Enzymes in Degradation of the Lignocellulosic Wastes 3030446700, 9783030446703

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Aparna B. Gunjal Neha N. Patil Sonali S. Shinde

Enzymes in Degradation of the Lignocellulosic Wastes

Enzymes in Degradation of the Lignocellulosic Wastes

Aparna B. Gunjal Neha N. Patil Sonali S. Shinde •

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Enzymes in Degradation of the Lignocellulosic Wastes

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Aparna B. Gunjal Department of Microbiology Dr. D. Y. Patil, Arts, Commerce and Science College Pimpri, Pune, Maharashtra, India

Neha N. Patil Department of Microbiology Annasaheb Magar Mahavidyalaya Pune, Maharashtra, India

Sonali S. Shinde Annasaheb Kulkarni Department of Biodiversity MES Abasaheb Garware College Pune, Maharashtra, India

ISBN 978-3-030-44670-3 ISBN 978-3-030-44671-0 https://doi.org/10.1007/978-3-030-44671-0

(eBook)

© Springer Nature Switzerland AG 2020 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Foreword by Dr. Chaitanya Kumar Jha

Waste generation and management in the world is a very serious issue from the point of environmental protection and human, animal and plant health. Huge amount of lignocellulosic wastes, viz. peanut shell, corn cob, rice straw, wheat straw, bagasse, press mud, and coconut husk are generated which is either incinerated or directly disposed to the landfills. The degradation of lignocellulosic wastes is not very easy and requires solution. The book Enzymes in Degradation of the Lignocellulosic Wastes gives detailed information and knowledge about the use of enzymes in degradation of lignocellulosic wastes. This book highlights the information of cellulase, hemicellulase, ligninase, pectinase, and lipase enzymes and the detail mechanisms of these enzymes in degradation of lignocellulosic wastes. This book gives the idea of different cellulase, hemicellulase-producing microorganisms, and their catalytic mechanisms to breakdown cellulose and hemicellulose, respectively. The detail role of ligninase enzymes, viz. laccases, peroxidases, manganese peroxidase, and versatile peroxidase (VP) for degradation of lignin, is also mentioned. The aspect of pectinase enzyme in degradation of lignocellulosic wastes is also focused, where the pectic substances and mechanism of action of the pectinolytic enzymes are described. The catalytic mechanism of lipases in degradation of lignocellulosic wastes and lipase-producing microorganisms is also described. This book also focuses on the assays methods for cellulase, hemicellulase, ligninase, pectinase, and lipase enzymes as well as applications of each of these enzymes. This book is useful to college students, researchers, and other scientists and is an excellent guide that provides solution for degradation of lignocellulosic wastes which is very important.

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Foreword by Dr. Chaitanya Kumar Jha

I have no doubt; this book will be an important milestone in this direction. I wish the authors all the very best!

Dr. Chaitanya Kumar Jha Assistant Professor Department of Microbiology Gujarat Arts and Science College Ahmedabad, Gujarat, India Dr. Chaitanya Kumar Jha is Assistant Professor and Head of Department of Microbiology at Gujarat Arts and Science College, Ahmedabad, Gujarat, India. He completed his Ph.D in Microbiology in 2011 from Gujarat University. He has 10 years of teaching experience. His research areas of interest are microbiology and biotechnology. He has 23 publications to his credit which includes research articles, book chapters and review articles. He has presented many research papers in National and International Conferences during which he has received best paper presentation award. He has also supervised M.Sc. and M.Phil students for their research work. He is a reviewer for the International Journals, viz. Cogent Food and Agriculture; Journal of Basic Microbiology; and 3 Biotech. He is also member of many societies, viz. Association of Microbiologists of India; Asian PGPR Society; and Indian Science Congress.

Foreword by Dr. Manojkumar Z. Chopda

Waste is generated in huge amount which is a serious problem. The lignocellulosic wastes, viz. peanut shell, corn cob, rice straw, wheat straw, bagasse, press mud, and coconut husk are generated which are either incinerated or directly disposed to the landfills. The degradation of lignocellulosic wastes is difficult and requires to be solved. The book Enzymes in Degradation of the Lignocellulosic Wastes gives information and knowledge about different enzymes which can degrade lignocellulosic wastes. This book highlights the use of enzymes, viz. cellulase, hemicellulase, ligninase, pectinase, and lipases and mechanisms of these enzymes in degradation of lignocellulosic wastes. This book gives the idea of different cellulase, hemicellulase-producing microorganisms and their catalytic mechanisms to breakdown cellulose and hemicellulose, respectively. The role of ligninase enzymes for degradation of lignin is also described. The aspect of pectinase enzyme in degradation of lignocellulosic wastes is also focused, where the pectic substances and mechanism of action of the pectinolytic enzymes is described. The lipase-producing microorganisms and mechanism of lipases in degradation of lignocellulosic wastes are also described. This book also focuses on assays methods for cellulase, hemicellulase, ligninase, pectinase, and lipase enzymes and applications of these enzymes. This book will be useful to college students, researchers, and other scientists and is an excellent guide that will provide solution for degradation of lignocellulosic wastes which is very important.

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Foreword by Dr. Manojkumar Z. Chopda

I wish the authors all the very best!

Dr. Manojkumar Z. Chopda Assistant Professor Department of Zoology Moolji Jaitha College Jalgaon, Maharashtra, India Dr. Manojkumar Z. Chopda is working as Assistant Professor, Department of Zoology at Moolji Jaitha College, Jalgaon, Maharashtra, India. He completed his Ph.D in Zoology in 2009 from North Maharashtra University. He has 15 years of teaching experience. His research areas of interest are medicinal plants and their biological activities and biodiversity. He has 55 publications to his credit which includes research articles, books, book chapters, and review articles. He has presented many research papers in National and International Conferences during which he has received best paper presentation award. Dr. Manojkumar Chopda has supervised many B. Sc., M.Sc., and Ph. D. students for their research work. He is also a reviewer for the National and International Journals. He has also completed Major Research Projects as Principal Investigator sanctioned by University Grants Commission, New Delhi, India. He is an expert committee member of Animal Dissection Reform, constituted by UGC, New Delhi.

Foreword by Dr. Dilan Sanjitha Rajapakshe

I am happy to give the foreword for the book Enzymes in Degradation of the Lignocellulosic Wastes. Different types of wastes are generated in the world which is a very serious issue from the point of environmental protection and human, animal, and plant health. In the category of wastes, huge amount of lignocellulosic wastes, viz. peanut shell, corn cob, rice straw, wheat straw, bagasse, press mud, and coconut husk are also generated. These lignocellulosic wastes are either incinerated or disposed to the landfills. The degradation of lignocellulosic wastes needs to be solved. This book Enzymes in Degradation of the Lignocellulosic Wastes describes in detail about the use of different enzymes, viz. cellulase, hemicellulase, ligninase, pectinase, and lipases for the degradation of lignocellulosic wastes. This book mentions different cellulase, hemicellulase-producing microorganisms, and their mechanisms to breakdown cellulose and hemicellulose, respectively. The ligninase enzymes, viz. laccases, peroxidases, manganese peroxidase, and versatile peroxidase (VP) for the degradation of lignin, are also highlighted. The important pectinase- and lipase-producing microorganisms are mentioned in this book. The catalytic mechanisms of action of the pectinolytic and lipases enzymes in degradation of lignocellulosic wastes are also described. This book also focuses on the assays methods for cellulase, hemicellulase, ligninase, pectinase, and lipase enzymes along with applications of each of these enzymes. This book is helpful to college students, researchers and other scientists, and also best guide that provides solution for degradation of lignocellulosic wastes.

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Foreword by Dr. Dilan Sanjitha Rajapakshe

I have no doubt; this book will be a real wonderful milestone in this direction. I wish the authors all the very best!

Dr. Dilan Sanjitha Rajapakshe Post Graduate Institute of Science University of Peradeniya Peradeniya, Sri Lanka Dr. Dilan Sanjitha Rajapakshe is Visiting Faculty at Department of Chemistry, Postgraduate Institute of Science, University of Peradeniya, Perideniya, Srilanka; Technological Studies, Uwa Wellassa University; and Australian College of Business and technology, Kandy, Sri Lanka. He completed his Ph.D in Chemistry in 2019 from University of Peradeniya, Peradeniya, Sri Lanka. He has three years of teaching experience. His research areas of interest are material chemistry; nanostructures; molecular biology; photochemistry; optics and renewable energy; and biotechnology. He has nine publications to his credit which includes research articles and book chapters. He has presented many research papers in National and International Conferences. He has also received travel grants, viz. Taiwan Travel Grant for Symposium on Energy, Environment and Technology at Taiwan; and ITEC Grant; SAKURA Grant and NSF Technology Grant for his research work. He has also supervised M.Sc. students for their research work. He has co-supervised many projects related to nanoscience and nanotechnology at the industrial level, viz. super-hydrophobic textiles in bulk content at Textured Jerseys Ltd., Sri Lanka; iron oxide nanomaterials from Galvanized Industrial Waste, LTL, Sri Lanka Transformers Ltd., Makola, Biyagama; treatment of industrial effluent using photocatalysts, at Brandix India Apparel City, Visakhapatnam, Andhra Pradesh, India, etc. He is also member of many societies, viz. Sri Lanka Academy of Young Scientists and Young Researcher’s Forum, University of Peradeniya, Sri Lanka.

Preface

The lignocellulosic wastes are generated in large amount and degradation of lignocellulosic wastes is a very serious issue and which needs to be given attention. Different lignocellulosic wastes are generated, viz. wheat bran, corn cob, sawdust, rice straw, coconut husk, bagasse, and peanut shell. The degradation of these lignocellulosic wastes needs to be solved. The book Enzymes in Degradation of the Lignocellulosic Wastes deals with the use of enzymes for the degradation of lignocellulosic wastes. The authors have contributed on various enzymes and their mechanisms for this aspect. Chapter 1 gives the introduction part which gives information on different lignocellulosic wastes and their components and problems in degradation of lignocellulosic wastes. The value-added products from lignocellulosic wastes are also mentioned. The classification of enzymes and their advantages in degradation of lignocellulosic wastes is described. The microorganism-degrading lignocellulosic wastes are also mentioned. Chapter 2 describes the role of cellulase in degradation of lignocellulosic wastes. The chapter here describes the structure of cellulose; cellulase production by fermentation; cellulose-degrading microorganisms; and enzymes which breakdown cellulose and cellulase systems of microorganisms. The chapter also focuses on cellulosomes to degrade cellulose; cellulose hydrolysis mechanisms; determination of cellulase activity; and applications of cellulase enzyme. Chapter 3 describes the role of hemicellulase in degradation of lignocellulosic wastes. The chapter mentions the structure of hemicellulose; families of hemicellulase enzyme; and hemicellulase production by microorganisms. The chapter also describes fungi in degradation of hemicellulose; role of transcriptional regulators in regulation of xylanolytic gene expression; hemicellulase enzymes and their activity; and applications of hemicellulase enzyme. Chapter 4 describes the role of ligninase for degradation of lignocellulosic wastes. Microorganisms produce ligninase enzymes, viz. lignin peroxidase, manganese peroxidase, versatile peroxidase, and laccase. These enzymes play an important role in lignin degradation, and this chapter describes the role of these enzymes for degradation of lignocellulosic wastes. xi

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Chapter 5 describes in detail the pectic substances and mechanism of various pectinase enzymes in degradation of lignocellulosic wastes. Chapter 6 describes the role of lipases enzyme in degradation of lignocellulosic wastes. The chapter describes the classification and structure of lipase enzyme. It also mentions catalytic mechanisms of lipase enzyme. The chapter also focuses on lipase-producing microorganisms; assays for lipase activity, and various applications of lipases enzyme. The target audience for this book will be students from schools, colleges, and universities and researchers working worldwide on lignocellulosic wastes. This book will provide a guide for the degradation of lignocellulosic wastes by the enzymes and also help researchers develop new ideas for the study of enzymes and their mechanisms in degradation of lignocellulosic wastes. Aparna B. Gunjal Assistant Professor Department of Microbiology Dr. D. Y. Patil, Arts, Commerce and Science College Pimpri, Pune, Maharashtra, India Neha N. Patil Head and Associate Professor Department of Microbiology Annasaheb Magar Mahavidyalaya Hadapsar, Pune, Maharashtra, India Sonali S. Shinde Assistant Professor Annasaheb Kulkarni Department of Biodiversity MES Abasaheb Garware College Pune, Maharashtra, India

Contents

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1 Wastes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.1 Lignocellulosic Wastes . . . . . . . . . . . . . . . . . . . . . 1.2 Lignocellulosic Wastes and Problems in Their Degradation . 1.3 Linkages Between Lignocellulose Components . . . . . . . . . . 1.4 Value-Added Products from Lignocellulosic Wastes . . . . . . 1.5 Types of Lignocellulosic Biomass . . . . . . . . . . . . . . . . . . . 1.5.1 Dry Grass . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.5.2 Forest Woody Feedstock . . . . . . . . . . . . . . . . . . . 1.5.3 Municipal Solid Wastes . . . . . . . . . . . . . . . . . . . . 1.5.4 Agricultural Residues . . . . . . . . . . . . . . . . . . . . . . 1.6 Bioprocessing of Lignocellulose Wastes . . . . . . . . . . . . . . . 1.7 Advantages of Enzymes in Degradation of Lignocellulose Wastes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.7.1 Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.7.2 Enzymes in Lignocellulose Degradation . . . . . . . . 1.8 Fungi-Degrading Lignocellulosic Wastes . . . . . . . . . . . . . . 1.8.1 Wood-Decaying Fungi . . . . . . . . . . . . . . . . . . . . . 1.8.2 Brown-Rot Fungi . . . . . . . . . . . . . . . . . . . . . . . . . 1.8.3 White-Rot Fungi . . . . . . . . . . . . . . . . . . . . . . . . . 1.8.4 Soft-Rot Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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2 Cellulase in Degradation of Lignocellulosic Wastes . . . . . . . 2.1 Cellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Microcrystalline Cellulose . . . . . . . . . . . . . . . . . . . . . . 2.3 Cellulose Structure, Function, and Properties . . . . . . . . 2.4 Cellulase Production . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4.1 Microorganisms Used for Cellulase Production 2.5 Enzymes Involved in Cellulose Breakdown . . . . . . . . .

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Cellulase Systems of Bacteria and Fungi . . . . . . . . . . . . 2.6.1 Bacterial Cellulase Systems . . . . . . . . . . . . . . . 2.6.2 Fungi Cellulase Systems . . . . . . . . . . . . . . . . . . 2.7 Microbial Sources of Cellulase Enzyme . . . . . . . . . . . . . 2.7.1 Bacteria Sources . . . . . . . . . . . . . . . . . . . . . . . 2.7.2 Actinobacteria Sources . . . . . . . . . . . . . . . . . . . 2.7.3 Fungi Sources . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Cellulosomes to Degrade Cellulose . . . . . . . . . . . . . . . . 2.8.1 Carbohydrate-Active enZymes (CAZymes) . . . . 2.8.2 Cellobiose Dehydrogenase (CDH) . . . . . . . . . . . 2.9 Cellulolytic Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . 2.10 Assay Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.10.1 Determination of Cellulase Activity (CMCase Assay) . . . . . . . . . . . . . . . . . . . . . . . 2.11 Physical and Biological Methods for Pretreatment of Cellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.11.1 Physical Methods for Pretreatment of Cellulose . 2.11.2 Biological Methods . . . . . . . . . . . . . . . . . . . . . 2.12 Mechanism of Cellulose Hydrolysis . . . . . . . . . . . . . . . . 2.12.1 Genes Related to Cellulose Degradation . . . . . . 2.13 Transcriptional Regulators Involved in Regulation of Cellulolytic Gene Expression (T. reesei) (Shida et al. 2016) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.13.1 Xylanase Regulator (Xyr1) . . . . . . . . . . . . . . . . 2.13.2 Activator of Cellulase Expression 2 (Ace2) . . . . 2.13.3 Activator of Cellulase Expression 1 (Ace1) . . . . 2.13.4 Beta-Glucosidase Regulator (BglR) . . . . . . . . . . 2.13.5 Activator of Cellulase Expression 3 (Ace3) . . . . 2.13.6 GH Families Involved in Cellulose Degradation . 2.14 Applications of Enzyme Cellulase . . . . . . . . . . . . . . . . . 2.14.1 Paper and Pulp Industries . . . . . . . . . . . . . . . . . 2.14.2 Textile Industry . . . . . . . . . . . . . . . . . . . . . . . . 2.14.3 Food and Feeds . . . . . . . . . . . . . . . . . . . . . . . . 2.14.4 Bioethanol . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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3 Hemicellulase in Degradation of Lignocellulosic Wastes . . . . . . . 3.1 Hemicellulose Structure and Property . . . . . . . . . . . . . . . . . . 3.2 Xylan Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Families of Enzyme Hemicellulase . . . . . . . . . . . . . . . . . . . 3.4 Hemicellulase Production by Microorganisms . . . . . . . . . . . . 3.5 Brown-Rot, White-Rot and Soft-Rot Fungi in Degradation of Hemicellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 Transcriptional Regulators Involved in Regulation of Xylanolytic Gene Expression (T. Reesei) (Shida et al. 2016) .

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Methods for Pretreatment of Hemicellulose . . . . . . . 3.7.1 Alkaline Pretreatment . . . . . . . . . . . . . . . . . 3.7.2 Wet Oxidation . . . . . . . . . . . . . . . . . . . . . . 3.7.3 Acid Pretreatment . . . . . . . . . . . . . . . . . . . . 3.7.4 Green Solvents . . . . . . . . . . . . . . . . . . . . . . 3.7.5 Steam-Explosion Pretreatment . . . . . . . . . . . 3.8 Enzymes for Hemicellulose Biodegradation . . . . . . . 3.8.1 Hemicellulase Enzymes . . . . . . . . . . . . . . . 3.9 Method for Measurement of Hemicellulase Enzyme Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.9.1 Laminarinase (Linton and Greenaway 2004) 3.9.2 Licheninase (Linton and Greenaway 2004) . 3.9.3 Xylanase (Linton and Greenaway 2004) . . . 3.10 Applications of Hemicellulase Enzyme . . . . . . . . . . 3.10.1 In Paper and Pulp Industries . . . . . . . . . . . . 3.10.2 Processing of Animal Feed . . . . . . . . . . . . . 3.10.3 Beverage Industry . . . . . . . . . . . . . . . . . . . 3.10.4 Bakery Industry . . . . . . . . . . . . . . . . . . . . . 3.10.5 Pharmaceutical Industry . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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4 Ligninase in Degradation of Lignocellulosic Wastes . . . . . 4.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Lignin Occurrence, Biogenesis, and Biodegradation . . 4.3 Mechanisms of Lignin Degradation . . . . . . . . . . . . . . 4.4 Enzymes Involved in the Degradation . . . . . . . . . . . . 4.4.1 Lignin Peroxidase (LiP) (EC 1.11.1.14) . . . . . 4.4.2 Manganese Peroxidase (MnP) (EC 1.11.1.13) 4.4.3 Versatile Peroxidase . . . . . . . . . . . . . . . . . . . 4.4.4 Cu-Containing Laccase . . . . . . . . . . . . . . . . . 4.5 Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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5 Pectinase in Degradation of Lignocellulosic Wastes 5.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 The Pectic Substances . . . . . . . . . . . . . . . . . . . 5.2.1 Protopectin . . . . . . . . . . . . . . . . . . . . 5.2.2 Pectinic Acids . . . . . . . . . . . . . . . . . . 5.2.3 Pectin or Pectins . . . . . . . . . . . . . . . . 5.2.4 Pectic Acid . . . . . . . . . . . . . . . . . . . . 5.3 Microbial Pectinolytic Enzymes . . . . . . . . . . . . 5.3.1 Esterases . . . . . . . . . . . . . . . . . . . . . . 5.3.2 Depolymerases . . . . . . . . . . . . . . . . . . 5.3.3 Protopectinases . . . . . . . . . . . . . . . . .

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5.4

Occurrence of Pectinolytic Enzymes . . . . . . . . . . . 5.4.1 Esterases . . . . . . . . . . . . . . . . . . . . . . . . . 5.4.2 Depolymerases . . . . . . . . . . . . . . . . . . . . . 5.5 Physicochemical and Biological Properties . . . . . . . 5.5.1 Physicochemical and Biological Properties of Esterases . . . . . . . . . . . . . . . . . . . . . . . 5.5.2 Physicochemical and Biological Properties of Depolymerases . . . . . . . . . . . . . . . . . . . 5.6 Assay Methods of Pectinolytic Enzymes . . . . . . . . 5.6.1 Assay Methods for Esterases . . . . . . . . . . . 5.6.2 Assay Methods for Depolymerases . . . . . . 5.7 Production of Pectinases . . . . . . . . . . . . . . . . . . . . 5.7.1 Production of Bacterial Pectinases . . . . . . . 5.7.2 Production of Fungal Pectinases . . . . . . . . 5.8 Pectinases as First Protein Product Made in Leaves 5.9 Application of Pectinases in Deconstruction of Lignocellulosic Wastes . . . . . . . . . . . . . . . . . . . 5.9.1 Potential Applications of Pectinases . . . . . 5.10 Special Approaches to Lignocellulosic Wastes . . . . 5.11 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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80 83 83 83 85 86 88 89

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90 91 95 96 96

6 Lipase in Degradation of Lignocellulosic Wastes . . . . . . . 6.1 Introduction of Lipases Enzyme . . . . . . . . . . . . . . . . 6.2 Structure of Lipase . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.1 Three-Dimensional Structure of Lipases . . . . . 6.3 Classification of Lipases . . . . . . . . . . . . . . . . . . . . . . 6.3.1 Bacterial Lipases . . . . . . . . . . . . . . . . . . . . . 6.4 Reactions Catalyzed by Lipase Enzyme . . . . . . . . . . . 6.4.1 Acidolysis . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4.2 Trans-esterification . . . . . . . . . . . . . . . . . . . . 6.4.3 Esterification . . . . . . . . . . . . . . . . . . . . . . . . 6.4.4 Aminolysis . . . . . . . . . . . . . . . . . . . . . . . . . 6.4.5 Hydrolysis . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4.6 Alcoholysis . . . . . . . . . . . . . . . . . . . . . . . . . 6.5 Catalytic Mechanism of Lipases . . . . . . . . . . . . . . . . 6.6 Lipase-Producing Bacteria and Fungi . . . . . . . . . . . . . 6.6.1 Lipases Enzyme by Solid-State Fermentation . 6.7 Assay for Lipase Enzyme (Amara et al. 2009) . . . . . . 6.8 Applications of Lipases Enzyme . . . . . . . . . . . . . . . . 6.8.1 Lipases for the Food and Agro-industrial Applications . . . . . . . . . . . . . . . . . . . . . . . . . 6.8.2 Dairy Industries . . . . . . . . . . . . . . . . . . . . . . 6.8.3 Baking Industries . . . . . . . . . . . . . . . . . . . . .

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105 105 105 106 106 107 107 107 108 108 108 108 108 108 109 109 109 110

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Contents

6.8.4 6.8.5 6.8.6 References . .

xvii

Human Milk Fat Substitutes . . . . . . . . . . . . . . . . . . . . . 111 Egg-Processing Industries . . . . . . . . . . . . . . . . . . . . . . . 111 Edible Oil Production . . . . . . . . . . . . . . . . . . . . . . . . . . 111 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 112

About the Authors

Dr. Aparna B. Gunjal has completed her B.Sc. from Annasaheb Magar Mahavidyalaya, Hadapsar; M.Sc. from Modern College Arts, Commerce and Science College, Ganeshkhind; and Ph.D. in Environmental Sciences subject from Savitribai Phule Pune University, Pune, Maharashtra, India. Currently, she is working as Assistant Professor in the Department of Microbiology at Dr. D. Y. Patil, Arts, Commerce & Science College, Pimpri, Pune, Maharashtra, India. She has 4 years of teaching and 18 years of research experience. Her research areas of expertise are solid waste management; plant growth-promoting rhizobacteria; e-waste management; bioremediation, etc. Aparna has 57 publications to her credit which includes research papers, review articles, books, and book chapters. She has also presented her research work in many National and International Conferences where she has received six awards for the best paper and poster presentations, including H. Khorana Award for the best paper presentation in the National Symposium on Recent Developments in Environmental Science and Technology held at Manonmaniam Sundaranar University, Alwarkurichi. Besides this, she has also received (a) DST-SERB Travel Grant under the category Young Scientist to attend the “International Conference on Solid Waste 2015: Knowledge Transfer For Sustainable Resource Management” at Hongkong SAR, China, from May 19 to 23, 2015; (b) Biotech Research Society of India Travel Grant to attend the International Conference on “Emerging Trends in Biotechnology for Waste Conversion (ETBWC-2017)” at NEERI, xix

xx

About the Authors

Nagpur in October 2017; and (c) Indian Academy of Sciences Travel Grant to attend the 83rd Annual Meeting of the Academy at North-Eastern Hill University (NEHU), Shillong in November 2017. Aparna has also received (a) Pune Municipal Corporation Award for excellent work in Environmental Sciences Research by the hands of Mayor of Pune city on August 15, 2015, (b) “The Elsevier Foundation-TWAS Sustainability Visiting Expert Programme” in 2018 to visit Sri Lanka, and (c) “Young Researcher with Innovative Technology” award for the paper titled “Formulation of symbiotic chocolates” in the National Symposium on “Recent Trends in Modern Biology and Biotechnology 2019” organized by Dr. D.Y. Patil Biotechnology and Bioinformatics, Dr. D.Y. Patil Vidyapeeth, Pune, in 2019. She has also worked on composting aspect as a Senior Researcher Assistant at Hongkong Baptist University, Hongkong, from 16.1.19 to 25.4.19. She is a member of many Institutions and Societies, viz. Asian Plant Growth-Promoting Rhizobacteria (PGPR) Society, Hyderabad; Life Member of Biotech Research Society, India (BRSI), Life Member of Indian Women Scientist’s Association (IWSA), Navi Mumbai, and Member of University Women’s Association, Pune (UWA), Affiliated to Indian Federation of University Women’s Association and International Federation of University Women. Aparna is a reviewer for many journals, viz., International Journal of Research in Environmental Science and Technology; International Journal of Environmental Sciences; Journal of Solid Waste Technology and Management; Frontiers in Microbiology; Environmental Sustainability; Current World Environment and guest editor for Biotech Express India Magazine; and also editorial board member of Journal of Environmental Science and Technology. She has also reviewed nine research papers. She has also guided around 35 postgraduate students in Microbiology and Environmental Science subjects.

About the Authors

xxi

Dr. Neha N. Patil is working as Associate Professor and Head, Department of Microbiology at PDEA’s Annasaheb Magar Mahavidyalaya, Pune, Maharashtra, India. She has completed her Master’s degree in Microbiology. She achieved her Ph.D. degree from Savitribai Phule Pune University, Pune, Maharashtra, India, where she worked on “Antimicrobial potential and enhancement strategies using Microbiospora sp.” Her research areas of interest include bioremediation, waste management, agriculture, algal biotechnology, nanotechnology, and nanoremediation. She has received grants from funding agencies like University Grant Commission, New Delhi, India and BCUD, Savitribai Phule Pune University, Pune. She has received best teacher award from the Department of Education Municipal Corporation of Pune, Maharashtra. She has also received young researcher award in 2019. She has received awards for best paper presentation in National and International Conferences. She has published around 25 research articles in scientific journals. She is a life member of several organizations like Association of Microbiologist’s Society of India, Indian Women Scientists’ Association, and Biotech research Society, India. She is a reviewer of number of journals like Current Nutrition and Food Science, Journal of Food Science, Journal of Microbiology, Biotechnology and Food Sciences, SN Applied Sciences, and Nepal Journal of Science and Technology. She has guided many M.Sc. students for their dissertation work. Sonali S. Shinde is working as Assistant Professor at Annasaheb Kulkarni, Department of Biodiversity, MES Abasaheb Garware College, Pune, Maharashtra, India. Sonali completed her bachelor’s degree in Industrial Microbiology and Masters in Microbiology. She teaches courses in microbial diversity, molecular biology, environment laws and patents to postgraduate students. She has supervised five master students for dissertation work on aflatoxin degradation, phytochemical analysis, and spatiotemporal variation in Musa (banana) leaves. She was associated with several research projects in the Institute of Bioinformatics and Biotechnology (IBB), National Center for Cell Sciences (NCCS) and Council of Scientific and

xxii

About the Authors

Industrial Research-Unit for Research and Development of Information Products (CSIR-URDIP), Pune, Maharashtra, India. She has worked in experimental microbiology, systems biology, and procedures of patenting an invention. She has her expertise in molecular modeling, docking, molecular simulation, and metabolic network construction to find the drug target. She has presented her research work in various National and International Conferences and has also published research papers in International Journals of repute. Her research areas of interest are natural product chemistry and interaction studies involving microbial diversity.

Chapter 1

Introduction

1.1 Wastes Human and industrial activities generate huge amount of wastes. The wastes generated are municipal solid wastes, industrial wastes, agricultural wastes, hazardous wastes, and e-wastes. There is very poor management of the wastes, and this is serious issue which needs to be considered. The wastes generated have bad impact on the human health and environment. The wastes generated are also difficult to degrade. Developed countries generate more wastes as compared to developing countries (Kathiravale and Muhd Yunus 2008).

1.1.1 Lignocellulosic Wastes Nearly 200 billion tons of lignocellulosic biomass is produced annually worldwide. About 75% are cereal lignocellulose residues. Sugarcane is the highest amount of residues with major wastes as leaves, stalk, bagasse, pressmud, etc. The annual production of cereal straw is 2.9 billion tons, where China alone produces 700 million tons per year. In India, around 600 million tons of residues are produced which includes 480 tons of crop residues and 120 tons of processing-based residues. In USA, about 1.3 billion tons of biomass is produced per year, where agricultural and forest residues produced are 998 and 368 million tons, respectively (USDA 2005). In Europe, 120 million tons of lignocellulosic biomass is produced per year (Philippoussis 2009). In Brazil, the lignocellulosic wastes generated are 350 million tons (Pereira et al. 2008). Africa and Australia, due to desert areas, generate less lignocellulosic wastes which are 100 and 40 million tons per year (Philippoussis 2009). Lignocellulose is a generic term which describes the constituents in most plants, viz. cellulose, hemicelluloses, and lignin (Chandel et al. 2007; Siqueira and Filho

© Springer Nature Switzerland AG 2020 A. B. Gunjal et al., Enzymes in Degradation of the Lignocellulosic Wastes, https://doi.org/10.1007/978-3-030-44671-0_1

1

2

1 Introduction

2010; Zhang et al. 2012). In cellulose, molecules are arranged regularly and gathered into bundles; hemicellulose is an acetylated arabinoxylan with minor amounts of galactose and mannose, and lignin is a phenolic polymer intermeshed and chemically bonded by non-covalent forces and covalent cross-linkages (Abe et al. 2018). Lignocellulose comprises many polysaccharides, phenolic polymers, and proteins and is fibrous part of plant material. Lignocellulosic biomass has distinctive physical and chemical characteristics. The main components of lignocellulose wastes are shown in Table 1.1. Countries like Brazil and USA have promoted domestic bioethanol production. “First-generation bioethanol” is made from sugar feedstock such as cane juice (in Brazil) and molasses (in India) or from starch-rich materials such as corn (in USA). Table 1.1 Components of lignocellulose biomass Lignocellulose wastes

Cellulose (wt%)

Hemicellulose (wt%)

Lignin (wt%)

References

Barley straw

33.8

21.9

13.8

Nigam et al. (2009)

Corn cob

33.7

31.9

6.1

Nigam et al. (2009)

Corn stalks

35.0

16.8

7.0

Nigam et al. (2009)

Cotton stalks

58.5

14.4

21.5

Nigam et al. (2009)

Oat straw

39.4

27.1

17.5

Nigam et al. (2009)

Rye straw

37.6

30.5

19.0

Nigam et al. (2009)

Soya stalks

34.5

24.8

19.8

Nigam et al. (2009)

Sugarcane bagasse

40.0

27.0

10.0

Nigam et al. (2009)

Sunflower stalks

42.1

29.7

13.4

Nigam et al. (2009)

Wheat straw

49.7

14.8

23.5

Santos et al. (2015)

Bagasse

25.0–45.0

28.0–32.0

15.0–25.0

Putro et al. (2016)

Rice husk

25.0–35.0

18.0–21.0

26.0–31.0

Kabenge et al. (2018), Wikee et al. (2017)

Paddy straw

34.8–36.4

23.6–30.3

10.4–11.0

Syazwanee et al. (2018)

Banana waste

13.2

14.8

14.0

John et al. (2006)

Sorghum straw

35.1

24.0

25.4

Vazquez et al. (2007)

1.1 Wastes

3

Bioethanol production is expected to increase by more than 100 billion lit by 2022 (Goldemberg and Guardabasi 2009). The raw materials are unable to meet the increasing demand for fuels (Hahn-Hagerdal et al. 2006). This led to interest in “second-generation ethanol” from non-food lignocellulosic materials, viz. agricultural residues, wood, paper, and municipal solid waste, and energy crops. In India, the interest in biofuels has increased. The fossil fuels are now blended with biodiesel and bioethanol. The worldwide availability of major lignocellulosic wastes—wheat straw, rice straw, corn straw, and sugarcane bagasse—is 354.34, 731.30, 128.02, and 180.73, respectively (Saini et al. 2015).

1.2 Lignocellulosic Wastes and Problems in Their Degradation Cellulose is major component of cell walls of plants. The repeating unit of the cellulose chain is cellobiose. It consists of intramolecular and intermolecular hydrogen bonds, which help to bind glucose units. Cellulose molecule exists as bundles which aggregate in the microfibrils order, i.e., crystalline and amorphous. After cellulose, hemicellulose is the next abundant polymer. Hemicellulose is amorphous in nature. Hemicellulose consists of many heteropolymers which include xylan, galactomannan, glucuronoxylan, arabinoxylan, glucomannan, and xyloglucan. Hardwood hemicelluloses contain mostly xylans, and softwood hemicelluloses contain mostly glucomannans. The heteropolymers of hemicellulose are composed of monosaccharide units. Hemicelluloses link cellulose fibers into microfibrils and also help cross-linking with lignin (Agbor et al. 2011; Scheller and Ulvskov 2010). Lignin is a three-dimensional polymer of phenylpropanoid units. The phenylpropane is the main block of lignin. Lignin gives strength to the plant tissues and individual fibers, and stiffness to the cell wall (Rubin 2008). The oxidative coupling of phenylpropane building blocks, viz. p-coumaryl, coniferyl, and sinapyl alcohols, makes the structure of lignin. Generally, lignocellulosic biomass comprises 35–50% cellulose, 20–35% hemicellulose, and 10–25% lignin, and the remaining are proteins, oils, and ash (Saha 2005). An enzymatic synergism is essential for hydrolysis of lignocellulose. The major lignin-degrading enzymes are heme-containing and H2 O2 -dependent lignin peroxidase (LiP), manganese peroxidase (MnP), and Cu-containing laccase (Hamid and Ur-Rehman 2009; Minussi et al. 2002; Sena-Martins et al. 2008). Laccases oxidize various substrates and at the same time reduces molecular oxygen to water; LiP oxidizes reducing substrates, and MnP oxidizes phenolic structures to phenoxyl radicals (Maciel et al. 2010). Lignocellulose-degrading enzymes are produced by many fungi.

4

1 Introduction

Table 1.2 Intra- and interpolymer linkages that form the individual polymers—lignin, cellulose, and hemicellulose—and between the polymers to form lignocellulose Intrapolymer linkages Ether bond

Lignin, (hemi) cellulose

Carbon to carbon

Lignin

Hydrogen

Cellulose

Ester bond

Hemicellulose

Interpolymer linkages Ether bond

Cellulose–lignin Hemicellulose–lignin

Ester bond

Hemicellulose–lignin

Hydrogen bond

Cellulose–hemicellulose Hemicellulose–lignin Cellulose–lignin

Lignocellulose wastes are hard to degrade. It is difficult in dissolving lignin without destroying it and some of its subunits. Lignin has three aromatic alcohols, viz. coniferyl alcohol, sinapyl and p-coumaryl, phenolic acids such as pcoumaric and ferulic acid. Lignin is further linked to both hemicelluloses and cellulose, and therefore, lignin is very hard to degrade. Cellulose also cannot easily undergo hydrolysis because of highly ordered crystalline structure (Pothiraj et al. 2006). The lignocellulolytic wastes are degraded by variety of bacteria and fungi (Kausar et al. 2010). There are four main types of bonds identified in the lignocellulose complex. Those are ether type of bonds, ester bonds, carbon-to-carbon bonds, and hydrogen bonds. These four bonds are the main types of bonds that provide linkages within the individual components of lignocellulose (intrapolymer linkages) and connect the different components to form the complex (interpolymer linkages). The position and bonding function of the latter linkages are summarized in Table 1.2 (Faulon et al. 1994).

1.3 Linkages Between Lignocellulose Components The ether bonds (70%) and carbon-to-carbon bonds (30%) connect building molecules within lignin polymer (Table 1.2). The ether bonds appear between allylic and aryl carbon atoms, or between aryl and aryl carbon atoms, or even between two allylic carbon atoms. The intra- and interpolymer linkages that form the individual polymers—lignin, cellulose, and hemicellulose—and between the polymers to form lignocellulose are shown in Table 1.2.

1.4 Value-Added Products from Lignocellulosic Wastes

5

Animal feed Pulp and paper

Lignocellulosic wastes

Composites

Enzymes Biofuels

Fig. 1.1 Value-added products from lignocellulosic wastes

1.4 Value-Added Products from Lignocellulosic Wastes The different value-added products from lignocellulosic wastes are shown in Fig. 1.1.

1.5 Types of Lignocellulosic Biomass 1.5.1 Dry Grass Grasses are used for the production of lignocellulosic biomass. The major herbaceous crops used are switchgrass (Panicum virgatum), Miscanthus (Miscanthus spp. Anderss), canary grass (Phalaris arundinacea), giant reed (Arundo donax L.), alfalfa (Medicago sativa L.), and Napier grass (Pennisetum purpureum). These grasses have environmental advantages over food crops (Faraco 2013). Switchgrass is particularly used as it is eco-friendly, available in large amount, resists diseases, and can give high yield of the product (Limayem and Ricke 2012). Miscanthus is also used as a combustible energy crop (Faraco 2013).

6

1 Introduction

1.5.2 Forest Woody Feedstock Lignocellulosic waste from forests includes residues from forests, sawdust from sawmills, woodchips and branches from dead trees, and fast-growing short-rotation forest trees.

1.5.3 Municipal Solid Wastes Municipal and industrial wastes contain lignocellulose material. The solid wastes include kitchen, garden, paper, and cardboard wastes.

1.5.4 Agricultural Residues Agro-industrial wastes are generated during processing of agricultural products. These include rice, wheat straw, corn cob, ground nut shells, and rice stalk. The agricultural residues consist mainly of cellulose, hemicellulose, and lignin (Nigam and Pandey 2009).

1.6 Bioprocessing of Lignocellulose Wastes Various technologies are available for conversion of lignocelluloses to ethanol and other products. Bioconversions of lignocellulosic materials to value-added products include pretreatment; hydrolysis of the polymers to sugars; bioutilization of these molecules to get chemical products; and separation and purification. It has been reported that the solid-state fermentation (SSF) is good alternative for different value-added products (e.g., enzymes) using lignocellulosic materials (Jecu 2000). The conditions for the growth of microorganisms under SSF are closer to their natural habitats which make it suitable for the production of enzymes and many other metabolites. SSF has more advantages in comparison to submerged fermentation.

1.7 Advantages of Enzymes in Degradation of Lignocellulose Wastes Enzymes have good specific activity. Higher stability, i.e., extended life-time; Enzymes can be reused.

1.7 Advantages of Enzymes in Degradation of Lignocellulose Wastes

7

Good flexibility for process configuration; Enzymes carry process with improved integration. Increased dry matter content because of less viscosity at high temperature.

1.7.1 Enzymes The microbial enzymes have uses in various industries, viz. food, agriculture, chemicals, medicine, and energy. Enzymes have gained wide importance because of reduced process time, require less energy input, cheap, non-toxic, and are ecofriendly (Choi et al. 2015; Li et al. 2012). Enzymes are biological molecules, proteinaceous in nature, and act as catalysts. The enzymes have biodegradation ability because they are eco-friendly (Choi et al. 2015; Illanes et al. 2012; Mojsov 2011). Enzymes are composed of amino acids with molecular mass ranging from kilodalton to megadalton. This specificity of enzyme to catalyze reactions between one types of chemical compound over the other provides the basis of its classification and name. Microbial enzymes are available in abundance, eco-friendly, and can work under varied physicochemical conditions. The market for industrial enzymes has surpassed the USD 7.1 billion, and five-year compound annual growth rate (CAGR) is around 8.2%. The market for food enzymes will rise and reach USD 2.94 billion by 2021. The maximum growth rate will be of detergent enzymes (CAGR of 11.3% in 2021) (Rigoldi et al. 2018).

1.7.1.1

Classification of Enzymes

Oxido Reductases The oxido reductases enzyme catalyzes oxidation and reduction reactions. The substrate that is oxidized is regarded as hydrogen donor. The systematic name is based on donor:acceptor oxidoreductase. The second figure in the code number of the oxidoreductases is the group in the hydrogen donor that undergoes oxidation, and the third figure is the acceptor involved.

Transferases Transferase enzymes transfer a methyl or a glycosyl group to other compounds. Mostly, the donor is a cofactor charged with the group to be transferred. The reaction involves oxidative deamination of amino acid linked with reductive amination of oxoacid.

8

1 Introduction

Hydrolases These enzymes cleave C–O, C–N, C–C, and phosphoric anhydride bonds by hydrolysis. The hydrolase enzymes transfer the particular group from their substrates to suitable acceptor.

Lyases Lyase enzyme splits C–C, C–O, C–N, and other bonds by removal, leaving double bonds or rings, or adding groups to double bonds. The pyridoxal phosphate, which is subclass of lyases, catalyzes removal of a β- or γ-substituent from α-amino acid and replaces it with other group.

Isomerases These enzymes catalyze geometric or structural changes within single molecule. Depending on the type of isomerism, they are called racemases, epimerases, cistrans-isomerases, isomerases, tautomerases, mutases, or cycloisomerases.

Ligases Ligase enzymes catalyze and join together two molecules along with the hydrolysis of a diphosphate bond in ATP.

1.7.2 Enzymes in Lignocellulose Degradation 1.7.2.1

Cellulases

Cellulases hydrolyze the glycoside bond present between the glucose residues in cellulose. Cellulose can be hydrolyzed by β-1, 4-endoglucanases, exoglucanases or 1, 4-β-cellobiosidase, and β-glucosidase.

1.7.2.2

Hemicellulases

Hemicellulases, e.g., xylanase, hydrolyze the xylan. Hemicellulases are extensively studied and degrade lignocellulose wastes.

1.7 Advantages of Enzymes in Degradation of Lignocellulose Wastes

1.7.2.3

9

Lignin Peroxidases

Lignin peroxidases are the heme glycoproteins and are important in lignin degradation. This enzyme breaks C–C bonds and oxidizes benzyl alcohols to aldehydes or ketones (Piontek et al. 2001). Lignin peroxidases act on phenolic and non-phenolic lignin substrates (Wong 2009). Mostly, fungi belonging to basidiomycetes have the ability to produce lignin peroxidases (Abbas et al. 2005; Wong 2009). The fungi producing lignin peroxidase enzyme are P. chrysosporium, T. versicolor, Pleurotus ostreatus (Sivakami et al. 2012), Panus sp., P. coccineus, Perenniporia medullapanis, and P. sanguineus (Pointing et al. 2005).

1.7.2.4

Manganese Peroxidases

Manganese peroxidase degrades the lignin mainly by attacking phenolic lignin component (Asgher et al. 2008). In the presence of H2 O2 , manganese peroxidase oxidizes the phenolic structures by converting Mn2+ to Mn3+ . Oxalate and malonate are the mediators that produce carbon-centered radicals, peroxyl radicals, and superoxide radicals and improve the effective lignin-degrading system (Asgher et al. 2008; Wong 2009). Manganese peroxidase is an essential factor to basidiomycetes and wood-decaying white-rot fungi. The manganese peroxidase enzyme producers are Agaricus bisporus (Lankinen et al. 2001), Lenzites betulinus (Hoshino et al. 2002), Panus tigrinus (Lisov et al. 2003), Nematoloma frowardii (Hilden et al. 2008), P. chrysosporium (Jarvinen et al. 2012), and Mucor racemosus (Bonugli-Santos et al. 2010).

1.7.2.5

Pectinases

The pectinolytic enzymes produced by both plants and microorganisms degrade the pectins. Pectin degradation involves depolymerization and de-esterification reactions. The fungi which produce pectinases enzyme are Aspergillus, Penicillium, Colletotrichum, Sclerotina, Fusarium, Trichoderma, Verticulum, Sclerotium, Geotrichum (Maldonado et al. 2002; Teixeira et al. 2000).

1.7.2.6

Laccases

Laccases are the copper-containing polyphenol oxidases. These oxidize phenolic compounds to phenoxy radicals which lead to cleavage of aryl-C. The laccaseproducing fungi are Trichoderma harzianum (Savoie et al. 2001), Trichoderma atroviride (Holker et al. 2002), Trichoderma longibrachiatum (Velazquez-Cedeiio et al., 2004), Trametes versicolor (Han et al. 2005), Lentinus tigrinus (Ferraroni et al. 2007), Trametes pubescens (Shleev et al. 2007), Cyathus bulleri (Salony et al. 2006), Paecilomyces sp. (Liang et al. 2007), P. chrysosporium (Viswanath et al. 2008), Lentines

10

1 Introduction

edodes (Shanmugam et al. 2009), Pleurotus ostreatus (Patel et al. 2009; Sivakami et al. 2012), Ganoderma lucidum (Li et al. 2001), Alternaria tenuissima (Abd El et al. 2015), and Trichoderma sp. (Brijwani and Vadlani 2011).

1.8 Fungi-Degrading Lignocellulosic Wastes 1.8.1 Wood-Decaying Fungi Fungi degrade lignocellulosic substrates very effectively. Fungi can be differentiated into different classes based on their spore structures, viz. ascomycetes, basidiomycetes, and deuteromycetes. The wood-decaying fungi use both enzymatic and non-enzymatic systems for the degradation of wood. The wood loses weight, strength, and density due to the action of these fungi. The fungi are divided into three groups, brown-rot, white-rot, and soft-rot fungi (Schwarze 2007).

1.8.2 Brown-Rot Fungi The brown-rot fungi reduce the strength of wood by 75% due to decomposition of cellulose and hemicellulose (Yelle et al. 2008). This turns wood into powder and is characterized by red brown in color. Poria incrassata, brown-rot fungi, has typical rhizomorphs to transport water from the soil to the wood, and the wood is further decayed by fungi. The examples of wood-decaying brown-rot fungi are Gloeophyllum trabeum, Fomitopsis lilacino-gilva, Laetiporus portentosus, Postia placenta, and Serpula lacrymans (Martinez et al. 2005; Wong 2009).

1.8.3 White-Rot Fungi The white-rot fungi produce ligninolytic enzymes, viz. laccase and lignin peroxidase, and manganese peroxidase produces H2 O2 essential for peroxidase activities. These enzymes break lignin in a synergistic manner. The examples of white-rot fungi are Ceriporiopsis subvermispora and Phlebia radiate (Fackler et al. 2007).

1.8.4 Soft-Rot Fungi Soft-rot fungi form cavity in the secondary walls of the wood cells. Soft-rot fungi degrade cellulose and hemicellulose.

References

11

References Abbas A, Koc H, Liu F, Tien M (2005) Fungal degradation of wood: initial proteomic analysis of extracellular proteins of Phanerochaete chrysosporium grown on oak substrate. Curr Genet 47:49–56 Abd El A, El-Shamy A, Atalla S, El-Diwany A, Hamed E (2015) Screening of fungal isolates for laccase enzyme production from marine sources. Res J Pharma Biol Chem Sci 6:221–228 Abe T, Lajide L, Owolabi B, Adebayo A (2018) Physico-chemical composition of lignocellulose biomass from Gliricidia sepium and Cola gigantea. Am J Innov Res Appl Sci 6:131–140 Agbor V, Cicek N, Sparling R, Berlin A, Levin D (2011) Biomass pretreatment: fundamentals toward application. Biotechnol Adv 29:675–685 Asgher M, Bhatti H, Ashraf M, Legge R (2008) Recent developments in biodegradation of industrial pollutants by white rot fungi and their enzyme system. Biodegradation 19:771–783 Bonugli-Santos R, Durrant L, DaSilva M, Sette L (2010) Production of laccase, manganese peroxidase and lignin peroxidase by Brazilian marine-derived fungi. Enzyme Microbial Technol 46:32–37 Brijwani K, Vadlani P (2011) Cellulolytic enzymes production via solid-state fermentation: effect of pretreatment methods on physico-chemical characteristics of substrate. Enzyme Res 2:120–128 Chandel A, Chan E, Rudravaram1 R, Narasu M, Rao L, Ravindra P (2007) Economics and environmental impact of bioethanol production technologies: an appraisal. Biotechnol Mol Biol Rev 2:14–32 Choi J, Han S, Kim H (2015) Industrial applications of enzyme biocatalysis: current status and future aspect. Biotechnol Adv 33:1443–1454 Fackler K, Schwanninger M, Gradinger C, Hinterstoisser B, Messner K (2007) Qualitative and quantitative changes of beech wood degraded by wood rotting basidiomycetes monitored by Fourier transform infrared spectroscopic methods and multivariate data analysis. FEMS Microbiol Lett 271:162–169 Faraco V (2013) Lignocellulose conversion: enzymatic and microbial tasks for bioethanol production. Springer, Berlin, pp 24–26 Faulon J, Carlson G, Hatcher P (1994) A three-dimensional model for lignocellulose from gymnospermous wood. Org Geochem 21:1169–1179 Ferraroni M, Myasoedova NM, Schmatchenko V, Leontievsky AA, Golovleva LA, Scozzafava A, Briganti F (2007) Crystal structure of a blue laccase from Lentinus tigrinus: evidences for intermediates in the molecular oxygen reductive splitting by multicopper oxidases. BMC Struct Biol 7:60 Goldemberg J, Guardabasi P (2009) Are biofuels a feasible option? Energy Policy 37:10–14 Hahn-Hagerdal B, Galbe M, Gorwa-Grauslund MF, Liden G, Zacchi G (2006) Bio-ethanol: the fuel of tomorrow from the residues of today. Trend Biotechnol 24:549–556 Hamid M, Ur-Rehman K (2009) Potential applications of peroxidases. Food Chem 115:1177–1186 Han M, Choi H, Song H (2005) Purification and characterization of laccase from the white rot fungus Trametes versicolor. J Microbiol 43:555–560 Hilden K, Bortfeldt R, Hofrichter M, Hatakka A, Lundell TK (2008) Molecular characterization of the basidiomycete isolate Nematoloma frowardii b19 and its manganese peroxidase places the fungus in the corticioid genus Phlebia. Microbiology 154:2371–2379 Holker U, Dohse J, Hofer M (2002) Extracellular laccases in ascomycetes Trichoderma atroviride and Trichoderma harzianum. Folia Microbiol 47:423–437 Hoshino F, Kajino T, Sugiyama H, Asami O, Takahashi H (2002) Thermally stable and hydrogen peroxide tolerant manganese peroxidase (MnP) from Lenzites betulinus. FEBS Lett 530:249–252 Illanes A, Cauerhff A, Wilson L et al (2012) Recent trends in biocatalysis engineering. Bioresour Technol 115:48–57 Jarvinen J, Taskila S, Isomäki R, Ojamo H (2012) Screening of white-rot fungi manganese peroxidases: a comparison between the specific activities of the enzyme from different native producers. AMB Express 2:1–9

12

1 Introduction

Jecu L (2000) Solid-state fermentation of agricultural wastes for endoglucanase production. Ind Crops Prod 11:1–5 John F, Monsalve G, Medina de Perez V, Ruiz Colorado A (2006) Ethanol production of banana shell and cassava starch. Dyna Rev Fac Nac Minas 73:21–27 Kabenge I, Omulo G, Banadda N, Seay J, Zziwa A, Kiggundu N (2018) Characterization of banana peels wastes as potential slow pyrolysis feedstock. J Sust Dev 11:14–24 Kathiravale S, Muhd Yunus M (2008) Waste to wealth. Asia Europe J 6:359–371 Kausar H, Sariah M, Saud H, Alam M, Ismail M (2010) Development of compatible lignocellulolytic fungal consortium for rapid composting of rice straw. Int Biodeterior Biodegradation 64:594–600 Lankinen V, Bonnen A, Anton L, Wood D, Kalkkinen N, Hatakka A, Thurston C (2001) Characteristics and N-terminal amino acid sequence of manganese peroxidase from solid substrate cultures of Agaricus bisporus. Appl Microbiol Biotechnol 55:170–176 Li P, Wang H, Liu G, Li X, Yao J (2001) The effect of carbon source succession on laccase activity in the co-culture process of Ganoderma lucidum and a yeast. Enzyme Microbial Technol 48:1–6 Li S, Yang X, Yang S et al (2012) Technology prospecting on enzymes: application, marketing and engineering. Comput Struct Biotechnol J 2:1–11 Liang Z, Han Y, Chu H (2007) A new thermo tolerant Paecilomyces species which produces laccase and a biform sporogenous structure. Fungal Diversity 27:95–102 Limayem A, Ricke S (2012) Lignocellulose biomass for bioethanol production: current perspectives, potential issues and future prospects. Prog Energy Combust Sci 38:449–467 Lisov A, Leontievsky A, Golovleva L (2003) Hybrid Mn-peroxidase from the ligninolytic fungus Panus tigrinus. Isolation, substrate specificity, and catalytic cycle. Biochemistry 68:1027–1035 Maciel M, Silva A, Ribeiro H (2010) Industrial and biotechnological applications of ligninolytic enzymes of basidiomycota: a review. Electron J Biotechnol 13:1–12 Maldonado M, Cáceres S, Galli E, Navarro A (2002) Regulation of the production of polygalacturonase by Aspergillus niger. Folia Microbiol 47:409–412 Martinez AT, Speranza M, Ruiz-Dueñas F et al (2005) Biodegradation of lignocellulosics: microbiological, chemical and enzymatic aspects of fungal attack to lignin. Int Microbiol 8:195–204 Minussi R, Pastore G, Duran N (2002) Potential applications of laccase in the food industry. Trends Food Sci Technol 13:205–216 Mojsov K (2011) Application of enzymes in the textile industry: a review. In: II International Congress engineering, ecology and materials in the processing industry, Jahorina, pp 230–238 Nigam P, Pandey A (2009) Production of organic acids from agro-industrial residues. Biotechnology for agro-industrial residues utilisation. Springer, Berlin, pp 37–38 Nigam PS, Gupta N, Anthwal A (2009) Pre-treatment of agro-industrial residues. In: Nigam P, Pandey A (eds) Biotechnology for agro-industrial residues utilization, 1st edn. Springer, Netherlands, pp 13–33 Patel H, Akshaya G, Shilpa G (2009) Effect of different culture conditions and inducers on production of laccase by a basidiomycete fungal isolate Pleurotus Ostreatus HP-1 under solid state fermentation. BioResour 4:268–284 Pereira N Jr, Couto M, Anna S (2008) Biomass of lignocellulosic composition for fuel ethanol production within the context of biorefinery. Rio de Janeiro: Escola de Quimica/UFRJ, Series on biotechnology, vol 2, pp 8–47 Philippoussis A (2009) Production of mushrooms using agro-industrial residues as substrates. In: Nigam P, Pandey A (eds) Biotechnology for agro-industrial residues utilization-utilization of agro-residues. Springer, Germany, pp 163–196 Piontek K, Smith A, Blodig W (2001) Lignin peroxidase structure and function. Biochem Soc Trans 29:111–116 Pointing S, Pelling A, Smith G, Hyde K, Reddy C (2005) Screening of basidiomycetes and xylariaceous fungi for lignin peroxidase and laccase gene-specific sequences. Mycol Res 109:115–124

References

13

Pothiraj C, Kanmani P, Balaji P (2006) Bioconversion of lignocellulose materials. Mycobiology 34:159–165 Putro J, Soetaredjo F, Lin S, Ju Y, Ismadji S (2016) Pretreatment and conversion of lignocellulose biomass into valuable chemicals. RSC Adv 6:46834–46852 Rigoldi F, Donini S, Redaelli A, Parisini E (2018) Review: engineering of thermostable enzymes for industrial applications. Appl Bioeng 2:1–17 Rubin E (2008) Genomics of cellulosic biofuels. Nature 454:841–845 Saha B (2005) Enzymes as biocatalysts for conversion of lignocellulosic biomass to fermentable sugars. In: Hou C (ed) Handbook of industrial biocatalysis. CRC Press, Boca Raton Saini J, Saini R, Tewari L (2015) Lignocellulosic agriculture wastes as biomass feedstocks for second-generation bioethanol production: concepts and recent developments. 3 Biotech 5:337– 353 Salony, Mishra S, Bisaria V (2006) Production and characterization of laccase from Cyathus bulleri and its use in decolourization of recalcitrant textile dyes. Appl Microbiol Biotechnol 71:646–653 Santos J, Sampedro R, Fillat U, Oliva JM et al (2015) Evaluating lignin-rich residues from biochemical ethanol production of wheat straw and olive tree pruning by FTIR and 2D-NMR. Int J Polym Sci 2015:1–11 Savoie J, Mata G, Mamoun M (2001) Variability in brown line formation and extracellular laccase production during interaction between white-rot basidiomycetes and Trichoderma harzianum biotype Th2. Mycologia 93:243–248 Scheller H, Ulvskov P (2010) Hemicelluloses. Annu Rev Plant Biol 61:263–289 Schwarze F (2007) Wood decay under the microscope. Fungal Biol Rev 21:133–170 Sena-Martins G, Almeida-Vara E, Duarte J (2008) Eco-friendly new products from enzymatically modified industrial lignins. Ind Crop Prod 27:189–195 Shanmugam S, Rajasekaran P, Joseph V (2009) Synthetic dye decolourization, textile dye and paper industrial effluent treatment using white rot fungi Lentines edodes. J Desalin Water Treat 4:143–147 Shleev S, Nikitina O, Christenson A, Reimann CT, Yaropolov A, Ruzgas T, Gorton L (2007) Characterization of two new multi forms of Trametes pubescens laccase. Bioorg Chem 35:35–49 Siqueira F, Filho E (2010) Plant cell wall as a substrate for the production of enzymes with industrial applications. Mini-Rev Org Chem 7:54–60 Sivakami V, Ramachandran B, Srivathsan J, Kesavaperumal G, Benila D, Kumar M (2012) Production and optimization of laccase and lignin peroxidase by newly isolated Pleurotus ostreatus LIG 19. J Microbiol Biotechnol Res 2:875–881 Syazwanee M, Shaziera A, Izzati M, Azwady A, Muskhazli A (2018) Improvement of delignification, desilication and cellulosic content availability in paddy straw via physico-chemical pretreatments. Annu Res Rev Biol 26:1–11 Teixeira M, Filho L, Duran N (2000) Carbon sources effect on pectinase production from Aspergillus japonicus 586. Braz J Microbiol 31:286–290 USDA-US DOE (2005) Biomass as feedstock for a bioenergy and bioproducts industry: the technical feasibility of a billion-ton annual supply Vazquez M, Oliva M, Téllez-Luis S, Ramírez J (2007) Hydrolysis of sorghum straw using phosphoric acid: evaluation of furfural production. Bioresour Technol 98:3053–3060 Velazquez-Cedeiio M, Farnet A, Ferre E (2004) Variations of lignocellulosic activities in dual cultures of Pleurotus ostreatus and Trichoderma longibrachiatum on unsterilized wheat straw. Mycologia 96:712–719 Viswanath B, Chandra S, Pallavi H, Reddy R (2008) Screening and assessment of laccase producing fungi isolated from different environmental samples. Afr J Biotechnol 7:1129–1133 Wikee S, Chumnunti P, Kanghae A, Chukeatirote A, Lumyong S, Faulds C (2017) Lignocellulolytic capability of endophytic Phyllosticta sp. J Bacteriol Mycol 4:1–7 Wong D (2009) Structure and action mechanism of ligninolytic enzymes. Appl Biochem Biotechnol 157:174–209

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Yelle D, Ralph J, Lu F, Hammel K (2008) Evidence for cleavage of lignin by a brown rot basidiomycete. Environ Microbiol 10:1844–1849 Zhang Z, Donaldson A, Ma X (2012) Advancements and future directions in enzyme technology for biomass conversion. Biotechnol Adv 30:913–919

Chapter 2

Cellulase in Degradation of Lignocellulosic Wastes

Abstract Cellulose is the major component in the cell wall of many plants. Degradation of cellulose is very difficult and needs to be solved. Cellulase is the enzyme in degradation of cellulose. The chapter here describes in detail the structure and properties of cellulose; cellulase production by solid-state and submerged state fermentation; cellulolytic microorganisms from extreme environments; enzymes involved in cellulose breakdown; and cellulase systems of bacteria and fungi. The chapter also focuses on microbial sources of cellulase enzymes; cellulosomes to degrade cellulose; determination of cellulase activity; pretreatment of cellulose; mechanisms of cellulose hydrolysis; genes related to cellulose degradation; and applications of cellulase enzyme. The chapter gives detailed information of cellulase enzyme in degradation of cellulose. This will help in the proper management of cellulosic wastes. Keyword Cellulase · Fermentation · Cell wall · Degradation · Lignocellulosic wastes

2.1 Cellulose Plants produce 180 billion tons of cellulose manufacture annually and are the most abundant polymer. Cellulose is the structural part of the primary cell wall of many green plants. The main occurrence of cellulose is the lignocellulosic substance. Cellulose-containing resources include agriculture residues, water plants, grasses, and other plant materials. The cellulose in cotton, flax, and chemical pulp is 90% and in wood 50% (Aitken 2012; Limayem and Ricke 2012). The composition of cellulose in different lignocellulosic raw materials and wastes is represented in Table 2.1. In Asia, 4.4 billion tons of solid and 790 million tons of municipal solid wastes are generated (Yoshizawa et al. 2004). Agricultural resources contribute 350 MT of wastes per year (Asokan et al. 2007). Cellulose is micrometer-sized particle comprised of microfibrils. Cellulose particles consist of crystalline and amorphous structures (Yang et al. 2011).

© Springer Nature Switzerland AG 2020 A. B. Gunjal et al., Enzymes in Degradation of the Lignocellulosic Wastes, https://doi.org/10.1007/978-3-030-44671-0_2

15

16 Table 2.1 Cellulose in different lignocellulosic raw materials and wastes

2 Cellulase in Degradation of Lignocellulosic Wastes Lignocellulosic raw materials and wastes

Cellulose (%)

Hardwood stems

40–55

Softwood stems

45–50

Rice straw

35–45

Wheat straw

38–45

Tobacco chops

22–30

Arundo donax

30–38

Miscanthus

35–40

Newspaper

40–55

2.2 Microcrystalline Cellulose Microcrystalline cellulose (MCC) market size will increase by 2024, growing at more than 7% compound annual growth rate. Growing demand in pharmaceuticals, cosmetic, and food industries is the key factor (Shokri and Adibkia 2013). MCC is generally used as a texturizer and emulsifier in pharmaceuticals. In food industries, it is used as anticaking agent and in dairy products and baked foods. In cosmetics, it is used as a thickening agent to give texture. The global personal care market share is predicted to surpass USD 17 billion by 2024, which is expected to favor MCC market size growth.

2.3 Cellulose Structure, Function, and Properties Cellulose is a fiber and long chain of linear polysaccharide with many d-glucose units linked by β-1,4-glycosidic bonds (Brown 1996). The acetal linkage is beta (β). Wood, paper, cotton, etc., contain fibrous cellulose. It is also present in plants as microfibrils with diameter 2–20 nm and length 100–40,000 nm. Cellulose is most abundant in nature. The individual chains adhere to each other by hydrogen bonding and van der Waals forces, and due to this cellulose is crystalline in structure. The structure of cellulose is represented in Fig. 2.1. Fig. 2.1 Structure of cellulose

2.3 Cellulose Structure, Function, and Properties

17

Cellulose can transform into different crystalline forms, viz., cellulose I, II, III, and IV (Kontturi 2005). The interactions are intrachain O–H–O bonds between glucose residues of the same chain and sheet; interchain O–H–O bonds between glucose residues of the same sheet but different chains; and intersheet C–H–O bonds in terms of van der Waals interactions. Cellulosic microfibrils and nanofibrils are twisted. Increase in the temperature or dehydration makes cellulose more aggregate and recalcitrant in nature (Taherzadeh and Jeihanipour 2012). The intermolecular hydrogen bonds give cellulose complex organization. Natural cellulose is cellulose I, which is crystalline microfibril with parallel chains. Two phases Iα and Iβ coexist in native cellulose I. Cellulose in regenerated cellulose fibers is cellulose II which is stable allomorph. Cellulose is tasteless, odorless, hydrophilic with the contact angle of 20–30, and insoluble in water and solvents. Under particular conditions, cellulose reacts with water. Cellulose can be broken to glucose units by acid or alkali treatment. The properties of cellulose depend on the degree of polymerization. Cellulose chain lengths in wood pulp are between 300 and 1700 units; while plant fibers and bacterial cellulose have chain lengths from 800 to 10,000 units (Klemm et al. 2005). Microbial cellulose has long chains in comparison with plant-derived cellulose. Cellulose has low flexibility due to polar molecules and intermolecular forces.

2.4 Cellulase Production The production of cellulase enzyme is by submerged fermentation using the fungi. The fungi used for cellulase production are, viz., Trichoderma reesei, T. longibrachiatum, and Aspergillus niger (Singhania et al. 2009). Different lignocellulosic wastes such as wheat bran, corn cob, and rice and wheat straw as substrates are used for cellulolytic enzyme production in both solid-state and submerged fermentations (Deswal et al. 2011; Dogaris et al. 2009; Kang et al. 2004). The cellulase production by solid-state fermentation (SSF) is cheaper in comparison with submerged fermentation (SmF). Fermentation is used for cellulase production, and T. reesei is commonly used in bioprocessing for cellulase production. There is a report on Aspergillus niger (Oyeleke et al. 2012) and Alternaria alternata (Goshadrou et al. 2011) using corn cob as substrate; Aspergillus oryzae (Brijwani and Vadlani 2011) and Trichoderma reesei (Herculano et al. 2011) with soybean hulls as substrate; and Aspergillus japonicus URM5620 using castor bean as substrate for cellulase production by solid-state fermentation (Herculano et al. 2011). Also, there is a report on bagasse powder as a substrate for cellulase production using thermostable yeast (Rai et al. 2012). The fungi including Trichoderma, Penicillium, Botrytis neurospora (Aro et al. 2005), Aspergillus niger and Aspergillus terreus, Rhizopus stolonifer (Pothiraj et al. 2006), Fusarium oxysporum (Heck et al. 2002; Ramanathan et al. 2010) are used for bioprocessing. Also, there is report on Aspergillus terreus for cellulase production with rice straw as a substrate under solid-state fermentation. The filter paper and the approved

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2 Cellulase in Degradation of Lignocellulosic Wastes

Table 2.2 Substrates used and fermentation type for cellulase production Bacteria

Substrates

Fermentation

References

Cellulomonas cellulans

Paddy straw

SmF

Mishra et al. (2007)

Bacillus coagulans

Palm fruit residue

SSF

Odeniyi et al. (2009)

Bacillus cereus

Palm kernel cake

SSF

Lah et al. (2012)

Bacillus sp.

Agricultural residue

SSF

Nizamudeen and Bajaj (2009)

Clostridium thermocellum

Paper pulp

SmF and SSF

Zhuang et al. (2007)

Cellulosimicrobium cellulans

Rice husk and rice straw

SSF

Lo et al. (2009)

activity were 9.73 and 10.96 U/g, respectively (Narra et al. 2012). Eichhornia crassipes (water hyacinth) has been reported for cellulase production, where the maximum cellulase activity was at 40 °C temperature and pH 5.0 (Sachin et al. 2011). The solid-state fermentation is used for endocellulase production using Aspergillus japonicus C03 (Facchini et al. 2011). Sorghum straw can also be used as a substrate for cellulase production (Mohite and Magar 2010). The cellulolytic enzymes are used synergistically. In endo–exo-energy model, endoglucanases act on random sites of the lignocellulosic chains and make sites available for cellobiohydrolases to attack. The amount of crystalline cellulose digested by cellulase is the cellulase activity (Pandey et al. 2001). The substrates used and fermentation type for cellulase production are shown in Table 2.2.

2.4.1 Microorganisms Used for Cellulase Production The common bacteria used in cellulase production are Bacillus (Mawadza et al. 2000), Streptomyces, Ruminococcus, Clostridium (Lopez-Contreras et al. 2004), Pseudomonas fluorescens (Sethi et al. 2013), Rhodothermus marinus, Acidothermus sp., Bacillus subtilis CEL PTK1 (Bai et al. 2012), Bacillus subtilis AS3 (Deka et al. 2011). The actinobacteria used in cellulase production are Cellulomonas fimi, C. bioazotea, Streptomyces sp. (Imran et al. 2016). The fungi used in cellulase production are Trichoderma reesei, T. longibrachiatum, T. harzianum, Aspergillus niger, A. nidulans, Fusarium solani, F. oxysporum, etc. (Imran et al. 2016; Sukumaran et al. 2005).

2.4.1.1

Cellulolytic Microorganisms from Extreme Environment

The microorganisms which can survive under extreme conditions are called extremophiles. The extreme conditions include low pH (acidophiles), alkaline pH

2.4 Cellulase Production

19

(alkalophiles), high salt concentrations (halophiles), tolerant to toxic metals (metallotolerant), etc. There are some cellulase-producing microorganisms which can survive under extreme environments, viz., gram-negative Antarctic bacterium Pseudoalteromonas haloplanktis from the seawater. This microorganism secretes enzyme which is psychrophilic cellulase, Cel5G, which displays good specific activity at low and moderate temperatures as well as high thermo-sensitivity with a reduction in intermolecular interactions. Extreme thermophilic cellulose-degrading microorganisms are of importance due to thermostable enzymes. The thermophilic cellulolytic microorganisms are Rhodothermus marinus, Acidothermus cellulolyticus, Caldicellulosiruptor sp., Clostridium sp., Geobacillus sp., Fervidobacterium sp., and Thermotoga sp. The cellulase enzyme produced by thermophilic bacterium Anaerocellum thermophilum also helps in degradation of lignocellulosic wastes.

2.5 Enzymes Involved in Cellulose Breakdown The cellulose is degraded by enzyme cellulases. Cellulases catalyze the hydrolysis of β-1, 4-glycosidic bonds of cellulose, and are converted to glucose (Bedford and Partridge 2001). The interaction between cellulolytic and non-cellulolytic microorganisms leads to degradation of cellulose with release of carbon dioxide and water under aerobic conditions, and carbon dioxide, methane, and water under anaerobic conditions (Leschine 1995). The fungal cellulases have a larger catalytic and a smaller cellulose binding domain (CBD). These domains are joined by a glycosylated linker peptide (Saloheimo et al. 1997). The CBD is necessary for cellulose degradation. There are three types of cellulases: Endoglucanases or endo-β-1,-4-glucanase break internal bonds and expose polysaccharide chains. Endoglucanases hydrolyze the bonds in amorphous zones. They decrease polymerization and attack cellobiohydrolase (Yuan et al. 2008). These enzymes have cellulose binding and catalytic domains. The catalytic domain is in the form of a crevice, and this helps to attack the cellulose chains everywhere. Cellobiohydrolases act from non-reducing or reducing ends with the release of cellobiose molecules. Cellobiohydrolases have catalytic site for hydrolysis reaction and cellulose binding domain for substrate binding (Bergmans et al. 1996). The catalytic domains of cellobiohydrolases are in the form of a tunnel (Lynd et al. 2002). The cellobiohydrolases allow access to glucose of cellulose chains (Vilanova et al. 2012). Cellobiase or β-glucosidase performs hydrolysis of cellobiose to give two glucose molecules. Cellobiase minimizes inhibition of cellobiohydrolases and endoglucanases by cellobiose. The synergistic action of three enzymes is important. Synergism is first between endo- and exo-β-glucanase and secondly between exo-β-glucanases which acts from reducing and non-reducing end.

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2 Cellulase in Degradation of Lignocellulosic Wastes

2.6 Cellulase Systems of Bacteria and Fungi 2.6.1 Bacterial Cellulase Systems Cellulolytic bacteria can survive both in the presence and in the absence of oxygen. The cellulolytic anaerobic bacteria, viz., C. thermocellum, C. cellulovorans, Acetivibrio cellulolyticus, produce high-molecular-weight complex called as cellulosome. The cellulosome of C. thermocellum comprises various endoglucanases and three exoglucanases. In case of anaerobic bacteria Acidothermus cellulolyticus, the cell bound-enzyme β-galactosidase helps in degradation of cellulose. The commonly studied cellulase system is of C. thermocellum. The cellulosome of this bacterium facilitates degradation of cellulose in the presence of Ca2+ element and dithiothreitol, a reducing agent. The cellulose degradation process is aerobic. Anaerobic process for cellulose degradation is only 5–10%. The top layer of soil contains more carbon. Further study is necessary to know the effects of global warming on decomposition of organic material present in soil which is necessary to understand atmospheric and climatic changes. Anaerobic bacteria also help to degrade cellulose. The cellulose is decomposed by anaerobes into CH4 , CO2 , and H2 O with the help of inorganic electron acceptors. The enzymes produced by cellulolytic microorganisms depolymerize cellulose with the formation of cellobiose, cellodextrins, and glucose. These fermentations produce hydrogen and organic acids, viz., acetate, propionate, butyrate. The hydrogen formed is utilized by methanogenic bacteria to reduce CO2 to CH4 .

2.6.2 Fungi Cellulase Systems The fungus which degrades cellulose efficiently is Trichoderma reesei because of the production of cellulase enzyme. Most of the cellulolytic microorganisms are fungi. Trichoderma reesei has five characterized β-1, 4-endoglucanases, two cellobiohydrolases, and two β-glucosidases (de Souza 2013). The most efficient aerobic bacteria-degrading celluloses are Cellulomonas, Pseudomonas, and Streptomyces sp. (Beguin and Aubert 1994). The fungal cellulases are mainly glycoproteins. Anaerobic cellulase-producing fungi belong to the genera Neocallimastix, Caccomyces, Orpinomyces, Piromyces, and Rurainorayces. Among these species, Neocallimastix frontalis has been studied in detail.

2.7 Microbial Sources of Cellulase Enzyme

21

2.7 Microbial Sources of Cellulase Enzyme 2.7.1 Bacteria Sources Many of the bacteria also produce the enzyme cellulase (Stern et al. 2015). The cellulase-producing bacteria are Bacillus, Cellulomonas, Streptomyces, Cytophaga, Cellvibrio, and Pseudomonas sp. In bacteria, cellulases are present as compact structures and attached to the cells (Juturu and Wu 2014). E. coli are used for recombinant cellulase protein production (Garvey et al. 2013). Bacillus subtilis has good endoglucanase activity and, therefore, is used to engineer recombinant cellulase strains which grow with cellulose as the only source of carbon (Zhang 2011). The genus Clostridium and Thermotoga are also reported to produce cellulase enzyme (Brunecky et al. 2012; Parisutham et al. 2014). The degradation of cellulose involves the action of variety of microorganisms.

2.7.2 Actinobacteria Sources The cellulase system of Thermobifida fusca has six extracellular cellulases (4 endocellulases and 2 exocellulases) and an intracellular β-glucosidase (Wilson 2012). The enzyme has two domains, viz., catalytic and carbohydrate-binding domains, joined by a linker peptide (Gomez-Del and Saadeddin 2014). Cellobiose plays a role as effector and dissociates CelR-DNA complex. The cellulases in C. fimi also have three endocellulases (CenA, CenB, and CenD), two exocellulases (CbhA and CbhB), and a processed endocellulase CenC (Wilson 2004). Microbispora bispora shows synthesis of six cellulases, with exo–exo- and endo–exo-synergism. Streptomyces coelicolor has 221 carbohydrate-active enzymes (CAZymes) or 154 glycosyl hydrolases (GHs) (Takasuka et al. 2013).

2.7.3 Fungi Sources Fungi are important cellulose-degrading microorganisms because of multicomponent, synergetic, cellulolytic, enzyme activity (Juturu and Wu 2014; Ulrich et al. 2008). T. reesei fungi produce cellulase and are most extensively studied (Wahlstrom et al. 2014). The cellulase mixtures of T. reesei consist of exoglucanases, which is 80%, and endoglucanases which is 15% of the total protein (Garvey et al. 2013). Many of the Aspergillus sp. has also been studied for cellulase production (Sakthi et al. 2011). Fungi from asco- and basidiomycetes produce extracellular enzymes to break down celluloses and convert into sugars. Fungal cellulase readily breaks down cellulose into two or three glucose units. Glucose is degraded and assimilated as glucose monomers (Chinedu et al. 2010).

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2 Cellulase in Degradation of Lignocellulosic Wastes

2.8 Cellulosomes to Degrade Cellulose Cellulosomes are multienzyme complexes produced by anaerobic bacteria and fungi (Berger et al. 2007). The cellulosome contains enzymes that have catalytic module together with a CBM. The cellulosomes are produced by Clostridium and Ruminococcus spp., and Chytridiomycetes (Blumer-Schuette et al. 2014). These proteins are found important to degrade cellulose (Zhou et al. 2009). Cellulosomes include subunits of endoglucanases, xylanases, and cellobiohydrolases. Scaffoldins are multidomain and functional proteins which help in interaction between catalytic proteins and glycosyl hydrolase domains (Bayer et al. 2004). The scaffoldin subunits contain many cohesion modules which bind to dockerin module. This interaction helps cellulosomes to degrade cellulose (Xiros et al. 2016). Most of the cellulose is degraded under anaerobic conditions. In case of most anaerobic microorganisms, the enzymes are organized into functional units which are cellulosomes. This arrangement is seen in Clostridium thermocellum. Due to this, the cellulolytic enzymes present at the interface between the cell and the substrate allow products of cellulolysis (e.g., cellobiose) to enter inside the bacterium. The cellulosomes of bacteria Acetivibrio cellulolyticus and Ruminococcus flavefaciens are more complex in comparison with C. thermocellum (Jindou et al. 2006; Xu et al. 2004). The cellulosomes of Acetivibrio cellulolyticus and Ruminococcus flavefaciens contain numerous cohesins and around 200 dockerin-bearing enzymes.

2.8.1 Carbohydrate-Active enZymes (CAZymes) Enzymes that take part in the synthesis, modification, or breakdown of glycoconjugates or complex polysaccharides are carbohydrate-active enzymes (CAZymes) (Cantarel et al. 2009). The CAZy database is divided into five classes, viz., glycoside hydrolases, glycosyl transferases, polysaccharide lyases, carbohydrate esterases, and non-catalytic carbohydrate-binding modules. There are 400 protein families divided into 6 classes in CAZy database. Glycosyltransferases (GTs) form glycosidic bonds and can generate glucoconjugates, oligosaccharides, and polysaccharides (Coutinho et al. 2003). Carbohydrate esterases (CEs) are a class of CAZymes that removes ester-based modifications by de-O or de-N acylation in a hydrolytic way. Polysaccharide lyases (PLs) cleave uronic acid by β-elimination (Lombard et al. 2010). Glycoside hydrolases (GHs) are the enzyme class with large no. of families. These enzymes hydrolyze glycosidic bonds between two carbohydrate moieties or one carbohydrate and one non-carbohydrate moiety. Lytic polysaccharide monooxygenase enzymes work in synergy with GHs (Horn et al. 2012). A non-catalytic class of proteins present in the CAZy database is the carbohydrate-binding modules (CBMs). CBMs are connected with CAZymes in multimodular structures and help in association with the substrate. The auxiliary activity (AA) proteins are newly added to the CAZy database (Levasseur et al. 2013).

2.8 Cellulosomes to Degrade Cellulose

23

2.8.2 Cellobiose Dehydrogenase (CDH) Cellobiose dehydrogenase (CDH) is exo-enzyme system of the fungus Phanerochaete chrysosporium that degrades wood. CDH is a catalytic activator for lytic polysaccharide monooxygenase (LPMO). LPMOs cleave the crystalline parts of cellulose. This provides attack points for cellulases. The CDH/LPMO system thus helps in degradation of cellulose (Langston et al. 2011; Phillips et al. 2011).

2.9 Cellulolytic Mechanisms Few of the enzymes in cellulosomes contain a cellulose binding module (CBM). Scaffoldin contains CBM, which helps the binding of complex to cellulose, and therefore, CBM plays an important role, particularly CBM 33. Aerobic fungi produce GH-7 cellulases. Aerobic fungi like Trichoderma reesei use the free enzyme mechanism to degrade cellulose, while brown-rot fungi secrete both cellulases and peroxidases (Martinez et al. 2005). The peroxidase produces peroxide, and –OH radicals oxidize the cellulose making it easy for the cellulases to degrade cellulose. The brown-rot fungus Postia placenta secretes a single endoglucanase (Martinez et al. 2009; Wilson 2004). The schematic representation of cellulase mechanism of action is shown in Fig. 2.2. Under alkaline conditions, cellulose degrades to form products, viz., αisosaccharinic acid (3-deoxy-2-C-(hydroxymethyl) erythro-pentanoic acids) and βisosaccharinic acid (Knill and Kennedy 2003). Celluloses degrade much more slowly under alkaline conditions. Cellulases secreted by some anaerobic cellulolytic bacteria contain CBM, e.g., Caldicellulosiruptor sp. These anaerobic cellulolytic bacteria have strong cell wall degradation system (Blumer-Schuette et al. 2010). In case of anaerobic cellulolytic bacteria, Fibrobacter succinogenes and Cytophaga hutchinsonii, cellulase genes present in the genome sequence are mostly endocellulases (Wilson 2008; Xie et al. 2007). These two organisms do not secrete free cellulases nor produce cellulosomes and therefore have novel mechanism for cellulose hydrolysis, but not known in detail (Qi et al. 2007). F. succinogenes grows faster on cellulose than other microorganisms (Fields et al. 2000). Many cellulolytic fungi contain numerous family GH-61 genes (Harris et al. 2010). In case of Thermobifida fusca, the organism when grown on cellulose secretes two proteins, viz., family 33 domain and family 33 CBM linked to a family 2 CBM. These two proteins bind to cellulose and help in cellulose hydrolysis (Moser et al. 2008). The cellulose degradation ability is different among each bacteria and fungi (Meryandini et al. 2009). In case of Trichoderma reesei, the breakdown of cellulose involves two steps: Cellulose

β-1,4 glucanase

−→

Cellobiose

β-glucosidase

−→

glucose

(2.1)

Substrate reducing end

Fig. 2.2 Schematic representation of cellulase mechanism of action

Cellulose attach to the substrate

Insert substrate into active site

Development of catalytic complex

Breakdown of Substrate

24 2 Cellulase in Degradation of Lignocellulosic Wastes

2.9 Cellulolytic Mechanisms

25

Moreira et al. (2011). Anaerobic microbial degradation of cellulosic materials also occurs (Lynd et al. 2002). Anaerobic microbial cellulose degradation occurs by number of steps, and the end products are CO2 and CH4 . During the oxidative cleavage of cellulose, CBM33 protein which is CelS2 cleaves cellulose and produces aldonic acids (Forsberg et al. 2011). GH61s are functionally similar to CBM33s (Beeson et al. 2012; Chandra et al. 2007; Li et al. 2012). The activity of GH61s enzyme is copper-dependent lytic polysaccharide monooxygenases. There are differences among GH61 sequences which is responsible functional differences (Li et al. 2012). C4 oxidation with formation of reducing ends is favorable. The cellulose degradation will produce monomeric and dimeric sugars (gluconic and cellobionic acid) (Cannella et al. 2012). GH61s accept electrons by the action of cellobiose dehydrogenase (Langston et al. 2011; Phillips et al. 2011).

2.10 Assay Methods 2.10.1 Determination of Cellulase Activity (CMCase Assay) CMCase assay is used to check cellulose degradation. One ml of the crude enzyme supernatant incubates with 1 ml of 1% CMC in 0.1 M sodium acetate buffer solution (pH 5.0) for 30 min at 63 °C. The resulting reducing sugars are determined according to Miller’s modified method of DNSA.

2.10.1.1

Total Cellulase (FPase) Assay

Total cellulase (FPase) activity is determined by the method of Gadgil et al. (1995). One ml of the crude enzyme supernatant incubates with 2 ml of 0.1 M citrate buffer (pH 4.8) containing 50 mg Whatman No. 1 filter paper. Incubate for 1 h at 50 °C, and estimate the reducing sugars which were determined according to Miller’s modified method of DNSA (Gadgil et al. 1995).

Miller’s Modified Method of DNSA The filtrate is collected from the fermentation media by centrifugation. Take 1 ml of filtrate in a test tube; add 1 ml of 1% cellulose solution and 1 ml of distilled water into it. Control tube is 1 ml of 1% cellulose solution and 2 ml of distilled water. Allow it to react for 30 min, and add 3 ml of DNSA reagent to each test tube. Heat the contents of test tubes in boiling water bath for 5 min, and then allow the

26

2 Cellulase in Degradation of Lignocellulosic Wastes Total cellulase activity

Individual cellulase activity

Endoglucanases activity

CMC

Exoglucanases activity

Avicel

β - glucosidase activity

Cellobiose

Reducing sugars

Reducing sugars

Fig. 2.3 Cellulase assays

test tubes to cool at room temperature. At the time of cooling, add 7 ml of freshly prepared 40% sodium potassium tartrate solution. Measure the absorbance at 540 nm by using spectrophotometer. The amount of reducing sugar was determined using standard graph (Miller 1972).

Determination of Cellulase Enzyme Assay The cellulase enzyme is assayed by determining the amount of glucose released from the substrates by Miller’s modified method of DNSA. One unit of cellulase activity is the amount of enzyme which liberates 10 ug of glucose in 30 min under optimized condition (Milala et al. 2005). The different cellulase assays are shown in Fig. 2.3 (Dashtban et al. 2010).

2.11 Physical and Biological Methods for Pretreatment of Cellulose

27

2.11 Physical and Biological Methods for Pretreatment of Cellulose 2.11.1 Physical Methods for Pretreatment of Cellulose The physical methods for pretreatment of cellulose are required before degradation of lignocellulose wastes. Physical methods reduce particle size and crystallinity, and also the degree of polymerization. The physical pretreatment will improve hydrolysis due to increase in surface area. Physical pretreatment methods of cellulose include milling and grinding, steam explosion, pyrolysis, and electrical pulses (Kim et al. 2003).

2.11.1.1

Acids

Acids used for pretreatment are hydrochloric, phosphoric, or sulfuric acid. The acid is proton source for the hydrolysis reaction, which breaks down sugar polymers in cellulose.

2.11.1.2

Alkalis

The alkalis used for cellulose pretreatment are sodium, potassium, calcium, and ammonium hydroxides.

2.11.1.3

Organic Solvents

The organic solvents used are alcohols, ketones, glycols, organic acids, phenols, esters, and ether. This pretreatment method involves the addition of a catalyst such as dilute sulfuric, oxalic, salicylic, and acetylsalicylic acids (Huijgen et al. 2011; Sun and Cheng 2002).

2.11.1.4

Ionic Liquids

Ionic liquids are salts which contain organic cations and inorganic anions. Anions interact with a hydroxyl group in cellulose (Janesko 2011). This results in the breakdown of cellulose.

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2 Cellulase in Degradation of Lignocellulosic Wastes

2.11.2 Biological Methods Biological methods use fungi and bacteria for the hydrolysis of cellulose. Brownand soft-rot fungi attack cellulose (Sun and Cheng 2002). The white-rot fungi used for pretreatment of cellulose are Ceriporiopsis vermispora, Dichomitus squalens, Pleurotus ostreatus, and Coriolus versicolor (Itoh et al. 2003). The advantages of biological pretreatment are low energy input, mild environmental conditions, and eco-friendly (Salvachua et al. 2011).

2.12 Mechanism of Cellulose Hydrolysis Cellulase enzyme hydrolyzes cellulose by β-1-4 glycosidic bonds of cellulose polymer. The hydrolysis product is glucose carried by synergetic action of three enzymes, endoglucanases, which cleave the internal bonds of the glycan chains. The reducing or non-reducing ends of cello oligosaccharides are further attacked by cellobiohydrolases (CBHs). CBH then hydrolyzes those chain ends to yield cellobiose. β-glucosidase hydrolyses cellobiose from the non-reducing ends of soluble cello oligosaccharides to give glucose (Jorgensen et al. 2007). The carbohydrate-binding module is a cellulose probe, which helps to bind the enzyme to the cellulose (Araki et al. 2010) and enhance the enzymatic activity. The reactions in degradation of cellulose are (Beaton et al. 2019): (i) Hydrolysis of cellulose into β-d-cellobiose

cellulose −→ β-d-cellobiose(CH2 O)

(2.2)

(ii) Hydrolysis of β-d-cellobiose and fermentation

CH2 O −→ organic acids + CO2 + H2

(2.3)

(iii) Hydrogenotrophic methanogenesis

4H2 + CO2 −→ CH4 + 2H2 O

(2.4)

4H2 + 2CO2 −→ CH4 + 2H2 O

(2.5)

(iv) Acetogenesis

2.12 Mechanism of Cellulose Hydrolysis

CO2 −→ CH3 COOH + 2H2 O

29

(2.6)

(v) Aceticlastic methanogenesis

CH3 COOH −→ CH4 + CO2

(2.7)

The cellulases which hydrolyze 1,4-β-glycosidic bonds are (Rabinovich et al. 2002): (i) (ii) (iii) (iv) (v) (vi) (vii)

Fungal endo-1,4-β-mannanases, and aerobic and anaerobic bacteria endoglucanases; Endoglucanases of actinomycetes, and aerobic and anaerobic bacteria; Exo-1,3-β-glucanases; endoglucanases/1,3-1,4-β-glucanases and 1,3-β-glucanases (yeast, Clostridium); Endoglucanases and mannanases of actinomycetes, aerobic and anaerobic bacteria, and anaerobic fungi; Endoglucanases of filamentous fungi and aerobic bacteria; Endo-1,6-β-glucanases; Cellodextrin phosphorylase or 1, 4-β-d-oligoglucan orthophosphate α-dglucosyl transferase;

(1, 4β-d-glucosyl)n + H3 PO4 −→(1, 4β-d-glucosyl)n − 1 + α-d-Glucose-1-P (2.8) (viii) Cellobiose epimerase. It was first reported in cells of Ruminococcus albus. It catalyzes the following reaction: Cellobiose −→ 4-O-β-d-glucosylmannose

(2.9)

The breakdown of cellulose to ethanol is represented in Fig. 2.4.

2.12.1 Genes Related to Cellulose Degradation The CAZyme contents of the genomes of basidiomycetes and Aspergillus species are same. Aspergillus and basidiomycetes have similar numbers of genes encoding CBHs (GH6 and -7). Aspergillus has more BGL genes in GH3 than basidiomycetes. The GH9-encoding genes are absent in Aspergillus. LPMO-encoding genes are more in basidiomycetes in comparison with Aspergillus (Zifcakova and Baldrian 2012).

pretreatment

Fig. 2.4 Breakdown of cellulose to ethanol

cellulose acids /enzymes

hydrolysis by

(solid state fermentation)

to ethanol (C2H5OH)

fermentation of sugars

Ethanol

distillation

and separation by

concentration

30 2 Cellulase in Degradation of Lignocellulosic Wastes

2.13 Transcriptional Regulators Involved in Regulation …

31

2.13 Transcriptional Regulators Involved in Regulation of Cellulolytic Gene Expression (T. reesei) (Shida et al. 2016) 2.13.1 Xylanase Regulator (Xyr1) The major activator and transcription factor for cellulase genes is Xyr1. The Xyr1 open-reading frame encodes Zn2Cys6-binuclear zinc cluster with 934 amino acids and has molecular weight 102 kDa. Xyr1 also controls the transcription of the gene which encodes d-xylose reductase (xyl1).

2.13.2 Activator of Cellulase Expression 2 (Ace2) Ace2 is another activator of cellulase which is isolated from cDNA library of T. reesei by yeast expression screening method.

2.13.3 Activator of Cellulase Expression 1 (Ace1) Ace1 is a class I zinc finger protein which has three Cys2His2-type zinc finger repressors and is isolated from cellulose-induced cDNA library of T. reesei.

2.13.4 Beta-Glucosidase Regulator (BglR) The BglR-encoding fungal-specific zinc binuclear cluster protein is BglR and plays a role in the activation of BGL genes.

2.13.5 Activator of Cellulase Expression 3 (Ace3) The Ace3 is present on upstream of Xyr1 in the induction mechanism of cellulase.

32

2 Cellulase in Degradation of Lignocellulosic Wastes

2.13.6 GH Families Involved in Cellulose Degradation A number of GH families are involved in cellulose degradation. The details of these families are mentioned below.

2.13.6.1

GH Family 3

The GH family 3 (GH3) is abundant in the CAZy database, which consists of 6000 enzymes. The family exhibits activities, viz., exo-acting β-d-glucosidases, α-l-arabinofuranosidases, β-d-xyloparanoside, and N-acetyl-β-d-glucosaminidases (Hamelinck et al. 2005; Pan et al. 2007; Xiros et al. 2016). Few GH3 enzymes catalyze glycosidic bond formation by reverse hydrolysis or trans-glycosylation (Lee et al. 2007). These enzymes also perform synthesis of functional glycosides from glycoside precursors (Bhat 2000). The exception is in case of Cellulomonas fimi Nag3, where glycosyl residues are singly removed from non-reducing ends by Koshland double-displacement mechanism (Fincher et al. 2017). There are two glucosyl-binding subsites on the active site of a GH3 enzyme.

2.13.6.2

GH Family 5

GH family 5 (GH5) belongs to Clan GH-A. There are about 51 GH5 threedimensional structures. The enzymes of GH5 consist of an amino acid chain which forms a (β/α)8 fold. This leads to formation of open groove which harbors the nucleophile Glu and acid/base Glu at the C terminus of β-strand 7 and β-strand 4, respectively. There is binding of carbohydrate substrate to the substrate-binding site from non-reducing end (− subsites) to reducing end (+ subsites) during the catalytic reaction.

2.13.6.3

GH Family 9

It is the second largest cellulase family and is mainly endoglucanases. This endoglucanase has a CBM of 3C family bound to C terminus of the catalytic domain. The cellulases of GH family 9 (GH9) are divided into EI and EII subgroups. EI subgroup has only bacterial cellulases, and EII consists of cellulases of bacterial and non-bacterial origin. The action of GH9 enzymes is by inversion of anomeric stereochemistry (Wilson and Urbanowicz 2017). In catalysis reaction, three amino acids are involved. The catalytic triad formed consists of a conserved Glu residue and two Asp residues.

2.13 Transcriptional Regulators Involved in Regulation … Table 2.3 Number of GH families in different bacterial sp. (Gibson et al. 2011)

Phylum

Species of

Proteobacteria

Agrobacterium

4

Ralstonia

5

Erwinia Pectobacterium

Actinobacteria

Table 2.4 No. of GH families in different fungal sp. (Gibson et al. 2011)

Total GH

5 15

Pseudomonas

9

Xanthomonas

20

Xylella

4

Clavibacter

9

Phylum

Species of

Ascomycota

Blumeria

Basidiomycota

2.13.6.4

33

Total GH 2

Botrytis

41

Sclerotinia

35

Fusarium

43

Magnaporthe

60

Trichoderma

22

Puccinia

23

Ustilago

6

Lytic Polysaccharide Monooxygenase (LPMO)

Cellulases are major part of the enzymes in glycoside hydrolase (GH) families, which hydrolyze the glycosidic bonds in glucose polymers. Studies show the enzymes break glycosidic bonds of glucose molecules through oxidation mechanism instead of hydrolysis (Forsberg et al. 2014; Langston et al. 2011; Vaaje-Kolstad et al. 2010). These enzymes are classified as lytic polysaccharide monooxygenase (Horn et al. 2012). The reaction carried by this enzyme is copper-dependent (Quinlan et al. 2011; Westereng et al. 2012). A number of GH families in different bacterial and fungal sp. are shown in Tables 2.3 and 2.4, respectively.

2.14 Applications of Enzyme Cellulase Cellulase has wide applications in various industries viz., textile, paper and pulp industries, etc.

34

2 Cellulase in Degradation of Lignocellulosic Wastes

2.14.1 Paper and Pulp Industries Cellulases are used for pulping and deinking of papers. Application of cellulase in pulping industries improves the physical properties such as interfiber bonding and mechanical strength (Chen et al. 2012) and avoids the use of hazardous chemicals. The deinking of papers involves enzymatic hydrolysis of carbohydrates, leading to the peeling of individual fibers (Kuhad et al. 1997). This deinking of waste papers improves the quality and brightness of recycled paper (Ibarra et al. 2012).

2.14.2 Textile Industry In textile industry, cellulases are used for biopolishing of cotton cloths and biostoning of denim jeans to give stonewashed look for denims. Cellulases hydrolyze the small fiber protrusions and remove the indigo dye attached to it during biostoning process. Cellulase replaces the conventional pumice stones and thus reduces the fiber damage caused in the process (Arja 2007). In biopolishing process, cellulases hydrolyze the small protrusions of the fibers, which give smooth and glossy appearance (Saravanan et al. 2009). Cellulases remove excess dye and impart color gradient to the fabrics (Kuhad et al. 2011). Cellulases have important applications in household laundry detergents to enhance fabric softness and brightness.

2.14.3 Food and Feeds Cellulases also have important applications in food industries. Cellulases along with other cell wall-degrading enzymes improve the taste and aroma of citrus fruits (Kuhad et al. 2011). Cellulases improve the liquor yield and aroma of beer and wine (Karmarkar and Ray 2010).

2.14.4 Bioethanol The extensive use of fossil fuels has resulted in reducing their natural reserves and causes environmental pollution. The focus is now on biofuels, especially bioethanol, which will replace use of fossil fuels (Msangi et al. 2007; Rosegrant et al. 2006). Cellulase is used in the production of bioethanol. Cellulases convert the cellulosic resources to glucose which is substrate for bioethanol production. Agriculture residues, viz., sugarcane bagasse, wheat straw, rice bran, corn stover, etc., are used as raw materials for bioethanol production, employing cellulases produced by fungi (Binod et al. 2010; Chen and Qiu 2010; Li et al. 2011).

References

35

References Aitken S (2012) Wood chemistry and secondary cell wall structure. FRST 210-Forest Biology II, The University of British Columbia, Vancouver Araki Y, Karita S, Tsuchiya T, Kondo M, Goto M (2010) Family 17 and 28 carbohydrate-binding modules discriminated different cell-wall sites in sweet potato roots. Biosci Biotechnol Biochem 74:802–805 Arja M (2007) Cellulases in the textile industry industrial enzymes. In: Industrial enzymes. Springer, Berlin, pp 51–63 Aro N, Pakula T, Penttila M (2005) Transcriptional regulation of plant cell wall degradation by filamentous fungi. FEMS Microbiol Revolut 29:719–739 Asokan P, Mohini S, Shyam A (2007) Solid wastes generation in India and their recycling potential in building materials. Build Environ 42:2311–2320 Bai S, Kumar M, Kumar D, Balashanmungam P, Bala M, Kalaichelvan P (2012) Cellulase production by Bacillus subtilis isolated from cow dung. Arch Appl Sci Res 4:269–279 Bayer E, Belaich J, Shoham Y, Lamed R (2004) The cellulosomes: multienzyme machines for degradation of plant cell wall polysaccharides. Annu Rev Microbiol 58:521–554 Beaton D, Pelletier P, Goulet R (2019) Microbial degradation of cellulosic material and gas generation: Implications for the management low and intermediate-level radioactive waste. Front Microbiol 10:1–13 Bedford M, Partridge G (2001) Enzymes in farm animal nutrition. In: Bedford M, Partridge G (eds) CAB International, Wallingford, pp 9–10 Beeson W, Phillips C, Cate J, Marletta M (2012) Oxidative cleavage of cellulose by fungal copperdependent polysaccharide monooxygenases. J Am Chem Soc 134:890–892 Beguin P, Aubert J (1994) The biological degradation of cellulose. FEMS Microbiol Rev 13:25–58 Berger E, Zhang D, Zverlov V, Schwarz W (2007) Two non cellulosomal cellulases of Clostridium thermocellum, Cel9I and Cel48Y, hydrolyse crystalline cellulose synergistically. FEMS Microbiol Lett 268:194–201 Bergmans M, Beldman G, Gruppen H, Voragen A (1996) Optimisation of the selective extraction of (glucurono) arabinoxylans from wheat bran: use of barium and calcium hydroxide solution at elevated temperatures. J Cereal Sci 23:235–245 Bhat M (2000) Cellulases and related enzymes in biotechnology. Biotechnol Adv 18:355–383 Binod P, Sindhu R, Singhania R, Vikram S, Devi L, Nagalakshmi S, Kurien N, Sukumaran R, Pandey A (2010) Bioethanol production from rice straw: an overview. Bioresour Technol 101:4767–4774 Blumer-Schuette S, Lewis D, Kelly R (2010) Phylogenetic, microbiological, and glycoside hydrolase diversities within the extremely thermophilic, plant biomass-degrading genus Caldicellulosiruptor. Appl Environ Microbiol 76:8084–8092 Blumer-Schuette S, Brown S, Sander K, Bayer E, Kataeva I, Zurawski J, Conway J, Adams M, Kelly R (2014) Thermophilic lignocellulose deconstruction. FEMS Microbiol Rev 38:393–448 Brijwani K, Vadlani P (2011) Cellulolytic enzymes production via solid state fermentation: effect of pretreatment methods on physiochemical characteristics of substrate. Enzyme Res 2011:1–10 Brown MR Jr (1996) The biosynthesis of cellulose. J Macromol Sci Part A Pure Appl Chem 33:1345–1373 Brunecky R, Alahuhta M, Bomble Y, Xu Q (2012) Structure and function of the Clostridium thermocellum cellobiohydrolase A X1-module repeat: enhancement through stabilization of the CbhA complex. Acta Crystallogr Sect D Biol Crystallogr 68:292–299 Cannella D, Hsieh C-w, Felby C, Jorgensen H (2012) Production and effect of aldonic acids during enzymatic hydrolysis of lignocellulose at high dry matter content. Biotechnol Biofuels 5:1–10 Cantarel B, Coutinho P, Rancurel C, Bernard T, Lombard V, Henrissat B (2009) The carbohydrateactive enzymes database (cazy): an expert resource for glycogenomics. Nucleic Acids Res 37:233–238 Chandra R, Bura R, Mabee W, Berlin A, Pan X, Saddler J (2007) Substrate pretreatment: the key to effective enzymatic hydrolysis of lignocellulosics? Adv Biochem Eng Biotechnol 108:67–93

36

2 Cellulase in Degradation of Lignocellulosic Wastes

Chen H, Qiu W (2010) Key technologies for bioethanol production from lignocellulose. Biotechnol Adv 28:556–562 Chen Y, Wan J, Zhang X, Ma Y, Wang Y (2012) Effect of beating on recycled properties of unbleached eucalyptus cellulose fiber. Carbohydr Polym 87:730–736 Chinedu S, Eni A, Adeniyi A, Ayangbemi J (2010) Assessment of growth and cellulase production of wild-type microfungi isolated from Ota, Nigeria. Asian J Plant Sci 9:118–125 Coutinho P, Deleury E, Davies G, Henrissat B (2003) An evolving hierarchical family classification for glycosyl transferases. J Mol Biol 328:307–317 Dashtban M, Maki M, Leung K, Mao C, Qin W (2010) Cellulase activities in biomass conversion: measurement methods and comparison. Crit Rev Biotechnol 30:302–309 Deka D, Bhargavi P, Sharma A, Goyal D, Jawed M, Goyal A (2011) Enhancement of cellulose activity from a new strain of Bacillus subtilis by medium optimization and analysis with various cellulosic substrates. Enzyme Res 2011:1–8 Deswal D, Khasa Y, Kuhad R (2011) Optimization of cellulase production by a brown rot fungus Fomitopsis sp. RCK2010 under solid state fermentation. Bioresour Technol 102:6065–6072 Dogaris I, Vakontios G, Kalogeris E, Mamma D, Kekos D (2009) Induction of cellulases and hemicellulases from Neurospora crassa under solid-state cultivation for bioconversion of sorghum bagasse into ethanol. Ind Crops Prod 29:404–411 Facchini F, Vici A, Reis V, Jorge J, Terenzi H, Reis R, Polizeli M (2011) Production of fibrolytic enzymes by Aspergillus japonicus C03 using agro-industrial residues with potential application as additives in animal feed. Bioproc Biosyst Eng 34:347–355 Fields M, Mallik S, Russell J (2000) Fibrobacter succinogenes S85 ferments ball-milled cellulose as fast as cellobiose until cellulose surface area is limiting. Appl Microbiol Biotechnol 54:570–574 Fincher G, Mark B, Brumer H (2017) In: Henrissat B (ed) Glycoside hydrolase family 3. CAZypedia Forsberg Z, Vaaje-Kolstad G, Westereng B, Bunaes AC, Stenstrom Y, MacKenzie A, Sorlie M, Horn S, Eijsink V (2011) Cleavage of cellulose by a CBM33 protein. Prot Sci 20:1479–1483 Forsberg Z, Mackenzie A, Sorlie M, Rohr AK, Helland R, Arvai A, Vaaje-Kolstad G, Eijsink V (2014) Structural and functional characterization of a conserved pair of bacterial celluloseoxidizing lytic polysaccharide monooxygenases. Proc Natl Acad Sci 111:8446–8451 Gadgil N, Daginawala H, Chakarabarti T, Khanna P (1995) Enhanced cellulase production by a mutant of Trichoderma reesei. Enzyme Microb Technol 17:942–946 Garvey M, Klose H, Fischer R et al (2013) Cellulases for biomass degradation: comparing recombinant cellulase expression platforms. Trends Biotechnol 31:581–593 Gibson D, King B, Hayes M, Bergstrom G (2011) Plant pathogens as a source of diverse enzymes for lignocellulose digestion. Curr Opin Microbiol 14:264–270 Gomez-Del P, Saadeddin A (2014) The cellulolytic system of Thermobifida fusca. Crit Rev Microbiol 40:236–247 Goshadrou A, Karimi K, Taherzadeh M (2011) Bioethanol production from sweet sorghum bagasse by Mucor hiemalis. Ind Crops Prod 34:1219–1225 Hamelinck C, van Hooijdonk G, Faaij A (2005) Ethanol from lignocellulosic biomass: technoeconomic performance in short-, middle- and long-term. Biomass Bioenerg 28:384–410 Harris P, Welner D, McFarland K, Re E, Navarro P, Brown K, Salbo R, Ding H, Vlasenko E, Merino S et al (2010) Stimulation of lignocellulosic biomass hydrolysis by proteins of glycoside hydrolase family 61: structure and function of a large, enigmatic family. Biochem 49:3305–3316 Heck J, Hertz P, Ayub M (2002) Cellulase and xylanase production by isolated Amazon Bacillus strains using soya been industrial residue based solid-state cultivation. Braz J Microbiol 33:213– 218 Herculano P, Porto T, Moreira K, Pinto G, Souza-Motta C, Porto A (2011) Cellulase production by Aspergillus japonicas URM5620 using waste from caster bean (Ricinus communis L.) under solid-state fermentation. Appl Biochem Biotechnol 165:1057–1067 Horn S, Vaaje-Kolstad G, Westereng B, Eijsink V (2012) Novel enzymes for the degradation of cellulose. Biotechnol Biofuels 5:45–56

References

37

Huijgen W, Smit A, Reith J, Uil HD (2011) Catalytic organosolv fractionation of willow wood and wheat straw as pretreatment for enzymatic cellulose hydrolysis. J Chem Technol Biotechnol 86:1428–1438 Ibarra D, Monte M, Blanco A, Martinez A, Martinez M (2012) Enzymatic deinking of secondary fibers: cellulases/hemicellulases versus laccase-mediator system. J Ind Microbiol Biotechnol 98:1684–1689 Imran M, Anwar Z, Irshad M, Asad M, Ashfaq H (2016) Cellulase production from species of fungi and bacteria from agricultural wastes and its utilization in industry: a review. Adv Enzyme Res 4:44–55 Itoh H, Wada M, Honda Y, Kuwahara M, Watanabe T (2003) Bioorganosolve pretreatments for simultaneous saccharification and fermentation of beech wood by ethanolysis and white rot fungi. J Biotechnol 103:273–280 Janesko B (2011) Modeling interactions between lignocellulose and ionic liquids using DFT-D. PCCP 13:11393–11401 Jindou S, Borovok I, Rincon M, Flint H, Antonopoulos D, Berg M, White B, Bayer E, Lamed R (2006) Conservation and divergence in cellulosome architecture between two strains of Ruminococcus flavefaciens. J Bacteriol 188:7971–7976 Jorgensen H, Kristensen J, Felby C (2007) Biofuels. Bioprod Biorefin 1:119–134 Juturu V, Wu J (2014) Microbial cellulases: engineering, production and applications. Renew Sustain Energy Rev 33:188–203 Kang S, Park LY, Hong J, Kim S (2004) Production of cellulases and hemicellulases by Aspergillus niger KK2 from lignocellulosic biomass. Bioresour Technol 91:153–156 Karmarkar M, Ray R (2010) Current trends in research and application of microbial cellulases. RJM 6:41–53 Kim J, Park C, Kim T-H, Lee M, Kim S, Kim S-W, Lee J (2003) Effects of various pretreatments for enhanced anaerobic digestion with waste activated sludge. J Biosci Bioeng 95:271–275 Klemm D, Heublein B, Hans-Peter F, Bohn A (2005) Cellulose: fascinating biopolymer and sustainable raw material. Chem Inform 44:3358–3393 Knill C, Kennedy J (2003) Degradation of cellulose under alkaline conditions. Carbohydr Polym 51:281–300 Kontturi E (2005) Surface chemistry of cellulose: from natural fibers to model surfaces. Technische Universiteit Eindhoven Kuhad R, Singh A, Eriksson K (1997) Microorganisms and enzymes involved in the degradation of plant fiber cell walls. Adv Biochem Eng Biotechnol 57:45–125 Kuhad R, Gupta R, Singh A (2011) Microbial cellulases and their industrial applications. Enzyme Res 2011:1–10 Lah T, Rahman N, Nama M (2012) Cellulase activity and glucose production by Bacillus cereus monoculture and co-culture utilizing palm kernel cake (PKC) under solid state fermentation. In: International conference on environment, energy and biotechnology, Singapore, pp 172–177 Langston J, Shaghasi T, Abbate E, Xu F, Vlasenko E, Sweeney M (2011) Oxidoreductive cellulose depolymerization by the enzymes cellobiose dehydrogenase and glycoside hydrolase 61. Appl Environ Microbiol 77:7007–7015 Lee J, Gwak K, Park J, Park M, Choi D, Kwon M, Choi I (2007) Biological pretreatment of softwood Pinus densiflora by three white rot fungi. J Microbiol 45:485–491 Leschine S (1995) Cellulose degradation in anaerobic environments. Annu Rev Microbiol 49:399– 426 Levasseur A, Drula E, Lombard V, Coutinho P, Henrissat B (2013) Expansion of the enzymatic repertoire of the CAZy database to integrate auxiliary redox enzymes. Biotechnol Biofuels 6:1–14 Li D, Li A, Papageorgiou A (2011) Cellulases from thermophilic fungi: Recent insights and biotechnological potential. Enzyme Res 2011:1–9 Li X, Beeson W, Phillips C, Marletta M, Cate J (2012) Structural basis for substrate targeting and catalysis by fungal polysaccharide monooxygenases. Structure 20:1051–1061

38

2 Cellulase in Degradation of Lignocellulosic Wastes

Limayem A, Ricke S (2012) Lignocellulosic biomass for bioethanol production: current perspectives, potential issues and future prospects. Prog Energy Comb Sci 38:449–467 Lo Y, Saratale G, Chen W, Bai M, Chang J (2009) Isolation of cellulose-hydrolytic bacteria and applications of the cellulolytic enzymes for cellulosic biohydrogen production. Enzyme Microb Technol 44:417–425 Lombard V, Bernard T, Rancurel C, Brumer H, Coutinho P, Henrissat B (2010) A hierarchical classification of polysaccharide lyases for glycolgenomics. Biochem J 432:437–444 Lopez-Contreras A, Gabor K, Martens A, Renckens B, Claassen P, van der Oost J, de Vos W (2004) Substrate-induced production and secretion of cellulases by Clostridium acetobutylicum. Appl Environ Microbiol 70:5238–5243 Lynd L, Weimer P, van Zyl W, Pretorius I (2002) Microbial cellulose utilization: fundamentals and biotechnology. Microbiol Mol Biol Rev 66:506–577 Mawadza C, Hatti-Kaul R, Zvauya R, Mattiasson B (2000) Purification and characterization of cellulases produced by two Bacillus strains. J Biotechnol 83:177–187 Martinez A, Speranza M, Ruiz-Duenas F, Ferreira P, Camarero S, Guillen F, Martınez M, Gutierrez A, del Rio J (2005) Biodegradation of lignocellulosics: microbial, chemical, and enzymatic aspects of the fungal attack of lignin. Int Microbiol 8:195–204 Martinez D, Challacombe J, Morgenstern I, Hibbett D, Schmoll M, Kubicek C, Ferreira P, RuizDuenas F, Martinez A, Kersten P et al (2009) Genome, transcriptome, and secretome analysis of wood decay fungus Postia placenta supports unique mechanisms of lignocellulose conversion. Proc Natl Acad Sci 106:1954–1959 Meryandini A, Widosari W, Maranatha B, Sunarti T, Rachmania N, Satria H (2009) Isolasi bakteri selulolitik dan karakterisasi enzimnya. Makara J Sci 13:33–39 Milala M, Shugaba A, Gidado A, Ene A, Wafar J (2005) Studies on the use of agricultural wastes for cellulase enzyme production by Aspergillus niger. Res J Agric Biol Sci 1:325–328 Miller G (1972) Use of Dinitrosalicylic acid reagent for determination of reducing sugar. Biotechnol Bioeng Symp 5:193–219 Mishra B, Pandey Lata A (2007) Lignocellulolytic enzymeproduction from submerged fermentation of paddy straw. Ind J Microbiol 47:176–179 Mohite B, Magar J (2010) Use of agricultural waste of cellulase production by Aspergillus niger with submerged and solid state fermentation. Bionano Front 2:189–192 Moreira L, Milanezi N, Filho E (2011) Enzymology of plant cell wall breakdown: an update. In: Buckeridge M, Goldman G (eds) Routes to cellulosic ethanol. Springer, New York, pp 73–96 Moser F, Irwin D, Chen S, Wilson D (2008) Regulation and characterization of Thermobifida fusca carbohydrate-binding module proteins E7 and E8. Biotechnol Bioeng 100:1066–1077 Msangi S, Sulser T, Rosegrant M, Valmonte-Santos R (2007) Global scenarios for biofuels: Impacts and implications for food security and water use. Paper presented at the 10th annual conference on global economic analysis, Purdue University, Indiana Narra M, Dixit G, Divecha J, Madamwar D, Shah A (2012) Production of cellulases by solid state fermentation with Aspergillus terreus and enzymatic hydrolysis of mild alkali-treated rice straw. Bioresour Technol 121:355–361 Nizamudeen S, Bajaj B (2009) A novel thermo-alkalitolerant endoglucanase production using costeffective agricultural residues as substrates by a newly isolated Bacillus sp. NZ. Food Technol Biotechnol 47:435–440 Odeniyi O, Onilude A, Ayodele M (2009) Production characteristics and properties of cellulase/polygalacturonase by a Bacillus coagulans strain from a fermenting palm-fruit industrial residue. Afr J Microbiol Res 3:407–417 Oyeleke S, Oyewole O, Egwim E, Dauda B, Ibeh E (2012) Cellulase and pectinase production potentials of Aspergillus niger isolated from corn cob. Bayero J Pure Appl Sci 5:78–83 Pan X, Xie D, Kang K, Yoon S, Saddler J (2007) Effect of organosolv ethanol pretreatment variables on physical characteristics of hybrid poplar substrates. Appl Biochem Biotechnol 137:367–377 Pandey A, Soccol C, Rodriguez-Leon J, Nigam P (2001) Solid state fermentation in biotechnology: fundamentals and applications. Asiatech Publishers, New Delhi, pp 1–221

References

39

Parisutham V, Kim T, Lee S (2014) Feasibilities of consolidated bioprocessing microbes: From pretreatment to biofuel production. Bioresour Technol 161:431–440 Phillips C, Beeson W, Cate J, Marletta M (2011) Cellobiose dehydrogenase and a copper-dependent polysaccharide monooxygenase potentiate cellulose degradation by Neurospora crassa. ACS Chem Biol 6:1399–1406 Pothiraj C, Balaji P, Eyini M (2006) Enhanced production of cellulases by various fungal cultures in solid state fermentation of cassava waste. Afr J Biotechnol 5:1882–1885 Qi M, Jun H, Forsberg C (2007) Characterization and synergistic interactions of Fibrobacter succinogenes glycoside hydrolases. Appl Environ Microbiol 73:6098–6105 Quinlan R, Sweeney M, Lo Leggio L, Otten H, Poulsen J, Johansen K, Krogh K, Jorgensen C, Tovborg M, Anthonsen A, Tryfona T, Walter C, Dupree P, Xu F, Davies G, Walton P (2011) Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Proc Natl Acad Sci 108:15079–15084 Rabinovich M, Melnik M, Bolobova A (2002) Microbial cellulases: a review. Appl Biochem Microbiol 38:305–321 Rai P, Tiwari S, Gaur R (2012) Optimization of process parameters for cellulase production by novel thermotolerant yeast. BioResources 7:5401–5414 Ramanathan G, Banupriya S, Abirami D (2010) Production and optimization of cellulase from Fusarium oxysporum. J Sci Ind Res 69:454–459 de Souza WR (2013) Microbial degradation of lignocellulosic biomass. In: Chandel A, da Silva S (eds) Sustainable degradation of degradation of lignocellulosic biomass. InTechOpen Rosegrant M, Msangi S, Sulser T, Valmonte-Santos R (2006) Biofuels and the global food balance. Bioenergy and agriculture: promises and challenges. International Food Policy Research Institute (IFPRI), Washington, DC Sachin T, Vishal G, Sandeep C, Rohan I, Ajinkya K, Sachin M, Meena P (2011) Production and characterization of cellulase by local fungal isolate of India using water hyacinth as carbon source and reuse of fungal biomass for dye degradation. Int J Eng Sci Technol 3:3236–3241 Sakthi S, Saranraj P, Rajasekar M (2011) Optimization for cellulase production by Aspergillus niger using paddy straw as substrate. Int J Adv Sci Tech Res 1:68–85 Saloheimo M, Nakari-Setala T, Tenkanen M, Penttila M (1997) cDNA cloning of a Trichoderma reesei cellulase and demonstration of endoglucanase activity by expression in yeast. Eur J Biochem 249:584–591 Salvachua D, Prieto A, Lppez-Abelairas M, Lu-Chau T, Martinez Á, Martínez M (2011) Fungal pretreatment: An alternative in second-generation ethanol from wheat straw. Bioresour Technol 102:7500–7506 Saravanan D, Vasanthi N, Ramachandran T (2009) A review on influential behavior of biopolishing on dyeability and certain physico-mechanical properties of cotton fabrics. Carbohydr Polym 76:1–7 Sethi S, Datta B, Gupta L, Gupta S (2013) Optimization of cellulose production from bacteria isolated from soil. ISRN Biotechnol 2013:1–7 Shida Y, Furukawa T, Ogasawara W (2016) Deciphering the molecular mechanisms behind cellulase production in Trichoderma reesei, the hyper-cellulolytic filamentous fungus. Biosci Biotechnol Biochem 80:1712–1729 Shokri J, Adibkia K (2013) Application of cellulose and cellulose derivatives in pharmaceutical industries. In: Cellulose-medical, pharmaceutical and electronic applications. IntechOpen, London, pp 47–66 Singhania R, Patel A, Soccol C, Pandey A (2009) Recent advances in solid-state fermentation. Biochem Eng J 44:13–18 Stern J, Kahn A, Vazana Y, Shamshoum M, Morais S, Lamed R, Bayer E (2015) Significance of relative position of cellulases in designer cellulosomes for optimized cellulolysis. PLoS ONE 10:e0127326 Sukumaran R, Singhania R, Pandey A (2005) Microbial celulases-production, application and challenges. J Sci Ind Res 64:832–844

40

2 Cellulase in Degradation of Lignocellulosic Wastes

Sun Y, Cheng J (2002) Hydrolysis of lignocellulosic materials for ethanol production: a review. Bioresour Technol 83:1–11 Taherzadeh M, Jeihanipour A (2012) Recalcitrance of lignocellulosic biomass to anaerobic digestion. In: Mudhoo A (ed) Biogas production: pretreatment methods in anaerobic digestion. Wiley, New York, pp 27–54 Takasuka T, Book A, Lewin G, Currie C, Fox B (2013) Aerobic deconstruction of cellulosic biomass by an insect-associated Streptomyces. Sci Rep 3:1–10 Ulrich A, Klimke G, Wirth S (2008) Diversity and activity of cellulose-decomposing bacteria, isolated from a sandy and a loamy soil after long-term manure application. Microb Ecol 55:512– 522 Vaaje-Kolstad G, Westereng B, Horn S, Liu Z, Zhai H, Sorlie M, Eijsink V (2010) An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science 330:219–222 Vilanova C, Marco G, Dominguez-Escriba L, Genoves S, Sentadreu V (2012) Bacteria from acidic to strongly alkaline insect midguts: Potential sources of extreme cellulolytic enzymes. Biomass Bioenergy 45:288–294 Wahlstrom R, Rahikainen J, Kruus K, Suurnakki A (2014) Cellulose hydrolysis and binding with Trichoderma reesei Cel5A and Cel7A and their core domains in ionic liquid solutions. Biotechnol Bioeng 111:726–733 Westereng B, Ishida T, Vaaje-Kolstad G, Wu M, Eijsink V (2012) The putative endoglucanase PcGH61D from Phanerochaete chrysosporiumis a metal-dependent oxidative enzyme that cleaves cellulose. PLoS ONE 6:1–11 Wilson D (2004) Studies of Thermobifida fusca plant cell wall degrading enzymes. Chem Rec 4:72–82 Wilson D (2008) Three microbial strategies for plant cell wall degradation. Ann N Y Acad Sci 1125:289–297 Wilson D (2012) Processive and nonprocessive cellulases for biofuel production—lessons from bacterial genomes and structural analysis. Appl Microbiol Biotechnol 93:497–502 Wilson D, Urbanowicz B (2017) Glycoside hydrolase family 9. In: Wilson D (ed). CAZypedia Xie G, Bruce D, Challacombe J, Chertkov O, Detter J, Gilna P, Han C, Lucas S, Misra M, Myers G et al (2007) Genome sequence of the cellulolytic gliding bacterium Cytophaga hutchinsonii. Appl Environ Microbiol 73:3536–3546 Xiros C, Topakas E, Christakopoulos P (2016) Hydrolysis and fermentation for cellulosic ethanol production. In: Lund D, Byrne J, Berndes G, Vasalos L (eds) Advances in bioenergy: the sustainability challenge. Wiley, New York, pp 11–31 Xu Q, Barak Y, Kenig R, Shoham Y, Bayer E, Lamed R (2004) A novel Acetivibrio cellulolyticus anchoring scaffoldin that bears divergent cohesins. J Bacteriol 186:5782–5789 Yang B, Dai Z, Ding S, Wyman C (2011) Enzymatic hydrolysis of cellulosic biomass. Biofuels 2:421–449 Yoshizawa S, Tanaka M, Shekhar A (2004) Global trends in waste generation. In: Gaballah I, Mishar B, Solozabal R, Tanaka M (eds) Recycling, Waste Treatment and Clean Technology. TMS Minerals, Metals and Materials Publishers , Spain, pp 1541–1552 Yuan J, Tiller K, Al-Ahmad H, Stewart N, Sttewart C (2008) Plants to power: bioenergy to fuel the future. Trends Plant Sci 13:421–429 Zhang Y (2011) Substrate channeling and enzyme complexes for biotechnological applications. Biotechnol Adv 29:715–725 Zhou F, Olman V, Xu Y (2009) Large-scale analyses of glycosylation in cellulases. Genomics Prot Bioinform 7:194–199 Zhuang J, Marchant M, Nokes S, Strobel H (2007) Economic analysis of cellulase production methods for bioethanol. Appl Eng Agric 23:679–687 Zifcakova L, Baldrian P (2012) Fungal polysaccharide monooxygenases: new players in the decomposition of cellulose. Fungal Ecol 5:481–489

Chapter 3

Hemicellulase in Degradation of Lignocellulosic Wastes

Abstract Hemicellulose is the major component in the cell wall of many plants. Degradation of hemicellulose is very difficult and needs to be solved. Hemicellulase is the enzyme in the degradation of hemicellulose. The chapter here describes in detail the structure and properties of hemicellulose; xylan degradation; families of hemicellulase enzyme; and hemicellulase production by microorganisms. The chapter also focuses on brown-rot, white-rot, and soft-rot fungi in degradation of hemicellulose; transcriptional regulators in regulation of xylanolytic gene expression; pretreatment of hemicellulose; hemicellulase enzymes; assays for hemicellulase enzyme activity; and applications of hemicellulase enzyme. The chapter gives detailed information of hemicellulase enzyme in degradation of hemicellulose. This will help in the proper management of hemicellulosic wastes. Keywords Hemicellulase · Xylan · Brown-rot fungi · Pretreatment · Polysaccharide · Monomers

3.1 Hemicellulose Structure and Property Hemicellulose is heterogeneous polysaccharide comprising of d-xylose, d-glucose, d-arabinose, d-galactose, d-mannose, d-4-O-methyl-glucuronic, d-galacturonic and d-glucuronic acid, and other ester-linked acetyl, coumaryl, and feruloyl moieties (Perez et al. 2002). Hemicellulose does not have packed crystalline structure and can undergo hydrolysis by enzymes (Saha 2003). Hemicelluloses are linked with cellulose and are found mainly in secondary wall of the cells. The hemicellulose in hardwoods and grasses is glucuronoxylan, and in softwoods is glucomannan. The sugars in hemicellulose are linked together with β-1,4 glycosidic bonds, and in some cases with β-1,3 glycosidic bonds (Perez e t al. 2002). Xylans are the most abundant hemicellulose (Mazeau et al. 2005). The structure of hemicellulose is represented in Fig. 3.1. Hemicellulose has hydrophilic property and is important component of the cell wall. Hemicelluloses are cross-linked and combine with the surface of the cellulose– microfibrils and are thus attached to the microfibrils. This forms hard fibers and interconnected networks of cells. © Springer Nature Switzerland AG 2020 A. B. Gunjal et al., Enzymes in Degradation of the Lignocellulosic Wastes, https://doi.org/10.1007/978-3-030-44671-0_3

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3 Hemicellulase in Degradation of Lignocellulosic Wastes

Fig. 3.1 Structure of hemicellulose

3.2 Xylan Degradation Degradation of xylan is by enzymes, viz. endoxylanases and endomannanases which hydrolyze backbone of xylans and glucomannan, respectively. The hydrolysis of xylans into monomers is by enzymes, viz. endo-1,4-β-xylanase, acetyl esterase, α-glucuronidase and β-xylosidase. The enzyme endo-1,4-β-xylanase forms xylan oligosaccharides and 1,4-β-xylosidase forms xylose. Tenkanen and co-workers stated that the enzymes from Trichoderma reesei synergistically will hydrolyze beech wood xylan. Therefore, the more efficient hydrolysis of xylan is by cooperative action of multiple isoenzyme systems. Xylanases are produced by many microorganisms and plants. Hemicellulose is difficult to degrade as compared to cellulose (Malherbe and Cloete 2002). Hemicellulose is biodegraded to sugars and acetic acid. Hemicellulose degradation also needs additional enzymes, viz. xylan esterases, ferulic and p-coumaricesterases, α-1-arabinofuranosidases, and α-4-O-methyl glucuronosidases, acting synergistically to efficiently hydrolyze wood xylans and mannans (Perez et al. 2002).

3.3 Families of Enzyme Hemicellulase

43

3.3 Families of Enzyme Hemicellulase Enzymes with hemicellulase activity are classified into glycosyl hydrolase (GH) families, viz. GH2, GH10, GH11, GH16, GH26, GH30, GH31, GH39, GH42, and GH43. The GH43 family is the largest among these in the carbohydrate-active enzymes (CAZy), with 4555 members (Jimenez et al. 2015). This family has many debranching enzymes which helps in the degradation of hemicellulose, specifically arabinoxylans and also helps in degradation of biopolymer. The CAZy family is specifically related to the degradation of hemicellulose.

3.4 Hemicellulase Production by Microorganisms The bacteria producing hemicellulase are Bacillus sp., Cellulomonas, Micrococcus, Staphylococcus, Thermotoga, Paenibacillus, Arthrobacter, Pseudoxanthomonas, and Rhodococcus (Shahi et al. 2016). The fungi producing hemicellulase are Fusarium, Cladosporium sp. (Del-Cid et al. 2014), Penicillium thomii (Palaniswamy et al. 2008), P. pinophilum (Lee et al. 2011; Li et al. 2006), A. niger (Sharma et al. 2015), Ceratocystis paradoxa, A. niger (Raghukumar et al. 2004), P. canescens (Burtseva et al. 2010), Alternaria alternata (Wipusaree et al. 2011), Geotrichum, Paecilomyces, Cephalosporium, and Trichoderma. Among the actinomycetes, the production of hemicellulase enzyme is by T. bifida, Cellulomonas fimi, Thermomonospora fusca, Actinomadura, Streptomyces flavogriseus (Saini et al. 2015), Thermonospora curvata, Thermonospora alba, Micromonospora, Microbispora bispora, Nocardia, Saccharomonospora viridis, Thermoactinomyces, Streptomyces lividans, Streptomyces violaceoruber, and Streptomyces aureofaciens (Saini et al. 2015). The other actinomycetes producing hemicellulase are Microtetraspora flexuosa, Streptomyces violaceoruber, Thermoactinomyces thalophilus, and Streptomyces thermocyanaeviolaceus.

3.5 Brown-Rot, White-Rot and Soft-Rot Fungi in Degradation of Hemicellulose The fungi in degradation of hemicellulose are divided into three categories: brownrot, white-rot and soft-rot fungi (Yelle et al. 2008). Brown-rot fungi breakdown hemicellulose present in the wood. The examples of brown-rot fungi are Gloeophyllum trabeum, Fomitopsis lilacino-gilva, Laetiporus portenosus, Postia placenta and Serpula lacrymans (Martinez et al. 2005; Wong 2009). The white-rot fungi are Ceriporiopsis subvermispora, Phlebia radiata, Rigidoporous lignosus, Dichornitus sqiualenis, Phellinuis pini and Inonotits diyophillis (Kantharaj et al. 2017).

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Xylanases is the major component of hemicellulases. The white-rot fungi Phanerochaete chrysosporium produces endoxylanases. The soft-rot fungi are also called as microfungi and degrade hemicellulose. The xylanase producing microorganisms are Cladosporium sp., P. thomii, P. pinophilum, A. niger, C. paradoxa, A. niger, P. canescens, etc. (Kantharaj et al. 2017).

3.6 Transcriptional Regulators Involved in Regulation of Xylanolytic Gene Expression (T. Reesei) (Shida et al. 2016) The transcriptional regulators in regulation of xylanolytic gene expression are the same as those involved in regulation of cellulolytic gene expression. These include xylanase regulator (Xyr1); activator of cellulase expression 2 (Ace2); activator of cellulase expression 1 (Ace1); beta-glucosidase regulator (BglR); and activator of cellulase expression 3 (Ace3).

3.7 Methods for Pretreatment of Hemicellulose 3.7.1 Alkaline Pretreatment Alkali pretreatment involves the use of bases, viz. sodium, potassium, calcium, and ammonium hydroxide, for the pretreatment of hemicellulose. The use of an alkali causes partial solvation of hemicellulose (McIntosh and Vancov 2010; Sills and Gossett 2011). Sodium hydroxide (Zhu et al. 2010) and lime are the alkali agents used for pretreatment of hemicellulose. The effectiveness of different alkaline solutions for pretreatment of hemicellulose in wheat straw has been studied. Zhao and co-workers (Zhao et al. 2008) showed the effectiveness of sodium hydroxide pretreatment for pretreatment of hemicellulose. The advantage of lime pretreatment is that it is very cheap.

3.7.2 Wet Oxidation Wet oxidation can be combined with other pretreatment methods, viz. steam explosion. The wet oxidation when combined with steam explosion is called wet explosion. This combination allows processing large particle sizes and operating at higher substrate loadings (Georgieva et al. 2008). This method has been found to be effective in pretreating biomass, viz. wheat straw, corn stover, sugarcane bagasse, cassava, peanuts, rye, canola, and fava beans.

3.7 Methods for Pretreatment of Hemicellulose

45

3.7.3 Acid Pretreatment Acid pretreatment involves the use of concentrated and diluted acids to break the rigid structure of hemicellulose. The commonly used acid is dilute sulfuric acid (H2 SO4 ). Other acids used for pretreatment of hemicellulose are hydrochloric (HCl), phosphoric (H3 PO4 ), nitric (HNO3 ), and triflulororacetic (TFA) acid. By acid pretreatment, hemicellulose is converted to sugars.

3.7.4 Green Solvents Many green solvents are also used for pretreatment of hemicellulose.

3.7.5 Steam-Explosion Pretreatment Steam-explosion pretreatment is also used for the pretreatment of hemicellulose, which uses both chemical and physical methods. The steam-explosion method subjects the material to high pressures and temperatures for a short time and then depressurizes the system which disrupts the structure of the fibrils of hemicellulose. The steam explosion in combination with acid catalysts helps improve hemicellulose hydrolysis. Dilute acids minimize retention time and temperature of operating systems. This helps in complete hydrolysis of hemicellulose. There are two steps in this process. The first is chemical pretreatment in the form of hydrolysis of the glycosidic bonds present in hemicellulose, which breaks the acetyl groups into acetic acid and helps in hemicellulose hydrolysis (Brodeur et al. 2011). The second is physical pretreatment which is rapid decompression of the system.

3.8 Enzymes for Hemicellulose Biodegradation The enzymes for hemicellulose biodegradation are xylan esterases, ferulic and pcoumaric esterases, α-1-arabinofuranosidases and α-4-O-methyl glucuronosidases. Xylan requires combination of hydrolytic enzymes for degradation. Endo-1,4β-xylanase (EC 3.2.1.8) produces oligosaccharides, and xylan 1,4-β-xylosidase produces xylose. The enzymes for hemicellulose degradation are shown in Fig. 3.2a, b. The hemicellulases and their EC number are represented in Table 3.1. The fungi which degrade hemicelluloses in soil are Chaetomium, Aspergillus, Penicillium, Trichoderma, Fusarium, Humicola, etc. The fungal enzymatic complex

Xylan

b

xylosidase

1,4 beta

acetyl esterase

esterases

α-1-arbionfuranosidase

methylglucrosidasse

α-1-O

Endomananases

O-acetylgalactoglucomanann

xylose and mannose

esterases

ferulic

β-xylosidase

p-coumaric

α-glucuronidase

esterases

xylan

Xylan and mannan

Fig. 3.2 a Enzymes in degradation of hemicellulose. b Enzymes in degradation of hemicellulose

endoxylanase

O-acetyl-4-O-methyl glucuronoxylan

xylose

xylanase

endo 1,4-beta-

a

46 3 Hemicellulase in Degradation of Lignocellulosic Wastes

3.8 Enzymes for Hemicellulose Biodegradation

47

Table 3.1 Hemicellulases and their EC number EC number

Recommended name

Systematic name

3.2.1.8

Endo-1,4-β-xylanase

1,4-β-d-Xylan xylanohydrolase

3.2.1.37

Xylan 1,4-β-d-xylosidase

1,4-β-d-Xylan xylohydrolase

3.2.1.32

Xylan endo-1,3-β-xylosidase

1,3-β-d-Xylan xylanohydrolase

3.2.1.72

Xylan 1,3-β-xylosidase

1,3-β-d-Xylan xylohydrolase

3.2.1.55

α-1-Arabinofuranosidase

α-l-Arabinofuranoside arabinofuranosidase

3.2.1.99

Arabinan endo-1,5-α-1-arabinosidase

1,5-α-l-Arabinan 1,5-α-l-arabinohydrolase

3.2.1.78

Mannan endo-1,4-β-mannosidase

1,4-β-d-Mannan mannanohydrolase

3.2.1.100

Mannan 1,4-β-mannobiosidase

1,4-β-d-Mannan mannobiohydrolase

3.2.1.101

Mannan endo-1,6-β-mannosidase

1,6-β-d-Mannan mannanhydrolase

Table 3.2 Fungal enzymatic complex for hemicellulose degradation (Rodrigo de Souza 2013) Hemicellulose polymer

Enzyme

Residue released

Xyloglucan/Xylan

α-arabinofuranosidases

l-arabinose

Xyloglucan

α-xylosidases

d-xylose

Xyloglucan

α-fucosidases

l-fucose

Xylan/galactomannans

α-galactosidases

d-galactose

Xylan

Feruloyl esterases

Ferulic acid

for hemicellulose degradation is shown in Table 3.2. The bacteria-degrading hemicelluloses in soil are Bacillus, Pseudomonas, Cytophaga, Vibrio, Erwinia, Streptomyces, etc.

3.8.1 Hemicellulase Enzymes 3.8.1.1

1,4,-β-D Xylanases

This hydrolyzes 1,4-β-d-xylopyranosyl linkages of d-xylans such as l-arabino-dglucrono-d-xylans and d-glucrono-d-xylans.

3.8.1.2

1,4-β-D-Mannanases

This enzyme hydrolyzes the 1,4-β-d-mannopyranosyl linkages of branched mannans (e.g., d-galacto-d-mannans), copolymer mannans (e.g., d-gluco- and d-galacto-dgluco-d-mannans) and linear d-mannans.

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3 Hemicellulase in Degradation of Lignocellulosic Wastes

1. Exo-1,4-β-d-Mannanases This enzyme degrades its substrate from non-reducing end of the molecule. 2. Endo-1,4-β-d-Mannanases Endo-β-mannanases degrade β-d-mannans to d-mannose and mannose oligosaccharides.

3.8.1.3

β-D-Galactanases

This enzyme degrades d-galactans and l-arabino-d-galactans. Endogalactanasesdegrading 1,4,-β-d-galactosyl linkages of d-galactans are called endo-1,4-βgalactanases and form d-galactose and galactose oligosaccharides. Endogalactanases attack 1,3-β-d-linked galactosyl bonds of arabinogalactans to give d-galactose and 1,3- and 1,6-linked β-d-galactose oligosaccharides. The main positive regulator of hemicellulase gene expression is xylanase regulator 1 (Xyr1) (Stricker et al. 2006, 2008), a zinc binuclear cluster protein binding to a GGCTAA-motif (Stricker et al. 2008). The hemicellulosic enzymes and their function in hemicellulose degradation are represented in Table 3.3. Table 3.3 Hemicellulosic enzymes and their function in hemicellulose degradation (Coughlan et al. 1993) Enzyme

Function

Endo-acting Endoarabinanase

Breakage of α-1,5-linked backbone of arabinan

Endogalactanase

Breakage of α-1,4- and α-1,3-linked backbone of arabinogalactans

Exo-acting α-l-Arabinosidase

Releases 1,3- or 1,2-α-l-arabinosyl from arabinoxylans and α-1,2 or 1,3-linked l-arabinose from arabinosides, arabinoxylans and xylo-oligomers

β-l-Arabinosidase

Releases β-linked l-arabinopyranosyl side chains of arabinogalactans

Exoxylanase

Releases xylose from non-reducing end of β-1,4-linked xylans

β-d-Galactosidase

Releases β-1,6-linked d-galactopyranosyl from side chains of arabinogalactans and galactoglucomannan

α-d-Glucuronidase

Releases α-1,2-linked d-glucopyranosuronic acid or 4-O-methylglucopyranosuronic acid from side chains of substituted oligomers released from glucuronoxylans

3.9 Method for Measurement of Hemicellulase Enzyme Activities

49

3.9 Method for Measurement of Hemicellulase Enzyme Activities The measurement of the hemicellulase enzymes, viz. laminarinase [endo-β-1,3glucanase (EC 3.2.1.39)], licheninase [endo-β-1,3; 1,4 glucanase (EC 3.2.1.73)], xylanase [endo-β-1,4-xylanase (EC 3.2.1.8), and 1,4-β-d-xylan xylanhydrolase (EC 3.2.1.37)] is given below.

3.9.1 Laminarinase (Linton and Greenaway 2004) Laminarinase activity is measured based on reducing sugars from the hydrolysis of laminarin. Sample 20 μl) + 50 μl of 1% (w/v) laminarin + 130 μl of 0.1 mol l−1 Na acetate buffer (pH 5.5).

Blank and sample are also run and incubated with agitation at 40 °C for 10 min.

Add 50 μl of 0.3 mol l−1 HCl to stop the reaction and neutralize with 10 μl of 2.5 mol l−1 K2 CO3 and measure the reducing sugars in a 10 μl aliquot.

3.9.2 Licheninase (Linton and Greenaway 2004) Activity of licheninase is measured by the reducing sugars produced from hydrolysis of lichenin.

Sample (20μ l) + 100 μl of 0.1% (w/v) lichenin + 80 μl of 0.1 mol l−1 Na acetate buffer (pH 5.5).

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3 Hemicellulase in Degradation of Lignocellulosic Wastes

Blank and standard are also run and incubated with agitation at 40 °C for 10 min.

Add 50 μl of 0.3 mol l−1 HCl to stop the reaction. Neutralize the mixture with 10 μl of 2.5 mol l−1 K2 CO3 .

Measure the reducing sugars in a 10 μl aliquot.

3.9.3 Xylanase (Linton and Greenaway 2004) Xylanase activity is measured based on the production of reducing sugars from the hydrolysis of xylan.

Sample (20 μl) incubate with 100 μl of 1% (w/v) xylan + 80 μl of 0.1 mol l−1 Na acetate buffer (pH 5.5).

Blank and standard are also run and incubated with agitation at 40 °C for 60 min.

Precipitate the protein with 50 μl of 0.3 mol l−1 HCl to stop the reaction and neutralize by the addition of 10 μl of 2.5 mol l−1 K2 CO3 .

3.9 Method for Measurement of Hemicellulase Enzyme Activities

51

Measure the reducing sugars in a 10 μl aliquot.

3.10 Applications of Hemicellulase Enzyme 3.10.1 In Paper and Pulp Industries Hemicellulase enzyme has application in the paper and pulp industry.

3.10.2 Processing of Animal Feed Arabinoxylans are partly water-soluble polysaccharides present in cereals like barley which is high viscous solution, and this makes it important in animal feeding. Xylanases are used for the pretreatment of arabinoxylan containing substrates. This improved the poultry feed quality (Babalola et al. 2006).

3.10.3 Beverage Industry The hemicellulose enzyme also has application in clarification of juices and wines. Hemicellulase enzyme has improved the liquefaction of fruits; recovery of essentials oils and vitamins, mineral salts, dyes, pigments etc.; reduced the viscosity and improved the hydrolysis of substances which affect the physicochemical properties (Polizeli 2005).

3.10.4 Bakery Industry Hemicellulases are used as dough strengtheners and to enhance the flour quality. The hemicellulase enzyme also increases volume of the baked bread (Camachom and Aguilar 2003).

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3 Hemicellulase in Degradation of Lignocellulosic Wastes

3.10.5 Pharmaceutical Industry The hemicellulase enzyme also has application in pharmaceutical industries.

References Babalola T, Apata D, Atteh J (2006) Effect of β-xylanase supplementation of boiled castor seed meal-based diets on the performance, nutrient absorbability and some blood constituents of pullet chicks. Trop Sci 46:216–223 Brodeur G, Yau E, Badal K, Collier J (2011) Chemical and physicochemical pretreatment of lignocellulosic biomass: a review. Enzyme Res 2011:1–17 Burtseva Y, Sova V, Pivkin M, Anastyuk S, Gorbach V, Zvyagintseva T (2010) Distribution of Oglycosylhydrolases in marine fungi of the sea of Japan and the sea of Okhotsk: characterization of exo-cellular N-acetyl-beta-D-glucosaminidase of the marine fungus Penicillium canescens. Appl Biochem Biotechnol 46:648–656 Camachom N, Aguilar O (2003) Production, purification, and characterization of a low-molecularmass xylanase from Aspergillus sp. and its application in baking. Appl Biochem Biotechnol 104:159–172 Coughlan M, Touhy M, Filho E, Puls J, Claeyssens M, Vrsanska M, Hughes M (1993) Enzymological aspects of microbial hemicellulases with emphasis on fungal systems. In: Coughlan M, Hazlewood G (eds) Hemicellulose and hemicellulases. Portland Press Research Monograph, London Del-Cid A, Ubilla P, Ravanal MC, Medina E, Vaca I, Levican G, Eyzaguirre J, Chavez R (2014) Cold-active xylanase produced by fungi associated with Antarctic marine sponges. Appl Biochem Biotechnol 172:524–532 Georgieva T, Mikkelsen M, Ahring B (2008) Ethanol production from wet-exploded wheat straw hydrolysate by thermophilic anaerobic bacterium Thermoanaerobacter BG1L1 in a continuous immobilized reactor. Appl Biochem Biotechnol 145:99–110 Jimenez D, Chaves-Moreno D, van Elsas J (2015) Unveiling the metabolic potential of two soilderived microbial consortia selected on wheat straw. Sci Rep 5:13845 Kantharaj P, Boobalan B, Sooriamuthu S, Mani R (2017) Lignocellulose degrading enzymes from fungi and their industrial applications. Int J Cur Res Rev 9:1–12 Lee J, Jang Y, Lee H, Lee S, Kim G, Kim J (2011) Screening for xylanase and β-xylosidase production from wood-inhabiting Penicillium strains for potential use in biotechnological applications. Holzforschung 66:267–271 Li Y, Liu Z, Cui F, Xu Y, Zhao H (2006) Production of xylanase from a newly isolated Penicillium sp. ZH-30. World J Microbiol Biotechnol 23:837–843 Linton S, Greenaway P (2004) Presence and properties of cellulase and hemicellulase enzymes of the gecarcinid land crabs Gecarcoidea natalis and Discoplax hirtipes. J Exper Biol 207:4095–4104 Malherbe S, Cloete T (2002) Lignocellulose biodegradation: fundamentals and applications. Rev Environ Sci Biotechnol 1:105–114 Martinez A, Speranza M, Ruiz-Duenas F, Ferreira P, Camarero S, Guillen F, Martinez M, Gutierrez A, del Rio JC (2005) Biodegradation of lignocellulosics: microbial, chemical and enzymatic aspects of fungal attack to lignin. Inter Microbiol 8:195–204 Mazeau K, Moine C, Krausz P, Gloaguen V (2005) Conformational analysis of xylan chains. Carbohydr Res 340:2752–2760 McIntosh S, Vancov T (2010) Enhanced enzyme saccharification of Sorghum bicolor straw using dilute alkali pretreatment. Bioresour Technol 101:6718–6727 Palaniswamy M, Pradeep B, Sathya R, Angayarkanni J (2008) Isolation, identification and screening of potential xylanolytic enzyme from little degrading fungi. Afr J Biotechnol 7:1978–1982

References

53

Perez J, Munoz-Dorado J, de la Rubia T, Martınez J (2002) Biodegradation and biological treatments of cellulose, hemicellulose and lignin: an overview. Int Microbiol 5:53–63 Polizeli M, Rizzatti A, Monti R, Terenz H, Jorge J, Amorim D (2005) Xylananses from fungi: properties and industrial applications. Appl Microbiol Biotechnol 67:577–591 Raghukumar C, Muraleedharan U, Gaud V, Mishra R (2004) Xylanases of marine fungi of potential use for biobleaching of paper pulp. J Ind Microbiol Biotechnol 31:433–441 Rodrigo de Souza W (2013) Microbial degradation of lignocellulosic biomass. In: Chandel A, Silverio da Silva S (eds) Sustainable degradation of lignocellulosic biomass-techniques, applications and commercialization. In-Tech Publishers, Rijeka, pp 207–248 Saha B (2003) Hemicellulose bioconversion. J Ind Microbiol Biotechnol 30:279–291 Saini A, Aggarwal N, Sharma A, Yadav A (2015) Actinomycetes: a source of lignocellulolytic enzymes. Enzyme Res 2015:1–15 Shahi N, Hasan A, Akhtar S, Siddiqui M, Sayeed U, Khan M (2016) Xylanase: a promising enzyme. J Chem Pharma Res 8:334–339 Sharma S, Vaid S, Bajaj B (2015) Screening of thermo-alkali stable fungal xylanases for potential industrial applications. Curr Res Microbiol Biotechnol 3:536–541 Shida Y, Furukawa T, Ogasawara (2016) Deciphering the molecular mechanisms behind cellulase production in Trichoderma ressei, the hyper-cellulolytic filamentous fungus. Biosci Biotechnol Biochem 80:1712–1729 Sills D, Gossett J (2011) Assessment of commercial hemicellulases for saccharification of alkaline pretreated perennial biomass. Bioresour Technol 102:1389–1398 Stricker A, Grosstessner-Hain K, Wurleitner E, Mach R (2006) Xyr1 (xylanase regulator 1) regulates both the hydrolytic enzyme system and D-xylose metabolism in Hypocrea jecorina. Eukaryot Cell 5:2128–2137 Stricker A, Mach R, de Graff L (2008) Regulation of transcription of cellulases- and hemicellulasesencoding genes in Aspergillus niger and Hypocrea jecorina (Trichoderma reesei). Appl Microbiol Biotechnol 78:211–220 Wipusaree N, Sihanonth P, Piapukiew J, Sangvanich P, Karnchanatat A (2011) Purification and characterization of a xylanase from the endophytic fungus Alternaria alternate isolated from the Thai medicinal plant, Croton oblongifolius Roxb. Afr J Microbiol Res 5:5697–5712 Wong D (2009) Structure and action mechanism of lignolytic enzymes. Appl Biochem Biotechnol 157:174–209 Yelle D, Ralph J, Lu F, Hammel K (2008) Evidence for cleavage of lignin by a brown-rot basidiomycete. Environ Microbiol 10:1844–1849 Zhao Y, Wang Y, Zhu J, Ragauskas A, Deng Y (2008) Enhanced enzymatic hydrolysis of spruce by alkaline pretreatment at low temperature. Biotechnol Bioengi 99:1320–1328 Zhu J, Wan C, Li Y (2010) Enhanced solid-state anaerobic digestion of corn stover by alkaline pretreatment. Bioresour Technol 101:7523–7528

Chapter 4

Ligninase in Degradation of Lignocellulosic Wastes

Abstract Agro-industrial waste industries are the largest polluting industries in the world with the potential application of biofuels or biosources. From the past several years, the worldwide economic and environmental pollution issues have been escalating research interest in the value of biosourced lignocellulosic biomass. The limited resource of fossil fuels and the rapid increase in energy demand has placed immense on the environment. Microbes are known to produce lignin modifying enzymes with high activity and specificity. Four enzymes namely lignin peroxidase, manganese peroxidase, versatile peroxidase, and Laccase are effective in lignin degradation. Dye decoloration is also reported activity together with it. Several fungal and bacterial species are discussed that may enhance production or lignin degradation. This chapter provides a general overview of the suitability of lignin-modifying enzymes used for different agro-industrial wastes and also deals with the use of these enzymes in the development of economic biocatalysts that are more sustainable. This may reduce harmful environmental impacts and improve the applications of enzymatic technology in the industry. Keywords White-rot fungi · Laccasse · Lignin peroxidases · Manganese peroxidases · Recalcitrant chemicals

4.1 Introduction With the rapid increase in the energy demand, the conventional resources mainly fossil fuels are becoming limited. This imbalance in demand to supply ratio has placed immense pressure on the environment indicating a prompt action for sustainable energy resources (Fernando et al. 2006). A techno-economic model was thus designed that was based on the assessment of the potential use of biofuels. This model provided invaluable guidance to research, investment and policy endeavors for value-added products. This study relied on experimentally derived or assumed parameters to estimate process performance which includes capital and investing cost, greenhouse gas emission, and biofuel yield. It aids in building a program that includes all of the unit operations and process flows needed as an input (corn, process water, etc.) into outputs like ethanol, CO2 , and electricity © Springer Nature Switzerland AG 2020 A. B. Gunjal et al., Enzymes in Degradation of the Lignocellulosic Wastes, https://doi.org/10.1007/978-3-030-44671-0_4

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(Klein-Marcuschamer et al. 2010). Lignocellulosic materials can be used as an alternative to fossil fuels partly due to the abundance and renewable nature of the material. Agricultural waste (wheat straw, grass hay) municipal solid waste or forestry wastes are rich in lignocellulosic material that can be used as low-cost feedstock alternatives. Increased biomass generation calls an ecological monitoring action. Industries releases in the form of recalcitrant compounds are difficult to degrade. The use of physical and chemical methods produced excess sludge and harmful chemicals that are not easily degradable. The biological mean of degradation is the use of microbes as whole or enzymes secreted for the treatment of aromatic compounds has gathered attention in recent years because of their ecofriendly nature. Various enzymes are employed for degrading ligninolytic waste namely termed as lignin-modifying enzymes (LME) that constitutes laccases and peroxidases (lignin peroxidase (LiP), manganese peroxidase (MnP), and versatile peroxidase (VP) and are considered as green catalysts of vast biotechnological importance.

4.2 Lignin Occurrence, Biogenesis, and Biodegradation Lignins are plant polymers derived from the hydroxycinnamyl alcohols or monolignols p-coumaryl, coniferyl, and sinapyl. It was a debatable question on what are lignins? Histochemical studies investigated on lignin as a plant polymer. Total lignin contents are determined by solvent extraction, alkali extraction, and more recent spectroscopic methods including NMR, infrared, and UV methods. Water-soluble and water-insoluble intermediate lignins degradation can be detected under solid-state and submerged fermentation. The intermediate can be extracted with polar solvent. Degradation of lignins to generate biofuels strongly depends on lignin’s chemical structure and fractionation process. These materials are decomposed in nature by several microorganisms like fungi and bacteria. Lignocellulosic biomass decomposition is carried out primarily by microbial involvement, utilizing it as carbon and nutrient/energy source for their growth. Lignocellulosic hydrolysis yield simple sugars that can be utilized for biofuel production. Microbes are known to break down the lignin and cellulose to utilize it as energy source. Few fungi and certain bacteria can also decolorize the cellulose. Lignin is a complex molecule with a non-repeating phenylpropane backbone biosynthesized by random polymerization of phenylpropanoid precursor with irregular structure. The two-dimensional structure of lignin is represented in Fig. 4.1. The major building blocks are monolignols or hydroxycinnamyl alcohols (Vanholme et al. 2010). Polymerization can be carried out in vitro to produce synthetic lignin or dehydrogenate polymerizes (DHP). The non-repeating structure is an important feature for biological delignification. Diversity of lignins is observed in the interunits making it difficult to beak the interunit bonds, complicating its use as a carbon and energy source for biological degradation. Degradation of lignins is only possible with few organisms that harbor the pathway to degrade it. The derivatives of lignin are shown is Fig. 4.2.

4.2 Lignin Occurrence, Biogenesis, and Biodegradation

57

Fig. 4.1 Two-dimensional structure of lignin polymer molecule

Fig. 4.2 Derivatives of lignin

White-rot basidiomycetes ( Hatakka et al. 2003; Leonowicz et al. 1999; Martinez et al. 2004; Nerud and Mišurcová 1996), actinomycetes, Trichoderma, Phanerochaete chrysosporium (Butler and Day 1998; Dosoretz et al. 1990; Venkatadri and Irvine 1990; Whitaker et al. 2017), Lentinus squarrosulus (Wuyep et al. 2003), Streptomyces cinnamomensis (Jing and Wang 2012), Xanthomonas sp. mixed white-rot fungi Schizophyllum commune, Bjerkandera adusta, and Fomitopsis palustris (Horisawa et al. 2019), Trichoderma (Vázquez et al. 2019), several bacterial and yeast species isolated from mangrove microflora (Philip et al. 2019) and Aspergillus are efficient lignin degraders (Bechem and Etaka 2018). Several fungal species are used in commercial purpose to decompose ligninolytic biomass. P. chrysosporium is studied in great detail and is a working model organism for studying lignin degradation. The genome sequence is annotated offering further insights on lignin degradative machinery and analogous genomic organization. It was observed that ten lignin peroxidases, five manganese peroxidases, and several other lignocellulolytic enzymes were encoded in its genome (Martinez et al. 2004). Several reports on P. chrysosporium which is an excellent decomposer of soft- and

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hardwood, branches, logs, leaves, etc., in forests suggest its commercial use. Chondrostereum purpureum, a plant fungal pathogen, produces an extensive catalog of lignocellulolytic enzymes (Reina et al. 2019).

4.3 Mechanisms of Lignin Degradation Fungi degrade lignin by secreting enzymes collectively termed ligninases or ligninmodifying enzymes which are classified into two categories: the heme peroxidases that include lignin peroxidases, manganese peroxidases, versatile peroxidases, and dye decolorizing peroxidases; the other group is laccases (phenol oxidases). The peroxidases are heme-containing enzymes with catalytic cycles that are activated by peroxide, accessory oxidases also generate hydrogen peroxide required for peroxidase activity, whereas laccases are multicopper oxidoreductases, an equally important enzyme required for degrading lignin completely. The major lignin-degrading enzymes are heme-containing and H2 O2 -dependent lignin peroxidase (LiP), manganese peroxidase (MnP), versatile peroxidases, and Cu-containing laccase. The lignin degradation by four different enzymes is shown in Fig. 4.3. This chapter will deal with the understanding of all the four enzymes involved in the degradation.

Fig. 4.3 Lignin degradation by four different enzymes

4.4 Enzymes Involved in the Degradation

59

4.4 Enzymes Involved in the Degradation 4.4.1 Lignin Peroxidase (LiP) (EC 1.11.1.14) Lignin peroxidase is secreted as an extracellular enzyme first time reported in 1982 in white-rot fungus P. chrysosporium as a secondary metabolite (Linko 1992). It is a glycoprotein with a molecular weight of approximately 41,000 that contains 1 mol of Fe protoporphyrin IX and exists as a series of isozymes (pI—3.2–4.0). Peroxidases are heme-containing enzymes that use hydrogen peroxide to oxidize a wide variety of organic and inorganic compounds. Several reviews on lignin peroxidase discusses on high redox potential and low optimum pH activity. It has low substrate specificity and reacts with a wide variety of related and unrelated lignin molecule. The highest lignin peroxidase activity was observed in solid state fermentation when incubated with Pleurotus ostreatus till the seventeenth day in dry potato peel waste, pretreated with distilled water, whereas cadmium stress in the solid-state fermentation reduced LiP activity (Zhao et al. 2015). The higher LiP activity was obtained by Ganoderma lucidum using pineapple leaves as a substrate (Hariharan and Nambisan 2013).

4.4.1.1

Production of Lignin Peroxidases

Several organisms produce enzymes mentioned above to degrade the lignin completely. White-rot basidiomycetes fungi secrete extracellular ligninase peroxidase (LiP) and manganese peroxidase (MnP) along with H2 O2 . The enzyme production in P. chrysosporium depends on several factors such as the presence of veratryl alcohol and surfactants, especially Tween 80 (Glumoff et al. 1990; Lewis and Yamamoto 1990). For optimum production of the enzyme it is important to obtain it in purified form. Liquid culture medium supplemented with glucose, ammonium tartrate, Mn2+ , Tween 80, and veratry alcohol together with the kirk medium recipe was selected and studied for its effects on enzyme production. Different concentration of carbon source affects the enzyme production. P. chrysosporium (ATCC 20696) when incubated for 7 days with 2, 5, and 10 g/lit glucose concentration was observed that 10 g/lit of glucose yielded higher enzyme production than at 2 and 5 g/lit (Wang et al. 2008), whereas P. chrysosporium (ATCC 24725), (ATCC 34541) and ME-446 showed lower enzyme production when grown at different glucose concentration (Kapich et al. 2004; Pazarlioglu et al. 2005; Xiong et al. 2008). Different nitrogen sources can also be optimized for higher enzyme activity. Ammonium tartarate in P. chrysosporium (ATCC 20696) was effective for higher enzyme activity (Wang et al. 2008). Inducer like surfactant Tween 20, Tween 80, polyoxyethylene oleate (Lestan et al. 1990), and Triton X-100 stimulated the production of LiP (Lestan et al. 1990; Wang et al. 2008). When Tween 80 was supplemented with 0–4.5 g/lit showed enhanced production of LiP as compared to control, indicating the importance of surfactant in LiP production (Wang et al. 2008). Surfactants are known to stimulate the growth of the spores and increase the bioavailability of non-polar substrate

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(Lestan et al. 1990; Usha et al. 2014; Zheng et al. 2019). Tween 80 was observed to alter the cell membrane structure and promote the permeation of lignolytic enzyme from the cell into the medium. Ligninase activity was also observed to increase when the media was supplemented with phospholipid like soybean asolectin and fatty acids. Addition of veratryl alcohol in P. chrysosporium culture was also observed to enhance the production of ligninolytic enzymes; the optimum however should not increase higher than 0.3 g/lit.

4.4.1.2

Degradation of Ligninocellulolytic Material

Lignins are difficult to degrade by several microbes; however, certain fungal species when deprived with nitrogen, sulfur, and carbon source are able to catalyze the oxidation of lignin to CO2 . It was also observed ligninolytic enzyme system of P. chrysosporium is formed under nitrogen starvation in the absence of lignin. Addition of veratryl alcohol to lignin peroxidases creates veratryl alcohol cation radical that attacks certain group of lignins. The enzyme catalyzes H2 O2 -dependant oxidation of a non-phenolic lignin compounds via the initial formation of aryl cation radical as the initial substrate. This reaction exhibits a stoichiometry of 1 H2 O2 consumed per aldehyde formed, suggested by steady-state kinetics study. One unit of activity is defined as 1 micromole of veratryl alcohol oxidized in 1 min, and the activities are reported in units/lit. The degradation can be achieved by immobilizing either the fungi or the enzyme with polystyrene carrier with a suitable surfactant and initiator (Wang and Ruckenstein 1994). Several fungi are studied to modify lignin and different range of pollutants by secreting extracellular ligninolytic systems. Of the ligninolytic system studied, purified forms of LiP have been found to directly oxidize xenobiotic compounds like aromatic hydrocarbons, chlorophenols, and dyes. LiP activity was stimulated from the protective activity of tryptophan against H2 O2 (Collins et al. 1997). LiPs are studied to oxidize lignin and generate several secreted metabolites like veratryl alcohol. Effective inducer veratryl alcohol is a secondary metabolite produced by ligninolytic cultures of white-rot fungi. LiP is oxidized by H2 O2 to form a two electron intermediates, a compound that oxidizes substrates by one electron, forming the more reduced enzyme intermediate. Another compound then oxidizes substrates leaving the enzyme to the ferric state. The nucleotide sequence of cDNA LiP shows that the proximal and distal histidines are conserved. Lignin peroxidases (LiP) has an unusually low pH optimum, 3.0 as an increase in pH may denature the activity of the enzyme. The enzyme can be separation and purification can be achieved by Fast Protein Liquid Chromatography (FPLC) that involves immobilization of the enzyme and depolymerization of the substrate.

4.4 Enzymes Involved in the Degradation

4.4.1.3

61

Assay for Lignin Peroxidases (LP)

Several LiP assay are mentioned in the literature every method has its own strength and limitations. In one of the method, lignin peroxidases are detected by oxidation of veratryl alcohol to veratraldehyde. The detection is based on UV spectrophotometer at 310 nm. Veratryl alcohol is the substrate for estimating lignin peroxidases. However, study has been done on comparison of Azure B and veratryl alcohol to determine the accurate method to estimate lignin peroxidase (Arora and Gill 2001). As veratryl alcohol is known to be contaminated with methyl vanillate and the assay is sensitive to turbidity. Azure B assay employ the accurate method of estimating lignin peroxidase (Archibald 1992; Arora and Gill 2001). The Azure B lignin peroxidases assay contains Azure B dye, H2 O2 , and sodium tartrate buffer (pH 4.5). The activity is monitored at 615 nm wavelength. Low levels of LP can be detected and does not react with non-LP-mediated substrate (Arora and Gill 2001; Canales et al. 1998). The assay for identification of peroxidase is represented in Table 4.1.

4.4.2 Manganese Peroxidase (MnP) (EC 1.11.1.13) Manganese peroxidase enzyme belongs to the family of oxidoreductases, and the heme group of the enzyme can oxidize Mn2+ to Mn3+ . The enzyme acts on both the phenolic and non phenolic group of lignin (Brink et al. 2019; Ralph et al. 2019). It also known to oxidizes NADPH, glutathione, dithiothreitol, and dihydroxymaleic acid to produce hydrogen peroxide. Several basidiomycetes species are known to be responsible for the formation of Mn3+ . Stability of the enzyme was determined at a pH of 2 in Ceriporiopsis subvermispora. The acid stable nature of the enzyme was confirmed by analyzing its three-dimensional crystal structure. Maximum MnP activity observed in steady-state fermentation by P. ostreatus was found in less than 20 days Table 4.1 Highly preferred assay for identification of peroxidase S. no.

Assay

Reaction mixture

λ (nm)

Reference

1

3,3 -diaminobenzidine

Horseradish peroxidase, O-dianisidine, gelation, sodium phosphate (pH 7.0), 3,3 -diaminobenzidine, H2 O2

465

Herzog and Fahimi (1973)

Malonate, MnSO4 , H2 O2 , 2, 6 DMP and phenol red, sodium tartarate buffer (pH 4.5)

468

Jong et al. (1992)

(DAB)

2

Manganese peroxidase (MnP)

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in dry potato peel waste, pretreated with distilled water was report. The enzyme production was maximum on the thirteenth day of incubation for when incubated with P. chrysosporium and Phanerochaete sp. (Rajan et al. 2010). In genome of Postia placenta brown rot fungi ten lignin peroxidases, five manganese peroxidases, and several other lignocellulolytic enzymes were encoded that indicate a rapid destruction and depolymerization of lignin (Martinez et al. 2004). Sometimes it is also observed that the extracellular product of lignocellulose diffuses from the enzyme surface and oxidizes the phenolic substrate to an organic pollutant (Cheng et al. 2013). Understanding the genome, proteome, and transcriptome offers an insight into diversification of lignocellulose-degrading mechanisms. Pleurotus pulmonarius fungi depend on Mn2+ for oxidizing phenolic and non-phenolic compounds to degrade several xenobiotic compounds like decolorizing of the dye anthraquinone, azo and Congo red (da Silva et al. 2016). Most of the ligninolytic enzymes acts as a natural ecosystem engineers and thus are reported to degrade 99% of phenanthrene and pyrene that are released into the environment as a result of incomplete combustion of polycyclic aromatic hydrocarbons (Agrawal et al. 2018). These ligninolytic enzymes thus have an important role in the transformation and mineralization of various organic pollutants (Wang et al. 2008). Manganese peroxidase enzyme finds the application in several fields that include pulp bleaching, biomechanical pulping, dye decolorization, bioremediation, and production of highly valuable chemicals from residual lignin from biorefineries, pulp, and paper side-streams (Jarvinen et al. 2012).

4.4.2.1

Production of Manganese Peroxidase

Activation of manganese peroxidase activity was detectable after induction with MnCl2 (Kuhar et al. 2016). The enzyme is highly specific to Mn2+ binding site, to catalyze the oxidation of Mn2+ ions to highly reactive Mn3+ ions. MnP has a high specificity for Mn2+ binding site shown in the crystal structure of manganese peroxidase from P. chrysosporium with a resolution of 2.0 Å (Sundaramoorthy et al. 1994). In the binding site of manganese peroxidase, there are three amino acid residues with an additional tryptophan residue on the enzyme surface that have been found required for the catalytic activity. Manganese peroxidase resembles lignin peroxidase to perform long-range electron transfer. The oxidation of Mn2+ ions is required by the enzyme to convert it to highly reactive Mn3+ ions. The Mn3+ in turn acts as a mediator to attack simple phenols, amines and phenolic lignin. MnP was first described in P. chrysosporium as an extracellular enzyme capable of decolorizing dye and decarboxylating vanilic acid. They are the key enzymes involved the lignin degradation system and organic pollutant degrader with genetic modification and expression system (Irie et al. 2001). The ability to synthesize manganese peroxidase in the groups of basidiomycetes, also observed in different habitats such as decaying sea grass, cooling tower wood, and brown coal. The catalytic cycle of manganese peroxidase is shown in Fig. 4.4.

4.4 Enzymes Involved in the Degradation

63

Fig. 4.4 Catalytic cycle of manganese peroxidase

4.4.3 Versatile Peroxidase Versatile peroxidase (EC 1.11.1.16) is also called as hybrid peroxidase or polyvalent peroxidase. Versatile peroxidases (VP) are extracellular hem protein. These are thirdtype lignin-modifying peroxidase comprising both LiP and MnP activities, thus are able to degrade a large number of substrate. It requires hydrogen peroxidase as electron acceptor to catalyze oxidative reactions with release of water molecule (Makela et al. 2015; Perez-Boada et al. 2005). VP is expressed in recombinant form several microbes that range from bacteria to fungi making it suitable in industrial application. VP was isolated from Physisporinus vitreus and expressed in E coli, together with lignin decomposition VP also had dye decoloration activity that requires Mn (II) to promote oxidation and dye decoloration (Liu et al. 2019). In Pleurotus sapidus VP, secretion is induced by the biogas residues used as a carbon and nitrogen source and capable of degrading suspended lignin organosolv particles (Schuttmann et al. 2014). Bjerkandera sp. (B33/3) isolated from straw pulping produced VP the study showed lignin transformation was possible directly without an external mediator (Moreira et al. 2007; Sridhar 2016). Whereas studies in Bjerkandera fumosa (VPBF) showed p-aminobenzoic acid is required for the increased VP activity and tween 80, NaN3 anthracene and fluorene decreased the activity (Pozdnyakova et al. 2013). In Bjerkandera adusta VP activity was inactivated by calcium depletion indicating its importance in the activity (Verdín et al. 2006). VP is purified by ammonia sulfate precipitation, ion exchange, and gel chromatography with 2,2 -azino-bis-(3ethylbenzothiazoline-6-sulfonic acid) (ABTS) as a substrate. Maximum activity in Pleurotus eryngii was observed at 50 degree Celsius and pH 3 (Chen et al. 2010).

4.4.4 Cu-Containing Laccase Phenol oxidase includes laccases (EC 1.10.3.2) that are classified as multi copper oxidases. Laccase oxidizes compounds with free alcohol group which forms phenoxy

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radicals. Laccase degrades lignin to sinapyl alcohol and further acts as a mediator in enzymatic hydrolysis of lignin (Zheng et al. 2019). Laccases are observed in plants, insects, and bacteria, but the most studied are from the fungi classified as white-rot fungi. This enzyme catalyzes the oxidation of a broad range of aromatic substrates where water is formed as a by-product. Production of laccase was enhanced by phenolic compounds contained in rice bran. Paper and pulp industry is an abundant feedstock rich in polysaccharides and also is the largest polluting industries in the world. This is a great concern as it releases harmful chlorine-based compound in the atmosphere. Various application and methods are employed to mitigate with the pollution. Laccase enzyme used in the degradation is explained in detail and is studied to have high ligninolytic potential. Most of the studies on laccase are confined to those produced by fungi and mushroom (Leonowicz et al. 2001). Several bacterial strains that show positive laccase production activities are also reported. The activity of laccase isolated from the soil bacteria of alphine showed large diversity. Soil metagenomic analysis and soil pedology demonstrated the method of acquisition of environmental enzyme discovery, linking soil stratigraphy to enzyme profiling for laccase gene activity (Dandare et al. 2019). Bacterial laccase is highly valuable and advantageous due to the bio-prospecting capabilities for industrial application and the ease of genetic modification or de novo assembly. A recently report on Kocuria sp. PBS-1 soil bacterium that has laccase activity together with dye degradation activity (Boruah et al. 2019) comparable to the laccase of Bacillus pumilus. The enzyme was observed to be active over a wide temperature range of 20–50 °C, with the half-life of 12 h indicating its thermal stability and use at industrial scale . High value of benzaldehyde chemicals were released with rice straw, kraft lignin, and organosolv lignin using Trametes versicolor and Caldalkalibacillus thermarum strain TA2.A1. Thermalkaliphilic laccase was isolated from these strain with Caldalkalibacillus thermarum strain TA2.A1 showed potential application of bacterial source of laccase for attempting conversion of benzaldehyde chemicals (Yang et al. 2019). The enzyme was also active at a broad range of pH revealing the thermo tolerant nature. Laccase enzyme extracted from Myrothecium verrucaria ITCC8447 was optimized by response surface methodology. The enzyme was found to be stable and active over an alkaline pH of 7–9 and a temperature range of 30–40 degree Celsius (Agrawal et al. 2019). Thermo-tolerant laccase isolated from Geobacillus sp. strain WSUCF1 has a half-life of 120 h and found active at 50 °C. Commercial cocktail of recombinant enzyme can be transformed in E coli and used on industrial scale (Rai et al. 2019). Lignin degradation activity by laccase is enhanced if white-rot fungi are grown in a chemically defined medium with cotton stalk extract. About 16% mineralization was observed on the 16th day of incubation than seen in the control tube was only 7%. The cotton stalk and aromatic compounds in the medium enhanced the laccase production. Wood and plant extract may contain some substances that may enhance the laccase enzyme production (Ardon et al. 1998). Pleurotus when grown on cotton straw, both the straw and lignin were degraded. It was fond of all laccase, oxidase, peroxidase and cellulose after 8–10 days rapidly declined. However, laccase activity was still reported after 21 days suggesting its essential role in the degradation of

4.4 Enzymes Involved in the Degradation

65

Table 4.2 Annotated ligninase enzymes from UniProt database Microorganism

Protein names

Development and Induction

References

Arthromyces ramosus

Peroxidase (EC-1.11.1.7)

–

Sawai-Hatanaka et al. (1995)

Coprinopsis cinerea

Peroxidase (EC-1.11.1.7)

–

Bausgaard et al. (1993)

Phanerochaete chrysosporium

Manganese peroxidase H3 (EC-1.11.1.13)

–

Stajich et al. (2010)

Phanerochaete chrysosporium

Ligninase A (EC-1.11.1.14)

–

Orth et al. (1994)

Phanerochaete chrysosporium

Manganese peroxidase H4 (EC-1.11.1.13)

Induced in wound-healing and by suberization factors

Gaskell et al. (1991)

Phanerochaete chrysosporium

Manganese peroxidase 1 (MnP-1) (EC-1.11.1.13)

Induced in wound-healing and by suberization factors

Godfrey et al. (1990)

Phanerochaete chrysosporium

Ligninase LG5, LG6 (EC-1.11.1.14)

–

Zhang et al. (1991)

Phanerochaete chrysosporium

Manganese peroxidase H5 (Fragment) (EC-1.11.1.13)

–

Pease and Tien (1992)

Phanerochaete chrysosporium

Ligninase LG2, B (EC-1.11.1.14)

LG2 is expressed during secondary metabolism and is triggered by nutrient limitation

Ritch et al. (1991; Ritch and Gold (1992)

Phanerochaete chrysosporium

Ligninase H2 (EC-1.11.1.14)

Expressed during secondary metabolism

Glumoff et al. (1990)

Phanerochaete chrysosporium

Ligninase LG (EC-1.11.1.14)

Expressed during secondary metabolism

Naidu and Reddy (1992)

Phlebia radiate

Manganese peroxidase 2 (MnP2) (EC-1.11.1.13)

Induced in wound-healing and by suberization factors

Hilden et al. (2006)

Phlebia radiate

Manganese peroxidase 3 (MnP3) (EC-1.11.1.13)

Induced in wound-healing and by suberization factors suberization

Hilden et al. (2005)

(continued)

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4 Ligninase in Degradation of Lignocellulosic Wastes

Table 4.2 (continued) Microorganism

Protein names

Development and Induction

References

Pleurotus eryngii

Versatile peroxidase VPL1, VPS1, VPL2 (EC- 1.11.1.16)

–

Ruiz-Duenas et al. (1999)

Taiwanofungus camphorates

Low-redox potential peroxidase (EC-1.11.1.7)

–

Huang et al. (2009)

Trametes versicolor

Ligninase A Fragment, B and C (EC-1.11.1.14)

Fragment C is expressed during secondary metabolism and is triggered by nutrient limitation

Johansson et al. (1993)

lignin. 3-Hydroxyanthranilic acid (3-HAA) is a potential laccase stimulator, isolated by hot extract of wheat straw. It is known to mediate the oxidation of non-phenolic part of lignin. The bioconversion efficiency was enhanced when guaiacyl lignin-type removal by laccase for guaiacyl-rich lignocellulosic biomass (Feng et al. 2019). Similarly, biomass that composes of forage grasses showed the role of lignin in biomass recalcitrance. White-rot fungi Pycnoporus cinnabarinus is also known to produce one isoform of laccase. In the study performed by Eggert (1996), showed that laccase less mutant greatly reduced the lignin degradation activity indicating the importance of laccase in lignin degradation. It was proved in Streptomyces coelicolor A3 mutant type, the importance of laccase in lignin degradation (Majumdar et al. 2014), whereas in some studies (Ander et al. 1990) showed the importance of multiple ligninolytic enzyme in the degradation of lignin Coriolus versicolor in combination of syringic acid supplemented with sweet sorghum bagasse. Pretreatment or saccharification of ligninolytic biomass like sugarcane bagasse enhances the process of biodegradation. In Pycnoporus sanguineus culture addition of coffee pulp was studied to enhance the laccase activity with urea used as a sole source of nitrogen source. Sole addition of coffee pulp increased the fungal biomass and laccase activity inducing two isoforms of the enzyme, whereas high amount of coffee inhibited laccase activity. The annotated ligninase enzymes from UniProt database are represented in Table 4.2.

4.5 Future Perspectives The rationale behind deciding the approach for purification is to achieve the maximum yield of the lignin-modifying enzymes with the highest catalytic activity and the potential purity. This chapter addresses recent developments related to and rated

4.5 Future Perspectives

67

to the global market for lignocellulose-degrading enzymes. The investigation of sustainable substrates, microorganisms, and fermentation strategies is to be evolved to achieve higher productivity and economic feasibility.

References Agrawal K, Bhardwaj N, Kumar B et al (2019) Process optimization, purification and characterization of alkaline stable white laccase from Myrothecium verrucaria ITCC-8447 and its application in delignification of agroresidues. Int J Biol Macromol 15:1042–1055 Agrawal N, Verma P, Shahi S (2018) Degradation of polycyclic aromatic hydrocarbons (phenanthrene and pyrene) by the ligninolytic fungi Ganoderma lucidum isolated from the hardwood stump. Biores Biopro 5:11 Ander P, Mishra C, Farrell R, Eriksson K (1990) Redox reactions in lignin degradation: Interactions between laccase, different peroxidases and cellobiose: quinone oxidoreductase. J Biotechnol 13:189–198 Archibald F (1992) A new assay for lignin-type peroxidases employing the dye Azure B. Appl Environ Microbiol 58:3110–3116 Ardon O, Kerem Z, Hadar Y (1998) Enhancement of lignin degradation and laccase activity in Pleurotus ostreatus by cotton stalk extract. Can J Microbiol 44:676–680 Arora D, Gill P (2001) Comparison of two assay procedures for lignin peroxidase. Enzyme Microb Technol 28:602–605 Baunsgaard L, Dalboge H, Houen G et al (1993) Amino acid sequence of Coprinus macrorhizus peroxidase and cDNA sequence encoding Coprinus cinereus peroxidase: a new family of fungal peroxidases. Eur J Biochem 1(213):605–611 Bechem E, Etaka S (2018) Screening of fungi isolated from household waste materials for ligninase activity. Int J Curr Res Biosci Plant Biol 5:1–28 Boruah P, Sarmah P, Das P, Goswami T (2019) Exploring the lignolytic potential of a new laccase producing strain Kocuria sp. PBS-1 and its application in bamboo pulp bleaching. Int Biodeterior Biodegrad 143:104726 Brink D, Ravi K, Lidén G, Gorwa-Grauslund M (2019) Mapping the diversity of microbial lignin catabolism: experiences from the eLignin database. Appl Microbiol Biotechnol 103:3979–4002 Butler M, Day A (1998) Destruction of fungal melanins by ligninases of Phanerochaete chrysosporium and other white rot fungi. Int J Plant Sci 159:989–995 Canales M, Lobos S, Vicuna R (1998) Molecular modeling of manganese peroxidase from the lignin-degrading fungus Cerioporiopsis subvermispora and structural comparison with other peroxidases. Electron J Biotechnol 1:25–96 Chen M, Yao S, Zhang H, Liang X (2010) Purification and characterization of a versatile peroxidase from edible mushroom Pleurotus eryngii. Chinese J Chem Eng 18:824–829 Cheng Z, Xiang-hua W, Ping N (2013) Continuous acid blue 45 decolorization by using a novel open fungal reactor system with ozone as the bactericide. Biochem Eng J 79:246–252 Collins P, Field J, Teunissen P, Dobson A (1997) Stabilization of lignin peroxidases in white rot fungi by tryptophan. Appl Environ Microbiol 63:2543–2548 da Silva B, Gomes Correa R, Kato C et al (2016) Characterization of a solvent-tolerant manganese peroxidase from Pleurotus pulmonarius and its application in dye decolorization. Curr Biotechnol 6:318–324 Dandare S, Young J, Kelleher B, Allen C (2019) The distribution of novel bacterial laccases in alpine paleosols is directly related to soil stratigraphy. Sci Total Environ 671:19–27 Dosoretz C, Chen H, Grethlein H (1990) Effect of environmental conditions on extracellular protease activity in lignolytic cultures of Phanerochaete chrysosporium. Appl Environ Microbiol 56:395– 400

68

4 Ligninase in Degradation of Lignocellulosic Wastes

Eggert C, Temp U, Dean J, Eriksson K (1996) A fungal metabolite mediates degradation of nonphenolic lignin structures and synthetic lignin by laccase. FEBS Lett 391:144–148 Feng N, Guo L, Ren H et al (2019) Changes in chemical structures of wheat straw auto-hydrolysis lignin by 3-hydroxyanthranilic acid as a laccase mediator. Int J Biol Macromol 122:210–215 Fernando S, Adhikari S, Chandrapal C, Murali N (2006) Biorefineries: current status, challenges, and future direction. Energy Fuels 20:1727–1737 Gaskell J, Dieperink E, Cullen D (1991) Genomic organization of lignin peroxidase genes of Phanerochaete chrysosporium. Nucleic Acids Res 19:599–603 Glumoff T, Harvey J, Molinari S et al (1990) Lignin peroxidase from Phanerochaete-chrysosporium molecular and kinetic characterization of isozymes. Eur J Biochem 187:515–520 Godfrey B, Mayfield M, Brown J, Gold M (1990) Characterization of a gene encoding a manganese peroxidase from Phanerochaete chrysosporium. Gene 93:119–124 Hariharan S, Nambisan P (2013) Optimization of lignin peroxidase, manganese peroxidase, and lac production from Ganoderma lucidum under solid state fermentation of pineapple leaf. BioResources 8:250–271 Hatakka A, Lundell T, Hofrichter M, Maijala P (2003) Manganese peroxidase and its role in the degradation of wood lignin. ACS Symp Series 855:230–243 Herzog V, Fahimi H (1973) A new sensitive colorimetric assay for peroxidase using 3,3 diaminobenzidine as hydrogen donor. Anal Biochem 55:554–562 Hilden K, Martinez A, Hatakka A, Lundell T (2005) The two manganese peroxidases Pr-MnP2 and Pr-MnP3 of Phlebia radiata, a lignin-degrading basidiomycete, are phylogenetically and structurally divergent. Fungal Genet Biol 42:403–419 Hilden K, Makela M, Hakala T et al (2006) Expression on wood, molecular cloning and characterization of three lignin peroxidase (LiP) encoding genes of the white rot fungus Phlebia radiata. Curr Genet 49:97–105 Horisawa S, Inoue A, Yamanaka Y (2019) Direct ethanol production from lignocellulosic materials by mixed culture of wood rot fungi Schizophyllum commune, Bjerkandera adusta, and Fomitopsis palustris. Fermentation 5:1–8 Huang S, Tzean S, Tsai B, Hsieh H (2009) Cloning and heterologous expression of a novel ligninolytic peroxidase gene from poroid brown-rot fungus Antrodia cinnamomea. Microbiol 155:424–433 Irie T, Honda Y, Watanabe T, Kuwahara M (2001) Homologous expression of recombinant manganese peroxidase genes in ligninolytic fungus Pleurotus ostreatus. Appl Microbiol Biotechnol 55:566–570 Jarvinen J, Taskila S, Isomaki R, Ojamo H (2012) Screening of white-rot fungi manganese peroxidases: A comparison between the specific activities of the enzyme from different native producers. AMB Express 2:1–9 Jing D, Wang J (2012) Controlling the simultaneous production of laccase and lignin peroxidase from Streptomyces cinnamomensis by medium formulation. Biotechnol Biofuels 5:1–7 Johansson T, Welinder K, Nyman P (1993) Isozymes of lignin peroxidase and manganese (II) peroxidase from the white-rot basidiomycete Trametes versicolor. II. Partial sequences, peptide maps, and amino acid and carbohydrate compositions. Arch Biochem Biophys 300:57–62 de Jong E, Field J, de Bont J (1992) Evidence for a new extracellular peroxidase manganese-inhibited peroxidase from the white-rot fungus Bjerkanderasp. BOS 55. FEBS Lett 299:107–110 Kapich A, Prior B, Botha A et al (2004) Effect of lignocellulose-containing substrates on production of ligninolytic peroxidases in submerged cultures of Phanerochaete chrysosporium ME-446. Enzyme Microb Technol 34:187–195 Klein-Marcuschamer D, Oleskowicz-Popiel P, Simmons B, Blanch H (2010) Technoeconomic analysis of biofuels: A wiki-based platform for lignocellulosic biorefineries. Biomass Bioenergy 34:1914–1921 Kuhar F, Castiglia V, Zamora J (2016) Detection of manganese peroxidase and other exoenzymes in four isolates of Geastrum (Geastrales) in pure culture. Rev Argent Microbiol 48:274–278

References

69

Leonowicz A, Cho N, Luterek J et al (2001) Fungal laccase: properties and activity on lignin. J Basic Microbiol 41:185–227 Leonowicz A, Matuszewska A, Luterek J et al (1999) Biodegradation of lignin by white rot fungi. Fungal Genet Biol 27:175–185 Lestan D, Strancar A, Perdih A (1990) Influence of some oils and surfactants on ligniolytic activity, growth and lipid fatty acids of Phanerochaete chrysosporium. Appl Microbiol Biotechnol 34:426– 428 Lewis N, Yamamoto E (1990) Lignin: Occurrence, biogenesis and biodegradation. Annu Rev Plant Physiol Plant Mol Biol 41:455–496 Linko S (1992) Production of Phanerochaete chrysosporium lignin peroxidase. Biotechnol Adv 40:494–498 Liu J, Zhang S, Shi Q et al (2019) Highly efficient oxidation of synthetic and natural lignin-related compounds by Physisporinus vitreus versatile peroxidase. Int Biodeterior Biodegrad 136:41–48 Majumdar S, Lukk T, Solbiati J et al (2014) Roles of small laccases from streptomyces in lignin degradation. Biochem 53(24):4047–4058 Makela M, Marinovi´c M, Nousiainen P et al (2015) Aromatic metabolism of filamentous fungi in relation to the presence of aromatic compounds in plant biomass. Adv Appl Microbiol 91:63–137 Martinez D, Larrondo L, Putnam N et al (2004) Genome sequence of the lignocellulose degrading fungus Phanerochaete chrysosporium strain RP78. Nat Biotechnol 22:695–700 Moreira P, Almeida-Vara E, Malcata F, Duarte J (2007) Lignin transformation by a versatile peroxidase from a novel Bjerkanderasp. strain. Int Biodeterior Biodegrad 59:234–238 Naidu P, Reddy C (1992) Nucleotide sequence of a new lignin peroxidase gene GLG3 from the white-rot fungus, Phanerochaete chrysosporium. Nucleic Acids Res 20:4124 Nerud F, Misurcova Z (1996) Distribution of ligninolytic enzymes in selected white-rot fungi. Folia Microbiol (Praha) 41:264–266 Orth A, Rzhetskaya M, Cullen D, Tien M (1994) Characterization of a cDNA encoding a manganese peroxidase from Phanerochaete chrysosporium: genomic organization of lignin and manganese peroxidase-encoding genes. Gene 148:161–165 Ozcirak S, Ozturk R (2017) Production of ligninolytic enzymes by solid state fermentation using Pleurotus ostreatus. Ann Agrar Sci 15:273–277 Pazarlioglu N, Urek R, Ergun F (2005) Biodecolourization of direct blue 15 by immobilized Phanerochaete chrysosporium. Process Biochem 40:1923–1929 Pease E, Tien M (1992) Heterogeneity and regulation of manganese peroxidases from Phanerochaete chrysosporium. J Bacteriol 174:3532–3540 Perez-Boada M, Ruiz-Dueñas F, Pogni R et al (2005) Versatile peroxidase oxidation of high redox potential aromatic compounds: Site-directed mutagenesis, spectroscopic and crystallographic investigation of three long-range electron transfer pathways. J Mol Biol 354:385–402 Philip R, Kachiprath B, Solomon S, Jayanath G (2019) Mangrove microflora as potential source of hydrolytic enzymes for commercial applications. Ind J Geo-Marine Sci 48:678–684 Pozdnyakova N, Makarov O, Chernyshova M et al (2013) Versatile peroxidase of Bjerkandera fumosa: substrate and inhibitor specificity. Enzyme Microb Technol 52:44–53 Rai R, Bibra M, Chadha B, Sani R (2019) Enhanced hydrolysis of lignocellulosic biomass with doping of a highly thermostable recombinant laccase. Int J Biol Macromol 137:232–237 Rajan A, Kurup J, Abraham T (2010) Solid state production of manganese peroxidases using arecanut husk as substrate. Brazi Arch Biol Technol 53:555–562 Ralph J, Lapierre C, Boerjan W (2019) Lignin structure and its engineering. Curr Opin Biotechnol 56:240–249 Reina R, Kellner H, Hess J et al (2019) Genome and secretome of Chondrostereum purpureum correspond to saprotrophic and phytopathogenic life styles. PLoS One Ritch T, Gold M (1992) Characterization of a highly expressed lignin peroxidase-encoding gene from the basidiomycete Phanerochaete chrysosporium. Gene 118:73–80 Ritch T, Nipper V, Akileswaran L et al (1991) Lignin peroxidase from the basidiomycete Phanerochaete chrysosporium is synthesized as a preproenzyme. Gene 14:e0212769

70

4 Ligninase in Degradation of Lignocellulosic Wastes

Ruiz-Duenas F, Martínez M, Martínez A (1999) Molecular characterization of a novel peroxidase isolated from the ligninolytic fungus Pleurotus eryngii. Mol Microbiol 31:223–235 Sawai-Hatanaka H, Ashikari T, Tanaka Y et al (1995) Cloning, sequencing, and Heterologous expression of a gene coding for Arthromyces Ramosus peroxidase. Biosci Biotechnol Biochem 59:1221–1228 Schuttmann I, Bouws H, Szweda R et al (2014) Induction, characterization, and heterologous expression of a carotenoid degrading versatile peroxidase from Pleurotus sapidus. J Mol Catal B Enzym 103:79–84 Sridhar M (2016) Versatile Peroxidases: Super peroxidases with potential biotechnological applications—a mini review. J Dairy Vet Anim Res 4:00116 Stajich J, Wilke S, Ahrén D et al (2010) Insights into evolution of multicellular fungi from the assembled chromosomes of the mushroom Coprinopsis cinerea (Coprinus cinereus). Proc Natl Acad Sci 107:11889–11894 Sundaramoorthy M, Kishi K, Gold M, Poulos T (1994) Preliminary crystallographic analysis of manganese peroxidase from Phanerochaete chrysosporium. J Mol Biol 238:845–848 Usha K, Praveen K, Reddy B (2014) Enhanced production of ligninolytic enzymes by a mushroom Stereum ostrea. Biotechnol Res Int 2014:1–9 Vanholme R, Demedts B, Morreel K et al (2010) Lignin biosynthesis and structure. Plant Physiol 153:895–905 Vazquez M, Cabrera E, Aceves M, Mallol J (2019) Cellulolytic and ligninolytic potential of new strains of fungi for the conversion of fibrous substrates. Biotechnol Res Innov 3:177–186 Venkatadri R, Irvine R (1990) Effect of agitation on ligninase activity and ligninase production of Phanerochaete chrysosporium. Appl Environ Microbiol 56:2684–2691 Verdín J, Pogni R, Baeza A et al (2006) Mechanism of versatile peroxidase inactivation by Ca2+ depletion. Biophys Chem 121:163–170 Walther I, Kalin M, Reiser J et al (1988) Molecular analysis of a Phanerochaete chrysosporium lignin peroxidase gene. Gene 70:127–137 Wang P, Hu X, Cook S et al (2008) Effect of culture conditions on the production of ligninolytic enzymes by white rot fungi Phanerochaete chrysosporium (ATCC 20696) and separation of its lignin peroxidase. World J Microbiol Biotechnol 24:2205–2212 Wang X, Ruckenstein E (1994) Immobilization of Phanerochaete chrysosporium on porous polyurethane particles with application to biodegradation of 2-chlorophenol. Biotechnol Tech 8:339–344 Whitaker B, Bauer J, Bever J, Clay K (2017) Negative plant-phyllosphere feedbacks in native Asteraceae hosts—a novel extension of the plant-soil feedback framework. Ecol Lett 20:1064– 1073 Wuyep P, Khan A, Nok A (2003) Production and regulation of lignin degrading enzymes from Lentinus squarrosulus (mont.) Singer and Psathyrella atroumbonata Pegler. Afr J Biotechnol 2:444–447 Xiong X, Wen X, Bai Y, Qian Y (2008) Effects of culture conditions on ligninolytic enzymes and protease production by Phanerochaete chrysosporium in air. J Environ Sci 20:94–100 Yang Y, Song W, Hur H et al (2019) Thermoalkaliphilic laccase treatment for enhanced production of high-value benzaldehyde chemicals from lignin. Int J Biol Macromol 124:200–208 Zhang Y, Reddy C, Rasooly A (1991) Cloning of several lignin peroxidase (LIP)-encoding genes: sequence analysis of the LIP6 gene from the white-rot basidiomycete, Phanerochaete chrysosporium. Gene 97:191–198 Zhao M, Zhang C, Zeng G et al (2015) Growth, metabolism of Phanerochaete chrysosporium and route of lignin degradation in response to cadmium stress in solid-state fermentation. Chemosphere 138:560–567 Zheng Y, Guo M, Zhou Q, Liu H (2019) Effect of lignin degradation product sinapyl alcohol on laccase catalysis during lignin degradation. Ind Crops Prod 139:1–9

Chapter 5

Pectinase in Degradation of Lignocellulosic Wastes

Abstract Pectin is one of the major plant cell wall components. Degradation of pectin is difficult, and pectinases enzyme can help in the degradation of pectin. The chapter described here mentions the pectic substances, viz. protopectin, pectinic acids, pectin or pectins, and pectic acids. The mechanism of action of the pectinolytic enzymes is also described along with the occurrence of pectinolytic enzymes. The physiochemical and biological properties of pectinases enzyme is also focused. The assay methods of pectinolytic enzymes and the production of pectinase enzymes by bacteria and fungi using fermentation are also described. The various applications of pectinases in the deconstruction of lignocellulosic wastes are also taken into consideration. This will help in understanding pectinolytic enzymes and their mechanisms of action which give insights on the role of pectinases in the degradation of lignocellulosic wastes. Keywords Polygalacturonase · Biofuels · Feedstock · Wastewater · Saccharification · Anti-inflammatory

5.1 Introduction The plant cell walls primarily consist of large biopolymers cellulose, hemicellulose, lignin, and pectin which provide the plant’s shape and support help to regulate physiological processes together defense responses and act as physical barriers to pathogen invasion (Aro et al. 2005). Pectin is the main and much complex plant cell wall components that have to be addressed by microorganisms invading the plant or utilize them for nutrition. Pectin is a heteropolysaccharide found in cell walls of terrestrial plants and middle lamella. Pectin is a major component of primary cell walls in dicots and non-graminaceous monocots, accounting for 30–35% of dry weight. It is also present in SCWs (in the middle lamella) and in grasses (Mohnen 2008). Pectin is a major component in cell wall construction and has a significant role in plant growth and development. It contributes in plant’s morphogenesis, fruit development, hydration of seeds, leaf abscission, defense, porosity, ion binding, cell adhesion, and cellular expansion (Mohnen 2008; Willats et al. 2011). © Springer Nature Switzerland AG 2020 A. B. Gunjal et al., Enzymes in Degradation of the Lignocellulosic Wastes, https://doi.org/10.1007/978-3-030-44671-0_5

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In nature, pectic structures are extremely diverse, depending on the origin of the plant and on the plant tissue. Three pectic substances are made up of three structural elements, viz. homogalacturonan, rhamnogalacturonan I, and substituted galacturonans. They have been isolated from primary cell walls, and their structures have been studied (O’Neill et al. 2004). They contain different degrees of their main component, d-galacturonic acid. Substituted galacturonans are characterized by the presence of saccharide appending residues branching from a backbone of dgalacturonic acid residues (Buchanan et al. 2015), whereas in rhamnogalacturonan I, the galacturonic acid residues are partly substituted by α-1-2 linked rhamnose residues. In addition, several side chains containing sugars, such as xylose, arabinose, glucose, fucose, mannose, or galactose, have been found to be linked to the main backbone structure (Muller-Maatsch et al. 2016). The galacturonic acid moieties in the backbone can also be esterified on the carboxylic acid moiety by methyl groups.

5.2 The Pectic Substances The complex colloidal carbohydrates found in or derived from plants contain a large proportion of anhydrogalacturonic acid units called as pectic substances. The carboxyl group of polygalacturonic acid to a certain extent is esterified by methyl groups. Sometimes they are partly or fully neutralized by one or more bases. “Pectin” is the term attributed to the family of oligosaccharide and polysaccharide which possess a high amount of galacturonic acid (65% as stipulated by Food and Agriculture Organization) (Brent et al. 2001). In the beginning of studies of pectic substances, there was great uncertainty regarding the naming of pectic substances. Now the definitions given by a committee of the American Society (1944) of these complex substances are commonly accepted, and the definitions have prevailed.

5.2.1 Protopectin Protopectin is the name coined for the water-insoluble parent pectic substance which occurs in plants, and from which pectic substances are produced. They are insoluble methylated polymer of galacturonic acid mainly found in raw fruits and vegetables.

5.2.2 Pectinic Acids Pectinic acids are defined as colloidal polygalacturonic acids containing an additional small proportion of methyl ester groups. Pectinic acids have characteristics of forming gels with sugar and acid under suitable conditions. It has been found that if the methoxyl content is low, then gel formation may take place with certain ions. The

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Fig. 5.1 Structure of pectinic acid

salts of pectinic acids formed are normal or acid salts of pectic acids. The structure of pectinic acid is shown in Fig. 5.1.

5.2.3 Pectin or Pectins Pectin or pectins term is coined for water-soluble pectinic acids of varying methyl ester content. They possess a different extent of neutralization. They can form gels with sugar and acid under suitable conditions. Pectinic acids are actually highmolecular-weight pectin. They are dispersible in water. The galacturonic acid chain possesses carboxyl groups that can undergo esterification with methanol. Pectins are classified as low-methoxyl or high-methoxyl pectins as per the degree of esterification. The low- and high-methoxyl groups have different properties. Gelling conditions for both are found to be different. Pectin consists of four different types of polysaccharides, and their structures are shown: Kdo, 3-deoxy-d-manno-2-octulosonic acid; DHA, 3-deoxy-d-lyxo-2heptulosaric acid (Harholt et al. 2010).

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5 Pectinase in Degradation of Lignocellulosic Wastes

Low-Methoxyl Pectins

Low-methoxyl pectins contain mostly free carboxyl groups. The carboxyl groups can be esterified to 20–40%. These methoxyl groups are accessible to form cross-links with divalent ions such as calcium.

5.2.3.2

High-Methoxyl Pectins

The esterified carboxyl groups are present in a high proportion (usually 50–58%) in high-methoxyl pectins. Most of the acid groups therefore are not available to form cross-links with divalent ions, so these pectins do not form gels.

5.2.4 Pectic Acid Pectic acid is the term designated to pectic substances comprising of colloidal polygalacturonic acid. The methyl ester groups are absent in pectic acid. Normal or acid pectate salts of pectic acid can be formed. A group of definitions given by Doesburg (1965) are considered by some workers to be more complete (McCready and Gee 1960). However, other workers considered the definitions given by the American Society (1944) (Fogarty and Kelly 1983; Kertesz 1951) to be more appropriate.

5.3 Microbial Pectinolytic Enzymes The plant-associated microorganisms produce pectinolytic enzymes. These pectinolytic enzymes play a considerable role in the decomposition and recycling of plant organic matter. Phytopathogenic microorganisms must penetrate carbohydrate barriers of host plant cell walls. Therefore, enzymes that are essential for the degradation of cell wall components are important for their infection to plants (Payasi et al. 2009). All of these enzymes are also known as cell wall-degrading enzymes (CWDEs). Among the best known microbial CWDEs are polygalacturonases (PG), pectin methylesterases (PME) and pectate lyases (PEL) (Lagaert et al. 2009). Pectic enzymes can cause plant tissue maceration, cell lysis, and modification of the cell wall structure, allowing other depolymerizing enzymes to act on their respective substrates. Several fungal pectin or pectate lyases from a variety of microbial species have been functionally characterized, such as the saprotrophic/opportunistic Aspergillus niger (Kusters-van Someren et al. 1992), Aspergillus oryzae (Kitamoto et al. 2001), Penicillium griseoroseum (Bazzolli et al. 2006), Penicillium occitanis (Trigui-Lahiani and Gargouri 2007), and the phytopathogenic fungi Glomerella cingulata (Templeton et al. 1994), Colletotrichum gloeosporioides, Colletotrichum

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lindemuthianum (Lara-Márquez et al. 2011), and Botrytis cinerea (Shah et al. 2009). PL1 is found out to be expanded gene families in Clonostachys rosea (Hypocreales, Bionectriaceae), a ubiquitous mycoparasitic fungus when comparative genome analysis was performed (Karlsson et al. 2015). The pectic enzymes that hydrolyze pectin have been categorized into two groups, esterases and depolymerases.

5.3.1 Esterases Pectin methyl esterase (EC.3.1.1.11) also called pectin esterase (systematic name: pectin pectyl hydrolase) cleaves methanol from esterified carboxyl groups to yield low-methoxyl pectin and polygalacturonic acid. High-methoxyl pectin is the preferred substrate. Pectin acetyl esterase (EC.3.1.1.6) hydrolyzes acetyl esters in HG regions of pectin (systematic name: acetic-ester acetyl hydrolase). Other frequent for esterases are C-esterase, acetic-ester hydrolase, chloroesterase, C-esterase, citrus acetyl esterase, and p-nitrophenyl acetate esterase.

5.3.2 Depolymerases The depolymerizing enzymes are either hydrolyzing pectin or pectic acid (polygalacturonic acid) or perform trans-elimination reaction on unsaturated polymethyl digalacturonates or pectic acid (Jayani et al. 2005). Depolymerases can also be divided into following subgroups: protopectinases acting on pectin; pectic acid (polygalacturonic acid); and oligo-d-galactosiduronates. Each subgroup can be further divided into two sub-subgroups: hydrolases and lyases, which can be further subdivided into endo- and exo-enzymes depending on the mode of action.

5.3.3 Protopectinases These enzymes degrade the insoluble protopectin and give rise to highly polymerized soluble pectin. The enzyme that catalyzes the solubilization of protopectin was originally named protopectinase (Flipphi et al. 1993; Leal and de Sa Nogueira 2004; Sakamoto et al. 1994). Pectinosinase is also synonymous with protopectinase. Protopectinase (PPase) catalyzes the following reaction: PPase

Protopectin + H2 O + −−→ Pectin (Insoluble)

(Soluble)

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5.3.3.1

5 Pectinase in Degradation of Lignocellulosic Wastes

Acting on Pectin

Polymethylgalacturonase Endo-polymethylgalacturonase (PMG) performs random hydrolysis of highly esterified to form oligomethylgalacturonates. Exo-polymethylgalacturonase (PMG) hydrolyzes pectin in a sequential fashion from the terminal end liberating monogalacturonates.

Polymethylgalacturonate Lyase Endo-polymethylgalacturonate lyase (PMGL) (EC.4.2.2.10) causes random cleavage in pectin by a trans-elimination process, forming a double bond between C4 and C5 of the galacturonic acid residues at the non-reducing end (systematic name: poly (methyl galactosiduronate) endo-lyase) and other common name is endo-pectin lyase. PMGL gives rise to unsaturated methyl oligo galacturonates by random trans-elimination of unsaturated poly (methyl) d-digalacturonate (Mayans et al. 1997; Mutenda et al. 2002). Unsaturated methyl monogalacturonates are released by action of exo-poly methyl d-galactosiduronate lyase by trans-elimination of unsaturated poly-(methyl-ddigalacturonate). It has systematic name poly (methoxygalactosiduronate) exo-lyase) also called exo-pectin lyase.

5.3.3.2

Acting on Polygalacturonic Acid or Pectic Acid

Polygalacturonase Endo-polygalacturonase (PG), EC.3.2.1.15, hydrolyzes polygalacturonic acid in a random fashion (systematic name: poly (1,4-α-d-galactosiduronate) glycanohydrolase) releasing oligogalacturonides (Latendresse 2013). They react with the inner site, i.e., the polygalacturonic acid regions.

Polygalacturonate Lyase Endo-polygalacturonate lyase (PGL) (EC.4.2.2.2) causes random cleavage in polygalacturonic acid by a trans-elimination process (systematic name: poly-(1,4-α-dgalactosiduronate) endo-lyase) commonly called as endo-pectate lyase. Exo-polygalacturonate lyase (EC.4.2.2.9) causes sequential cleavage in polygalacturonic acid by trans-elimination process (systematic name: poly (1,4-α galactosiduronate) exo-lyase) commonly called as exo-pectate lyase. The eliminative

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cleavage of an unsaturated disaccharide from the reducing end of homogalacturonan is catalyzed by the enzyme. It is the backbone of smooth regions of pectate, also known as de-esterified pectin.

5.3.3.3

Acting on Oligo-D-Galactosiduronates

Oligogalacturonase (OG) or Exo-Poly-α-Galacturonosidase (EC.3.2.1.82) Enzyme Commission Synonyms are exo-polygalacturonosidase, exopolygalacturanosidase, poly (1,4-α-d-galactosiduronate) digalacturonohydrolase. The bacterial oligogalacturonase has been reported to hydrolyze the second α1,4-glycosidic bond from the non-reducing end of polygalacturonate, releasing digalacturonate (Latendresse 2013).

Oligogalacturonate Lyase (OGL) (EC.4.2.2.6) Oligogalacturonate lyase catalyzes the cleavage of unsaturated oligo-dgalactosiduronate by a trans-elimination process. It has been assigned systematic name as oligo-d-galactosiduronate lyase. It is also known as unsaturated oligogalacturonate transeliminase, OGTE. Oligogalacturonate lyase (OGL) catalyzes eliminative removal of unsaturated terminal residues from oligosaccharides of d-galacturonate.

5.4 Occurrence of Pectinolytic Enzymes The pectinolytic enzymes are widely found in bacteria, fungi, and plants. They are also found in higher plants, parasitic plants, and some plant-parasitic nematodes. They have been screened out from various sources to hydrolyze pectins to produce valuable end products that possess immense applications in various industrial processes.

5.4.1 Esterases Pectin methyl esterase (EC.3.1.1.11) Pectin acetyl esterase (EC.3.1.1.6). Pectin esterases are found in plants, fungi, and plant pathogenic bacteria (Hasunuma et al. 2003). It has been reported in some species like Rhodotorula sp., Phytophthora infestans, Erwinia chrysanthemi B341 (Pitkanen et al. 1992), Saccharomyces

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cerevisiae (Gainvors et al. 1994), Lachnospira pectinoschiza (Cornick et al. 1994), Pseudomonas solanacearum (Schell et al. 1994), A. niger (Maldonaldo and Saad 1998) Lactobacillus lactis subsp. Cremoris (Karam and Belarbi 1995), Penicillium frequentans (Kawano et al. 1999), E. chrysanthemi 3604 (Laurent et al. 2000), P. occitanis (Hadj-Taieb et al. 2002), Aspergillus japonicus (Semenova et al. 2003), and others. There are many reports of presence of PE in plants like Carica papaya (Fayyaz et al. 1993; Innocenzo and Lajalo 2001), Lycopersicum esculentum (Warrilow et al. 1994), Prunus malus (Macdonald and Evans 1996), Vitis vinifera (Corredig et al. 2000), Citrus sp. (Arias and Burns 2002), Pouteria sapota (Arenas-Ocampo et al. 2003), and Malpighia glabra L. (Assis et al. 2004).

5.4.2 Depolymerases 5.4.2.1

Protopectinases

A-type PPase was detected in the culture filtrates of yeast and yeast-like fungi. Kluyveromyces fragilis IFO 0288, Galactomyces reesei L., and Trichosporon penicillatum SNO-3 produce protopectinases and are referred to as PPase-F, -L, and -S, respectively. Bacillus subtilis IFO 12,113, B. subtilis IFO 3134 (Sakai and Sakamoto 1990), and Trametes sp. (Sakai et al. 1993) produce B-type PPases and are referred to as PPase- B, -C, and -T, respectively. An extensive range of Bacillus sp. producing B-type PPases has also been reported in the culture filtrate (Sakai 1992).

5.4.2.2

Acting on Pectin

Polymethylgalacturonase Endo-polymethylgalacturonase (PMG) Exo-polymethylgalacturonase (PMG).

Polymethylgalacturonate Lyase Endo-polymethylgalacturonate lyase (PMGL) (EC.4.2.2.10). Polymethylgalacturonate lyases (pectin lyases or PMGLs) are found to be produced by few microorganisms as indicated by the literature survey. They have also been reported to be produced by Aspergillus japonicus, Penicillium paxilli, Penicillium sp. (Sapunova et al. 1995), Pythium splendens (Chen et al. 1998), Pichia pinus (Moharib et al. 2000), Aspergillus sp. (Sunnotel and Nigam 2002), and Thermoascus auratniacus (Martins et al. 2002).

5.4 Occurrence of Pectinolytic Enzymes

5.4.2.3

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Acting on Pectic Acid (polygalacturonic Acid)

Polygalacturonase Endo-polygalacturonase (PG) (EC.3.2.1.15) Exo-PG-1 (EC.3.2.1.67) Exo-poly methyl d-galactosiduronate hydrolases (EC.3.2.1.82). Endo-PGases are extensively distributed among fungi, bacteria, and numerous types of yeasts. They are present in higher plants and some plant-parasitic nematodes (Sakai et al. 1993). They have been screened out in many microorganisms, including Aureobasidium pullulans, Rhizoctonia solani Kuhn, Fusarium moniliforme, Neurospora crassa (Polizeli et al. 1991), Rhizopus stolonifer, Aspergillus sp. (Nagai et al. 2000), Thermomyces lanuginosus (Kumar and Palanivelu 1999), and Peacilomyces clavisporus (Souza et al. 2003). Endo-PGases have furthermore been cloned and studied in a great number of microbial species (Centis et al. 1996; Centis et al. 1997; Gainvors et al. 2000; Gao et al. 1996; Guo et al. 1996; Laing and Pretorius 1993; Naumov et al. 2001; Reymond et al. 1994). In contrast, exo-PGases occur in few microbial species. They have been reported in Erwinia carotovora, Agrobacterium tumefaciens (Rodrigues-Palenzuela et al. 1991), Bacteroides thetaiotaomicron (Tierny et al. 1994), E. chrysanthemi (Koboyashi et al. 2001), Alternaria mali (Nozaki et al. 1997), Fusarium oxysporum (Maceira et al. 1997), Ralstonia solanacearum (Huang and Allen 1997), and Bacillus sp. (Koboyashi et al. 2001). Exo-PGases are differentiated into two types: fungal exo-PGases, which produce monogalacturonic acid as the main end product; and the bacterial exo-PGases, which produce digalacturonic acid as the main end product (Sakai et al. 1993). Occurrence of PGases in plants has also been searched out (Alonso et al. 2003).

Polygalacturonate Lyase Endopolygalacturonate lyase (PGL) (EC.4.2.2.2). Polygalacturonate lyases (pectate lyases or PGLs) are reported in many bacteria and a few pathogenic fungi. They produce endo-PGLs being largely found than exoPGLs. The food spoilage and soft-rot bacteria and fungi have been reported to be producing PGLs. They have been reported in B. thetaiotaomicron, E. carotovora, and E. chrysanthemi (Favey et al. 1992; Shevchik et al. 1997).

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5.4.2.4

Acting on Oligo-D-Galactosiduronates

Oligogalacturonate Lyase (OGL) (EC.4.2.2.6) Colletotrichum lindemuthianum, Amucala sp. (Bruhlmann 1995), Pseudomonas syringae pv. Glycinea (Margo et al. 1994), Colletotrichum magna (Wattad et al. 1995), Bacillus sp. (Koboyashi et al. 1999; Singh et al. 1999a, b; Takao et al. 2000), Bacillus sp. DT-7 (Kashyap et al. 2000), C. gloeosporioides (Drori et al. 2003; Yakoby et al. 2000).

5.5 Physicochemical and Biological Properties 5.5.1 Physicochemical and Biological Properties of Esterases Pectin methylesterase (EC.3.1.1.11) Pectin acetylesterase (EC.3.1.1.6). The enzyme pectin esterase acts on methoxyl group adjacent to free carboxyl groups. Its activity is the highest about 65–75% methylated pectin. PE action has a very little effect on viscosity of pectin containing solutions unless divalent ions are present, which increases viscosity due to cross-linking. PEs is highly specific enzyme. Some PEs attack only at the reducing chain, while others attack the non-reducing end (Sakai et al. 1993). The molecular weights of most PEs are in the range of 35–50 kDa. The pH values at which PEs are active range from 4.0 to 8.0. Fungal PEs has a lower pH optimum than that of bacterial origin. Optimum temperature range for maximal activity for majority of PEs is 40–50 °C. Immobilization studies have also been carried out on PEs (Assis et al. 2004). Two isoforms of PE, isolated from A. japonicus, have a pI of 3.8 and are best active at 50 °C (Hasunuma et al. 2003). Two different PEs, viz. PmeA, an extracellular enzyme, and PmeB, an outer membrane protein (Shevchik et al. 1996), have been isolated from E. chrysanthemi. A PE reported from E. chrysanthemi 3937 showed the best activity at alkaline pH and 50 °C (Laurent et al. 2000).

5.5.2 Physicochemical and Biological Properties of Depolymerases 5.5.2.1

Protopectinases

A-type PPases are found to share common biological properties and have analogous molecular weight of 30 kDa. PPase-F is an acidic protein, and PPase-L and -S are

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basic proteins. Pectin is released from protopectin from various origins by activity of protopectinase enzyme. The enzymes catalyze the hydrolysis of polygalacturonic acid; they decrease the viscosity, slightly increasing the reducing value of the reaction medium containing polygalacturonic acid. PPase-B, -C, and -T have molecular weights of 45, 30, and 55 kDa, respectively. PPase-B and -C have an isoelectric point (pI) of around 9.0, whereas PPase-T has a pI of 8.1 (Sakai 1992). PPase-B, -C, and -T act on protopectin from various citrus fruit peels and other plant tissues, releasing pectin (Sakai 1992).

Acting on Pectin Polymethylgalacturonase Endo-polymethylgalacturonase (PMG) Exo-polymethylgalacturonase (PMG). Bacillus sp. strain BR1390 has PMG with a unique feature among bacterial pectinases with an optimum pH and temperature of 6.0 and 60 °C, respectively. Polymethylgalacturonase enzymes were studied to have 0.066 μmol min−1 (Vmax) and 2.51 mg ml−1 (Km) kinetic parameters using Hanes–Woolf linearized plot. The Km value of the PMG was lower than any Bacillus sp. pectinases reported so far (KlugSantner et al. 2006; Kobayashi et al. 1999; 2001; Nadaroglu et al. 2010; Payasi et al. 2009; van Dyk et al. 2010; Yadav et al. 2009). Among the different ions tested, Ca2+ , Cu2+ , Mn2+ , and Mg2+ stimulated the PMG activity that is in contrast with pectin lyase from Bacillus pumilus (Nadaroglu et al. 2010). The fungal and tomato pectinases do not require Ca2+ for their activity as required by polymethylgalacturonase. Due to the blocking function of Hg2+ and Zn2+ ions on thiol groups, they significantly inhibit the PMG activity. The studies have been carried out on involvement of thiol group in PMG active site as in case of B. subtilis polygalacturonase (Ahlawat et al. 2008). The enzyme is reported to be a metal-dependent pectinase as it was inhibited by 5 mM EDTA. The PMG activity was not affected by SDS, but was stimulated by Tween 80. These studies suggest high detergent stability similar to Tetracoccosporium sp. polygalacturonase (Aminzadeh et al. 2007).

Polymethylgalacturonate Lyase Endo-Polymethylgalacturonate Lyase (PMGL) (EC.4.2.2.10) Activity of PMGLs is stimulated by Ca2+ and other cations due to the masking of negative charges on polymethylgalacturonates (Dinnella et al. 1995). Interestingly, endo-PMGL is the only enzyme known to be able to cleave, without the prior action of other enzymes, the a-1,4-glycosidic bonds of highly esterified pectins (Dinnella et al. 1995). Two PMGLs, L1 and L2, from A. pullulans LV-10, showed optimal activity at pH 5.0 and 7.5, respectively, and temperature 40 °C.

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Acting on Pectic Acid (polygalacturonic Acid) Polygalacturonase Endo-polygalacturonase (PG) (EC.3.2.1.15) Exo-PG-1 (EC.3.2.1.67) Exo-poly methyl d-galactosiduronate hydrolases (EC.3.2.1.82). PGases isolated from different microbial sources differ markedly from each other with respect to their physicochemical and biological properties and their mode of action. Among the PGases obtained from different microbial sources, most have the optimal pH range of 3.5–5.5 and optimal temperature range of 30–50 °C. Two endoPGases (PG I and PG II), isolated from A. niger, have optimal pH range of 3.8–4.3 and 3.0–4.6, respectively (Singh and Rao 2002). Most of the PGase reported, work efficiently in acidic pH range, but a few alkaline PGases have been also reported from Bacillus licheniformis (Rehman et al. 2012) and F. oxysporum f. sp. lycopersci (Pietro and Roncero 1996) with optimum pH of 11.0. Four isoenzymes, viz. PG I, PG II, PG III, and PG IV, with same molecular weight but differing in their isoelectric points have been studied (Barnby et al. 1990). Although most of the PGases work in a temperature range of 30–40 °C, but a few PGases, which can catalyze the hydrolysis of pectic substances at higher temperatures, have also been isolated from Bacillus licheniformis (Rehman et al. 2012), Bacillus sp. KSM-P443 (Koboyashi et al. 2001), and F. oxysporum f. sp. lycopersci (Pietro and Roncero 1996). Polygalacturonate Lyase Endo-polygalacturonate lyase (PGL) (EC.4.2.2.2). Most of the pectin lyases have been reported from microorganisms, and there are scanty reports of their presence in plants and animals (Whitaker 1990). In bacteria, lyases are the largest group of pectinolytic enzymes and are directly involved in plant pathogenicity (Dixit et al. 2004). PGLs have an absolute requirement for Ca2+ ions (Margo et al. 1994), and hence, chelating agents such as EDTA act as their inhibitors. Most of the lyases have molecular weights ranging between 30 and 40 kDa, with isoelectric point ranging from 7.0–11.0. They have pH optima in the alkaline range (7.5–10.0) and optimum temperature of 40–50 °C. It was found that a thermostable exo-PGL from Bacillus sp., which showed maximum activity at pH 11.0 and temperature 69 °C and was dependent on Ca2+ for its activity. Thermostable lyases have also been reported from Bacillus sp. TS 47, T. auratniacus (Martins et al. 2002), and Fusarium monoiliforme (Dixit et al. 2004). Immobilized lyase from A. japonicus had a Km value of 0.16 and was best active at pH 6.0 and 55 °C. Many purification studies have also been carried out on lyases, which led to increase in specific activities of the enzymes with significant recoveries (Bruhlmann 1995).

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Acting on Oligo-d-Galactosiduronates Oligogalacturonate Lyase (OGL) (EC.4.2.2.6) In the bacterium Dickeya dadantii 3937 (E. chrysanthemi strain 3937), the gene ogl encodes a cytoplasmic oligogalacturonate lyase (Ogl) which cleaves unsaturated and saturated oligogalacturonides to their monomeric components (4S,5R)4,5-dihydroxy-2,6-dioxohexanoate and d-galacturonate. The action of this enzyme on a saturated digalacturonide substrate produces equimolar amounts of (4S,5R)-4,5dihydroxy-2,6-dioxohexanoate and d-galacturonate, whereas its action on an unsaturated digalacturonate substrate produces two molecules of (4S,5R)-4,5-dihydroxy2,6-dioxohexanoate. The gene encoding Ogl has been cloned from D. dadantii 3937 (E. chrysanthemi strain 3937). In contrast to the extracellular pectate lyases, Ca2+ is unable to restore the activity of Ogl, while several other cations, including Co2+ , Mn2+ , and Ni2+ , can activate oligo galacturonate lyase (Jann et al. 2008; Shevchik et al. 1999).

5.6 Assay Methods of Pectinolytic Enzymes 5.6.1 Assay Methods for Esterases Pectin methylesterase (EC.3.1.1.11) Pectin acetylesterase (EC.3.1.1.6). PE activity is extensively studied by gel diffusion assay (Downie et al. 1998). The principle that underlines the assay is the increase in binding of ruthenium red to pectin, as the number of methyl esters attached to the pectin decreases. The standard curve is constructed from the log-transformed commercial enzyme activity versus stained zone diameter. The unit of activity in nano- or picokatals is calculated from the curve constructed. The sensitivity, specificity, and simplicity of this PE assay are finer to other methods. A pH-stat method is also employed to calculate PE activity because ionization of the carboxyl group of the product releases a proton, which causes a change in pH.

5.6.2 Assay Methods for Depolymerases 5.6.2.1

Protopectinases

PPase activity assay is done by carbazole-sulfuric acid method. The amount of pectic substance liberated from protopectin is measured. The standard curve of dgalacturonic acid is constructed and pectin concentration is measured. One unit of

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protopectinase activity is defined as the moles of protopectin converted per unit time into pectic substance corresponding to 1 mmol of d-galacturonic acid per ml of reaction mixture.

Acting on Pectin Polymethylgalacturonase Endo-polymethylgalacturonase (PMG) Exo-polymethylgalacturonase (PMG). Polymethylgalacturonate Lyase Endo-polymethylgalacturonate lyase (PMGL) (EC.4.2.2.10)—the most convenient method of the following activity of lyases is to measure the increase in absorbance at 235 nm due to formation of the D 4:5 double bonds produced at the non-reducing ends of the unsaturated products (Liao et al. 1999; Whitaker 1990). The molar extinction coefficients for PMGL are 5.5 × 103 M−1 cm−1 . One unit of enzyme activity is defined as the amount of enzyme that releases 1 mmol of unsaturated product per min under assay conditions. Reducing group methods are also useful in determining the lyase activity. Viscosity reduction method in conjunction with a reducing group method or along with intermediate product analysis by HPLC or GC can be used to distinguish between endo- and exo-splitting enzymes.

Acting on Pectic Acid (polygalacturonic Acid) Polygalacturonase Endo-polygalacturonase (PG) (EC.3.2.1.15) Exo-PG-1 (EC.3.2.1. 67) Exo-poly methyl d-galactosiduronate hydrolases (EC.3.2.1.82). PGase activity involves the rate of increase in number of reducing groups during the course of the reaction and the decrease in viscosity of the substrate solution. The amount of reducing sugar is then measured by colorimetric methods, viz. 3,5dinitrosalicylate reagent method and the arsenomolybdate–copper reagent method. Under standard assay conditions, one unit of enzyme activity is defined as the enzyme required releasing 1 mmol ml−1 min−1 galacturonic acid. The PGase activity can also be determined by viscosity reduction measurements. The unit of enzyme activity is mostly selected as the amount of enzyme required for attaining a certain decrease of viscosity per unit time. However, this method has met with limited success. There is no direct correlation between viscosity reduction and number of glycosidic bonds hydrolyzed. The third method of PGase activity estimation is the cup-plate method. The substrate is added to molten agar and then

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solidified. The cups are cut out and are filled with the enzyme solution. After specific incubation time, the zones of degraded substrate are stained with iodine and quantified. Polygalacturonate Lyase Endopolygalacturonate lyase (PGL) (EC.4.2.2.2). The PGL activity is determined as in method for PMGL except for the fact that the molar extinction coefficients for PGL are 4.6 × 103 M−1 cm−1 .

Acting on Oligo-d-Galactosiduronates Oligogalacturonate lyase (OGL) (EC.4.2.2.6). Oligogalacturonate lyase activity is determined by monitoring spectrophotometrically the formation of unsaturated products from substrate at 230 nm. The standard assay mixture is made up of 50 mM Tris–HCl (pH 8.5), 0.1 mM CaCl2 , and 0.5 g of PGA l−1 . The appearance of products is monitored at 37 °C over a period of 1 min (at 6 s intervals). The molar extinction coefficient of unsaturated oligogalacturonates is 5200. One unit of activity is defined as the amount of enzyme required to produce 1 mmol of unsaturated product min−1 (Jahn et al. 2008; Shevchik et al. 1999a).

5.7 Production of Pectinases The microorganisms are primary source of industrial enzymes. About 50% originate from fungi and yeast, 35% from bacteria, and 15% are either of plant or animal origin (Anisa and Girish 2014). The pectinases are being produced by diverse types of microorganisms (Angayarkanni et al. 2002; Hoondal et al. 2002; Kapoor et al. 2001; Mohamadi et al. 2014; Servili et al. 1992; Sharma and Satyanarayana 2012; Sharma et al. 2013). An individual organism can produce pectinase along with additional industrially important enzymes (Kaur et al. 2011; Singh et al. 2015). Pectinases can be produced by both submerged and solid-state fermentation (SSF). The conditions for production of pectinases are different as per microbial isolate and substrate utilized for production. To reduce production costs, agro-waste sources, such as fruit waste and vegetable waste, have been used to produce pectinase using microbial solid-state fermentation. Bacterial pectinase is of choice over fungal pectinase as optimization of fermentation process is easy. Also, modern techniques can be easily applied to pectinase produced by bacteria improving production yield.

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Pectinases can be classified into acidic and alkaline enzymes on the basis of optimal pH required for enzymatic activity. Alkaline enzymes are nearly produced by bacteria and comprise numerous applications, such as textile processing; pharmaceutical uses; and leather, detergent, and paper production.

5.7.1 Production of Bacterial Pectinases 5.7.1.1

Bacterial Pectinases from Bacillus Sp.

Enzyme pectin lyase (EC.4.2.2.10) has been reported to be produced extracellularly by soil isolate, Bacillus sp. DT7 in considerable amounts. Bacillus sp. DT7 produced increased amount of pectin lyase (53 units/ml) after optimizing growth conditions in submerged fermentation. This mesophilic Bacillus sp. DT7 interestingly produces an alkalophilic and thermo-tolerant pectinase and hence has tremendous potential in textile industry and plant tissue maceration. The isolate has been also grown by solid-state fermentation on wheat bran for production of pectinases. Highest enzyme production of 8050 U/g dry substrate was obtained. The wheat bran is a cheap and readily available by-product; the production of pectinase using SSF will be very much economical. This enzyme has been successfully used for degumming of fiber crops (Kashyap et al. 2000, 2001, 2003), and the low cost of its production may further broaden the scopes for its use in industries involved in treatment of fiber crop. The other isolate Bacillus sp. AD 1 produces polygalacturonase (PGase) by submerged fermentation (SmF). The fermentation has been optimized employing pectin and citrus wastes as the sole carbon source. The enzyme was extremely thermostable, and 75% of the activity was restored even after the exposure of the enzyme protein at 80 °C for 90 min. The enzyme was 86% stable in a broad range of pH of 4–9 (Dey et al. 2011). The fermentation conditions for pectinase production by various microorganisms are represented in Table 5.1.

5.7.1.2

Bacterial Pectinases from Gram-Negative Microorganisms

A study was done to enrich and isolate the potential bacterial strain from the natural fruit reservoirs producing industrially important pectinase enzyme. Scale-up of production will lead to treatment of fruit wastes for production of reducing sugars from rotten apple and orange peels (Kumar and Sharma 2012).

5.7.1.3

Bacterial Pectinases from Erwinia carotovora sp.

Erwinia carotovora (Eca) is a member of the Enterobacteriaceae and is the causal agent of soft-rot disease in vegetable crops in temperate regions. The pathogenicity

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Table 5.1 Fermentation conditions for pectinase production by various microorganisms Microorganism

Substrate

Fermentation

References

Type

Temperature (°C)

pH

A. niger A 138

Sucrose

SmF

32

4.5

Friedrich et al. (1990)

Penicillium veridicatum RFC3

Bagasse

SSF

30

–

Silva et al. (2002)

Bacillus subtilis

Pectin

SmF

50

7.0

Swain and Ray (2010)

Thermomucor indicae-seudaticae

Wheat bran

SSF

45

–

Martin et al. (2010)

Pseudozyma sp. SPJ

Citrus peel

SSF

32

7.0

Sharma and Satyanarayana (2012)

Erwinia carotovora

Pectin

SmF

35

5.2

Kothari and Baig (2013)

Streptomyces sp.

Pectin

SmF

30

8.5

Das et al. (2013)

is related to the production of a variety of extracellular pectolytic enzymes of which pectate lyase (PL) and polygalacturonase (PG) are considered to be the most important. Hence, the isolate is screened for pectinases activity by submerged fermentation studies. The polygalactouronase is produced by the isolate with maximum activity at 35 °C and pH 5.2 (Kothari and Baig 2013).

5.7.1.4

Bacterial Pectinases from Streptomyces sp.

The mangrove isolate of Streptomyces sp. GHBA10 was used for production of polygalacturonase (PG). Submerged fermentation was carried out. The specific activity of the purified pectinase was determined to be 2610 U/mg of enzyme protein. Purification gave rise to 3.5-fold increase in specific activity of the enzyme. Also, another actinomycetes Streptomyces lydicus was found to be a potent producer of polygalacturonase by submerged fermentation. The enzyme was employed for the treatment of banana fibers and was found to be successful in separating the fibers which have application production of textiles, handicrafts, or banana paper (Jacob et al. 2008). An alkalophilic Streptomyces sp. RCK-SC produces thermostable alkaline pectinase. The half-life of pectinase was 3 h at 70 °C. Pectinase was stable at alkaline pH ranging from 6.0–9.0 for more than 8 h at room temperature retaining more than 50% of its activity. In an immobilized cell system containing polyurethane foam (PUF), the pectinase production was enhanced by 32% (101,000 IU l−1 ) compared to submerged fermentation. The enzyme has great potential due to the economic viability

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and commercial capability of pectinase in degumming of bast fibers and treatment of alkaline pectic wastewater (Kuhad et al. 2004).

5.7.2 Production of Fungal Pectinases 5.7.2.1

Production of Pectinases by Pseudozyma sp.

Production of high titers of an alkaline, extracellular, and thermo-tolerant pectinase by yeast Pseudozyma sp. SPJ has been carried out under solid-state fermentation. Pectinase production about 1215.66 IU/g dry substrate was achieved. Citrus peel, the inexpensive agro-industrial residue used as substrate, was experienced to be unrivaled.

5.7.2.2

Production of Pectinases by Pleurotus sajor-caju

Onion juice waste was studied for production of Pleurotus sajor-caju and pectinases. The onion waste residue is not suitable for fodder or landfill due to the rapid growth of phytopathogens. Hence, P. sajor-caju was grown on the onion waste to produce commercially important edible mushroom with several health benefits, such as immunomodulation, hypoglycemic, antioxidant, antithrombotic, anti-inflammatory, antimicrobial activities, antitumor activities, the ability to reduce blood pressure, and the ability to reduce the concentration of low-density lipoproteins along with production of pectinases enzyme production. The pectinases enzyme exhibited activity maximum at pH 6 and temperature 80 °C (Pereira et al. 2017).

5.7.2.3

Production of Pectinases by Penicillium and Aspergillus sp.

Production of pectinase by fungi using fruit waste such as orange peels was studied. Along with pectinase, the fungal isolates also produced cellulase. It was found out that highest production of polygalacturonase and endoglucanase by Penicillium atrovenetum was observed at pH 5, 40 °C, and at 0.2% ammonium persulfate. Polygalacturonase and endoglucanase production by Aspergillus flavus occurred at pH 5.5, temperature 40 °C, and 0.25% ammonium persulfate. A. oryzae produced the two enzymes at pH 5.5, temperature 35 °C, and 0.2% ammonium persulfate. The cocktail of the fungal isolates can be employed for bioconversion of agro-industrial wastes in to highly economic pectinases production (Johnson et al. 2012).

5.7 Production of Pectinases

5.7.2.4

89

Production of Pectinases by Thermomucor indicae-ieudaticae N31

Lignocellulosic waste wheat bran and orange peels were studied as economical feasible process for pectinase production by a Brazilian thermophilic fungus Thermomucor indicae-seudaticae N31 in solid-state and submerged fermentation. Under SSF conditions T. indicae-seudaticae N31 produced maximum of 120 U/ml of exo-PG. The fermentation medium was made by a mixture of wheat bran and orange bagasse (1:1). The initial moisture was kept at 70%. This fungus produced low quantities of exo-PG, i.e., 13.6 U/ml in submerged fermentation (SmF). The crude PG from SmF was found out to be more thermostable than that produced by SSF method. It also displayed higher stability in acidic pH (Martin et al. 2010).

5.7.2.5

Production of Polygalacturonase by Penicillium citrinum

Polygalacturonase enzyme produced by gamma-irradiated Penicillium citrinum was studied in solid-state fermentation (SSF) using sugar beet pulp as substrate. The gamma irradiations potentiate the productivity of the enzyme to its maximum value (152.2 U/gdfs) post exposure to 0.7 kGy. The optimum pH and temperature of the enzyme were found to be 6.0 and temperature 40 °C, respectively. The enzyme was found to be stable in the pH range 4–8 and showed high stability at temperature range 20–60 °C (El-Batal et al. 2013).

5.8 Pectinases as First Protein Product Made in Leaves Biological deconstruction of lignocellulosic wastes needs large quantities of enzymes. The unaffordable costs of current microbial enzymes restrict their widespread use in diverse industrial biological applications. Decades-old microbial production systems require prohibitively expensive fermentation facilities, purification from host cells, formulation to increase concentration and stability, and cold storage/transportation. Therefore, there is a great need to explore novel production technologies that could eliminate these prohibitively expensive enzyme processes. While the cost of enzyme production, mark-up by industries and production capacity are often debated, launching of leaf enzyme products, through PhylloZyme funded by investors who own textile or microbial enzyme industries, underscores the need for new platform technologies. The plant seeds are primary source of genetically modified commercial products. The first protein product made in leaves for commercial use has been produced. Leaf pectinases have been formulated as eight different liquid commercial microbial enzyme products for textile or juice industry applications. The leaf pectinases are of great advantage being functional in broad pH or temperature ranges as crude leaf extracts. Many commercial enzyme products showed considerable activity loss at

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alkaline pH or higher temperature during important textile applications. The leaf pectinase powder is found to possess shelf life of 16 months at ambient temperature without loss of enzyme activity. Today’s available commercial liquid enzymes require cold storage or transportation. As compared to the Fraunhofer method, the pectinase enzyme yield from the greenhouse plants was double. Thus, this leaf-production platform leads to a novel, economical approach for enzyme production as there is no requirement of fermentation, purification, concentration, formulation, and cold chain (Daniell et al. 2019).

5.9 Application of Pectinases in Deconstruction of Lignocellulosic Wastes Lignocellulosic biomass presents several features that confer recalcitrance, such as crystalline cellulose which precludes decomposition by enzymes (Rastogi and Shrivastava 2017); highly complex hemicelluloses and pectin, which demands a huge number of enzymes; the high degree of lignin adsorption to proteins that inhibits enzymatic activity (Huang et al. 2011); and the complex cross-linkages between phenolic and polysaccharide components (Lygin et al. 2011; Oliveira et al. 2016). Biological, chemical, and physical pretreatments can reduce the crystallinity of lignocellulose and break various linkages, drastically reducing its complexity (Kou et al. 2017). Biological pretreatments have been widely studied and have demonstrated some advantages over chemical and physical pretreatments such as a low demand for energy, environmental friendliness, and low levels of toxic products (Wei et al. 2015). Biological pretreatments include in-vivo application of microorganisms (Balat and Balat 2009). For highly complex pectin structures deconstruction, pectinases are of immense importance (Mota et al. 2018). Pectinases have a potential application in improving ethanol production from various feedstocks, as pectinase treatment requires less energy and produces no inhibitory factors (Chen et al. 2012; Kashyap et al. 2001; Kataria and Ghosh 2011; Oberoi et al. 2011).

5.9 Application of Pectinases in Deconstruction …

91

5.9.1 Potential Applications of Pectinases 5.9.1.1

Production of Reducing Sugars from Lignocellulosic Wastes

Coffee Pulp Waste (CPW) for Production of Reducing Sugars from Lignocellulosic Wastes Biological methods using bacteria and fungi are regarded as more economically viable and environmentally friendly alternatives for improving lignocellulosic degradation. There is a significant amount of lignocellulosic wastes generated during coffee manufacturing process. It is a prospective source of reducing sugars. On the other hand, it contains lignin as a matrix polymer. It is associated with pectin which covers the cellulosic microfibrils. This renders it difficult to be digested during the bioprocess. The biological pretreatment of coffee pulp waste by using a co-culture of B. subtilis (BS), A. niger (AN), or Trichoderma reesei (TR) in reducing lignin and pectin has been reported. The results showed when using a co-culture of AN and TR with a ratio of 1:1 (v/v) and of BS and TR with a ratio of 2:1 (v/v), the lignin and pectin removal was 99.9%. With the addition of surfactant during hydrolysis of pretreated CPW using a AN:TV (2:1) co-culture, the yield of reducing sugar estimated to be higher than that of the control. Use of PEG 4000 as a surfactant had a positive effect on enhancing the yield of reducing sugar from coffee pulp waste (Ishwanto et al. 2019).

Production of Glucose from Corn Stover The cocktail of cellulase with other auxiliary enzymes and chemicals was studied by Yu et al. for improving the conversion rate of cellulose of corn stover to glucose (Yu et al. 2015). The highest increasing rate of glucan-to-glucose conversion (44.9%) was achieved at the optimum concentration of β-glucosidase (377 μg/g cellulose), and sodium thiosulphate (1 mg/ml sodium thiosulphate).

Higher Reducing Sugars from Rice Straw and Corn Stovers The value-added products, viz. bio-ethanol, bioplastic, γ-valerolactone (GVL), 5hydroxymethylfuroic acid (HMF) and levulinic acid, are produced from saccharification of lignocellulosic biomass. Efficient fungal biomass-degrading enzyme mixtures for saccharification of local lignocellulosic feedstock were developed. Higher reducing sugars were obtained using BCC5776 enzyme cocktail as compared to Acremonium cellulose from hydrolysis of pretreated rice straw and corn stovers. The reducing sugar will be further employed as sugar feedstock for production of ethanol or commodity chemicals (Wanmolee et al. 2014).

92

5.9.1.2

5 Pectinase in Degradation of Lignocellulosic Wastes

Conversion of Lignocellulosic Waste into Biofuels

Lignocellulosic Agave Residues into Liquid Biofuels Thousands of tons of solid residues per year are generated by Agave-based alcoholic beverage companies in Mexico. This agave is being hoped for biofuel production due to their abundance and favorable sustainability characteristics. Ammonia fiber expansion (AFEX)-pretreated agave residues can be effectively hydrolyzed at high solids loading using an optimized commercial enzyme cocktail. Ammonia fiber expansion (AFEX™ )1 is a leading pretreatment that can increase sugar conversion in an effective and sustainable manner. AFEX is a “dry-to-dry” alkaline treatment that requires minimum water inputs and does not generate liquid waste streams unlike most thermochemical pretreatments. The enzyme cocktail comprises of (i) Cellic® CTec3 comprised cellulase enzymes expressed in T. reesei (including endo- and exo-cellobiohydrolases, accessory activities, bacterial beta-glucosidase, and minor amounts of hemicellulases), (ii) Cellic® HTec3 (composed of xylanase and xylosidase activities as well as auxiliary enzyme activities) and (iii) Multifect® Pectinase (composed of diverse hemicellulase activities such as arabinofuranosidase, xylan esterase, pectinase, pectin lyase, alpha-galactosidase, mannanase, mannosidase, and other activities). Ethanol production after pretreatment by AFEX using enzyme cocktail has been optimized. For 6% glucan loading (17–19% TS), the enzyme cocktail has been standardized as of A. tequilana bagasse were 78:22:0, for A. salmiana bagasse, they were 67:27:6; for A. tequilana leaf, they were 80:18:2; and for A. salmiana leaf, they were 78:10:12. The total enzyme concentration was kept constant at 20 mg protein/g glucan. An optimized commercial enzyme cocktail (25 mg protein/g glucan) can be used for hydrolysis of AFEX-pretreated agave residues at high solids loading. The hydrolysis resulted in production of 85% sugar conversions and 40 g/l bio-ethanol titers. These results show that AFEX technology has considerable potential to convert lignocellulosic agave residues to bio-based fuels and chemicals in a biorefinery (Flores-Gómez et al. 2018).

Production of Biogas from Pectin and Lignocelluloses Waste26 Has Bee99A1 Patent EP2661499A1 comprises process for producing biogas from pectin and lignocellulose containing material. They had employed beet pulp waste for production of biogas. The patent process can be applied for the material comprises or is derived from potato pulp, sweet potato pulp, cassava pulp, sugar beet pulp, apple pulp, pear pulp, banana pulp, orange pomace, grape pomace, lemon pulp, pineapple pulp as well as waste residue from carrots, cereal straw, wheat straw, palm fronds, palm fruits, empty palm fruit bunches, palm residues, switchgrass, Miscanthus, rice hulls, municipal solid waste, industrial organic waste, office paper, and bagasse of sugar cane or mixtures thereof.

5.9 Application of Pectinases in Deconstruction …

93

Conversion of the Lignocellulosic Biomass Arundo Donax into Fuel Ethanol A novel pectate lyase, named Paenxyl Pel, was identified from Paenibacillus xylanilyticus 2-6L3 isolated from mature compost obtained from agro-industrial wastes. The pectate lyase from P. xylanoliticus showed unusual activity combining traits of pectate lyase and pectin lyase. Its cocktail with an improved variant of GH51 a-larabinofuranosidase from Pleurotus ostreatus by directed evolution has been applied for enzymatic saccharification of Arundo donax (perennial cane) and production of fuel ethanol (Giacobbe 2015).

Biofuel Production by Thermatoga the Thermophilic Plant Biomass Degraders Thermophilic plant biomass degraders fall into one of two categories: cellulosomal (i.e., multi-enzyme complexes) and non-cellulosomal (i.e., “free” enzyme systems) Thermotoga species are typically thought to use a free-enzyme system. Hyperthermophilic microorganisms are of considerable industrial interest because they are natural sources of heat-stable enzymes useful in biotechnology. Thermotoga maritima grows optimally at 80 °C (maximal growth at 90 °C), and the genus represents one of the most thermophilic eubacterial genera yet identified. T. maritima metabolizes a variety of polysaccharides, such as xylan, starch, cellulose, and pectin. This family of microbes facilitates the isolation of highly thermostable enzymes that can be used as valuable catalysts for industrial purposes. The use of thermostable pectinolytic enzymes will lead to thermophilic lignocelluloses deconstruction for producing biofuel form high pectin content wastes which are poorly soluble and viscous at normal processing temperatures.

5.9.1.3

Retting and Degumming Lignocellulosic Wastes

Retting of flax (Linum usitatissimum L.) for production of linen for textiles and fibers for thinner papers (Akin et al. 2007). Flax (L. usitatissimum L.) is an important commercial crop that supplies both linseed and bast fibers for multiple applications. Retting, which is a microbial process, separates industrially useful bast fibers from non-fiber stem tissues. While several methods (i.e., water- and dew-retting) are used to ret flax, more recently enzymes have been evaluated to replace methods used currently.

Retting and Degumming of Plant Bast Fibers Plant bast fibers are the soft fibers formed in groups outside the xylem, phloem or pericycle, e.g., ramie and sun hemp. The textile making needs the removal of gum from fibers before it is used for further process. The degumming treatment performed using

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5 Pectinase in Degradation of Lignocellulosic Wastes

chemicals causes pollution, increases toxicity, and is non-biodegradable. Biotechnological degumming process employing pectinases in combination with xylanases is a green technology and economic alternative to the said problem (Kapoor et al. 2001). In an application study, enzymatic depolymerization of ramie was performed for fiber degumming by an alkaline polygalacturonase (PN32) from Bacillus paralicheniformis CBS32 isolated from kimchi (Rahman et al. 2019). As compared to the commercial pectinase, the pectinase produced by PN32 showed 28% higher depolymerization.

Fibers from Banana Waste with Extract of Pectinase Banana fibers are lightweight, soft fibers obtained from the pseudo-stem of the banana. After harvesting the fruit, the stem of the banana is discarded as waste material. The banana fiber can be used for the production of the Kraft paper (Jacob et al. 2008). Jocob et al. (2008) performed a study of degradation of the fiber of dried banana and hand stripped with crude extract of pectinase from the S. lydicus lineage and observed that a gradual release of reducing sugar was an effective treatment.

5.9.1.4

Improving Nutritional Quality of Animal Feed

The enzyme cocktail comprising of pectinases are used for the production of animal feeds. Usually, a feed enzyme preparation is a multi-enzyme cocktail containing glucanases, xylanases, proteinases, pectinases, and amylases. The addition of enzyme cocktail reduces viscosity of lignocellulosic wastes used for animal feed production. This enhanced absorption of nutrients because of nutrients liberated by hydrolysis of non-degradable fibers, or blocked nutrients liberated from these fibers. Recent studies in broilers conducted with and without the use of antibiotics suggest that feed additives significantly modify the immune-derived inflammatory response under stress conditions resulting in better mortality, weight gain, feed conversion, and bone strength in the broilers (Petersen 2001). Studies have been carried out to evaluate the action of commercial and noncommercial cellulase and pectinase on rice husk and Tifton 85 hay hydrolyses for aiming at improving substrates’ nutritional quality for animal feed.

5.9.1.5

Treatment of Pectic Wastewater

The pectinaceous materials present in wastewater from the citrus-processing industry are difficult for decomposition by microorganisms by the activated-sludge treatment. Studies on new wastewater treatment process by using an alkalophillic microorganism have been done. Alkalophilic Bacillus sp. (GIR 621) was reported to produce an extracellular endo-pectate lyase in alkaline media at pH 10.0. The treatment of wastewater was found to be useful using alkalophilic Bacillus sp. (GIR 621). For

5.9 Application of Pectinases in Deconstruction …

95

treatment of wastewater from citrus-processing industries, various processes have been investigated, which include physical dewatering, spray irrigation, chemical coagulation, direct activated-sludge treatment, and chemical hydrolysis followed by methane fermentation, but these processes have low efficiency. The chemical resistance of the pectic substances causes high treatment cost, long treatment periods, and process complexity. A soft-rot pathogen, E. carotovora (FERM P7576), which secrets endo-pectate lyase, has been reported to be useful in the pretreatment of pectinaceous wastewater. Pretreatment of the wastewaters from citrus-processing industries with pectinolytic enzymes leads to effective removal of pectinaceous materials. The process is suitable for decomposition by activated-sludge treatment (Beg et al. 2000).

5.9.1.6

Detoxification of Tobacco Stalk Waste

The processing of raw materials such as tobacco stalk (TS) generates huge quantities of agro-industrial waste residues. The problem is more in a country like China which has abundant crop residue. It is expected that more than one hundred thousand metric tons of tobacco stalk are discarded annually in China (Zhong et al. 2010). The tobacco industries in China have to search for eco-friendly process for the disposal of tobacco waste due to strict environmental legislation. The use of tobacco stalk will minimize the soil pollution due to its improper disposal and also generate additional income for the tobacco producers (Hu et al. 2015).

5.10 Special Approaches to Lignocellulosic Wastes The insufficient technology for biofuel production from lignocellulosic materials has given rise to inefficient and costly process for the production of fermentable sugars from the complex lignocellulosic raw materials. Hence, studies were conducted to produce enzyme cocktail comprising of endoglucanases, exo-glucanase, pectate lyases, cutinase, swollenin, xylanase, acetyl xylan esterase, beta-glucosidase, and lipase derived from Escherichia coli or tobacco chloroplast. For this purpose, the genes from bacteria or fungi were expressed in E. coli or tobacco chloroplasts. Pectate lyases or endoglucanase can be produced up to 49, 64, and 10, 751 million units annually per acre of tobacco based on observed expression levels. Plant production cost of endoglucanase is 3100-fold, and pectate lyase is 1057 or 1480-fold lower than the same recombinant enzymes sold commercially, produced via fermentation. The recombinant enzymes sold commercially, produced by fermentation are high-cost processes about 3100-fold, and pectate lyase is 1057 or 1480-fold when compared to plant production. Chloroplast-derived enzymes are thermostable and possess enzyme activity at a wide pH range as compared to the enzymes expressed in E. coli. The enzymes expressed in E. coli need purification, whereas the plant crude

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5 Pectinase in Degradation of Lignocellulosic Wastes

extracts showed higher enzyme activity without purification. With the higher concentration of proteins produced in plant chloroplast, the enzymes can be used directly without incurring purification costs. Chloroplast-derived crude extract enzyme cocktails yielded higher yields of glucose from filter paper, pinewood, or citrus peel than commercial cocktails. Furthermore, pectate lyase trans-plastomic plants showed enhanced resistance to Erwinia soft rot. The plant-derived enzyme cocktails for production of fermentable sugars from lignocellulosic biomass are a novel process now available. Limitations of higher cost and lower production capacity of fermentation systems are addressed by chloroplast-derived enzyme cocktails (Verma et al. 2010).

5.11 Conclusion Biotechnological solutions or environmental sustainability are modern studies that help in the growth of the nation and are an advantage for the wellbeing of human beings for the present and future generations. Deteriorating reserves of fossil fuel and petroleum derivatives, rising oil prices, alarm about environmental impact, and supply insecurity now need environmentally sustainable energy sources. The alternative approach to petrochemical production of fuels and chemicals is through microbial fermentation of plant material as renewable feedstock. Lignocellulose waste has candidature as the most prevalent and rich source of carbon in nature. It is now targeted as the source that could provide a sufficient amount of feedstock to assure the global energy and chemical needs in a renewable manner. There is the enormous potential of pectinase in various sectors of industries whenever degradation of pectin is needed. There is a need of research in screening of microorganisms for pectinase production. Extensive studies have to be carried on the optimization of conditions for the production of microbial pectinases. Study of the molecular aspects of pectinases and engineering of enzymes that are stouter with respect to their pH and temperature kinetics is needed. Studies on protein engineering and site-directed mutagenesis need to be focused in the coming times.

References Ahlawat S, Mandhan R, Dhiman S, Kumar R, Sharma J (2008) Potential application of alkaline pectinase from Bacilllus subtilis SS in pulp and paper industry. Appl Biochem Biotechnol 149:287–293 Akin D, Condon B, Sohn FJ, Dodd R, Rigsby L (2007) Optimization for enzyme-retting of flax with pectate lyase. Ind Crops Prod 25:136–146 Alonso J, Canet W, Hawell N, Alique R (2003) Purification and characterization of carrot (Daucas carota L) pectinesterase. J Sci Food Agric 83:1600–1606 Aminzadeh S, Naderi-Manesh H, Khadesh K (2007) Isolation and characterization of polygalacturonase produced by Tetracoccosporium sp. Iran J Chem Eng 26:47–54 Angayarkanni J, Palaniswamy M, Murugesan S, Swaminathan K (2002) Improvement of tea leaves fermentation with Aspergillus sp. pectinase. J Biosci Bioeng 94:299–303

References

97

Anisa S, Girish K (2014) Pectinolytic Activity of Rhizopus sp. and Trichoderma viride. Int J Res Pure Appl Microbiol 4:28–31 Arenas-Ocampo M, Evangelista-Lozano S, Arana-Errasquin R, Jimenez-Aparicio A, Davila-Ortız G (2003) Softening and biochemical changes of sapote mamey fruit (Pouteria sapota) at different development and ripening stages. J Food Chem 27:91–95 Arias C, Burns J (2002) A pectinmethylesterase gene associated with a heatstable. J Agric Food Chem 50:3465–3472 Aro N, Pakula T, Penttila M (2005) Transcriptional regulation of plant cell wall degradation by filamentous fungi. FEMS Microbiol Rev 29:719–739 Assis S, Fernandes P, Ferreira B, Trevisan H, Cabral J, Oliveira O (2004) Screening of supports for the immobilization of pectin-methylesterase from acerola (Malpighia glabra L.). J Chem Technol Biotechnol 79:277–280 Balat M, Balat H (2009) Recent trends in global production and utilization of bio-ethanol fuel. Appl Energ 86:2273–2282 Barnby F, Morpeth F, Pyle D (1990) Endopolygalacturonase production from Kluyveromyces marxianus. I Resolution, purification and partial characterization of the enzyme. Enzyme Microbial Technol 12:891–897 Bazzolli D, Ribon A, de Queiroz MV, de Araújo E (2006) Molecular characterization and expression profile of pectin-lyase-encoding genes from Penicillium. Can J Microbiol 52:1070–1077 Beg Q, Bhushan B, Kapoor M, Hoondal G (2000) Production and characterization of thermostable xylanase and pectinase from Streptomyces sp. QG-11-3. J Ind Microbiol Biotechnol 24:396–402 Brent R, O’Neill M, Debra M (2001) Pectins: structure, biosynthesis, and oligogalacturonide-related signaling. Phytochemistry 57:929–967 Bruhlmann F (1995) Production and characterization of an extracellular pectate lyase from an Amycolata sp. Appl Environ Microbiol 61:3580–3585 Buchanan B, Gruissem W, Jones R (eds) (2015) Biochemistry and molecular biology of plants. American Society of Plant Physiologists, Wiley, Blackwell Centis S, Dumas B, Fournier J, Marolda M, Esquerré-Tugayé M (1996) Isolation and sequence analysis of Clpgl, a gene coding for an endopolygalacturonase of the phytopathogenic fungus Colletotrichum lindemuthianum. Gene 170:125–129 Centis S, Guillas I, Sejalon N, Esquerre-Tugaye M (1997) Endo-polygalacturonase gene from Colletotrichum lindemuthianum: cloning of CLPG2 and comparison of its expression to that of CLPG1 during saprophytic and parasitic growth of the fungus. Mol Plant Microbe Inter 10:769– 775 Chen W, Hsieh H, Tseng T (1998) Purification and characterization of a pectin lyase from Pythium splendens infected cucumber fruits. Botanical Bull Acad Sinica 39:181–186 Chen Q, Jin Y, Zhang G et al (2012) Improving production of bioethanol from duckweed (Landoltia punctata) by pectinase pretreatment. Energies 5:3019–3032 Cornick N, Jensen N, Stahl D, Hartman P, Allison M (1994) Lachnospira pectinoschiza sp. nov., an anaerobic pectinophile from the pig intestine. Int J Syst Bacteriol 44:87–93 Corredig M, Kerr W, Wicker L (2000) Separation of thermostable pectinmethylesterase. J Agric Food Chem 48:4918–4923 Daniell H, Ribeiro T, Lin S, Saha P, McMichael C, Chowdhary R, Agarwal A (2019) Validation of leaf and microbial pectinases: commercial launching of a new platform technology. Plant Biotechnol J 17:1154–1166 Das A, Bhattacharya S, Reddy N, Sundara R (2013) Improved production and purification of pectinase from Streptomyces sp. GHBA10 isolated from Valapattanam mangrove habitat, Kerala. India. Int Res J Biol Sci 2:16–22 Dey A, Karmakar M, Ray R (2011) Extracellular thermostable polygalacturonase from Bacillus sp. AD1. Der Pharm Lett 3:358–367 Dinnella C, Lanzarini G, Ercolessi P (1995) Preparation and properties of an immobilized solubleinsoluble pectinlyase. Process Biochem 30:151–157

98

5 Pectinase in Degradation of Lignocellulosic Wastes

Dixit V, Kumar A, Pant A, Khan M (2004) Low molecular mass pectate lyase from Fusarium moniliforme: similar modes of chemical and thermal denaturation. Biochem Biophys Res Commun 315:477–484 Doesburg J (1965) PecUc substances m fresh and preserved fruits and vegetables. Communication No. 25 Wageningen, The Netherlands Downie B, Dirk L, Hadfield K, Wilkins T, Bennet A, Bradford K (1998) A gel diffusion assay for quantification of pectinmethylesterase activity. Anal Biochem 264:149–157 Drori N, Kramer-Haimovich H, Rollins J, Dinoor A, Okon Y, Pines O, Prusky D (2003) External pH and nitrogen source affect secretion of pectate lyase by Colletotrichum gloeosporioides. Appl Environ Microbiol 69:3258–3262 El-Batal A, Osman E, Shaima I (2013) Optimization and characterization of polygalacturonase enzyme produced by gamma irradiated Penicillium citrinum. J Chem Pharm Res 5:336–347 Favey S, Bourson C, Bertheou Y, Kotoujansky A, Boccora M (1992) Purification of the acidic pectate lyase of Erwinia chrysanthemi 3937 and sequence analysis of the corresponding gene. J Gen Microbiol 138:499–508 Fayyaz A, Asbi B, Ghazali H, Che Man T, Jinap S (1993) Pectinesterase extraction from papaya. Food Chem 47:183–185 Flipphi M, Panneman H, van der Veen P, Visser J, de Graaff L (1993) Molecular cloning, expression and structure of the endo-1,5-alpha-l-arabinase gene of Aspergillus niger. Appl Microbiol Biotechnol 40:318–326 Flores-Gómez C, Escamilla Silva E, Zhong C, Dale B, da Costa Sousa L, Balan V (2018) Conversion of lignocellulosic agave residues into liquid biofuels using an AFEX™ -based biorefinery. Biotechnol Biofuels 11:1–18 Fogarty W, Kelly C (1983) Pectin enzymes. In: Fogarty W (ed) Microbial enzymes and biotechnology. Applied Science, London, pp 131–182 Friedrich J, Cimerman A, Steiner W (1990) Production of pectolytic enzymes by Aspergillus niger, effect of inoculum size and potassium hexacianoferrate II-trihydrate. Appl Microbiol Biotechnol 33:377–381 Gainvors A, Frezier V, Lemaresquier H, Lequart C, Aigle M, Belarbi A (1994) Detection of polygalacturonase, pectin lyase and pectinesterase activities in Saccharomyces cerevisiae strain. Yeast 10:1311–1319 Gainvors A, Nedjaoum N, Gognies S, Muzart M, Nedjma M, Belarbi A (2000) Purification and characterization of acidic endo-polygalacturonase encoded by the PGL-1 gene from Saccharomyces cerevisiae. FEMS Microbiol Lett 183:131–135 Gao S, Choi G, Shain L, Nuss D (1996) Cloning and targeted disruption of enpg-1, encoding the major in-vitro extracellular endopolygalacturonase of the chestnut blight fungus, Cryphonectria parasitica. Appl Environ Microbiol 62:1984–1990 Giacobbe S (2015) New wild-type and engineered biocatalysts and biosystems for conversion of the lignocellulosic biomass Arundo donax into fuel ethanol. Ph.D. Thesis Guo W, Gonzalez-Candelas L, Kolattukudy P (1996) Identification of a novel pelD gene expressed uniquely in plant by Fusarium solani f. sp. pisi (Nectria haematococca, mating type VI) and characterization of its protein as an endo-pectate lyase. Arch Biochem Biophys 332:305–312 Hadj-Taieb N, Ayadi M, Trigui S, Bouabdollah F, Gargouri A (2002) Hyper production of pectinase activities by fully constitutive mutant (CT1) of Penicillium occitanis. Enzyme Microbial Technol 30:662–666 Harholt J, Suttangkaku A, Henrik V, Scheller H (2010) Biosynthesis of pectin 1: update on pectin biosynthesis. Plant Physiol 153:384–395 Hasunuma T, Fukusaki E, Kobayashi A (2003) Methanol production is enhanced by expression of an Aspergillus niger pectin methylesterase in tobacco cells. J Biotechnol 106:45–52 Herron S, Jurnak F (2003) Mechanistic lessons from structural studies of the pectate lyases. In: Voragen F, Schols H, Visser R (eds) Advances in pectin and pectinase research. Springer, Netherlands, Dordrecht, pp 221–233

References

99

Hoondal G, Tewari R, Tewari R, Dahiya N, Beg Q (2002) Microbial alkaline pectinases and their industrial applications: a review. Appl Microbiol Biotechnol 5:409–418 Hu R, Wang J, Li H, Ni H, Chen Y, Zhang Y, Xiang S, Li H (2015) Simultaneous extraction of nicotine and solanesol from waste tobacco materials by the column chromatographic extraction method and their separation and purification. Sep Purif Technol 146:1–7 Huang Q, Allen C (1997) An exo-poly-a-d-galacturonosidase, Peh B, is required for wild type virulence of Ralstonia solanacearum. J Bacteriol 179:7369–7378 Huang R, Su R, Qi W et al (2011) Bioconversion of lignocellulose into bioethanol process intensification and mechanism research. BioEnerg Res 4:225–245 Innocenzo M, Lajalo F (2001) Effect of gamma irradiation on softening changes and enzyme activities during ripening of papaya fruit. J Food Biochem 25:19–27 Iswanto T, Hendrianie N, Shovitri M, Altway A, Widjaja T (2019) The effect of mixed biological pretreatment and PEG 4000 on reducing sugar production from coffee pulp waste. Int J Technol 10:453–462 Jacob N, Niladevi K, Anisha G, Prema P (2008) Hydrolysis of pectin: an enzymatic approach and its application in banana fiber processing. Microbiol Res 163:538–544 Jahn C, Charkowski A, Willis D (2008) Evaluation of isolation methods and RNA integrity for bacterial RNA quantitation. J Microbiol Methods 75:318–324 Jayani R, Saxena S, Gupta R (2005) Microbial pectinolytic enzymes: a review. Process Biochem 40:2931–2944 Johnson A, Ayodele O, Afolake O, Christiana O (2012) Production of cellulase and pectinase from orange peels by fungi. Nat Sci 10:107–112 Kapoor M, Beg Q, Bhushan B, Singh K, Dadhich K, Hoondal G (2001) Application of an alkaline and thermostable polygalacturonase from Bacillus Sp. MG-Cp-2 in degumming of Ramie (Boehmeria nivea) and Sunn Hemp (Crotalaria juncea) bast fibers. Process Biochem 36:803–807 Karam N, Belarbi A (1995) Detection of polygalacturonase and pectinesterases in lactic acid bacteria. World J Microbiol Biotechnol 11:559–563 Karlsson M, Durling M, Choi J, Kosawang C et al (2015) Insights on the evolution of mycoparasitism from the genome of Clonostachys rosea. Genome Biol Evol 27:465–480 Kashyap D, Chandra S, Kaul A, Tewari R (2000) Production, purification and characterization of pectinase from a Bacillus sp. DT7. World J Microbiol Biotechnol 16:277–282 Kashyap D, Vohra P, Chopra S, Tewari R (2001) Applications of pectinases in the commercial sector: a review. Bioresour Technol 77:215–227 Kashyap D, Soni S, Tewari R (2003) Enhanced production of pectinase by Bacillus sp. DT7 using solid state fermentation. Bioresour Technol 88:251–254 Kataria R, Ghosh S (2011) Saccharification of Kans grass using enzyme mixture from Trichoderma reesei for bioethanol production. Bioresource Technol 102:9970–9975 Kaur A, Mahajan R, Singh A, Garg G, Sharma J (2011) A novel and cost effective methodology for qualitative screening of alkalo-thermophilic cellulase free xylano-pectinolytic microorganisms using agricultural wastes. World J Microbiol Biotechnol 27:459–463 Kawano C, Chellegatti M, Said S, Fonseca M (1999) Comparative study of intracellular and extracellular pectinases produced by Penicillium frequentans. Biotechnol Appl Biochem 29:133–140 Kertesz Z (1951) The pectin substances. J Amer Pharma Assoc 41:54 (Interscience Publishers, New York) Kitamoto N, Yoshino-Yasuda S, Ohmiya K, Tsukagoshi N (2001) A second pectin lyase gene (pel2) from Aspergillus oryzae KBN616: its sequence analysis and overexpression, and characterization of the gene products. J Biosci Bioeng 91:378–381 Klug-Santner B, Schnitzhofer W, Vrsanska M, Weber J, Agrawal P, Nierstrasz V, Guebitz G (2006) Purification and characterization of a new bioscouring pectate lyase from Bacillus pumilus BK2. J Biotechnol 121:390–401

100

5 Pectinase in Degradation of Lignocellulosic Wastes

Kobayashi T, K. Koike T, Yoshimatsu N, Higaki A, Suzumatsu, Ozawa T (1999) Purification and properties of a low-molecular weight, high-alkaline pectate lyase from an alkaliphilic strain of Bacillus. Biosci Biotechnol Biochem 63:72–75 Koboyashi T, Higaki N, Yajima N, Suzumatsu A, Haghihara H, Kawai S, Ito S (2001) Purification and properties of a galacturonic acid releasing exopolygalacturonase from a strain of Bacillus. Biosci Biotechnol Biochem 65:842–847 Kothari M, Baig M (2013) Production and characterization of extracellular polygalacturonase by Erwinia carotovora MTCC 1428. Int J Adv Biotechnol Res 4:981–998 Kou L, Song Y, Zhang X et al (2017) Comparison of four types of energy grasses as lignocellulosic feedstock for the production of bio-ethanol. Bioresource Technol 241:424–429 Kuhad R, Kapoor M, Rustagi R (2004) Enhanced production of an alkaline pectinase from Streptomyces sp. RCK-SC by whole-cell immobilization and solid-state cultivation. World J Microbiol Biotechnol 20:257–263 Kumar S, Palanivelu P (1999) Purification and characterization of an exoplolygalacturonase from the thermophilic fungus, Thermomyces lanuginosus. World J Microbiol Biotechnol 15:643–646 Kumar A, Sharma R (2012) Production of alkaline pectinase by bacteria isolated from decomposing fruit materials. J Phytopathol 4:1–5 Kusters-van Someren M, Flipphi M, de Graaff L, van den Broeck H, Kester H et al (1992) Characterization of the Aspergillus niger pelB gene: structure and regulation of expression. Mol Gen Genet 234:113–120 Lagaert S, Belien T, Volckaert G (2009) Plant cell walls: protecting the barrier from degradation by microbial enzymes. Semin Cell Dev Bio 20:1064–1073 Laing E, Pretorius I (1993) Co-expression of an Erwinia crysanthemi pectate lyase-encoding gene (pel E) and an E. carotovora polygalacturonase-encoding gene (peh 1) in Saccharomyces cerevisiae. Appl Microbiol Biotechnol 75:149–158 Lara-Márquez A, Zavala-Páramo MG, López-Romero E, Calderón-Cortés N, Lopez-Gomez R, Conejo-Saucedo U et al (2011) Cloning and characterization of a pectin lyase gene from Colletotrichum lindemuthianum and comparative phylogenetic/structural analyses with genes from phytopathogenic and saprophytic/opportunistic microorganisms. BMC Microbiol 11:1–14 Latendresse M (2013) Computing Gibbs free energy of compounds and reactions in MetaCyc Laurent F, Kotoujansky A, Bertheau Y (2000) Overproduction in Escherichia coli of the pectin methylesterase A from Erwinia chrysanthemi 3937: one-step purification, biochemical characterization, and production of polyclonal antibodies. Can J Microbiol 46:474–480 Leal T, de Sa-Nogueira I (2004) Purification, characterization and functional analysis of an endoarabinanase (AbnA) from Bacillus subtilis. FEMS Microbiol Lett 241:41–48 Liao C, Revear L, Hotchkiss A, Savary B (1999) Genetic and biochemical characterization of an exo polygalacturonase and a pectate lyase from Yersinia enterolitica. Can J Microbiol 45:396–403 Lygin A, Upton J, Dohleman F et al (2011) Composition of cell wall phenolics and polysaccharides of the potential bioenergy crop—Miscanthus. Global Change Biol Bioenergy 3:333–345 Macdonald H, Evans R (1996) Purification and properties of apple pectinesterase. J Sci Food Agric 70:321–326 Maceira F, Pietro A, Roncero M (1997) Purification and characterization of a novel exopolygalacturonase from Fusarium oxysporum f. sp. lycopersici. FEMS Microbiol Lett 154:37–43 Maldonaldo M, Saad A (1998) Production of pectinesterase and polygalacturonase by Aspergillus niger in submerged and solid state systems. J Ind Microbiol Biotechnol 20:34–38 Margo P, Varvaro L, Chilosi G, Avanzo C, Balestra G (1994) Pectinolytic enzymes produced by Pseudomonas syringae pv. glycinea. FEMS Microbiol Lett 117:1–6 Martins E, Silva D, Da Silva R, Gomes E (2002) Solid state production of thermostable pectinases from thermophilic Thermoascus aurantiacus. Process Biochem 37:949–954 Martin N, Guez M, Sette LD, Da Silva R, Gomes E (2010) Pectinase production by a Brazilian thermophilic fungus Thermomucor indicae-seudaticae N31 in solid-state and submerged fermentation. Microbiol 79:306–313

References

101

Mayans O, Scott M, Connerton I, Gravesen T et al (1997) Two crystal structures of pectin lyase A from Aspergillus reveal a pH driven conformational change and striking divergence in the substrate-binding clefts of pectin and pectate lyases. Struct Lond Engl 5:677–689 McCready R, Gee M (1960) Plant pectin analysis, determination of pectic substances by paper chromatography. J Agric Food Chem 8:510–513 Mohamadi A, Shahbazi S, Behgar M, Fard S, Askari H (2014) A study of pectinase enzyme activity changes in gamma-irradiated Trichoderma reesei mutants. Intl J Farm Alli Sci 23:555–561 Moharib S, El-Sayed S, Jwanny E (2000) Evaluation of enzymes produced by yeast. Nahrung 44:47–51 Mohnen D (2008) Pectin structure and biosynthesis. Curr Opin Plant Biol 11:266–277 Mota T, de Oliveira D, Marchiosi R, Ferrarese-Filho O, dos Santos W (2018) Plant cell wall composition and enzymatic deconstruction. Aims Bioeng 5:63–77 Muller-Maatsch J, Bencivenni M, Caligiani A, Tedeschi T et al (2016) Pectin content and composition from different food waste streams. Food Chem 201:37–45 Mutenda K, Korner R, Christensen T, Mikkelsen J, Roepstorff P (2002) Application of mass spectrometry to determine the activity and specificity of pectin lyase A. Carbohydr Res 337:1217–1227 Nadaroglu H, Taskın E, Adıgüzel A, Güllüce M, Demir N (2010) Production of a novel pectin lyase from Bacillus pumilus (P9), purification and characterization and fruit juice application. Romanian Biotechnol Lett 15:5167–5176 Nagai M, Katsuragi T, Terashita T, Yoshikawa K, Sakai T (2000) Purification and characterization of an endo-polygalacturonase from Aspergillus awamori. Biosci Biotechnol Biochem 64:1729–1732 Naumov G, Naumova E, Aigle M, Masneuf I, Belarbi A (2001) Genetic reidentification of the pectinolytic yeast strain SCPP as Saccharomyces bayanus var. uvarum. Appl Microbiol Biotechnol 55:108–111 Nozaki K, Miyairi K, Hizumi S, Fukui Y, Okuno T (1997) Novel exopolygalacturonases produced by Alternaria mali. Biosci Biotechnol Biochem 61:75–80 Oberoi H, Vadlani P, Nanjundaswamy A et al (2011) Enhanced ethanol production from Kinnow mandarin (Citrus reticulata) waste via a statistically optimized simultaneous saccharification and fermentation process. Bioresource Technol 102:1593–1601 Oliveira D, Finger-Teixeira A, Mota T et al (2016) Ferulic acid: A key component in grass lignocellulose recalcitrance to hydrolysis. Plant Biotechnol J 13:1224–1232 O’Neill M, Ishii T, Albersheim P, Darvill A (2004) Rhamnogalacturonan II: Structure and function of a borate cross-linked cell wall pectic polysaccharide. Annu Rev Plant Biol 55:109–139 Payasi A, Sanwal R, Sanwal G (2009) Microbial pectate lyases: characterization and enzymological properties. World J Microbiol Biotechnol 25:1–14 Pereira G, Cipriani M, Wisbeck E, Souza O et al (2017) Onion juice waste for production of Pleurotus sajor-caju and pectinases. Food Bioprod Process 106:11–18 Petersen S (2001) Enzymes to upgrade plant nutrients. Feed Mix 9:12–15 Pietro A, Roncero M (1996) Purification and characterization of an exo-polygalacturonase from the tomato vascular wilt pathogen Fusarium oxysporum f.sp. lycopersici. FEMS Microbiol Lett 145:295–298 Pitkanen K, Heikinheimo R, Pakkanen R (1992) Purification and characterization of Erwinia chrysatnthemi B374 pectin methylesterase produced by Bacillus subtilis. Enzyme Microbial Technol 14:832–836 Polizeli M, Jorge J, Terenzi H (1991) Pectinase production by Neurospora crassa: purification and biochemical characterization of extracellular polygalacturonase activity. J Gen Microbiol 137:1815–1823 Rahman M, Choi Y, Kim Y, Park C, Yoo J (2019) Production of novel polygalacturonase from Bacillus paralicheniformis CBS32 and application to depolymerization of ramie fiber. Polymers 11:1–20

102

5 Pectinase in Degradation of Lignocellulosic Wastes

Rastogi M, Shrivastava S (2017) Recent advances in second generation bioethanol production: an insight to pretreatment, saccharification and fermentation processes. Renew Sust Energ Rev 80:330–340 Rehman H, Qader S, Aman A (2012) Polygalacturonase: production of pectin depolymerising enzyme from Bacillus licheniformis KIBGE IB-21. Carbohydr Polym 90:387–391 Reymond P, Deleage G, Rascie C, Fevre M (1994) Cloning and sequence analysis of a polygalacturonase-encoding gene from the phytopathogenic fungus Sclerotinia sclerotiorum. Gene 146:233–237 Rodrigues-Palenzuela P, Burr T, Collmer A (1991) Polygalacturonase is a virulence factor in Agrobacterium tumifaciens biovar 3. J Bacteriol 173:6547–6552 Sakai T, Sakamoto T (1990) Purification and some properties of a protopectin-solubilizing enzyme that has potent activity on sugar beet protopectin. Agric Biol Chem 53:1213–1223 Sakai T (1992) Degradation of pectins. In: Winkelmann G (ed) Microbial degradation of natural products. VCH, Weinheim, pp 57–81 Sakai T, Sakamoto T, Hallaert J, Vandamme E (1993) Pectin, pectinase and protopectinase: production, properties and applications. Adv Appl Microbiol 39:231–294 Sakamoto T, Hours R, Sakai T (1994) Purification, characterization and production of two pectictranseliminases with protopectinase activity from Bacillus subtilis. Biosci Biotechnol Biochem 58:353–358 Sapunova L, Mikhailova R, Lobanok A (1995) Properties of pectin lyase preparations from the genus Penicillium. Appl Microbiol Biochem 31:435–438 Schell M, Denny T, Huang J (1994) Extracellular virulence factors of Pseudomonas solanacearum: role in disease and their regulation. In: Kado C, Crosa J (eds) Molecular mechanism of bacterial virulence. Kluwer Academic Press, The Netherlands, pp 311–324 Semenova M, Grishutin S, Gusakov A, Okunev O, Sinitsyu A (2003) Isolation and properties of pectinases from the fungus Aspergillus japonicus. Biochem 68:559–569 Servili M, Begliomini A, Montedoro G, Petruccioli M, Federici F (1992) Utilisation of a yeast pectinase in olive oil extraction and red wine making processes. J Sci Food Agric 58:253–260 Shah P, Gutierrez-Sanchez G, Orlando R, Bergmann C (2009) A proteomic study of pectin-degrading enzymes secreted by Botrytis cinerea grown in liquid culture. Proteomics 9:3126–3135 Sharma D, Satyanarayana T (2012) Biotechnological potential of agro residues for economical production of thermoalkalistable pectinase by Bacillus pumilus dcsr1 by solid-state fermentation and its efficacy in the treatment of ramie fibers. Enz Res 2012:1–18 Sharma N, Rathore M, Sharma M (2013) Microbial pectinase: sources, characterization and applications. Rev Environ Sci Biotechnol 12:45–60 Shevchik V, Condemine G, Hugouvieux-Cotte-Pattat N, Robert-Baudouy J (1996) Characterization of pectin methylesterase B, an outer membrane lipoprotein of Erwinia chrysanthemi 3937. Mol Microbiol 19:455–466 Shevchik V, Robert-Baudouy J, Hugouvieux-Cotte-Pattat N (1997) Pectate lyase Pel 1 of Erwinia chrysanthemi 3937 belongs to a new family. J Bacteriol 179:7321–7330 Shevchik V, Condemine G, Robert-Baudouy J, Hugouvieux-Cote-Pattat N (1999) The exopolygalacturonate lyase PelW and the oligogalacturonate lyase Ogl, two cytoplasmic enzymes of pectin catabolism in Erwinia chrysanthemi 3937. J Bacteriol 181:3912–3919 Silva D, Martins E, Da Silva R, Gomes E (2002) Pectinase production by Penicillium viridicatum RFC3 by solid state fermentation using agricultural wastes and agro-industrial by-products. Braz J Microbiol 33:318–324 Singh S, Plattner H, Diekmann H (1999a) Exo polygalacturonate lyase from a thermophilic Bacillus sp. Enzyme Microbial Technol 25:420–425 Singh S, Ramakrishna M, Rao A (1999b) Optimization of downstream processing parameters for the recovery of pectinase from the fermented broth of Aspergillus carbonarious. Process Biochem 35:411–417

References

103

Singh S, Rao A (2002) A simple fractionation protocol for, and a comprehensive study of the molecular properties of two major endopolygalacturonases from Aspergillus niger. Biotechnol Appl Biochem 35:115–123 Singh A, Kaur A, Dua A, Ritu M (2015) An efficient and improved methodology for the screening of industrially valuable xylano-pectino-cellulolytic microbes. Enz Res 2015:1–7 Souza J, Silva T, Maia M, Teixeira M (2003) Screening of fungal strains for pectinolytic activity: endopolygalacturonase production by Peacilomyces clavisporus 2A.UMIDA.1. Process Biochem 4:455–458 Sunnotel O, Nigam P (2002) Pectinolytic activity of bacteria isolated from soil and two fungal strains during submerged fermentation. World J Microbiol Biotechnol 18:835–839 Swain M, Ray R (2010) Production, characterization and application of a thermostable exopolygalacturonase by Bacilus subtilis CM5. Food Biotechnol 24:37–50 Takao M, Nakaniwa T, Yoshikawa K, Terashita T, Sakai T (2000) Purification and characterization of thermostable pectate lyase with protopectinases activity from thermophilic Bacillus sp. TS 47. Biosci Biotechnol Biochem 64:2360–2367 Templeton M, Sharrock K, Bowen J, Crowhurst R, Rikkerink E (1994) The pectin lyase-encoding gene (pnl) family from Glomerella cingulata: Characterization of pnlA and its expression in yeast. Gene 142:141–146 Tierny Y, Bechet M, Joncquiert J, Dubourguier H, Guillaume J (1994) Molecular cloning and expression in Escherichia coli of genes encoding pectate lyase and pectin methylesterase activities from Bacteroides thetaiotaomicron. J Microbiol 76:592–602 Trigui-Lahiani H, Gargouri A (2007) Cloning, genomic organisation and mRNA expression of a pectin lyase gene from a mutant strain of Penicillium occitanis. Gene 388:54–60 van Dyk J, Sakka M, Sakka K, Pletschke B (2010) Characterisation of the multi-enzyme complex xylanase activity from Bacillus licheniformis SVD1. Enzym Microb Technol 47:174–177 Verma D, Kanagaraj A, Jin S, Singh N, Kolattukudy P, Daniell H (2010) Chloroplast-derived enzyme cocktails hydrolyse lignocellulosic biomass and release fermentable sugars. Faculty Biblio 8:332– 350 Wanmolee W, Sornlake N, Laosiripojana CV (2014) Development of efficient fungal biomassdegrading enzyme mixtures for saccharification of local lignocellulosic feedstock. Inter J Energy Power Eng 8:69–73 Warrilow A, Turner R, Jones M (1994) A novel form of pectinesterase in tomato. Phytochem 35:862–872 Wattad C, Freeman S, Dinoor A, Prusky D (1995) A nonpathogenic mutant of Colletotrichum magna is deficient in extracellular secretion of pectate lyase. Mol Plant-Microbe Interact 8:621–626 Wei Y, Li X, Yu L et al (2015) Mesophilic anaerobic co-digestion of cattle manure and corn stover with biological and chemical pretreatment. Bioresource Technol 198:431–436 Whitaker J (1990) Microbial pectinolytic enzymes. In: Fogarty W, Kelly C (eds) Microbial enzymes and biotechnology, 2nd edn. Elsevier Science, London, pp 133–176 Willats W, McCartney L, Mackie W, Knox J (2011) Pectin: cell biology and prospects for functional analysis. Plant Mol Biol 47:9–27 Yadav P, Singh V, Yadav S, Yadav K, Yadav D (2009) In-silico analysis of pectin lyase and pectinase sequences. Biochem 74:1049–1055 Yakoby N, Kobiler I, Dinoor A, Prusky D (2000) pH regulation of pectate lyase secretion modulates the attack of Colletotrichum gloeosporioides in avocado fruits. Appl Environ Microbiol 66:1026– 1030 Yu X, Liu Y, Meng J, Cheng Q, Zhang Z, Cui Y, Liu J, Teng L, Lu J, Meng Q, Ren X (2015) Optimization of enzyme complexes for efficient hydrolysis of corn stover to produce glucose. Pak J Pharm Sci 8:1115–1120 Zhong W, Zhu C, Shu M, Sun K, Zhao L, Wang C (2010) Degradation of nicotine in tobacco waste extract by newly isolated Pseudomonas sp. ZUTSKD. Bioresour Technol 10:6935–6941

Chapter 6

Lipase in Degradation of Lignocellulosic Wastes

Abstract Lipases enzyme are very important in the degradation of lignocellulosic wastes. The chapter here describes in detail the structure of lipase; classification of lipase enzyme including bacterial lipases; and reactions catalyzed by lipase enzyme. The chapter also focuses on active and catalytic residues of lipase enzyme; the catalytic mechanism of lipases; lipase-producing bacteria and fungi; and assays for lipase enzyme activity and applications of lipases enzyme. The chapter gives detailed information on lipase enzyme in degradation of lignocellulose wastes. This will help in the proper management of lignocellulosic waste. Keywords Lipase · Triacylglycerol · Active site · Fatty acids · Esterification · Biocatalysts

6.1 Introduction of Lipases Enzyme Lipases are the biocatalysts having many applications in food, dairy, detergent, and pharmaceutical industries (Ray 2015). Lipases are triacylglycerol acyl hydrolases (E.C. 3.1.1.3) which are involved in hydrolysis of triacylglycerol to glycerol and fatty acids. They are produced by plants, animals, and microorganisms (Thakur 2012). Microbial lipases can remain active under extreme temperature, pH, and organic solvents. Lipases are glycoproteins, and some bacterial lipases are lipoproteins. They act at the interface created by a hydrophobic liquid substrate in a hydrophilic liquid medium (Hong and Chang 1998).

6.2 Structure of Lipase The lipases enzyme has α/β-hydrolase fold, a catalytic triad (Ser-His-Asp/Glu) and lid to cover the active site. The lid is displaced during activation to make available the active site to the substrate. Lipases have molecular weight from 20 to about 60 kDa (Mehta et al. 2017). The lipases are both intracellular and extracellular. Intracellular

© Springer Nature Switzerland AG 2020 A. B. Gunjal et al., Enzymes in Degradation of the Lignocellulosic Wastes, https://doi.org/10.1007/978-3-030-44671-0_6

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lipases refer to the use of lipases while it is present in producing organism (RoblesMedina et al. 2009) and extracellular lipases are the lipases initially extracted from the organism producing it (Andualema and Gessesse 2012).

6.2.1 Three-Dimensional Structure of Lipases The three-dimensional structure reveals that lipase has a central β-sheet with eight various β-strands (β1 –β8 ) which are connected by 6 α-helices. The 3-D structure reveals that the molecular weights of lipase enzyme are from 19 to 60 kDa. The 3-D structure has α/β-hydrolase folding with a helical segment that covers the active site. The lipid aggregates help to open the lid and enhances enzyme activity, called as interfacial activation (Andualema and Gessesse 2012).

6.2.1.1

The Active and Catalytic Residue of Lipase Enzyme

The active site of lipase has three catalytic residues, viz. catalytic acid residue containing aspartate or glutamate; nucleophilic residue containing serine and histidine residue. The nucleophilic residue is seen in a highly conserved pentapeptide Gly-Xaa-Ser-Xaa-Gly (Ollis et al. 1992). The lipid breakdown initiates with the binding of a nucleophilic serine residue of lipase enzyme. The carbonyl group of the ester makes a bond with the oxygen of the serine hydroxyl group (Jaeger et al. 1999). This forms tetrahedral intermediate which is stabilized by hydrogen bonds between NH-groups and oxyanion. A proton is donated to weak ester oxygen bond by histidine residue and is cleaved. A covalent intermediate is formed. A water molecule breaks down the covalent intermediate serine–acyl complex and fatty acids are released (Jaeger et al. 1999). In the absence of water molecule, reverse esterification reaction takes place which forms an ester bond.

6.3 Classification of Lipases The lipases are distributed into three classes, viz. GX, GGGX, and Y (Fischer and Pleiss 2003). Yeasts and fungal lipases are grouped into five subclasses: two in the GX class, two in the GGGX class, and one in the Y class (Gupta et al. 2015). The lipases based on lipase engineering database are shown in Fig. 6.1.

6.3 Classification of Lipases

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Yeast and fungal lipases

GGGX class

GX class

Y

class

Yarrowia lipolytica antartica

Filamentous fungi lipase-like

Candida lipase

lipase A-like

Candida rugosa lipase-like

Candida antartica lipase B-like

Fig. 6.1 Lipases based on lipase engineering database (Gupta et al. 2015)

6.3.1 Bacterial Lipases There are eight families of bacterial lipases. Family I lipase is divided into eight subfamilies. The family II lipase enzyme is also called GDSL family and membrane esterase. In family II lipases, the conserved motifs are Gly-Asp-Ser-(Leu). Lipases from cold-loving microorganisms are included in family III. Family IV lipase is also called hormone-sensitive lipase family. Family V includes lipases from pyschrophiles, thermophiles, and mesophiles microorganisms. Family VI lipases are carboxylesterases with molecular weight 23–26 KDa, while family VII lipases have molecular weight of 50–65 KDa. The family VIII is similar to β-lactamases class C with enzyme residues Gly-Xaa-Ser-Ala-Gly conserved motif.

6.4 Reactions Catalyzed by Lipase Enzyme 6.4.1 Acidolysis It is a reaction between an acid and ester.

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6.4.2 Trans-esterification It is hydrolysis of triglycerides with the help of alcohol, where the products formed are methyl ester and glycerol.

6.4.3 Esterification Esterification is a reaction between alcohol and acid to form ester.

6.4.4 Aminolysis Aminolysis is the conversion of amines and alcohols to form amides and esters.

6.4.5 Hydrolysis Degradation of triglycerides into fatty acids and glycerol.

6.4.6 Alcoholysis It is a reaction between triolein and oleyl alcohol to form wax esters.

6.5 Catalytic Mechanism of Lipases The mechanism of the lipase is to catalyze ester hydrolysis. This involves a nucleophilic attack of the serine on the carbonyl carbon of the ester bond, to yield a covalent acyl–enzyme and release of diacylglycerol (Miranda et al. 2015). A second nucleophilic attack forms a carboxylic acid. Many nucleophilic compounds break the acyl–enzyme intermediate (Adlercreutz 2013). Lipases can perform transesterification, esterification, and acidolysis reactions. Lipases hydrolyze esters to form di- and mono-acylglycerols, fatty acids, and glycerol. Lipases also carry the reverse reactions of esterification, interesterification, and trans-esterification.

6.6 Lipase-Producing Bacteria and Fungi

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6.6 Lipase-Producing Bacteria and Fungi The bacterial lipase producers are Acinetobacter radioresistens (Li et al. 2005), Pseudomonas sp. (Kiran et al. 2008), Bacillus sp. (Ertugrul et al. 2007), Geobacillus sp., Burkholderia cepacia (Fernandes et al. 2007), and Pseudomonas aeruginosa (Mahanta et al. 2008; Ruchi et al. 2008), Acinetobacter sp., Arthrobacter sp., Achromobacter sp. The fungal lipase producers are Aspergillus niger (Dutra et al. 2008), Rhizopus sp. (Martinez-Ruiz et al. 2008), Penicillium citrinum (D’Annibale et al. 2006), Fusarium sp., Alternaria sp., etc. These lipase-producing microorganisms have been isolated from palm fruit, raw milk, meat, fermented sausages, crude-oil-contaminated areas, hot springs, soil, contaminated water sources, etc. (Gunasekaran and Das 2005).

6.6.1 Lipases Enzyme by Solid-State Fermentation The selection of a substrate for the solid-state fermentation process is important. The solid support which has important nutrients for the microorganisms is a good substrate. The most suitable substrate for lipases is the synthetic materials containing glyceride moieties. The fungi involving the production of lipases using solid-state fermentation are Trichoderma sp., Penicillium sp., Rhizopus sp., and Aspergillus sp. Lipases from different microorganisms have been produced using SSF using various solid substrates, such as lipase from Penicillium restrictum in babassu cake (Azeredo et al. 2007); lipase from P. simplicissimum in soybean cake, and castor bean cake (Gutarra et al. 2009); lipase from Candida rugosa in rice flour; lipase from Rhizopus homothallicus in sugarcane bagasse (Rodriguez et al. 2006); lipase from A. niger in wheat bran and sesame seed cake (Mahadik et al. 2002); and lipase from Rhizopus rhizopodiformis and Rhizomucor pusillus in olive oil cake and sugarcane bagasse (Cordova et al. 1998).

6.7 Assay for Lipase Enzyme (Amara et al. 2009) Lipase assays depend on the measure of fatty acids liberated. In the colorimetric method, there is use of chromogenic substrates which liberates colored product, whereas spectrophotometric method is to measure lipase enzyme spectrophotometrically.

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Prepare the substrate mixture by adding 200 μl tween 20–40 mg p-nitrophenyl palmitate and dissolve in 10 ml DMSO.

Determine the enzyme activity by adding 40 μl from the supernatant to 500 μl from 50 mM Tris HCl, pH 8.0.

Add different volumes of substrate which contain 0.4, 0.8, 2.0, and 2.8 mg/ml p-nitrophenyl palmitate to start the enzyme reaction.

Measure the activity for each substrate concentration at 37, 60, and 75 °C. The activities are determined by the rate of p-nitrophenyl palmitate produced measured at 405 nm.

Measure the increase in absorbance at varied time intervals. One unit is the amount of enzyme which liberates of 1 pmol p-nitrophenyl/min (Sullivan et al. 1999).

6.8 Applications of Lipases Enzyme 6.8.1 Lipases for the Food and Agro-industrial Applications Lipases have applications in the food industries. In vegetal oil processing, lipases enhance oil yield. Lipases improve the emulsification properties of egg yolk lipids. Lipases are used to create new functional foods.

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6.8.2 Dairy Industries In dairy industries, lipases catalyze the hydrolysis of milk fats and impart flavors to cheese (Jooyandeh et al. 2009).

6.8.3 Baking Industries In baking industries, lipases have application for producing emulsifying lipids in situ. Lipases also increase the flavor of bakery food items through esterification; the loaf volume for the shelf life of bakery food items to be good; and also help to improve their texture and softness (Robert 2015).

6.8.4 Human Milk Fat Substitutes Human milk fat (HMF) contains lipids, viz. oleic (30–35%), palmitic (20–30%), linoleic (7–14%), and stearic acids (5.7–8%). HMF substitutes are obtained by sn-1, 3 lipase-catalyzed acidolysis of tripalmitin, butterfat, palm oil, and palm stearin with free fatty acids (FFA) from various sources.

6.8.5 Egg-Processing Industries Eggs are functional ingredients for the food industries. Lipases play a role in improving the emulsifying properties of egg lipids.

6.8.6 Edible Oil Production Edible oils from plants have applications in food, feed, and fuel industries. During refining vegetal oils, the impurities phospholipids need to be removed. For long period, thermo-chemical processes were used to remove phospholipids from crude oil. Now, the lipases have attracted the attention to remove the impurities phospholipids from the crude oil.

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References Adlercreutz P (2013) Immobilisation and application of lipases in organic media. Chem Soc Rev 42:6406–6436 Amara AA, Salem SR, Shabeb MA (2009) Biodetergent: the possibility to use bacterial protease and lipase as detergent. Glob J Biotechnol Biochem 4:104–114 Andualema B, Gessesse A (2012) Microbial lipases and their industrial applications: review. Biotechnology 11:100–118 Azeredo L, Gomes P, Sant’Anna G Jr, Castilho L, Freire D (2007) Production and regulation of lipase activity from Penicillium restrictum in submerged and solid-state fermentations. Curr Microbiol 54:361–365 Cordova J, Nemmaoui M, Ismaïli-Alaoui M, Morin RA, Raimbault M, Benjilali B (1998) Lipase production by solid state fermentation of olive cake and sugar cane bagasse. J Mol Catal B 5:75–78 D’Annibale A, Sermanni G, Federici F, Petruccioli M (2006) Olive-oil wastewaters: a promising substrate for microbial lipase production. Bioresour Technol 97:1828–1833 Dutra J, Terzi S, Bevilaqua J, Damaso M, Couri S, Langone M, Senna L (2008) Lipase production in solid state fermentation monitoring biomass growth of Aspergillus niger using digital image processing. Appl Biochem Biotechnol 147:63–75 Ertugrul S, Donmez G, Takac S (2007) Isolation of lipase producing Bacillus sp. from olive mill wastewater and improving its enzyme activity. J Hazard Mater 149:720–724 Fernandes M, Saad E, Meira J, Ramos L, Mitchell D, Krieger N (2007) Esterification and transesterification reactions catalyzed by addition of fermented solids to organic reaction media. J Mol Catal B Enzym 44:8–13 Fischer M, Pleiss J (2003) The lipase engineering database: a navigation and analysis tool for protein families. Nucleic Acid Res 31:319–321 Gunasekaran V, Das D (2005) Lipase fermentation: progress and prospects. Ind J Biotechnol 4:437– 445 Gupta R, Kumari A, Syal P, Singh Y (2015) Molecular and functional diversity of yeast and fungal lipases: their role in biotechnology and cellular physiology. Prog Lipid Res 57:40–54 Gutarra M, Godoy M, Maugeri F, Rodrigues M, Freire D, Castilho L (2009) Production of an acidic and thermostable lipase of the mesophilic fungus Penicillium simplicissimum by solid-state fermentation. Bioresource Technol 100:5249–5254 Hong Mc, Chang Mc (1998) Purification and characterization of a alkaline lipase from a newly isolated Acinetobacter radioresistens CMC1. Biotech Lett 20:1027–1029 Jaeger K, Dijkstra B, Reetz M (1999) Bacterial biocatalysts: molecular biology, three-dimensional structures and biotechnological applications of lipases. Annu Rev Microbiol 53:315–351 Jooyandeh H, Amarjeet K, Minhas K (2009) Lipases in dairy industry: a review. Food Sci Technol 46:181–189 Kiran G, Shanmughapriya S, Jayalakshmi J, Selvin J, Gandhimathi R, Sivaramakrishnan S, Arunkumar M, Thangavelu T, Natarajaseenivasan K (2008) Optimization of extracellular psychrophilic alkaline lipase produced by marine Pseudomonas sp. (MSI057). Bioprocess Biosyst Eng 31:483–492 Li C, Chen S, Cheng C, Chen T (2005) Production of Acinetobacter radioresistens lipase with repeated fed-batch culture. Biochem Eng J 25:195–199 Mahadik N, Puntambekar U, Bastawde K, Khire J, Gokhale D (2002) Production of acidic lipase by Aspergillus niger in solid state fermentation. Process Biochem 38:715–721 Mahanta N, Gupta A, Khare S (2008) Production of protease and lipase by solvent tolerant Pseudomonas aeruginosa PseA in solid-state fermentation using Jatropha curcas seed cake as substrate. Bioresour Technol 99:1729–1735 Martinez-Ruiz A, Garcia H, Saucedo-Castaneda G, Favela-Torres E (2008) Organic phase synthesis of ethyl oleate using lipases produced by solid-state fermentation. Appl Biochem Biotechnol 151:393–401 Mehta A, Bodh U, Gupta R (2017) Fungal lipases: a review. J Biotech Res 8:58–77

References

113

Miranda A, Miranda L, Souza R (2015) Lipases: valuable catalysts for dynamic kinetic resolutions. Biotechnol Adv 33:372–393 Ollis D, Cheah E, Cygler M, Dijkstra B, Frolow F, Franken S, Harel M, Remington S, Silman I, Schrag J, Sussman J, Verschueren K, Goldman A (1992) The α/β hydrolase fold. Protein Eng Des Sel 5:197–211 Ray S (2015) Application of extracellular microbial lipase—a review. Int J Res Biotechnol Biochem 5:6–12 Robert H (2015) Lipases in baking. Adebiotech, Lipinov Romainville Robles-Medina A, Gonzalez-Moreno P, Esteban-Cerdan L, Molina-Grima E (2009) Biocatalysis; towards ever greener biodiesel production. Biotechnol Adv 27:398–408 Rodriguez J, Mateos J, Nungaray J, Gonzalez V, Bhagnagar T, Roussos S, Cordova J, Baratti J (2006) Improving lipase production by nutrient source modification using Rhizopus homothallicus cultured in solid state fermentation. Process Biochem 41:2264–2269 Ruchi G, Anshu G, Khare S (2008) Lipase from solvent tolerant Pseudomonas aeruginosa strain: production optimization by response surface methodology and application. Bioresour Technol 99:4796–4802 Sullivan E, Leahy J, Colwell R (1999) Cloning and sequence analysis of the lipase chaperoneencoding genes from Acinetobacter calcoaceticus RAG-1 and redefinition of a proteobacterial lipase family and an analogous lipase chaperone family. Gene 230:277–286 Thakur S (2012) Lipases, its sources, properties and applications: a Review. Int J Sci Eng Res 3:1–29