Applied Microbiology and Bioengineering [1 ed.] 9780128154076, 9780128155486

Table of contents :
Front Matter
Copyright
Contributors
About the Editor
Author Introduction
Preface
Microbial Tyrosinases: A Novel Enzyme, Structural Features, and Applications
Introduction
Mechanism of Tyrosinase Action
Potential Applications of Tyrosinase
Tyrosinase in Browning Reactions
Tyrosinase in Resistance to Stress and Defense Against Pathogen
Tyrosinase in Bioremediation
Tyrosinase as Biosensor
Tyrosinase in Biosynthetic Processes
l -DOPA Production
Microbial Tyrosinases
Characteristics of Microbial Tyrosinases
Structural and Active Site Characteristics Features of Microbial Tyrosinase
Molecular Characteristics of Microbial Tyrosinase
Conclusions
References
Production of Bioethanol From Sugarcane Bagasse: Current Approaches and Perspectives
Introduction
Sugarcane Bagasse
Various Microorganisms Degrading Bagasse
Pretreatment of Bagasse
Mechanical Comminution
Pyrolysis
Steam Pretreatment or Steam Explosion
Acid Pretreatment
Alkaline Pretreatment
Thermal Pretreatment
Biological Pretreatment
Ammonia Fiber Explosion (AFEX) and Carbon Dioxide (CO 2) Explosion Pretreatment
Wet Oxidation Pretreatment
Ozonolysis Pretreatment
Organosolvent Pretreatment
Cellulose Hydrolysis
Detoxification
Ethanol Production by Fermentation
Application of Bagasse in the Production of Other Value-Added Products
Conclusions
Acknowledgments
Conflict of Interest
References
Further Reading
Improvement of the Amino-Sugar-1-Phosphate Acetyltransferase Activity of the Archaeal Bifunctional Protein
Introduction
Identification of the ST0452 Protein and Its Sugar-1-P NTase Activity
Characterization of the Amsugar-1-P AcTase Activity of the ST0452 Protein
Characterization by Introduction of Substitution Mutations
Characterization via C-Terminal Truncation
Conclusion
Acknowledgments
References
Recent Bio-Processing Technologies for Value Added Horticultural Products
Introduction
Horticultural Produce and Postharvest Losses
Scope and Applications of Bio-Processing
Quality of Processed Products
Conclusion
References
Saccharomyces cerevisiae as Potential Probiotic: Strategies for Isolation and Selection
Introduction
Screening and Characterization of Saccharomyces cerevisiae
Isolation and Characterization
Sources of Saccharomyces cerevisiae
Medium Preparation
Isolation
Determination of Sugar Assimilation Property
Molecular Characterization and Phylogenetic Analysis
In Vitro Studies to Determine the Probiotic Characteristics
Antibacterial and Antifungal Drug Sensitivity
Antagonistic Activity Against Human Pathogens
Co-Culture Activity of Probiotics With Normal Intestinal Flora
Stress Tolerance
Determination of Pathogenicity
Adhesion Assay
Screening for Imunostimulating Properties
Antioxidant Property
Anticancer Properties
Total Glutathione
Free Radical Scavenging Property of Probiotics
Nitric Oxide Scavenging Property of Probiotic Yeast
Hydroxyl Scavenging Property of Probiotic Yeast
Fe2+ Chelating Property of Probiotic Yeast
Splenocyte Proliferation Activity
Advanced Technologies for Selection of Probiotic Saccharomyces cerevisiae
RT-PCR
RT-qPCR
Fluorescence In Situ Hybridization
Fluorescent Activated Cell Sorting (FACS)
Conclusion
Acknowledgment
References
Ecological and Biotechnological Aspects of Methylobacterium mesophilicum
Introduction
General Aspects of Methylobacterium mesophilicum
Methylobacterium mesophilicum Associated With Plants
Conclusions and Future Perspectives
References
Smart Actuators for Innovative Biomedical Applications: An Interactive Overview
Introduction
Smart Actuators
Ionic EAP
Electronic EAP
CNT (Carbon Nano Tube) Based Sensors
Enzyme-Based Biosensor
Integrated Biosensors
Implantable Biosensors
Operating Principle of Implantable Biosensors
Applications of Smart Implanted Biosensor
Smart Textile Biosensor
Applications of Smart Textile Biosensors
Surface Plasmon Resonance
Principle of Surface Plasmon Resonance
Working of Surface Plasmon Resonance
Application of Surface Plasmon Resonance
Applications of Smart Actuators in Biomedical Field
Drug-Delivery Management Using Smart Micropump
Smart Microrobot Arm for Handling of Small Samples
Stepper Motor Actuator for Biomedical Applications Using Ionic Polymer-Metal Composite
Microvalves for Controlled Direction and Delivery of Fluid
Conclusion
References
Recent Developments and Future Prospects of Natural and Synthetic Antitubercular Peptide Drugs
Introduction
Computational and Experimental Approaches to Design Peptide Libraries
In Vitro Techniques
In Silico Techniques: TOOLS, Databases and Webservers for in Silico Design, Screening and Selection of Antimicrobia ...
Types of Peptides and Their Classification
Natural Antitubercular Peptides
Antimycobacterial Peptides Derived From Human Cells
Antimycobacterial Peptides Derived From Microbes and Other Natural Sources
Synthetic/Bioengineered Antitubercular Peptides
Case Studies on Antitubercular Peptides
Applications and Current Developments
Challenges, Limitations and Solutions
Future Directions
Conclusion
Acknowledgments
References
Design and Development of Antibiotic Fermentation Using Different Processing Strategies: Challenges and Perspectives
The Genesis of Penicillin
Penicillin as β-Lactam: Products and Derivatives
Production Strategies for Penicillin
The Challenges of β-Lactam Antibiotics
Cephalosporins
Production Strategies for Cephalosporins
Structural Modifications for Improvement in β-Lactam Antibiotics
Antimicrobial Peptides
Anti-MRSA Daptomycin Production
Production Strategies for Daptomycin
Cofactor(s) Manipulation and Utilization
Precursor and Their Utilization
Significance of Fed Batch Strategy
Metabolic Flux Analysis Model
Morphological Modifications and Immobilization Strategy
Conclusions
References
Further Reading
Current Perspectives and Future Strategies for Fructooligosaccharides Production Through Membrane Bioreactor
Introduction
Market Value of Oligosaccharides
Human Gut Microbiota and Oligosaccharides
Enzymatic Production of Oligosaccharides
Fructooligosaccharide
Production of FOS
Fructosyltransferase
β - d -Fructofuranosidase
Enzymatic Production of FOS
Whole-Cell Synthesis of FOS
Immobilized Biocatalysis (Membrane-Based Bioreactor)
Purification of FOS
Conclusion
Acknowledgment
Conflict of Interest
References
Effectual Bioprocess Development for Protein Production
Introduction
Industrial Hosts Used for Protein Production
E. coli
Yeast/Fungi
Mammalian Cells
Insect Cells
Bioprocess Technologies
Conventional Technologies
Cell-Support Systems
Modern Technologies
Process Analytical Technology
Scale-Out Technology
Types of Bioreactors for Modern Technologies
Wavy Bioreactor
Rocker Bioreactor
Pneumatic Bioreactor
Ready-to-Use Technologies
Use of High Throughput Process Devices
Software
Multivariate Analysis
Sensors
Data Storage
Single-Use Technology
Continuous Processing
Approaches to Economical Production
Future Perspective
Acknowledgment
Conflict of Interest
References
Microalgae: A Way Forward Approach Towards Wastewater Treatment and Bio-Fuel Production
Introduction
Wastewater Treatment With Microalgae and Subsequent Biodiesel Production
Microalgae Harvesting Techniques for Efficient Biofuel Production
Summary
References
Index
A
B
C
D
E
F
G
H
I
J
L
M
N
O
P
Q
R
S
T
U
V
W
X
Y

Citation preview

Applied Microbiology and Bioengineering An Interdisciplinary Approach

Applied Microbiology and Bioengineering An Interdisciplinary Approach

Edited by

Pratyoosh Shukla Department of Microbiology, Enzyme Technology and Protein Bioinformatics Laboratory, Maharshi Dayanand University, Rohtak, Haryana, India

Academic Press is an imprint of Elsevier 125 London Wall, London EC2Y 5AS, United Kingdom 525 B Street, Suite 1650, San Diego, CA 92101, United States 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom © 2019 Elsevier Inc. 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. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN 978-0-12-815407-6 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Andre Gerhard Wolff Acquisition Editor: Kattie Washington Editorial Project Manager: Sam W. Young Production Project Manager: Sreejith Viswanathan Cover Designer: Vicky Pearson Esser Typeset by SPi Global, India

Contributors

Numbers in parentheses indicate the pages on which the authors’ contributions begin.

Pragati Agarwal (3), Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, India Rajeev Agrawal (101), Department of Mechanical Engineering, Malaviya National Institute of Technology, Jaipur, India Archana Anthappagudem (71), Department of Microbiology, University College of Science, Osmania University, Hyderabad, India Welington Luiz Araújo (87), Institute of Biomedical Science, Department of Microbiology, University of São Paulo, São Paulo, Brazil Chiranjib Banerjee (229), Department of Environmental Science and Engineering, Indian Institute of Technology (Indian School of Mines) Dhanbad, Dhanbad, India Srinivas Banoth (71), Department of Microbiology, University College of Science, Osmania University, Hyderabad, India Bhima Bhukya (71), Department of Microbiology, University College of Science, Osmania University, Hyderabad, India Aline Aparecida Camargo-Neves (87), Institute of Biomedical Science, Department of Microbiology, University of São Paulo, São Paulo, Brazil Jong-Chan Chae (21), Division of Biotechnology, Chonbuk National University, Iksan, Republic of Korea Ipsita Chakravarty (163), School of Biochemical Engineering, Indian Institute of Technology (Banaras Hindu University), Varanasi, India Arun Kumar Dangi (203), Enzyme Technology and Protein Bioinformatics Laboratory, Department of Microbiology, Maharshi Dayanand University, Rohtak, India Kashyap Kumar Dubey (185), Department of Biotechnology, University Institute of Engineering and Technology, Maharshi Dayanand University, Rohtak, India; Department of Biotechnology, Central University of Haryana, Mahendergarh, India Shailja Dwivedi (203), Advanced Biotech Lab, Ipca Laboratories Ltd, Mumbai, India Neelam Garg (21), Department of Microbiology, Kurukshetra University, Kurukshetra, India Sanjeev K. Gupta (203), Advanced Biotech Lab, Ipca Laboratories Ltd, Mumbai, India Amandeep Kaur Kahlon (121), Special Centre for Molecular Medicine, Jawaharlal Nehru University, New Delhi, India xi

xii  Contributors Nitish Kaushik (101), Department of Production Engineering, Birla Institute of Technology, Mesra, Ranchi, India Yutaka Kawarabayasi (43), National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba, Japan Ch. Koteswarapavan (101), Department of Production Engineering, Birla Institute of Technology, Mesra, Ranchi, India Punit Kumar (185), Department of Biotechnology, University Institute of Engineering and Technology, Maharshi Dayanand University, Rohtak, India Niwas Kumar (229), Department of Environmental Science and Engineering, Indian Institute of Technology (Indian School of Mines) Dhanbad, Dhanbad, India Vipin Kumar (229), Department of Environmental Science and Engineering, Indian Institute of Technology (Indian School of Mines) Dhanbad, Dhanbad, India Subir Kundu (163), School of Biochemical Engineering, Indian Institute of Technology (Banaras Hindu University), Varanasi, India Kanika Kundu (163), Chemistry Section, MMV, Banaras Hindu University, Varanasi, India Kui-Jae Lee (21), Division of Biotechnology, Chonbuk National University, Iksan, Republic of Korea Prakhar Matre (101), Department of Production Engineering, Birla Institute of Technology, Mesra, Ranchi, India Sikandar I. Mulla (21), Division of Biotechnology, Chonbuk National University, Iksan, Republic of Korea Sangeeta Negi (229), NMC Biolab, New Mexico Consortium, Los Alamos, NM, United States Vinod Kumar Nigam (57), Department of Bio-Engineering, Birla Institute of Technology, Mesra, Ranchi, India Sumedha Ojha (163), School of Biochemical Engineering, Indian Institute of Technology (Banaras Hindu University), Varanasi, India Madhurya Ray (229), Department of Environmental Science and Engineering, Indian Institute of Technology (Indian School of Mines) Dhanbad, Dhanbad, India Ashok Sharma (121), Biotechnology Division, CSIR-Central Institute of Medicinal and Aromatic Plants (CIMAP), Lucknow, India Pratyoosh Shukla (203), Enzyme Technology and Protein Bioinformatics Laboratory, Department of Microbiology, Maharshi Dayanand University, Rohtak, India Mukta Singh (3), Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, India Jyoti Singh (3), Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, India R.P. Singh (3), Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, India

Contributors xiii

Madhu Smita (203), Patent Facilitation Centre (PFC), Technology Information Forecasting and Assessment Council (TIFAC), New Delhi, India Amit Kumar Tiwari (57), Department of Chemical Engineering, Birla Institute of Technology, Mesra, Ranchi, India Shubhandra Tripathi (121), Biotechnology Division, CSIR-Central Institute of Medicinal and Aromatic Plants (CIMAP), Lucknow, India Swati Tyagi (21), Division of Biotechnology, Chonbuk National University, Iksan, Republic of Korea

About the Editor Pratyoosh Shukla, Ph.D., is Professor at and Head of Department of Microbiology, Maharshi Dayanand University, Rohtak, India. He is also General Secretary of Association of Microbiologists of India (AMI), at 75  year’s old it is a world-reknowned scientific organization. His research interests are in the fields of enzyme technology, protein bioinformatics and microbial biotechnology. He has accumulated more than 18  years of research and teaching experience from reputed universities throughout India and postdoctoral experience from Durban University of Technology, Durban, South Africa and has worked as a Research Professor at University of Cincinnati under the Indo-US Science and Technology Forum (IUSSTF) and American Society of Microbiology (ASM) program. He has written 8 books, 22 book chapters, and has published more than 98 peer-reviewed papers in reputed national and international journals. He has received several awards, including: Prof. S.B. Saksena, F.N.A., Award in life sciences (1999); Best Presentation Award (Senior Category-2006) by NCSTC, India; NRF-DUT PDF Award in Enzyme Biotechnology (2008), Danisco, India; an award in probiotics and enzyme technology (2010); Indo-US Professorship Award (2014) by Indo-US Science and Technology Forum (IUSSTF) and American Society of Microbiology (ASM); AMI-Alembic Award (2015); and Bentham Science Ambassador Award (2018). He was also selected as a scientist to participate in the Southern Ocean Expedition (2011) and is a Fellow of Academy of Microbiological Sciences (2017). He is currently serving as Editor for the Indian Journal of Microbiology (Springer), as Associate Editor for BMC Microbiology, as Academic Editor for PLOS One (3Biotech), and as an editorial board member and a reviewer for many other journals. He has also been Editor in Chief for the Internet Journal of Microbiology, Internet Scientific Publishers, USA (2008–2009) and was Special Editor in Chief in JASES, Indonesia for Environmental Biotechnology for a special issue.

xv

Author Introduction R.P. Singh Professor, Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee 247667, Uttarakhand, India E-mail: [email protected] R.P. Singh is senior Professor in the Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, Uttarakhand, India. His research interests are in the areas of: microbial bioconversion for production of value-added products, biomass utilization for energy generation, enzyme engineering, bioremediation, and nanobiotechnology.

Swati Tyagi Division of Biotechnology, Chonbuk National University, Iksan 54597, Republic of Korea E-mail: [email protected] Swati Tyagi graduated in Microbiology from Kurukshetra University (Haryana) in 2012 and specializes in enzyme technology, fermentation, and alcohol technology. Presently she is pursuing her PhD from the Division of Biotechnology, Chonbuk National University, Republic of Korea. Her current research interest is in plant-pathogen interaction and elucidation of the mechanism at molecular level.

xvii

xviii  Author Introduction

Jong-Chan Chae Professor, Division of Biotechnology, Chonbuk National University, Iksan 54597, Republic of Korea E-mail: [email protected] Jong-Chan Chae is a Professor in the Division of Biotechnology, Chonbuk National University, Republic of Korea. His research interest is in the microbial transformation of chemical structures and the elucidation of the mechanism at molecular level.

Yutaka Kawarabayasi Senior Researcher, National Institute of Advanced Industrial Science and Technology (AIST), Higashi 1-1-1, Tsukuba, Ibaraki 305-8566, Japan E-mail: [email protected] Yutaka Kawarabayasi is a Senior Researcher at the Bioproduction Research Institute, National Institute of Advanced Industrial Science and Technology, Tsukuba (Ibaraki), Japan. His research interest is in the identification of useful thermostable enzymes from the genomic data of thermophilic archaea, especially enzymes catalyzing sugar metabolism.

Vinod Kumar Nigam Associate Professor in Department of BioEngineering, Birla Institute of Technology, Mesra, Ranchi 835215, Jharkhand E-mail: [email protected] Vinod Kumar Nigam is working as an Associate Professor in Department of BioEngineering, Birla Institute of Technology, Mesra, Ranchi, Jharkhand, India. He is involved in both teaching and research activities guiding several UG, PG, and PhD students. His areas of research include process biotechnology, enzyme technology, agricultural

Author Introduction xix

biotechnology, food processing, etc. He has several government-funded projects to his credit and has published more than 50 research papers in peer reviewed journals of international repute.

Bhima Bhukya Associate Professor & Head, Department of Microbiology, Osmania University, Hyderabad 500 007, India E-mail: [email protected]; bhima. [email protected] Bhima Bhukya is an Associate Professor of Microbiology in Osmania University. His present research interest lies in microbial biotechnology (probiotics, animal nutrition, biofuel). He has a total 17 years of research and teaching experience including 3  years of undergraduate (veterinary) and 14  years of postgraduate (MSc) teaching. Dr. Bhima is a life member of AMI, ISCA, BRSI, and SAB societies. He was also the recipient of Best Teacher Award (2015) from the state government.

Aline Aparecida Camargo-Neves University of Sao Paulo (USP), São Paulo, São Paulo, Brazil E-mail: [email protected] Aline Aparecida Camargo-Neves is a collaborative researcher at the University of São Paulo, Institute of Biomedical Sciences, Brazil. Her research interest in plant-­ microbe interaction has been applied to biotechnology production.

xx  Author Introduction

Welington Luiz De Araújo University of Sao Paulo (USP), São Paulo, São Paulo, Brazil E-mail: [email protected] Welington Luiz De Araújo is an Associate Professor at the University of São Paulo, Institute of Biomedical Sciences, Brazil. His research interest is in the study of the microbial diversity of different environments, especially in the interior of plants.

Rajeev Agrawal Associate Professor at Department of Mechanical Engineering, Malaviya National Institute of Technology, Jaipur, India Email: [email protected] Rajeev Agrawal has more than 15 years of professional experience. He is actively involved in bringing industry orientation to the engineering education system in India working with several industry and statutory bodies and other organizations. Dr. Agrawal's research work focuses on process-improvement initiatives for productivity enhancement. He has demonstrated his research capabilities in terms of research papers that he has published and presented (more than 60 research papers) and various sponsored and industrial projects. Dr. Agrawal's current research projects include automation in manufacturing, lean six sigma, robotics, supply chain design, and reconfigurable manufacturing systems.

Author Introduction xxi

Amandeep Kaur Kahlon Post-Doctoral Fellow, Special Centre for Molecular Medicine (SCMM), Jawaharlal Nehru University (JNU), New Delhi 110067, India E-mail: [email protected] Amandeep Kaur Kahlon, graduated with a BSc in Life Sciences (2001) with Zoology, Botany and Chemistry from University of Delhi, and an MSc with specialization in Toxicology (2004) from Jamia Hamdard University, New Delhi. She completed her PhD in Biotechnology (2013) at CSIR-Central Institute of Medicinal and Aromatic Plants (CIMAP), Lucknow affiliated to Jawaharlal Nehru University, New Delhi. She worked at the International Centre for Genetic Engineering and Biotechnology (ICGEB) from 2014 to 2016 as a Research Associate. She has postdoctoral work experience in the field of peptide drugs discovery and host-pathogen interactions. Since 2016 she has been working as D.S Kothari Postdoctoral Fellow at SCMM, JNU, New Delhi. Her current research interest is in the discovery, synthesis, and mechanism of interaction of identified peptide inhibitors targeting Mycobacterium tuberculosis.

Subir Kundu Senior Professor and Coordinator, School of Biochemical Engineering, Indian Institute of Technology (Banaras Hindu University), Varanasi, Varanasi 221005, Uttar Pradesh, India E-mail: [email protected]; skundu. [email protected] Subir Kundu is the Senior Professor and the Coordinator of the School of Biochemical Engineering, IIT(BHU), Varanasi, India. He is a Biochemical Engineer and his research contributions span from the development of several life-saving antibiotics, bioenergy, nano-biotechnology, probiotics, to crucial fields of biochemical engineering. He is a pioneer in antibiotic fermentation. His current research focuses on the development of antimicrobial peptides and the ­production of lipopeptide antibiotics to combat drug resistance.

xxii  Author Introduction

Kashyap Kumar Dubey Department of Biotechnology, Bioprocess Engineering Laboratory, Department of Biotechnology, Central University of Haryana, Mahendergarh 123031, Haryana, India E-mail: [email protected] Kashyap Kumar Dubey is working as Associate Professor and Head of the Department of Biotechnology, Central University of Haryana, Mahendergarh (Haryana), India. His research interests are the development of bioprocess technology for the production of secondary metabolites and wastewater treatment.

Punit Kumar Department of Biotechnology, University Institute of Engineering and Technology (UIET), Maharshi Dayanand University, Rohtak 124001, Haryana, India E-mail: [email protected] Punit Kumar is working as a resource person in the Department of Biotechnology, University Institute of Engineering and Technology, MDU Rohtak (Haryana), India. He has completed his PhD from Mararshi Dayanand University Rohtak, Haryana. His research interests are the isolation and purification of biomolecules from microbial sources.

Pratyoosh Shukla Professor and Head, Department of Microbiology, Maharshi Dayanand University, Rohtak 124001, Haryana, India E-mail: [email protected] Pratyoosh Shukla has an MSc in Applied Microbiology Biotechnology from Dr. H.S. Gour University, Sagar, India and a PhD in Microbiology. He is presently working as Professor and Head of the Department of Microbiology, Maharshi Dayanand University, Rohtak, India. He was

Author Introduction xxiii

awarded with Indo-US Visiting research Professorship by the American Society of Microbiology (ASM) in 2014. His current interests are enzyme technology, systems biology, and protein bioinformatics.

Chiranjib Banerjee Assistant Professor (INSPIRE Faculty), Department of Environmental Science & Engineering, Indian Institute of Technology (ISM) Dhanbad, India Email: [email protected]; [email protected] Chiranjib Banerjee has completed his PhD from Birla Institute of Technology, Mesra, India. After completing a PhD in renewable bioenergy, he was awarded the prestigious INSPIRE Faculty award from the Indian government's Department of Science & Technology (DST). Presently he is working as an Assistant Professor at the DST-INSPIRE Faculty in the Department of Environmental Science & Engineering, Indian Institute of Technology (Indian School of Mines), Dhanbad, India. His research interests are in the fields of environmental biotechnology and algal biotechnology.

Vipin Kumar Assistant Professor, Department of Environmental Science & Engineering, Indian Institute of Technology (ISM) Dhanbad, India Email: [email protected] Vipin Kumar is an Assistant Professor in the Department of Environmental Science and Engineering at Indian Institute of Technology (Indian School of Mines), Dhanbad, India. He completed his postgraduate in Microbiology and Doctorate in Environmental Biology. Dr. Kumar's research interests include microbial remediation of pollutants, biological treatment of solid waste, etc. Earlier, he served as a Research Associate and Senior Research Fellow in Indian government's Department of Biotechnology (DBT), New Delhi. He has published many international and national scientific research papers.

xxiv  Author Introduction

Madhurya Ray Junior Research Fellow, Department of Environmental Science and Engineering, Indian Institute of Technology (Indian School of Mines) Dhanbad, India Email: [email protected] Madhurya Ray completed her M. Tech in Environmental Science and Engineering from the Indian Institute of Technology (Indian School of Mines) Dhanbad in 2017. She is currently a junior research fellow (JRF) at the Department of Environmental Science and Engineering, IIT(ISM) Dhanbad. Her research work focuses on the microbial degradation of pollutants.

Niwas Kumar Senior Research Fellow, Department of Environmental Science and Engineering, Indian Institute of Technology (Indian School of Mines) Dhanbad, India Email: [email protected] Niwas Kumar is a senior research fellow (SRF) at the Department of Environmental Science and Engineering, Indian Institute of Technology (Indian School of Mines), Dhanbad. He currently holds a nationallevel scholarship (ICMR). His research work focuses on algal bio-energy.

Sangeeta Negi Research Scientist, New Mexico Consortium, NMC Biological Laboratory, 100 Entrada Drive, Los Alamos, NM 87544, United States Email: [email protected] Sangeeta Negi is currently working as a Research Scientist at the New Mexico Consortium. She completed her PhD in Applied Microbiology and Biotechnology at Dr. Hari Singh Gour University, Sagar, India. She is currently working on different aspects of the efficient functioning of the photosynthetic antenna systems and will

Author Introduction xxv

use this knowledge to increase the photosynthetic efficiency of the green algae Chlamydomonas reinhardtii.

Arun Kumar Dangi Enzyme Technology and Protein Bioinformatics Laboratory, Department of Microbiology, Maharshi Dayanand University, Rohtak, Haryana, India Email: [email protected] Puneet Kumar Singh post-graduated in Biotechnology in 2010 from Kurukshetra University, Kurushetra, and gained his Doctoral Degree in Microbial Biotechnology from Panjab University, Chandigarh, India. His current research interests are in recombinant DNA technology, systems biology, protein engineering, and bioremediation.

Madhu Smita Women Scientist-C, Patent Facilitation Centre, TIFAC, New Delhi, India E-mail: [email protected] Madhu Smita, graduated in Botany, Zoology and Chemistry from Punjabi University, Patiala (Punjab) in 2008 and in 2010 specialized in Industrial Microbiology at the Department of Biotechnology, Thapar University, Patiala (Punjab). She gained her PhD from Panjab University, Chandigarh with the main focus being on both upstreaming and downstreaming processes. Presently she is a Women Scientist in PFC, TIFAC, New Delhi. Her current interest is the amalgamation of IPR in the field of continuous bioprocessing technologies.

xxvi  Author Introduction

Sanjeev K. Gupta General Manager and Head-Advanced Biotech Lab, Ipca Laboratories Ltd., Plot#125, Kandivli Ind. Estate, Kandivli (West)-400067, Mumbai-India Email: [email protected] Sanjeev K. Gupta has gained a PhD in Microbiology (cell engineering) and an MSc in Applied Microbiology and Biotechnology. At present he is the General Manager and Head at the Advanced Biotech Lab (R&D-Biosimilar), Ipca Laboratories Ltd., Mumbai, India. He has 18 years of industrial experience and has been working since 2000 on the development of “Biosimilars” including monoclonal antibodies. His core expertise lies into molecular biology, cell line, process development for recombinant therapeutic molecule development.

Shailja Dwivedi Research Executive-Advanced Biotech Lab, Ipca Laboratories Ltd., Plot#125, Kandivli Ind. Estate, Kandivli (West)-400067, Mumbai, India Email: [email protected] Shailja Dwivedi has gained a PhD in Biotechnology (Metabolic engineering) and an MSc in Biotechnology. At present she is working as a Research Executive at the Advanced Biotech Lab (Biosimilar), Ipca Laboratories Ltd., Mumbai, India. She has 3 years of industrial experience and is working on the development of “Biosimilars”, including monoclonal antibodies. Her core expertise lies in the molecular biology, cell line engineering, and cell banking and characterization.

Preface This book is on interdisciplinary applied microbiology and b­ ioengineering and contains an outstanding compilation of the various research aspects of ­ microbiology and bioengineering that highlight recent advances and ­innovations. It presents the most interesting topics in modern b­ iotechnology, which are d­ivided into three sections: enzymes in bioprocessing; human health, microbial physiology and biomedical applications; and bioprocess technology. The most significant attribute of this book is that it presents the most upto-date areas of microbiology with special emphasis on the biotechnological advancements in interdisciplinary research into microbiology with other subdisciplines of bioengineering. The first section of this book inclines toward enzyme production and enzyme engineering. The second section includes some of the most important current topics, such as probiotics, smart actuators for biomedical application, and the natural and synthetic antitubercular peptide drugs that have become vital for human health. The last section covers some of the most remarkable aspects of current microbiology, such as bioprocess technologies for biofuel, therapeutic proteins, synbiotic foods, and value-added horticulture products. This book will be a valuable resource for senior undergraduate and graduate students, researchers, professionals and other interested individuals or groups working in the interdisciplinary areas.

xxvii

Chapter 1

Microbial Tyrosinases: A Novel Enzyme, Structural Features, and Applications Pragati Agarwal, Mukta Singh, Jyoti Singh, R.P. Singh Department of Biotechnology, Indian Institute of Technology Roorkee, Roorkee, India

1. INTRODUCTION Tyrosinase (EC 1.14.18.1) is a copper containing metalloenzyme and is widely distributed throughout the phylogenetic scale [1–3]. The enzyme is almost universally distributed through every lifeform from bacteria to mammals [4], as shown in Fig. 1. The enzyme is a member of a large family of proteins termed type-3 copper proteins and is a key protein primarily involved in the initial steps of the melanin biosynthesis pathway [5–7]. Other important members of this family are catecholoxidase, phenoloxidase, and hemocyanin, and all are vital for numerous important physiological functions in animals, plants, fungi, and bacteria. Tyrosinase occurs in mammals, invertebrates, plants, and microorganisms, where it is involved in several critical biological functions, such as skin pigmentation in mammals, the primary immune response and wound-healing systems in plants and fungi [8,9], the browning reactions in fruits and vegetables, the host defense system in arthropods, the differentiation of reproductive organs and spore formation in fungi [1,10–12], and the sclerotization of the cuticle in arthropods after molting or injury. Tyrosinase is a multifunctional metalloenzyme that catalyzes the o-hydroxylation of monophenols to o-diphenols (cresolase or monophenolase activity) and the subsequent oxidation of the latter to reactive o-quinones (catecholoxidase or diphenolase activity) using oxygen [13]. Further, these o-quinones polymerize into brown-black pigments through a series of enzymatic and nonenzymatic reactions [8,14,15]. These two activities are the basis for the widespread biotechnological and industrial applications of tyrosinase, for instance biosensors for the monitoring of the phenolic content in wastewater [16,17] and food products; in environmental technology for the bioremediation of phenol-containing wastewater and contaminated soils [18,19]; in pharmaceutical industries for the biotransformation of l-tyrosine to Applied Microbiology and Bioengineering. https://doi.org/10.1016/B978-0-12-815407-6.00001-0 © 2019 Elsevier Inc. All rights reserved.

3

4  SECTION | I  Enzymes in Bioprocessing

Enzymatic browning of fruits

Melanin formation in fungi L-Tyrosinase

Enzymatic browning of vegetables

Melanin formation in human

FIG. 1  Ubiquitous distribution of tyrosinase.

l-DOPA, which is the preferred drug for Parkinson’s disease [20,21]; and in the cosmetic and food industries due to either undesirable or beneficial oxidative browning reactions [3].

2.  MECHANISM OF TYROSINASE ACTION Tyrosinase catalyzes the conversion of l-tyrosine to l-DOPA and then to dopachrome, which is subsequently polymerized spontaneously to melanin via a series of reactions [7,22]. Thus tyrosinase is a bi-functional enzyme catalyzing the o-mono-hydroxylation of monophenols (phenols) to their corresponding o-diphenols (o-cresolase or monophenolase) and their subsequent oxidation (catechol oxidase or diphenolase) into reactive o-quinones [6,11,12]. Molecular oxygen is used as an electron acceptor and it is reduced to water in both the reactions. Subsequently, the resulting o-quinones undergo nonenzymatic oxidoreduction reactions with various nucleophilic moieties, producing intermediates that auto-polymerize spontaneously in dark-brown pigments, such as melanins [10,23], as demonstrated in Fig. 2 The monophenolase activity is the most critical and is the initial rate-determining reaction in the process [24]. The catalytic mechanism of type-3 copper proteins has been studied extensively for the past two decades, but the precise reaction mechanism of tyrosinase action is still unclear. Using the previous structural studies as a basis it has been hypothesized that to initiate the reaction requires dioxygen moiety, which links the two copper centers by forming peroxide linkage. Next, the hydroxyl group of the substrate is proposed to be deprotonated by the peroxide

Microbial Tyrosinases  Chapter | 1  5 COOH HO

NH2

Tyrosine

HO

COOH NH2

HO Tyrosinase (Monophenolase)

O

COOH NH2

O Tyrosinase (Diphenolase)

DOPA

Dopaquinone

Tyrosinase

HO

O

Melanin HO Polymerization

N H

O

N H

COOH

Dopachrome FIG. 2  Catalytic cycle of tyrosinase.

ion bound between CuA and CuB at the start of the reaction. Subsequently, an ortho‑carbon atom of the substrate approaches the peroxide and the substrate is ortho-hydroxylated. Finally, the diphenolic compound adduct undergoes oxidation in the presence of second oxygen, which results in an o-quinone product [25]. The other hypothesis states that the dinuclear copper center of the enzyme binds with one molecule of atmospheric oxygen [1,2], and inserts it in a position ortho to an existing hydroxyl group onto the aromatic ring (electrophilic attack) followed by the oxidation of the diphenol to the corresponding quinone [26]. This results in a reduced copper content being available for phenol and cosubstrate dioxygen binding and for further turnover. Thus during the catalytic cycle the type-3 copper center of tyrosinase can adopt three different functional forms: the oxy-state [Cu(II)―O2― Cu(II)], deoxy-state [Cu(I)•Cu(I)] and the met-state [Cu(II)―OH― OH―Cu(II)]. The valences of the two copper atoms change from Cu(I) to Cu(II) reversibly, i.e., it gets oxidized and then again gets reduced. In the oxy-state the molecular oxygen is reversibly bound as peroxide between the two copper atoms. In the met-state the two copper atoms are bridged through hydroxo ions. According to present conception both the met-state and the oxy-state of tyrosinase enable the diphenol oxidase activity whereas the mono-hydroxylase reaction requires the oxy-state [8]. Met tyrosinase cannot hydroxylate monophenols. A two-electron reduction yields deoxy-tyrosinase, which consecutively binds dioxygen to its binuclear copper site giving rise to the active oxy-form. Then one Cu(II) of the oxy-tyrosinase copper pair binds the monophenol hydroxyl group. This binding step is followed by the ortho hydroxylation of the substrate. Thus both met- and oxy-tyrosinase can oxidize o-diphenol into o-quinone while the enzyme is recovered as the deoxy- or met-form respectively. Since deoxy-tyrosinase is transformed into the oxy form by dioxygen, thus 2 moles of o-diphenol can be oxidized into o-quinone through a cyclic process, at the cost of one molecule of dioxygen [27]. In

6  SECTION | I  Enzymes in Bioprocessing

the tyrosinase-catalyzed reactions a lag phase for the oxidation reaction of monophenols is generally present, during which the oxy-form of tyrosinase is generated from the met-form [28].

3.  POTENTIAL APPLICATIONS OF TYROSINASE The capability to catalyze the hydroxylation of monophenols and to oxidize diphenols make tyrosinase a potentially attractive source for several biotechnological and industrial applications [29]. The enzyme had demonstrated its effectiveness in environmental technology for the detoxification of wastewater and soil that are contaminated with phenolic compounds [19,30–32]; as biosensors for the monitoring of phenols; in pharmaceutical industries for the synthesis of l-DOPA from l-tyrosine, which is the drug of choice for the treatment of Parkinson’s disease; and in cosmetic and food industries for the biosynthesis of melanin and other phenolic polymers, such as lignin, flavonoids, and tannin [6,7,11], etc., as discussed in Fig. 3. Presently, the immobilized tyrosinase appears to be a promising candidate for the removal of phenols from industrial wastewaters [33,34].

3.1  Tyrosinase in Browning Reactions Enzymatic browning is a consequence of the tyrosinase-catalyzed oxidation of phenolic substrates into quinones and these reactive quinones undergo further reactions (oxidations and polymerizations) spontaneously leading to high molecular mass; the dark pigments called melanins, which account for skin and hair pigmentation; undesirable fruit or vegetable browning [35]; wound healing in plants and arthropods; and enzymatic browning following bruising, cutting, or other injury to the plant cell [1]. Biormediation of toxic phenolic compounds

L-DOPA therapy for Parkinson’s treatment

Tyrosinase Defense responses in fungi

Biosensing of phenolic compounds Melanin biosynthesis FIG. 3  Potential industrial applications of tyrosinase.

Microbial Tyrosinases  Chapter | 1  7

3.2  Tyrosinase in Resistance to Stress and Defense Against Pathogen Due to their fungistatic, bacteriostatic, and antiviral properties, melanin and its intermediates are the key components of wound healing and the primary immune response of certain invertebrate phyla, especially arthropods [36,37], plants, and fungi. In plants, brown polyphenolic catechol melanins [38] are assumed to protect the damaged plant part from pathogens and insects [39,40]. In insects and other arthropods tyrosinase is involved in sclerotization of the exoskeleton after molting or injury and in protection against other organisms by encapsulating them in melanin [41]. Fungal pigments are involved in the development and stability of spores [3].

3.3  Tyrosinase in Bioremediation The presence of phenols and dyes in ecosystems presents serious environmental and health concerns. For several years the use of oxidizing enzymes from fungi has been recognized as an eco-friendly biological solution for the treatment of wastes containing toxic phenolic compounds [14,42]. Tyrosinase has been demonstrated to remove these toxic compounds, for instance chlorophenol, fluorophenol, and catechols, from the environment by oxidizing them. In a general manner, tyrosinase requires only molecular oxygen as a cofactor for their detoxification [14]. The activation of phenols by tyrosinase tends to lead to their polymerization and subsequent precipitation allowing their easy removal during waste-water treatment. Purified preparations of mushroom tyrosinase had been shown to be effective in the precipitation and biotransformation of aqueous phenols [33] and chlorophenols [30]. The extracellular tyrosine-inducible tyrosinase from the Zygomycete Amylomyces rousii had been applied for the efficient degradation of pentachlorophenol, a recalcitrant xenobiotic used as a pesticide [19]. Tyrosinase has also been suggested as playing a role in the degradation of dyes such as Direct Orange 39, Reactive Yellow 107, Reactive Red 198, Reactive Black 5 and Direct Blue 71, discharged mainly in textile wastewaters.

3.4  Tyrosinase as Biosensor Biosensors based on tyrosinase activity could be used for detecting phenol concentrations in industrial effluents, food products, and cosmetics, etc. Phenols, especially chlorophenols and fluorophenols, have been defined as dangerous pollutants due to their toxicity and longer existence in the environment. Phenol is a widely exercised chemical and byproduct of numerous industries. Waste phenols are primarily released into the environment by the resins industry, oil refineries, wood-pulp industry (as they are vital components of natural compounds, e.g., lignin and tannin), and as a result of the degradation of pesticides with phenolic skeleton. Thus for the analysis of these noxious phenolic derivatives, biosensors based on tyrosinase are gaining popularity as an

8  SECTION | I  Enzymes in Bioprocessing

a­ lternative technique. Tyrosinase is capable of oxidizing phenols and thus can detect different types of monophenolic and diphenolic compounds. Presently, tyrosinases from various sources are being immobilized onto different support matrices to facilitate the biosensing of phenols, e.g., Seo et al. [43] had developed a biosensor with immobilized mushroom tyrosinase, enabling the quantification of phenols and the elimination of related compounds in water and soil samples. Several other workers have also immobilized tyrosinase onto ZnO nanoparticles [17] and polystyrene microplate [44] in order to use it as biosensor.

3.5  Tyrosinase in Biosynthetic Processes Tyrosinase is a vital enzyme for the biosynthesis of phenolic polymers such as lignin, flavonoids, melanin, and tannin, and for l-DOPA, which is a potent drug for the treatment of Parkinson’s disease and for myocardium neurogenic injury. Tyrosinase is the key regulatory enzyme involved in the initial rate-limiting step of the biosynthesis of the polyphenolic melanin pigments that are distributed in all living organisms. Melanins synthesized by tyrosinase action have numerous applications, e.g., as protectives against free radicals, gamma rays, dehydration, and extreme temperatures, and they act as cation exchangers and carriers for drugs, antioxidants, antiviral agents, and immunogens [22,45]; they also contribute to the fungal spore formation and in cell-wall resistance against hydrolytic enzymes. In mammals melanogenesis is responsible for skin, eye, and hair pigmentation, and for protecting the skin via absorption of UV radiation [46,47].

3.5.1  l-DOPA Production l-DOPA is an amino acid analogue and precursor of dopamine that is used as the most potent symptomatic drug treatment for Parkinson’s disease (PD), a degenerative disorder that is primarily associated with diminished levels of the neurotransmitter dopamine (DA) in the brain. This may be because either it is not being synthesized or it is being produced at critically lower levels by the dopaminergic neurons of the brain, alternatively the dopamine may be malfunctioning. The reduced levels of dopamine causes rigidity, tremors, slowness of speech and eventually dementia [48]. PD is a long-term disorder of the central nervous system that mainly affects the motor system. It is the second most common neurodegenerative disorder after Alzheimer's disease and was present in 53 million people and resulted in about 103,000 deaths globally in 2013 (Global Burden of Disease Study 2013). The average life expectancy following the diagnosis of this chronic and progressive movement disorder is between 7 and 14 years [49]. Although its symptoms are related to the depletion of dopamine in corpus striatum, the administration of dopamine is ineffective in its treatment as it

Microbial Tyrosinases  Chapter | 1  9

cannot cross the blood-brain barrier. Levodopa has been the most widely used treatment for over 30 years. Fig. 4 represents the dopaminergic synapse and the action of l-DOPA. Dr. William S. Knowles, a Nobel laureate (2001), demonstrated for the first time the asymmetric synthesis of l-DOPA in 1977, which is now called the Monsanto process [50]. l-DOPA is currently synthesized chemically for pharmaceutical use [51], which involves multiple steps that are time consuming, eco-unfriendly and uneconomical. In contrast to chemical production, biotechnological production of l-DOPA is an eco-friendly process and results in enhanced recovery of the product using a relatively simple process. l-Tyrosine is a precursor of l-DOPA and l-DOPA is biotechnologically synthesized from l-tyrosine by a one-step process of oxidation catalyzed by tyrosinase [52]. Few attempts have been made to produce l-DOPA from fungal species such as Aspergillus oryzae [20,53], Acremonium rutilum [21] and Penicillium jensenii [54].

L-Tyrosine

Dopaminergic synapse

L-DOPA

Dopamine Presynaptic terminal Autoreceptor

Dopamine degradation

COMT

Dopamine transporter

MAO Postsynaptic membrane Dopamine receptor Signal transduction

FIG. 4  Dopaminergic synapse, the principal site of action of current Parkinson’s disease treatments.

10  SECTION | I  Enzymes in Bioprocessing

4.  MICROBIAL TYROSINASES Few studies have been carried out to investigate the molecular and functional characteristics of tyrosinase from various prokaryotic and eukaryotic sources. The commercially available tyrosinase is isolated from edible mushroom, i.e., Agaricus bisporus, and from Streptomyces species. Bacterial tyrosinases have been studied mainly in Pseudomonas, Bacillus, Myrothecium, Miriococcum, and Streptomyces [55]. Streptomyces tyrosinases are the most thoroughly characterized tyrosinase of bacterial origin [56]. The first bacterial tyrosinases were purified from the cell extracts of Streptomyces nigrifaciens [57] and Streptomyces glaucescens [58]. The Streptomyces tyrosinase is a monomeric extracellular protein with a low molecular weight of 30 kDa and is involved in melanin production. A heat-inducible tyrosinase from Bacillus thuringiensis with a molecular weight of 14 kDa has been purified [59] and it had one of the lowest molecular masses among all the known tyrosinases from various sources. A thermostable tyrosinase with optimal activity at 70°C and pH 9.5 has been isolated from Thermomicrobium roseum [60]. The first crystal structure of tyrosinase was determined from Streptomyces castaneoglobisporus and was complexed with the caddie protein at 1.2–1.8 Å resolution by Matoba et al. [56]. Afterwards a tyrosinase structure was determined from Bacillus megaterium at a resolution of 2.0–2.3 Å, which is represented in Fig. 5. This revealed the presence of two Cu(II) ions aiding as the major cofactors within the active site, which are coordinated by six conserved histidine residues. Tyrosinases have been isolated and characterized from fungi Neurospora crassa [5,61], A. bisporus [62–65], Lentinula edodes [66], Ascovaginospora [67], Trametes, white-rot fungus Pycnoporus [10,68], Trichoderma [69], A. rutilum [21], A. oryzae [6,7,70], P. jensenii [54], and Aspergillus niger [71–73]. Among these, tyrosinases from A. bisporus, N. crassa, and A. oryzae are the most extensively investigated fungal tyrosinases [11], from both structural and

FIG. 5  Crystal structure of Bacillus megatarium and Agaricus bisporus.

Microbial Tyrosinases  Chapter | 1  11

functional points of view. The tyrosinase-encoding gene had been cloned from fungi such as A. oryzae [6,7], Pycnoporus sanguineus [10], A. rutilum [21], and P. jensenii [54]. Tyrosinase characterization from the edible mushroom A. bisporus led to new insights into its biochemical and molecular characteristics. The tyrosinase-encoding cDNA of A. bisporus was cloned and characterized using primers designed on the basis of sequence homologies in the copper-binding domains from a number of plant and fungal tyrosinases [65]. Obata et al. [6] had cloned a novel tyrosinase-encoding gene (melB) from A. oryzae in pUC119 and overexpressed it in pGMB1 vector. Rao et al. [7] had cloned and overexpressed tyrosinase from A. oryzae in Yarrowia lipolytica. Halaouli et al. [10] had cloned and characterized a tyrosinase gene from the P. sanguineus and overexpressed in A. niger. The crystal structure of tyrosinase from A. bisporus was determined at 2.3 Å resolution in deoxy-form [25] as shown in Fig. 5, while the structure of A. oryzae was solved by Fujieda et al. [74]. Sanchez-Ferrer et al. [4] showed its involvement in the melanin synthesis pathway and that it was capable of bringing about biotransformation of a single amino acid l-tyrosine to a biologically active molecule l-DOPA by hydroxylation reaction. The transformation of l-tyrosine to l-DOPA was also achieved by A. rutilum, which was isolated from a decomposed banana stud [21]. Rao et al. [7] cloned and overexpressed tyrosinase from A. oryzae in Yarrowia lipolytica and observed that the clone overexpressing the tyrosinase was able to transform l-tyrosine to l-DOPA in the reaction mixture. P. jensenii, isolated from the soil was found to be a potent tyrosinase producer under shake-culture conditions and was capable of bringing about biotransformation of l-tyrosine to l-DOPA [54].

5.  CHARACTERISTICS OF MICROBIAL TYROSINASES 5.1  Structural and Active Site Characteristics Features of Microbial Tyrosinase The structural data on microbial tyrosinases have been limited and the first highresolution three-dimensional crystal structure of tyrosinase was determined from Streptomyces castaneoglobisporus complexed with the caddie protein at 1.2–1.8 Å resolution by Matoba [56]. Tyrosinase is considered as a metalloenzyme [75] having a diamagnetic spin-coupled copper pair in the active center [76], with copper being very compactly bound to the apoprotein. Tyrosinase is abundant in mammals, plants, fungi, and bacteria, and there are a few universal structural determinants present in all tyrosinases; the di‑copper-binding ligands and other active-site residues establish significant interactions to uphold the globular folding. Each copper ion is coordinated by a peptide motif consisting of three conserved histidine residues (His) [77] thus called binuclear type-3 copper center as demonstrated in Fig. 6. This copper pair, called CuA and CuB, present in the holo-enzyme is the site at which tyrosinase interacts with both

12  SECTION | I  Enzymes in Bioprocessing

FIG. 6  (A) Superimposed tertiary structures of Bacillus megatarium (cyan) and Agaricus bisporus (magenta), (B) alignment of the dinuclear copper center of B. megatarium and A. bisporus tyrosinase. Spheres represent Cu-A and Cu-B copper ions at active site. Each of the two copper atoms (brown) is coordinated by three histidines. Structures are generated using chimera (http:// www.cgl.ucsf.edu/chimera/).

molecular oxygen and its phenolic substrates. Each atom of the binuclear copper cluster is identified to form ligands with three nitrogen atoms provided by histidine side-chains [78]. This active site structure with each of the two copper ion bound by imidazole rings of three histidine residues is the most prominent attribute and is well conserved throughout the phylogenetic range [35,79–82]. When sequences of tyrosinases from bacteria and from funguses are aligned and compared, the central copper-binding domains, i.e., two histidine triads, were the sites that were absolutely conserved, as specified in Fig.  7. Structural comparison of these two tyrosinases further depicted the complete

FIG.  7  Sequence alignment of tyrosinases from Bacillus megaterium (GenBank accession ACC86108.1) and Agaricus bisporus (GenBank accession CAA11562.1) indicating conserved histidines (red dot). Alignment is generated using ClustalW (http://www.ebi.ac.uk/Tools/msa/clustalw2/).

Microbial Tyrosinases  Chapter | 1  13

correspondence of these histidines to form active site domains (Fig. 6). In this arrangement the overall configuration of a tyrosinase active site is effectively maintained by electrostatic and cation-π interactions between the helical segments [82]. The second His at the CuA motif is the only flexible because it is located in a loop structure in the protein, whereas the other five His are located in the surrounding α-helical fragments [83,84]. Principally, this complete catalytic center consists of a hydrophobic pocket within a helix bundle comprising two densely packed antiparallel α-helix pairs. This four α-helix bundle motif bears the active site along with the two histidine-coordinated copper atoms [82]. In the inactive form of the enzyme, the antiferromagnetically coupled copper-ions pair is penta-coordinated in a distorted square pyramidal geometry. Depending on the copper-ion valence and bond with molecular oxygen, three different intermediate states of the tyrosinase active site have been reported: the oxidized Cu(II) containing met-form (CuII―CuII), the oxygenated oxy-form (CuII―O2―CuII) and the reduced Cu(I) containing deoxy-form (CuI― CuI) [79]. The latent form of tyrosinase is found to be a combination of 85% met- and 15% oxy-forms [85]. In the oxy-form molecular oxygen is proposed to be coordinated as peroxide in a side-on bridging between the two copper atoms. It has been suggested that monophenols would dock to CuA and o-diphenols would dock to CuB at the tyrosinase active site [80]. Oxygen binding induces alteration in the valency of the copper atoms, which are in the Cu(I) state in the deoxy form but become Cu(II) upon oxygenation. This change results in the blue color developed by several type-3 copper proteins upon oxygenation, such as in hemocyanin. Oxy-tyrosinase sequentially decays back to met-tyrosinase when the peroxide is lost in the reaction. The met-form is converted into deoxyform by a two-electron reduction and the resultant deoxy-form reversibly fixes molecular oxygen leading to the oxy-form. In the absence of any substrate met is the primary form. The deprotonated monophenol binds to CuB, followed by the addition of oxygen to the ring in ortho position through an association to CuA, enabled by the liberation of the flexible His [56]. This residue has also been proposed to be involved in the proton shift, i.e., deprotonation of the monophenolic substrate [82,86].

5.2  Molecular Characteristics of Microbial Tyrosinase The molecular weight of bacterial and fungal tyrosinase shows considerable diversity. Early studies on the tyrosinase holo-enzyme from the common mushroom A. bisporus suggested that the enzyme is a hetero-tetramer H2L2 of 569 amino acids, consisting of two heavy polypeptide chains and two light chains encoding a protein with a molecular weight of 64 kDa [65]. In the case of N. crassa tyrosinase, the mature protein (46 kDa) is 407 amino acids long, which results from the cleavage of 213 amino acids (21 kDa) from the C-terminal end of a pro-tyrosinase 620 amino acids long. Table 1 shows the main molecular features of several fungal and bacterial tyrosinases. The Streptomyces tyrosinase is

14  SECTION | I  Enzymes in Bioprocessing

TABLE 1  Salient Characteristic of Microbial Tyrosinases Source

No. of amino acid residues

Molecular Protein mass (KDa) accession no. Reference

Agaricus bisporus

569

64

CAA11562.1

Wichers et al. [65]

Neurospora crassa

620

65

M33271

Kupper et al. [5]

Aspergillus orvzae

539

67

D37929

Fujita et al. [87]

Lentinula edodes

618

70

BAB71736.1

Kanda [66]

Pycnoporus sanguineus

618

75

AAX46018.1

Halaouli [68]

Trichoderma reesei

561

61

CAL90884.1

Selinheimo [69]

Marinomonas mediterranea

675

75

AAF7583

Lopez-Serrano et al. [88]

Rhizobium etli

609

68

NP659960

Lerch [2]

Ralstonia solanacearum

412

44

NP519622

Gelder et al. [1]

Bacillus thuringiensis

151

17

AAW29015

Lerch and Ettlinger [58]

Streptomyces antibioticus

272

31

P07524

Bernan et al. [89]

Streptomyces griseus

306

36

Q9ZN72

Endo et al. [90]

a monomeric extracellular protein with a low molecular weight of 30 kDa and a heat-inducible tyrosinase from B. thuringiensis with a molecular weight of 14 kDa has been purified [59] which has one of the lowest molecular masses among all the known tyrosinases from various sources.

6. CONCLUSIONS Tyrosinases are exploited for a variety of biotechnological and environmental applications and thus have attracted various groups actively engaged in molecular characterization and bioengineering studies. The present article has summarized the key structural, molecular, and functional features of tyrosinases derived from various bacterial and fungal species. Recent findings have revealed several important insights into the structural and molecular characteristics of the microbial tyrosinases and have highlighted their resemblance to other type-3 copper metalloproteins. Further, elucidation of an insight into its biochemical

Microbial Tyrosinases  Chapter | 1  15

characterization may enable the development of a simpler and more industrially feasible microbial system for achieving higher yields and more cost-effective production of tyrosinase and for exploring its potential in food, pharmaceutical, and environmental sectors.

REFERENCES [1] Gelder CWV, Flurkey WH, Wichers HJ. Sequence and structural features of plant and fungal tyrosinases. Phytochemistry 1997;45:1309–23. [2] Lerch  K. Tyrosinase: molecular and active-site structure. ACS Symp Ser 1995;600: 64–80. [3] Mayer AM, Harel E. Polyphenol oxidases in plants. Phytochemistry 1978;18:193–215. [4] Sanchez-Ferrer  A, Laveda  F, Garcla-Carmona  F. Partial purification of soluble potato polyphenol oxidase by partitioning in an aqueous two-phase system. J Agric Food Chem 1993;41:1219–24. [5] Kupper  U, Niedermann  DM, Travaglini  G, Lerch  K. Isolation and characterization of the tyrosinase gene from Neurospora crassa. J Biol Chem 1989;264:17250–8. [6] Obata H, Ishida H, Hata Y, Kawato A, Abe Y, Akao T, Akita O, Ichishima E. Cloning of a novel tyrosinase-encoding gene (melB) from Aspergillus oryzae and its over expression in solid-state culture (rice koji). J Biosci Bioeng 2004;97:400–5. [7] Rao A, Pimprikar P, Bendigiri C, Kumar AR, Zinjarde S. Cloning and expression of a tyrosinase from Aspergillus oryzae in Yarrowia lipolytica: application in L-DOPA biotransformation. Appl Microbiol Biotechnol 2011;92:951–9. [8] Claus H, Decker H. Bacterial tyrosinases. Syst Appl Microbiol 2006;29:3–14. [9] Muller WE, Grebenjuk VA, Thakur NL, Thakur AN, Batel R, Krasko A, Muller IM, Breter HJ. Oxygen-controlled bacterial growth in the sponge Suberites domuncula: toward a molecular understanding of the symbiotic relationships between sponge and bacteria. Appl Environ Microbiol 2004;70:2332–41. [10] Halaouli S, Record E, Casalot L, Hamdi M, Sigoillot JC, Asther M, Lomascolo A. Cloning and characterization of a tyrosinase gene from the white-rot fungus Pycnoporus sanguineus and overproduction of the recombinant protein in Aspergillus niger. Appl Microbiol Biotechnol 2006;70:580–9. [11] Konishi YK, Tsuji M, Hatana S, Asanuma M, Kakuta D, Kawano T, Mukouyama EB, Goto H, Suzuki H. Purification, characterization, and molecular cloning of tyrosinase from Pholiota nameko. Biosci Biotechnol Biochem 2007;71:1752–60. [12] Yi LN, Ming CW, Li JQ, Ping QQ, Lai RF. Molecular cloning and expression of polyphenoloxidase genes from the mushroom, Agaricus bisporus. Agric Sci China 2011;10:185–94. [13] Labus K, Turek A, Liesiene J, Bryjak J. Efficient Agaricus bisporus tyrosinase immobilization on cellulose-based carriers. Biochem Eng J 2011;56:232–40. [14] Burton SG. Oxidizing enzymes as biocatalysts. Trends Biotechnol 2003;21:543–9. [15] Wan X, Chai B, Liao Y, Su Y, Ye T, Shen P, Chen X. Molecular and biochemical characterization of a distinct tyrosinase involved in melanin production from Aeromonas media. Appl Microbiol Biotechnol 2009;82:261–9. [16] Atlow  SC, Banadonna-Aparo  L, Klibanov  AM. Dephenolization of industrial wastewaters catalyzed by polyphenol oxidase. Biotechnol Bioeng 1984;26:599–603. [17] Li YF, Liu ZM, Liu YL, Yang YH, Shen GL, Yu RQ. Mediator-free phenol biosensor based on immobilizing tyrosinase to ZnO nanoparticles. Anal Biochem 2006;349:33–40.

16  SECTION | I  Enzymes in Bioprocessing [18] Claus H, Filip Z. Behavior of phenoloxidases in the presence of clays and other soil-related adsorbents. Appl Microbiol Biotechnol 1988;28:506–11. [19] Montiel  AM, Fernandez  FJ, Marcial  J, Soriano  J, Barrios-Gonzales  J, Tomasini  A. A fungal phenoloxidase (tyrosinase) involved in pentachlorophenol degradation. Biotechnol Lett 2004;26:1353–7. [20] Ali S, Shultz JL, Haq I. High performance microbiological transformation of L-tyrosine to L-DOPA by Yarrowia lipolytica NRRL-143. BMC Biotechnol 2007;7:50. [21] Krishnaveni  R, Rathod  V, Thakur  MS, Neelgund  YF. Transformation of L-tyrosine to L-DOPA by a novel fungus, Acremonium rutilum under submerged fermentation. Curr Microbiol 2009;58:122–8. [22] Wang G, Aazaz A, Peng Z, Shen P. Cloning and over expression of a tyrosinase gene mel from Pseudomonas maltophila. FEMS Microbiol Lett 2000;185:23–7. [23] Soler-Rivas C, Jolivet S, Arpin N, Olivier JM, Wichers HJ. Biochemical and physiological aspects of brown blotch disease of Agaricus bisporus. FEMS Microbiol Rev 1999;23:591–614. [24] Rodriguez-Lopez JN, Tudela J, Varon R, Garcia-Carmona F, Garcia-Canovas F. Analysis of a kinetic model for melanin biosynthesis pathway. J Biol Chem 1992;267:3801–10. [25] Ismaya  WT, Rozeboom  HJ, Weijn  A, Mes  JJ, Fusetti  F, Wichers  HJ, Dijkstra  BW. Crystal structure of Agaricus bisporus mushroom tyrosinase: identity of the tetramer subunits and interaction with Tropolone. Biochemistry 2011;50:5477–86. [26] Mayer AM. Polyphenol oxidases in plants and fungi: going places? A review. Phytochemistry 2006;67:2318–31. [27] Jolivet J, Arpin N, Wichers HJ, Pellon G. Agaricus bisporus browning: a review. Mycol Res 1998;102:1459–83. [28] Cooksey CJ, Garratt PJ, Land EJ, Pavel S, Ramsden CA, Riley PA, Smit NP. Evidence of the indirect formation of the catecholic intermediate substrate responsible for the auto-activation kinetics of tyrosinase. J Biol Chem 1997;272:26226–35. [29] Sanchez-Amat A, Lucas-Elio P, Fernandez E, Garcia-Borron JC, Solano F. Molecular cloning and functional characterization of a unique multipotent polyphenol oxidase from Marinomonas mediterranea. Biochim Biophys Acta 2001;1547:104–16. [30] Ikehata K, Nicell JA. Color and toxicity removal following tyrosinase-catalysed oxidation of phenols. Biotechnol Prog 2000;16:533–40. [31] Lee  SG, Rao  HS, Hong  SP, Kim  EH, Sung  MH. Production of L-DOPA by thermostable tyrosine phenol-lyase of a thermophilic Symbiobacterium spp. over expressed in recombinant Escherichia coli. J Microbial Biotechnol 1996;6:98–102. [32] Nagatsu  T, Sawada  M. L-DOPA therapy for Parkinson’s disease: past, present, and future. Parkinsonism Relat Disord 2009;15:S3–8. [33] Ensuncho L, Alvarez-Cuenca M, Legge RL. Removal of aqueous phenol using immobilized enzymes in a bench scale and pilot scale three-phase fluidized bed reactor. Bioprocess Biosyst Eng 2005;27:185–91. [34] Krastanov A. Removal of phenols from mixtures by co-immobilized laccase/tyrosinase and polyclar adsorption. J Ind Microbiol Biotechnol 2000;24:383–8. [35] Martinez  MV, Whitaker  JR. The biochemistry and control of enzymatic browning. Trends Food Sci Technol 1995;6:195–200. [36] Ashida M, Brey P. Role of the integument in insect defense: prophenoloxidase cascade in the cuticular matrix. Proc Natl Acad Sci U S A 1995;92:10698–702. [37] Soderhall K, Cerenius L. Role of the prophenoloxidase activating system in invertebrate immunity. Curr Opin Immunol 1998;10:23–8. [38] Pierpoint WS. o-Quinones formed in plant extracts. J Biochem 1969;112:609–16.

Microbial Tyrosinases  Chapter | 1  17 [39] Dervall BJ. Phenolase and pectic enzyme activity in the chocolate spot disease of beans. Nature 1961;189:311. [40] Walker  JRL, Ferrar  PH. Diphenol oxidases, enzyme-catalysed browning and plant disease resistance. Biotechnol Genet Eng Rev 1998;15:457–97. [41] Sugumaran M. Comparative biochemistry of eumelanogenesis and the protective roles of phenoloxidase and melanin in insects. Pigment Cell Res 2002;15:2–9. [42] Karam J, Nicell JA. Potential applications of enzymes in waste treatment. J Chem Technol Biotechnol 1997;69:141–53. [43] Seo SY, Sharma VK, Sharma N. Mushroom tyrosinase: recent prospects. J Agric Food Chem 2003;51:2837–53. [44] Saini AS, Kumar J, Melo JS. Microplate based optical biosensor for L-DOPA using tyrosinase from Amorphophallus campanulatus. Anal Chim Acta 2014;849:50–6. [45] Nosanchuk  JD, Casadevall  A. The contribution of melanin to microbial pathogensis. Cell Microbiol 2003;5:203–23. [46] Del-Marmol  V, Beermann  F. Tyrosinase and related proteins in mammalian pigmentation. FEBS Lett 1996;381:165–8. [47] Hearing  VJ, Tsukamoto  K. Enzymatic control of pigmentation in mammals. FASEB J 1991;5:2902–9. [48] Surwase SN, Patil SA, Apine OA, Jadhav JP. Efficient microbial conversion of L-tyrosine to L-DOPA by Brevundimonas sp. SGJ. Appl Biochem Biotechnol 2012;167:1015–28. [49] Sveinbjornsdottir  S. The clinical symptoms of Parkinson's disease. J Neurochem 2016;139:318–24. [50] Knowles WS, Sabacky MJ, Vineyard BD. Monsanto Company. Asymmetric catalysis. US Patent 4008281. 1974 Oct 15. [51] Lewitt PA. Levodopa therapeutics for Parkinson’s disease: new developments. Parkinsonism Relat Disord 2009;15:S31–4. [52] Algieri  C, Donato  L, Bonacci  P, Giorno  L. Tyrosinase immobilized on polyamide tubular membrane for the L-DOPA production: total recycle and continuous reactor study. Biochem Eng J 2012;66:14–9. [53] Haneda K, Watanabe S, Takeda I. Synthesis of L-3,4-dihydroxyphenylalanine from L-tyrosine by microorganisms. Appl Microbiol 1971;22:721–2. [54] Raval KM, Vaswani PS, Majumder DR. Biotransformation of a single amino-acid L-tyrosine into a bioactive molecule L-DOPA. Int J Sci Res Publ 2012;2:2250. [55] McMahon AM, Doyle EM, Brooks SJ, O’Connor KE. Biochemical characterisation of the coexisting tyrosinase and laccase in the soil bacterium Pseudomonas putida F6. Enzyme Microb Technol 2007;40:1435–41. [56] Matoba  Y, Kumagai  T, Yamamoto  A, Yoshitsu  H, Sugiyama  M. Crystallographic evidence that the dinuclear copper center of tyrosinase is flexible during catalysis. J Biol Chem 2006;281:8981–90. [57] Nambudiri AMD, Bhat JV. Conversion of p-cumarate into caffeate by Streptomyces nigrifaciens. J Biochem 1972;130:425–33. [58] Lerch K, Ettlinger L. Purification of a tyrosinase from Streptomyces glaucescens. Eur J Biochem 1972;31:427–37. [59] Liu N, Zhang T, Wang YJ, Huang JH, Ou P, Shen A. A heat inducible tyrosinase with distinct properties from Bacillus thuringiensis. Lett Appl Microbiol 2004;3:407–12. [60] Kong  KH, Hong  MP, Choi  SS, Kim  YT, Cho  SH. Purification and characterization of a highly stable tyrosinase from Thermomicrobium roseum. Biotechnol Appl Biochem 2000;31:113–8.

18  SECTION | I  Enzymes in Bioprocessing [61] Lerch K. Neurospora tyrosinase: structural, spectroscopic and catalytic properties. Mol Cell Biochem 1983;52:125–38. [62] Nakamura T, Sho S, Ogura Y. On the purification and properties of mushroom tyrosinase. J Biochem 1966;59:481–6. [63] Robb DA, Gutteridge S. The polypeptide composition of two fungal tyrosinases. Phytochemistry 1981;20:1481–5. [64] Wichers  HJ, Gerritsen  YAM, Chapelon  CGJ. Tyrosinase isoforms from the fruitbodies of Agaricus bisporus. Phytochemistry 1996;43:333–7. [65] Wichers HJ, Recourt K, Hendriks M, Ebbelaar CEM, Biancone G, Hoeberichts FA, Mooibroek H, Soler-Rivas C. Cloning, expression and characterisation of two tyrosinase cDNAs from Agaricus bisporus. Appl Microbiol Biotechnol 2003;61:336–41. [66] Kanda K, Sato T, Ishii S, Enei H, Ejiri S. Purification and properties of tyrosinase isozymes from gill of Lentinus edodes fruiting bodies. Biosci Biotechnol Biochem 1996;60:1273–8. [67] Abdel-Raheem A, Shearer CA. Extracellular enzyme production by freshwater ascomycetes. Fungal Divers 2002;11:1–19. [68] Halaouli  S, Asther  M, Kruus  K, Guo  L, Hamdi  M, Sigoillot  JC, Asther  M, Lomascolo  A. Characterization of a new tyrosinase from Pycnoporus species with high potential for food technological applications. J Appl Microbiol 2005;98:332–43. [69] Selinheimo E, Saloheimo M, Ahola E, Westerholm-Parvinen A, Kalkkinen N, Buchert J, Kruus K. Production and characterization of a secreted, C-terminally processed tyrosinase from the filamentous fungus Trichoderma reesei. J Fed Soc Biochem Mol Biol 2006;273:4322–35. [70] Nakamura  M, Nakajima  T, Ohba  Y, Yamauchi  S, Lee  BR, Ichishima  E. Identification of copper ligands in Aspergillus oryzae tyrosinase by site-directed mutagenesis. J Biochem 2000;360:537–45. [71] Agarwal P, Dubey S, Singh M, Singh RP. Aspergillus niger PA2 Tyrosinase covalently immobilized on a novel eco-friendly bio-composite of chitosan-gelatin and its evaluation for L-DOPA production. Front Microbiol 2016;7:2088. [72] Agarwal P, Pareek N, Dubey S, Singh J, Singh RP. Aspergillus niger PA2: a novel strain for extracellular biotransformation of L-tyrosine into L-DOPA. Amino Acids 2016;48:1253–62. [73] Agarwal P, Singh J, Singh RP. Molecular cloning and characteristic features of a novel extracellular tyrosinase from Aspergillus niger PA2. Appl Biochem Biotechnol 2016;182:1–15. [74] Fujieda N, Yabuta S, Ikeda T, Oyama T, Muraki N, Kurisu G, Itoh S. Crystal structures of copper-depleted and copper-bound fungal pro-tyrosinase: insights into endogenous cysteinedependent copper incorporation. J Biol Chem 2013;288:22128–40. [75] Vallee BL. Zinc and metalloenzymes. Adv Protein Chem 1955;10:317–84. [76] Klabunde T, Eicken C, Sacchettini JC, Krebs B. Crystal structure of a plant catechol oxidase containing a dicopper center. Nat Struct Biol 1998;5:. [77] Jackman MP, Hajnal A, Lerch K. Albino mutants of Streptomyces glaucescens tyrosinase. J Biochem 1991;274:. [78] Jimeinez M, Maloy WL, Hearing VJ. Specific identification of an authentic clone for mammalian tyrosinase. J Biol Chem 1989;264:3397–404. [79] Decker H, Tuczek F. Tyrosinase/catecholoxidase activity of hemocyanins: structural basis and molecular mechanism. Trends Biochem Sci 2000;25:392–7. [80] Olivares C, Garcia-Borron JC, Solano F. Identification of active site residues involved in metal cofactor binding and stereo-specific substrate recognition in mammalian tyrosinase. Implications to the catalytic cycle. J Biochem 2002;41:679–86. [81] Selinheimo E, NiEidhin D, Steffensen C, Nielsen J, Lomascolo A, Halaouli S. Comparison of the characteristics of fungal and plant tyrosinases. J Biotechnol 2007;130:471–80.

Microbial Tyrosinases  Chapter | 1  19 [82] Olivares C, Solano F. New insights into the active site structure and catalytic mechanism of tyrosinase and its related proteins. Pigment Cell Melanoma Res 2009;22:750–60. [83] Garcia-Borron JC, Solano F. Molecular anatomy of tyrosinase and its related proteins: beyond the histidine-bound metal catalytic center. Pigment Cell Res 2002;15:162–73. [84] Inoue T, Shiota Y, Yoshizawa K. Quantum chemical approach to the mechanism for the biological conversion of tyrosine to dopaquinone. J Am Chem Soc 2008;130:16890–7. [85] Jolley RL, Evans LH, Makino N, Mason HS. Oxytyrosinase. J Biol Chem 1974;249:335–45. [86] Li  Y, Wang  Y, Jiang  H, Deng  J. Crystal structure of Manducasexta prophenoloxidase provides insights into the mechanism of type 3 copper enzymes. Proc Nat Acad Sci U S A 2009;106:17002–6. [87] Fujita Y, Uraga Y, Ichisima E. Molecular cloning and nucleotide sequence of the protyrosinase gene, melO, from Aspergillus oryzae and expression of the gene in yeast cells. Biochim Biophys Acta 1995;1261:151–4. [88] Lopez-Serrano  D, Sanchez-Amat  A, Solano  F. Cloning and molecular characterization of a SDS-activated tyrosinase from Marinomonas mediterranea. Pigment Cell Res 2002;15:104–11. [89] Bernan V, Fipula D, Herber W, Katz E. The nucleotide sequence of the tyrosinase gene from Streptomyces antibioticus and characterization of the gene product. Gene 1985;37:101–10. [90] Endo  K, Hayashi  Y, Hibi  T, Hosono  K, Beppu  T, Ueda  K. Enzymological chracterization of EpoA, a laccase like phenol oxidase produced by Streptomyces griseus. J Biochem 2003;133:671–7.

Chapter 2

Production of Bioethanol From Sugarcane Bagasse: Current Approaches and Perspectives Swati Tyagi*, Kui-Jae Lee*, Sikandar I. Mulla*, Neelam Garg†, Jong-Chan Chae* *

Division of Biotechnology, Chonbuk National University, Iksan, Republic of Korea Department of Microbiology, Kurukshetra University, Kurukshetra, India

†

1. INTRODUCTION The whole world is facing two big challenges: (1) to provide enough energy sources, and (2) to protect the environment. The current issues of fuel demands and intense climate change are directing the world’s attention toward finding alternative energy sources that can reduce our dependency on fossil fuels [1,2]. The world economy depends on the usage of fossil fuels because it is the main source of energy (about 80%) [3,4]. Uncontrolled usage of natural fuels has a negative impact on nature because of environmental pollution and the emission of a huge amount of GHG resulting in global warming [5]. However, energy consumption is ever increasing because of the race toward industrial development [6,7]. In addition, another main drawback of fossil fuel usage is that it is a finite resource and will be depleted in the near future. In 2016 a BP Statistical Review of World Energy reported that the majority of our natural fuels would be used up in next 50 years. The emission of GHG not only contributes to global warming, but also has other deleterious impacts on mammalian life. For these reasons over the last decade many efforts have been made around the world to find clean and sustainable energy sources that could substitute the fossil fuels [8–10]. In this respect biofuel is getting more attention because it is produced from lignocellulosic/cellulosic biomass via the alcoholic fermentation of sugar [11,12]. Lignocellulosic biomass (LCM) consisting of cellulose (~50%), hemicelluloses (~30%) and lignin (~20%), and it is a potential renewable energy source in the form of biofuel, especially bioethanol [13]. Researchers have classified LCM into six major groups on the basis of origin and usage: (1) from agriculture waste, (2) from hardwood, (3) from softwood, (4) from cellulose-based Applied Microbiology and Bioengineering. https://doi.org/10.1016/B978-0-12-815407-6.00002-2 © 2019 Elsevier Inc. All rights reserved.

21

22  SECTION | I  Enzymes in Bioprocessing

waste materials, (5) from herbaceous biomass (alfalfa stems and switchgrass), and (6) from municipal solid wastes [14–16]. A comparative analysis of various lignocellulosic materials is given in Table 1. Statistics say that in dry weight almost 52 million tons of hardwood, 145 million tons of processed wood/softwood, 47 million tons of agriculture waste, 64 million tons of herbaceous biomass, and 60 million tons of biomass from the biofuel industry are released annually into the environment [30]. The composition and moisture content of LCM might differ depending on area, agriculture practices, processing and storage time (Table 1). Agriculture-based lignocellulosic wastes like sugarcane bagasse (SCB) are attracting attention worldwide as a potential substitute for fossil fuels. SCB, usually considered as a waste, is one of the major residues derived from the sugar and alcohol industry. Most SCB is used as fuel for the boilers that equip the factories [31]. However, biotechnology has enabled it to be turned into a desirable and inexpensive substrate for the large-scale production of ethanol. In addition, the ready availability of SCB and the low cost has rendered it attractive as a raw material for ethanol production. India is one of the world’s largest producers of sugarcane, harvesting 357 million tons per annum. Nearly 60% of the sugarcane is consumed for the production of sugar. The residual after production is called bagasse containing up to 35% of pith [32]. In order to recycle the generated SCB it needs to be considered as an appropriate substrate for bioconversion to ethanol instead of being utilized as a burning resource to produce energy, since it contains high carbohydrate and low lignin. SCB is a rich source of glucose (hexose) and xylose (pentose), which are six and five‑carbon sugars, respectively. The cellulose and hemicellulose fraction of SCB is hydrolyzed to produce pentose and hexose sugars. Whereas resultant hexose sugars are easily utilized by microorganisms to produce bioethanol, few microorganisms showed the ability to utilize pentose sugars as a growth substrate [33]. For the highest production of bioethanol, microorganisms that can utilize all kind of sugars, especially hexoses and pentoses, are considered as seed organisms. Saccharomyces cerevisiae is able to ferment hexose sugars, while Pachysolen tannophilus is a promising pentose-fermenting organism [34–37]. Therefore a mixed culture of those microorganisms could be considered for the efficient conversion of SCB to bioethanol. SCB may prove a potential alternative substrate for bioethanol production and use of bioethanol as a transportation biofuel could be a sustainable choice for safe energy production thus decreasing GHG emissions [5,36]. Despite these advantages, production of bioethanol on a commercial scale still faces obstacles because of the relatively low yield and high production costs and the fact that a considerable quantity of SCB is required owing to the limited efficiency of the process. In addition, hydrolysis of bagasse requires a huge quantity of cellulase enzymes, which is expensive and not economically feasible [11,38].

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  23

TABLE 1  Comparative Analysis of Various Lignocellulosic Materials for Bioethanol Production Content (Dry wt%)

Lignocellulosic Material

Cellulose

Hemicellulose

Lignin

References

Corn stover

40.00

29.60

23.00

[17]

Corn cobs

45

35

15

[18]

Banana waste

13.2

14.8

14

[19]

Rice husks

36.70

20.05

21.30

[20]

Barley husks

21.40

36.62

19.20

[20]

Sweet sorghum

21

27

45

[21]

Rye straw

41.10

30.20

22.90

[22]

Oat straw

39.40

27.10

17.50

[23]

Rice straw

32.1

24

18

[18]

Wheat straw

32.90

24.00

8.90

[23]

Corn stalks

35.00

16.80

NA

[23]

Sponge gourd fibers

66.59

17.44

15.46

[24]

Cotton stalks

58.50

14.40

21.50

[23]

Soya stalks

34.50

24.80

7.00–19.80

[23]

Sunflower stalks

42.10

29.70

13.40

[23]

Grasses

25–40

25–50

10–30

[25]

Sugarcane bagasse

40.00

27.00

10.00

[23]

Ethiopian mustard

32.70

21.90

18.70

[26]

Flax shives

47.70

17.00

26.60

[26]

Hemp hurds

37.40

27.60

18.00

[26]

Alfalfa stems

27.50

23.00

15.80

[26]

Switch grass

31.98

25.19

18.13

[27]

Newspaper

61.30

9.80

12.00

[28]

Waste papers from chemical

60–70

10–20

5–10

[29]

Primary waste water solids pulps

8–15

NA

24–29

[29]

NA, not available.

24  SECTION | I  Enzymes in Bioprocessing

Therefore cutting-edge technologies are required for the economical production of saccharifying enzymes and improvements in hydrolysis. In this chapter we aimed to review the methods and the processes used for the production of bioethanol from SCB.

2.  SUGARCANE BAGASSE SCB contains a large amount of cellulose, along with hemicelluloses, lignin, and waxes (Table 2). Cellulose is a homopolysaccharide composed of β-d-glucopyranose units linked together by (1→4)-glycosidic bond. The basic unit of the cellulose is cellobiose which consists of long chains of 5000–10,000 glucose units [39,40]. Hemicellulose comprises a group of various complex carbohydrates, such as xylose, arabinose, uronic acid, 4-O-methyl glucuronic acid, and galacturonic acid, whereas lignin is a complex aromatic polymer connected with cellulose fibers. Due to the high yield (~80 tons/ha) and the high carbohydrate and low ash (~2%) content when compared with other agricultural wastes, SCB is considered a rich energy source [41–45]. In addition, SCB (a complex polymer of cellulose, hemicelluloses, and lignin) is abundant, renewable, and an attractive low-cost feedstock, which makes it a versatile starting material for several processes, such as fermentation, biocatalysis, and chemo-catalysis, to generate value-added products including biofuels, biopolymers, and other useful chemicals.

3.  VARIOUS MICROORGANISMS DEGRADING BAGASSE The process of conversion of SCB to bioethanol requires microorganisms that are capable of utilizing various sugars. For decades a number of microorganisms including bacteria, fungi, and yeasts have been cultivated on bagasse. Although, filamentous white-rot fungus belonging to basidiomycete has been mainly used by the bioethanol industries [7], other fungi, such as brown-rot and soft-rot, also have been used to degrade lignocellulosic biomass. Usually white-rot and soft-rot fungi are used to degrade hemicellulose and lignin, whereas brown-rot fungus was mainly used for cellulose degradation [46]. Other fungal species such as Trichoderma sp. [47], Aspergillus terreus [48], TABLE 2  Composition of Sugarcane Bagasse Component

Composition (%)

Cellulose

32–55

Hemicelluloses

26.7–32

Lignin

19–24

Waxes

0.8–6

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  25

Cyathus stercoreus [49], Lentinus squarrosulus [50], Lentinus edodes [51], Pleurotus sp. [52], Penicillium camemberti [53], etc., have been reported to degrade 45%–75% of hemicellulose and 65%–80% of lignin. The only drawback in the application of diverse microorganisms is that they cannot metabolize xylose [54]. One possible way to overcome this problem is the genetic engineering of these microorganisms. By inserting the genes responsible for xylose metabolism into the genomes this would allow them to ferment all the sugars during SCB hydrolysis and bioethanol production [55,56]. The introduction of genes for xylose metabolism into S. cerevisiae has been reported [37,57]. Although the S. cerevisiae (Baker’s yeast) used in the studies did not naturally have the capability to ferment pentose sugars, pentose sugars (especially xylose) were successfully utilized by the engineered microorganisms [37]. However, even after a decade this area remains a great challenge for molecular biologists and engineers. It is important for microorganisms to improve a tolerance to the high concentrations of metabolites formed during the fermentation steps, which might inhibit microbial growth. The various xylose-fermenting microbes that have been reported comprise bacteria, yeasts, and fungi [44]. Researchers are no longer solely concerned with finding novel organisms that have the xylose-fermenting ability, they also want to understand the xylose metabolism. Hence xylose degrading bacteria [58] have also gained attention, because their growth rate is substantially faster than yeast. With the recent advances in microbial biotechnology, it has been found that these bacteria (e.g. Zymobacter palmae) have great potential to produce bioethanol more economically.

4.  PRETREATMENT OF BAGASSE Production of bioethanol from SCB comprises five key stages: (1) pretreatment of the biomass, (2) cellulose hydrolysis, (3) fermentation, (4) separation, and (5) treatment of the effluent (Fig. 1). In order to facilitate cellulose hydrolysis a number of pretreatment techniques have been developed. These techniques include various physical, chemical, physiochemical, and biological pretreatments [59–61]. Physical pretreatment includes comminution and hydrothermolysis whereas chemical pretreatment is performed using acid-, alkali-, ozone-, and solvent-based methods. Physiochemical pretreatment is the combination of physical and chemical methods such as steam explosion and ammonia fiber explosion. Alternatively, biological pretreatment uses microorganisms and/or microbial enzyme systems. The primary goal of these pretreatments is to disrupt the complex structure of SCB and its main components, like celluloses, hemicelluloses, and lignin, into simple sugars, which are useful resources in the bioconversion processes [62]. A number of different pretreatment processes had been traditionally used, but none were efficient enough to give good results. Processes now employ a number of innovative methods that are capable of dispersing the three main constituents of SCB (Table 3).

26  SECTION | I  Enzymes in Bioprocessing

Sugarcane bagasse Crushing Acid/alkali treatment

Ethanol purification

Pretreatment method

Bioethanol

Sugarcane bagasse Distillation

Microorganisms

Yeast

Enzyme production unit

Enzyme production method

Fermentation unit

Fermentation method

Waste product

FIG. 1  Schematic representation of bioethanol production from sugarcane bagasse.

TABLE 3  Methods and Results of Delignification Technologies Method

Pretreatment Conditions

Results

References

Steam explosion

1.3 MPa, 190°C for 15 min

7.9 ± 9.1% lignin solubilization

[63]

1% (w/v) H2SO4 at 180°C

74% glucose

[62]

Solid liquid ratio 1:10 (w/v), 100°C for 1 h

92.7 ± 3.9% lignin solubilization

[63]

0.75% (w/v) NaOH at 121°C for 15 min

90.43% glucan, 65.11% xylan

[64]

Diluted ammonia (15%, w/v) at 170°C for 60 min

82.7% glucose, 47.3% xylose

[65]

2% H2O2 with 1.5% NaOH, 121°C, 15 min

Increased cellulose level 1.2 times and decreased hemicelluloses content 8.5 times in sugarcane bagasse

[66]

Alkaline pretreatment

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  27

TABLE 3  Methods and Results of Delignification Technologies—cont’d Method Acid pretreatment

Ammonia fiber expansion

Organosolv

Liquid hot water

Wet oxidation alkaline

Pretreatment Conditions

Results

References

1% H2SO4 and enzymatic hydrolysis after microwave alkali treatment (1% NaOH)

0.83 g reducing sugars/g dry sugarcane bagasse

[67]

1.5% (v/v) H2SO4 at 161°C for 5 min

85.5% glucose

[68]

1% (w/w) H2SO4 treatment at 180°C for 10 min

63.17% glucan, 75.12% xylan

[69]

2 kg ammonia + 1.5 kg water/kg dry bagasse, 140°C, 30 min

85% glucan conversion by cellulases and 95%– 98% xylan conversion by hemicellulases in bagasse and cane leaf residue

[70]

AFEX pretreatment at 135°C for 45 min

90% total reducing sugar

[71]

30% ethanol with NaOH (v/v), 60 min, 195°C, preceded by dilute-acid pretreatment

Residual solid material from sugarcane bagasse containing 67.3% glucose (w/w)

[72]

Using ethanol as organic solvent

89.7% sugar recovery

[73]

190–230°C, rapid immersed percolation (45 s to 4 min)

50% solubilization of sugarcane bagasse and leaves (all of the hemicellulose, 60% of the acid insoluble lignin and less than 10% of the cellulose)

[74]

Pretreated at 210°C for 10 min

64.55% glucose

[75]

195°C, 15 min

Solid material with 70% cellulose with solubilization of 93% hemicelluloses and 50% lignin

[76]

Pretreatment at 185°C with oxygen (0.6 MPa)

87.4% glucose

[77]

28  SECTION | I  Enzymes in Bioprocessing

4.1  Mechanical Comminution The comminution of SCB is a method used to reduce the crystallinity of cellulose. Mechanical comminution of SCB is performed by finely grinding or fragmenting it into small pieces (15–30 mm) and then milling these pieces still further to reduce the size to 0.2–2.0 mm [59]. Although ordinary ball milling can be used vibratory ball mills are known to be more effective. Improved digestibility of the biomass and reduced cellulose crystallinity of spruce and aspen chips were obtained when vibratory ball milling was carried out [59,78]. The effect of mechanical comminution on various agricultural-based materials depends on the final particle size and the features [79].

4.2 Pyrolysis Pyrolysis (also known as thermal degradation) is a method in which SCB biomass is exposed to a high temperature (between 500°C and 800°C) without an oxidizing agent. At high temperatures carbohydrates quickly generate a number of gaseous by-products along with pyrolysis oil and leftover char [59]. At lower temperatures decomposition of SCB by pyrolysis is slowed and fewer volatile products are formed. However, additional treatment with a mild acid such as 1.0 N sulfuric acid (H2SO4) at 97°C for 2.5 h can improve the degradation rate up to 80%–85% [80]. A good oxygen supply can also improve the efficiency of the process [59,81].

4.3  Steam Pretreatment or Steam Explosion The steam explosion method is an efficient process and widely used for the pretreatment of the plant cell wall [82]. In this method fragmented biomass is treated with highly saturated steam under pressure (at 160–260°C with a pressure of 0.69–4.83 MPa) and then the pressure is slowly reduced to an atmospheric level. Due to the high temperature and pressure, lignin and hemicelluloses are decomposed into simpler molecules which in turn can be utilized via cellulose hydrolysis [56,83]. Exposure time, temperature, pressure, size of SCB materials, and moisture can all affect the efficiency of steam explosion. Exposure time and temperature are almost inversely proportional to each other. For example, a high temperature and a short exposure time (270°C, 1 min), or a low temperature and a longer exposure time (190°C, 10 min) result in similar solubilization rates. Some researchers support the use of a lower temperature and a longer exposure time because fewer toxic byproducts are produced under these conditions [84,85]. Additionally, improved enzymatic hydrolysis efficiency (~90%) has been reported when SCB is treated by steam explosion [31,83–85]. Steam explosion is reported to be more effective for hardwoods and agricultural residues than for softwoods. A major limitation of this method is the incomplete destruction of xylan and lignin thus generating toxic byproducts,

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  29

which can inhibit the growth of fermenting microorganisms. Because of the formation of inhibitory compounds, a washing step is needed to remove them. Thus additional cost and effort are required [86,87].

4.4  Acid Pretreatment Various types of acid have been used for the pretreatment of different biomasses including SCB [59] and a few of the acids that are commonly used are discussed here. Hydrolysis using H2SO4, hydrochloric acid (HCl) or acetic acid (CH3COOH) is usually called acid hydrolysis or prehydrolysis. Hemicellulose is the substrate that is sought from acid treatment. As it is more easily degraded than cellulose and lignin, two phases are obtained at the end of the experiment: a liquid phase containing simple sugars and a solid phase containing an unaltered portion of lignin and cellulose, which is processed further [88]. Depending on the kind of acid and the experimental conditions, in addition to the reducing sugars, byproducts such as furfural and 5-hydroxymethylfurfural (HMF) can be formed [88]. In the fermentation process these byproducts are toxic to the fermenting microorganisms and can arrest the process if present in high concentrations. However, if the concentrations of the toxic compounds are low, hydrolysate can serve as an enriched medium for fermentation [88]. SCB hydrolysis with H2SO4 yielded 24.5 g/L of total sugar containing 11 g/L glucose, 2.22 g/L arabinose, 2.48 g/L acetic acid, and 0.12 g/L furfural [86]. Some studies showed that the use of HCl for pretreatment of SCB yielded higher levels of sugars compared with other acids. However, its deleterious effects on the environment and corrosive properties have limited its application [89]. Phosphoric acid (H3PO4) also can be used for the pretreatment of SCB. After SCB is treated with phosphoric acid and sodium hydroxide for neutralization, salts of sodium and phosphate are formed in hydrolysate as byproducts, which could be utilized by the SCB fermenting microorganisms [90]. Therefore filtration to remove the phosphate-sulfate salt is not needed thus highlighting the cost-effectiveness of the process. Recently, a combination of acetic acid (CH3COOH) and hydrogen peroxide (H2O2) was reported as an effective pretreatment method to remove lignin from SCB [91].

4.5  Alkaline Pretreatment While acid treatment targets the hemicellulose fraction of SCB, alkaline pretreatment digests the lignin fraction of SCB [5]. Alkali pretreatment hydrolyzes the cell-wall constituents and breaks the crystalline structure of cellulose [91]. The treatment of SCB generates alkali-soluble residue (mainly cellulose), which can be used for producing value-added products such as bioethanol by fermentation [40]. Treatment with 1%–3% sodium hydroxide (NaOH) aqueous solutions stimulated a significant dissolution of the hemicellulose and the lignin in SCB into simple sugars [40]. The greatest degradation of hemicellulose

30  SECTION | I  Enzymes in Bioprocessing

and lignin occurred when NaOH was used in comparison with other alkaline agents such as sodium carbonate (Na2CO3), ammonium hydroxide (NH4OH), calcium hydroxide [Ca(OH)2], and H2O2. Fermentation yield was also higher in NaOH treatment [92,93]. It was also reported that pretreatment with 2.5% NaOH resulted in the maximum production of cellulose (81%) and delignification (68.5%) of sugarcane bagasse [94].

4.6  Thermal Pretreatment Thermal pretreatment demonstrates tremendous benefits, such as low energy requirements and the production of fewer toxic compounds and growth inhibitors when compared with the mechanical and chemical processes [95]. During thermal pretreatment SCB is heated to very high temperatures (in excess of 150–180°C) and high pressures (5 MPa). At this high temperature hemicellulose and lignin are sequentially solubilized. The combination of thermal and chemical treatments of SCB improves the solubility of hemicellulose [96]. Boussarsar et  al. [92] reported that higher hemicellulose solubilization occurred when SCB was treated using the hydrothermal process. Further transformation of sugars (glucan, xylan oligomers, and polymers with large chains) into toxic byproducts was also minimal in this process [87]. Rocha et al. [97] reported that steam pretreatment of SCB at 1.3 MPa, 190°C, and 50% moisture hydrolyzed hemicelluloses, cellulose, and lignin with an average of ~85%, ~18%, and ~10%, respectively. However, a few of the toxic inhibitory compounds, such as furfural and carboxylic acid, were also found. Ribeiro et al. [95] examined the anaerobic digestion of the hemicellulose hydrolysate formed after thermal pretreatment of SCB and showed that 20.5% of HMF and 11.5% of fufural were removed.

4.7  Biological Pretreatment Basidiomycetes capable of degrading lignin were found to be the most effective for biological pretreatment [98]. In addition, the fungus Phlebia sp. was also used as a biological agent for the pretreatment of SCB and was reported to degrade about 60% of the lignin and 10% of the hemicellulose [45]. Other microorganisms like Phanerochaete chrysosporium, Phanerochaete sordida, Ceriporia sp., Aspergillus terreus, Trichoderma sp., Cyathus stercoreus, Lentinus squarrosulus, Lentinus edodes, Trametes pubescens, Penicillium camemberti, Phanerochaete chrysosporium, Ceriporiopsis subvermispora, Phellimus pini, and Pleurotus sp. were also reported as being used for the degradation of bagasse [98,99]. White-rot fungi are known to degrade lignin while brown rots mainly attack cellulose. However, soft-rot fungi can metabolize both cellulose and lignin [7,48]. Edible mushrooms Agrocybe aegerita, Volvariella volvacea and Pleurotus sp. also have been reported to hydrolyze agricultural waste with an efficiency of 40%–90% [52,100].

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  31

In biological pretreatment, microorganisms and their enzymes degrade SCB components including lignin, cellulose, and hemicellulose [101,102]. Biological pretreatments have several advantages over chemical and physical methods because of the low energy requirement, negligible waste production, and the lack of a negative impact on the environment. Nowadays, commercially available microbial enzymes are also employed for hydrolysis of SCB. The white-rot fungi produce the extracellular enzymes responsible for lignin degradation, such as lignin peroxidases, polyphenol oxidases, laccases, and manganese-dependent peroxidases. In addition, other proteins including H2O2-producing enzymes and quinine-reducing enzymes were reported to be involved in lignin degradation [60,83]. These enzymes catalyze oxidation of lignin units generating aromatic radicals. Generally, lignin is not utilized as a sole carbon source but through a secondary metabolic process when the availability of nitrogen, carbon, or sulfur is limited. Hence, lignin-degradation processes require additional cosubstrates, such as cellulose, hemicellulose or glucose [103]. There are, however, some major drawbacks, such as the time needed for fermentation, the failure to ferment pentose sugars, and the toxic effect of the growth inhibitors on the fermenting organisms [103]. To overcome these limitations, newly developed bioengineering techniques, such as zinc finger nucleases (ZFN), transcription activation like effector nucleases (TALEN), and clustered regularly interspaced short palindromic repeats (CRISPR), have been used to edit the genome of target organism [104,105]. Exploiting these techniques, genetically modified organisms can utilize sugars (pentoses and hexoses) and have an improved capability to digest cellulose, hemicellulose, and lignin [105].

4.8  Ammonia Fiber Explosion (AFEX) and Carbon Dioxide (CO2) Explosion Pretreatment Ammonia fiber explosion (AFEX) method is analogous to steam pretreatment. In this method, ammonia is passed over SCB at a high temperature and pressure, and the pressure is then gradually dropped. During the process ammonia reacts with lignin, depolymerizes it and breaks lignin-cellulose linkages. In enzymatic hydrolysis, lignin concentration affects the process. However, removal of lignin with AFEX can drop the enzyme requirement and thus make it economical [106]. A number of LCMs have been treated using this method [29,107–110]. Similarly, carbon dioxide can be used for the pretreatment of SCB. During the reaction CO2 is transformed into carbonic acid (a weak acid), which improves the hydrolysis process similarly to acid treatment [91].

4.9  Wet Oxidation Pretreatment During wet oxidation pretreatment SCB is treated with water at high temperatures in the presence of oxygen. In this process two reactions take place: a lowtemperature hydrolytic reaction and a high-temperature oxidation reaction [77].

32  SECTION | I  Enzymes in Bioprocessing

The major advantage of this method is that no byproducts, such as furfural and HMF, are generated [111,112]. However, the use of oxygen makes this method more expensive. Ramos [111] reported that the treatment with hot water (at 200–230°C for 15 min) hydrolyzed almost 60% of SCB. Further acid treatment resulted in degradation of hemicellulose (~90%) into monomeric sugars [113]. For wet oxidation three different types of hot-water reactors are available: (1) flow type, in which hot water is passed over a solid biomass with a longer residence time dissolving hemicellulose and lignin; (2) the cocurrent reactor, in which biomass is exposed to a high temperature followed by other pretreatments and then cooling; and (3) countercurrent type, in which biomass and water run in opposite directions and dissolve the SCB components [113,114]. This method can reduce the removal and conditioning cost [113,114].

4.10  Ozonolysis Pretreatment Pretreatment of SCB also can be done by ozone. Ozonolysis can degrade the lignin content of the whole biomass but does not have any effect on cellulose and hemicellulose [115]. This method can be advantageous if lignin removal is the primary goal. In addition, no toxic byproduct is formed during the process of delignification [115,116].

4.11  Organosolvent Pretreatment This method uses organic solvents, methanol, ethanol, acetone, and ethyleneglycol with the addition of catalytic agents to disrupt lignin and hemicellulose linkages [117]. Degradation of LCM is arrested by the high concentration of lignin and the crystallinity of cellulose [118]. The use of organic solvents overcomes these issues through delignification and reducing the crystallinity of cellulose, which makes further cellulose hydrolysis easier. Consequently, it lowers the overall cost of the process [81,119–121]. Glycerol-acid pretreatment of SCB at 80–120°C for 60 min showed 68.5% digestibility of SCB and 0.57 g/L ethanol production in a final step [122]. Methods could be applied depending on the type of LCM and experimental conditions [123,124]. Table 3 shows the effects of the pretreatment processes that have great potential for ethanol production.

5.  CELLULOSE HYDROLYSIS Cellulose obtained from sachharification is further hydrolyzed by either microbial degradation or by cellulolytic enzymes. Enzymatic hydrolysis decomposes the cellulosic biomass of hydrolysate by cellulase activity and improves the resultant sugar yield [100,125]. Although enzymatic hydrolysis is a significantly slower process, it has the advantage of no byproduct being formed during the process. Krishnan et al. [71] used cellulase and cellobiase of Trichoderma reesei to hydrolyze sugarcane after alkaline delignification [71,126]. Before

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  33

e­ nzymatic hydrolysis, pretreatment is necessary to expose the fibers for enzyme accessibility. Martín et al. [77] reported that steam pretreated SCB after hydrolysis with a combination of various enzymes gave similar results to chemically treated bagasse. Hernández-Salas et al. [88] also used a combination of different enzymes (Celluclast, Novozymes, and Viscozyme L. in alkaline enzymatic treatment) to process SCB and agave. Ammonia pretreated bagasse with citrate buffer and sodium azide gave the maximum sugar yield when supplemented with cellulase [127].

6. DETOXIFICATION During the pretreatment of SCB a number of toxic compounds such as acids, furfural, HMF, and many phenolic compounds are produced along with sugars [127]. Because these compounds subsequently inhibit the fermentation, detoxification of hydrolysates is important to improve the fermentation process. A number of detoxification methods are employed to remove the toxic compounds from the hydrolysate. Enzyme-mediated detoxification is one of the novel approaches in which enzymes such as phenol oxidase, laccase, etc., are used. Calcium hydroxide overliming, pH modifications, ion resin exchange, and active charcoal are other methods that are applied in detoxifying the SCB hydrolysate [127,128]. In order to detoxify the hydrolysate using a neutralization method, chemicals that can neutralize the acidic content are added to the hydrolysate thus allowing salt formation. The generated salt is then removed by filtration. Other toxic compounds are also removed during this process if they are present in low concentrations. The addition of activated charcoal can detoxify the hydrolysate by decreasing acetic content. The ion exchange method is also efficient at removing toxic compounds by up to 80%. An excess amount of lime is used to remove the toxic byproducts and acid content, such as H2SO4, from the lysate. A limitation of the overliming method is the massive sugar loss, which is due to the formation of nonfermentable products [129]. Electrodialysis is another important method of detoxification in which ionic species are separated from aqueous and uncharged components by applying electrically charged membranes [83,130]. SCB detoxification by boiling and electrodialysis resulted in an improved fermentation process [130]. Electrodialysis accomplished almost 90% removal of acetic acid, but sugar loss was less than 5% [127].

7.  ETHANOL PRODUCTION BY FERMENTATION Fermentation of hydrolysate from lignocellulosic residues is necessary to produce alcohol. For this purpose both pentose-utilizing yeast strains [131–133] and pentose-utilizing negative yeast strains (S. cerevisiae) have been used [37,126]. Microbial strains like Escherichia coli, Zymomonas mobilis, and S. cerevisiae, which are capable of utilizing pentose and hexose sugars, have

34  SECTION | I  Enzymes in Bioprocessing

been constructed using recombinant DNA technology, which has made the process efficient [89,134,135]. SCB fermentation can be classified into two groups: solid-state fermentation and liquid fermentation. Furthermore, liquid fermentation can be carried out using two methods: by using whole bagasse as a substrate or by using SCB hydrolysate as substrate. Hernández-Salas et  al. [88] fermented the cellulosic hydrolysates by liquid fermentation with S. cerevisiae strain and reported a total yield of 32% alcohol. Velmurugan and Muthukumar [128] also fermented SCB hydrolysate with S. cerevisiae and produced 8.11 g/L of ethanol. On the other hand, Cheng et al. [130] fermented SCB hydrolysate and produced 0.36 g/g of sugars. Additionally, when S. cerevisiae MTCC 174 was used to ferment SCB, it yielded 15.4 g/L of ethanol [83,136].

8.  APPLICATION OF BAGASSE IN THE PRODUCTION OF OTHER VALUE-ADDED PRODUCTS Several value-added products, e.g., bioethanol, acids, single-cell protein (SCP), etc., can be produced by using SCB in the submerged fermentation system [137–139]. In addition, SCB hydrolysate can be used to produce cellulolytic enzymes [5]. Several white-rot fungi were used successfully to degrade SCB for enzyme production [140]. Another important application of SCB is the production of SCP as a cattle feed. Researchers tried to use one or more organisms together for synchronized sachharification and fermentation processes, which could minimize the cost of the process and improve the yield of the final product [5]. A mixed microbial culture was used for saccharifying SCB that yielded SCP with 35.5% crude protein and showed 69.8% digestibility in rumens [5]. A number of patents have been filed on the production of bioethanol from LCMs like SCB [132,140–144]. When reviewing resource recycling, the indications are that bioethanol production from wastes has great potential in the industry.

9. CONCLUSIONS Tropical climate conditions in many countries allow the cultivation of sugarcane crops, which are a great resource for bioethanol production due to the high cellulose content. A huge amount of sugarcane is produced worldwide and it is being used for sugar extraction as well as bioenergy production. Bioethanol is economically viable if the production yield is high from low-quantity SCB. Currently, the basic challenge for scientists is to achieve a high yield of ethanol via economically feasible pretreatment/hydrolysis processes. Pretreatment or hydrolysis of SCB is a critical step in bioethanol production. However, to date, a standard pretreatment process is not available, although a combination of methods, such as physical or chemical treatment followed by enzymatic hydrolysis, are applied to make SCB more useful. The processes using these methods are still expensive so that the practical applicability is limited. Therefore i­ nnovative

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  35

bioconversion technologies are required to improve the yield of bioethanol production so that usage of bioethanol as an alternative and sustainable fuel can reduce dependence on natural and nonrenewable fossil fuels thus mitigating GHG emissions.

ACKNOWLEDGMENTS This work, doctoral fellowship to the first author Swati Tyagi is supported by BK21 plus programme and National Research Foundation of Korea (NRF) Grant funded by the Korea government (MOE) (No. 2016R1D1A1B03935614).

CONFLICT OF INTEREST The author declares that there is no conflict of interest.

REFERENCES [1] Bagewadi ZK, Mulla SI, Ninnekar HZ. Purification and characterization of endo beta-1,4-dglucanase from Trichoderma harzianum strain HZN11 and its application in production of bioethanol from sweet sorghum bagasse. 3 Biotech 2016;6:101. [2] Bagewadi ZK, Mulla SI, Ninnekar HZ. Optimization of laccase production and its application in delignification of biomass. Int J Recycl Org Waste Agric 2017;6:351–65. [3] Goldemberg J. Ethanol for a sustainable energy future. Science 2007;315:808–10. [4] Saratale GD, Saratale RG, Lo YC, Chang JS. Multicomponent cellulase production by Cellulomonas biazotea NCIM-2550 and its applications for cellulosic biohydrogen production. Biotechnol Prog 2010;26:406–16. [5] Sanchez OJ, Cardona CA. Trends in biotechnological production of fuel ethanol from different feedstocks. Bioresour Technol 2008;99:5270–95. [6] Farzad  S, Mandegari  MA, Guo  M, Haigh  KF, Shah  N, Gorgens  JF. Multi-product biorefineries from lignocelluloses: a pathway to revitalisation of the sugar industry? Biotechnol Biofuels 2017;10:87. [7] Pandey  A, Soccol  CR, Nigam  P, Soccol  VT. Biotechnological potential of agro-industrial residues. I: sugarcane bagasse. Bioresour Technol 2000;74:69–80. [8] International renewable energy agency (IRNA). Renewable energy statistics. Abu Dhabi, UAE: International Renewable Energy Agency; 2017ISBN: 978-92-9260-033-4. [9] Demirbas A. Importance of biodiesel as transportation fuel. Energy Policy 2007;35:4661–70. [10] Ragauskas AJ, Nagy M, Kim DH, Eckert CA, Hallett JP, Liotta CL. From wood to fuels: Integrating biofuels and pulp production. Ind Biotechnol 2006;2:55–65. [11] Tiwari R, Nain L, Labrou NE, Shukla P. Bioprospecting of functional cellulases from metagenome for second generation biofuel production: a review. Crit Rev Microbiol 2018;44: 244–57. [12] Zhang  M, Shukla  P, Ayyachamy  M, Permaul  K, Singh  S. Improved bioethanol production through simultaneous saccharification and fermentation of lignocellulosic agricultural wastes by Kluyveromyces marxianus 6556. World J Microb Biot 2010;26:1041–6. [13] Bagewadi  ZK, Mulla  SI, Ninnekar  HZ. Optimization of endoglucanase production from Trichoderma harzianum strain HZN11 by central composite design under response surface methodology. Biomass Conv Bioref 2017. https://doi.org/10.1007/s13399-017-0285-3.

36  SECTION | I  Enzymes in Bioprocessing [14] Karp SG, Faraco V, Amore A, Letti LA, Soccol VT, Soccol CR. Statistical optimization of laccase production and delignification of sugarcane bagasse by Pleurotus ostreatus in solidstate fermentation. Biomed Res Int 2015;2015:181204. [15] Levin DB, Islam R, Cicek N, Sparling R. Hydrogen production by Clostridium thermocellum 27405 from cellulosic biomass substrates. Int J Hydrogen Energy 2006;31:1496–503. [16] Patel H, Gupte A. Optimization of different culture conditions for enhanced laccase production and its purification from Tricholoma giganteum AGHP. Bioresour Bioprocess 2016;3:11. [17] Sassner P, Galbe M, Zacchi G. Techno-economic evaluation of bioethanol production from three different lignocellulosic materials. Biomass Bioenergy 2008;32:422–30. [18] Prasad S, Singh A, Joshi HC. Ethanol as an alternative fuel from agricultural, industrial and urban residues. Resour Conserv Recyl 2007;50:1–39. [19] John F, Monsalve G, Medina PIV, Ruiz CAA. Ethanol production of banan shell and cassava starch. Dynamis 2006;73:21–7. [20] Parajo JC, Garrote G, Cruz JM, Dominguez H. Production of xylooligosaccharides by autohydrolysis of lignocellulosic materials. Trends Food Sci Technol 2004;15:115–20. [21] Kim M, Day DF. Composition of sugar cane, energy cane, and sweet sorghum suitable for ethanol production at Louisiana sugar mills. J Ind Microbiol Biotechnol 2011;38:803–7. [22] Gullon B, Yanez R, Alonso JL, Parajo JC. Production of oligosaccharides and sugars from rye straw: a kinetic approach. Bioresour Technol 2010;101:6676–84. [23] Nigam PS, Gupta N, Anthwal A. Pre-treatment of agro-industrial residues. In: Nigam PS, Pandey A, editors. Biotechnology for agro-industrial residues utilization. the Netherlands: Springer; 2009. p. 13–33. [24] Guimaraes  JL, Frollini  E, da Silva  CG, Wypych  F, Satyanarayana  KG. Characterization of banana, sugarcane bagasse and sponge gourd fibers of Brazil. Ind Crop Prod 2009;30: 407–15. [25] Malherbe S, Cloete T. Lignocellulose biodegradation: fundamentals and applications. Rev Environ Sci Biotechnol 2002;1:105–14. [26] Gonzalez-Garcia  S, Moreira  MT, Feijoo  G. Comparative environmental performance of lignocellulosic ethanol from different feedstocks. Renew Sustain Energy Rev 2010;14: 2077–85. [27] Hamelinck  CN, van Hooijdonk  G, Faaij  APC. Ethanol from lignocellulosic biomass: techno-economic performance in short-, middle- and long-term. Biomass Bioenergy 2005;28: 384–410. [28] Kim  SB, Moon  NK. Enzymatic digestibility of used newspaper treated with aqueous ­ammonia-hydrogen peroxide solution. Appl Biochem Biotechnol 2003;105:365–73. [29] Sun Y, Cheng JY. Hydrolysis of lignocellulosic materials for ethanol production: a review. Bioresour Technol 2002;83:1–11. [30] Duff SJB, Murray WD. Bioconversion of forest products industry waste cellulosics to fuel ethanol: a review. Bioresour Technol 1996;55:1–33. [31] Wilkins MR, Widmer WW, Grohmann K, Cameron RG. Hydrolysis of grapefruit peel waste with cellulase and pectinase enzymes. Bioresour Technol 2007;98:1596–601. [32] Jain  RK, Thakur  VV, Pandey  D, Adhikari  DK, Dixit  AK, Mathur  RM. Bioethanol from bagasse pith a lignocellulosic waste biomass from paper/sugar industry. IPPTA J 2011;23: 169–73. [33] Zhang Z, Schwartz S, Wagner L, Miller W. A greedy algorithm for aligning DNA sequences. J Comput Biol 2000;7:203–14. [34] Hahn-Hägerdal B, Karhumaa KFC, Spencer-Martins I, Gorwa-Grauslund MF. Towards industrial pentose-fermenting yeast strains. Appl Microbiol Biotechnol 2007;74:937–53.

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  37 [35] Peng F, Ren JL, Xu F, Bian J, Peng P, Sun RC. Comparative study of hemicelluloses obtained by graded ethanol precipitation from sugarcane bagasse. J Agric Food Chem 2009;57: 6305–17. [36] You YZ, Li PF, Lei FH, Xing Y, Jiang JX. Enhancement of ethanol production from green liquor-ethanol-pretreated sugarcane bagasse by glucose-xylose cofermentation at high solid loadings with mixed Saccharomyces cerevisiae strains. Biotechnol Biofuels 2017;10:92. [37] Dangi AK, Dubey KK, Shukla P. Strategies to improve Saccharomyces cerevisiae: technological advancements and evolutionary engineering. Indian J Microbiol 2017;57:378–86. [38] Saha BC. Hemicellulose bioconversion. J Ind Microbiol Biotechnol 2003;30:279–91. [39] Buruiana CT, Garrote G, Vizireanu C. Bioethanol production from residual lignocellulosic materials: a review—part  1. Ann Univ Dunarea de Jos Galati Fascicle VI: Food Technol 2013;37:9–24. [40] Mohan D, Pittman CU, Steele PH. Pyrolysis of wood/biomass for bio-oil: a critical review. Energy Fuel 2006;20:848–89. [41] Cardona CA, Quintero JA, Paz IC. Production of bioethanol from sugarcane bagasse: status and perspectives. Bioresour Technol 2010;101:4754–66. [42] Gao MA, Xu F, Li SR, Ji XC, Chen SF, Zhang DQ. Effect of SC-CO2 pretreatment in increasing rice straw biomass conversion. Biosyst Eng 2010;106:470–5. [43] Ingram LO, Aldrich HC, Borges ACC, Causey TB, Martinez A, Morales F, Zhou SD. Enteric bacterial catalysts for fuel ethanol production. Biotechnol Prog 1999;15:855–66. [44] Olsson L, HahnHagerdal B. Fermentation of lignocellulosic hydrolysates for ethanol production. Enzyme Microb Technol 1996;18:312–31. [45] Zhao L, Riensche E, Menzer R, Blum L, Stolten D. A parametric study of CO2/N2 gas separation membrane processes for post-combustion capture. J Membr Sci 2008;325:284–94. [46] Goodell B, Qian Y, Jellison J. Fungal decay of wood: Soft rot-brown rot-white-rot. In: Schultz T, Nicholas D, Militz H, Freeman MH, Goodell B, editors. Development of commercial wood preservatives: efficacy, environmental, and health issues. Washington: ACS Symposium Series; 2008. p. 9–31. [47] Perez M, Torrades F, Garcia-Hortal JA, Domenech X, Peral J. Removal of organic contaminants in paper pulp treatment effluents under Fenton and photo-Fenton conditions. Appl Catal B: Environ 2002;36:63–74. [48] Emtiazi G, Satarii M, Mazaherion F. The utilization of aniline, chlorinated aniline, and aniline blue as the only source of nitrogen by fungi in water. Water Res 2001;35:1219–24. [49] Keller  FA, Hamilton  JE, Nguyen  QA. Microbial pretreatment of biomass—potential for reducing severity of thermochemical biomass pretreatment. Appl Biochem Biotech 2003;105:27–41. [50] Shide  EG, Wuyep  PA, Nok  AJ. Studies on the degradation of wood sawdust by Lentinus squarrosulus (Mont.) Singer. Afr J Biotechnol 2004;3:395–8. [51] Songulashvili  G, Elisashvili  V, Penninckx  M, Metreveli  E, Hadar  Y, Aladashvili  N, Asatiani M. Bioconversion of plant raw materials in value-added products by Lentinus edodes (Berk.) Singer and Pleurotus spp. Int J Med Mushrooms 2005;7:467–8. [52] Ragunathan  R, Swaminathan  K. Bioconversion of lignocellulosic agro-wastes by fungus, Pleurotus spp. Biol Membr 2004;30:1–6. [53] Taseli BK. Fungal treatment of hemp-based pulp and paper mill wastes. Afr J Biotechnol 2008;7:286–9. [54] Eliasson A, Christensson C, Wahlbom CF, Hahn-Hagerdal B. Anaerobic xylose fermentation by recombinant Saccharomyces cerevisiae carrying XYL1, XYL2, and XKS1 in mineral medium chemostat cultures. Appl Environ Microbiol 2000;66:3381–6.

38  SECTION | I  Enzymes in Bioprocessing [55] Fernandes S, Murray P. Metabolic engineering for improved microbial pentose fermentation. Bioeng Bugs 2010;1:424–8. [56] Cheng  J, Zhu  M. A novel anaerobic co-culture system for bio-hydrogen production from sugarcane bagasse. Bioresour Technol 2013;144:623–31. [57] Laser M, Schulman D, Allen SG, Lichwa J, Antal MJ, Lynd LR. A comparison of liquid hot water and steam pretreatments of sugar cane bagasse for bioconversion to ethanol. Bioresour Technol 2002;81:33–44. [58] Yanase H, Sato D, Yamamoto K, Matsuda S, Yamamoto S, Okamoto K. Genetic engineering of Zymobacter palmae for production of ethanol from xylose. Appl Environ Microbiol 2007;73:2592–9. [59] Kumar AK, Sharma S. Recent updates on different methods of pretreatment of lignocellulosic feedstocks: a review. Bioresour Bioprocess 2017;4:7. [60] Miller GL. Use of Di-nitrosalicylic acid reagent for determination of reducing sugar. Anal Chem 1959;31:426–8. [61] Wingren  A, Galbe  M, Zacchi  G. Techno-economic evaluation of producing ethanol from softwood: comparison of SSF and SHF and identification of bottlenecks. Biotechnol Prog 2003;19:1109–17. [62] Cadoche L, Lopez GD. Assessment of size-reduction as a preliminary step in the production of ethanol from lignocellulosic wastes. Biol Waste 1989;30:153–7. [63] Bertoti AR, Luporini S, Esperidiao MCA. Effects of acetylation in vapor phase and mercerization on the properties of sugarcane fibers. Carbohydr Polym 2009;77:20–4. [64] Kuglarz M, Gunnarsson IB, Svensson SE, Prade T, Johansson E, Angelidaki I. Ethanol production from industrial hemp: effect of combined dilute acid/steam pretreatment and economic aspects. Bioresour Technol 2014;163:236–43. [65] Wang ZY, Keshwani DR, Redding AP, Cheng JJ. Sodium hydroxide pretreatment and enzymatic hydrolysis of coastal Bermuda grass. Bioresour Technol 2010;101:3583–5. [66] Zhang HD, Wu SB. Dilute ammonia pretreatment of sugarcane bagasse followed by enzymatic hydrolysis to sugars. Cellulose 2014;21:1341–9. [67] Aguiar  MM, Ferreira  LFR, Monteiro  RTR. Use of vinasse and sugarcane bagasse for the production of enzymes by lignocellulolytic fungi. Braz Arch Biol Technol 2010;53:1245–54. [68] Binod P, Satyanagalakshmi K, Sindhu R, Janu KU, Sukumaran RK, Pandey A. Short duration microwave assisted pretreatment enhances the enzymatic saccharification and fermentable sugar yield from sugarcane bagasse. Renew Energy 2012;37:109–16. [69] Chiesa S, Gnansounou E. Use of empty fruit bunches from the oil palm for bioethanol production: a thorough comparison between dilute acid and dilute alkali pretreatment. Bioresour Technol 2014;159:355–64. [70] Lu XB, Zhang YM, Angelidaki I. Optimization of H2SO4-catalyzed hydrothermal pretreatment of rapeseed straw for bioconversion to ethanol: focusing on pretreatment at high solids content. Bioresour Technol 2009;100:3048–53. [71] Krishnan C, Sousa LD, Jin MJ, Chang LP, Dale BE, Balan V. Alkali-based AFEX pretreatment for the conversion of sugarcane bagasse and cane leaf residues to ethanol. Biotechnol Bioeng 2010;107:441–50. [72] Petri R, Schmidt-Dannert C. Dealing with complexity: evolutionary engineering and genome shuffling. Curr Opin Biotechol 2004;15:298–304. [73] Mesa  L, Gonzalez  E, Cara  C, Gonzalez  M, Castro  E, Mussatto  SI. The effect of organosolv pretreatment variables on enzymatic hydrolysis of sugarcane bagasse. Chem Eng J 2011;168:1157–62.

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  39 [74] Perez-Cantu L, Schreiber A, Schutt F, Saake B, Kirsch C, Smirnova I. Comparison of pretreatment methods for rye straw in the second generation biorefinery: effect on cellulose, hemicellulose and lignin recovery. Bioresour Technol 2013;142:428–35. [75] Allen SG, Kam LC, Zemann AJ, Antal MJ. Fractionation of sugar cane with hot, compressed, liquid water. Ind Eng Chem Res 1996;35:2709–15. [76] Wan CX, Zhou YG, Li YB. Liquid hot water and alkaline pretreatment of soybean straw for improving cellulose digestibility. Bioresour Technol 2011;102:6254–9. [77] Martin C, Klinke HB, Thomsen AB. Wet oxidation as a pretreatment method for enhancing the enzymatic convertibility of sugarcane bagasse. Enzyme Microb Technol 2007;40:426–32. [78] Zaldivar J, Nielsen J, Olsson L. Fuel ethanol production from lignocellulose: a challenge for metabolic engineering and process integration. Appl Microbiol Biotechnol 2001;56:17–34. [79] Shafizadeh F, Bradbury AGW. Thermal-degradation of cellulose in air and nitrogen at lowtemperatures. J Appl Polym Sci 1979;23:1431–42. [80] Yang B, Wyman CE. Pretreatment: the key to unlocking low-cost cellulosic ethanol. Biofuels Bioprod Biorefin 2008;2:26–40. [81] Fan LT, Gharpuray MM, Lee YH. Cellulose hydrolysis biotechnology monographs. Berlin: Springer; 198721–119. [82] Pitarelo AP, da Fonseca CS, Chiarello LM, Girio FM, Ramos LP. Ethanol production from sugarcane bagasse using phosphoric acid-catalyzed steam explosion. J Brazil Chem Soc 2016;27:1889–98. [83] Ghose TK, Bisaria VS. Studies on the mechanism of enzymatic-hydrolysis of cellulosic substances. Biotechnol Bioeng 1979;21:131–46. [84] Gamez S, Gonzalez-Cabriales JJ, Ramirez JA, Garrote G, Vazquez M. Study of the hydrolysis of sugar cane bagasse using phosphoric acid. J Food Eng 2006;74:78–88. [85] Wright JD. Ethanol from biomass by enzymatic-hydrolysis. Chem Eng Prog 1988;84:62–74. [86] Pattra S, Sangyoka S, Boonmee M, Reungsang A. Bio-hydrogen production from the fermentation of sugarcane bagasse hydrolysate by Clostridium butyricum. Int J Hydrogen Energy 2008;33:5256–65. [87] Kumar  V, Dangi  AK, Shukla  P. Engineering thermostable microbial xylanases toward its industrial applications. Mol Biotechnol 2018;60:226–35. [88] Hernandez-Salas  JM, Villa-Ramirez  MS, Veloz-Rendon  JS, Rivera-Hernandez  KN, Gonzalez-Cesar  RA, Plascencia-Espinosa  MA, Trejo-Estrada  SR. Comparative hydrolysis and fermentation of sugarcane and agave bagasse. Bioresour Technol 2009;100:1238–45. [89] Rezende CA, de Lima MA, Maziero P, deAzevedo ER, Garcia W, Polikarpov I. Chemical and morphological characterization of sugarcane bagasse submitted to a delignification process for enhanced enzymatic digestibility. Biotechnol Biofuels 2011;4:1–18. [90] Geddes  CC, Peterson  JJ, Roslander  C, Zacchi  G, Mullinnix  MT, Shanmugam  KT, Ingram  LO. Optimizing the saccharification of sugar cane bagasse using dilute phosphoric acid followed by fungal cellulases. Bioresour Technol 2010;101:1851–7. [91] Cheng CB, Wang JZ, Shen DK, Xue JT, Guan SP, Gu S, Luo KH. Catalytic oxidation of lignin in solvent systems for production of renewable chemicals: a review. Polymers 2017;9:240. [92] Boussarsar  H, Roge  B, Mathlouthi  M. Optimization of sugarcane bagasse conversion by hydrothermal treatment for the recovery of xylose. Bioresour Technol 2009;100:6537–42. [93] Kurakake  M, Ooshima  H, Harano  Y. Pretreatment of bagasse by UCT-solvent for the enzymatic-hydrolysis. Appl Biochem Biotech 1991;27:111–21. [94] Iram M, Asghar U, Irfan M, Huma Z, Jamil S, Nadeem M, et al. Production of bioethanol from sugarcane bagasse using yeast strains: a kinetic study. Energy Sources Part A: Recovery Util Environ Eff 2018;40:364–72.

40  SECTION | I  Enzymes in Bioprocessing [95] Ribeiro FR, Passos F, Gurgel LVA, Baeta BEL, de Aquino SF. Anaerobic digestion of hemicellulose hydrolysate produced after hydrothermal pretreatment of sugarcane bagasse in UASB reactor. Sci Total Environ 2017;584:1108–13. [96] Sendelius J. Steam pretreatment optimisation for sugarcane bagasse in bioethanol production. Sweden: Lund university; 2005. [97] Rocha  GJD, Martin  C, Soares  IB, Maior  AMS, Baudel  HM, de Abreu  CAM. Dilute mixed-acid pretreatment of sugarcane bagasse for ethanol production. Biomass Bioenergy 2011;35:663–70. [98] Cadete RM, Lopes MR, Rosa CA. Yeasts associated with decomposing plant material and rotting wood. In: Buzzini P, Lachance MA, Yurkov A, editors. Yeasts in natural ecosystems: diversity. Cham: Springer; 2017. [99] Li Y, Chen J, Lun SY. Biotechnological production of pyruvic acid. Appl Microbiol Biotechnol 2001;57:451–9. [100] Meshartree M, Dale BE, Craig WK. Comparison of steam and ammonia pretreatment for enzymatic-hydrolysis of cellulose. Appl Microbiol Biotechnol 1988;29:462–8. [101] Kumar V, Chhabra D, Shukla P. Xylanase production from Thermomyces lanuginosus VAPS24 using low cost agro-industrial residues via hybrid optimization tools and its potential use for saccharification. Bioresour Technol 2017;243:1009–19. [102] Tiwari  R, Singh  S, Shukla  P, Nain  L. Novel cold temperature active β-glucosidase from Pseudomonas lutea BG8 suitable for simultaneous saccharification and fermentation. RSC Adv 2014;4:58108–15. [103] Viswanath B, Rajesh B, Janardhan A, Kumar AP, Narasimha G. Fungal laccases and their applications in bioremediation. Enzyme Res 2014;2014:163242. [104] Gaj  T, Gersbach  CA, Barbas  CF. ZFN, TALEN, and CRISPR/Cas-based methods for genome engineering. Trends Biotechnol 2013;31:397–405. [105] Welker  CM, Balasubramanian  VK, Petti  C, Rai  KM, DeBolt  S, Mendu  V. Engineering plant biomass lignin content and composition for biofuels and bioproducts. Energies 2015;8:7654–76. [106] Vermaas JV, Petridis L, Qi XH, Schulz R, Lindner B, Smith JC. Mechanism of lignin inhibition of enzymatic biomass deconstruction. Biotechnol Biofuels 2015;8:217. [107] Hendriks ATWM, Zeeman G. Pretreatments to enhance the digestibility of lignocellulosic biomass. Bioresour Technol 2009;100:10–8. [108] Holtzapple  MT, Humphrey  AE, Taylor  JD. Energy-requirements for the size-reduction of poplar and aspen wood. Biotechnol Bioeng 1989;33:207–10. [109] Reshamwala  S, Shawky  BT, Dale  BE. Ethanol-production from enzymatic hydrolysates of afex-treated coastal bermudagrass and switchgrass. Appl Biochem Biotech 1995;5152:43–55. [110] Tengerdy RP, Nagy JG. Increasing the feed value of forestry waste by ammonia freeze explosion treatment. Biol Waste 1988;25:149–53. [111] Ramos LP. The chemistry involved in the steam treatment of lignocellulosic materials. Quim Nova 2003;26:863–71. [112] Vonsivers  M, Zacchi  G, Olsson  L, Hahnhagerdal  B. Cost-analysis of ethanol-production from willow using recombinant Escherichia coli. Biotechnol Prog 1994;10:555–60. [113] Bhagia S, Muchero W, Kumar R, Tuskan GA, Wyman CE. Natural genetic variability reduces recalcitrance in poplar. Biotechnol Biofuels 2016;9:106. [114] Selig M, Weiss N, Ji Y. Enzymatic saccharification of lignocellulosic biomass. Laboratory analytical procedure. Golden: US DOE National Renewable Energy Laboratory; 20081–8.

Production of Bioethanol From Sugarcane Bagasse  Chapter | 2  41 [115] Travaini R, Otero MDM, Coca M, Da-Silva R, Bolado S. Sugarcane bagasse ozonolysis pretreatment: Effect on enzymatic digestibility and inhibitory compound formation. Bioresour Technol 2013;133:332–9. [116] Balat  M, Balat  H, Oz  C. Progress in bioethanol processing. Prog Energy Combust Sci 2008;34:551–73. [117] Gauss WE, Suzuki S, Takagi M. Manufacture of alcohol from cellulosic material using plural fermenters. 1976. Google Patents. [118] Shevchenko SM, Bailey GW. The mystery of the lignin-carbohydrate complex: a computational approach. J Mol Struct: Theochem 1996;364:197–208. [119] Krishna SH, Prasanthi K, Chowdary GV, Ayyanna C. Simultaneous saccharification and fermentation of pretreated sugar cane leaves to ethanol. Process Biochem 1998;33:825–30. [120] Lynd LR, Cushman JH, Nichols RJ, Wyman CE. Fuel ethanol from cellulosic biomass. Science 1991;251:1318–23. [121] Mosier N, Hendrickson R, Ho N, Sedlak M, Ladisch MR. Optimization of pH controlled liquid hot water pretreatment of corn stover. Bioresour Technol 2005;96:1986–93. [122] Hilares RT, Swerts MP, Ahmed MA, Ramos L, da Silva SS, Santost JC. Organosolv pretreatment of sugar cane bagasse for bioethanol production. Ind Eng Chem Res 2017;56:3833–8. [123] Chandel AK, Antunes FF, Anjos V, Bell MJ, Rodrigues LN, Singh OV, da Silva SS. Ultrastructural mapping of sugarcane bagasse after oxalic acid fiber expansion (OAFEX) and ethanol production by Candida shehatae and Saccharomyces cerevisiae. Biotechnol Biofuels 2013;6:4. [124] Martin C, Thomsen MH, Hauggaard-Nielsen H, Belindathomsen A. Wet oxidation pretreatment, enzymatic hydrolysis and simultaneous saccharification and fermentation of cloverryegrass mixtures. Bioresour Technol 2008;99:8777–82. [125] Carvalho  W, Canilha  L, de Silva  SS. Semi-continuous xylitol bioproduction in sugarcane bagasse hydrolysate: effect of nutritional supplementation. Braz J Pharm Sci 2007;43:47–53. [126] Carvalheiro F, Duarte LC, Lopes S, Parajo JC, Pereira H, Girio FM. Evaluation of the detoxification of brewery's spent grain hydrolysate for xylitol production by Debaryomyces hansenii CCMI 941. Process Biochem 2005;40:1215–23. [127] Mohagheghi A, Evans K, Chou YC, Zhang M. Cofermentation of glucose, xylose, and arabinose by genomic DNA-integrated xylose/arabinose fermenting strain of Zymomonas mobilis AX101. Appl Biochem Biotechnol 2002;98-100:885–98. [128] Velmurugan R, Muthukumar K. Utilization of sugarcane bagasse for bioethanol production: sono-assisted acid hydrolysis approach. Bioresour Technol 2011;102:7119–23. [129] Diaz  MJ, Cara  C, Ruiz  E, Romero  I, Moya  M, Castro  E. Hydrothermal pre-treatment of rapeseed straw. Bioresour Technol 2010;101:2428–35. [130] Cheng  KK, Cai  BY, Zhang  JA, Ling  HZ, Zhou  YH, Ge  JP, Xu  JM. Sugarcane bagasse hemicellulose hydrolysate for ethanol production by acid recovery process. Biochem Eng J 2008;38:105–9. [131] Azhar SHM, Abdullaa R, Jambo SA, Marbawi H, Gansau JA, Mohd Faik AA, Francis Rodrigues  KF. Yeasts in sustainable bioethanol production: a review. Biochem Biophys Rep 2017;10:52–61. [132] Babu BV, Gupta S. Adsorption of Cr(VI) using activated neem leaves: kinetic studies. Adsorption 2008;14:85–92. [133] Perez JA, Ballesteros I, Ballesteros M, Saez F, Negro MJ, Manzanares P. Optimizing liquid hot water pretreatment conditions to enhance sugar recovery from wheat straw for fuel-­ ethanol production. Fuel 2008;87:3640–7.

42  SECTION | I  Enzymes in Bioprocessing [134] Roberto  IC, Lacis  LS, Barbosa  MFS, Demancilha  IM. Utilization of sugar-cane bagasse hemicellulosic hydrolysate by Pichia-Stipitis for the production of ethanol. Process Biochem 1991;26:15–21. [135] Van Zyl C, Prior BA, Du Preez JC. Production of ethanol from sugar cane bagasse hemiceilulose hydrolyzate by Pichia stipitis. Appl Biochem Biotechnol 1988;17:357–69. [136] Singh A, Sharma P, Saran AK, Singh N, Bishnoi NR. Comparative study on ethanol production from pretreated sugarcane bagasse using immobilized Saccharomyces cerevisiae on various matrices. Renew Energy 2013;50:488–93. [137] Christen P, Villegas E, Revah S. Growth and aroma production by Ceratocystis fimbriata in various fermentation media. Biotechnol Lett 1994;16:1183–8. [138] DiPardo  J. Outlook for biomass ethanol production and demand. In: Energy information administration. Washington, DC: US Department of Energy; 2000. [139] Wood PM. Pathways for production of Fenton's reagent by wood-rotting fungi. FEMS Microbiol Rev 1994;13:313–20. [140] Breccia  JD, Sineriz  F, Baigori  MD, Castro  GR, HattiKaul  R. Purification and characterization of a thermostable xylanase from Bacillus amyloliquefaciens. Enzyme Microb Tech 1998;22:42–9. [141] Delmas  M, Mlayah  BB. Process for producing bioethanol from lignocellulosic plant raw material. 2013. Google Patents. [142] Dien BS, Nichols NN, O’Bryan PJ, Bothast RJ. Development of new ethanologenic Escherichia coli strains for fermentation of lignocellulosic biomass. Appl Biochem Biotechnol 2000;84–86:181–96. [143] Kumbhar PS, Babu M, Sabale TR, Joshi SW, Pillanti RR. Preparation of ethanol from biomass and sugarcane based feed stocks. 2017. Google Patents. [144] Sant’Anna LMM, Pereira N, Bitancur GJV, Bevilaqua JV, da Conceicao Gomes A, Menezes EP. Process for producing ethanol from a hydrolysate of the hemicellulose fraction of sugarcane bagasse in a press reactor. 2014. Google Patents.

FURTHER READING [145] Biswas R, Uellendahl H, Ahring BK. Wet explosion pretreatment of sugarcane bagasse for enhanced enzymatic hydrolysis. Biomass Bioenergy 2014;61:104–13.

Chapter 3

Improvement of the Amino-Sugar-1-Phosphate Acetyltransferase Activity of the Archaeal Bifunctional Protein Yutaka Kawarabayasi National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba, Japan

1. INTRODUCTION The monomer form of a carbohydrate molecule is usually used for energy production, meanwhile its polymer structures fulfil a variety of important biological functions: energy storage, providing a barrier between the internal and external environment of the cells, and recognition of foreign cells. The nucleotide-sugar molecule (e.g., GDP-mannose), which is synthesized from NTP and sugar1-phosphate (Sugar-1-P) by the function of sugar-1-phosphate nucleotidylyltransferase (Sugar-1-P NTase), is used as the sole substrate for construction of the polymer structure of carbohydrate. Among a variety of nucleotide-sugar molecules, uridine diphosphate N-acetylglucosamine (UDP-GlcNAc) is one of the most important substrate compounds and it is particularly important for incorporation of GlcNAc moiety into the polymer structure of carbohydrate. It is well known that from bacteria to eukaryote UDP-GlcNAc is generally constructed from fructose-6-phosphate (Frc-6-P) via a four-step reaction. Archaea were distinguished from bacteria approximately 40 years ago on the basis of 16S rRNA gene sequences [1]. Many species of archaea have been isolated from hydro- or geothermal environments, indicating that these archaea species specifically grow only at high temperatures. The proteins identified from these thermophilic archaea exhibit extreme stability at high temperatures and under other stressful conditions. This stability is advantageous for functional analyses and determination of the crystal structure, as well as for industrial applications. Applied Microbiology and Bioengineering. https://doi.org/10.1016/B978-0-12-815407-6.00003-4 © 2019 Elsevier Inc. All rights reserved.

43

44  SECTION | I  Enzymes in Bioprocessing

Archaea are known as chimeric microorganisms possessing features similar to both bacteria and eukarya. Archaea possess unique metabolic pathways, such as the nonphosphorylated Entner-Doudoroff pathway for glucose metabolism in species of the genus Sulfolobus [2]. Experimental analyses of enzymes identified from an acidothermophlic archaeon Sulfolobus tokodaii [3] indicated that they exhibit unique properties, such as the ability to accept multiple substrates and utilize nontypical metal ions [4]. The uniqueness of archaea species and their enzymes suggests that many as-yet unidentified novel and useful enzymes or proteins are encoded within the archaeal genome. The size of the archaeal genome already determined is relatively small compared with the bacteria and eukarya, which suggests that fewer genes are encoded in the archaeal genome. From the entire genomic data of the thermophilic archaeon Sulfolobus tokodaii determined around 15 years ago [5], the ST0452 protein was identified as a bifunctional protein with multiple Sugar-1-P NTase and multiple amino-sugar-1-phosphate acetyltransferase (amSugar-1-P AcTase) activities [4,6]. This review describes the overview of the activities including the Nacetylglucosamine-1-phosphate uridyltransferase (GlcNAc-1-P UTase) activity encoded within the N-terminus of the ST0452 protein, and characterization plus improvement of the amSugar-1-P AcTase activity encoded within the C-terminal domain of the bifunctional ST0452 protein.

2.  IDENTIFICATION OF THE ST0452 PROTEIN AND ITS SUGAR-1-P NTASE ACTIVITY A putative 401 amino acid long ST0452 protein was originally predicted as a glucose-1-phosphate thymidylyltransferase (Glc-1-P TTase) based on sequence similarity [4]. Recombinant ST0452 protein produced in Escherichia coli exhibited multiple Sugar-1-P NTase activities, as shown in Table 1 [4]. Detailed analyses of the Sugar-1-P NTase activities of the ST0452 protein indicated that the optimal pH and temperature were respectively 8.0°C and 90°C [4]. The Sugar-1-P NTase activity of the ST0452 protein also accepts multiple metal cations, with the highest activity in the presence of Co2+ [4]. Analyses of the kinetic parameters of the GlcNAc-1-P UTase activity of the ST0452 protein indicated that both the apparent Km and kcat values are lower than those of E. coli GlmU [7]. As proteins with a low Km value can accept and utilize low-­ concentration substrates, this property is convenient for industrial or other applications. However, low kcat values are indicative of slower turnover rates, which is a hindrance with respect to applications. Thus improvement of the Sugar-1-P NTase activity of the ST0452 protein was attempted by generating targeted mutants substituted to Ala or Phe [7]. Five of the 11 ST0452 substitution mutants constructed exhibited increased GlcNAc-1-P UTase activity in the presence of high concentrations of substrate (Fig. 1) [7], suggesting that the activity of archaeal thermostable enzymes can be improved by introducing single substitution mutation within the reaction center.

The Archaeal Bifunctional Protein  Chapter | 3  45

TABLE 1  Substrate Specificity of the Sugar-1-P NTase Activity of the ST0452 Protein Nucleotide Substrates

Sugar-1-P Substrates

Relative Activity (%)

dTTP

α-d-glucose-1-phosphate

100

dATP

35

dCTP

7

dGTP

1

UTP

130

ATP/CTP/GTP

ND

dTTP

UTP

N-acetylglucosamine-1phosphate

320

α-d-glucosamine-1-phosphate

ND

α-d-galactose-1-phosphate

ND

α-d-mannose-1-phosphate

ND

N-acetylglucosamine-1phosphate

540

α-d-glucosamine-1-phosphate

ND

α-d-galactose-1-phosphate

ND

α-d-mannose-1-phosphate

ND

Relative activity is shown as a percentage of the activity detected for reaction using Glc-1-P and TTP as the substrate. ND indicates nondetectable.

It was expected that the combination of two single substitution mutations to Ala or Phe in the mutant ST0452 protein might increase its GlcNAc-1-P UTase activity. All recombinant double-mutant proteins constructed were successfully expressed in E. coli as thermostable and soluble forms. The Sugar1-P NTase activity of these proteins was measured; however, unfortunately, all double-mutant proteins exhibited drastically decreased activity (Fig.  2) [8]. This means that the introduction of two first-generation single mutations might result in the failure of appropriate interactions between substrate and proteins. To counteract this problem a saturation-mutagenesis strategy was applied to the 97th amino acid residue of the ST0452 protein, which indicated the highest GlcNAc-1-P UTase activity when converted to Ala. Of 19 mutant proteins constructed, six mutant proteins exhibited an increase in both the GlcNAc-1-P UTase and Glc-1-P UTase activities, and eight mutant proteins indicated only

46  SECTION | I  Enzymes in Bioprocessing

Relative activity

2.0

1.0

0 ST0452

G9A

T80A

Y97A

Y97F

K147A

FIG.  1  N-acetylglucosamine-1-phosphate uridyltransferase (GlcNAc-1-P UTase) activity of the first-generation mutant ST0452 proteins under optimized conditions. GlcNAc-1-P UTase activities of each mutant protein were measured in the reaction solution containing 100 μM UTP and 10 mM GlcNAc-1-P. The relative activity is expressed as a ratio of the activity detected on the wild-type ST0452 protein under the same conditions. All experiments were repeated four times.

2.0

Relative activity

1.5

1.0

0.5

0

C

1

2

3

4

5

6

7

8

9

FIG.  2  N-acetylglucosamine-1-phosphate uridyltransferase (GlcNAc-1-P UTase) activity of the wild-type and double-mutant ST0452 proteins. Enzymatic activities were measured for 2 min at 80°C. The relative activity is expressed as a ratio of the activity measured for the wild-type ST0452 protein. Lane C, wild-type ST0452 protein; Lane 1, G9A/T80A; Lane 2, G9A/Y97A; Lane 3, G9A/Y97F; Lane 4, G9A/K147A; Lane 5, T80A/Y97A; Lane 6, T80A/Y97F; Lane 7, T80A/K147A; Lane 8, Y97A/K147A; Lane 9, Y97F/K147A. All experiments were repeated three times.

The Archaeal Bifunctional Protein  Chapter | 3  47

Relative activity

5.0 4.0 3.0 2.0 1.0 0

Y A C D

E F G H I

K L M N P Q R S T V W

FIG.  3  N-acetylglucosamine-1-phosphate uridyltransferase (GlcNAc-1-P UTase) and glucose1-phosphate uridyltransferase (Glc-1-P UTase) activities of the wild-type and Y97 series singlemutant ST0452 proteins. The relative activities of GlcNAc-1-P UTase activity (open bars) and Glc-1-P UTase (hatched bars) are shown. The relative activity is expressed as a ratio of each activity measured in the wild-type ST0452 protein. Lane Y, wild-type ST0452 protein; Lane A, Y97A; Lane C, Y97C; Lane D, Y97D; Lane E, Y97E; Lane F, Y97F; Lane G, Y97G; Lane H, Y97H; Lane I, Y97I; Lane K, Y97K; Lane L, Y97L; Lane M, Y97M; Lane N, Y97N; Lane P, Y97P; Lane Q, Y97Q; Lane R, Y97R; Lane S, Y97S; Lane T, Y97T; Lane V, Y97V; Lane W, Y97W. All experiments were repeated three times. The symbols shown in squares are the mutant proteins indicating an increase in both GlcNAc-1-P UTase and Glc-1-P UTase activities. The underlined symbols are the mutant proteins demonstrating only increased GlcNAc-1-P UTase activity.

increased GlcNAc-1-P UTase activity, although the remaining five mutant proteins showed decreased activities (Fig. 3) [8]. The Y97N mutant protein exhibited a four-times higher GlcNAc-1-P UTase activity (the highest activity among mutant proteins constructed as this series) than that of the wild-type ST0452 protein [8]. These observations indicate the possibility of further increasing of the enzymatic activity in the ST0452 protein by introducing another type of mutation. In the following section the results of an experimental analyses obtained for another activity of the bifunctional ST0452 protein will be described.

3.  CHARACTERIZATION OF THE AMSUGAR-1-P ACTASE ACTIVITY OF THE ST0452 PROTEIN Repeat motif sequences with an LßH structural feature were detected in the long C-terminal region of the ST0452 protein, as shown in Fig. 4. Since these repeat motif sequences are usually found within acyl- or acetyltransferase enzymes, the acetyltransferase activity of the ST0452 protein was examined. Glucosamine-1-phosphate acetyltransferase (GlcN-1-P AcTase) activity of the ST0452 protein was evaluated by two independent experiments; detection of the conversion of acetyl coenzyme A (acetyl-CoA), which is used as a donor of acetyl group in this reaction, to CoA and formation of UDP-GlcNAc [6]. Conversion of acetyl-CoA to CoA was observed only when the ST0452 protein

48  SECTION | I  Enzymes in Bioprocessing

FIG.  4  Amino acid sequence of the C-terminal region of the ST0452 protein. Amino acid sequence from residue 234 to the C-terminus of the ST0452 protein is aligned for each turn of the LßH structure. The residues in parentheses comprise the extra loop region. Residues with asterisks were chosen for substitution mutation. The numerals shown to the right side of each line indicate the coordination of residues shown at the right end of each line.

was added to the reaction mixture [6]. Only when GlcN-1-P was converted to GlcNAc-1-P by the function of the ST0452 acetyltransferase activity, in the presence of UTP the GlcNAc-1-P UTase activity of the ST0452 protein could construct UDP-GlcNAc from GlcNAc-1-P, a product of the acetyltransferase reaction. Detection of the UDP-GlcNAc produced in the reaction mixture containing GlcN-1-P, acetyl-CoA, and UTP therefore indicates the presence of acetyltransferase activity. As expected UDP-GlcNAc production was detected in the presence of UTP and acetyl-CoA (Fig. 5), confirming that the ST0452 protein exhibits the GlcN-1-P AcTase activity [6]. The substrates shown in Table  2 were analyzed in order to characterize the substrate specificity of the ST0452 acetyltransferase activity. Within the biosynthetic pathway of UDP-GlcNAc from Frc-6-P, GlcN-1-P is acetylated in the bacterial pathway; conversely, in eukaryotes, acetylation occurs only on glucosamine-6-phosphate (GlcN-6-P). To evaluate which type of acetylation reaction is catalyzed by the ST0452 protein, acceptability of two distinct compounds, GlcN-1-P and GlcN-6-P, were analyzed as substrate. Conversion of acetyl-CoA to CoA was not detected when GlcN-6-P was added as the substrate, but CoA production was detected only when GlcN-1-P was used as the

The Archaeal Bifunctional Protein  Chapter | 3  49

(A)

(B)

UD

UD P Ga -Gl lNA cNA c c

P-

(C)

UT

P

FIG.  5  High-performance liquid chromatography elution profile of the products of the glucosamine-1-phosphate acetyltransferase (GlcN-1-P AcTase) activity of the ST0452 protein. The reaction proceeded at 80°C for 10 min in 10 μL of standard acetyltransferase reaction solution [50 mM Tris-HCl (pH 7.5), 2 mM MgCl2, and 50 ng of the recombinant ST0452 protein] with 0.1 mM UTP and 2 mM GlcN-1-P (A) or with 0.1 mM UTP, 2 mM acetyl-CoA, and 2 mM GlcN-1-P (B). Elution profile of a mixture of standard UTP, UDP-GlcNAc, and UDP-GalNAc (C).

substrate, revealing that a bacteria-type UDP-GlcNAc biosynthetic pathway is present in this archaeon. Surprisingly, conversion of acetyl-CoA to CoA was detected when galactosamine-1-phosphate (GalN-1-P) was added as the substrate, as shown in Table 2. The galactosamine-1-phosphate acetyltransferase (GalN-1-P AcTase) activity of the ST0452 protein was confirmed by exploiting the coupling activity of the same protein’s Sugar-1-P NTase activity. As shown in Fig. 6, the expected peak was detected at the same elution position as the standard UDP-N-acetylgalactosamine (UDP-GalNAc), revealing that the ST0452 protein also possesses GalN-1-P AcTase activity [6].

50  SECTION | I  Enzymes in Bioprocessing

TABLE 2  Substrate Specificity of the Acetyltransferase Activity of the ST0452 Protein Substrates

Relative Activity (%)

Glucosamine-1-phosphate

100

Glucosamine-6-phosphate

ND

Galactosamine-1-phosphate

51

Relative activity is shown as a percentage of the activity detected for Glucosamine-1-phosphate. ND indicates nondetectable.

(A)

(B)

UD

UD P Ga -Gl lNA cNA c c

P-

(C)

UT

P

FIG.  6  High-performance liquid chromatography elution profile of the products of the galactosamine-1-phosphate acetyltransferase (GalN-1-P AcTase) activity of the ST0452 protein. The reaction proceeded at 80°C for 10 min in 10 μL of standard acetyltransferase reaction solution [50 mM Tris-HCl (pH 7.5), 2 mM MgCl2, and 50 ng of the recombinant ST0452 protein] with 0.1 mM UTP and 2 mM GalN-1-P (A) or with 0.1 mM UTP, 2 mM acetyl-CoA, and 2 mM GalN-1-P (B). Elution profile of a mixture of standard UTP, UDP-GlcNAc, and UDP-GalNAc (C).

The Archaeal Bifunctional Protein  Chapter | 3  51

These analyses also indicated that the ST0452 protein might exhibit GalNAc1-P UTase activity. Confirmation of the forward direction of this reaction requires GalNAc-1-P, but this compound is not commercially available. Therefore the reverse direction of the expected GalNAc-1-P UTase activity in the ST0452 protein was analyzed. As shown in Fig. 7, production of UTP from UDP-GalNAc and PPi was detected, revealing that the ST0452 protein possesses GalNAc-1-P UTase activity in addition to GlcNAc-1-P UTase and Glc-1-P TTase activities [6]. The amSugar-1-P AcTase activity of the ST0452 protein was then fully characterized.

(A)

(B)

(C)

UTP

UDP-GalNAc

FIG.  7  High-performance liquid chromatography elution profile of the products of the reverse N-acetylgalactosamine-1-phosphate uridyltransferase (GalNAc-1-P UTase) reaction. Reaction solution [50 mM Tris-HCl (pH 7.5), 2 mM MgCl2, 0.5 mM pyrophosphate, 5 mM UDP-GalNAc and 50 ng of the recombinant ST0452 protein] was incubated for 0 min (A), 5 min (B), or 10 min (C). The elution positions of standard UTP and UDP-GalNAc are shown by arrows.

52  SECTION | I  Enzymes in Bioprocessing

4.  CHARACTERIZATION BY INTRODUCTION OF SUBSTITUTION MUTATIONS To elucidate the roles of the amino acid residues located within the amSugar1-P AcTase reaction center of the ST0452 protein, substitution mutations were introduced into targeting residues predicted as being important based on the three-dimensional structure of the ST0452 protein modeled from the determined three-dimensional structure of bacterial GlmU proteins [9]. Five residues (residues at 308th, 311th, 331st, 377th and 340th positions) were chosen as targets for Ala substitution (Fig. 4). The GlcN-1-P AcTase and GalN-1-P AcTase activities of the constructed mutant ST0452 proteins were analyzed. All constructed mutant proteins exhibited decreased GalN-1-P AcTase activity (Table  3) [10]. And mutant proteins substituted with an Ala at 308th and 331st residues resulted in a decrease in both the GlcN-1-P and GalN-1-P AcTase activities [10]. It was hypothesized based on these observations that residues at 308th and 311st positions play an important or critical role in the AcTase activity of the ST0452 protein. Substitution of 308th His residue with an Ala reduced both AcTase activities to