 0.657 mg/g (dry mass of F. vesiculosus blade) against an organic solvent (acetone) extraction yield 0.699 mg/g (dry mass of F.

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vesiculosus blade) [130]. Extracted cellulose of red algae (Gelidiella acerosa) was hydrolyzed using an optimized hydrolysis period, temperature, and dose of a commercial cellulase (Novozymes), was carried out in a small-scale trial, and the results suggested that a ton of fresh biomass could yield 0.3 0.7 kg of R-phycoerythrin, 0.1 0.3 kg of R-phycocyanin, 1.2 4.8 kg of lipids, 28.4 94.4 kg of agar, 4.4 41.9 kg of cellulose, and 3.1 3.6 kiloliters of mineral solution, also a efficiency bioethanol conversion of 89.08% [14]. Red macroalgae, Gelidium pusillum was also studied for an optimized R-phycoerythrin extraction, using an enzymatic cocktail of Agarase, Cellulase, and Xylanase in two proportions 80:40:80 and 40:80:80 (U/g biomass fresh weight, respectively), pH 5.8, and temperature of 25 C were found to improve R-phycoerythrin extraction [131]. Optimal parameters for R-phycoerythrin extraction from different macroalgae have been carried out, such as Mastocarpus stellatus [132], Furcellaria lumbricalis [133], and G. verrucosa [124].

17.7 CONCLUSIONS AND PERSPECTIVES Macroalgae is a promising renewable feedstock, and the third generation of biorefinery offers a wide platform to develop technologies to exploit it effectively, but the relatively costly downstream processing is still one of the biggest issues to overcome. Macroalgae biomass contains high-value low-volume bioproducts that can be obtained to broaden the incomes and in order the third generation of biorefinery to compete directly with fossil fuels, the process must be completely profitable; enzyme technology offers a wide array of uses but nowadays is limited to commercial enzymes and commercial genetic engineered strains. Essentially, all processes for macroalgal biorefining involve the depolymerization of the algae cell wall—although depolymerization may be partial or full, but the elucidation of the polysaccharide structure, key fermentation genes, manipulation of enzyme interactions and modification of microorganisms to enhance metabolic features will ease the use of the biomass more effectively or obtain high-valued compounds applicable across multiple industries. The technology to use the biomass as a source of enzymes, high-value products, the eventual reduction of costs regarding genetic manipulation technologies, and specialized strains may imply a reduction in the final costs of production, granting a positive impact on the perception of biofuels in the fuel market, boosting its availability to the consumer. Efforts are being made to fully understand enzymatic interactions and application regarding macroalgae, to develop scalable and profitable processes for macroalgal biorefining as an important part of the new bioeconomy substituting fossil fuel-based processes.

ACKNOWLEDGMENT This work was financially supported by the Innovation Incentive Program (PEI)—Mexican Science and Technology Council (SEP-CONACYT) with the Project (Ref. PEI-251186), titled: Study of the variation temporary space of Fucoidan from Sargassum SP. Abraham Lara thanks to the National Council for Science and Technology (CONACYT, Mexico) for his Master Fellowship support (grant number: 732281/8932).

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[98] M.J.L. Solis, S. Draeger, T.E.E. Dela Cruz, Marine-derived fungi from Kappaphycusalvarezii and K. striatum as potential causative agents of ice-ice disease in farmed seaweeds, Bot. Mar. 53 (2010) 587 594. Available from: https://doi.org/10.1515/BOT.2010.071. [99] D.W. Jeong, J.E. Hyeon, Y.C. Joo, et al., Integration of bacterial expansin on agarolytic complexes to enhance the degrading activity of red algae by control of gelling properties, Mar. Biotechnol. 20 (2018) 1 9. Available from: https://doi.org/10.1007/s10126-017-9782-4. [100] R. Karray, M. Hamza, S. Sayadi, Production and characterization of enzymatic cocktail produced by Aspergillus niger using green macroalgae as nitrogen source and its application in the pre-treatment for biogas production from Ulva rigida, Bioresour. Technol. 216 (2016) 622 628. Available from: https:// doi.org/10.1016/j.biortech.2016.05.067. [101] M.T. Ces´ario, M.M.R. da Fonseca, M.M. Marques, M.C.M.D. de Almeida, Marine algal carbohydrates as carbon sources for the production of biochemicals and biomaterials, Biotechnol. Adv. 36 (2018) 798 817. Available from: https://doi.org/10.1016/j.biotechadv.2018.02.006. [102] S. Amamou, C. Sambusiti, F. Monlau, et al., Mechano-enzymatic deconstruction with a new enzymatic cocktail to enhance enzymatic hydrolysis and bioethanol fermentation of two macroalgae species, Molecules 23 (2018) E174. Available from: https://doi.org/10.3390/molecules23010174. [103] S. Ichikawa, M. Ichihara, T. Ito, et al., Glucose production from cellulose through biological simultaneous enzyme production and saccharification using recombinant bacteria expressing the β-glucosidase gene, J. Biosci. Bioeng. 127 (2019) 340 344. Available from: https://doi.org/ 10.1016/j.jbiosc.2018.08.008. [104] B. Ndaba, I. Chiyanzu, S. Marx, N-Butanol derived from biochemical and chemical routes: a review, Biotechnol. Rep. 8 (2015) 1 9. Available from: https://doi.org/10.1016/j.btre.2015.08.001. [105] Renewable Fuels Association. Ethanol Industry Outlook. ,https://ethanolrfa.org/wp-content/uploads/ 2019/02/RFA2019Outlook.pdf., 2019 (accessed 29.05.19). [106] T. Potts, J. Du, M. Paul, et al., The production of butanol from jamaica bay macro algae, Environ. Prog. Sustain. Energy 31 (2012) 29 36. Available from: https://doi.org/10.1002/ep. [107] H. van der Wal, B.L.H.M. Sperber, B. Houweling-Tan, et al., Production of acetone, butanol, and ethanol from biomass of the green seaweed Ulva lactuca, Bioresour. Technol. 128 (2013) 431 437. Available from: https://doi.org/10.1016/j.biortech.2012.10.094. [108] M.H. Huesemann, L.J. Kuo, L. Urquhart, et al., Acetone-butanol fermentation of marine macroalgae, Bioresour. Technol. 108 (2012) 305 309. Available from: https://doi.org/10.1016/j.biortech.2011.12.148. [109] H. Huang, H. Liu, Y.R. Gan, Genetic modification of critical enzymes and involved genes in butanol biosynthesis from biomass, Biotechnol. Adv. 28 (2010) 651 657. Available from: https://doi.org/10.1016/j. biotechadv.2010.05.015. [110] J. Herlet, W.H. Schwarz, V.V. Zverlov, et al., Addition of β-galactosidase boosts the xyloglucan degradation capability of endoglucanase Cel9D from Clostridium thermocellum, Biotechnol. Biofuels 11 (2018) 238. Available from: https://doi.org/10.1186/s13068-018-1242-5. [111] F. Mingardon, S. Perret, A. Belaich, et al., Heterologous production, assembly, and secretion of a minicellulosome by Clostridium acetobutylicum ATCC 824, Appl. Environ. Microbiol. 71 (2005) 1215 1222. Available from: https://doi.org/10.1128/AEM.71.3.1215-1222.2005. [112] E. Garcia-Ruiz, A. Badur, C.V. Rao, H. Zhao, Characterization and engineering of seaweed degrading enzymes for biofuels and biochemicals production, in: P.C.K. Lau (Ed.), Quality Living Through Chemurgy and Green Chemistry, Springer, Berlin, Heidelberg, 2016, pp. 99 128. [113] W.G. Kidanu, P.T. Trang, H.H. Yoon, Hydrogen and volatile fatty acids production from marine macroalgae by anaerobic fermentation, Biotechnol. Bioprocess Eng. 22 (2017) 612 619. Available from: https://doi.org/10.1007/s12257-017-0258-1.

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[129] T. Wang, R. Jo´nsdo´ttir, H.G. Kristinsson, et al., Enzyme-enhanced extraction of antioxidant ingredients from red algae Palmaria palmata, LWT Food Sci. Technol. 43 (2010) 1387 1393. Available from: https://doi.org/10.1016/j.lwt.2010.05.010. [130] E. Shannon, N. Abu-Ghannam, Enzymatic extraction of fucoxanthin from brown seaweeds, Int. J. Food Sci. Technol. 53 (2018) 2195 2204. Available from: https://doi.org/10.1111/ijfs.13808. [131] R. Mittal, K.S.M.S. Raghavarao, Extraction of R-phycoerythrin from marine macro-algae, Gelidium pusillum, employing consortia of enzymes, Algal Res. 34 (2018) 1 11. Available from: https://doi.org/ 10.1016/j.algal.2018.07.002. [132] H.P.T. Nguyen, M. Moranc¸ais, J. Fleurence, J. Dumay, Mastocarpus stellatus as a source of R-phycoerythrin: optimization of enzyme assisted extraction using response surface methodology, J. Appl. Phycol. 29 (2017) 1563 1570. Available from: https://doi.org/10.1007/s10811-016-1024-z. [133] M. Saluri, M. Kaldma¨e, R. Tuvikene, Extraction and quantification of phycobiliproteins from the red alga Furcellaria lumbricalis, Algal Res. 37 (2019) 115 123. Available from: https://doi.org/10.1016/j. algal.2018.11.013.

CHAPTER

ENZYMATIC SYSTEMS FOR THE DEVELOPMENT OF JUICE CLARIFICATION STRATEGIES

18

Lokesh Kumar Narnoliya1, Jyoti Singh Jadaun2, Manisha Chownk1 and Sudhir P. Singh1 1

Center of Innovative and Applied Bioprocessing (CIAB), Mohali, Punjab, India 2Botany Department, Dayanand Girls Postgraduate College, Kanpur, Uttar Pradesh, India

18.1 INTRODUCTION The juice is a popular healthy drink, prepared by extraction of fruits and vegetables. The juice is an appetizer, thirst quencher, and refreshing agent and it is composed of water, sugars, organic acids, aroma and flavor compounds, vitamins, minerals, pectic substances, and pigments with a trace amount of proteins and fats [1,2]. In India, B76% of fruits are consumed in raw form, and only B4% is being processed [3]. Conservation of fruits is essential, but it is tedious due to their perishable nature. In the Indian context, around 20% of fruits are spoiled before their utilization. The easiest and economical way to handle this problem is fruit juice. It provides the nutritious components of fruits and vegetables throughout the year and covers an array of consumers. Nowadays, probiotics, as well as prebiotics, are also supplemented with juice, which enhances the importance of juice for human nutrition system [4,5]. The most common steps of fruit juicing are extraction, clarification, and stabilization [1,2]. The major hindrance in the marketability of any juice is its turbidity and cloudiness, which is due to the presence of pectin, cellulose, hemicelluloses, lignin, starch, protein, tannin, and metals. Juice clarification makes the juice more attractive, healthy, and increases its marketability as it removes the turbidity and cloudiness [6]. Earlier, fruit juice clarification was performed by physical
chemical adsorbents and/or filtration technology, but this approach has disadvantages as processed juice is not stable and produces pronounced haziness as well as browning [7]. The enzymatic treatment is the most effective procedure for removing haziness, turbidity, and cloudiness. The cellulases, pectinases, hemicellulases (mainly xylanases), and laccases are the key enzymes, which are applied for fruit juice clarification (Table 18.1). A combination of pectinases, cellulases, and hemicellulases is collectively known as macerating enzymes [3]. These enzymes have wide applications, and nowadays, they are also used in the management of agro-industrial waste, functional food production, bioactive molecule production, and textile industries [15,16]. Although these enzymes are present in plants, the primary source of these enzymes is microbes. These enzymes can be identified through transcriptomic, genomic, and metagenomic approach [17
21]. Pectinase is employed alone or in combination as macerating enzyme for juice clarification. The cellulases and hemicellulases are other key enzymes of juice clarification industries. Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00018-1 © 2020 Elsevier B.V. All rights reserved.

397

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CHAPTER 18 ENZYMATIC SYSTEMS FOR THE DEVELOPMENT

Table 18.1 Immobilized enzymes and their parameters used for juice clarification in the juice industry. S. No.

Enzyme immobilized

1.

Pectin methylesterase

2.

3.

Mixture of pectic enzymes Xylanase

4.

Matrix used for immobilization

Fruit juice clarified

2% sodium alginate by cross-linking with 2% CaCl2 solution Polyacrylonitrile (PAN) beads

Bael (Aegle marmelos) juice Apple juice

Pectinase

1,3,5-Triazinefunctionalized silica encapsulated magnetic nanoparticles Polyvinyl alcohol gel

5.

Xylanase

Silica

6.

Pectinase

7.

Pectinase

Celite cross-linked with glutaraldehyde PAN copolymer membrane

Apple, Pineapple and Orange juice Apple and Pomegranate Mosambi juice Pineapple juice


Optimum pH

Optimum temperature

References

3.6
4.0



40 C

[8]

4.0

50 C

[9]

6.5

60 C

[10]

4.0

45 C
50 C

[11]

7.5

55 C

[12]

5.5

45 C

[13]

5.5

60 C

[14]

Besides these, recently use of laccase for juice clarification has increased because it also significantly removes haziness, turbidity, and cloudiness [7]. Although, having remarkable advantages, the application of enzymes in juice clarification has certain limitations at industrial scale related to operational conditions such as long-term stability, difficult recovery, and reusability of the enzyme. The immobilization technology seems like a potential tool to overcome these limitations, so this is in practice at industrial scale for juice clarification. Generally, this approach improves the catalytic properties of enzyme in terms of their thermal stability and tolerance against high pH [22]. Thus, not only free enzymes but also their immobilized constructs may be workable at commercial scale for juice clarification. Immobilization of enzyme offers the reusability of biocatalyst for multiple cycles, which enhances the product yield remarkably [23]. Although there are several technologies available for juice clarification, still there is a need for an affordable process, which can be adopted at the industrial scale. CRISPR-Cas is the emerging technology, and it can be used for the development of modified enzymes, which has value at a commercial scale. CRISPR-Cas is successfully applied in various areas of health, agriculture, food, and bioengineering [24]. So, we can expect that soon CRISPR-Cas-based technology will be available for application in juice clarification industries. The regulatory systems of the critical enzymes should be known, so enzyme engineering can be performed through their regulatory factors such as transcription factor, miRNA, and lncRNA. In this chapter, we will discuss the enzymatic process of juice clarification.

18.2 WHY JUICE CLARIFICATION?

399

18.2 WHY JUICE CLARIFICATION? Juices are the extracted product of edible fruits, and unprocessed juice has haziness, turbidity, and cloudiness, which suppress its market value as the appearance of the product is the main feature of a product, which attracts customers (Fig. 18.1). Unprocessed juice contains several undesirable

FIGURE 18.1 A schematic diagram showing the juice extraction procedure from fruits.

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CHAPTER 18 ENZYMATIC SYSTEMS FOR THE DEVELOPMENT

substances, which are not compatible with its taste and nutritious value. Before its availability in the market for consumers, it is necessary to clarify any juice product to enhance its appearance and nutritive value. Juice clarification is essential because haze and sediment forming substances spoil the juice at the time of their storage [1,25]. Some other components are also present in almost all types of fruit juices, which are needed to be removed before serving to the consumer, such as proteins, polyphenols, oxidized phenolics, tannins, and metals [25]. Thus juice clarification step is a necessary and crucial step in juice industries.

18.3 JUICE CLARIFICATION STRATEGIES Several strategies are applied for juice clarification in juice industries, which include mechanical, chemical, and enzymatic approaches. Although all of the above techniques can be used up to a certain extent, enzymatic process is most commonly used and more effective technique. Here, we will discuss in detail about enzymatic approaches for juice clarification.

18.3.1 MECHANICAL PROCESS In this process, juices are clarified without any enzyme or chemical, and only mechanical methods are applied for juices clarification.

18.3.1.1 Straining or screening This is a primary process of juice clarification, and it can be applied for any juice. It consumes less time and energy. In this approach, juices are clarified mechanically by using a muslin cloth or sieve, which removes any suspended substances such as broken fruit tissue, seed, peel, large pectic substances, and protein from the colloidal suspension of juice [26]. Although this technique is not able to provide complete clarification, it makes other processes such as chemical or enzymatic treatment very easy and more efficient.

18.3.1.2 Centrifugation and finishing It is the most comfortable procedure for the separation of solid substances of juice. In this technique, solid particles of unprocessed juice are separated by centrifugal force [27]. Finishing method is used to separate pulp, seed, and rag from fruits such as citrus. A finisher contained a rotating auger inside a cylinder screen with a hole of approximately 0.020
0.030 inches in diameter. The finishing procedure depends on the condition, softness, and pulp content of citrus fruit [28].

18.3.1.3 Clarification by freezing and heating Freezing or heating are other procedures for mechanical juice clarification [29]. Generally, these methods are applied for cherry, grape, mulberry, apple, and pomegranate juice clarification [29
32]. Raw unprocessed grape juice contained tartarate along with pulp and skin, which can be removed by freezing and thawing. Heating of any juice coagulates colloidal material that settles down after cooling, and all of that settled material can be separated by filter press or centrifugation.

18.4 ENZYMES USED FOR JUICE CLARIFICATION

401

18.3.1.4 Physical finings Fruit juices can be clarified by using several fining agents such as bentonite, kaolin, diatomaceous earth, and clay. Generally, 0.5%
1% of fining agents are mixed with fruit juices and then passed through the filter press [33,34]. Ultrafiltration is also used to separate colloidal particles of juices based on their molecular weight with minimum loss of nutrients. Usually, before processing for the ultrafiltration process, the viscosity of juice should be reduced by enzymatic degradation, which produces a satisfactory clear juice [35].

18.3.2 CHEMICAL PROCESS Several chemicals are also used in the fruit juice clarification, such as gelatin, albumin, casein, tannin, and polyvinylpolypyrrolidone. Most of the fruit juices contain hydrolyzable tannins and anthocyanins, which need to be removed and gelatin is an efficient chemical used for this process. The turbidity and haze formation can also be removed by chitosan and xanthan gum in pomegranate, apple, and bayberry juices [34,36]. The charged particles of juices can be neutralized by gelatin and casein, which further generate insoluble precipitate with colloidal substances of juices [37]. The pomegranate juice is clarified using several chemicals such as protein-based (albumin, casein, and gelatin) and polysaccharide-based (chitosan and xanthan gum). Juice clarification efficiency is estimated by analyzing total phenolics, hydrolyzable tannins, anthocyanins, and antioxidant activity. In this study, it is observed that protein-based agents more efficiently reduce total phenolics, hydrolyzable tannins, and anthocyanins content in comparison to natural sedimentation and polysaccharide-based agents [36].

18.3.3 ENZYMATIC PROCESS In juice industry, application of enzymes is significant for process optimization, reducing energy cost, enhanced nutritional quality, safety, improved quality of juice, development of new products and new applications. The enzymatic method is the most widely used technique at commercial scale in the fruit juice industries because mechanical and chemical processes are unable to produce an acceptable level of clarity in the juices, which is one of the significant criteria of good quality juice. Besides this, mechanical and chemical processes have several other significant drawbacks like these processes are time-consuming, inefficient, and more laborious. Addition of fining agents may slightly change the flavor of juices, which is unacceptable for consumers. Pectinases are the most widely used enzymes for significant juice recovery as well as clarification [38]. The enzymatic degradation of colloidal substances depends on the nature of enzyme or combination of enzymes, incubation time, pH and agitation, the concentration of enzyme, and optimum temperature [3].

18.4 ENZYMES USED FOR JUICE CLARIFICATION The pectic enzymes [pectinesterase (PE), polygalacturonase (PG), and pectinlyase (PL)], cellulases, laccases, and amylases are the key enzymes used for juice clarification. Other enzymes such as

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CHAPTER 18 ENZYMATIC SYSTEMS FOR THE DEVELOPMENT

hemicellulase, xylanase, glucanase, glucosidase, amyloglucosidase, fructozyme, phenolases, dextranase, protease, and arabinase are also used for juice clarification [38]. Here, we will describe these enzymes and their application in the juice industry for juice clarification.

18.4.1 PECTIC ENZYMES Pectin, a high molecular weight complex heteropolysaccharide, is consisted of galacturonans (polymers of galacturonic acid), arabinans (polymers of arabinose), galactans (polymers of galactose), rhamnogalacturonans (mixed polymers of rhamnose and galacturonic acid), and arabinogalactans (mixed polymers of arabinose and galactose). A major portion of pectin is present in the middle lamella of the cell wall of plants and is also present in primary cell wall in a trace amount [3,39,40]. Pectin-degrading enzymes collectively known as pectic enzymes and PE, PG, and PL are the elemental enzymes of this group. Pectic enzymes efficiently degrade pectic substances to produce clarified juice. Apart from clarification, pectic enzymes are also used for juice extraction, stabilization of the juice color during storage, yield improvement, releasing flavor, proteins, etc. [3]. Nowadays, pectic enzymes cover approximately 20% of the worldwide total enzyme market. The pectic enzymes are naturally produced by bacteria, fungi, yeast, and plants [1]. Aspergillus niger and Aspergillus aculeatus fungi are the main sources of commercial pectolytic enzyme production at industrial scale. Pectinex Ultra SP-L, Pectinol, and Panzym are the commercial pectinases prepared from microorganisms for juice and food industries [41].

18.4.1.1 Pectinesterase Generally, PE has two classes, first is the pectin methylesterase and second the pectin acetylesterase. Pectin methylesterase is the chief enzyme, so it is commonly known as pectinesterase. Pectase, pectin methoxylase, pectindemethoxylase, and pectolipase are the synonyms of pectin methylesterase [3]. Pectin methylesterase belongs to hydrolase group and removes the methoxyl group from pectin. Further, de-esterified pectin is precipitated out due to complex formation with calcium, and finally, the clear juice is obtained [3].

18.4.1.2 Polygalacturonase It hydrolyzes α(l-4) bond between D-galacturonic acid units of pectin. Generally, four types of PG enzymes exist in nature based on substrate acceptance and how they hydrolyze substrate. Exosplitting PG acts on the end side, and endo-splitting hydrolyzes internal bonds of pectin substrate. A group of PG catalyzes pectin-like substrates such as poly [α(1-4)-D-methyl galacturonic acid] and another group of enzymes catalyze pectic acid-like substrates such as poly[α(1-4)-D-galacturonic acid] [3]. Yuan et al. reported the use of low-temperature active endo PG enzyme for the clarification of apple juice at pH 3.5 and a temperature of 40 C for 1 h. The resultant juice had reduced intrinsic viscosity by 4.5% and an increased transmittance of 71.8% [42].

18.4.1.3 Pectinlyase PL (trans-eliminative) performs nonhydrolytic breakdown of pectin by breaking the glycosidic linkages at two conjugated carbons and produces unsaturated products [3]. They can be endopolygalacturonate lyase, exo-polygalacturonate lyase, endo-polymethylgalacturonate, and exo-polymethyl galacturonate lyase based on the pattern of action and the substrate acted upon by them [43].

18.4 ENZYMES USED FOR JUICE CLARIFICATION

403

Generally, all the PL enzymes require Ca21 for their catalytic activity, while PG does not require any metal ion for enzymatic activity [40].

18.4.2 CELLULASE Cellulose is the structural component of the plant cell wall, and it is the most abundant constituent or organic polymer of the plant on earth [44]. Cellulose is an organic polysaccharide consisting of a linear chain of several β (1-4) glucosidic linkages of D-glucose units [38]. It is a complex polymer; therefore its breakdown is difficult and needs additional efforts. Cellulose degrading enzymes are known as cellulases, which is a group of enzymes such as exoglucanases, endoglucanase, and β-glucosidase. To obtain the complete degradation of cellulose, all the enzymes are needed to work synergistically. Exoglucanases are also known as cellobiohydrolase or exo-β-glucanase and endoglucanase known as carboxymethyl cellulase or endo-β-glucanase [38,45]. The amorphous part of cellulose is cleaved by the combined enzymatic process of endo- and exo-glucanase, and crystalline part is cleaved by β-glucosidase. The final end product of all the enzymatic processes is glucose. Normally, amorphous region of cellulose is hydrolyzed more easily than crystalline region [38]. The exoglucanase break β-1,4 glucosidic bonds from the end region of cellulose and endoglucanase cleave internal β-1,4 glucosidic bonds, while β-glucosidase releases glucose from cellobiose and short cello-oligosaccharides. Cellulase enzyme is lacking in animals, including humans; therefore they are unable to digest plant materials. Cellulase enzyme has wide applications in various areas such as the textile industry, laundry detergents, oil extraction, pulp and paper industry, biofuel production, and biomass degradation. It plays a crucial role in the processing of food and juices at the industrial scale. It helps in the production of oligosaccharides and low-calorie food substituents, which are used in functional food ingredients [16,46]. After pectin, cellulose is second in this list of turbidity and haziness-producing agents in juices. Therefore, cellulases play a vital role in the area of fruits and vegetable juices clarification. Predominantly, fungi such as A. niger, Aspergillus nidulans, and Aspergillus oryzae are the main sources of commercial microbial cellulases [3,47].

18.4.3 LACCASE Ligninolytic enzyme laccase is a polyphenol oxidase enzyme (p-diphenol: dioxygen oxidoreductases; EC 1.10.3.2) that oxidizes various phenolics and nonphenolic lignin compounds using molecular oxygen as the electron acceptor [48]. Laccase involves in lignin degradation along with lignin peroxidases, manganese peroxidases (MnPs), and some other peroxidases. In recent years, they have gained attention due to their environmental pollutant degrading capacities, although they are highly recalcitrant [21]. In addition, a novel role is dedicated to them as fruit juice-clearing agents, and that is why the food industry is paying attention to these enzymes [49]. They widely occur in nature, but fungal laccases (e.g., white-rot fungus) are considered to be of higher significance due to their ability to act on wooden components [50]. Laccases are designed to degrade lignin component of woods so that the other components of wood such as cellulose and hemicellulose are easily accessible to enzymatic degradation. They have applications in various industries due to their low substrate specificity and laccases accept

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CHAPTER 18 ENZYMATIC SYSTEMS FOR THE DEVELOPMENT

polyaromatic hydrocarbons, xenobiotic compounds, and textile dyes as their substrates. They require only oxygen and substrate to carry out the reaction [51]. The polyphenols and proteins react to form a haze, which is quite undesirable and negatively affects the commercial value of the juice. Although there are several chemical
physical processes are available to avoid that, the use of enzymes is always preferred for their economical, environment-friendly, and simple approach [52]. Juice processing industries have been using the enzyme-mediated clarification of fruit juices for many years. There are many reports available, which describe that the use of laccases improves the color and flavor stability of the fruit juices. Laccase was used by Maier and Dietrich for the clarification of apple juice, and they found that it could be worked along with ultrafiltration without the addition of finishing agents [52]. Also, similar results were reported by Stutz, where the juice was clarified by laccase, which resulted in its light color and stability [53]. It is also reported that the addition of ascorbic acid and sulfites along with laccase enzyme to the juice increased the flavor stability as well as the color [54]. The clarity and browning of apple juice were improved by Pseudomonas putida LUA15.1 in a similar study, which also resulted in the enhanced sensory profile of the juice [55]. The capacity of laccase to dehaze the pomegranate juice was applied in a study along with ultrafiltration; laccase was able to reduce the phenol content of the juice by 40% [56]. A fungus Abortiporus biennis isolated from soil was characterized to produce laccase with properties to be thermo- and pH stable. It was used to reduce the phenolic contents in litchi juice [57]. In a recent report, a laccase enzyme from Bacillus atrophaeus (BaLc) was isolated and physico-kinetically characterized. Further, this BaLc enzyme was immobilized on magnetic nanoparticles and applied for banana pseudo-stem, sorghum stem, and apple fruit juice clarification. The immobilized BaLc enzyme retained 60% of the residual activity after 10 consecutive reaction cycles of 2,20 -azino-bis(3-ethylbenzothiazoline-6sulfonic acid) oxidation. The free and immobilized biocatalyst, provide 41%
58% of phenol reduction, 41%
58% decolorization, 50%
59% turbidity reduction in banana pseudo-stem and sweet sorghum stalk and apple fruit juices [21]. In recent years, the enzyme has been immobilized and applied in food industries for its enhanced juice-clearing properties (Fig. 18.2). In another study, laccase was immobilized on coconut fibers activated with glutaraldehyde. The enzyme from Tinea versicolor was immobilized, and it was used for apple juice clarification, which resulted in the lightning of juice color by 61% 6 1% and clarified the juice by 29% 6 1% [58]. In another approach, POXA1b laccase from Pleurotus ostreatus was immobilized on epoxy-activated poly(methacrylate) beads, which had immobilization efficiency of 98%. The final product was then used for juice clarification, which led to a reduction of 45% phenol as well as the vinyl guaiacol content without affecting the flavanone content [7]. However, there are several reports, which suggest that the use of immobilized laccase enzyme for the clarification of fruit juices negatively affected the color of the juice. It led to more browning upon storage as compared to the conventional methods [54,59]. Sammartino et al. suggested that the use of immobilized/free enzyme for the clarification of apple juice led to an unstable product than the conventionally treated juices (SO2, metabisulfite, bentonite) [60]. Many reports revealed that the use of ultrafiltration in addition to laccase for the dehazing purpose could solve the storage problem altogether [52,61]. In a nutshell, using laccase enzyme in conjugation with ultrafiltration can be beneficial than conventional methods, although different levels of standardizations are required.

18.4 ENZYMES USED FOR JUICE CLARIFICATION

405

FIGURE 18.2 An illustration of enzymatic methods applicable for juice clarification process. (A) Free enzymes, (B) immobilized enzymes, and (C) coimmobilized enzymes.

18.4.4 AMYLASE The 25% coverage of the total enzyme market by amylase shows how significant amylases are at the industrial level. The α-amylases are a class of enzymes, which catalyze the hydrolysis of internal α-1,4-glycosidic linkages in starch, leading to the formation of small molecules such as glucose, maltose, and maltotriose units [62]. Amylases are such efficient catalysts that they have almost completely replaced the chemical processes for hydrolyzing starch in industries. There are many sources of amylases, including animals, plants, and microorganisms, but amylases of microbial origin (fungi, bacteria, yeasts) are always preferred as they are easy to manipulate and are readily available for easier production in large quantities [63]. Amylases have applications in various industries, which include food, paper, textile, pharmaceutical, fermentation, detergent, brewing, and distilling industries [63,64].

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For the purpose of industrial applications, many studies have reported thermostable amylases. The most thermostable amylase enzyme was isolated from Bacillus licheniformis, which is still used in many industrial processes. The enzyme is stable at 90 C for several hours. It is widely used in the bakery industry due to being maltogenic in nature [62]. For the conversion of starch to fructose and glucose syrup, α-amylase from Bacillus stearothermophilus or B. licheniformis has been used widely in recent years. Over 90% of liquid detergents contain amylase enzyme isolated either from Bacillus or Aspergillus. Another amylase from Pyrococcus woesei is stable in a wide temperature range from 40 C to 130 C at pH 5.5 [65]. Besides these, many sources of amylase have been isolated in recent years due to their potential applications in various processes. A thermostable amylase was isolated from Bacillus sp. CICIM 304. This was stable in the range of 30 C
70 C and had the optimal temperature at 70 C in alkaline pH. This enzyme was stable over a wide range of pH 5.5
9.0 [66]. Another thermotolerant amylase has been reported to be produced by Geobacillus sp. The bacterium was isolated from hot waters of geothermal springs. The enzyme isolated from Geobacillus sp. was stable at alkaline pH and temperature of 90 C. The purified fraction of the protein was obtained by diethylaminoethyl-cellulose column and Sephadex G-150 gel filtration chromatography [67]. Besides these, E. coli and Bacillus amylases have also been used in fuel production where starch is converted into fermentable sugars, which are further converted into alcohol by yeast [68]. The α-amylase produced by Aspergillus niger is extensively used in commercial processes such as manufacturing of soy sauce, organic acids such as citric and acetic acids. Fungal amylases are preferred in the food industry because of their generally regarded as safe status [69]. Amylases have been used in fruit juice clarification for quite a long time now. There are various reports available using only amylases or in conjugation with other enzymes (pectinases, xylanase, etc.) for this purpose. A report suggested the use of amylase and pectinase mixture (0.02% and 0.084%) for the clarification of banana juice at acidic pH of 4 and different process times for pectinase and amylase (60 C; 1 h for amylase and 43.2 C for 80 min for pectinase). The resultant juice had the clarity of 0.006 Abs, the viscosity of 1.89 cps, and turbidity of 0.92 NTU [70]. Another study revealed the use of equal quantities of amylase and pectinase (0.025%) for the clarification of kiwi fruit juice at an acidic pH of 3.52 and the temperature of 50 C for 2 h. The resultant juice was reported to be clear with less haziness as compared to control samples [71]. A similar study was conducted by Kothari et al. where they used an enzyme mixture of pectinase, amylase, and cellulase and studied the clarification of apple juice by measuring percent clarity and reducing sugars produced. The study showed improved clarity of juice as compared to juice treated with independent enzymes. Another study reported the use of PG and amylase to clarify the apple juice [72]. The pectinase and amylase enzymes were isolated from Aspergillus awamori by using apple pomace, and mosambi orange (Citrus sinensis var. mosambi) peels as solid substrate and A. oryzae, respectively, using wheat bran by solid-state fermentation. Both enzymes were used in combination to clarify apple juice. In the presence of 1% PG and 4% α-amylase, a maximum clarity of 97% was achieved after 2 h of incubation at 50 C in the presence of 10 mM calcium chloride. The total phenolic content of the apple juice was found to be reduced by 19.8%, and higher radical scavenging property was observed [73]. In another report, effects of pH, heat, and calcium ions on the amylase activities were observed using commercial enzymes for the apple juice clarification. It was found that in the presence of citrate ions, the enzyme activity was retained even after heat treatment. After the enzyme was treated to 60 C and 65 C, they regain their activity by the addition of

18.5 CONCLUSIONS AND PERSPECTIVES

407

calcium chloride in the media. The possible implications of these studies are in the apple juice processing industries where higher temperature is required for processing [74]. A study reported the use of an α-amylase of fungus (Rhizopus stolonifer) for Soursop juice clarification. The final results showed that juice treated with a low concentration of extracted enzyme and less incubation time was more viscous as compared with the juice containing a high level of amylase and higher incubation time [75]. In a similar study, mushroom compost was used to isolate Bacillus subtilis M13, which was then further used to extract α-amylase. The enzyme worked at a pH 9 and 45 C temperature, which was used to increase 60% and 55% of apple and kiwi juice clarity with enhanced color, taste, flavor, and overall acceptability, respectively [76].

18.4.5 OTHERS There are many other enzymes used for juice clarification purposes, which include xylanases, pullulanase, etc. but they are not widely used as compared to abovementioned enzymes [38,77]. In a study reported by Nagar et al., xylanase was isolated from Bacillus pumilus SV-85S, characterized in detail, and was further used to demonstrate its juice-clearing properties. The purified enzyme was used to treat apple, pineapple, and tomato juice, and it was found that treatment led to an increase in the juice yield by 25%, 10%, and 20%, respectively. The increase in the transmittance was found to be 22%, 19%, and 14%, respectively [77]. One report suggested the use of 0.1% pectinase along with β-galactosidase, chitinase, and transgalactosidase for the purpose of Carambola juice clarification even after treatment for only 30 min at pH 4, which resulted in the maximum clarity and minimum turbidity and absorbance [78]. Sapodilla juice was treated with 0.1% crude pectinase in combination with cellulase and hemicellulose at pH 4.7 and temperature of 40 C for 2 h where the final product showed the lowest turbidity and low viscosity with higher transmittance [79].

18.5 CONCLUSIONS AND PERSPECTIVES Enzymes became an indispensable tool in almost every industry in the past few years. The biological nature of enzymes, along with secure handling, manipulation, low-cost features, allowed researchers to expand their application fields. Fruit juice industries earlier relied on chemical processes for the clarification purposes, but the introduction of highly efficient enzymes led to faster, economical, environment-friendly processes. Enzymes such as pectinase, amylase, laccase, PG, and xylanase have proved their efficiency in lowering turbidity, absorbance and improving transmittance, flavor, and overall acceptability. However, the type of enzyme to be used depends on many factors such as the type of substrate, pH, and temperature, and each process requires optimization to get the best result. The use of immobilized enzymes has provided a boost to the fruit juice industries. Many routes are available to immobilize enzymes, and cellulose fibers are already used for this purpose. The combinations of two or more enzymes in a single reaction have also been used to dehaze the juice. However, the immobilization of two or more enzymes on a single matrix can lead to a better biocatalyst, which can be used on multiple substrates. Although reaction conditions have to be

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standardized; eventually, the final product would be highly efficient, economical, and user-friendly. Companies such as biocatalysts have been using enzyme products such as Pectinase 831L, Pectinase 947L, amylase 826MDP, and Pectinase 743L extensively and are paving the way for the enzyme products manufacturing companies and researchers alike. Despite, having several juice clarification technologies through an enzymatic process, there is still a need to improve these enzymes, which can be significantly used at commercial scale.

ACKNOWLEDGMENTS The authors acknowledge the Department of Biotechnology (DBT), Government of India. LKN and MC acknowledge Science and Engineering Research Board (SERB) N-PDF fellowships, PDF/2015/000662 and PDF/2016/002550, respectively.

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CHAPTER

BIOCATALYST SYSTEMS FOR XYLOOLIGOSACCHARIDES PRODUCTION FROM LIGNOCELLULOSIC BIOMASS AND THEIR USES

19

Michele Michelin and Jose´ A. Teixeira CEB—Centre of Biological Engineering, University of Minho, Campus Gualtar, Braga, Portugal

19.1 INTRODUCTION Industrial processes and agricultural practices generate a huge amount of lignocellulosic materials (LCM) that are considered as “waste,” for example, sugarcane bagasse (SCB), corn stover, corncob, wheat straw, etc. These materials are often left in the field or burned for energy generation in some industries. The reuse of these materials not only avoids some environmental concerns caused by their inappropriate disposal, but also provides an additional value to these residues. LCM are rich in lignocellulose, generally made of 30% 50% cellulose, 15% 35% hemicellulose, and 10% 20% lignin. The presence of these components gives LCM a very significant biotechnological value [1,2]. Currently, the use of LCM has been very explored, due to future needs in fields of energy, chemicals, and food production. Thus, it is important to develop technologies that explore the utilization of the underutilized components in LCM, such as hemicelluloses [3]. Xylan is one of the important hemicelluloses of LCM. The conversion of xylan into valueadded products has been shown quite promising for the exploration of a variety of agricultural residues. Different xylan-rich LCM are being explored for the possibility to use them for the generation of XOS. Additionally, the availability of hemicellulose as a by-product of second-generation ethanol processing can further stimulate their production [4,5]. To date, a great number of LCM were evaluated for XOS production, such as SCB [4,6 9], cotton stalk [10], corn stover [11,12], corncob [1,11,13 15], cotton stalk [10], rice husks [16,17], wheat bran [18], and brewer’s spent grain [19,20].

19.2 XYLAN: SOURCES AND STRUCTURES Xylan is the main constituent of hemicelluloses, which together with cellulose constitute the major components of LCM. Although xylan comprises up to 50% of some grasses and cereal grains Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00019-3 © 2020 Elsevier B.V. All rights reserved.

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biomass, the main focus has been on agricultural crop residues, which are an abundant, inexpensive, and renewable source of lignocellulose in nature [21]. It is found in large amounts in hardwoods (15% 30% of the cell wall) and softwoods (7% 10%), as well as in annual plants (,30%) [22]. Xylan consists of a linear polymer of β-D-xylose units (nearly 90%) linked by β-1,4-glycosidic linkages, substituted with different groups, such as arabinofuranosyl, glucuronopyranosyl, acetyl, feruloyl, and/or p-coumaroyl residues. The frequency of these substituents varies according to the source of feedstocks [2,22,23] and results in XOS with varied biological properties [24]. In nature, hemicelluloses are usually found as complex structures made of more than one polymer. The most common are glucuronoxylans and arabinoglucuronoxylans, as well as glucomannans, arabinogalactans, and galactoglucomannans [23,25,26]. Wood xylans comprise arabino-4-O-methylglucuronoxylan in softwoods and O-acetyl-4-O-methylglucuronoxylan in hardwoods, while xylans in annual plants and in grasses are normally arabinoxylans [22]. Xylan from LCM has been used to obtain different value-added products, including XOS.

19.3 XYLOOLIGOSACCHARIDES XOS are sugar oligomers, obtained from hydrolysis of xylan, composed of xylose units [1,27,28]. The molecular formula of XOS is C5nH8n12O4n11 being n 5 2 6 and they are known as xylobiose, xylotriose, xylotetrose, and so on [21,29]. Furthermore, according to some authors, oligomers with xylose degree of polymerization (DP) # 20 can also be considered XOS [30]. They are considered as nondigestible food ingredients, and only XOS with xylose DP 2 4 present potential as ingredient for functional foods [4,27,31]. XOS can be naturally found in various food products, such as honey, fruit, milk, and vegetables [27], or can be obtained through the hydrolysis of xylans, commonly found in LCM, such as corncob, SCB, wheat straw, and others [21,29]. The structural features of XOS depend not only on the origin of xylan but also on the production process [8,32]. XOS present many functional properties, besides nontoxicity and not metabolized by human digestive system [33,34].

19.4 PRODUCTION OF XYLOOLIGOSACCHARIDES XOS can be produced by various methods from xylan from LCM. These methods involve autohydrolysis of LCM at high temperature and pressure, chemical methods (e.g., acid hydrolysis or alkali extraction), and enzymatic hydrolysis of a susceptible substrate or a combination of chemical and enzymatic methods [21]. XOS have been mainly produced by acid hydrolysis from LCM. However, this process is considered environmentally unfriendly and expensive due to the purification steps involved. Other approaches to XOS production include transglycosylation reactions via glycosyltranferases and glycosynthase β-xylosidases. However, the need for pure monosaccharides as raw materials makes them expensive and unviable [35,36]. XOS production through enzymatic method is a promising strategy due to the high specificity of xylanase enzymes to the xylan substrate. In general, the enzymatic method consists in the

19.4 PRODUCTION OF XYLOOLIGOSACCHARIDES

415

employment of xylanolytic enzymes, mainly endoxylanase, to depolymerize xylan and produce XOS [37,38]. However, XOS production through direct enzymatic hydrolysis is suitable only for susceptible xylan-containing materials, such as citrus peels [39]. Thus the pretreatment of the LCM before the enzymatic hydrolysis has shown to be a fundamental step for surpassing structural and steric barriers (e.g., lignin presence), improving the access of enzyme and allowing a more efficient hydrolysis [40]. In addition, an efficient pretreatment can lower downstream pressure by making LCM more enzymatically accessible [41]. Fig. 19.1 presents a schematic representation of XOS production from agricultural crop residues through a combination of chemical and enzymatic methods. Therefore, XOS production has been explored in two stages: xylan extraction from LCM followed by enzymatic hydrolysis [7,10]. Currently, several types of pretreatment (including physicochemical, chemical, biological, or a combination of various processes) have been studied for xylan

FIGURE 19.1 Schematic representation of xylooligosaccharides (XOS) production from lignocellulosic materials through a combination of chemical and enzymatic methods

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extraction prior to the enzymatic hydrolysis for the purpose of improving the enzyme accessibility and digestibility. The choice of the pretreatment used for LCM must be cost-effective with minimal power, heat, and chemical requirements and must also be environmentally friendly [1,42]. Xylan is usually extracted from LCM through alkali pretreatment, such as NaOH or KOH, and then it is converted to XOS by the action of endoxylanase enzyme having little or no exoxylanase activity, such as β-xylosidase [10]. The use of xylanases with little or no β-xylosidase activity is preferred, aiming to minimize or avoid the formation of xylose in the XOS mixture [8,27,41]. Unlike autohydrolysis, this method is preferred as it does not produce undesirable by-products in the XOS mixture, such as furfural (due to low reaction temperature) or high quantity of monosaccharides (xylose), and does not require special apparatus. Thus high-purity XOS are produced, that are easily recovered, which reduces the downstream processing costs [1,10,36 38]. Table 19.1 presents some works that describe the XOS production by enzymatic hydrolysis of xylan. In the enzymatic production of XOS, free or immobilized xylanases can be used directly

Table 19.1 Production of XOS through enzymatic process using lignocellulosic biomass as a xylan source. Xylan source CC SCB

SCB SCB Mahogany and MWS CC WB, SCB, BS, and RH CC

SF

XOS production

DP

References

Partially purified xylanase from Aspergillus foetidus MTCC 4898 Recombinant endo-1,4-β-D-xylanase A from Bacillus subtilis strain 168

116.64 mg/g

2 5

[1]

119 mg/g

Mainly 3 5

[4]

Recombinant endo-β-1,4-xylanase in Pichia pastoris β-xylosidase-free endoxylanase from B. subtilis KCX006 Purified xylanase from Clostridium strain BOH3

59.6 mg/g

2 3

[9]

94.95 mg/g

2 4

[38]

89.5 and 67.6 mg/g, respectively 32.8 mg/g

2 5

[43]

Mainly 2 3 2 and 4

[44]

196 mg/g

2 4

[46]

288.36 mg/g

2 4

[47]

Pretreatment

Xylanase source

Mild alkali treatment Hydrogen peroxide and acetic acid Alkali Aqueous ammonia Mild alkali conditions Ultrahigh pressure Alkaline extraction

Acidic electrolyzed water Not performed

Endoxylanase from Streptomyces thermovulgaris TISTR1948 Crude extract from B. subtilis KCX006 containing endoxylanase and debranching α-L-arabinofuranosidase and esterase Purified recombinant xylanase (Xyn10CD18) Xylanase and arabinofuranosidase from genetically modified strains of Aspergillus nidulans A773

Max 665.2 mg/g from SCB

[45]

CC, Corncob; WB, wheat bran; SCB, sugarcane bagasse; BS, bamboo shoots; RH, rice husk; SF, soybean fiber; MWS, mango wood sawdust; XOS, xylooligosaccharides; DP, degree of polymerization.

19.4 PRODUCTION OF XYLOOLIGOSACCHARIDES

417

into the reaction medium [30]. Several factors affect XOS yield from xylan, such as xylan source, enzyme activity, hydrolysis time, and temperature and pH of incubation [21].

19.4.1 XYLANOLYTIC ENZYMES Complete hydrolysis of xylan is attributed to several different classes of enzymes like endo-, exoand debranching enzymes, that act synergistically. Xylanases are produced by fungi, bacteria, marine algae, yeast, insect, seeds, protozoans, crustaceans, snails, etc.; however, filamentous fungi are the main commercial sources [23]. Xylanase (EC 3.2.1.8) is the main endoxylanase that cleaves the internal glycosidic linkages in the xylan backbone, producing initially XOS, and afterward, smaller molecules such as xylotriose, xylobiose and xylose may be produced. Xylan is not hydrolyzed randomly, the linkages that will be hydrolyzed depend on the nature of the substrate, for example, the branching degree, chain length, and the presence of substituents [23,48]. Thus, depending on the source of xylan used for the production of XOS, the XOS structure varies in DP, monomeric units, and types of linkages [24]. The use of endoxylanases with low exoxylanase and/or β-glucosidase activity is preferred, as commented previously. Exoxylanases hydrolyze long-chain xylo-oligomers from the reducing end to produce short-chain xylo-oligomers and afterwards xylose [49], and β-xylosidase (EC 3.2.1.37), an exoxylanase, hydrolyzes short-chain XOS and xylobiose produced by the action of endoxylanase and release free D-xylose [23,48]. The degree of hydrolysis of xylan from LCM by endoxylanase has been shown to be increased by synergistic action of debranching enzymes such as α-L-arabinofuranosidase, α-glucuronidases, esterases, and β-D-glucosidase [37,50], since the xylan substituents sterically hinder the action of most endoxylanases [51]. α-L-Arabinofuranosidases (EC 3.2.1.55) remove substituted arabinose residues at positions 2 and 3 of the xylan backbone. This enzyme jointly with arabinanases (EC 3.2.1.99) hydrolyzes side chains of arabinan linked to the xylan backbone to arabinose [48,52,53]. α-Glucuronidases (EC 3.2.1.139) hydrolyze the α-1,2-glycosidic linkages of the 4-O-methylglucuronic or D-glucuronic acid side chain from the xylan backbone in glucuronoxylan [48,54]. Regarding the esterases, acetyl xylan esterases (EC 3.1.1.72) remove O-acetyl substituents from C2 and/or C3 position of xylose residues in acetyl xylan. Feruloyl esterases (EC 3.1.1.73) and pcoumaric acid esterases (EC 3.1.1.) hydrolyze ester linkages in xylan, between the arabinose side chains and ferulic acid or p-coumaric acid residues, respectively [23,48,53,54]. Glucuronoyl esterases (EC 3.1.1.-) hydrolyze ester linkages between D-glucuronic or 4-O-methyl-D-glucuronic acid residues of glucuronoxylans [48,55].

19.4.2 PURIFICATION OF XYLOOLIGOSACCHARIDES After XOS production, the most important step is the purification process, where the high molecular weight oligomers and the sugars of low molecular weight that are not of interest are separated from the target product. For this goal, vacuum evaporation, solvent extraction, adsorption, or chromatographic methods can be used [10,28,41]. However, these methods are considered impractical and economically inviable for production of XOS in large scale or attain low purity level.

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Membrane technologies such as ultrafiltration and nanofiltration have shown to be an attractive alternative for refining and concentrating XOS. They have been used successfully for processing XOS [10,56 58] due to their low energy requirements, their relatively easy scale-up, and easy manipulation of operational variables. For example, Apikar et al. [10] used ultrafiltration process to purify XOS produced through enzymatic hydrolysis. For that, membranes with different molecular weight cut-off (1, 3, and 10 kDa) were used. Complete removal of unhydrolyzed xylan and xylanase was achieved without losing any oligosaccharides with DP 1 5 using 10 kDa membrane. Yuan et al. [56] used membranes of nanofiltration to concentrate XOS obtained through enzymatic hydrolysis of xylan from steam-pretreated corncob. However, several steps of purification may be required to obtain high-purity XOS [28].

19.5 BIOLOGICAL PROPERTIES OF XYLOOLIGOSACCHARIDES The importance of XOS as ingredient for functional foods or pharmaceuticals is increasing in the current scenario as it presents a variety of health benefits. XOS have been recognized as prebiotics that have the ability to stimulate the growth and proliferation of a selective group of beneficial bacteria, such as Bifidobacteria and Lactobacilli, in the human intestine. The growth of these probiotic bacteria prevents the growth of harmful bacteria [37,59], decreasing the risk of many colonic diseases such as inflammatory bowel disease, acute ulcerative colitis, and Crohn’s disease and keeping the intestinal microflora healthy [60,61]. They modulate the immune system of the host by increasing the production of antimicrobial substances to equilibrate the microbiota of the intestine [3,60,62]. Regarding its efficacy in improving the gastrointestinal health, XOS are reported to be better than fructooligosaccharides [63]. XOS, as nondigestible oligosaccharides, remain stable in acidic media at high temperature [21,28,30]. Because of this acid stability, XOS are not degraded by the acids of the stomach or absorbed in the small intestine. Thus, they go to the large intestine colon, where they are fermented by the beneficial bacteria to short-chain fatty acids (SCFAs), such as acetate, lactate, butyrate, and propionate. SCFA can modulate some aspects of metabolic activity, such as bowel homeostasis, colonocyte function, renal physiology, the immune system, blood lipids, energy gain, and appetite [64]. These compounds are crucial for intestinal health, contributing to lower the pH (protecting against acid-sensitive enteropathogens) and improving calcium absorption, and may be related to the prevention of intestinal infection, reducing the risk of colon cancer and improving the intestinal health [8,28,30,65 68]. For example, Reddy and Krishnan [38] reported the prebiotic effect of XOS mixture produced from SCB for the growth of three strains of bifidobacteria, namely Bifidobacterium infantis, Bifidobacterium breve, and Bifidobacterium bifidum. All strains produced a mixture of SCFAs in different proportions. Around 40 mM total SCFAs were produced by the strains from XOS mixture, being the main SCFA: acetate, formate and small amounts of lactate. Therefore, this work showed that XOS obtained from SCB present potential as prebiotics for the proliferation of bifidobacteria. XOS obtained from pretreated sawdust also showed a great prebiotic effect on Lactobacilli and Bifidobacteria [43].

19.6 XYLOOLIGOSACCHARIDES APPLICATIONS

419

Several studies have reported the health benefits of using XOS in maintaining the gastrointestinal health, due to the selective growth of beneficial colon microbiota, which include increased absorption of minerals from the large intestine and lipid metabolism, as commented previously, reduction of procarcinogenic enzyme in the gastrointestinal tract, the reduction of blood cholesterol levels, protection against cardiovascular disorders, regulation of insulin secretion from the pancreas, antioxidant activity, and antimicrobial, antiallergy, antiinfection and antiinflammatory properties [1,8,21,27,28,30,68 72]. Moreover, these bioactive compounds are believed to relieve symptoms of type 2 diabetes mellitus, cancer and stress [21,35,36,73 75]. Regarding the antioxidant properties, Rashad et al. [36] produced XOS mixtures from different agricultural residues pretreated by alkali (corncobs, SCB, rice straw, wheat bran, orange peels, mango peels, sawdust, and water hyacinth plant) by using crude xylanase from Bacillus amyloliquefaciens NRRL B-14393. XOS mixtures from orange peels presented 96% of antioxidant activity, while the total phenolic content (TPC) was 156.32 mg GAE/L extract. XOS mixtures from mango peels also presented high antioxidant activity (76.84%), as well as TPC (133,74 mg GAE/L extract), both at 1000 μg/mL concentration. The antioxidant activities and TPC presented by the other XOS mixtures varied between 10% and 20% and 11.59 49.72 mg GAE/L extract, at various concentrations. The high amount of TPC in XOS mixtures from orange and mango peels may explain the relatively higher antioxidant activities in these XOS mixtures compared to the others. Thus, xylanase from B. amyloliquefaciens NRRL B-14393 presents a high potential for production of XOS with high phenolic content, as well as antioxidant activity. Clinical tests have reported no adverse effects of XOS [76,77]. The health benefits of XOS are mainly related to their action on the gastrointestinal flora [27]. XOS also present application in pharmaceutics, as active components of symbiotic preparations [8,9,78].

19.6 XYLOOLIGOSACCHARIDES APPLICATIONS The most important application of XOS is related to their use as ingredients for functional foods. It can be blended in drinks and food, such as milk, soft drinks, tea or cocoa drinks, nutritive preparations, yogurts, fermented dairy products, candies, cakes, biscuits, jellies, jam and honey products, and special health food preparations for elder people and children [28]. There is growing studies showing that some ingredients for functional foods can present a beneficial impact on several intestinal-related diseases and lifestyle and age-related dysfunctions [24]. The growing search for these foods can be justified by the high health care costs, the increased life expectancy, and the desire of the elderly to improve their life quality [79,80]. XOS present acceptable organoleptic property [33] and are valuable, low-calorie food sweetener with prebiotic, noncarcinogenic, and antifreezing properties, that can be used as additive in drinks and food products [19,28,36,81]. XOS with DP within range of 2 4 are preferred for use as food ingredients. Xylobiose (DP 5 2) is considered an XOS, in terms of food applications, because its sweetness corresponds to 30% that of sucrose and it has no flavor [27]. Moderate sweetening power and the wide pH and temperature stability, as well as inhibitory properties against starch retrogradation, make that XOS-added foods have improved nutritional and sensory properties [7,10,82].

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Besides food applications, XOS can be used as ingredients in pharmaceuticals, products for diabetics, feed formulations, or agricultural products.

19.7 MARKET AND SAFETY ASPECTS OF XYLOOLIGOSACCHARIDES A recent analysis performed by the Market Study Reports [83] shows that the application spectrum of the XOS market involves medicine and health products, food and drinks, feed, and others. Application of XOS in food products, as prebiotics and fortifying agent, is because of their properties, described previously, such as high stability, resistance to stomach acid pH and digestion and absorption in the upper gastrointestinal tract, fermentation to SCFA by beneficial bacteria in colon, and acceptable organoleptic properties [38]. Recently, a mixture of XOS as a novel food (NF) has been approved by the EFSA Panel on Dietetic Products, Nutrition and Allergies (NDA), for use by the general population. The NF is obtained from corncob through enzymatic hydrolysis and subsequent purification. The oligosaccharides contained in NF are resistant to human digestive enzymes and are fermented by beneficial bacteria from colon. The proposal is to blend it to a diversity of foods such as fruit jelly, chocolates, bakery and dairy products, and soy drinks. Safety of XOS as NF is in accordance with Regulation (EU) 2015/2283 [84]. An example of commercially available XOS is the PreticX, a non-GMO corncob-derived XOS prebiotic from AIDP Inc., which is manufactured by LongLive and has previously achieved FDA’s GRAS status [85]. Longlive, together with Kangwei, HFsugar, Henan Shengtai, YIBIN YATAI, HBTX, YuHua, ShunTian are between the main manufacturers of the global XOS market [83]. Consumers can purchase PreticX prebiotics as health food supplements on nutritional websites for Bh12.5 22 per 100 g of product. The worldwide market for XOS is expected to reach 130 million US$ in 2025, from 94 million US$ in 2018, at a CAGR (Compound Annual Growth Rate) of roughly 4.1% during the forecast period, according to a new study [83].

19.8 CONCLUSION AND PERSPECTIVES XOS have attracted the attention of the researchers and consumers due to their great health benefits. They can be obtained through xylan sources, such as LCM, through different methods, including enzymatic hydrolysis, using xylanolytic enzymes. Thus, researches involving the isolation of new microorganism producers of xylanases, the development of efficient and viable strategies to pretreat LCM to recover xylan, and the use of these xylanases in the extracted xylan for XOS production are a fundamental approach for functional food field. Besides, research to make them available at a affordable price to a large number of consumers is a challenge. Therefore, each step of the process for their production that contributes to the total price needs to be evaluated. In this sense, approaches such as the XOS production from widely available agricultural crop residues which are rich in hemicellulose has been explored, due to the high cost of the commercial xylan.

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CHAPTER

BIOCATALYSIS FOR CASCADE REACTIONS TO PRODUCE HIGH-VALUE CHEMICALS

20

Ji Liu, Kaiyuan Tian and Zhi Li Department of Chemical & Biomolecular Engineering, National University of Singapore, Singapore

20.1 INTRODUCTION 20.1.1 WHAT IS A CASCADE REACTION? For a multistep chemical synthesis, individual reactions are often carried out in a stepwise manner. The product from the previous step is isolated and purified from the reaction mixture before being used for the subsequent reaction in a different reaction vessel (Fig. 20.1). Such a process is widely used in chemical synthesis due to incompatible reaction conditions between steps and poor catalytic specificity and selectivity of chemical catalysts. Some chemical reactions, such as reduction and oxidation, require extreme and somehow opposite conditions. In comparison, a cascade reaction performs two or more consecutive reactions in single reaction vessel. The intermediates from previous step are used directly for the subsequent reactions without further isolation. Such a process is often employed for biocatalytic reactions as biocatalysts often have good substrate specificity, chemo- and stereoselectivity and share similar reaction conditions.

20.1.2 ADVANTAGES OF CASCADE BIOCATALYSIS Unlike a stepwise workflow, a cascade reaction performs multiple compatible reaction steps in a single reaction compartment; thus it avoids the isolation and purification of synthetic intermediates and greatly simplifies the operational procedure, minimizing the production cost, use of toxic reagents, waste generation, and material loss. Cascade reactions may have less issues with mass transfer as catalysts are in close proximity to each other, enabling better accessibility to the reaction intermediates. Such features have attracted tremendous interest in both academia and industry as a green and sustainable substitution to some problematic chemical syntheses. Biocatalytic cascades also benefit from the excellent chemo-, regio- and stereoselectivity of biocatalysts. This offers an intriguing approach to selectively handle multiple chemically similar functional groups or mixtures of stereoisomers. A simple installation of the biocatalyst could replace its chemical counterpart that relies on cumbersome protection-deprotection procedures or sophisticated/expensive metal catalyst [1]. Compared with single-step biocatalysis, conducting cascade reactions may also provide engineering solutions to enzyme inhibition by the reaction intermediate Biomass, Biofuels, Biochemicals. DOI: https://doi.org/10.1016/B978-0-12-819820-9.00020-X © 2020 Elsevier B.V. All rights reserved.

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FIGURE 20.1 Schematic illustration of stepwise synthesis and cascade reaction. S, Substrate; P, product; C, catalyst.

(s), self-sufficient cofactor recycling, and minimize the accumulation of unstable intermediate and byproduct formation [2,3], providing much improved synthetic efficiency.

20.1.3 PRODUCTION OF HIGH-VALUE CHEMICALS VIA CASCADE BIOCATALYSIS On the flip side, biocatalysts typically have high preparation cost. To make biocatalytic cascade production of chemicals economically sensible, the focus of the field and therefore this review is on the production of valuable chemicals such as natural products and pharmaceutical molecules. A cascade that shows broad substrate scope is also considered in this chapter as it is meaningful for practical applications such as drug discovery and development, which aims at diversity-oriented synthesis to explore potential chemical space. Last but not least, the preparation of commodity or platform chemicals from nature-sourced biomass or waste materials offers alternatives to contemporary petrochemical production and great potential for more green and sustainable production in the future. In the past decades, cascade biocatalysis has attracted tremendous interest as a promising technology for green and sustainable chemical production. Combining different enzyme chemistries such as reduction, oxidation, isomerization, hydrolysis, and condensation, many cascades have been developed, converting petroleum or bio-sourced feedstock to valuable chemicals. Herein, we review some basic concepts of cascade biocatalysis and recent examples of chemical synthesis via biocatalytic cascade reaction.

20.2 DESIGNS OF BIOCATALYTIC CASCADE REACTIONS 20.2.1 TOPOLOGY OF CASCADE 20.2.1.1 Linear cascade A linear cascade (Fig. 20.2A) is the most common type and provides a simple synthetic solution for a substrate product pair [4]. In this design, two or more consecutive reactions are conducted in a same reaction vessel with the initial substrate (S) and two catalysts (C1 and C2), so that the intermediate from the first reaction (P1) is converted by the subsequent catalytic reaction to the product P2. The reactions in the cascade may be carried out in a simultaneous or sequential manner

20.2 DESIGNS OF BIOCATALYTIC CASCADE REACTIONS

429

FIGURE 20.2 Common topology of biocatalytic cascade reactions, including (A) linear cascade, (B) cyclic cascade, (C) orthogonal cascade, (D) coupled cascade, (E) double-coupled cascade, (F) convergent cascade, and (G)divergent cascade. S, Substrate; P, product; C, catalyst.

depending on the catalytic conditions and specificity of the catalysts. Most of the biocatalytic cascades described in this chapter are linear cascades.

20.2.1.2 Cyclic cascade The design of a cyclic cascade is shown in Fig. 20.2B. At least two substrates are first converted by the first reaction with a nonselective catalyst C1 to give product (P). The second step is catalyzed by a selective catalyst C2, which converts P back to substrate S2, finishing the cascade in a circle. Through this process, S1 is continuously removed until exhausted; thus a high percentage of S2 is accumulated. Cyclic cascade is mainly used for the specific conversion of different chemical or stereoisomers, such as deracemization and epimerization [5 9].

20.2.1.3 Orthogonal cascade Orthogonal design (Fig. 20.2C) starts with a reaction producing two intermediates P1 and P2. Subsequent catalysis specifically converts only one of the intermediates to the desired product P3. This cascade is often seen for coproduct removal in transaminase-catalyzed transamination to shift the reaction equilibrium toward the formation of the amine product [10]. Orthogonal cascades can also be used for the derivatization of carboxylic acid from lipase-catalyzed ester hydrolysis [11] or lyase-catalyzed deamination [12].

20.2.1.4 Coupled cascade In a coupled cascade, two reactions may be connected through a shared substrate and product (Fig. 20.2D). For instance, during the conversion of S1 to P1, a cosubstrate S2 is used and a coproduct P2 is produced. Meanwhile, a parallel S3 to P3 conversion is present, which requires P2 as cosubstrate, converting it back to S2. The second reaction is thus coupled to the first and does not start until P2 is generated. In reactions that require the use of cofactor or cosubstrate, a coupled

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CHAPTER 20 BIOCATALYSIS FOR CASCADE REACTIONS

cascade is often used to provide cofactor regeneration via coupled reaction [2,10]. To simplify the reaction system, a cascade may be double-coupled with the reaction intermediate and the cosubstrate/coproduct is shared between two reaction steps (Fig. 20.2E). Practically, the cascade only requires both catalyst, a stoichiometric amount of S1 and catalytic amount of S2. The conversion of S1 to P3 is conducted in tandem with S2 being recycled between the two reactions. A well-known example of a double-coupled cascade is the hydrogen-borrowing cascade for alcohol to amine conversion [13 15].

20.2.1.5 Convergent and divergent cascade Convergent cascade (Fig. 20.2F) design includes two or more reactions that yield the same product from different substrates [16,17], providing good flexibility for a desired chemical production. Furthermore, if the two reactions have complementary cofactor requirement, interreaction cofactor recycling can be obtained as in the coupled cascade [17,18]. Divergent cascade occurs when multiple catalytic activities are present in the first reaction step, resulting in two different products P1 and P2 (Fig. 20.2G). The subsequent reaction by C2 may convert both P1 and P2 to the final product P3. Alternatively, a specific catalyst C3 can be used for the P2 to P3 conversion. Compared with other types of cascades, divergent cascade may provide a flexible catalytic system for complex chemical processes such as oxidation of furfural [16] or multihydroxylation of steroids [19].

20.2.1.6 Modularized cascade Inspired by biological metabolic pathways, the abovementioned cascades can be modularized to form specific functional catalytic units, which can be integrated to give more complex cascade reactions, such as node-based branched cascades [2,20 22] or catalytic networks [23]. For a desired chemical product, individual modules can be combined in a plug and play manner to provide on-demand synthesis from initial substrates. Biocatalytic cascades can also be integrated into the cell metabolism of a host cell strain to form artificial pathways. Such a strategy significantly increases the diversity of production as more branches could be installed to a chemical node within a pathway, leading to new types of products [24]. Catalytic modules can also allow the utilization of cheap and readily available petroleum or biomass-based feedstock to generate platform or commodity chemicals. These long catalytic pathways of the cascade usually use engineered whole-cell catalysts to rely on the cell’s indigenous cofactors to reduce production cost.

20.2.2 PATHWAY DESIGN AND REACTION ENGINEERING Given a target chemical synthesis, various cascades may be feasible, taking different pathways from starting substrates to the final product. However, a few important factors should be considered for efficient cascade production. Ideally, catalysts for individual reaction steps should be compatible with each other. As different biocatalysts may have unique optimal temperature, pH, ion strength, or other conditions for best catalytic performance, it is important to establish a set of common optimal conditions to achieve efficient overall production. Individual catalytic reactions in the cascade should not interfere with each other but only react according to the designed pathway. This allows cascade reactions to occur simultaneously in one-pot, providing simple operation. In the

20.3 BIOCATALYSTS

431

presence of undesired activity of catalysts that leads to side-reactions [25] or incompatible reaction conditions [26], the cascade may be performed in a sequential manner. The upstream reaction is first completed, and then the next catalyst or reaction condition is introduced to start the subsequent catalytic step. Cascade reactions are often performed in aqueous buffer that provides suitable conditions for enzyme or cell catalysts without the use of any organic solvent. However, some substrates may come with low water solubility, and cosolvents may be used to enhance the substrate concentration for efficient production. On other occasions, multiphasic reaction systems (organic aqueous, solid liquid, etc.) are employed to provide high substrate concentration, reduce substrate toxicity or inhibition to the biocatalyst, and convenient in situ product separation. In order to achieve efficient production of designed cascade reactions, the rate-limiting step should be enhanced. This can be done with the development of more active and stable catalysts, optimization of catalytic conditions, or simply increased catalyst loading. If the cascade contains any thermodynamically equilibrated reaction, such as transaminase-catalyzed amination, coproduct removal is usually used to shift the equilibrium toward the desired product. Reversible reactions such as ADH-catalyzed alcohol reduction/oxidation should also be controlled to avoid undesired reverse reaction against the overall production. Once the cascade is tested and validated in laboratory scale, special reactors may be designed for further optimization and scale-up of the cascade. This includes commonly used fix-bed reactors or membrane reactors for convenient catalyst recycling, or even flow reactors providing real-time reaction and separation process.

20.3 BIOCATALYSTS 20.3.1 ENZYMES Enzymes are catalytic proteins that catalyze a wide range of chemical reactions, including substitution, reduction, oxidation, isomerization, hydrolysis, and condensation. They are often used in cascade reactions at an early stage of development to explore the feasibility of the design. Enzymes typically have excellent specificity and selectivity, and thus, are able to produce clean products free of by-products. The high preparation and operation expense of free enzymes, however, restrict the use of enzymes at the laboratory level. Over the years, more and more new enzymes have been discovered, which greatly expand the toolbox for cascade design. Enzymes with good operational stability, broad substrate scope, or novel chemistry are particularly interesting to construct more versatile [2,27] or unique cascades [28].

20.3.2 WHOLE-CELL Whole-cell catalysis is an intriguing alternative to enzyme catalysis built on the modern gene cloning and expression technology. By overexpressing required enzymes in host cells, researchers are able to construct highly active cell catalysts for a set of target cascade reactions, without implementing costly enzyme purification. Moreover, the cell membrane and cytoplasm provide enzymes a more stable and natural environment, which helps to retain the activity and stability of the enzymes, and thus, may improve the overall catalytic performance. The use of whole-cell catalysts

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also allows us to tap on the cell’s metabolism for substrates or cofactors. Inspired by natural metabolic pathways, a few cascade reactions were successfully achieved with genetic engineering to afford artificial pathways producing chemicals from natural cell metabolites [21,24,29]. The downside of using whole-cell as catalyst, however, is the existence of undesired catalytic activity from cellular enzymes. Cell metabolites may enter the reaction system resulting in contamination. In some cases, cells could also intake the substrate or product as a nutritional source for cell metabolism. Consequently, resting cells are usually harvested after cultivation and prepared for the targeted catalysis. Similar to free enzymes, it is difficult to recycle whole-cell catalysts, which leads to high production cost.

20.3.3 FUSED ENZYMES Enzyme fusion is a great way to produce multiple enzymes. Connected by flexible peptide linkers, the enzymes with closely coupled functions could be expressed and prepared together to catalyze cascade reactions with improved expression, catalytic activity, and stability [30]. This strategy is mainly used in a coupled cascade (Fig. 20.2D), such as redox enzymes fused with a cofactor regeneration partner [31], and double-coupled cascade (Fig. 20.2E), where mass transfer of intermediates between enzymes greatly affects overall catalytic performance [11]. The ratio of different enzyme components could also be conveniently adjusted to balance catalytic steps of different activities. Enzyme fusion may even help to design efficient biocatalysts in nanoscale that tune up the diffusion and transport of cascade intermediates to enhance catalysis via substrate channeling [32].

20.3.4 IMMOBILIZED ENZYMES Compared with free enzymes, enzyme immobilization on a solid supporting carrier could significantly improve the operational stability and reduce catalyst cost through enzyme recycling. Thus far, many types of immobilization techniques and carriers have been developed and greatly facilitated the application of the technology. For cascade reactions, different enzymes could be coimmobilized on the same carrier, providing efficient mass transfer [3,33,34], or on different carriers that could be separated under different conditions. Different immobilized enzymes can also be connected in a certain sequence to explore the most optimal combination for cascade reactions [35]. Immobilized enzymes are the most often used biocatalyst in the industry. However, the extra cost is imposed for carrier preparation, enzyme isolation, and immobilization. A high recycling number is typically required for catalysis with immobilized enzyme to achieve economical chemical production [36,37].

20.4 EXAMPLES OF CASCADE BIOCATALYSIS FOR THE PRODUCTION OF VALUABLE CHEMICALS Over the years, cascade biocatalysis has been widely adapted by chemists to produce a great diversity of chemicals [4,13,24,38]. Herein, we review some relevant applications of cascade biocatalysis in the production of valuable chemicals, including chiral chemicals (diols, amino alcohols, hydroxyacids, amino acids, and amines) and aroma chemicals (lactones, aldehydes, and alcohols).

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The cascades are either designed to replace problematic chemical approaches or to provide sustainable synthetic routes from cheap or renewable feedstock.

20.4.1 CASCADE BIOCATALYSIS FROM STYRENES Styrene and aryl-substituted styrenes and are petroleum-based commodity chemicals. They are widely used for polymers and are readily available with a market price as low as 1.2$/kg. Inspired by the natural biological degradation of styrene, Wu et al. developed a cascade biocatalysis using Escherichia coli coexpressing styrene monooxygenase (SMO) with terminal-selective epoxide hydrolase (EH) SpEH and subterminal selective StEH to produce enantiopure (S)- and (R)-vicinal diols, respectively (Fig. 20.3A). The cascade was performed in an organic aqueous biphasic

FIGURE 20.3 Cascade biocatalysis for the conversion of styrenes to (A) vicinal diols, (B) amino alcohols, (C) hydroxy acids, and (D) amino acids. SMO, Styrene monooxygenase; SpEH, epoxide hydrolase from Sphingomonas sp. HXN-200; StEH, epoxide hydrolase from tomato; AlkJ, alcohol dehydrogenase from Pseudomonas putida; EcALDH, phenylacetaldehyde dehydrogenase from Escherichia coli; CvTA, transaminase from Chromobacterium violaceum; AlaDH, alanine dehydrogenase from Bacillus subtilis; HMO, hydroxymandelate oxidase from Streptomyces coelicolor; EcaTA, branch chain amino acid transaminase from E. coli; DpgAT, D-phenylglycine aminotransferase.

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system to reduce toxic effect of oxides, affording 15 (S)- and (R)-diols in high ee values (mostly .95%). The cascade could also be used to convert four nonstyrene-type aryl olefins to give the corresponding chiral diols in high yield and ee values [39]. Stemming from the SMO-EH cascade, three more biocatalytic cascade modules were designed and combined in E. coli cells to provide multiple whole-cell catalyzed cascade reactions (Fig. 20.3) [20]. In the first extended cascade, the (S)-diol intermediates were converted to aldehydes through a terminal alcohol oxidation by AlkJ. Subsequent transamination of aldehydes with CvTA using alanine as amino donor-afforded enantiopure (S)-amino alcohols with 91% 99% ee (Fig. 20.3B). In an alternative design, the aldehyde products from SMO-EH-AlkJ cascade were further oxidized with an aldehyde dehydrogenase (EcALDH), giving rise to the corresponding (S)-hydroxyacids as final products with 71% 97% conversion and 96% 99% ee (Fig. 20.3C). The cascade was further extended to give a six-step cascade reaction (Fig. 20.3D), by combining a hydroxymandelate oxidase-catalyzed subterminal alcohol oxidation and an enantioselective transamination by S-selective EcaTA. This allowed efficient transformation of (S)-hydroxyacids from the SMO-EH-AlkJEcALDH cascade to optically pure (S)-amino acids (ee .99%). Similarly, (R)-amino acids could be produced in 92% 99% ee when the cascade was combined with an R-selective transaminase (DpgAT) [22]. The versatility of the SMO-based cascade was further improved through the introduction of new enzymatic catalysis (Fig. 20.4). Styrene oxide isomerase was used for the isomerization of epoxides from the SMO-catalyzed epoxidation to give aldehyde intermediates, which could be oxidized to

FIGURE 20.4 Cascade biocatalysis for the conversion of (A) styrenes to phenylacetic acids, phenylethylamines, and phenethyl alcohols and (B) α-methylstyrene to (S)-2-phenyl propionic acid and (S)-2-phenyl propylamine. SMO, Styrene monooxygenase; SOI, styrene oxide isomerase; EcALDH, phenylacetaldehyde dehydrogenase from Escherichia coli; PAR, phenylacetaldehyde reductase from tomato; CvTA, transaminase from Chromobacterium violaceum; VfTA, transaminase from Vibrio fluvialis.

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give substituted phenylacetic acids or converted to alcohols and amines through carbonyl reduction and transamination catalyzed by an alcohol dehydrogenase (PAR) and transaminase (CvTA), respectively. In this cascade, both intermediates (aldehydes) and products (acids, alcohols, and amines) are important aroma compounds that are widely used in food, cosmetic, and fine chemical industries [2,40]. This cascade could also be used to prepare enantiopure (S)-carboxylic acids from α-methyl styrenes that are important building blocks of nonsteroidal antiinflammatory drugs (Fig. 20.4B). Similarly, (S)-amines could be obtained from α-methyl styrenes. A highly active and enantioselective transaminase mutant was used to rapidly convert aldehydes to amines, minimizing the spontaneous racemization of the aldehydes [2].

20.4.2 CASCADE BIOCATALYSIS FROM L-PHENYLALANINE Using highly renewable biological feedstock for chemical production is a highly sought technology of tremendous potential to replace petroleum chemicals. An early example was demonstrated through a two-step cascade conversion of renewable L-phenylalanine to produce styrene (Fig. 20.5A) [12]. In the first reaction catalyzed by phenylalanine ammonia lyase (PAL), L-

FIGURE 20.5 Cascade biocatalysis for the conversion of L-phenylalanine to (A) styrene; (B) vicinal diols, mandelic acid, and amino acids; and (C) benzylamine. SMO, Styrene monooxygenase; SpEH, epoxide hydrolase from Sphingomonas sp. HXN-200; StEH, epoxide hydrolase from tomato; AlkJ, alcohol dehydrogenase from Pseudomonas putida; EcALDH, phenylacetaldehyde dehydrogenase from Escherichia coli; CvTA, transaminase from Chromobacterium violaceum; AlaDH, alanine dehydrogenase from Bacillus subtilis; SMDH, S-mandelic acid dehydrogenase from P. putida; EcaTA, branch chain amino acid transaminase from E. coli; DpgAT, D-phenylglycine aminotransferase; BFD, benzoylformate decarboxylase.

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phenylalanine was deaminized. The resulting cinnamic acid was then converted to styrene via a decarboxylation by phenylacrylic acid decarboxylase. In combination with the abovementioned cascade biocatalysis from styrene (Fig. 20.3), a fourstep cascade was established to prepare (S)- and (R)-phenethyl diols from L-phenylalanine (Fig. 20.5). Similarly, enantiopure (S)-mandelic acid, (S)-phenylglycine, and (R)-phenylglycine could be produced from L-phenylalanine through extending the cascade reactions with established catalytic modules. This strategy was successfully used to develop a nine-step cascade biocatalysis to produce a commodity chemical benzylamine from renewable L-phenylalanine substrate (Fig. 20.5C). The keto-acid intermediate was obtained through the previously established cascade and was then converted to benzylamine via a decarboxylation by benzoylformate decarboxylase and subsequent transamination by CvTA. This allowed the production of benzylamine from feedstock with great efficiency, affording 57% isolated yield from 60 mM L-phenylalanine substrate [21], providing an alternative for chemical production. Taking advantage of the development of metabolic engineering, the metabolic pathway to Lphenylalanine in E. coli could be greatly enhanced, affording high production titer through fermentation with a simple and renewable carbon source such as glucose or glycerol [41]. The L-phenylalanine in the fermentation media could be directly used for the cascade biocatalysis, providing more flexibility for profitable chemical production. Such an example was demonstrated in the biocatalytic cascade production of (S)-mandelic acid from L-phenylalanine after fermentation [41]. Through reaction engineering and catalyst recycling, 328 mM product was yielded. Through this strategy, 2-phenylethanol was also synthesized from commercially sourced L-phenylalanine [25] or that produced by fermentation [42].

20.4.3 CASCADE BIOCATALYSIS FOR THE SYNTHESIS OF AMINES 20.4.3.1 Cascade through alcohol oxidation and reductive amination Chiral amines are important compounds for pharmaceutical synthesis. Compared with chemical catalysis, enzymatic amination of carbonyl group provides a simpler and highly selective approach to prepare chiral amines. For instance, cascade formation of ketone or aldehyde intermediates and subsequent transamination has been widely used for many amine syntheses from cheap and readily available alcohol or carboxylic acid substrates (Fig. 20.6). As shown, two stereocomplementary

FIGURE 20.6 Cascade for the amination of alcohols with two stereocomplementary alcohol dehydrogenases and a ω-transaminase.

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alcohol dehydrogenases can be coupled with a ωTA for the conversion of racemic alcohols to enantiopure amines via a ketone intermediate [13]. In order to drive the equilibrium toward the formation of the amine, the pyruvate is continuously recycled to L-alanine. In one such example, a two-step cascade reaction was established by combining acetohydroxy acid synthase I and a (S)- or (R)-selective transaminase to prepare enantiopure pseudoephedrine directly from benzaldehyde and pyruvate acid [43]. With the discovery and development of new reductive amination enzymes such as amine dehydrogenase, reductive aminase, and imine reductase, numerous new cascade biocatalyses have been established for the synthesis of amines from alcohols. Here, we review three recent examples. The new development of amine dehydrogenases opened the possibility for combining alcohol dehydrogenases and amine dehydrogenases in a hydrogen-borrowing cascade. In the first such study, two stereocomplementary alcohol dehydrogenases were used in tandem for the oxidation of racemic alcohols to the ketone intermediates, while reductive amination of the ketones was carried out by an amine dehydrogenase to afford (R)-amines in up to 96% yield and .99% ee (Fig. 20.7) [14]. This cascade relied on the elegant strategy of shuttling hydrogen for consecutive oxidation and reduction steps via a nicotinamide cofactor, whereas ammonia was employed as an amine donor, thus producing only water as a by-product. This system was further simplified by replacing enantioselective ADHs with a nonenantioselective enzyme, thus requiring only two enzymes for the H-borrowing cascade [44]. The use of two stereocomplementary alcohol dehydrogenases was, however, a limiting factor for the cascade; hence another study used a single nonenantioselective alcohol dehydrogenase for the oxidation of alcohols to ketones. The authors also replaced the amine dehydrogenase in the second step with a newly discovered reductive aminase from Aspergillus oryzae, which also relied on a nicotinamide cofactor for hydrogen transfer, thus maintaining the hydrogen-borrowing nature of the previous cascade (Fig. 20.8A) [45]. This new two-enzyme cascade biocatalysis converted

FIGURE 20.7 Hydrogen-borrowing cascade for the asymmetric amination of alcohols with two stereocomplementary alcohol dehydrogenases and an enantioselective amine dehydrogenase.

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FIGURE 20.8 Hydrogen-borrowing cascade with a nonenantioselective alcohol dehydrogenase and a reductive aminase for the production of secondary amines from (A) racemic secondary alcohols and (B) primary alcohols.

FIGURE 20.9 Enzymatic cascade involving the irreversible oxidation of primary alcohols with an alcohol oxidase followed by the reductive amination with a reductive aminase to produce secondary amines.

racemic alcohols to the corresponding chiral secondary amines in up to 84% conversion and up to .97% ee. Fortuitously, the same enzymes could also catalyze the amination of primary alcohols to produce achiral secondary amines with up to 99% conversion (Fig. 20.8B). Since the oxidation of alcohols by alcohol dehydrogenases and the amination of carbonyls by reductive aminases are reversible, the final yield of amines was limited by equilibrium, which had to be overcome by using a large excess of amine donor (up to 50 times). To overcome this, a third study employed an alcohol oxidase AcCO6 for the irreversible and efficient oxidation of primary alcohols to aldehydes, followed by reductive amination catalyzed by the same reductive aminase in a previous study (Fig. 20.9) [46]. Since the alcohol oxidase was flavin adenine dinucleotide (FAD)dependent instead of a nicotinamide cofactor-dependent enzyme, this system was not a hydrogenborrowing cascade. However, glucose dehydrogenase was used to recycle NADPH for the reductive amination step, with the addition of glucose as a coupled substrate. This cascade could catalyze the

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amination of primary alcohols to secondary amines with up to .99% conversion with just a moderate excess of amine donor (2 4 times). The production efficiency of the cascade could be further improved by using alcohol oxidase for the alcohol oxidation for large-scale applications [46]. Whole-cell combining with immobilized AmDH and cofactor recycling enzyme could also be used to improve the efficiency of ADHAmDH cascade for the preparation of amphetamine [3].

20.4.3.2 Cascade through alcohol activation and nucleophilic substitution Another notable cascade biocatalysis converting alcohol to amine relies on alcohol activation and nucleophilic substitution, as demonstrated in the preparation of unnatural amino acids from L-serine [26]. Serine acetyltransferase was first used for the acetylation of the hydroxyl group of L-serine with acetyl-CoA as acetyl donor. The resulted O-acetylserine was then substituted with nucleophiles by using O-acetylserine sulfhydrylase, which accepts azide, cyanide, thiols, and a broad range of amines for the substitution (Fig. 20.10) [47,48].

20.4.3.3 Cascade through imine formation and asymmetric C N double bond reduction Imine reductases (IREDs) provide another biocatalytic route for accessing optically pure amines via the reduction of imines. One example of a cascade uses a carboxylic acid reductase to reduce a keto acid substrate into a ketoaldehyde; then a transaminase catalyzes the amination of the ketoaldehyde into a keto amine, which spontaneously turns into a cyclic imine. Finally, an (R)- or (S)-selective IRED reduces the imine into an enantiopure cyclic amine (Fig. 20.11) [49]. This cascade could synthesize a range of di-substituted piperidines and pyrrolidines from the corresponding keto acids with up to 95% yield and in up to 98% ee and de values.

FIGURE 20.10 One-pot substitution of OH group of L-serine with pyrazole to synthesize β-pyrazol-1-yl-L-alanine via simultaneous cascade reactions by using Escherichia coli cells expressing serine acetyltransferase and immobilized O-acetylserine sulfhydrylase (CysK) for amine substitution.

FIGURE 20.11 Cascade biocatalysis for the conversion of keto acids to cyclic amines with imine reductase-catalyzed asymmetric reduction.

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20.4.4 CASCADE BIOCATALYSIS FOR THE SYNTHESIS OF LACTONES Chiral lactones are aroma and biologically active chemicals that are widely used in the industry. Chemically produced lactones are readily available but considered as inferior to natural counterparts for food, health care, and cosmetic applications. Furthermore, different stereoisomers of chiral lactones may have different odor and biological properties, resulting in different market values. Enzymatically, lactones can be produced via Baeyer Villiger oxidation of cyclic ketones with Baeyer Villiger monooxygenase (BVMO) [50] or intramolecular cyclization of hydroxy acids via various redox reactions [51 53], leading to two major types of biocatalytic cascade productions. The first type of cascade relies on the asymmetric reduction of α,β-unsaturated cyclic ketone substrates and enantioselective Baeyer Villiger oxidation. In one example, R-selective ene-reductase (ER) in the resting cells of Acinetobacter sp. RS1 is able to fully reduce the C C double bond of two alkene substrates to give enantiomerically enriched alkyl-substituted cyclopentanone (Fig. 20.12). E. coli cells with overexpressed BVMO and a cofactor regeneration enzyme was then used to convert the two ketone intermediates to the corresponding (R)lactones in 97% and 98% ee, with 83% and 66% isolated yield, respectively [25]. Extended cascades were developed to incorporate alcohol dehydrogenase (ADH)-catalyzed nonenantioselective alcohol oxidation that allowed the use of racemic cyclic alcohols as substrates [54,55] or to combine with enzymatic hydrolysis and amination to produce hydroxy acids or amino acids [56]. In a proof of concept study, the ER-BVMO cascade was combined with oxyfunctionalization catalyzed by Pseudomonas putida to convert limonene from waste orange peels to carvolactone [57]. An alternative cascade biocatalysis is based on the enzymatic formation of chiral hydroxyl carboxylate precursor, which goes through self-cyclization to give the corresponding chiral lactone. This design was well demonstrated in a three-step cascade reaction from ethyl 4-oxopent-2-enoate type of substrates. Through the highly enantioselective asymmetric C C bond reduction by enoate reductase and carbonyl reduction by an ADH, chiral hydroxyl esters were generated and simultaneously cyclized to give enantiopure lactones with 98% 99% ee [58]. The BVMO-catalyzed and ADH-catalyzed lactone formation could even be combined to give a convergent cascade biocatalysis with self-sufficient cofactor regeneration [17,18]. As shown in the Fig. 20.13, horse liver ADH catalyzes a double oxidation of diol substrates, producing NAD(P) H, which fuels the Baeyer Villiger oxidation of cyclic ketone running in parallel. Through the careful selection of substrates, the same lactone product could be obtained through this convergent cascade.

FIGURE 20.12 Preparation of (R)-lactones from enone substrates through cascade C C double bond reduction by Acinetobacter sp. and Baeyer Villiger oxidation by Escherichia coli (CPMO-GDH).

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FIGURE 20.13 (A) Preparation of chiral lactones from linear ester substrates through a cascade enoate reduction catalyzed by ER, ADH-catalyzed ketone reduction, and spontaneous cyclization. (B) A convergent cascade catalyzed with BVMO and ADH to produce lactones from ketone and diol.

20.4.4.1 Cyclic alcohols Drug and pharmaceutical molecules usually contain multiple ring structures that help to orient and stabilize pharmacologically functional structures [59]. Some ring structures may also carry chiral functional groups that link to other parts and determine the overall shape of the molecules. Furthermore, drug molecules often contain ring structures with clustered functional groups, presenting significant synthetic challenges [59]. Cascade enzymatic syntheses of chiral cyclic molecules often rely on asymmetric C O bond-forming and derivatization. In one of the reported cascade preparation, meso-cyclic epoxides were firstly hydrolyzed by EH to give trans-(1R,2R)-vicinal diols. A highly enantioselective ADH was then used for R-selective alcohol oxidation, converting diols to the corresponding hydroxyl ketones with 98% 99% ee [60]. This cascade was successfully achieved with both enzymatic and whole-cell catalytic systems with engineered E. coli (SpEH) and E. coli (BDHA) cells (Fig. 20.14A). Functional groups may also be introduced to cyclic rings via hydroxylation with P450 monooxygenases. As shown in Fig. 20.14B, D-glucono-1,5-lactone is hydroxylated by a P450-BM3 mutant giving a racemic hydroxyl ester, which further goes through alcohol oxidation and enantioselective enoate reduction to afford R- or S-product [61]. In another example, two highly active and enantioselective P450 mutants were developed for a three-step cascade dihydroxylation of cyclohexane. Each mutant is able to catalyze a tandem hydroxylation, alcohol oxidation, and asymmetric hydroxylation cascade reactions, producing the corresponding enantiomer of hydroxyl ketones. Alternative cascade biocatalysis to install chiral functional groups at the vicinal position of the ring involves the highly enantioselective reduction of cyclic ketones with ketone reductase (KRED) [62]. Racemic ketonitriles were selectively reduced by different enantioselective KREDs. Combined with nonselective hydrolysis of the resulted hydroxynitriles by Rhodococcus rhodochrous cells, each of the four stereoisomers was produced in high dr and ee values (Fig. 20.14C).

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FIGURE 20.14 (A) Preparation of chiral hydroxyl ketones through a cascade consisting of epoxide hydrolysis and enantioselective alcohol oxidation. (B) Three-step oxidative cascade to prepare hydroxyl ketones catalyzed by a single P450 mutant. (C) Reduction-hydrolysis cascade for the conversion of cyclic ketonitriles to hydroxy acids.

20.5 CONCLUSIONS AND PERSPECTIVES Due to its great simplicity, cascade biocatalysis has become a valuable tool for organic chemists to prepare high-value chemicals from cheap and readily available feedstock. It is particularly useful for the synthesis of complex compounds that may have multiple functional groups or chiral centers. Using highly selective and mild biocatalyst, it even allows the direct combination of reductive and oxidative reactions in the same reaction vessel. In some problematic chemical syntheses, biocatalytic reactions could be combined in a cascade to replace their chemical counterparts. Two or three steps of biocatalysis could also be performed to provide an alternative synthetic route. Immobilized enzymes are typically used for such applications due to their tolerance to organic solvents and compatibility with the equipment setup in typical organic synthesis. This strategy was used in the cascade production of chiral amines, lactones, and other chiral molecules. Alternatively, whole-cells coexpressing functional enzymes may be used as immediate catalysts. This is typically used for cascades with long reaction pathways to reduce catalyst cost. Cell metabolism could also provide cascade reactions with necessary cofactors or even initial substrates. This allowed the production of many chiral or other valuable chemicals from styrenes and L-phenylalanine. Developments of new synthetic cascade reactions are always limited by the available catalysts and catalytic reactions in the toolbox. On one hand, new catalysts with different substrate scope, catalytic specificity, and regio- and stereoselectivity provide useful add-ons to the existing cascade

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biocatalysis designs, expanding the range of application. For instance, the catalytic performance of the existing enzymes could be tuned to the desired level through enzyme engineering. Whole-cell catalysts could be developed by overexpressing hydroxylating enzymes with efficient cofactor regeneration system in a host cell strain through molecular cloning approaches. Enzymes with improved operational stability could be obtained through immobilization and conveniently recycled at reduced temperatures or through magnetic separation. On the other hand, discovery of novel biocatalytic reactions allows new synthetic routes and leads to new chemical productions. For instance, aryl substituents may also be introduced from upstream L-phenylalanine in the cascade. Starting from five substituted cinnamic acids that can be easily prepared via Grignard reaction, a cascade biocatalysis consisting of nonenantioselective amination catalyzed by PAL and enantioselective deracemization, the corresponding L-phenylalanine derivatives were obtained in 99% ee. An alternative cascade biocatalysis was designed to produce substituted L-phenylalanine from substituted benzylaldehyde and glycine. The cascade was initiated with aldol condensation and deamination catalyzed by threonine aldolase (TA) and threonine deaminase (TD), which was enantioselective converted to L-amino acid products by R-selective transaminase.

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FURTHER READING B.R. Lukito, S. Wu, H.J.J. Saw, Z. Li, One-pot production of natural 2-phenylethanol from L-phenylalanine via cascade biotransformations, ChemCatChem. 11 (2019) 831 840. Available from: https://doi.org/10.1002/ cctc.201801613.

Index Note: Page numbers followed by “f” and “t” refer to figures and tables, respectively.

A Ab initio structure-prediction calculations, 98 Abortiporus biennis, 404 Absorbance-activated droplet sorter (AADS), 71
72, 111
112 Acetone, butanol, and ethanol (ABE) fermentation, 383
384 enzymatic engineering for, 384 Acetyl xylan esterases, 417 Acid
base catalysis, 5 Acid proteases, 21 Activated carbon, 197 Adenosine triphosphate (ATP), 23 Advanced computational modeling, 58 Agarose gel, 39 Agarplate-based assays, 154
155 Agrobacterium tumefaciens NRRL B11291, 58
59 Alcohol dehydrogenases (ADH), 146 Alcohol lysis, 5 Alginate, 194 Alkaline hot spring, 92 Alkalizing hydroxides, 292 Amino acid, 3 Amino-dipeptidase, 20 Ammonium sulfate, 38
39 Amylases, 17
18, 42, 345
346, 405
407 thermostability, 18 Ancestral sequence reconstruction (ASR), 75 Animal-derived enzymes, 31 Antimicrobial peptides (AMPs), 134
135 Artemisinic acid, 343 Arthrobacter sp., 33
34 Aspergillus sp., 251
252 A. aculeatus, 402 A. aculeatus β-glucosidase, 322
323 A. nidulans, 253
255, 403 A. niger, 18
19, 402
403, 406 A. niger α-amylase, 205 A. oryzae, 403 A. oryzae alkaline protease genes, 33
34 A. oryzae (AspRedAm), 341 Avidin, 40
41

B Bacillus sp., 21 B. atrophaeus (BaLc), 404 B. badius phenylalanine dehydrogenase (PheDH), 160 B. cereus (BcLeuDH), 341
342 B. licheniformis, 406

B. licheniformis (BLA), 93
94 B. pumilus SV-85S, 407 B. stearothermophilus, 406 B. stearothermophilus leucine dehydrogenase (LeuDH), 159
160 B. subtilis, 94, 345 B. subtilis lipase, 70 thermostabilization of, 71f Baeyer
Villiger oxidation of cyclic ketones, 440 BCR-ABL1 fusion protein, 119
120 Beta-factor (B-factor) analysis, 76 B-factor profiles for protein thermostability, 76 Bifidobacterium sp. B. bifidum, 285, 418 B. breve, 418 B. infantis, 418 B. longum KACC 91563, 316
317 Bimolecular fluorescence complementation (BiFC) technique, 136 Biobutanol, 383
384 Biocatalysis engineering, 50, 103 advantages of, 50 phases, 50 Biocatalysts, 49
50 Biocatalytic cascades, 427 designs of, 428
431 convergent cascade, 430 coupled cascades, 429
430 cyclic cascade, 429 divergent cascade, 430 linear cascade, 428
429 modularized cascade, 430 orthogonal cascades, 429 examples, 432
441 for conversion of keto acids to cyclic amines, 439f of epoxide hydrolysis, 442f from L-phenylalanine, 435
436, 435f from styrenes, 433
435, 433f for synthesis of amines, 436
439 for synthesis of lactones, 440
441 through alcohol activation and nucleophilic substitution, 439 through alcohol oxidation and reductive amination, 436
439 through imine formation and asymmetric C
N double bond reduction, 439 pathway design and reaction engineering, 430
431 topology of, 429f

449

450

Index

Biodiesel, 352 Bioethanol, 245, 351
352 Biofuels, 335
336 Biohydrogen, 352 Biomass, 246
247 processing enzymes, 247
249 Biorefinery and biorefining, 245
246, 363
365 first and second generation, 364
365 third generation, 365 Biosynthesis of boceprevir, telaprevir, and atazanavir, 342f of natural products, 343 of parthenolide, 343f of plant-derived natural products, 344f Biotransformations, 335
336 in biofuels industry, 348
352 classification, 349
351 of chemicals, 346
348 in food processing, 343
348 in pharmaceutical industry, 336
343 BRaunschweig ENzyme DAtabase (BRENDA), 12

C Calcium alginate-entrapped peroxidase, 206 CaM-binding peptide of myosin light chain kinase (MLCKp), 136 Candida sp. C. antarctica lipase B (CALB), 336
337, 337f C. boidinii (CbFDH), 341
342 C. boidinii xylose reductase (CbXR), 58 C. glabrata (CgKR1), 338 C. rugosa Lipase1 (Lip1), 74 Cannabidiol (CBD), 343 Carbohydrases, 322
323 Carbohydrate-active enzyme (CAZy) database, 261 Carrageenan, 195 Cascade biocatalysis, 427
428 production of high-value chemicals via, 428 Cascade reaction, 427, 428f Catalytic mechanism of, 3 Celite, 196 Cell-free enzymes, 335 Cell-free protein translation systems, 111 Cell metabolites, 432 Cellobiohydrolases (CBHs), 18 Cellobiose dehydrogenase, 15
16 Cellobiose 2-epimerases, 323
324 Cellulases, 18
19, 248
249, 397, 403 accessory and auxiliary enzymes, 256
263 auxiliary enzymes, 261
262 β-glucosidase, 256
257 laccases, 260 xylanase, 257
259 for bioconversion, 249
250

encoding genes present in microorganisms, 254t enhanced production, 254t Cellulolytic filamentous fungi, 253 Cellulose, 195, 246, 350
351 Cellulose-binding module (CBM), 248
249 Cellulose/carbohydrate-binding domain (CBD), 248
249 Cellulosic biofuel industry, 19 Ceramics, 196 Cetyltrimethyl ammonium bromide (NH4Br), 38
39 Charcoal, 197 Chimeric or fusion protein applications of for biomass degradation, 133
134 as biopesticides, 134
135 biosensor (protein switching), 136 as biotherapeutic agent, 135
136 artificial or synthesized or recombinant, 120
121 methods used for, 121f, 122
132 domain insertion, 129
132 end-to-end fusion, 122
129 linker, 122
128 posttranslational conjugation, 132 natural, 119
120 phenological or physiological role, 120 Chiral alcohols, 337
340 Chiral amines and amino acids, 340
342 Chiral carboxylic acids, 336
337 Chitin, 195 Chitosan, 195 Chlamydomonas reinhardtii fermentation, 352 Chlorella reinhardtii biomasses, 352 Chromogenic/fluorogenic substrate surrogate-based assays, 152
153 Chrysosporium lucknowense cellulases, 250 Clostridium acetobutylicum xyn B, 259 ClustalW alignment tool, 96
98 Codeinone, 343 Cofactors, 4 Collagen, 195 Combinatorial active-site saturation test (CAST), 151 Combinatorial sequence-optimization calculations, 98 Combi-protein-coated microcrystals (Combi-PCMCs), 51
53 Computational disulfide engineering, 79
80 Computational engineering, of protein structures, 58 Computational molecular simulation, 106 Condensation, 5 Conducting polymers, 198 Conjugated linoleic acid (CLA), 319 Consensus analysis (CA), 74 Copper-containing oxidoreductases, 15 Coupled enzyme assays, 153
154 Covalent catalysis, 6 Covalent immobilization, 202 CRISPR-Cas, 398

Index

Cross-linked enzyme aggregates (CLEAs), 172
175, 177, 199, 228
229 Cross-linked enzyme crystals (CLECs), 51
53 Cross-linked protein-coated microcrystals (CL-PCMC), 51
53 Cross-linking system, 132, 172
175, 199, 202 Cryogel, 196 Cryogenic electron microscopy (CryoEM), 112 Cyan fluorescent protein (CFP), 136 Cyclic alcohols, 441 CYP101D2, 57
58 Cysteine proteases, 21 Cytidine triphosphate, 23

D Darwin’s theory of evolution and natural selection, 148 Database of Aligned Structural Homologs (DASH), 105 Debye
Waller factor, 76 Decarboxylases, 22 Dehydrogenases, 15 Dekkera bruxellensis, 322 De novo enzyme designing, 96 De novo protein design, 78
79, 98, 109 Deoxyribonucleotide triphosphates (dNTPs), 149 β-D-fructofuranoside (FFase), 275
276 Directed evolution, 69
72, 106
108, 222 DNA shuffling, 107 for enantioselectivity of enzymes, 148
155 high-throughput screening of enantioselective enzymes, 152
155 mutant library construction, 149
151 of enzymes, 93
95, 94f random mutagenesis technique, 107 in vivo continuous, 108 Disulfide by Design (DdD), 79 DNase I nuclease, 130 DNA shuffling, 70, 149
150, 150f Docosahexaenoic acid (DHA), 319 Docosapentaenoic acid, 319 Domain insertion library, 129
132, 131f construction of, 130
132 Drosophila nuclear transport factor-2-related (Dntf-2r) gene, 119
120 Dye-decolorizing peroxidases (DyPs), 15 Dynamic reductive kinetic resolution (DYRKR), 339
340, 339f

E Easy-to-use sequence-based engineering method, 74 Eicosapentaenoic acid (EPA), 319 Enantioselective biotransformations, 145
146 applications of, 146f Enantioselective enzymes, 145 Endoamylases, 17

451

Endoglucanases, 105
106, 248 Endoxylanases, 417 Engineered transposon, 132 Enterobacter aerogenes, 352 Entrapment system, 199 Enzymatic-assisted extraction (EAE), 385 Enzymatic transgalactosylation, 274 Enzyme Commission (EC), 11, 32
33 number, 12 Enzyme conjugation, support systems for biopolymers, 194
196 conducting polymers, 198 inorganic support materials, 196
197 organic support materials, 193 smart polymers, 198 Enzyme engineering, 218 strategies for thermostabilization, 68
82 Enzyme immobilization, 51
55, 192, 192f, 193t advantages of, 192
193 applications, 203
206 biomedical, 206 in bioremediation methods and wastewater treatment, 233 in brewing industry, 204 in cosmetic industry, 235 in detergent industry, 205 in food industry, 203 in food processing, 230
232 medical, 232 in pharmaceutical industry, 203
204 in pulp and paper industry, 205 in textile industry, 206, 234
235 valorization of food wastes, 234 basic methods and submethods of, 52f beneficial steps of, 226 chemical methods, 202 covalent binding, 202 cross-linking, 202 functionalized materials-based cues at nanolevel, 51
53 history of, 192 impacts of, 223
229 activity changes, 224 changes in features, 224 diffusional effects, 225 specificity and selectivity, 224
225 materials used, 229 matrices, 229, 230t merits and demerits of methods, 53t physical methods, 200
202 ionic binding, 201 microencapsulation, 201
202 physical adsorption, 200
201 physical entrapment, 201
202 preimmobilization factors and actions, 225
226

452

Index

Enzyme immobilization (Continued) selected methods of, 226
229 adsorbing method, 227 covalent bonding method, 227
228 cross-linking techniques, 228
229 entrapment method, 228 strategies, 217
218 support systems employed in, 194f techniques, 172
175, 198
199, 199f Enzymes, 3 active site of, 4 adsorption binding of, 175 animal-derived, 31 applications of, 41
42 as biocatalyzers, 12
13, 219
223, 431 in bioethanol production, 372
373 for biofuels, 372
385 in biogas production, 384
385 bioinformatics analysis, 96 for biomass degradation, 124t biomass-processing, 247
249 building block of, 3 catalytic mechanism of, 3 acid
base catalysis, 5 basics of, 4
5 catalytic reaction, 12 chirality, 49
50 covalent catalysis, 6 electrostatic catalysis, 5 essential component for execution of, 4 historical overview of, 5 induced-fit model, 13
14 keyhole-lock-key model, 13
14 lock-and-key model, 13
14 mechanistic view of, 5
6 selected-fit model, 13
14 types of selectivities, 49
50 commercial, 309 covalent binding of, 176 cross-linking of, 177
178 databases, 12, 13t denaturation of, 219 destabilization of, 219
220 differences between protein and, 3
4 enantioselectivity of, 148 of amine dehydrogenase, 159
161, 160f directed evolution, 148
155 examples, 155
161 invertion of P450pyr monooxygenase, 158
159, 159f of P450pyr monooxygenase, 156
158, 156f entrapment or encapsulation of, 176
177 extracellular, 36 in food industry and their potential applications, 173t fusion, 432

genetic engineering of, 377t improvement, computational approach, 96
98, 97f market scenario, 262
263 microbial, 309 from microbial system, 31
32 production, 34
40, 35f nomenclature and classification, 11
12, 32
33 production of, 33
41, 366
372 from brown algae, 370
371 from green algae, 369
372 from macroalgae and derivate polysaccharides, 368t from plants and animals, 40
41, 41t from red algae, 371
372 semi-solid-state fermentation, 367
368 solid-state fermentation, 367
368 upstream processes for, 34 in production of functional foods, 310
321, 310f carbohydrate-modifying enzymes, 312
317, 313t genetically modified enzymes, 322
324 L-asparaginase, 317
318 lipases, 318
320 phytases, 320
321 proteases, 310
312, 311t tannase, 320 using specific enzyme-producing starters, 321
322 relationship between structure and function, 147 stabilization of, 219
222 through partitioning, 225 structural dynamics of, 6
7 thermostable, 67, 70, 72f Enzymes catalyzing production fructooligosaccharides, 314
315 resistant starch, 316 xylooligosaccharides, 315
316 Enzyme stability, in different pH range molecular mechanism, 92
93 Error-prone polymerase chain reaction, 70, 149 Escherichia coli, 33
34, 345 Esterases, 417 Esterification, 5 Ethanol, 245 Eupergit C, 193, 205 Exoglucanase, 248 Exoxylanases, 417 Extremophilic microorganisms, 67 Extremostable proteins, 68

F FastML, 75 Fed-batch culture, 40
41 Fermentation process, 35
37 Feruloyl esterases, 417 Filter paper activity (FPU), 249
250 First-generation biofuels, 349

Index

FLAG peptide, 137 Flavin-containing oxidases, 15 Flexible linkers, 123 Fluorescence-activated cell sorting (FACS), 155 based screening and isolation, 111 Fluorescence-activated droplet sorting, 71
72 Flurbiprofen, 336
337 FoldX, 77
78 Food chemical production, 348 Fo¨rster resonance energy transfer (FRET), 136 FRET probe-based high-throughput screening, 71
72 Fossil fuels, 245 Fourth-generation biofuels, 351 Fractionation of oligosaccharides, 292 Fructooligosaccharides (FOS), 289
290, 290f, 347 Fructosyltransferase, 275 Fruit juicing, 397 Functional foods, 309 enzymes in production of, 310
321, 310f carbohydrate-modifying enzymes, 312
317, 313t genetically modified enzymes, 322
324 L-asparaginase, 317
318 lipases, 318
320 phytases, 320
321 proteases, 310
312, 311t tannase, 320 using specific enzyme-producing starters, 321
322 revenue of functional food market worldwide, 274f Functionalized multiwall carbon nanotubes (f-MWCNTs), 198 Fungal amylases, 406 Fungal oxidoreductases, 15 Fused enzymes, 432

G Galactooligosaccharides (GOS), 276
285, 347
348 β-Galactosidase, 180, 274
275, 313
314 Gelatin, 195 Genencor, 262
263 Generally recognized as safe (GRAS), 34, 171 Gene shuffling, 149
150, 150f Genetically modified enzymes, 322
324 Genetic engineering, 252
256 of enzymes, 377t for bioethanol and biofuel production, 373 Geobacillus sp., 406 GH domain, 19 Glass, 197 Glucose isomerase, 23, 346
347 β-Glucosidases, 256
257, 275, 316
317 α-Glucuronidases, 417 Glutathione S-transferase protein, 137 Glutathione transferases (GSTs), 16 β-Glycosidases, 275 Glycoside hydrolases, 276

453

Graphene nanosheet, 180 Green algae, 365
366, 369
372 Green chemistry
mediated biological process, 171 Green fluorescent protein (GFP), 136 Greenhouse gas (GHG) emissions, 245 Guanosine triphosphate, 23

H Hairpin ribozymes, 7 Hammerhead hepatitis δ virus (HDV), 7 Heme-containing peroxygenases, 15 Hemicellulose, 246, 350
351 degrading enzymes, 257
258 High-throughput screening methods, 111
112 of enantioselective enzymes, 152
155 Horseradish peroxidase (HRP), 153
154 Humicola sp., 251
252 H. insolens cellulase system, 18
19 Hydratases, 22 Hydrocolloids, extraction of, 385
386 Hydrogels, 196 Hydrogen-borrowing cascade, 437f, 438f Hydrolases, 17
21, 33 Hydrolysis, 5

I Ibuprofen, 336
337 Imine reductases (IREDs), 439 Immobilized enzyme, 172, 432 Imprinted CLEAs (iCLEAs), 51
53 Industrial enzymes stabilization of, 219
222 grouping methods, 221 in nonconventional solvents, 223 pH value, 220
221 temperature, 221 Intermolecular and intramolecular hydrogen bonding, 246 International Union for Pure and Applied Chemistry, 32
33 International Union of Biochemistry and Molecular Biology (IUBMB), 11, 32
33 Inulinases, 276 Inverse-cascade principle, 365 Isomaltooligosaccharides (IMO), 276, 290
291, 291f Isomerases, 22
23, 33 Iterative saturation mutagenesis (ISM), 70, 151, 152f

J Jeotgalibacillus malaysiensis, 317 Jingwei (jgw) gene family of Drosophila, 119
120 Juice clarification, 397
398 enzymes used for, 405f amylase, 405
407 cellulose, 403 chitinase, 407

454

Index

Juice clarification (Continued) β-galactosidase, 407 laccase, 403
404 pectins, 402
403 pullulanase, 407 transgalactosidase, 407 xylanases, 407 immobilized enzymes and their parameters used for, 398t rationale for, 399
400 technologies available for, 398 centrifugation and finishing, 400 chemical processes, 401 enzymatic process, 401 finings, 401 freezing and heating, 400 straining or screening, 400 Juice extraction procedure, 399f Jumping genes, 132

K Ketoprofen, 336
337 Ketoreductases (KREDs), 337, 441 Kluyveromyces lactis, 285 Koshland Jr., Daniel, 147

L Laccases, 16, 260, 403
404 immobilization, 176 Lactobacillus sp. L. acidophilus L1, 319 L. delbrueckii, 345 L. fermentum (LfSDR1), 338
339 L. plantarum, 285 L. plantarum ATCC 8014 cells, 319 L. reuteri (ATCC 23272 and ATCC 55739) strains, 319 Lactosucrose, 288
289 Lactulose, 286
287, 286f α-L-Arabinofuranosidases, 417 L-asparaginase, 317
318 Leucine-rich repeat transmembrane neuronal 2 (LRRTM2), 74 Ligases, 23
24, 33 DNA, 23
24 reactions catalyzed by, 24t Lignin, 246
247 degrading enzymes, 260 -modifying peroxidases (LMPs), 15 peroxidases (LIPs), 15 Lignocellulose saccharification, 250 Lignocellulosic biomass, 245
247 bioconversion using cellulases, 249
250 enzymes-mediated bioconversion of, 247 microbial degradation of, 248 pretreatment of, 247 Lignocellulosic materials (LCM), 413

Linkers, 122
128 artificial or synthesized, 123
128 cleavable, 127 photo, 128 protease-sensitive, 127 in vivo, 127 designing tools and databases, 128 flexible, 123 naturally occurring, 123 rigid, 123
127 Linoleic acid (LA) isomers, 319 α-linolenic acid, 319 Lipase-cutinase (Lip-Cut) chimeric enzyme, 120
121 Lipases (triacylglycerol acylhydrolases), 20, 42, 235, 318
320 Litopenaeus vannamei hepatopancreas, 345 Lock-and-key theory, 147, 147f Lyases, 21
22, 33 Lytic polysaccharide monooxygenase (LPMO), 261
262 AA9 LPMO proteins, 261
262, 262t AA10 LPMO proteins, 261
262 Lytic polysaccharide monooxygenases (LPMOs), 15
16, 248

M Macroalgae biofuels from, 374t biomass, 363, 364f brown algae, 365
366, 370
371 enzymatic saccharification of, 380 green algae, 365
366, 369
372 enzymatic saccharification of, 373
382 polysaccharides, 365
366 red algae, 365
366, 371
372 enzymatic saccharification of, 380
382 as a source of high-value products, 365
366 Macromolecular CLEAs (M-CLEAs), 51
53 MAFFT (multiple alignment using fast Fourier transform), 105 Magnetic CLEAs (mCLEAs), 51
53 Manganese peroxidases (MNPs), 15 “Medium-throughput” assays, 155 Membrane nanofiltration, 293 Membrane technology, 293
294 Metal
organic frameworks (MOFs), 51
55 Microbial enzyme production, 34
40, 35f cell-free system, 37 cell permeabilization, 37 cell rupture methods, 37 cross-flow microfiltration, 37 by fermentation, 35
37 formulation of, 40 localization of enzyme, 36 purification, 38
40 affinity chromatography, 39

Index

bovine pancreatic nucleases treatment, 38
39 efficient elution of proteins, 39 hydrophobic chromatography, 39 ion-exchange chromatography, 39 protein fractionation, 38
39 salting out precipitation, 38
39 size-exclusion chromatography, 39 recovery of enzymes, 37
38 ultrafiltration, 37
38 Modeller tool, 96
98 Molecular dynamics (MD), 3, 76
77 Monte Carlo-based sampling protocol, 79 MrBayes, 75 MuDel transposon, 133f Multiple sequence alignment (MSA), 74, 105 Multiple Sequence Comparison by Log-Expectation (MUSCLE), 105 Multiplex inverse PCR, 131
132 Multipurpose CLEAs (multi-CLEAs), 51
53 Muscle alignment tool, 96
98 Myeciliophthora, 251
252

N Nanobiocatalyst, 175 bioprocessing through, 235 in food processing, 178
180 from Thermomyces lanuginosus, 179 Nanocarrier-bound biocatalyst, 172 Nanofiltration membranes, 293
294 Nano-immobilization, 172
178 Natural products, biosynthesis of, 343 N-benzyl-pyrrolidine, 156
157 Neurospora crassa, 253
255 Niabella soli strain DSM 19437, 119
120 NNK single-site mutagenesis, 161 Nonhomologous isofunctional enzymes (NISE), 11 Noscapine, 343 Novozymes, 262
263 Nuclear magnetic resonance (NMR) spectrometry, 105
106

O Ochrobactrum anthropi SV3, 58
59 β-O-4-ether bonds, 260 Omega-3 fatty acids, fish oil hydrolysis for, 178
179 Overlap extension polymerase chain reaction (PCR), 128
129 Oxcarbazepine, 338f Oxidoreductases, 33, 42 Oxidoreductases or oxireductases, 15
16

P Papain, 40
41 p-coumaric acid esterases, 417 Pectin acetylesterase, 402 Pectinase, 397
398

455

Pectinlyase, 402
403 Pectin methylesterase, 402 Pectins, 195
196, 402
403 Penicillin acylases (PA), 92
93 Penicillium sp., 251
252, 321
322 P. glaucum mold, 145 Peroxidases, 15 Phanerochaete chrysosporium, 260 pH-dependent stability, of enzymes, 92
93 Phenolic compounds, extraction of, 386 pH indicator assays, 153 pH value of reaction media, 220
221 Phylogenetic analysis by maximum likelihood (PAML), 75 Phylogenetic analysis using parsimony (PAUP), 75 Phyre tool, 96
98 Phytases, 320
321 Pichia pastoris, 345 GS115, 33
34 Pigments, extraction of, 386
387 Plant and animal-derived food proteases, 323 Pleurotus sp., 16 P. ostreatus, 260 Plexaura homomalla, 119
120 Poly-acrylic acid-co-2-acrylamido-2-methylpropanesulfonic acid (PAA-co-AMPS) polymer, 346
347 Poly-acrylic acid (PAA), 346
347 Poly [bis(pmethylphenoxy)] phosphazene (PMPPh), 206 Polyethylene glycol-dextran (PEG/dextran), 38 Polyethylene glycol-salt (PEG/salt), 38 Polyethyleneimine, 38
39 Polygalacturonase, 402 Polygalacturonate lyase, 22 Polygonum cuspidatum, 322 Poly (lactic acid) (PLA), 201
202 Poly (lactic-co-glycolic acid) (PLGA), 201
202 Polymethyl galacturonate lyase, 22 Polyphenol oxidases (PPOs), 16 Polypropylene-based hydrophobic granules/Accurel EP-100, 200
201 Polyunsaturated fatty acids (PUFA), 318
319 Polyvinyl alcohol (PVA) cryogels, 196 Porous CLEAs (p-CLEAs), 51
53 Prebiotic galactotrisaccharides (GOS3), 317 Prebiotics, 273, 347
348 beneficial activities of, 273
274 downstream processing of, 292
294 enzymes involved in production of, 281t β-galactosidase, 274
275 glycoside hydrolases, 276 inulinases, 276 fermentation of, 273
274 global status of, 294
295 production of, 276
291 enzymes involved in production of

456

Index

Prebiotics (Continued) β-D-fructofuranoside (FFase), 275
276 fructosyltransferase, 275 β-glucosidases, 275 β-glycosidases, 275 fructooligosaccharides, 289
290, 290f galactooligosaccharides (GOS), 276
285 isomaltooligosaccharides, 290
291 lactose, 277t lactosucrose, 288
289 lactulose, 286
287, 286f tagatose, 287
288, 288f PreticX prebiotics, 420 Procheck tool, 96
98 Profens, 336
337 Propionibacterium acnes isomerase, 319 Proteases, 20
21, 310
312, 311t Protein-coated microcrystals (PCMCs), 51
53, 197 Protein Data Bank, 105 Protein engineering, 51
59, 103, 120, 221
222 advanced technologies for, 109
112, 110f cell-free translation systems, 111 cryogenic electron microscopy (CryoEM), 112 high-throughput screening methods, 111
112 spatial organization, 109
110 surface display systems, 111 approaches, 104f computational molecular simulation, 106 multiple sequence alignment, 105 rational design, 104
106 structure-guided design, 105
106 structure-assisted, 56
58 thermostability, 68f via directed evolution and rational design, 58
59 Proteins, 3
4 stabilization, software for monitor flexibility or predicting mutation for, 73t tag, 137 Pseudoalteromonas atlantica, 315
316 Pseudomonas sp. P. cepacia lipase CLEA, 177
178 P. oryzihabitans, 317 P. putida, 404 Pyridoxal-50 -phosphate, 340
341

R Racemic ketonitriles, 441 Random mutagenesis, 149
150, 251
252 semi-rational design, 151 Rational and semi-rational design, 72
78, 72f Rational design method, 96
98, 97t Recombinant Cel7A of Penicillium funiculosum (rPfCel7A), 255
256

Reconstructing evolutionary adaptive path (REAP) approach, 75 Reconstruction of Expression Regulatory Network (REXRN) technology, 255 Resistant Starch (RS), 316 Resveratrol, 316
317 Rhizopus oryzae, 176 lipase, 319 Ribozymes, 7
8 naturally available, 7 Rigidify flexible site (RFS) approach, 75 Rigid linkers, 123
127 RNA
protein complex, 7
8 Root mean square fluctuation (RMSF), 76
77 Rosetta_ddg, 77
78 RosettaVIP, 78
79 R-selective transaminase (R-ATA), 33
34 Rubber transferase, 16

S S1 nuclease, 130
131 Saccharification by enzymatic complexes, 382
383 Saccharomyces cerevisiae, 292
293, 322
323 Screening strategies, for evolved enzymes, 95 Second-generation biofuels, 349
351 Semi-rational design, 108
109 Semi-solid-state fermentation, 367
368 Sequence-based engineering, 72
75 Sequence saturation mutagenesis (SeSaM) method, 70 Serine acetyltransferase, 439 Serine proteases, 21 Short-chain dehydrogenase/reductase (SDRs), 339f Short-chain fatty acids (SCFAs), 313
314 Silica, 197 Simulated protein sequences database (SAliBASE), 105 Sitagliptin, 340
341 Site-directed mutagenesis, 72, 252 Site-saturation mutagenesis, 72 (S)-Licarbazepine, 337
338 Smart polymers, 198 (S)-N-benzyl 3-hydroxypyrrolidine, 156
157 Solid
liquid separation, 37 Solid-state fermentation, 367
368 Spliceosome, 7
8 Splicing, 7
8 Starch, 195 Stearidonic acid, 319 Strain improvement via mutation, 250
256 genetic engineering, 252
256 site-directed mutagenesis, 252 via mutagenesis, 251
252, 251t Streptomyces griseus, 345 Streptomycin sulfate, 38
39 Structure-assisted protein engineering, 56
58

Index

Structure-based engineering, 75
78 Structure-guided design, 105
106 Structure-guided sequence-based engineering, 78 Styrene monooxygenase (SMO)-based cascade, 433
435 Submerged fermentation (SmF), 19 Substrate engineering, 55
56, 56f Sucrose, 289
290 Sulfolobus solfataricus, 92 Supercritical fluid technology, 293 Support matrix system, 198 Surface display systems, 111 Surface-functionalized nanoparticle, 179 Synlinker, 128

T Tagatose, 287
288, 288f Tanacetum parthenium, 343 Tannase, 320 Taxadiene, 343 T-Coffee (Tree-based Consistency Objective Function for Alignment Evaluation), 96
98, 105 Tetrahydrocannabinol (THC), 343 Thebaine, 343 Thermomyces lanuginosus, 315
316, 336 Thermostable enzyme, 67, 70, 72f Thermotoga sp. T. maritima β-glucosidase, 322
323 T. maritima MSB8, 127 T. naphthophila RKU-10, 317 Thioester hydrolases, 17 Third-generation biofuels, 351 Thunnus alalunga, 345 Toxicodendron verniciflua lacquer, 16 Trametes versicolor, 260 Transesterification, 352 Transferases, 16
17, 33 Translocases, 24 Trichoderma sp. T. reesei, 18
19, 248
249, 253 endoxylanase II, 259 M2C38 endoxylanases, 259 T. viride cellulase, 205 Trypsin, 345 Tyrosinases, 16

U Ultrasound-assisted extraction (UAE), 385
386 Unspecific peroxygenase (UPO), 15 Uridine triphosphate, 23

V Verify-3D tool, 96
98

W What-check tool, 96
98 Whole-cell catalysis, 431
432 Whole-cell one-pot transformation processes, 347

X Xylan, 19, 413 sources and structures, 413
414 Xylanases, 19
20, 257
259, 417 from bacterial, archaeal, and fungal origin, 258 genes encoding, 258
259 genetic engineering of, 258
259 of GH10 and GH11 family, 258 patented industrial, 259t types of xylan, 258 Xylanolytic enzymes, 417 Xylooligosaccharides (XOS), 19, 347
348, 414 applications, 419
420 biological properties of, 418
419 antioxidant properties, 419 health benefits of, 419 as nondigestible oligosaccharides, 418 prebiotic effects, 418 market and safety aspects of, 420 prebiotics, 315 production of, 414
418, 415f, 416t purification of, 417
418 Xylose isomerase, 23

Y Yarrowia lipolytica lipase, 200
201 Yellow fluorescent protein (YFP), 136 Yersinia mollaretii phytase (Ymphytase), 70

Z Zeolites, 196

457