Recent Advancement in White Biotechnology Through Fungi: Volume 3: Perspective for Sustainable Environments [1st ed. 2019] 978-3-030-25505-3, 978-3-030-25506-0

Table of contents :
Front Matter ....Pages i-xxvii
Secretomics of Wood-Degrading Fungi and Anaerobic Rumen Fungi Associated with Biodegradation of Recalcitrant Plant Biomass (Nasib Singh, Joginder Singh)....Pages 1-16
Bioremediation: New Prospects for Environmental Cleaning by Fungal Enzymes (Neha Vishnoi, Sonal Dixit)....Pages 17-52
Genetic Diversity of Methylotrophic Yeast and Their Impact on Environments (Manish Kumar, Raghvendra Saxena, Pankaj Kumar Rai, Rajesh Singh Tomar, Neelam Yadav, Kusam Lata Rana et al.)....Pages 53-71
White Rot Fungi and Their Enzymes for the Treatment of Industrial Dye Effluents (Dhevagi Periasamy, Sudhakarn Mani, Ramya Ambikapathi)....Pages 73-100
Pleurotus ostreatus: A Biofactory for Lignin-Degrading Enzymes of Diverse Industrial Applications (Hesham El Enshasy, Farid Agouillal, Zarani Mat, Roslinda Abd Malek, Siti Zulaiha Hanapi, Ong Mei Leng et al.)....Pages 101-152
Extracellular Fungal Peroxidases and Laccases for Waste Treatment: Recent Improvement (Shanmugapriya S., G. Manivannan, Selvakumar Gopal, Sivakumar Natesan)....Pages 153-187
Fungal Enzymes for Bioremediation of Contaminated Soil (Prem Chandra, Enespa)....Pages 189-215
Bioremediation of Polycyclic Aromatic Hydrocarbons (PAHs) Contaminated Soil Through Fungal Communities (Ulises Conejo-Saucedo, Darío R. Olicón-Hernández, Tatiana Robledo-Mahón, Haley P. Stein, Concepción Calvo, Elisabet Aranda)....Pages 217-236
Role of Fungal Enzymes for Bioremediation of Hazardous Chemicals (Nitika Singh, Abhishek Kumar, Bechan Sharma)....Pages 237-256
Biotechnological Applications of β-Glucosidases in Biomass Degradation (Sushma Mishra, Deepika Goyal, Amit Kumar, Prem Kumar Dantu)....Pages 257-281
Role of Fungi in Climate Change Abatement Through Carbon Sequestration (Sandeep K. Malyan, Amit Kumar, Shahar Baram, Jagdeesh Kumar, Swati Singh, Smita S. Kumar et al.)....Pages 283-295
Microbial Enzymes and Their Application in Pulp and Paper Industry (Abdulhadi Yakubu, Upasana Saikia, Ashish Vyas)....Pages 297-317
Arbuscular Mycorrhizal Fungi-Mediated Mycoremediation of Saline Soil: Current Knowledge and Future Prospects (Dileep Kumar, Priyanka Priyanka, Pramendra Yadav, Anurag Yadav, Kusum Yadav)....Pages 319-348
Fungal Enzymes for Bioconversion of Lignocellulosic Biomass (Subhadeep Mondal, Suman Kumar Halder, Keshab Chandra Mondal)....Pages 349-380
Bioconversion of Biomass to Biofuel Using Fungal Consortium (Pavana Jyothi Cherukuri, Rajani Chowdary Akkina)....Pages 381-396
Role of Fungi in the Removal of Heavy Metals and Dyes from Wastewater by Biosorption Processes (Ajay Kumar, Vineet Kumar, Joginder Singh)....Pages 397-418
Impact of Arbuscular Mycorrhizal Fungi (AMF) in Global Sustainable Environments (Sanjeev Kumar, Joginder Singh)....Pages 419-436
Fungal Phytoremediation of Heavy Metal-Contaminated Resources: Current Scenario and Future Prospects (Amit Kumar, Ashish K. Chaturvedi, Kritika Yadav, K. P. Arunkumar, Sandeep K. Malyan, P. Raja et al.)....Pages 437-461
Fungal Enzymes for Bioremediation of Xenobiotic Compounds (Peter Baker, Araven Tiroumalechetty, Rajinikanth Mohan)....Pages 463-489
Fungal White Biotechnology: Conclusion and Future Prospects (Ajar Nath Yadav)....Pages 491-498
Back Matter ....Pages 499-511

Citation preview

Fungal Biology

Ajar Nath Yadav Sangram Singh Shashank Mishra Arti Gupta Editors

Recent Advancement in White Biotechnology Through Fungi Volume 3: Perspective for Sustainable Environments

Fungal Biology

Series Editors Vijai Kumar Gupta Department of Chemistry and Biotechnology Tallinn University of Technology Akadeemia tee, Tallinn, Estonia Maria G. Tuohy School of Natural Sciences National University of Ireland Galway Galway, Ireland

About the Series Fungal biology has an integral role to play in the development of the biotechnology and biomedical sectors. It has become a subject of increasing importance as new fungi and their associated biomolecules are identified. The interaction between fungi and their environment is central to many natural processes that occur in the biosphere. The hosts and habitats of these eukaryotic microorganisms are very diverse; fungi are present in every ecosystem on Earth. The fungal kingdom is equally diverse, consisting of seven different known phyla. Yet detailed knowledge is limited to relatively few species. The relationship between fungi and humans has been characterized by the juxtaposed viewpoints of fungi as infectious agents of much dread and their exploitation as highly versatile systems for a range of economically important biotechnological applications. Understanding the biology of different fungi in diverse ecosystems as well as their interactions with living and non-living is essential to underpin effective and innovative technological developments. This series will provide a detailed compendium of methods and information used to investigate different aspects of mycology, including fungal biology and biochemistry, genetics, phylogenetics, genomics, proteomics, molecular enzymology, and biotechnological applications in a manner that reflects the many recent developments of relevance to researchers and scientists investigating the Kingdom Fungi. Rapid screening techniques based on screening specific regions in the DNA of fungi have been used in species comparison and identification, and are now being extended across fungal phyla. The majorities of fungi are multicellular eukaryotic systems and therefore may be excellent model systems by which to answer fundamental biological questions. A greater understanding of the cell biology of these versatile eukaryotes will underpin efforts to engineer certain fungal species to provide novel cell factories for production of proteins for pharmaceutical applications. Renewed interest in all aspects of the biology and biotechnology of fungi may also enable the development of “one pot” microbial cell factories to meet consumer energy needs in the 21st century. To realize this potential and to truly understand the diversity and biology of these eukaryotes, continued development of scientific tools and techniques is essential. As a professional reference, this series will be very helpful to all people who work with fungi and should be useful both to academic institutions and research teams, as well as to teachers, and graduate and postgraduate students with its information on the continuous developments in fungal biology with the publication of each volume. More information about this series at http://www.springer.com/series/11224

Ajar Nath Yadav  •  Sangram Singh Shashank Mishra  •  Arti Gupta Editors

Recent Advancement in White Biotechnology Through Fungi Volume 3: Perspective for Sustainable Environments

Editors Ajar Nath Yadav Department of Biotechnology Akal College of Agriculture Eternal University, Baru Sahib Sirmour, Himachal Pradesh, India Shashank Mishra Biotech Park Lucknow, Uttar Pradesh, India

Sangram Singh Dr. Ram Manohar Lohia Avadh University Faizabad, Uttar Pradesh, India Arti Gupta Department of Zoology Dr. Ram Manohar Lohia Avadh University Gonda, Uttar Pradesh, India

ISSN 2198-7777     ISSN 2198-7785 (electronic) Fungal Biology ISBN 978-3-030-25505-3    ISBN 978-3-030-25506-0 (eBook) https://doi.org/10.1007/978-3-030-25506-0 © Springer Nature Switzerland AG 2019 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, express 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

White biotechnology is industrial biotechnology dealing with the potential application of microbes and their enzymes for different processes in industry, agriculture, pharmaceuticals, and environments. The main application of white biotechnology is commercial production of various useful organic substances, such as acetic acid, citric acid, acetone, and glycerin, and antibiotics like penicillin, streptomycin, and mitomycin. The potential applications of fungi and fungal enzymes include being environment-friendly solutions as an alternative to sustainable environments. One of the first goals on white biotechnology’s agenda has been the production of biodegradable plastics and textiles. White biotechnology can provide an unlimited and pure source of enzymes as an alternative to the harsh chemicals traditionally used in industry for accelerating chemical reactions. Enzymes are found in naturally occurring microorganisms, such as bacteria, fungi, and yeast, all of which may or may not be genetically modified. In the twenty-first century, humans acquired skills to harness fungi to protect human health (through antibiotics, antimicrobials, immunosuppressive agents, value-added products, etc.), which led to industrial-scale production of enzymes, alkaloids, detergents, acids, and biosurfactants. Human knowledge of white biotechnology has evolved to the point where today products derived from white biotechnology often display better performance, higher sustainability, and more commercially viable characteristics than products created from traditional chemical procedures. Since then, white biotechnology has steadily developed and now plays a key role in several industrial sectors, providing both high-value nutraceuticals and pharmaceutical products. The fungal strains and bioactive compounds also play an important role in environmental cleaning. The present book volume on “Recent Advancement in White Biotechnology Through Fungi” Volume 3: Perspective for Sustainable Environments is a very timely publication, which provides state-of-the-art information in the area of white biotechnology, broadly involving fungi and fungal-based products for sustainable environments. The book volume comprises 20 chapters. Chapter 1 by Singh and Singh describes fungal secretomes and their potential application for biodegradation of lignocelluloses and biopolymers for sustainable environments. Chapter 2 presented by Vishnoi and Dixit highlights eco-friendly techniques and fungal v

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enzymes used for bioremediation for environmental cleaning. Chapter 3 by Kumar et al. describes the genetic diversity of methylotrophic yeast and their impact on environments. Chapter 4 by Periasamy et al. highlights the opportunities and challenges of white-rot fungi and their enzymes for the treatment of industrial dye effluents. Enshasy et  al. describe the production of lignin-degrading enzymes by Pleurotus ostreatus and their different industrial applications in Chap. 5. Chapter 6 details the functional attributes of fungal peroxidase and laccase enzymes for waste treatments. Chandra and Enespa highlight the recent advancements in enzymes from different fungal communities for bioremediation in Chap. 7. In Chap, 8, Conejo-Saucedo and colleagues describe in detail the bioremediation of polycyclic aromatic hydrocarbon-contaminated soil through fungal communities. Singh et al. highlight the fungal production of novel enzymes and bioactive compounds for bioremediation of hazardous chemicals in Chap. 9. Mishra et al. explain β-glucosidases produced by fungal communities and their potential biotechnological applications in biomass degradation for the sustainable environments in Chap. 10. The roles of fungi in climate change abatement through carbon sequestration have been described by Malyan et al. in Chap. 11. Chapter 12 by Vyas and Yakubu describes microbial enzymes and their applications in pulp the and paper industry. Kumar et al. highlight the arbuscular mycorrhizal fungi-mediated mycoremediation of saline soil in Chap. 13. Mondal et al. discuss the fungal enzymes for bioconversion of lignocellulosic biomass in Chap. 14. Bioconversion of biomass to biofuel using fungal consortium is discussed in Chap. 15 by Cherukuri and Akkina. Kumar et al. describe the roles of fungi in the removal of heavy metals and dyes from wastewater by biosorption processes in Chap. 16. Kumar and Singh explain the impact of arbuscular mycorrhizal fungi in global sustainable environments in Chap. 17. In Chap. 18 Kumar et al. describe the current scenario and future prospects of fungal phytoremediation of heavy metal-contaminated resources. Baker et al. highlight the bioremediation of xenobiotic compounds using fungal enzymes in Chap. 19. Finally, in Chap. 20, the conclusion and future prospects on fungal white biotechnology are described by Ajar Nath Yadav. Overall, great efforts have been carried out by Dr. Ajar Nath Yadav, his editorial team, and scientists from different countries to compile this book as a highly unique and up-to-date source on fungal white biotechnology for the students, researchers, scientists, and academicians. I hope that the readers will find this book highly useful and interesting during their pursuit of fungal biotechnology.

Vice Chancellor Eternal University Baru Sahib, Himachal Pradesh, India

Dr. H. S. Dhaliwal

Foreword

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Dr. H. S. Dhaliwal is presently the Vice Chancellor of Eternal University, Baru Sahib, Himachal Pradesh, India. Dr. Dhaliwal holds a Ph.D. in Genetics from the University of California, Riverside, USA (1975). He has 40 years of research, teaching, and administrative experience in various capacities. Dr. Dhaliwal is a Professor of Biotechnology at Eternal University, Baru Sahib, from 2011 to present. He had worked as Professor of Biotechnology at IIT, Roorkee (2003– 2011); Founding Director of Biotechnology Centre, Punjab Agricultural University, Ludhiana (1992–2003); Senior Scientist and Wheat Breeder-cum-Director at PAU’s Regional Research Station, Gurdaspur (1979– 1990); Research Fellow at FMI, Basel, Switzerland (1976–1979); and D.F. Jones Postdoctoral Fellow at the University of California, Riverside, USA (1975–1976). Dr. Dhaliwal was elected as Fellow at the National Academy of Agricultural Sciences, India, (1992); worked as Visiting Professor at the Department of Plant Pathology, Kansas State University, Kansas, USA (1989); and was elected as Senior Research Fellow at CIMMYT, Mexico (1987). He has many national and international awards under his name such as the Pesticide India Award from the Indian Society of Mycology and Plant Pathology in 1988 and a cash award from the Federation of Indian Chambers of Commerce and Industry (FICCI) in 1985. He has to his credit more than 400 publications including 250 research papers, 12 reviews, 15 chapters contributed to books, 105 papers presented in meetings and conferences and abstracted, 18 popular articles, and 2 books/ bulletins/manuals. His important research contributions are identification of new species of wild diploid wheat Triticum urartu and gathering of evidences to implicate T. urartu as one of the parents of polyploid wheat; serving as team leader in the development of seven wheat varieties, viz., PBW 54, PBW 120, PBW 138, PBW 175, PBW 222, PBW 226, and PBW 299, approved for cultivation in Punjab and North Western Plains Zone of India; molecular marker-assisted pyramiding of bacterial blight resistance genes Xa21 and Xa13 and the green revolution semi-­dwarfing gene sd1 in Dehraduni Basmati; and development of elite wheat lines biofortified for grain rich in iron and zinc through wide hybridization with related non-­progenitor wild wheat species

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and molecular breeding. Dr. Dhaliwal made a significant contribution to the development of life and epidemiology life cycle of Tilletia indica fungus, the causal organism of Karnal bunt disease of wheat, and development of Karnal bunt-resistant wheat cultivar. Dr. Dhaliwal had the membership of several task forces and committees of the Department of Biotechnology, Ministry of Science and Technology, Government of India, New Delhi; served as Chairman of the Project Monitoring Committee for Wheat Quality Breeding, Department of Biotechnology, Ministry of Science and Technology, Government of India (2007–2010), and Chairman of the Project Monitoring Committee in “Agri-biotechnology,” Department of Biotechnology, Government of India, New Delhi (2014–2016); and presently is a Member of the newly constituted Expert Committee for DBT-UDSC Partnership Centre on Genetic Manipulation of Crop Plants at UDSC, New Delhi (2016 onward).

Foreword

Perspective for environmental sustainability is to maintain the persona of an environment for people and other species. Some of the issues that create major environmental sustainability problems include destruction of the living environments (habitats) of native species, release of polluting chemicals and other materials into the environment, emission of greenhouse gases into the atmosphere than can cause climate change, and depletion of low-cost oil and other fossil fuels. One promising alternative treatment strategy to incineration is bioremediation which is to exploit the ability of microorganisms to remove pollutants from contaminated sites like water, air, and climate. Fungi are among the most potential candidates of bioremediation as they are natural decomposers of waste matter and secrete several extracellular enzymes capable of decomposing lignin and cellulose. Fungi possess the biochemical and ecological capacity to degrade environmental organic chemicals and to decrease the risk associated with metals, metalloids, and radionuclides, either by chemical alteration or by influencing chemical bioavailability. CEO Biotech Park, Lucknow, UP, India

Prof. Pramod Tandon

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Preface

White biotechnology is drawing much attention as a solution to producing enzymes and bioactive compounds for bioremediation for environmental cleaning through eco-friendly techniques such as the use of fungi to synthesize hydrolytic enzymes and compounds for plant growth promotion, biocontrol, and other processes for agriculture, medicine, industry, pharmaceuticals, and allied sectors. White fungal biotechnology is an emerging field in science that supports the revealing of novel and vital biotechnological components. The fungi Aspergillus, Bipolaris, Cordyceps, Fusarium, Gaeumannomyces, Myceliophthora, Penicillium, Phoma, Piriformospora, Pleurotus, Trichoderma, and Xylaria are highly important fungal groups which can be utilized for production of different antibiotics, enzymes, pigments, and peptides useful in different processes of environmental cleaning. The present book on “Recent Advancement in White Biotechnology Through Fungi” Volume 3: Perspective for Sustainable Environments covers agriculturally and industrially important fungi producing enzymes and bioactive compounds having the potential application for cleaning polluted environments. The fungal community from different habitats such as those from extreme habitats and plant-associated fungi having capability to produce extracellular enzymes, secondary metabolites, and bioactive compounds for diverse processes are targeted to be applied in therapeutics, diagnostics, bioremediation, agriculture, industries, and environments. This book volume will be immensely useful to those working in biological sciences, especially to microbiologists, microbial biotechnologists, biochemists, researchers, and scientists of fungal biotechnology. We are honored that the leading scientists who have extensive, in-depth experience and expertise in fungal system and microbial biotechnology took the time and effort to develop these outstanding chapters. Each chapter is written by internationally recognized researchers/scientists so the reader is given an up-to-date and detailed account of our knowledge of white biotechnology and innumerable environmental applications of fungi. We are grateful to the many people who helped to bring this book to light. Editors wish to thank Dr. Eric Stannard, Senior Editor, Botany, Springer; Dr. Vijai Kumar Gupta and Prof. Maria G.  Tuohy, Series Editors, Fungal Biology, Springer; and Ms. Saveetha Balasundaram and Mr. Rahul Kumar, Project Coordinators, Springer, xi

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for their generous assistance, constant support, and patience in initializing the volume. Dr. Ajar Nath Yadav gives special thanks to his exquisite wife Ms. Neelam Yadav for her constant support and motivation in putting everything together. Dr. Yadav also gives special thanks to his esteemed friends, well-wishers, colleagues, and senior faculty members of Eternal University, Baru Sahib, India. Baru Sahib, Himachal Pradesh, India  Ajar Nath Yadav  Lucknow, Uttar Pradesh, India  Shashank Mishra  Faizabad, Uttar Pradesh, India  Sangram Singh  Gonda, Uttar Pradesh, India  Arti Gupta

Contents

1 Secretomics of Wood-Degrading Fungi and Anaerobic Rumen Fungi Associated with Biodegradation of Recalcitrant Plant Biomass ������������������������������������������������������������������������������������������    1 Nasib Singh and Joginder Singh 2 Bioremediation: New Prospects for Environmental Cleaning by Fungal Enzymes������������������������������������������������������������������   17 Neha Vishnoi and Sonal Dixit 3 Genetic Diversity of Methylotrophic Yeast and Their Impact on Environments��������������������������������������������������������������������������������������   53 Manish Kumar, Raghvendra Saxena, Pankaj Kumar Rai, Rajesh Singh Tomar, Neelam Yadav, Kusam Lata Rana, Divjot Kour, and Ajar Nath Yadav 4 White Rot Fungi and Their Enzymes for the Treatment of Industrial Dye Effluents����������������������������������������������������������������������   73 Dhevagi Periasamy, Sudhakarn Mani, and Ramya Ambikapathi 5 Pleurotus ostreatus: A Biofactory for Lignin-Degrading Enzymes of Diverse Industrial Applications������������������������������������������  101 Hesham El Enshasy, Farid Agouillal, Zarani Mat, Roslinda Abd Malek, Siti Zulaiha Hanapi, Ong Mei Leng, Daniel Joe Dailin, and Dalia Sukmawati 6 Extracellular Fungal Peroxidases and Laccases for Waste Treatment: Recent Improvement ����������������������������������������������������������  153 Shanmugapriya S., Manivannan G., Selvakumar Gopal, and Sivakumar Natesan 7 Fungal Enzymes for Bioremediation of Contaminated Soil����������������  189 Prem Chandra and Enespa

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8 Bioremediation of Polycyclic Aromatic Hydrocarbons (PAHs) Contaminated Soil Through Fungal Communities������������������������������  217 Ulises Conejo-Saucedo, Darío R. Olicón-Hernández, Tatiana Robledo-Mahón, Haley P. Stein, Concepción Calvo, and Elisabet Aranda 9 Role of Fungal Enzymes for Bioremediation of Hazardous Chemicals��������������������������������������������������������������������������������������������������  237 Nitika Singh, Abhishek Kumar, and Bechan Sharma 10 Biotechnological Applications of β-Glucosidases in Biomass Degradation����������������������������������������������������������������������������������������������  257 Sushma Mishra, Deepika Goyal, Amit Kumar, and Prem Kumar Dantu 11 Role of Fungi in Climate Change Abatement Through Carbon Sequestration������������������������������������������������������������������������������  283 Sandeep K. Malyan, Amit Kumar, Shahar Baram, Jagdeesh Kumar, Swati Singh, Smita S. Kumar, and Ajar Nath Yadav 12 Microbial Enzymes and Their Application in Pulp and Paper Industry����������������������������������������������������������������������������������  297 Abdulhadi Yakubu, Upasana Saikia, and Ashish Vyas 13 Arbuscular Mycorrhizal Fungi-Mediated Mycoremediation of Saline Soil: Current Knowledge and Future Prospects ������������������  319 Dileep Kumar, Priyanka Priyanka, Pramendra Yadav, Anurag Yadav, and Kusum Yadav 14 Fungal Enzymes for Bioconversion of Lignocellulosic Biomass����������  349 Subhadeep Mondal, Suman Kumar Halder, and Keshab Chandra Mondal 15 Bioconversion of Biomass to Biofuel Using Fungal Consortium ��������  381 Pavana Jyothi Cherukuri and Rajani Chowdary Akkina 16 Role of Fungi in the Removal of Heavy Metals and Dyes from Wastewater by Biosorption Processes������������������������������������������  397 Ajay Kumar, Vineet Kumar, and Joginder Singh 17 Impact of Arbuscular Mycorrhizal Fungi (AMF) in Global Sustainable Environments������������������������������������������������������  419 Sanjeev Kumar and Joginder Singh 18 Fungal Phytoremediation of Heavy Metal-­Contaminated Resources: Current Scenario and Future Prospects����������������������������  437 Amit Kumar, Ashish K. Chaturvedi, Kritika Yadav, K. P. Arunkumar, Sandeep K. Malyan, P. Raja, Ram Kumar, Shakeel Ahmad Khan, Krishna Kumar Yadav, Kusam Lata Rana, Divjot Kour, Neelam Yadav, and Ajar Nath Yadav

Contents

xv

19 Fungal Enzymes for Bioremediation of Xenobiotic Compounds��������  463 Peter Baker, Araven Tiroumalechetty, and Rajinikanth Mohan 20 Fungal White Biotechnology: Conclusion and Future Prospects��������  491 Ajar Nath Yadav Index������������������������������������������������������������������������������������������������������������������  499

Contributors

Farid  Agouillal  Research Unit on Analysis and Technological Development in Environment (URADTE), Centre de Recherche Scientifique et Technique en Analyses Physico-Chimiques (CRAPC), Tipaza, Algeria Rajani  Chowdary  Akkina  Department of Microbiology & Food Science and Technology, Institute of Science, GITAM (Deed to be University), Visakhapatnam, Andhra Pradesh, India Ramya  Ambikapathi  Department of Environmental Sciences, Tamil Nadu Agricultural University, Coimbatore, India Elisabet  Aranda  Department of Microbiology, Institute of Water Research, University of Granada, Granada, Spain K. P. Arunkumar  Central Muga Eri Research and Training Institute, Central Silk Board, Jorhat, Assam, India Peter Baker  Department of Biology, Colgate University, Hamilton, NY, USA Shahar Baram  Institute of Soil, Water and Environmental Sciences, The Volcani Research Center, Agricultural Research Organization (ARO), Rishon LeZion, Israel Concepción  Calvo  Department of Microbiology, Institute of Water Research, University of Granada, Granada, Spain Prem  Chandra  Department of Environmental Microbiology, School for Environmental Sciences, Babasaheb Bhimrao Ambedkar (A Central) University, Lucknow, Uttar Pradesh, India Ashish  K.  Chaturvedi  Water Management (Agriculture) Division, Centre for Water Resources Development and Management, Kozhikode, Kerala, India Pavana  Jyothi  Cherukuri  Department of Microbiology & Food Science and Technology, Institute of Science, GITAM (Deed to be University), Visakhapatnam, Andhra Pradesh, India xvii

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Contributors

Ulises Conejo-Saucedo  Department of Microbiology, Institute of Water Research, University of Granada, Granada, Spain Daniel  Joe  Dailin  Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia School of Chemical and Energy Engineering, Faculty of Engineering, Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia Prem  Kumar  Dantu  Department of Botany, Dayalbagh Educational Institute, Deemed University, Dayalbagh, Agra, India Sonal Dixit  Department of Botany, University of Lucknow, Lucknow, India Enespa  Department of Plant Pathology, School of Agriculture, MPDC, University of Lucknow, Lucknow, Uttar Pradesh, India Hesham  El Enshasy  Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia School of Chemical and Energy Engineering, Faculty of Engineering, Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia City of Scientific Research and Technology Applications, New Burg Al Arab, Alexandria, Egypt Selvakumar Gopal  Department of Microbiology, Alagappa University, Karaikudi, Tamil Nadu, India Deepika Goyal  Department of Botany, Dayalbagh Educational Institute, Deemed University, Dayalbagh, Agra, India Suman  Kumar  Halder  Department of Microbiology, Vidyasagar University, Midnapore, West Bengal, India Siti  Zulaiha  Hanapi  Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia Shakeel  Ahmad  Khan  Centre for Environment Science and Climate Resilient Agriculture, ICAR-Indian Agricultural Research Institute, New Delhi, India Divjot Kour  Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, Sirmour, Himachal Pradesh, India Abhishek Kumar  Department of Biochemistry, Faculty of Science, University of Allahabad, Allahabad, India Ajay  Kumar  School of Bioengineering and Biosciences, Lovely Professional University, Phagwara, Punjab, India Amit Kumar  Host Plant Section, Central Muga Eri Research & Training Institute, Central Silk Board, Lahdoigarh, Jorhat, Assam, India Dileep  Kumar  Department of Biochemistry, University of Lucknow, Lucknow, Uttar Pradesh, India

Contributors

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Jagdeesh  Kumar  Department of Hydrology, Indian Institute of Technology Roorkee, Roorkee, Uttarakhand, India Manish  Kumar  Amity Institute of Biotechnology, Amity University, Gwalior, India Ram Kumar  Centre for Environment Science and Climate Resilient Agriculture, ICAR-Indian Agricultural Research Institute, New Delhi, India Sanjeev Kumar  Department of Genetics and Plant Breeding, Lovely Professional University, Jalandhar, India Smita S. Kumar  Center for Rural Development and Technology, Indian Institute of Technology Delhi, New Delhi, India Vineet  Kumar  School of Bioengineering and Biosciences, Lovely Professional University, Phagwara, Punjab, India Ong Mei Leng  Harita Go Green Sdn. Bhd., Johor Bahru, Johor, Malaysia Roslinda  Abd  Malek  Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia Sandeep  K.  Malyan  Institute of Soil, Water and Environmental Sciences, The Volcani Research Center, Agricultural Research Organization (ARO), Rishon LeZion, Israel Sudhakarn Mani  Department of Environmental Sciences, Tamil Nadu Agricultural University, Coimbatore, India Manivannan G.  Department of Microbiology and Biotechnology, SVN College, Madurai, Tamil Nadu, India Zarani  Mat  Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia Sushma Mishra  Department of Botany, Dayalbagh Educational Institute, Deemed University, Dayalbagh, Agra, India Rajinikanth Mohan  Department of Biology, Colgate University, Hamilton, NY, USA Department of Biology, Mercyhurst University, Erie, PA, USA Keshab  Chandra  Mondal  Department of Microbiology, Vidyasagar University, Midnapore, West Bengal, India Subhadeep  Mondal  Department of Microbiology, Vidyasagar University, Midnapore, West Bengal, India Darío  R.  Olicón-Hernández  Department of Microbiology, Institute of Water Research, University of Granada, Granada, Spain Dhevagi  Periasamy  Department of Environmental Sciences, Tamil Nadu Agricultural University, Coimbatore, India

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Contributors

Priyanka  Priyanka  Department of Biochemistry, University of Lucknow, Lucknow, Uttar Pradesh, India Pankaj  Kumar  Rai  Department of Biotechnology, Invertis University, Bareilly, Uttar Pradesh, India P. Raja  ICAR-IISWC, Regional Centre, Ooty, Tamil Nadu, India Kusam  Lata  Rana  Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, Sirmour, Himachal Pradesh, India Tatiana  Robledo-Mahón  Department of Microbiology, Institute of Water Research, University of Granada, Granada, Spain Upasana  Saikia  Department of Microbiology, School of Bioscience and Bioengineering, Lovely Professional University, Phagwara, Punjab, India Raghvendra Saxena  Amity Institute of Biotechnology, Amity University, Gwalior, India Shanmugapriya  S.  Department of Molecular Microbiology, School Biotechnology, Madurai Kamaraj University, Madurai, Tamil Nadu, India

of

Bechan Sharma  Department of Biochemistry, Faculty of Science, University of Allahabad, Allahabad, India Joginder  Singh  Department of Biotechnology, School of Bioengineering and Biosciences, Lovely Professional University, Jalandhar, Phagwara, Punjab, India Nasib Singh  Department of Microbiology, Akal College of Basic Sciences, Eternal University, Baru Sahib, Himachal Pradesh, India Nitika  Singh  Department of Biochemistry, Faculty of Science, University of Allahabad, Allahabad, India Swati  Singh  Department of Environmental Science, Chaudhary Charan Singh University, Meerut, Uttar Pradesh, India Sivakumar  Natesan  Department of Molecular Microbiology, School of Biotechnology, Madurai Kamaraj University, Madurai, Tamil Nadu, India Haley P. Stein  Department of Microbiology, Institute of Water Research, University of Granada, Granada, Spain Dalia  Sukmawati  Faculty of Mathematics and Natural Sciences, Universitas Negeri Jakarta, Jakarta, Indonesia Araven Tiroumalechetty  Department of Biology, Colgate University, Hamilton, NY, USA Rajesh  Singh  Tomar  Amity Institute of Biotechnology, Amity University, Gwalior, India

Contributors

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Neha  Vishnoi  Department of Environmental Sciences, Babasaheb Bhimrao Ambedkar University, Lucknow, India Ashish  Vyas  Department of Microbiology, School of Bioscience and Bioengineering, Lovely Professional University, Phagwara, Punjab, India Ajar  Nath  Yadav  Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, Sirmour, Himachal Pradesh, India Anurag  Yadav  Department of Microbiology, College of Basic Science and Humanities, Sardarkrushinagar Dantiwada Agricultural University, Banaskantha, Gujarat, India Krishna  Kumar  Yadav  Institute of Environment and Development Studies, Bundelkhand University, Jhansi, Uttar Pradesh, India Kritika  Yadav  Department of Botany, Dayalbagh Educational Institute, Agra, Uttar Pradesh, India Kusum  Yadav  Department of Biochemistry, University of Lucknow, Lucknow, Uttar Pradesh, India Neelam Yadav  Gopi Nath P.G. College, Veer Bahadur Singh Purvanchal University, Deoli-Salamatpur, Ghazipur, Uttar Pradesh, India Pramendra  Yadav  Department of Biochemistry, University of Lucknow, Lucknow, Uttar Pradesh, India Abdulhadi  Yakubu  Department of Science Laboratory Technology, College of Science and Technology, Jigawa State Polytechnic, Dutse, Nigeria

About the Editors

Ajar  Nath  Yadav  is an Assistant Professor in the Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, Himachal Pradesh, India. He has 4 years of teaching and 10 years of research experiences in the field of Industrial Biotechnology, Microbial Biotechnology, Microbial Diversity, and Plant-Microbe Interactions. Dr. Yadav obtained his doctorate degree in Microbial Biotechnology, jointly from the Indian Agricultural Research Institute, New Delhi, and Birla Institute of Technology, Mesra, Ranchi, India; M.Sc. (Biotechnology) from Bundelkhand University, India; and B.Sc. (CBZ) from the University of Allahabad, India. Dr. Yadav has 111 publications, which include 39 research papers, 15 review papers, 11 books, 1 Laboratory manual, 33 book chapters, 8 popular articles, 7 editorials, 2 technical reports, and 1 patent with h-index of 25, i10-index of 57, and 1760 citations (Google Scholar). Dr. Yadav has published 109 research communications in different international and national conferences. Dr. Yadav has got 12 Best Paper Presentation Awards, one Young Scientist Award (NASI-Swarna Jayanti Puraskar), and three certificates of excellence in reviewing awards. Dr. Yadav received the “Outstanding Teacher Award” in the 6th Annual Convocation in 2018 by Eternal University, Baru Sahib, Himachal Pradesh. Dr. Yadav has a long-standing interest in teaching at the UG, PG, and Ph.D. level and is involved in taking courses in Agriculture Microbiology, Bacteriology, Bioprocess Engineering and Technology, Environmental Microbiology, Industrial Microbiology, and Microbial xxiii

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About the Editors

Biotechnology. Dr. Yadav is currently handling two projects, one funded by the Department of Environment, Science and Technology (DEST), Shimla, entitled “Development of Microbial Consortium as Bioinoculants for Drought and Low Temperature Growing Crops for Organic Farming in Himachal Pradesh” as Principal Investigator and another funded by HP Council for Science, Technology and Environment (HIMCOSTE) on “value-added products” as Co-PI. He also worked as an Organizing Committee Member for seven international conferences/symposia in the related field. Presently he is guiding four scholars for Ph.D. degree and one scholar for M.Sc. dissertation. To his credit ~6700 microbes (archaea, bacteria, and fungi) were isolated from diverse sources and ~550 potential and efficient microbes were deposited at culture collection of the National Bureau of Agriculturally Important Microorganisms (NBAIM), Mau, India. He has deposited 2386 nucleotide sequences and 3 whole genome sequences (Bacillus thuringiensis AKS47, Arthrobacter agilis L77, and Halolamina pelagica CDK2) and 2 transcriptomes to NCBI GenBank databases: in public domain. Dr. Yadav and group have developed a method for screening of archaea for phosphorus solubilization for the first time. He has been serving as an Editor/ Editorial Board Member and Reviewer for more than 35 national and international peer-reviewed journals. He has lifetime memberships of the Association of Microbiologists of India; Indian Science Congress Association, India; and National Academy of Sciences, India. Please visit https://sites.google.com/site/ajarbiotech/ for more details. Sangram  Singh  is an Associate Professor in the Department of Biochemistry, Dr. Ram Manohar Lohia Avadh University, Faizabad, India, and has 11 years of teaching and 14 years of research experiences in the field of Applied Biochemistry. Dr. Singh obtained his Ph.D. in Biochemistry and M.Sc. in Biochemistry from Dr. Ram Manohar Lohia Avadh University, Faizabad, India. Dr. Singh has published 34 national and international research papers, 2 books, and 2 book chapters. He has presented nine papers in different national and international symposia/seminars/ conferences/workshops.

About the Editors

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Shashank  Mishra  is presently working as Scientist “C,” Biotech Park, Lucknow, Uttar Pradesh, India. He obtained his doctorate degree in science (“Industrial Biotechnology”) in 2015 from Birla Institute of Technology, Mesra, Ranchi, India; M.Phil. (Biotechnology) in 2008 from Alagappa University, Tamil Nadu, India; M.Sc. (Botany) in 2005 from Dr. R.M.L.  Avadh University, Ayodhya, India; M.Sc. (Biotechnology) in 2004 from Barkatullah University, Bhopal, India; and B.Sc. (Botany and Chemistry) in 2001 from Dr. R.M.L.  Avadh University, Ayodhya, India. He has made pioneering contributions in the area of Microbial Biotechnology, Natural Product Synthesis, and Environmental Microbiology for Food, Pharmaceutical, and Human Health. To his credit he has 23 publications (7 research papers, 2 review articles, 3 books, and 11 book chapters) in different reputed international and national journals and publishers with 138 citations, h-index of 5, and i10-index of 4 (Google Scholar). He has reported for the first time high concentration of phenolic compounds by optimizing various parameters and published in peer-reviewed and refereed international journals. He has published 16 abstracts in different conferences/symposia/workshops. He has presented 16 papers (12 poster + 04 oral) in conferences/symposia and got 1 Best Poster Presentation Award. Dr. Mishra has contributed in organizing seven conferences/workshops. He has deposited three nucleotide sequences to NCBI GenBank databases: in public domain. Dr. Mishra and group have isolated and characterized three microbes (bacteria and microalgae) from tulsi and paddy plantation site and transformed ferulic acid into value-added phenolic compounds, viz., vanillin, vanillic acid, and 4-vinylguaiacol. He has a long-standing interest in teaching at the UG, PG, and Ph.D. level and is involved in taking courses in Industrial Biotechnology, Bioprocess Engineering and Technology, Environmental Biotechnology, Environmental Microbiology, Industrial Microbiology, Microbial Biotechnology, and Techniques in Microbiology and Biotechnology. He is a reviewer in six international journals including BMC Microbiology, Indian Phytopathology, PLOS One,

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About the Editors

Scientific Reports and Archive of Phytopathology and Plant Protection, and 3 Biotech. He has lifetime memberships of the Association of Microbiologists of India (AMI) and Vigyan Bharati (VIBHA).

Arti  Gupta  is an Assistant Professor in the Department of Zoology, Sri Avadh Raj Singh Smarak Degree College, Bishunpur Bairiya, Gonda, India. Dr. Arti Gupta received her B.Sc. in Botany, Zoology, and Chemistry in 2001 and got her M.Sc. in Biotechnology in 2003 from Chaudhary Charan Singh University, Meerut, India. Dr. Arti Gupta obtained her Ph.D. from Mahatma Jyotiba Phule Rohilkhand University, Bareilly, India, in 2010 in Animal Science. Dr. Gupta started her career in 2004 with teaching for graduate and post-graduate students of Biotechnology from D.A.V. (P.G.) College, Muzaffarnagar. In 2005, she was appointed as a Research Intern at the Central Drug Research Institute, Uttar Pradesh. In 2010, she was appointed as Teaching Associate at the Govind Ballabh Pant Engineering College, Pauri Garhwal, Uttarakhand. In 2012, she worked as Scientist with Sun Agrigenetics Pvt. Ltd., Vadodara, Gujarat, and has 9 years of teaching and 11 years of research experiences in the field of Animal Biotechnology, Molecular Plant Biotechnology, Molecular Animal Biotechnology, Bioprocess Technology, and Microbiology. Dr. Gupta has published one monograph, has edited three Springer Nature Switzerland books and is currently editing other Springer Nature books, has published 22 national and international research papers, and has attended 36 national and international symposia/seminars/ conferences/workshops. Dr. Gupta has been awarded University Topper (Gold Medal), M.Sc. (Biotech.), by Ch. C.  S. University, Meerut; Young Scientist Award (Gold Medal) by the Zoological Society of India, Lucknow; Best Poster Presenter by the Asian Journal of Experimental Science, Jaipur; Best Poster Presenter by the International Consortium of Contemporary Biologists (ICCB) and Madhawi-Shyam Educational Trust (MSET), Ranchi; Fellowship Award by the International Consortium of Contemporary Biologists (ICCB) and Madhawi-Shyam Educational Trust

About the Editors

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(MSET); and Dr. V.P. Agarwal Gold Medal by D.A.V. (P.G.) College, Muzaffarnagar. Dr. Gupta has lifetime memberships of the Indian Science Congress Association, Biotech Research Society of India, Zoological Society of India, and International Consortium of Contemporary Biologists.

Chapter 1

Secretomics of Wood-Degrading Fungi and Anaerobic Rumen Fungi Associated with Biodegradation of Recalcitrant Plant Biomass Nasib Singh and Joginder Singh

1.1  Introduction Lignocellulose is a widely available recalcitrant plant biopolymer composed of polymeric polysaccharides cellulose, hemicellulose, and heteropolymeric lignin (Lewis and Yamamoto 1990; Eastwood et al. 2011; Bugg et al. 2011; Janusz et al. 2017; dos Santos et  al. 2018; Bissaro et  al. 2018; Brink et  al. 2019; Ralph et  al. 2019). According to an estimate, 550 billion tons of carbon are present in vegetation in terrestrial ecosystems including forest ecosystems where dead wood is the major form of the plant biomass (Siegenthaler and Sarmiento 1993; Krah et al. 2018). This recalcitrant plant biomass is recognized as the most abundant carbon source in terrestrial ecosystem. In woody plants (angiosperms and gymnosperms), cellulose generally constitutes 40–50% of the dry weight, whereas the amount of hemicelluloses and lignin ranges from 15% to 30% (Krah et al. 2018; Adesogan et al. 2019). Cellulose is a macropolymer of numerous glucose units attached linearly by β-1,4-­ glycosidic linkages. It is responsible for rigidity and crystalline form of plant cell walls (Baldrian and Valaskova 2008; McFarlane et  al. 2014; Bissaro et  al. 2018). Hemicelluloses, on the other hand, are complex and heterogeneous plant polysaccharides consisting of xylose, mannose, arabinose, glucose, galactose, and sugar acids. Xyloglucans, xylans, mannans, and glucomannans are the main examples of hemicelluloses (Scheller and Ulvskov 2010). These are known to strengthen the plant cell wall by filling the voids around cellulose fibrils and interacting with lignin.

N. Singh (*) Department of Microbiology, Akal College of Basic Sciences, Eternal University, Baru Sahib, Himachal Pradesh, India J. Singh Department of Biotechnology, School of Bioengineering and Biosciences, Lovely Professional University, Jalandhar, Phagwara, Punjab, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_1

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Hemicelluloses are considered as the second most abundant polysaccharide in the nature (Saha 2003). Lignin is complex, aromatic, heteropolymeric, and most indigestible parts of the plant cell wall exhibiting almost complete resistance to hydrolytic degradation (Lewis and Yamamoto 1990; Janusz et al. 2017; Brink et al. 2019; Ralph et al. 2019). It contributes 15–30% of dry weight of vascular plant cell walls (Lewis and Yamamoto 1990; Gall et al. 2017). It is a phenolic polymer composed mainly of p-coumaryl, coniferyl, and sinapyl alcohols (Janusz et al. 2017; dos Santos et al. 2018; Brink et al. 2019; Ralph et al. 2019). Lignin confers rigidity to the plant cell walls and inhibits hydrolytic attacks on adjacent cellulose and hemicellulose (Lewis and Yamamoto 1990; Gall et  al. 2017; Ralph et  al. 2019). Lignocellulose-rich forest waste, dead woods, agro-food industry wastes, and leftover crop residues offer a sustainable, eco-friendly, and abundant resource for industrial-scale production of green energy and biofuels.

1.2  Degradation and Depolymerization of Plant Biomass The aromatic nature of lignin is a major obstacle for biodegradation and mineralization of lignocellulose (Lewis and Yamamoto 1990; Janusz et al. 2017; Bissaro et al. 2018; Brink et al. 2019). Lignin and its phenolic derivatives are known to inhibit lignocellulolytic enzymes by adsorption or deactivation. In the biological world, fungi are considered to be the most prolific producers of lignocellulolytic enzymes (Blanchette 1991; Conesa et  al. 2001; Bouws et  al. 2008; Eastwood et  al. 2011; Girard et al. 2013; Edwards et al. 2017; Kameshwar et al. 2019; Yadav et al. 2019a, b). These eukaryotic organisms play indispensable role in plant matter decomposition, carbon cycle, and overall nutrients recycling. Most fungi are aerobic and facultative anaerobic with the exception of strict anaerobic species present in the rumen of herbivorous animals (Sirohi et al. 2012; Edwards et al. 2017; Hooker et al. 2019). Filamentous fungi, in particular, secrete abundant enzymes to break down plant matter and complex materials in the environment which is in turn absorbed through hypha walls and utilized further for their growth and maintenance (Bouws et  al. 2008; Eastwood et al. 2011; Girard et al. 2013; Edwards et al. 2017). Their involvement in plant matter decomposition, association with plant roots as mycorrhiza, association with photobionts in lichens, and presence in rumen and intestine of wood-decaying insects are well established, widely reported, and extensively reviewed earlier by several investigators. Fungi and bacteria are indispensably involved in degradation of recalcitrant plant biomass, thus contributing massively to carbon cycle in various ecosystems (Cragg et al. 2015; Janusz et al. 2017). Surprisingly, animals such as ruminants, termites, millipedes, and terrestrial isopods are dependent on their respective microbiomes for lignocellulose degradation as none of them encode the entire enzymatic repertoire required for lignocellulose degradation (Sirohi et  al. 2012; Edwards et  al. 2017; Kameshwar et  al. 2019). In this chapter, we will emphasize the roles and secretomes of wood-degrading fungi and anaerobic rumen fungi associated with

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plant biomass degradation. Fungi are among the most diversified and widely studied Natural Biomass Utilization Systems (NBUS; Eastwood et al. 2011; Girard et al. 2013). Several fungi are equipped with enzymes which empower them to harvest energy and nutrition from plant biomass, which otherwise indigestible for other living organisms (Girard et al. 2013; Edwards et al. 2017; Janusz et al. 2017). Several fungal species from class Agaricomycetes (Basidiomycota) are closely associated with wood decay in different ecological niches (Krah et al. 2018). The saprotrophic members of this class cause decay of dead woods as either white rot or brown rot. The white rot fungi (WRF) and brown rot fungi (BRF) are the prominent colonizers which degrade lignocellulose cell wall components of compact wood logs, branches, and stumps (Martinez et al. 2004, 2009; Girard et al. 2013; Edwards et al. 2017; Janusz et al. 2017; SistaKameshwar and Qin 2018; Reina et al. 2019) (Fig. 1.1). In addition to white and brown rot, other wood-decaying manifestations, viz., soft rot and gray rot, are also exhibited by some members of Basidiomycota and Ascomycota (Riley et al. 2014). Anaerobic rumen fungi (ARF) account for 8–20% of total microbial biomass of rumen and alimentary tract of herbivorous mammals (Sirohi et al. 2012; Edwards et al. 2017; Kameshwar et al. 2019; Hooker et al. 2019). First described in 1975 by Colin Orpin (Youssef et al. 2013), these obligate anaerobic, filamentous, and motile zoospore-forming fungi are assigned to phylum Neocallimastigomycota. Despite their low numbers (106 per ml of rumen fluid), ARF are key players in lignocellulose degradation in the rumen as these physically penetrate and disrupt the plant cell walls and thus facilitate rapid growth of fibrolytic bacteria leading to optimal degradation and utilization of lignocellulosic biomass (Sirohi et al. 2012; Youssef et al. 2013;

Fig. 1.1  Association of fungi with degradation and depolymerization of woods, fodder, and crop residues

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Solomon et al. 2016; Haitjema et al. 2017; Edwards et al. 2017; Hooker et al. 2019). At present, only 11 genera have been cultured from rumen ecosystem of various herbivorous animals. These genera are Neocallimastix, Anaeromyces, Caecomyces, Cyllamyces, Orpinomyces, Piromyces, Buwchfawromyces, Feramyces, Oontomyces, Pecoramyces, and Liebetanzomyces (Sirohi et al. 2012; Edwards et al. 2017; Hooker et al. 2019; Li et al. 2019). In the next section, we have discussed the metabolic capabilities of wood-degrading fungi and ARF involved in plant biomass degradation and mineralization.

1.3  S  ecretomics and Mechanism of Lignocellulose Biodegradation Secretome, a term coined by Tjalsma et al. (2000), represents all the proteins and cellular machineries which are secreted outside the plasma membrane into the environment or extracellular matrix by a cell (McCotter et al. 2016). Plants, bacteria, and fungi exhibit their unique and substrate-specific secretome under different environmental conditions. Fungal secretome, therefore, essentially comprises extracellular enzymes which are released exterior to the cell wall, usually in the presence of lignocellulosic plant matter (Bouws et al. 2008; Eastwood et al. 2011; Girard et al. 2013; Kameshwar et al. 2019). In the past decade, advances in protein identification techniques and genome sequencing have enabled detailed investigation of the secretomes of many saprophytic, pathogenic, and symbiotic fungal species revealing rich, diverse, and highly specific enzymatic profiles. The deconstruction and mineralization of recalcitrant and indigestible plant biomass require the synergistic and cooperative action of several hydrolytic, oxidative, and nonhydrolytic enzymes (Blanchette 1991; Girard et  al. 2013; Cragg et  al. 2015; Edwards et al. 2017; Janusz et al. 2017; Bissaro et al. 2018). An extensive knowledge of fungal secretomes involved in recalcitrant plant biomass degradation is of immense significance in the present scenario where increasing emphasis is devoted toward sustainable bioeconomy. In the coming sections, we describe different carbohydrate-active enzymes (CAZymes) involved in lignocellulose degradation (Table  1.1 and Fig.  1.2). According to CAZy database, there are six types of CAZymes, i.e., glycoside hydrolase (GH), carbohydrate esterase (CE), glycosyltransferase (GT), polysaccharide lyases (PL), auxiliary activity (AA), and carbohydrate-binding domains (Lombard et al. 2014). It is estimated that the proportion of secreted proteins in fungal species ranges from 4 to 14% (Lowe and Howlett 2012). Degradation of recalcitrant plant biomass, viz., dead wood, wheat straw, fodder, etc., is accomplished by highly coordinated and synergistic actions of multiple CAZymes exhibiting a combination of oxidative, hydrolytic, and non-hydrolytic activities (Bugg et al. 2011; Lombard et al. 2014; Janusz et al. 2017; SistaKameshwar and Qin 2018; Bissaro et  al. 2018). The most difficult part of plant cell wall is lignin and it needs to be degraded before enzyme can access cellulose and

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Table 1.1 The enzymatic repertoire of wood-degrading fungi and anaerobic rumen fungi associated with degradation of recalcitrant lignocellulosic plant biomass Plant cell wall component Cellulose

Hemicellulose

Lignin

Enzymes involved Endo-1,4-β-D-glucanase (EC 3.2.1.4) Exo-1,4-β-D-glucanase (EC 3.2.1.91) Cellobiohydrolase (EC 3.2.1.91) β-Glucosidase (EC 3.2.1.21) Cellobiose dehydrogenase Glucose 1-oxidase Pyranose 2-oxidase Gluco-oligosaccharide oxidases PQQ-dependent pyranose dehydrogenase Lytic polysaccharide monooxygenases (LPMOs) Endo-β-1,4-xylanases (EC 3.2.1.8) β-D-xylosidase (EC 3.2.1.37) 1,4-β-D-endo-mannanases (EC 3.2.1.78) 1,4-β-D-mannosidases (EC 3.2.1.25) β-1,4-Galactosidase Galactomannan acetyl esterase Arabinoxylan arabinofuranohydrolase α-L-Arabinofuranosidases α-Glucuronidases Feruloyl esterases Acetyl esterases 4-O-Methyl glucuronoyl methylesterases Xyloglucan transferase/hydrolases α-Xylosidases Galactose 6-oxidase Lytic xylan oxidase Laccases (EC 1.10.3.2) Lignin peroxidases (EC 1.11.1.14) Manganese peroxidases (EC 1.11.1.13) Versatile peroxidases (EC 1.11.1.16) Dye-decolorizing peroxidase (EC 1.11.1.19) Glyoxal oxidase (EC 1.2.3.5) Aryl alcohol oxidases (EC 1.1.3.7) Pyranose 2-oxidase (EC 1.1.3.10) Cellobiose dehydrogenase (EC 1.1.99.18) Glucose oxidase (EC 1.1.3.4) Chloroperoxidases (EC 1.11.1.10) Glucose dehydrogenase (EC 1.1.99.10) Aromatic peroxygenases (EC 1.11.2.1)

CAZy families GH1, GH3, GH5_5, GH5_7, GH5_9, GH5_12, GH5_45, GH5_48 GH5_74, GH5_131, GH5_148, GH6, GH7, AA3_2, AA3_4, AA7, AA8-AA3_1(CBM1), AA8-AA12-CBM1, AA9, AA10, AA11, AA13, AA14, AA15

GH2, GH5_7, GH5_26, GH27, GH36, GH35, GH3, GH39, GH43, GH52, GH10, GH11, GH62, GH51, GH54, GH67, GH115, GH12, GH16, GH74, GH31, AA5_2, AA14, CE1, CE1–CE7, CE12, CE16, GE/GCE, CE15

AA1, AA2, AA3, AA5, AA6

CAZy carbohydrate-active enzymes, CBM cellulose-binding domains, GHs glycoside hydrolases, CE carbohydrate esterases, AA auxiliary activities, LPMOs lytic polysaccharide monooxygenases Sources: Lewis and Yamamoto (1990), Eastwood et al. (2011), Bugg et al. (2011), Lombard et al. (2014), Hori et al. (2014), Janusz et al. (2017), dos Santos et al. (2018), Bissaro et al. (2018), Brink et al. (2019), Ralph et al. (2019)

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Fig. 1.2  The carbohydrate-active enzymes involved in degradation and depolymerization of recalcitrant plant biomass consisting primarily of lignin, cellulose, and hemicellulose. White rot fungi, brown rot fungi, and anaerobic rumen fungi secrete an array either all or some of essentially involved lignocellulolytic enzymes on variable substrates under in situ and in  vitro conditions. Lignocellulose degradation is achieved by concerted and simultaneous action of several hydrolytic enzymes, oxidoreductases, peroxidases, free radicals, and other reaction mediators

hemicellulose components. This activity is achieved in white rot fungi by extracellular enzymes, viz., oxidoreductases (AA2), lignin peroxidases, laccases, manganese peroxidases, versatile peroxidases, copper radical oxidases (AA5), dye-decolorizing peroxidases, and phenol-oxidizing multicopper oxidases (AA1), and a number of mediators, e.g., reactive oxygen species, free radicals, and aromatic intermediates (Martinez et al. 2004; Kuuskeri et al. 2016; Janusz et al. 2017; Bissaro et  al. 2018; dos Santos et  al. 2018). A detailed list of these enzymes is provided in Table 1.1. Both lignin-­modifying enzymes and lignin-degrading auxiliary enzymes are involved in lignin degradation process (Bugg et al. 2011; Janusz et al. 2017; Bissaro et al. 2018). On the other hand, cellulose and hemicellulose degradation into disaccharides and monosaccharides is accomplished with the help

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of cellobiohydrolases (GH families GH6 and GH7), β-glucosidases (GH1 and GH3), endoglucanases (GH5, GH9, GH12, GH44, and GH45), lytic polysaccharide monooxygenases (AA9), polysaccharide lyases, and carbohydrate esterases (Kuuskeri et al. 2016; Janusz et al. 2017; Bissaro et al. 2018). Exoglucanases from GH family, GH6, GH7, and GH48, attack cellulose fibrils at the ends of the chain. In case of hemicelluloses, it is endo-­hemicellulases, exo-hemicellulases, and accessory enzymes which cleave chains at various positions and locations (Table 1.1). Compositional details of lignocellulosic plant biomass, microorganisms involved in its biodegradation, and associated enzymatic repertoire have also been extensively reviewed earlier by Blanchette (1991), Eastwood et al. (2011), Girard et al. (2013), Lombard et al. (2014), Guerriero et al. (2015), Cragg et al. (2015), Janusz et al. (2017), Gall et al. (2017), Edwards et al. (2017), Bissaro et al. (2018), dos Santos et al. (2018), and Hooker et al. (2019). Fungal secretome studies are increasingly facilitated by accelerated genome sequencing availability of advanced software, databases, algorithms, analytical tools, prediction models, and improved proteomic approaches (Table 1.2). As depicted in Fig. 1.2, lignin is acted upon by oxidative enzymes (laccase, lignin peroxidase, versatile peroxidase, manganese peroxidase) and auxillary activity redox enzymes (glyoxal oxidase, pyranose oxidase, aryl alcohol oxidase, methanol oxidase; Hori et al. 2014; Adesogan et al. 2019; Brink et al. 2019; Ralph et al. 2019). During lignin depolymerization by fungi, laccases and manganese peroxidases mainly act on phenols, and the nonphenolic lignin components are attacked by lignin peroxidase, whereas versatile peroxidases act on both (Bugg et al. 2011; Janusz et  al. 2017; Brink et  al. 2019; Ralph et  al. 2019). In addition to numerous well-­ characterized hydrolytic enzymes, lignocellulolytic fungi exhibit simultaneous secretion of an array of lytic polysaccharide monooxygenases (LPMOs) and several other oxidoreductases (Bissaro et al. 2018). LPMOs are a class of copper-dependent enzymes classified as auxiliary activities (AA) and belong to families AA9, AA10, AA11, AA13, AA14, and AA15 of CAZy (Bissaro et  al. 2018). LPMOs are known to hydroxylate carbons at scissile glycosidic bonds. Interestingly, H2O2 plays a crucial role in catalytic activity of LPMOs (Bugg et al. 2011; Janusz et al. 2017; Bissaro et al. 2018).

1.4  Wood-Degrading Fungi The number of species of potent wood-degrading fungi is more than 1000; however, the actual numbers are expected to be much higher. These fungi mainly exhibit saprotrophic mode of nutrition but sometimes may be showing parasitic attributes in forest ecosystems (Janusz et al. 2017). Ongoing research based on transcriptome and secretome has offered considerable insights on enzymatic machinery, and lignocellulose-­degrading capabilities of several fungal species adapted to saprophytic, plant pathogenic, symbiotic, anaerobic, and endosymbiotic life styles (SistaKameshwar and Qin 2018). Secretomic studies offer deeper insights into

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Table 1.2  List of some important fungal genome sequences, secretomes, transcriptome databases, and software packages Database/authority/Genome Description CAZy database Carbohydrate-active enzymes database MycoCosm 1000 Fungal Genomes Project Genome sequence of Phanerochaete chrysosporium, a white rot fungus Genome sequence of Postia placenta, a brown rot fungus Genome sequence of Orpinomyces C1A FunSecKB

First-ever genome sequence of wood-­ decaying fungi First genome sequence of brown rot fungi

Weblink http://www.cazy.org/ https://genome.jgi.doe.gov/mycocosm/ home Ohm et al. (2014) https://genome.jgi.doe.gov/mycocosm/ home, http://genome.jgi-psf.org/ Pchrysosporium2_2, Martinez et al. (2004) Martinez et al. (2009)

Genome sequence of Youssef et al. (2013) anaerobic rumen fungus Fungal Secretome http://bioinformatics.ysu.edu/ Knowledge Base secretomes/fungi.php http://bioinformatics.ysu.edu/ FunSecKB2 The Fungal Secretome secretomes/fungi2/index.php and Subcellular Proteome Knowledge Base 2.1 SRA NCBI Sequence Read https://submit.ncbi.nlm.nih.gov/subs/sra Archive FungiDB – http://fungidb.org/fungidb/ www.elignindatabase.com eLignin eLignin Microbial Database Brink et al. (2019) FSD The Fungal Secretome http://fsd.snu.ac.kr/index.php?a=view Database Software packages used in transcriptome and secretome data analysis and prediction 1. SignalP v4.1 2. SecretomeP v1.027 3. TargetP v1.1 4. TMHMM v2.0 5. ProtComp v9.0

mechanistic details of lignocellulose degradation by various species of bacteria and fungi in their respective natural habitats or under symbiotic associations with higher organisms. These studies have the potential to elucidate the life style adaptation and survival mechanisms of wood-degrading fungi on highly recalcitrant plant biomass as sole carbon source. In addition, secretomic analyses can facilitate our search for novel enzymes and exploitable secretary pathways essentially needed to establish industrial-scale and sustainable biofuels production. Here, we specifically focus on wood-decaying fungi and anaerobic rumen fungi well recognized for their exceptional lignocellulolytic potential. The challenging task of plant biomass depolymerization in natural environments is primarily accomplished by filamentous fungi.

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Wood-degrading fungi mostly belong to the class Agaricomycetes of phylum Basidiomycota, although some members of Ascomycota are also of considerable significance. Lignocellulolytic activity of wood-degrading fungi is manifested through concerted action of oxidoreductases, peroxidases, glycoside hydrolases, exoglucanases, endoglucanases, xylanases, and a number of other CAZymes (Gaskell et al. 2016; Bissaro et al. 2018). In addition, the role of H2O2 and reactive oxygen species is also crucial. The most extensively studied (in terms of transcriptome and secretome) wood-degrading fungi are Phanerochaete chrysosporium, Phanerochaete carnosa, Phlebia tremellosa, Trametes versicolor, Phlebia radiata, Bjerkandera adusta, Irpex lacteus, Gloeophyllum trabeum, Agaricus bisporus, Stropharia coronilla, Agrocybe praecox, Chondrostereum purpureum, Heterobasidium annosum, Ceriporiopsis subvermispora, Phellinus pini, Lentinula edodes, Hericium clathroides, Pleurotus ostreatus, Obba rivulosa, Postia placenta, Piptoporus betulinus, Serpula lacrimans, Fomitopsis lilacinogilva, Ganoderma lucidum, Laetiporus portentosus, Fomitiporia mediterranea, Pycnoporus cinnabarinus, Dichomitus squalens, Punctularia strigosozonata, Botrytis cinerea, Stereum hirsutum, Pleurotus eryngii, Fibroporia radiculosa, Wolfiporia cocos, Dacryopinax primogenitus, Daedalea quercina, Laetiporus sulphurous, Neolentinus lepideus, Calocera cornea, Fistulina hepatica, Hydnomerulius pinastri, and Coniophora puteana (Riley et al. 2014; Presley and Schilling 2017; SistaKameshwar and Qin 2018).

1.4.1  Secretomes of White Rot Fungi (WRF) WRF have the unique distinction of being the only microorganism in the biological world to completely degrade lignin, cellulose, and hemicellulose simultaneously (Manavalan et al. 2015; Xie et al. 2016). WRF are members of Basidiomycota, the largest phylum of kingdom Fungi (Blanchette 1991; Martinez et  al. 2004; Krah et  al. 2018). During lignin degradation by WRF, the crystalline cellulose with a bleached appearance is left behind which appears as white rot. This is why these are called white rot fungi. WRF initiate lignin depolymerization by producing large amount of free radicals mediated by oxidases and peroxidases (Martinez et  al. 2004). With the help of a consortium of enzymes, WRF carry out mineralization of lignocellulose plant matter to carbon dioxide and thereby ensure continuity of carbon cycle in the habitats characterized by forest litter, fallen trees, wooden stumps, wood logs, etc. In return, WRF fulfil their energy and nutrition requirements from the same substrate (Blanchette 1991; Martinez et al. 2004; Baldrian and Valaskova 2008; Eastwood et al. 2011; Krah et al. 2018; Bissaro et al. 2018). WRF from phylum Basidiomycota are reported to efficiently degrade lignin, cellulose, and hemicellulose present in the plant cell wall manifested through co-­ secretory and synergistic action of hydrolytic, oxidative, and non-hydrolytic enzymes, thus leaving a bleached fibrous residue (Krah et  al. (2018). WRF can uniquely attack lignin barrier first before gaining access to cellulose and hemicelluloses of plant cell wall. WRF also produce extracellular reactive oxygen

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species mediated by peroxidases which facilitate physical disruption of crystalline lignocellulose biomass leading to greater access for degradative enzymes (Martinez et al. 2004; Bissaro et al. 2018). P. chrysosporium is a model organism for studying lignin degradation. Its genome sequence is published and offers greater insights on lignin-­degradative enzymatic machinery and corresponding genomic organization (Martinez et al. 2004; Ohm et al. 2014). It is considered as the most prolific producer of CAZy, especially laccases, lignin peroxidases, manganese peroxidases, and LPMOs among various WRF species (Singh and Chen 2008). P. chrysosporium RP78 genome encodes >240 putative CAZymes belonging to 69 distinct families. Among these, 166 were glycoside hydrolases, 57 glycosyltransferases, 40 putative endoglucanases (GH5, GH9, GH12, GH61, and GH74), 14 carbohydrate esterases, at least 9 β-glucosidases, and 7 exocellobiohydrolases (Martinez et  al. 2004). Further, ten lignin peroxidases, five manganese peroxidases, and several other lignocellulolytic enzymes were encoded by its genome (Martinez et al. 2004). P. chrysosporium is an excellent decomposer of soft- and hardwood, branches, logs, leaves, etc. in forests. The secretomes of a number of other WRF growing in situ or on various substrates under in vitro conditions are now available in the literature. As compared to P. chrysosporium, which carry out simultaneous degradation of cellulose, hemicellulose, and lignin, Ceriporiopsis subvermispora exhibit unique and selective characteristic of lignin removal before initiating the cellulose degradation (Hori et al. 2014). Chondrostereum purpureum, a basidiomycetous fungus, produces an extensive repertoire of lignocellulolytic enzymes (Reina et  al. 2019). Almost 50% of CAZy encoded by its genome belongs to GHs (GH5, GH6, GH7, GH10, GH11, GH12, GH16, GH30), CEs, and cellulose-binding domains (CBMs) which specifically target cellulose and hemicelluloses. In addition, 153 oxidoreductases, lignin-­modifying enzymes, and auxillary activity (AA1, AA3) enzymes are encoded by its genome under variable culture conditions (Reina et al. 2019). Similarly, several families of CBMs and CEs were found to be encoded by C. purpureum genome. Further, expression of 81 oxidoreductases was recorded under substrate-specific culture conditions (Reina et al. 2019). Secretome analysis of Pleurotus eryngii, an edible mushroom and white rot fungi, revealed the production of seven glucanases, cellobiohydrolase, cellulose 1,4-beta-cellobiosidase, glucosidases, 22 glycoside hydrolase (GH families GH1, GH6, GH12, GH16, GH17, GH24, GH31, GH32, GH35, GH43, GH44, GH51, GH61, GH74, GH76, GH78, GH79, GH88, GH92, GH95), and CBM of family 21. These findings conclusively established the rich cellulolytic enzymes repertoire in P. eryngii under different substrate conditions (Xie et al. 2016). In another study, Kuuskeri et al. (2016) studied the secretory enzyme profile of Phlebia radiata cultured in solid state on spruce wood. The transcriptomic and secretomic analyses indicated expression and secretion of oxidoreductase, glyoxal oxidases, alcohol oxidases, cellobiohydrolases (GH6 and GH7), LPMO (AA9), lignin peroxidases, and acetyl xylan esterase. Prominent upregulation of genes whose products are involved in wood decay was observed at different growth periods.

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1.4.2  Secretome of Brown Rot Fungi (BRF) BRF are basidiomycetous fungi which occur as common pests of plants in conifer-­ dominated woodlands (Presley and Schilling 2017). BRF decompose wood via glycoside hydrolase-mediated saccharification and free radical oxidation (Baldrian and Valaskova 2008). However, compared to WRF, BRF preferentially degrade cellulose and hemicellulose whereas lignin is not depolymerized significantly thus leaving behind a brownish residue with fragmented appearance (Krah et al. 2018). Genomic analysis revealed evolution of BRF from WRF by gradual loss of genes which encode for ligninolytic peroxidases (Januszet al. 2017). Absence of class II peroxidases and cellobiohydrolases was reported in Wolfiporia cocos (Gaskell et al. (2016). The important species of BRF are Gloeophyllum trabeum, Postia placenta, Piptoporus betulinus, Serpula lacrimans, Fomitopsis lilacinogilva, Laetiporus portentosus, Wolfiporia cocos, Fibroporia radiculosa, Dacryopinax primogenitus, Daedalea quercina, Laetiporus sulphurous, Neolentinus lepideus, Calocera cornea, Fistulina hepatica, Hydnomerulius pinastri, and Coniophora puteana (Krah et al. 2018; SistaKameshwar and Qin 2018). Most of BRF are aerobic in nature and contribute a little inside digestive tracts of herbivorous mammals. WRF vary significantly in terms of substrate specificity and mechanisms of action. The genomic, phenotypic, and phylogenetic basis of these variations is yet to be fully understood. Presley and Schilling (2017) studied in vitro degradation of spruce wafers by two BRF, namely, Serpula lacrymans and Gloeophyllum trabeum, using a proteomic approach. Upon initial colonization, oxidoreductase diversity was observed first followed by higher glycoside hydrolase activity at later stages. Their findings suggest significant variations in their oxidoreductase profiles as indicated by presence of putative copper radical oxidase in S. lacrymans but absence in G. trabeum. On the other hand, GMC oxidoreductase and a xyloglucan-specific AA9 family protein were produced by G. trabeum but not by S. lacrymans. S. lacrymans exhibited higher mannanase activity compared to G. trabeum which showed elevated xylanase production. Interestingly, GH6 and cellobiohydrolases (CBHs) were not detected in case of S. lacrymans. As compared to 93 proteins identified in S. lacrymans, the protein counts were 209 in G. trabeum. Overall analysis of their secretomes indicates a two-step brown rot decay mechanism manifested through entirely different biochemical routes. In another BRF species Postia placenta from phylum Basidiomycota, 242 putative CAZY-encoding genes were reported (Martinez et al. 2009). Among these, the number of GHs, CEs, glycosyltransferases, and polysaccharide lyases were 144, 10, 75, and 6, respectively. In addition, expansin-like proteins, laminarinases, chitinases, endoxylanases, β-xylosidases, and L-α-arabinofuranosidases were identified. However, the enzymes/proteins involved in lignin degradation, viz., lignin peroxidase, manganese peroxidase, exocellobiohydrolases, versatile peroxidase, cellulose-binding domains, and cellulose-binding endoglucanases, were entirely absent. This unique secretome profile substantiated the non-action of P. placenta against lignin components of plant biomass. Gaskell et al. (2016) determined 64 glycoside hydrolases

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from Wolfiporia cocos growing on different media containing glucose, purified crystalline cellulose, lodgepole pine, and aspen. More than fourfold upregulation of hemicellulase-, endoxylanase-, and chitinase-encoding genes was observed. Additionally, there was upregulation of genes involved in oxidative depolymerization of cellulose.

1.5  Secretome of Anaerobic Rumen Fungi (ARF) ARF have indispensable role in digestion of recalcitrant lignocellulosic feed materials in digestive system of herbivorous animals. CAZymes in ARF exist either as free enzymes or as cellulosome, a multiprotein complex (Haitjema et  al. 2017). The genome sequencing of four species of Neocallimastigomycota suggests that many of these CAZymes have been acquired by horizontal gene transfer from rumen bacteria (Youssef et al. 2013; Haitjema et al. 2017). The extensive CAZyme repertoire, cellulosome, and extracellular proteases produced by Neocallimastigomycetes may help these microbes compete with other rumen inhabitants for limited nutrients (Youssef et al. 2013; Haitjema et al. 2017). As described earlier in Sect. 1.3, only 11 genera have been cultured and exhaustively investigated for secretome analyses. Due to their strict anaerobic lifestyles, in vitro studies are limited on these fungi. Still, a number of studies have offered insights in secretome profiles of ARF. Wang et  al. (2011) identified 25 families of glycosyl hydrolases (GHs) from anaerobic rumen fungus Neocallimastix patriciarum W5 culture anaerobically on substrate mixture comprising rice straw, napier grass, and sugarcane bagasse. Transcriptome and secretome analysis revealed 25 putative GH families dominated by GH6 (15%), GH10 (9.5%), GH5 (9.1%), and GH43 (9.1%). The main CAZymes were cellobiohydrolase (EC 3.2.1.91), endoglucanase (EC 3.2.1.4), and xylanases. The number of cellulases and hemicellulases was found to be higher in N. patriciarum W5 as compared to other plant matter-degrading fungi. The genome sequencing of Orpinomyces sp. strain C1A by Youssef et al. (2013) revealed an efficient lignocellulolytic enzymes repertoire comprising 357 glycoside hydrolase genes, 92 carbohydrate esterases genes, and 24 polysaccharide lyases genes. Interestingly, 220 genes with fungal dockerin domain and 103 genes harboring carbohydrate-binding module domains were also identified. Further, expansion of cellulolytic and hemicellulolytic families, viz., GH6, GH9, Gh10, GH11, Gh43, GH45, and GH48, and reduction or complete loss of families GH7, GH16, GH18, GH28, and GH61 were also observed (Youssef et al. 2013). The genes attributing for an efficient and extensive glycoside hydrolase machinery of this rumen fungus are believed to be acquired through horizontal gene transfer from multiple ruminal bacteria present in rumen. A previous study also reported presence of cellulase (GH family 48) containing two C-terminal fungal dockerin domains from Piromyces equi (Steenbakkers et al. 2002). Kameshwar and Qin (2018) compared the genome-wide annotations of five ARF, namely, Neocallismatix californiae, Anaeromyces robustus, Orpinomyces sp., Piromyces sp. E2, and Piromyces finnis. Findings of this comprehensive analysis

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revealed that ARF have the highest number of CAZyme-encoding genes compared to other fungi. Moreover, the presence of genes for cellulosomes and carbohydrate transport and metabolism strongly supported their remarkable polysaccharide-­ degrading abilities. Surprisingly, the genes encoding for lignin-degrading auxiliary activity enzymes, such as lignin peroxidase, laccase, manganese peroxidase, versatile peroxidase, aryl alcohol oxidase, glyoxal oxidases, and glucose oxidases, were completely lacking in their genomes. Among these five ARF species, Neocallimastix californiae was found to possess the highest number of genes coding for CAZymes involved in cellulolytic, hemicellulolytic, and pectinolytic activities (Kameshwar and Qin 2018). A comparative analysis of transcriptome of four ARF, i.e., Anaeromyces mucronatus YE505, Neocallimastix frontalis 27, Orpinomyces joyonii SG4, and Piromyces rhizinflata YM600 revealed that 8.1–11.2% of the entire transcriptome were predicted CAZymes with highest in O. joyonii. About 40–44% of the CAZymes-encoding contigs had one or more carbohydrate-binding modules (Gruninger et al. 2018).

1.6  Conclusion In spite of the availability of enormous amount of plant matter, woody substances, crop residues, and agro-food industry by-products as an attractive renewable resource, its industrial-scale utilization remains limited due to inherent structural complexity and recalcitrance. The controlled decomposition of biomass in general and of lignocellulose in particular involves a wide diversity of enzymatic activities and chemical reactions, which are far from fully elucidated. Moreover, our knowledge of the fungal secretion pathway is still at an early stage. Wood-degrading fungi and anaerobic rumen fungi can accomplish this daunting task with remarkable efficiency in natural environments using their specialized and sophisticated biomolecular machinery. The fungal secretomes have been explored to find enzymes and enzyme combinations for paper, textile, and food manufacturing industries. Similarly, low cost and sustainable processes for plant biomass conversion to biofuels can revolutionize industrial and environmental microbiology. The study of secretomes of novel fungal genera/species is interestingly poised to elucidate novel enzymes suitable for efficient plant cell wall degradation which can be exploited for commercial biotechnological applications. White rot fungi in particular have tremendous potential for biotechnological applications, bioremediation, pulp and paper industries, and effluents treatment in different industrial settings. These fungi possess remarkable potential for implementing eco-friendly, sustainable, and consolidated biological processing of lignocellulosic biomass for biofuel and biorefining industries and bioremediation processes. Acknowledgments  NS is grateful to The Chancellor, Eternal University, for their financial support and infrastructural facilities. The authors are thankful to Dr. Sumit Singh Dagar for his valuable and expert suggestions.

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Reina R, Kellner H, Hess J, Jehmlich N, García-Romera I, Aranda E, Hofrichter M, Liers C (2019) Genome and secretome of Chondrostereum purpureum correspond to saprotrophic and phytopathogenic life styles. PLoS One 14:e0212769. https://doi.org/10.1371/journal.pone.0212769 Riley R, Salamov AA, Brown DW, Nagy LG, Floudas D, Held BW et  al (2014) Extensive sampling of basidiomycete genomes demonstrates inadequacy of the white-rot/brown-rot paradigm for wood decay fungi. Proc Natl Acad Sci U S A 111:9923–9928.  https://doi. org/10.1073/pnas.1400592111 Saha BC (2003) Hemicellulose bioconversion. J  Ind Microbiol Biotechnol 30:279–291.  https:// doi.org/10.1007/s10295-003-0049-x Scheller HV, Ulvskov P (2010) Hemicelluloses. Annu Rev Plant Biol 61:263–289. https://doi. org/10.1146/annurev-arplant-042809-112315 Siegenthaler U, Sarmiento JL (1993) Atmospheric carbon dioxide and the ocean. Nature 365:119– 125. https://doi.org/10.1038/365119a0 Singh D, Chen S (2008) The white-rot fungus Phanerochaete chrysosporium: conditions for the production of lignin-degrading enzymes. Appl Microbiol Biotechnol 81:399–417. https://doi. org/10.1007/s00253-008-1706-9 Sirohi SK, Singh N, Dagar SS, Puniya AK (2012) Molecular tools for deciphering the microbial community structure and diversity in rumen ecosystem. Appl Microbiol Biotechnol 95:1135– 1154. https://doi.org/10.1007/s00253-012-4262-2 SistaKameshwar AK, Qin W (2018) Comparative study of genome-wide plant biomass-degrading CAZymes in white rot, brown rot and soft rot fungi. Mycology 9:93–105. https://doi.org/10.1 080/21501203.2017.1419296 Steenbakkers P, Freelove A, Van Cranenbroek B, Sweegers B, Harhangi H, Vogels G, Hazlewood G, Gilbert H, Op den Camp H (2002) The major component of the cellulosomes of anaerobic fungi from the genus Piromyces is a family 48 glycoside hydrolase. DNA Seq 13:313–320. https://doi.org/10.1080/1042517021000024191 Solomon KV, Haitjema CH, Henske JK, Gilmore SP, Borges-Rivera D, Lipzen A et  al (2016) Early-branching gut fungi possess a large, comprehensive array of biomass-degrading enzymes. Science 351:1192–1195. https://doi.org/10.1126/science.aad1431 Tjalsma H, Bolhuis A, Jongbloed JD, Bron S, van Dijl JM (2000) Signal peptide-dependent protein transport in Bacillus subtilis: a genome-based survey of the secretome. Microbiol Mol Biol Rev 64:515–547. https://doi.org/10.1128/MMBR.64.3.515-547.2000 Wang TY, Chen HL, Lu MJ, Chen YC, Sung HM, Mao CT, Cho HY, Ke HM, Hwa TY et al (2011) Functional characterization of cellulases identified from the cow rumen fungus Neocallimastix patriciarum W5 by transcriptomic and secretomic analyses. Biotechnol Biofuels 4:24. https:// doi.org/10.1186/1754-6834-4-24 Xie C, Yan L, Gong W, Zhu Z, Tan S, Chen D, Hu Z, Peng Y (2016) Effects of different substrates on lignocellulosic enzyme expression, enzyme activity, substrate utilization and biological efficiency of Pleurotus eryngii. Cell Physiol Biochem 39:1479–1494. https://doi. org/10.1159/000447851 Yadav AN, Mishra S, Singh S, Gupta A (2019a) Recent advancement in white biotechnology through fungi Volume 1: diversity and enzymes perspectives. Springer International Publishing, Cham Yadav AN, Mishra S, Singh S, Gupta A (2019b) Recent advancement in white biotechnology through fungi. Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham Youssef NH, Couger MB, Struchtemeyer CG, Liggenstoffer AS, Prade RA, Najar FZ, Atiyeh HK, Wilkins MR, Elshahed MS (2013) The genome of the anaerobic fungus Orpinomyces sp. strain C1A reveals the unique evolutionary history of a remarkable plant biomass degrader. Appl Environ Microbiol 79:4620–4634. https://doi.org/10.1128/AEM.00821-13

Chapter 2

Bioremediation: New Prospects for Environmental Cleaning by Fungal Enzymes Neha Vishnoi and Sonal Dixit

2.1  Introduction The planet ‘Earth’ is endowed with rich wealth of natural resources such as forests, wildlife, land, soil, air, water, wind, plants and animals. One of the major problems in present scenario faced by the earth is the contamination of soil, water, and air with toxic chemicals released from numerous human activities. The battle between human and natural resources started when humans began living a stable life rather than a nomadic life. The use, overuse, and now the misuse of natural resources since the beginning of the civilization have led to its depletion to an extent that today half of our natural wealth are either depleted or at the edge of depletion (Gosavi et al. 2004). In early times, it was believed that our land and its resources are in abundance and will remain available for decades. However, today, existing state of our resources shows carelessness and negligence in using them. There are various anthropogenic activities like use of chemical fertilizers in agriculture, release of industrial waste, burning of fossil fuels, etc. which point toward the exploitation of natural resources. Rapid industrialization and the extensive use of pesticides and chemicals in agriculture and various fields have been proved to be the major cause of environmental pollution. The continuous use of organic compounds has become a serious threat to our eco system. Various industrial and anthropogenic activities have resulted in increased polluted sites due to ignorance regarding production and proper use and disposal of hazardous substances. There are various conventional techniques developed to decontaminate the contaminated sites. Conventional technique mainly

N. Vishnoi Department of Environmental Sciences, Babasaheb Bhimrao Ambedkar University, Lucknow, India S. Dixit (*) Department of Botany, University of Lucknow, Lucknow, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_2

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removes the contaminated soil to a landfill or covers the contaminated sites but this may create major risks in the excavation, handling, and transport of hazardous material. At the same time, these methods are quite expensive and difficult to find an alternative landfill sites for the final disposal of material. Other techniques like usage of chemicals for decomposition, UV oxidation and dechlorination, and incineration techniques have been employed and found to be effective in reducing the level of contaminants chlorinated solvents, petroleum, ketones, polyaromatic hydrocarbons (PAHs), trinitrotoluene (TNT), inorganic nitrogen, explosives, pesticides, herbicides and heavy metals, but at the same time, they show several demerits such as technical complexity, lack of public acceptance, and increased contaminants exposure to site workers and nearby residents (Vidali 2001). Various physicochemical treatments such as coagulation with alum or lime followed by adsorption on powdered activated carbon (PAC) are reported to yield high removal efficiency for phenolics. But, on the other hand, these processes generate large volume of hazardous sludge also and do not lead to the complete degradation of the contaminants (Mehta and Chavan 2009). To overcome these problems, an alternative method known as bioremediation has been used to completely remove or transform the contaminants into some biodegradable forms. In this technique, natural biological activity has been used to clean environment and its resources by degrading various contaminants. It is proved to be a safer, cleaner, economical, and eco-friendly technology which received a high public acceptance and can often be carried out at any site. According to Van Dillewijn et al. (2007), “bioremediation” is defined as the process by which various biological agents primarily microorganisms used to degrade or detoxify environmental contaminants. Bioremediation agents were defined as microbiological cultures, enzyme, and nutrient additives that significantly accelerate the rate of biodegradation to mitigate the effect of various contaminants. Bioremediation has been proved to be more advantageous than conventional methods due to high efficiency, cost-effectiveness, minimal release of chemical and biological sludge, selectivity to specific metals, regeneration of biosorbent, and the possibility of metal recovery (Kratochvil and Volesky 1998). A huge number of microbial enzymes and plant enzymes have been notified to be implicated in the biodegradation of harmful organic contaminants. Several studies have proved that enzymes from white-rot fungi are effectual degraders of so many organic pollutants like dyes, PAHs, phenols, polychlorinated biphenyls (PCBs), and TNT (Yadav et  al. 2019a, b). Numerous enzymes such as cellulase, lipase, and protease prove to have the capability to lessen solid contents and number of pathogens and enhance deflocculation in the sludge. Enzymes like hydrolases and oxidoreductases are used widely in bioremediation. Extracellular enzymes are those which are either secreted by the microorganisms (e.g., lignin peroxidase (LiP) from white-rot fungi) or which go through the aqueous stage during an aerobic submerged fermentation procedure (Bhargava et al. 2003). The extracellular enzymes have been used in several industries for various applications, but currently they are used widely for enhancing bioremediation practices. Each and every enzyme generally has only one particular function, for example, to lessen the activation energy for the degradation of an intramolecular bond, but few enzymes can act on wide variety

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of varying substrates (Fullbrook 1996; Gianfreda and Rao 2004). These second types of enzymes are mainly suitable for bioremediation purposes. These enzymes are capable to react with xenobiotic and synthetic materials and so convert them from a recalcitrant state to biodegradable form (Bollag 1992; Gianfreda and Rao 2004). Extracellular enzymes allow more competent treatment practices for biodegradable matter because they are capable of accelerating the rate of degradation of these substances. The application of enzymes is gaining much attention because enzymes can carry out the equivalent or even better function as compared to harsher chemicals and they also do not produce any type of hazardous waste. While enzymes are possibly little expensive because of the associated cost of extraction and purification process, they can on the other hand tend to be very cost-effective as they reduce waste disposal and heating requirements (Godfrey and Reichelt 1996; Gianfreda and Rao 2004). Fungus acts as an important agent in bioremediation because of their vigorous morphology and varied metabolic ability. Bioremediation process by using fungi as a tool for harmful organic remediation is an eco-friendly and sustainable way for clean-up of contaminated sites. Profound investigation toward this area would greatly assist in the development of advanced bioprocess technologies to decrease the level of contamination as well as to find some novel valuable materials. This chapter emphasizes on the importance of fungal enzymes for efficient bioremediation of a large variety of environmental pollutant. We will discuss in detail the multiple approach used by fungi for detoxification of various noxious and recalcitrant compounds including prominent fungal enzymes such as oxidoreductase, laccases, catalases, peroxidases, etc.

2.2  Bioremediation Bioremediation is considered as one of the safer, cleaner, cost-effective, and eco-­ friendly technologies for decontaminating sites which are contaminated with varied range of pollutants. Various biological processes are involved for the removal/ reduction of hazardous chemicals present in the environment (Kour et al. 2019a, b; Gianfreda and Rao 2004). The term “bioremediation” refers to all biochemical reactions of natural attenuation which include all biotic and abiotic processes used to minimize contaminant levels. This practice uses various biological agents such as higher plants, bacteria, algae, yeast, fungi, and enzymes to degrade environmental pollutants. Bioremediation can be further divided into phytoremediation and microbial bioremediation (Fig. 2.1). Phytoremediation refers to the in situ utilization of plants, their enzymes, roots, and associated microorganisms to degrade pollutants present in soil, water, and air. Microbial bioremediation involves the use of microorganisms for conversion and removal of pollutants from various environmental systems. The technique of bioremediation chiefly relies on microbes producing enzymes utilized to detoxify the contaminants and changes them to harmless products. Environmental conditions play a crucial role for microbial bioremediation

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BIOREMEDIATION (clean up of pollutants by living organisms)

MICROBIAL BIOREMEDIATION (remediation of pollutants by using microbes)

PHYTOREMEDIATION (remediation of pollutants by using plants)

ENZYMATIC BIOREMEDIATION (remediation of pollutants by enzymes secreted by organisms)

Oxidoreductase

Lyases

Peroxidase

Hydrolase

Dehalogenase

Fig. 2.1  An overview of types of bioremediation

as optimal conditions are necessary for the growth and vital activities of microorganisms. Effective bioremediation often involves the alteration of environmental conditions to permit growth of microbes and degradation to carry on at a faster rate. Bioremediation techniques can be classified as in situ and ex situ on the basis of removal and transportation of wastes (Fig. 2.2). In situ bioremediation technique involves treatment of contaminated material at the same site, whereas ex situ involves complete removal of contaminated material from one site to another site where it has been treated using biological agents. When both the methods have been compared, it was found that the rate of biodegradation and consistency of process outcome differ between in situ and ex situ methods. The cost of ex situ bioremediation is relatively high as compared to in situ due to the cost of excavation of contaminated samples for treatment. Both the bioremediation methods depend essentially on microbial metabolism, but in situ methods are favored over ex situ for ecological restoration of contaminated site (Vidali 2001). Microbes are ubiquitous on the biosphere due to their remarkable metabolic activity and ability to grow in a diversified environmental condition (Yadav et al. 2015a, b, 2017b). The nutritional versatility of microbes can also be exploited for breakdown of contaminants. Microorganisms act as an effective tool for pollutant decontamination in soil, water, and sediments as they are advantageous over other remediation procedures. They restore the original condition of natural surroundings and prevent further pollution (Demnerova et al. 2005). Several strains of microbes

2  Bioremediation: New Prospects for Environmental Cleaning by Fungal Enzymes Fig. 2.2  An outline of different methods of bioremediation

Intrinsic bioremediation

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Engineered bioremediation

In situ

(on site bioremediation)

Bioremediation

Ex situ

(off site bioremediation)

Solid Phase

Slurry Phase

are known to act as agents of bioremediation under laboratory conditions. Microbes can survive in extreme conditions of environment but effective bioremediation with enzymatic processes usually prefer optimal conditions which are difficult to achieve outside the laboratory (Bernhard-Reversat and Schwartz 1997; Vidali 2001; Dua et al. 2002; Dana and Bauder 2011). Optimal conditions can be achieved easily in ex situ technique than in situ technique. Therefore, in situ bioremediation by enzymes may be considered as less effective and consumes more time and requires a high concentration of enzyme when compared with ex situ bioremediation. The growth of microbes gets affected by numerous factors like pH, temperature, oxygen, soil structure, moisture, proper amount of nutrients, poor bioavailability of contaminants and presence of other toxic compounds. Bioremediation methods usually take place under aerobic conditions, but anaerobic conditions may also allow degradation of recalcitrant compounds aided by microbes. Both intracellular and extracellular enzymes produced by various groups of bacteria and fungi participate in the degradation of xenobiotics (Hammel 1997; Vidali 2001). The process of bioremediation depends on the metabolic potential of microorganisms, accessibility, and bioavailability of pollutant for removal/detoxification of pollutant (Antizar-Ladislao et al. 2008). The process of remediation can be accelerated by the addition of various microorganisms (called seeding or inoculation) to a contaminated site to enhance rate of biodegradation. The inoculums may be a blend of non-indigenous microbes (specially selected and cultivated for their various

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pollutant-­degrading characteristics) from various contaminated environments, or it may be a mixture of microbes selected from the site to be remediated or ­mass-­cultured in the laboratory. Addition of nutrients along with seeding process accelerates the process of bioremediation (Boopathy 2000). With the advances in biotechnology, bioremediation has become one of the significant developing fields of ecological restoration that utilizes microorganisms to minimize the concentration and toxicity of various chemical pollutants such as heavy metals, dyes, pesticides, etc. Considerable efforts have been devoted for developing low-cost removal technologies that can effectively immobilize dissolved toxic metals. Different types of biomass have been tested to remove and/or recover valuable metals for their possible reuse as reported by several authors (Say et al. 2001; Adhiya et al. 2002; Sheng et al. 2004). A strain of Pseudomonas putida was registered in 1974 as the first patent agent for biological remediation to degrade petroleum (Prescott et al. 2002). About 70 microbial genera were reported in 1991 (US Congress 1991) to degrade petroleum compounds and almost an equal number has been added to the list in the successive two decades (Kumar et al. 2011).

2.2.1  Types of Bioremediation Bioremediation can occur naturally on its own (natural attenuation or intrinsic bioremediation) or can be spurred via addition of fertilizers for the enhancement of bioavailability within the medium (biostimulation). There are different types of techniques under bioremediation processes such as biostimulation, bioattenuation, bioaugmentation, bioventing, bioleaching, bioreactor and piles (Li and Li 2011). 2.2.1.1  Biostimulation In this type of technique, specific nutrients at the polluted sites are injected to stimulate the activity of indigenous microbes. It relies upon the stimulation of indigenous or naturally existing microbial population by supplying fertilizers, growth supplements and trace minerals and by providing other environmental requirements like pH, temperature, and oxygen to accelerate their metabolism rate and pathway (Kumar et  al. 2011; Adams et  al. 2015). The presence of minimal amount of pollutant can also behave as stimulant by turning on the operons for bioremediation enzymes. This type of strategic pathway is mostly continued with the addition of nutrients and oxygen in order to help indigenous microorganisms. These nutrients fulfil the nutritional demand of the microbes and are considered as the basic building blocks of life. They permit microbial growth by providing the basic requirement like energy, cell biomass, and enzymes to decontaminate the polluted site. They also need some essential components like nitrogen, phosphorous and carbon (Madhavi and Mohini 2012).

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2.2.1.2  Bioattenuation It is also known as natural attenuation which means elimination of varying concentrations of contaminants from surroundings. It is a physical phenomenon and takes place through biological process via dispersion, dilution, diffusion, volatilization, and sorption/desorption by chemical reactions like ion exchange, complexation and abiotic transformation. Natural attenuation can also be termed as intrinsic remediation or biotransformation (Mulligana and Yong 2004). The nature can perform its work in four different ways to decontaminate the environment with chemicals. First, tiny bugs or microbes living in the soil and groundwater utilize some chemicals in order to fulfil their food requirement. They transform these contaminants into water and harmless gases on complete digestion. Second, chemicals are held by the soil through sorption process which restricts their movement and ultimately does not cause groundwater pollution. This process is not used in cleaning the chemicals but creating obstruction in the flow of chemicals to other site. Third, when contaminants move through soil and groundwater, their dilution occurs as it mixes with clean water. This reduces the level of contamination. Fourth, the pollutants like oil and various solvents easily convert their state from liquid to gas and are ultimately destroyed by the sunlight. When the natural attenuation does not work quickly or completely, then bioremediation can be accelerated by the process of either biostimulation or bioaugmentation (Li et al. 2010). 2.2.1.3  Bioaugmentation When natural or engineered population of microorganism responsible for degradation of contaminants is added to augment or enhance the biodegradative capability of indigenous of microbes on the contaminated site, the process is known as bioaugmentation. The growth and degradation capacity of naturally present microbes can be increased by their collection from remediation site, then culturing, making genetic modification, and then returning to the same site. A study by Niu et  al. (2009) suggested that all vital microbes are present in their sites where soil and groundwater are contaminated with chlorinated ethenes such as tetrachloroethylene and trichloroethylene which are used to ensure that the in situ microorganisms can completely convert these contaminants to non-toxic ethylene and chloride. Bioaugmentation is the process in which engineered microbes are added as a bioremediator in a system in such a manner so as to completely eradicate complex pollutants in a fast manner. Genetically engineered microbes (GEMs) having diverse metabolic profile of detoxification proved to be the suitable candidates for the degradation of a wide range of environmental pollutant effectively (Malik and Ahmed 2012; Alwan et al. 2013; Gomez and Sartaj 2014). Natural species do not degrade all the compounds quickly, so to facilitate their degradation, species must be modified at gene level through DNA manipulation. The capacity of degradation in genetically engineered microbes is more than natural species and they highly compete

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with the indigenous species, predators and also with various abiotic factors. Due to enhanced degradation properties, genetically engineered microbes have shown potential for bioremediation of soil, groundwater and activated sludge contaminated with a broad range of toxicants (Sayler and Ripp 2000; Thapa et  al. 2012). Researchers are continuously diverting their focal point toward the ways to augment contaminated sites with various other non-native microbes especially genetically modified microorganisms. The recombinant DNA and other molecular biological techniques have shown certain advantages: (i) enabled amplification, disruption, and modification of the targeted genes that encode the enzymes of metabolic pathways, (ii) minimized bottlenecks pathway, (iii) enhanced redox and energy generation, and (iv) played important role in recruiting heterologous genes to give new characteristics (Liu et al. 2006). It can be said that the process of bioaugmentation will open a new series of possibilities for future process of bioremediation. Genetically engineered microorganisms (GEMs) are those whose genetic material has been altered by using various techniques of genetic engineering inspired by natural or artificial genetic exchange between microbes. This process is known as recombinant DNA technology. Recombinant DNA techniques or natural genetic material exchange between organisms resulted in the development of recombinant living organisms. In the present scenario, scientists are successful in inserting the suitable gene for the production of particular pollutant-degrading enzyme (Jain et al. 2010). Recently, a number of opportunities come forward for the improvement of degradation ability using genetic engineering strategies. The rate-limiting steps in metabolic pathways can be increased and altered or new pathways incorporated in microbial strains to enhance their degradation capacities for xenobiotics. Four strategies are followed in genetic engineering microbes: (1) modification of enzyme specificity and affinity, (2) pathway construction and regulation, (3) bioprocess development, monitoring, and control, and (4) bioaffinity bioreporter sensor applications for chemical sensing, toxicity reduction, and end-point analysis. Vital genes of bacteria are carried on a single chromosome but genes for enzymes which are needed for the catabolism of some of the contaminants may be carried on plasmids. Plasmids have been implicated in the catabolism. Therefore, GEMs can be used efficiently for biodegradation process to optimize the eradication of hazardous unwanted wastes, which symbolizes a research frontier with broad implications in the coming future (Jain et al. 2011; Kulshreshtha 2013). The major advantage of GEMs is that they accelerate the recovery of contaminated sites, increase substrate degradation, display a high catalytic or utilization capacity with small amount of cell innoculum, and create eco-friendly environmental conditions by the mechanism of detoxification. The major drawbacks of using genetic engineered microbes are that they are never used in traditional procedure and the risk associated with the release of dead cellular mass. Biotic and abiotic factors are directly correlated and also play significant role in the functioning of microbes. Introduction of foreign modified strain to the system leads to deleterious effects on the original structural composition, occurrence, and functional properties of microbes.

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2.2.1.4  Bioventing Bioventing is involved in venting of oxygen through soil to stimulate growth of natural or introduced bacteria and fungus in the soil by providing oxygen to existing soil microorganisms; indeed, it is functional in aerobically degradable compounds. Bioventing uses low airflow rates to provide only enough oxygen to sustain microbial activity. Oxygen is most commonly supplied through direct air injection into residual contamination in soil by means of wells. Adsorbed fuel residuals and volatile compounds are biodegraded which thereafter move slowly through biologically active soil. Some researchers like Agarry and Latinwo (2015) proved bioventing to be an effective technique in bioremediation of petroleum-contaminated soil. 2.2.1.5  Biopiles In this technique, soil is excavated and then spread in the form of piles. Biopiles are also known with other names such as biocells, bioheaps, biomounds, and compost piles which are employed for reducing petroleum contaminants in excavated soils during the course of biodegradation. In this technique, air is supplied to the biopile system during a system of piping and pumps that either forces air into the pile under positive pressure or draws air through the pile under negative pressure (Delille et al. 2008). The microbial activity is enhanced through microbial respiration which results in the accelerated degradation of adsorbed petroleum contaminants (Emami et al. 2012; Kumar et al. 2016).

2.2.2  Limitations of Bioremediation Bioremediation is restricted to only those compounds that are degraded by biological means and not to those which cannot be subjected to degradation. Important concern should be taken to check the toxicity and persistent nature of products synthesized during biodegradation. The biological methods are extremely specific with culture requirements and at times are difficult to extrapolate the results from lab to field. The time duration for these methods is comparatively more than for excavation treatment. There are various factors affecting the process of bioremediation such as depletion of preferential substrates, lack of nutrients, toxicity and solubility of contaminants, oxidation or reduction potential, and microbial interaction. The outcome of each degradation process depends on microbes (biomass concentration, population diversity, and enzyme activities), substrate (physicochemical characteristics, molecular structure, and concentration), and a range of environmental factors (pH, temperature, moisture content, availability of electron acceptors, and carbon and energy sources). These parameters affect the acclimation period of microbes to the substrate. The molecular structure and contaminant concentration have been shown to strongly affect the feasibility of bioremediation. The type of

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microbial transformation depends on whether the compound serves as a primary, secondary, or co-metabolic substrate (Boopathy 2000). All the contaminants are not easily treated, accumulated, or degraded by bioremediation using microorganisms, and the effects of microorganisms on the metal leaching associated with phytoremediation have yet not received proper attention (Neagoe et al. 2009).

2.3  Fungi as Bioremediator Mycoremediation or fungal remediation is a type of bioremediation which uses fungi to remove the pollutants from the environment. Mycoremediation utilizes fungal mycelium to expel the waste material from the contaminated site. Fungal colonization can occur in a variety of habitats, i.e., fresh water and marine with complex soil lattice. Fungi can flourish in varying extreme climatic conditions and germinate through the dispersal of its spores in the atmosphere (Anastasi et al. 2013). There are several reports which demonstrate that some fungi also grow well in industrial effluent treatment plants (Zhang et  al. 2013; Badia-Fabregat et  al. 2015). Their potential to survive in diverse habitats and secreting multiple enzymes enable them to be the best candidate for bioremediation. The fungal mycelia secrete different types of extracellular enzymes and acids that are utilized to convert the lignin and cellulose into simpler forms. Fungal species are specific for a particular pollutant which is an important feature for fungal remediation. White-rot fungi like Phanaerochaete chrysosporium and Polyporus sp. which are also known as ligninolytic fungi are employed for the removal of numerous persistent or toxic environmental contaminants such as petroleum hydrocarbons, PAHs, explosives, PCBs, and organochlorine pesticides and thus prove to be the best candidate for bioremediation (Wu and Yu 2007; Ayu et al. 2011). They are reported for the degradation of numerous organopollutants as they secrete an enzyme which acts on lignin. White-rot fungi secrete extracellular enzymes during the decaying process of lignin so they have been proved to be the potential agents for oxidizing pollutants. A bird’s nest ligninolytic fungus known as Cyathus bulleri which colonizes on dead herbaceous stems, wood chips, dung, sticks, and other woody debris are found cost-effectively appropriate for the degradation of lignin  as it  produces a single laccase, an internal peptide which shows resemblance with enzyme  laccase of white-rot fungi with differences in proportions of some basic and hydrophobic amino acids. Laccases are copper-containing enzymes using molecular oxygen as the electron acceptor for catalyzing the oxidation of a varying range of phenolic and aromatic amine compounds (Garg et  al. 2008). Many workers have showed that the fungi belonging to these groups were utilized for the oxidation of lignin which shows structural similarity with PAH compounds by the secretion of extracellular enzymes via mineralization with carbon dioxide as end product (Levin et al. 2003; Mai et al. 2004). The process of degradation is not yet decisively mapped but is expected that intracellular and extracellular enzymes are responsible for oxidizing different

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p­ ollutants under different environmental conditions. Fungal enzymes have also been utilized for the degradation and decolorization of azo dyes (Vidali 2001; Husain 2006). Metals like uranium, cadmium, zinc, copper, and cobalt are biosorbed by microbes through electrostatic interactions between the metal ions present in solutions and cell walls (Bai and Abraham 2001). Chromium biosorption in fungi like Ganoderma lucidum and Aspergillus niger takes place through ion exchange mechanism (Muraleedharan and Venkobachar 1990; Ahluwalia and Goyal 2006). Pleurotus ostreatus (tasty oyster mushroom) exhibit a broad range of applications in bioremediation. A report by Favero et al. (1991) concluded that it is used in remediation of soil contaminated with diesel oil and can be used in the conversion of 95% PAH to non-toxic components. Fungi which act as wood degraders are particularly effective in degrading aromatic pollutants and chlorinated compounds (Arıca et al. 2004). It is found that the indigenous microbe also participates with the fungi for the mineralization of pollutants into carbon dioxide and water. Yeasts possess the property to resist under unfavorable environmental conditions and can accumulate metals and metalloids which are vital for structural and catalytic functions (Chatterjee et al. 2011). Some examples of yeast species possessing degrading capabilities are proved to be suitable representatives of remediation of pollutants from environment: Candida, Clavispora, Debaryomyces, Leucosporidium, Pichia, Rhodosporidium, Rhodotorula, Sporidiobolus, Sporobolomyces, Stephanoascus, Trichosporon, and Yarrowia (Csutak et al. 2010). Dye decolorization from industrial effluents by yeasts mainly occurs through three mechanisms: biosorption, bioaccumulation, and biodegradation (Donmez 2002). Candida utilis accumulates metal ions and radionuclides from the environment (Kujan et al. 2006). Bioadsorption of lead and accumulation of free and complexed silver ions in yeast Rhodotorula mucilaginosa has been achieved by metabolism-dependent and metabolism-­independent processes (Gomes et al. 2002). A comparative study was conducted by Ksheminska et al. (2003) in which Pichia guilliermondii was used to check uptake potential of chromium and it was concluded that chromium tolerance increases on addition of riboflavin.

2.3.1  White-Rot Fungi These are considered as important agents for biodegradation of lignin-containing material in environment and therefore also contribute largely to the carbon recycling. Previous studies showed the bioremediation prospective of white-rot fungi like Bjerkandera adjustam, P. chysosporium, Pleurotus sp., and Trametes versicolor in the degradation of recalcitrant compounds by secreting various ligninolytic enzymes like laccases and peroxidases, which results in bioaccumulation, toxicity to aquatic life, and deleterious effects to human beings (dos Santos Bazanella et al. 2013). White-rot fungi secrete ligninolytic enzymes that have been utilized for the detoxification of a range of organic contaminants like pesticides from polluted

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wastewaters (Rodríguez-Rodríguez et al. 2013). Extracellular ligninolytic enzymes of fungi are also capable for the adsorption of various dyes. This property of white-­ rot fungi established them as a governing power in the field of dye removal or decolorization as confirmed by the decolorization of Direct Blue 14 by different species of Pleurotus (Singh et al. 2013a) and Remazol Brilliant Blue-R by Agaricomycete (a white-rot fungus from Amazon forest) (dos Santos et al. 2015). Various fungal species like Coriolus versicolor, Hirschioporus larincinus, Inonotus hispidus, P. chrysosporium, and Phlebia tremellosa have been previously examined for the decolorization of different dye effluents (Jebapriya and Gnanadoss 2013). White-rot fungi like Lentinus tigrinus and T. versicolor were reported for the decontamination of soil contaminated with cresolate via. bioaugmentation process (Lladó et  al. 2013). Residues of petroleum hydrocarbons and PAHs with high molecular weight are mainly responsible for the pollution of cresolate soil. This problem of petroleum residue contamination can be solved by significant degradation of these residues through biostimulation by means of lignocellulosic substrate together with bioaugmentation of fungi. However, there is a usual chance with this treatment that it may support the growth and intensification of local microorganisms which can rule over the augmented microbes. This can pressurize the requirement to authenticate such kind of study at a small level prior to the field applications. Besides these applications of ligninolytic enzymes for bioremediation of a range of substances, laccases have also been utilized for the degradation of substituted organic compounds with improved removal efficiencies (Cutright and Erdem 2012; Fan et al. 2013; Purnomo et al. 2013). Considering the significance of such features in bioremediation, laccase production was maximized in T. versicolor and P. ostreatus by solid-state fermentation on orange peels. Further examination of bioremediation capabilities of these white-rot fungi revealed enhanced potential for PAHs such as phenanthrene and pyrene removal (Rosales et al. 2013). For an improved perceptive and utilization of bioremediation potential of fungi completely, genomic-level research of these fungi is needed.

2.3.2  Marine Fungi The application of marine fungi in bioremediation of heavy metals and hydrocarbons along with their capability to produce biosurfactants, secondary metabolites, polyunsaturated fatty acids, polysaccharides, and novel enzymes are well acknowledged a long time ago (Damare et al. 2012). The adaptation ability of marine fungi toward extremophilic conditions like high salinity and pH variations proved an advantage in comparison to terrestrial fungal species. The effectiveness of marine microorganism in metal removal provides an additional opportunity of using these extremophilic microbes for bioremediation purposes. Thatoi et al. (2013) examined the application of mangroves marine fungus for their enormous ecological and biotechnological functions like production of novel enzymes, drugs, biopesticides, biodiesel and their potential use in bioremediation. Numerous aspects related to fungi

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have been examined to increase the bioremediation potential of fungi for removal of hazardous and persistent organic contaminants. The noteworthy role of enzymes obtained from marine fungi and its biotechnological significance were demonstrated by Bonugli-Santos et  al. (2015). Bioremediation properties of enzymes of marine basidiomycetes obtained from marine sponges and Cerrena unicolor (marine white-rot basidiomycete) were successfully examined for decolorization of dyes Remazol Brilliant Blue-R and anthraquinone Reactive Blue 4, respectively (Bonugli-Santos et  al. 2012). Divya et  al. (2013) documented distinctive feature of marine fungi Trichoderma viride Pers NFCCI-2745, which was isolated from a phenolic-polluted estuary, for production of salinity- and phenol-tolerant laccase enzyme. Biotransformation of persistent organic contaminants like PCB 118 by marine fungi Penicillium sp. and Trichoderma harzianum can be affected by biostimulation and bioaugmentation process (Verma et al. 2012; Gao et al. 2013; Vacondio et al. 2015). Several other marine-originated fungi like Aspergillus, Penicillium, and Mucor and slime mold also showed effective bioremediation properties in removal of crude oil; however, higher concentration of crude oil was observed to be toxic for fungi (Hickey 2013).

2.3.3  Extremophilic Fungi Microbes colonizing extreme environments are very crucial for various industrial applications as they produce extremophilic enzymes which depict several unique properties such as tolerance against temperature, pH and other extreme environmental conditions (Yadav 2018, 2019; Yadav et al. 2016, 2017b; Neifar et al. 2015). Fungi showed tolerance against high level of toxicants present in the effluent treatment plant and utilized for numerous bioremediation applications. These properties make them potent candidates for economical, eco-friendly bioconversions of raw materials into non-toxic forms used in food industries, leather processing, textiles manufacture, and animal feed preparation (Nigam 2013). Microbes have been employed for the removal of heavy metal contaminants from environment. Biosynthesis of nanoparticles provides a safer pathway for metal removal (Sinha et al. 2014). A psychrophilic fungus named Cryptococcus sp. isolated from deep-­ sea sediments showed tolerance and growth in presence of elevated levels of heavy metals concentration up to 100  mg/L ZnSO4, CuSO4, Pb(CH3COO)2, and CdCl2 (Singh et al. 2013b). Many hydrolytic enzymes were reported which showed activity under extremophilic conditions such as high salinity and crude oil concentration which is a result of drilling waste from oil belts (Yadav et al. 2017a, 2018). Laccases were observed to be responsible for bioremediation activity in Pestalotiopsis palmarum in the presence of wheat bran and production of lignin peroxidases occurred when heavy crude oil was present in high concentration and it was utilized by fungi as the sole carbon and energy source (Betancor et al. 2013; Naranjo-Briceno et al. 2013). On the other hand, enzymes like chitinases synthesized by a psychrophilic fungus named

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Lecanicillium muscarium was employed for increasing the activity of insecticides which showed capability to act on insect chitin exoskeleton (Li et  al. 2013; Narayanan et  al. 2013). Not only literature about the bioremediation studies by extremophilic fungi is important but their isolation from extreme habitats such as a deep biosphere represented by fumarolic ice caves on Antartica’s Mt. Erebus can also be used for identifying unique properties of fungi capable of utilizing energy sources other than photosynthesis and also provides information about possible anthropogenic sources of contamination of such extreme places (Connel and Staudigel 2013).

2.3.4  Symbiotic Fungi with Plants Fungi are known to build mutual and close association with plants in order to rise above the barrier of limited growth under varied environmental conditions. A symbiotic relationship between plants and fungi is represented by arbuscular mycorrhizal fungi (AMF) (Yadav et al. 2019a). In this association, fungus accelerates the degradation of pollutants by providing higher surface area for absorption through its hyphae and spores and thereby converts the pollutants into mobile form and binds to the root. Fester (2013) observed AMF colonization in the root sample of plants which are utilized for phytoremediation of polluted groundwater in a constructed wetland. Numerous arbuscular mycorrhizal fungi such as Aspergillus nidulans, Bjerkandera adusta, Trametes hirsuta, T. viride, Funalia trogii, Irpex lacteus, and P. ostreatus could thrive in the toxic environment of dyes and were employed for decolorization of textile industry effluents (Tegli et  al. 2013). Arbuscular mycorrhizal fungus named Rhizophagus custos showed elevated level of tolerance against anthracene, a polyaromatic hydrocarbon, and converts it to the less toxic anthraquinone by-product (Aranda et al. 2013). Quinoa plants showed enhanced uptake of 137 Cesium on loamy soil after being inoculated with a commercial AM product which also represents mycorrhizal association due to root colonization (Vinichuk et al. 2013). Ectomycorrhizal fungi such as Suillus bovinus and Rhizopogon roseolus representing association with Pinus have been reported for the removal of calcium from the surroundings. Environmental factors such as pH, type of nutrient and temperature also play a crucial role in the decontamination process (Sousa et  al. 2014). Other uses of such fungi have been targeted at overcoming technical barriers of algal bio-fuels and photosynthetic biorefineries by cocultivation of microalgae and fungi for the complete removal of single algal cells from fermentation medium. This allowed their extraction and harvesting in a simple manner by simple filtration with result of increased biomass, lipids, and bio-product yields (Xie et al. 2013). There are several advantages of coculture studies for bioremediation, but their applications are complex and require immense understanding about the interaction between diversified metabolic pathways from different organisms.

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2.4  Fungal Enzymes in Bioremediation Common enzymes of fungal origin used in bioremediation include amylases, catalases, cellulases, laccases, lipases, peroxidases, proteases, xylanases, etc. which have potential industrial applications and are used in managing organic waste (Marco et al. 2013). These fungal enzymes may be utilized for hydrolysis of polymeric materials, for example, cellulose, protein, lipid, starch, and xylan from organic wastes such as kitchen waste, vegetable market waste, leaf, litter, etc. The hydrolyzed substances can be further used for composting or for manufacture of significant products such as volatile fatty acids and biogas (Khardenavis et al. 2013; Hattori et al. 2015). The group of white-rot fungi synthesizes various types of ligninolytic enzymes which rely on the fungal species and environmental conditions, which cannot only be used for lignin degradation of ligno-cellulosic materials but also in the remediation of different recalcitrant pollutants including dyes. These enzymes alter azo dye configuration by damaging the chromophoric linkages which lead to the synthesis of phenoxyl radicals in the reactions (Gül and Dönmez 2013). Ligninolytic enzymes produced by the group of white-rot fungi have been divided into two groups: peroxidases (lignin peroxidases and manganese peroxidases) and laccases (Jebapriya and Gnanadoss 2013; He et al. 2015). Several enzymes secreted by white-rot basidiomycetes such as laccases and some peroxidases are very well known for the degradation of persistent organic xenobiotics (Ikehata 2015). Such enzymes are also secreted by extremophilic fungi which can be used for remediation of pollutants under extreme conditions like high salinity, extra-heavy crude oil contamination, etc. Currently, much attention has been given on the modification of enzymes from white-rot fungi through recombinant gene expression techniques and protein engineering process, which can be used effectively and eco-friendly for remediation of toxic pollutants (Fonseca et al. 2013; Sakaki et al. 2013; Syed et al. 2013; Wong et al. 2013). An overview of common fungal enzymes used for bioremediation of different pollutants is given in Table 2.1.

2.4.1  Enzymological Background Enzymes are biological moieties which act as a catalyst and modify a reaction’s rate by providing suitable conditions which reduce the activation energy of the reaction, without being used with substrates of the reaction. Enzymes are formed naturally by almost every known organism to help in the processes such as cell synthesis, digestion, and metabolism (Madigan et al. 2003). An enzyme might comprise a protein or a glycoprotein and a polypeptide moiety. The segments of an enzyme which are directly involved with the catalytic process are known as active sites (Fig. 2.3). An enzyme may have other groups associated with the active sites through either covalent or noncovalent bonds which are crucial for catalytic activity. The protein

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Table 2.1  An overview of the potential of common fungal enzymes for bioremediation SN 1.

Enzyme Lignin Peroxidase (LiP)

Compound Pyrene

Fungi Phanerochaete chrysosporium

References Hammel (1997)

Aminodinitrotoluenes

P. chrysosporium

Bentazon (herbicide)

P. chrysosporium

Trichlorophenol

Cotiolos versicolor, Panus tigrinus Pleurotus pulmonarius

Cameron et al. (2000) Castillo et al. (2000) Leontievsky et al. (2000) Lau et al. (2003)

Poly aromatic hydrocarbon (PAH) Delor 106 [Polychlorinated biphenyl (PCB)] Phencyclidine (PCP)

2.

Manganese Peroxidase (MnP)

Trametes versicolor

Remazol Brilliant Blue R

T. versicolor, Trametes sp. Irpex lacteus

PAH

P. chrysosporium

Trinitrotoluene (TNT)

TNT

Nematolloma frowardii Stropharia rugosoannulata C. versicolor, P. tigrinus P. chrysosporium

PAH Delor 106 (PCB)

P. pulmonarius T. versicolor

Remazol Brilliant Blue R

I. lacteus

PCP

T. versicolor, Trametes sp. Bjercandera adusta

Trichlorophenol

PAH

P. pulmonarius Di(2-ethylhexyl) phthalate, heavy metals, total petroleum hydrocarbons Bentazon P. chrysosporium PAH

P. chrysosporium

Reactive black 5, Veratryl alcohol

Ceriporiopsis subvermispora

Novotny et al. (2004) Ford et al. (2007) Novotny et al. (2004) Wang et al. (2009) Scheibner and Hofrichler (1998) Leontievsky et al. (2000) Cameron et al. (2000) Lau et al. (2003) Novotny et al. (2004) Novotny et al. (2004) Ford et al. (2007) Valentin et al. (2007) Chiu et al. (2009) Castillo et al. (2000) Wang et al. (2009) Fernández-­ Fueyo et al. (2014) (continued)

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Table 2.1 (continued) SN 3.

Enzyme Laccase

Compound 2,6 dimethoxy phenol

Fungi Pleurotus ostriatus

Phenolic compounds

C. versicolor

Trichlorophenol

C. versicolor, P. tigrinus C. versicolor

2,4-dichlorophenol, pentachlorophenol Bromophenol blue, Malachite green, Orange G, Amalanth TNT

Pycnoporus sanguineus

Trametes villosa

PAH P. pulmonarius Anthracene and benzo pyrene T. versicolor 4- hydroxybiphenyl Delor 106 (PCB)

T. versicolor, Pleurotus ostriatus T. versicolor

Remazol brilliant blue R

I. lacteus

PCP

T. versicolor, Trametes sp. T. versicolor

Atrazine (herbicide)

P. pulmonarius Di(2-ethylhexyl) phthalate, heavy metals, total petroleum hydrocarbons Flexographic inks T. villosa, Coriolopsis rigida, Pycnoporus coccineus Myceliopthora thermophila Pesticides P. chrysosporium, T. versicolor, Pleurotus sp., B. adjusta Phenols T. versicolor PCB

Doratomyces nanus, D. purpureofuscus, D. verrucisporus, Myceliophthora thermophila, Phoma eupyrena, Thermoascus crustaceus

References Hublik and Schinner (2000) Davis and Burns (1992) Leontievsky et al. (2000) Ullah et al. (2000) Mayer and Staples (2002)

Wang et al. (2002) Lau et al. (2003) Dodor et al. (2004) Keum and Li (2004) Novotny et al. (2004) Novotny et al. (2004) Ford et al. (2007) Bastos and Magan (2009) Chiu et al. (2009) Fillat et al. (2012)

dos Santos Bazanella et al. (2013) Margot et al. (2013) Mouhamadou et al. (2013)

(continued)

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Table 2.1 (continued) SN

Enzyme

Compound Phenylurea herbicide diuron

Fungi Mortierella

Gasoline

Exophiala xenobiotica

Anthracene

Armillaria sp.

Naphthalene

Pleurotus eryngii

Organic pollutants

White-rot basidiomycetes Aspergillus sp., Curvularia sp., Drechslera sp., Fusarium sp., Lasiodiplodia sp., Mucor sp., Rhizopus sp., Tricoderma sp. Aspergillus sp., Curvularia sp., Acrimonium sp., Pythyme sp. A. flavus

4.

Lipase

PAHs

5.

Catalase

Heavy metals

6.

Peroxidase

Malondialdehyde

A. foetidus

Heavy metals

Penicillium sp. Rhizopus sp. A. niger, A. foetidus, T. viride, A. sojae, Geotrichum candidium, Penicillium sp., Pycnoporus cinnabarinus Trichoderma sp. White-rot fungi B. adusta, Ceriporia metamorphosa, Ganoderma sp. P. chrysosporium, T. versicolor, Pleurotus sp., B. adjusta White-rot basidiomycetes

Textile dye decolorization

Dye decolorization

Pesticides

Organic pollutants

References Ellegaard-Jensen et al. (2013) Isola et al. (2013) Hadibarata et al. (2013) Hadibarata et al. (2013) Ikehata (2015) Lladó et al. (2013) Chang et al. (2015) Balaji et al. (2014)

Akhtar et al. (2013)

Kurniati et al. (2014) Chakraborty et al. 2013 Deshmukh et al. (2016) Jebapriya and Gnanadoss (2013)

Ma et al. (2014)

dos Santos Bazanella et al. (2013) Ikehata (2015)

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Complex of enzyme & substrate

Substrate Active sites Reaction products

Enzyme

Enzyme

Fig. 2.3  A schematic diagram showing mechanism of enzymatic reactions

or glycoprotein part in an enzyme is known as the apoenzyme, and the nonprotein moiety is called the prosthetic group. The combination of both apoenzyme and prosthetic group forms the holoenzyme. Enzyme names are given according to the role of the enzyme and type of the reaction catalyzed (Lehninger et al. 2004). The identification of a particular enzyme is possible through its enzyme commission (E.C.) number. The assignment of E.C. numbers is described in guidelines set out by the International Union of Biochemistry. All known enzymes fall into one of these six categories. The six main divisions are (1) the oxidoreductases, (2) the transferases, (3) the hydrolases, (4) the lyases, (5) the isomerases, and (6) the ligases (synthetases). Oxidoreductases catalyze the transfer of electrons and protons from a donor to an acceptor. Transferases catalyze the transfer of a functional group from a donor to an acceptor. Hydrolases facilitate the cleavage of C–C, C–O, C–N and other bonds by water. Lyases catalyze the cleavage of these same bonds by elimination, leaving double bonds (or, in the reverse mode, catalyze the addition of groups across double bonds). Isomerases facilitate geometric or structural rearrangements or isomerizations. Finally, ligases catalyze the joining of two molecules (Lehninger et al. 2004).

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2.4.1.1  Oxidoreductases The oxidoreductase enzyme facilitates detoxification of organic pollutants by different fungi via oxidative coupling mechanism (Gianfreda et  al. 1999). Fungal enzymes cleave the chemical bonds of substrates through biochemical reactions releasing energy and assist the transfer of electrons from a reduced organic substrate to another chemical compound which acts as an acceptor. The contaminants are finally oxidized to non-toxic compounds during the process of oxidation coupled with reduction. Oxidoreductases through the process of detoxification convert toxic xenobiotics into harmless compounds. Xenobiotics like phenolic or anilinic compounds get transformed into non-toxic form through the process of polymerization, copolymerization, or binding to humic entities (Park et al. 2006). Oxygenases represent the oxidoreductase group of enzymes. Organic compounds mainly of halogenated nature like herbicides, insecticides, fungicides, plasticizers and intermediates used for the synthesis of chemicals consist the largest groups of environmental pollutants ubiquitous in nature. These pollutants are degraded by specific oxygenases. The dehalogenation reactions of halogenated methanes, ethanes, and ethylenes are mediated by oxygenases in combination with multifunctional enzymes (Fetzner and Lingens 1994). Chlorinated phenolic compounds are considered to be the most abundant recalcitrant wastes found in the effluents generated by the paper and pulp industry. It generates mainly recalcitrant wastes in their effluent. The partial degradation of lignin during pulp bleaching process results in the formation of these recalcitrant compounds. Fungal populations are considered to be appropriate candidate for the decontamination of sites contaminated with chlorinated phenolic compounds. The production of extracellular oxidoreductase enzymes like laccase, manganese peroxidase and lignin peroxidase by the action of fungi is useful for bioremediation. Filamentous fungi penetrate the soil pollutants more efficiently than bacteria (Rubilar et al. 2008). 2.4.1.2  Laccases Laccases represent a group of copper-based enzymes synthesized by fungi. They utilize copper as a cofactor and molecular oxygen as cosubstrate which further gets reduced to water during oxidation process of xenobiotics (Gianfreda et  al. 1999; Mai et al. 2000). Laccases are known to occur in multiple. Various isoenzyme forms of laccases have been reported. Each enzyme is coded by an individual gene. The expression of gene depends on the nature of inducer (Giardina et al. 1995; Rezende et al. 2005). Most of the phenolic, non-phenolic, and aromatic compounds are oxidized by laccases and an increase in their activity about 20 times was observed in the fungi like T. versicolor (Margot et al. 2013). Both intracellular and extracellular laccases were produced by many genera of fungi which catalyze the oxidation of ortho and paradiphenols, aminophenols, polyphenols, polyamines, lignins, aryl

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diamines, and some inorganic ions (Mai et al. 2000; Ullah et al. 2000; Rodríguez Couto and Toca Herrera 2006). In spite of oxidation of phenolic and methoxyphenolic acids, laccase also performs the function of decarboxylation and demethylation. These enzymes participate in the depolymerization of many aromatic compounds. This enables them to be potent candidates to function as a biocatalyst in the production of numerous dyes and colorants, thereby reducing the cost and energy expenditure in their synthesis with no residual material (Polak and Jarosz-­ Wilkolazka 2012). These compounds are also utilized by microbes as a nutrient source or repolymerized to humic materials by laccase (Kim et al. 2002). Laccases signify an appealing group of ubiquitous, oxidoreductase enzymes among biological agents contributing great prospective in biotechnological and bioremediation applications (Gianfreda et al. 1999). Their non-specific property for varying range of substrates ultimately makes them ideal catalyst for numerous industrial applications. Enzymes have been explored to be used extensively for their efficient bioremediation potential (Vishwanath et  al. 2014). Fillat et  al. (2012) demonstrated deinking process in printed paper on recycling by the enzyme laccases produced by the three fungi belonging to group basidiomycetes (Trametes villosa, Coriolopsis rigida, Pycnoporus coccineus) and other to the ascomycetes (Myceliopthora thermophila) in the presence of synthetic mediators. Due to the synthetic origin of the textile dyes, they remain unaltered in the environment on exposure to sunlight, water, and other external factors and thus subsequently impart toxicity to the ecosystem. Fungal marine laccases were used for the color removal, detoxification, and mineralization of Reactive Blue 4 dye in the studies of Verma et al. (2012) and Vishwanath et al. (2014) for the first time. An endocrine disrupting chemical named Bisphenol A is degraded by laccase isolated from Fusarium incarnatum as reported by Chhaya and Gupte (2013). Laccases were found to be produced at low temperature in Himalayan region by some extremophilic fungi like Penicillium pinophilum (Dhakar et  al. 2014). Less literature is available about the mechanism of laccase under extreme conditions. However, only few laccases from fungus belonging to the group ascomycetes like Melanocarpus albomyces and Thielavia arenaria were explored with crystal nature. They differ from other fungus by having conserved ‘C-terminal plug’ most likely used in proton transfer processes (Kallio et al. 2011). Changes in pH can be directly correlated with the substrate specificity and enzyme affinity. An inhibition in the production of laccase was observed by the presence of various chemicals like halides, azides, cyanides, and hydroxide (Xu 1996). The synthesis of laccase in fungi is sensitive to the concentration of nitrogen. High levels of nitrogen are directly proportional to the production of recombinant. It can be produced by either homologous or heterologous means (Gianfreda et al. 1999). Laccases show remarkable potential in bioremediation, but their utilization is limited due to their low shelf-life. Immobilization or tailoring of enzymes is considered as an appropriate method in order to overcome this problem and enhancing their efficiency (Mate et al. 2011; Torres-Salas et al. 2013; Patel et al. 2014).

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2.4.1.3  Catalase The accumulation of reactive oxygen species (ROS) in biological systems results in the damage at cellular level. These species cause deleterious effects to cellular macromolecules. Fungi produce monofunctional catalases and bifunctional peroxidase/ catalase enzymes for defense mechanism at primary level in order to cope up with the toxicity of reactive oxygen species. Lindane inhibits the production of catalase and thus accelerates the generation of reactive oxygen species which ultimately declines the growth of Saccharomyces cerevisiae (Pita et al. 2013). The production of reactive oxygen species in microbes has been induced by the presence of several heavy metals like lead, copper, cadmium, zinc, etc. Several studies have shown that presence of heavy metals on the generation of reactive oxygen species is directly correlated, and it leads to the production of anti-oxidative enzymes. Studies of Chakraborty et  al. (2013) demonstrated that fungus Aspergillus foetidus showed tolerance against 200  mg/L concentration of Pb (II) by the production of anti-­ oxidative enzymes including laccase for the detoxification of malondialdehyde and hydrogen peroxide. A study conducted by Mitra et al. (2014) showed that an increase in tolerance level of Aspergillus sp. against oxidative stress induced by the presence of varying concentrations of heavy metals such as 100 mg/L Cu(II) and 750 mg/L Zn(II) is achieved by the production of catalase. Complete literature about the effect of heavy metals on fungal physiology is not yet known. An inhibition in catalase and peroxidase activity and increase in cytochrome P450 (CYP450) activity were observed in the case of P. chrysosporium when exposed to 50–100 μM concentration of cadmium or lead (Zhang et al. 2015). An increase in the catalase activity was observed in the case of fungal consortia A. niger, Penicillium sp., and Rhizopus sp. when exposed to heavy metals like lead and copper at 50 mg/L (Thippeswamy et al. 2014). In a study by Lin et  al. (2009), it was demonstrated that catalase activity could be employed effectively for monitoring bioremediation potential of fungus, and it decreases with increase in the oil concentration in sites contaminated with oil residues. Catalase proved to be an effective tool for providing tolerance capacity to fungi for decontamination of contaminated sites. Thus fungi producing enzymes play a significant role in the bioremediation of contaminated sites with metals. 2.4.1.4  Peroxidases Peroxidases catalyze the oxidation of lignin and other phenolic compounds at the expense of hydrogen peroxide in the presence of a mediator. These peroxidases belong to proteins of heme and nonheme nature. The heme peroxidases are divided into two broad groups. First group is found in animals, plants, fungi, and prokaryotes while the second group peroxidases have been subdivided into three distinct classes on the basis of sequence comparison. Class I represents enzymes of intracellular nature including yeast cytochrome c peroxidase, ascorbate peroxidase from plant origin, and gene-duplicated catalase peroxidases from bacteria. Class II represents fungal-secreted heme peroxidases from P. chrysosporium like lignin

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peroxidases and manganese peroxidases. Lignin peroxidases and manganese peroxidases were reported for the utilization of hydrogen peroxidase and manganese in carrying out their vital activities. These enzymes were reported for the degradation of recalcitrant toxic compounds by the group of white-rot and basidiomycetes fungi. Versatile peroxidase enzymes are specific for a broad range of substrates. They oxidize both phenolic and non-phenolic compounds and thus they are highly efficient for bioremediation (Karigar and Rao 2011). Degradation of lignin in wood was reported by the class II peroxidases. Class III represents the secretory plant peroxidases such as those from horseradish, barley, or soybean. These peroxidases behave as a biosynthetic enzyme involved in the plant cell wall formation and lignification processes (Hiner et al. 2002; Koua et al. 2009). Non-heme peroxidases are not evolutionarily linked and form five independent families. They include thiol peroxidases, alkyl hydroperoxidases, non-heme haloperoxidase, manganese catalase, and NADH peroxidases. Thiol peroxidases represent the largest family when compared with others. They also include two subfamilies such as glutathione peroxidases and peroxy redoxins (Koua et al. 2009). Another group of heme peroxidases were reported as dye-decolorizing peroxidases (DyPs) and unspecific peroxygenases (UPO) which utilize hydrogen peroxide for catalyzing oxidations of varying non-phenolic lignin compounds and other organic compounds. This group does not fit in the above classification system (Liers et al. 2013; Strittmatter et al. 2013; Hofrichter and Ullrich 2014). One of such peroxidases produced by B. adusta was declared for the disruption of the phthalocyanine ring in phthalocyanine dyes. This has been achieved by the cleavage of azo bond and ultimately leading to the decolorization of azo and phthalocyanine dyes. Five fungal dye-decolorizing peroxidases of high redox nature were demonstrated by Liers et al. (2013) which possess catalytic properties of both lignin peroxidases and versatile peroxidases as confirmed from their capability to oxidize non-phenolic aromatic compounds and Reactive Black B. The study highlighted the need to further distinguish peroxidase activities in crude enzyme mixtures of fungi posing difficulty in classifying the dye-decolorizing peroxidases from lignin peroxidases and versatile peroxidases. Such type of classification based on catalytic specificity was suggested only after the refinement of the different enzymes. 2.4.1.4.1  Lignin Peroxidases They belong to the group of heme proteins secreted mainly by the members of white-rot fungi during their secondary metabolism. These enzymes degrade lignin and other phenolic compounds in the presence of cosubstrate H2O2 and veratryl alcohol which acts as a mediator. During the course of reaction, H2O2 gains electrons from lignin peroxidases and thus reduced to H2O. The oxidized form of lignin peroxidases gained electron from veratryl alcohol and thereafter returns to its original reduced state with the formation of veratryl aldehyde. It gets reduced back to veratryl alcohol by gaining an electron from substrate. This ultimately results in the oxidation of halogenated phenolic compounds, polycyclic aromatic compounds and

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other aromatic compounds followed by a series of nonenzymatic reactions (Yoshida 1998; Ten Have and Teunissen 2001). Lignin peroxidases play a crucial role in the biodegradation of lignin which is a constituent of the plant cell. It also oxidizes aromatic compounds with redox potentials more than 1.4 V (NHE) by the single-­ electron abstraction mechanism. The exact redox mechanism is still not properly understood (Piontek et al. 2001). 2.4.1.4.2  Manganese Peroxidases They are an heme group of enzymes produced extracellularly from the basidiomycetes group of fungus responsible for the degradation of lignin which oxidizes Mn2+ to Mn3+ in a multistep manner. Mn2+ initiates the production of manganese peroxidases and also acts as a substrate for it. Manganese peroxidases generate Mn3+ which plays an important role in the oxidation of varying phenolic compounds by acting as a mediator (Ten Have and Teunissen 2001). Recently, Ceriporiopsis subvermispora was used for the production of enzyme manganese peroxidases which were modified through genetic manipulation to increase the stability even at acidic pH 2. The acid-stable nature of Mn2+ and its high oxidizing activity were confirmed by analyzing its crystal structure as a scaffold. An engineered enzyme of stable nature was found for oxidizing Reactive Black 5 and veratryl alcohol (Fernández-­ Fueyo et al. 2014). 2.4.1.4.3  Versatile Peroxidases Versatile peroxidase enzymes are found capable of oxidizing Mn2+, methoxybenzenes, and phenolic aromatic substrates. Versatile peroxidases are specific for a broad range of substrates. They oxidize substrates in the absence of manganese when compared with other group of peroxidases. Both phenolic and non-phenolic lignin model dimmers were found to be oxidized by VP (Ruiz-Dueñas et al. 2007). A highly well-organized versatile peroxidase production system in excess was formulated for various biotechnological applications in industries and removal of xenobiotic compounds of recalcitrant nature from the environment (Mai et al. 2000; Tsukihara et al. 2006).

2.4.2  Advantages of Enzymatic Bioremediation Enzymes are adapted to act in a varied range of environmental conditions without alteration in their active nature (Ahuja et al. 2004; Gianfreda and Rao 2004). An example depicts that protease enzyme is capable of functioning in the pH range 4–11 but the experiment retards at pH 11. The enzyme was found still to be active at this moment with temperature lesser than 20 °C and greater than 70 °C

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(Whiteley et al. 2002). Enzymes are found to work effectively in diverse environments especially when they present in immobilized form. This property makes the enzymes more opposed to hard environmental conditions and can be recycled and recovered when they are not needed (Gianfreda and Rao 2004). Several workers found that xenobiotics like phenol, PCBs, herbicides and dyes were degraded by the laccase (Ullah et al. 2000; Dodor et al. 2004; Novotný et al. 1997; Mougin et al. 2000; Mayer and Staples 2002). Enzymatic degradation of several xenobiotic compounds was found to be a feasible method, but at the same time, the rate of reaction for each compound should be minimized for a system (Alvarez-Cohen and Speitel Jr 2001). Enzymatic treatment of contaminated site is an eco-friendly process over conventional methods. These procedures are relatively time-consuming in comparison to conventional methods without imparting deleterious effect on the environment. Enzymes are proteins with degradable nature, i.e., when they are not recovered, they are degraded in the nature without leaving any harmful residues. So, enzymatic processes are found to be an attractive alternative for the decontamination of contaminated sites.

2.4.3  Disadvantages of Enzymes Although enzymatic technology is very promising, it has limitations. Microbes can reproduce and increase their population in order to consume a large amount of substrate, but extracellular enzymes cannot. Enzymes cannot reproduce themselves, meaning that any increase in enzyme population must come from outside of the system (i.e., humans adding more enzymes to the system). It has been noticed that enzymes lose their reactive property after their interaction with the pollutants and thus get converted into inactive entities. (Gianfreda and Rao 2004). The concentration of enzyme is dependent on the reaction rate so their concentrations must be checked and controlled properly so as to optimize enzyme kinetics for site-specific situations. Enzymes do not possess the reproductive nature so they are not able to adapt themselves as microbes during mutation. Mutations allow microorganisms to withstand harsh conditions and able to metabolize new substrates (Madigan et al. 2003). Enzymes can withstand varying range of environmental conditions only within their limits of adaptation. The main drawback of utilizing extracellular enzymes for bioremediation is their high costs (Duran and Esposito 2000). The cost of producing enzymes employed for bioremediation in pure state is relatively high as purity directly correlates with their specific functions and showing no adverse effects. The cost of synthesis of enzyme solutions in crude form is comparatively cheaper, but it tends to show side effects (Fullbrook 1996). The reduction of production expenditure can be achieved by the selection of cheaper substrates for the growth of microbes in the production of enzymes. The focus of advancement in technology is in the process for the replication of bacteria and fungi used for the synthesis of enzymes (Ahuja et al. 2004; Gianfreda and Rao 2004).

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2.4.4  Scope of Enzymatic Bioremediation For bioremediation of various pollutants, fungi have now been proved to play a noteworthy role. Variety of toxic pollutants, for instance, dyes, pesticides, pulp and paper industry effluents, persistent organic pollutants (POPs), PAHs, petroleum hydrocarbons, leather tanning effluents, etc., can be degraded by using different fungal enzymes. Various fungi like Acrimonium, Aspergillus, and Curvularia were reported for metal tolerance and metal removal ability (Akhtar et  al. 2013). Few basidiomycetes such as T. versicolor and white-rot fungi P. ostreatus have been studied for degradation of PAHs in solid-state fermentation during growth phase on agro-industrial wastes (Rosales et al. 2013). Pollutant from coffee industries like coffee pulp can also be remediated by using fungi such as Aspergillus restrictus, Chrysosporium keratinophilum, Fusarium solani, Gliocladium roseum, Penicillium and Stemphylium under controlled environmental conditions with additional nutrients input (Nayak et  al. 2013). A variety of fungi of different groups including white-rot fungi, Aspergillus, Penicillium, etc. have been reported to degrade and decolorize diversified pollutants from various industries like sugar mill, textile dye, paper and pulp industry, and leather tanning industry, which indicate that fungi can be used for a range of substrates (Bennett et  al. 2013; Buvaneswari et  al. 2013; Duarte et al. 2013; Jebapriya and Gnanadoss 2013; Reya et al. 2013; Huang et al. 2014). Bioremediation of PAHs in presence of A. niger and P. chrysosporium revealed considerable removal of petroleum hydrocarbons from petrol- and diesel-­ polluted soil (Maruthi et al. 2013). Some studies showed the complete remediation of pesticides such as chlorpyrifos and also its metabolite 3,5,6-trichloro-2-pyridinol (TCP) from contaminated soils by using fungi A. niger JAS1 without any supplementary nutrient input (Silambarasan and Abraham 2013). The degradation of metabolite TCP by chlorpyrifos-degrading fungal strain was a considerable finding in view of the antimicrobial property and catabolite repression character showed by TCP.

2.5  Conclusion and Future Prospects Though there have been several reports on the bioremediation potential of fungal species, profound evaluation of the versatile function of fungi in remediation of xenobiotic compounds suggesting the descriptive mechanism of fungi for attaining this task is missing. In the present chapter, we have tried to elaborate the role of fungal enzymes in bioremediation of synthetic compounds with special prominence to organic pollutant. Besides enzymes like peroxidases and laccases, some stress response proteins like ABC transporters also play an active role in the removal of toxic contaminants in fungi, and there is a demand for exploring these genes for further applications. Thus, a necessity arises to search new techniques such as genetically modified microorganisms or making consortia by combining plants,

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bacteria and fungi for providing appealing opportunities in bioremediation technique. A continuous investigation for the new biological form is required for proper management of increasing pollution and contamination. Therefore, bioremediation is still considered as a developing technology to regulate the day-to-day environmental problems faced by humans residing in an area. Acknowledgments  The author Sonal Dixit acknowledges DSKPDF Cell, Pune, India, and University Grant Commission, New Delhi, India, for the financial assistance in the form of D.S. Kothari Postdoctoral Fellowship (F4-2/2006 (BSR)/BL/15-16/0156). There are no conflicts of interest among authors.

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Chapter 3

Genetic Diversity of Methylotrophic Yeast and Their Impact on Environments Manish Kumar, Raghvendra Saxena, Pankaj Kumar Rai, Rajesh Singh Tomar, Neelam Yadav, Kusam Lata Rana, Divjot Kour, and Ajar Nath Yadav

3.1  Introduction In the environment, abundant reduced C1-compounds are available. Reduced carbon compounds, such as methane and methanol, utilised by methylotrophs, which have the ability to utilise C1-compounds as the sole source of carbon and energy, also appear to be cosmopolitan in nature. There is a remarkable difference between prokaryotic methylotrophs and eukaryotic methylotrophs. Pokaryotic methylotrophs utilize carbon substrate like methanol, methane and methylamine while methanol is used as carbon substrate and methylamine as nitrogen source by eukaryotic methylotrophs. Methylotrophic yeast comprises genera like Pichia, candida and some other related genera close to Pichia i.e. Kuraishia, Ogataea and Komagataella (Yurimoto et al. 2011). Different lineages of methylotrophic yeast utilising methanol as sole source of carbon and energy were documented and described (de Koning and Harder 1992). The similar metabolic pathway for methanol utilisation was followed by all methylotrophic yeasts, and they composed of several enzymes localised in peroxisomes which proliferate during growth of yeast in methanol (Veenhuis et al. 1992; Yurimoto

M. Kumar · R. Saxena · R. S. Tomar Amity Institute of Biotechnology, Amity University, Gwalior, India P. K. Rai Department of Biotechnology, Invertis University, Bareilly, Uttar Pradesh, India N. Yadav Gopi Nath P.G. College, Veer Bahadur Singh Purvanchal University, Deoli-Salamatpur, Ghazipur, Uttar Pradesh, India K. L. Rana · D. Kour · A. N. Yadav (*) Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, Sirmour, Himachal Pradesh, India e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_3

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et  al. 2002). The methylotrophic yeast with regulated promoters of methanol oxidising genes is reported to be used in the study of production of recombinant proteins as well as industrial proteins (Ravin et al. 2013). The diversity of methylotrophic yeast involved in glycerol metabolism was also discussed in one of the earlier reports in which methylotrophic strains such as Candida boidinii no. 2201, Hansenula ofunaensis and Hansenula polymorpha DL-1 were identified with specific enzyme activity of glycerol kinase (GK), glycerol dehydrogenase (GDH) and both GK and GDH, respectively (Tani and Yamada 1987). Methanol is considered as a very recent alternative carbon source that replaces the petroleum and coal (Olah 2005). It is considered that methanol is formed with the combination of CO and H2 or from CO2 with the help of H2 gas. Methane, an abundant natural carbon substrate produces CO and H2. Since methanol is a cheaper substrate, it can be used as feedstock for different biochemical and biotechnological processes. Methane oxidizing bacterial communities are responsible for release of methanol from methane and further triggers the decomposition of lignins and pectins containg methylester and methoxyl groups respectively (Mitsui et al. 2003; Nakagawa et al. 2000). Methylotrophs and methylotrophic yeasts oxodise CO2. Thus, methylotrophs play indispensable roles in the global carbon cycle between methane and CO2 called “the methane cycle”. A thorough understanding of the molecular basis of methylotrophy is needed not only to better understand the global methane cycle but also to permit more efficient use of methanol as a renewable carbon source. In the recycling of carbon in the environment, methylotrophic yeast plays a very crucial role. The ability of C. Boidinii to grow over pectin as a substrate shows its methylotrophic metabolism (Nakagawa et  al. 2000). The intensive researches explain the beneficial relationship between plants and methylotrophic microbial communities (Kumar et al. 2019; Meena et al. 2012; Verma et al. 2013, 2014, 2015a, b, 2016a, b; Yadav 2009). Moreover, the interaction between plants and methylotrophic yeasts has not been characterised and very less report or documentation is available (Limtong et al. 2005; van der Klei et al. 2006). Therefore, keeping in view the importance of the methylotrophic yeast in the agriculture, industry and environments, the present chapter deals with the impact of methylotrophic yeast in environments and describes recent insights into the molecular basis of yeast methylotrophy.

3.2  Genetic Diversity of Methylotrophic Yeast Yeasts are well known for their beneficial activity for humankind by their exploitation and application in the food, beverages and in the production of various types of biochemicals. Since they contain a significant content of vitamin B, amino acids and minerals, they can be used as a food supplement also. Moreover, methylotrophic yeast can be used in the gene regulation study in eukaryotes and as biofactories for heterologous and homologous proteins (Cremata and Díaz 1999; Negruţă et  al. 2010) (Table  3.1). This group of yeast is able to survive by metabolising

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Table 3.1  Details of various strains of methylotrophic yeast and their genetic diversity Methylotrophic yeast Candida parapolymorpha Candida rishirensis

Gene for identification/ functionality/phylogeny D1/D2 region of LSU rRNA gene D1/D2 domain of LSU rRNA gene Formate dehydrogenase

Isolation source YM agar, 2% malt agar, Soil of Rishiri island Hansenula Methanol-grown polymorpha yeast cells Hansenula Dihydroxyacetone synthase Methanol-grown polymorpha yeast cells Hansenula Formaldehyde dehydrogenase Methanol-grown polymorpha yeast cells Hansenula Whole genome sequence Methanol- and polymorpha DL1 analysis Glucose-grown yeast cells Hansenula HpELO1, a fatty acid elongase – polymorpha gene Kluyveromyces D1/D2 domain of LSU rRNA – gene Culture media Komagataella phaffii Methanol-inducible gene expression with methanol Komagataella phaffii KpMit1 transcription factor Culture media gene expression with methanol Meyerozyma candida D1/D2 domain of LSU rRNA – gene YM agar, 2% Ogataea angusta SSU rRNA gene, internal malt agar, transcribed spacers (ITS) cornmeal agar, including 5.8S rRNA gene, and the D1/D2 region of LSU and V8 juice agar rRNA gene Ogataea D1/D2 domains of the Soil and tree chonburiensis large-subunit rDNA sequence exudates Ogataea D1/D2 large-subunit ribosomal From tree falcaomoraisii DNA exudates Ogataea glucozyma, SSU and LLU rRNA – Ogataea haglerorum D1/D2 LSU rRNA gene, sp. ITS1-5.8S-ITS2, and translation elongation factor-1α (EF-1 α) Ogataea henricii, SSU and LLU rRNA – Ogataea minuta SSU and LLU rRNA – Ogataea minuta, SSU and LLU rRNA – Ogataea D1/D2 domains of the Soil and tree nakhonphanomensis large-subunit rDNA sequence exudates Ogataea D1/D2 large-subunit ribosomal Leaf and rotten nitratoaversa DNA and ITS sequence wood

Reference Suh and Zhou (2010) Nakase et al. (2010) Ravin et al. (2013) Ravin et al. (2013) Ravin et al. (2013) Ravin et al. (2013)

Prasitchoke et al. (2007) Negruţă et al. (2010) Ohsawa et al. (2018) Leão-Helder et al. (2003) Negruţă et al. (2010) Suh and Zhou (2010)

Limtong et al. (2008) Morais et al. (2004) Naumov et al. (2018) Naumov et al. (2017)

Naumov et al. (2018) Naumov et al. (2018) Naumov et al. (2018) Limtong et al. (2008) Péter et al. (2008) (continued)

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Table 3.1 (continued) Methylotrophic yeast Ogataea philodendra, Ogataea polymorpha, Ogataea thermomethanolica TBRC656 Ogataea thermomethanolica TBRC656 Ogataea, Millerozyma Pichia pastoris Pichia pastoris Pichia pastoris Pichia pastoris Pichia pastoris Pichia pastoris Pichia pastoris Pichia pastoris Pichia sp. Pichia sp. N002 Pichia sp. N069 Pichia sp. PT31T Sacharomyces sp. Trichosporon cutaneum

Gene for identification/ functionality/phylogeny SSU and LLU rRNA

Isolation source Reference – Naumov et al. (2018)

SSU and LLU rRNA Gene encoding Hac1 transcription factor

– Grown in culture media

Naumov et al. (2018) Phithakrotchanakoon et al. (2018)

Novel gene expression based on maltase (mal) gene

–

Puseenam et al. (2018)

D1/D2 domain of LSU rRNA gene DAS (Dihydroxyacetone synthase) FLD1 (Formaldehyde dehydrogenase) THL1 (Thiamine biosynthesis gene) ADH1 (Alcohol dehydrogenase) AOX1 gene (Alcohol dehydrogenase) ICL1 (isocitrate lyase) Used as recombinant gene expression system AOX1 and AOX2 genes encoding alcohol oxidase D1/D2 domain of LSU rRNA gene D1/D2 domain of LSU rRNA gene D1/D2 domain of LSU rRNA gene D1/D2 domain of LSU rRNA gene D1/D2 domain of LSU rRNA gene Ribosomal DNA-based characterisation

–

Negruţă et al. (2010)

–

Ahmad et al. (2014)

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Ahmad et al. (2014)

–

Ahmad et al. (2014)

–

Ahmad et al. (2014)

–

Ahmad et al. (2014)

– Ahmad et al. (2014) Methanol-grown Young et al. (2012) yeast cells – Cereghino and Cregg (2000) – Negruţă et al. (2010) Soil

Limtong et al. (2005)

Soil

Limtong et al. (2005)

Soil

Limtong et al. (2005)

–

Negruţă et al. (2010)

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monocarbonic compounds such as formaldehyde and methanol (Kaszycki et  al. 2001). Methylotrophic yeasts are able to grow on extract of woods and other pectic material especially in fruits and vegetable products (Craveri et al. 1976). The woody materials are the source of methanol because of the presence of metoxi chain in

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Fig. 3.1  Illustration of methylotrophic yeast diversity based on genes involved in their metabolism and physiology

lignin. Figure 3.1 illustrates the diversity of methylotrophic yeast based on genes involved in their metabolism and physiology. Two methylotrophic yeast strains H. polymorpha and P. pastoris were utilised in the production of heterologous proteins. Apart from this, H. polymorpha is used for studying gene regulation of enzymes associated with abiotic stress tolerance, methanol metabolism, heavy metal resistance and nitrate assimilation. They are widely used in the methanol-contaminated waste water treatment also (Kaszycki and Koloczek 2002; Kaszycki et al. 2001). The methylotrophic yeast has the ability to grow in extreme environment also. In one of the investigations, three novel strains of thermotolerant methylotrophic yeast, which belong to genus Pichia, were reported. The methylotrophic strains were designated as N002, N069 and PT31T. The Pichia strains were isolated from soil (taken from Thailand) enriched with methanol. Thermotolerant yeast is found to grow at 20 °C (minimum temp.) but no limit for the maximum temperature for growth (Arthur and Watson 1976). According to this definition, methylotrophic yeast growing at 20 °C up to a temp. of 37 °C will be called as a thermotolerant methylotrophic yeast (Limtong et al. 2005). The presence of helmet-/hat-shaped ascospores, multilateral budding, ubiquinon

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Q-7 and negative for Diazonium blue B and urease reactions are the basic characteristics of genus Pichia. They do not have ballistospore and arthrospores also. The phylogenetic analysis based on D1/D2 domain of large subunit rDNA revealed the closeness with Pichia dorogensis. Because of the differences in phenotypic appearance, the above three strains were designated as novel species of Pichia and the name proposed was Pichia thermomethanolica sp. nov. The type strain is PT31T (Limtong et al. 2005). In a report, whole genome sequencing of a methylotrophic yeast H. polymorpha was performed and total transcripts were analysed from the yeast culture grown in methanol and glucose as well. A total of 9 Mb size of genome was sequenced for Hansenula polymorpha DL1. In a transcriptome analysis of H. polymorpha under methanol growth condition, 40% genome expression has shown the identified unregulated and abundant gene expression through RNA-seq analysis along with alternate splicing events. From seven chromosomes of H. polymorpha, different proteins of subtelomeric region were identified and the evolutionary relationship established revealed the closeness of H. polymorpha with both non-methylotrophic and methylotrophic yeast Dekkera bruxellensis and Pichia pastoris respectively.  In the investigation, phylogenetic analysis indicated the methylotrophic evolutionary pattern in filamentous fungi and yeasts (Ravin et al. 2013). Evolutionary analysis based on methanol-utilising pathway genes evaluated the relatedness of methylotrophic yeasts using NCBI nucleotide database and associated tools. Different genes involved in methanol utilisation pathway such as AOX (alcohol oxidase), DAS (dihydroxyacetone synthase), FDH (format dehydrogenase) and DAK (dihydroxyacetone kinase) were identified against BLAST searches. Using fast minimum evolution algorithm, phylogenetic tree was prepared to see the evolutionary relatedness. The phylogenetic analysis based on these genes and MEGA software revealed the position of methylotrophic yeasts (Okonechnikov et al. 2012). Methylotrophic yeast was reported to be a suitable expression system also. To enhance the yields of complex proteins having unnatural amino acids, a recombinant gene expression system was developed in methylotrophic yeast Pichia pastoris. In the investigation, it was emphasised that by modulating aaRS level, the optimization of expression of unnatural amino acids in the methylotrophic host cell was done and better than as reported earlier in Saccharomyces cerevisiae (Young et al. 2012). Earlier, S. cerevisiae was considered to be specific for unnatural amino acids, but in this investigation, it was explained that a mutant of recombinant human serum albumin with p-phenylalanine is expressed efficiently in the methylotrophic yeast system and therefore allows the higher production of complex proteins whose gene expression is difficult in the existing systems (Young et al. 2012). In a study, the alcohol oxidase activity was analysed in Pichia pastoris on the basis of two genes AOX1 and AOX2 and their expression analysis in the cell (Tschopp et al. 1987). The AOX1 gene expression was observed undetectable when cells are grown in the media with carbon sources other than methanol (Cregg et al. 1989). In a study, the genes encoding polyunsaturated fatty acids are targeted for the

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identification and phylogenetic analysis of methylotrophic yeast Hansenula polymorpha. In the investigation, gene encoding fatty acid elongase, HpELO1, was identified and characterised. The HpELO1 gene encoding a protein has 319 long amino acids, and it contains 5 different conserved membrane-spanning regions in yeast Elo protein family. The phylogenetic analysis based on amino acid sequences revealed that HpELO1 gene is an orthologue of S. cerevisiae ELO3. ELO3 gene is responsible for the elongation of VLCFAs (very long chain fatty acids) (Fang et al. 2017; Hong et al. 2019; Prasitchoke et al. 2007; Řezanka et al. 2018). To see the clear classification and taxonomy of Hansenula (Ogataea) polymorpha and other related species, the phylogenetic relatedness was observed based on conserved gene sequences (Suh and Zhou 2010; Yoo et al. 2019). The phylogenetic analysis was done based on ribosomal gene sequences of ITS and D1/D2 region of LSU rRNA gene, and this molecular analysis revealed that most of the O. Polymorpha strains were different phylogenetically from type strain of Pichia angusta (ATCC 14755), Ogataea thermophila was found to be evolutionarily related to O. Polymorpha and two novel strains of Candida (ATCC 26012 and ATCC 58401) were close to O. polymorpha. The character-based method was applied to construct the phylogenetic tree. The maximum likelihood-based and parsimonious trees were constructed by taking sequences of ITS and D1/D2 region of LSU rRNA gene (>1 Kb). The result showed the close evolutionary relatedness of O. angusta and C. parapolymorpha with O. polymorpha along with some significant evolutionary distances in the tree constructed. A very closest matching and relatedness was observed in case of O. thermophila and O. polymorpha with 100% bootstrap value and therefore were grouped in the same clade (Suh and Zhou 2010). The SSU (small subunit) and LSU (large subunit) ribosomal gene sequence-­based study revealed that five different methylotrophic yeast species of Ogataea genus, that is, O. henricii, O. philodendra, O. glucozyma, O. minuta and O. polymorpha were reported earlier but a variety of Ogataea minuta var. nonfermentas was distantly related with genus Pichia (Kurtzman et al. 2008; Naumov et al. 2018; Yamada et al. 1994). Thereafter, O. minuta var. nonfermentas was reclassified separately as O. nonfermentas (Kurtzman and Robnett 2010). The multigene-based identification and phylogenetic analysis illustrated more than 37 species of methylotrophic yeast and recently 67 species of Ogataea genus were reported (Kurtzman 2009; Lu et al. 2017). The multigene analysis involved different types of genes for the study such as LSU rRNA, SSU rRNA, elongation factor EF-1α and mitochondrial SSU rRNA gene. The multigene analysis revealed differentiation among genera such as Pachysolen, Nakazawaea, Ambrosiozyma, Komagataella, Phaffomyces and Ogataea (Kurtzman and Robnett 2010). The pioneer of yeast molecular phylogeny, K.P. Kurtzman, described the phylogenetic classification of yeasts based on multiple genes and evolutionary relationship was observed. In one of the investigations, a total of thirteen methylotrophic yeast strains utilising methanol as a carbon substrate (forming ascospore) were isolated and identified from the sap exudates of a tree Sclerobium sp. from Costa Rica and Brazil. Their characterisation for the identification and phylogenetic study was based on the sequence analysis of D1/D2 large subunit rDNA.  The ribosomal gene sequence

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analysis and neighbour joining method of phylogenetic tree construction revealed their identification as predominant species of Ogataea genus (syn. Pichia) and Candida sp. Later, few new isolates were identified as Ogataea falcaomoraisii (Morais et al. 2004). Three methylotrophic yeast strains were isolated from leaf and rotten wood samples of temperate forest in Hungary. The strains were found to be nitrate negative and assimilating methanol as a source of carbon. The D1/D2 large subunit rRNA gene sequence analysis grouped these strains in a clade of Ogataea sp. The strains have similar sequences and differ from genetically related and close species Pichia pilisensis. A novel methylotrophic yeast species Ogataea nitratoaversa was proposed to accommodate these nitrate-negative yeast strains. The variation in the D1/ D2 and ITS sequences was observed due to several substitutions. Since the investigation does not allow the inclusion of nitrate-negative strains, they were named as Ogataea yamada, maeda and mikata (Péter et al. 2008). In one of the earlier studies, two novel thermotolerant methylotrophic yeast strains were reported from soil and tree exudates from Thailand. The biochemical and phenotypic characterisation included the nitrate assimilation, hat-shaped ascospore formation, ubiquinone study, multilateral budding, urease reaction and other observations to identify the strains. The sequence analysis of D1/D2 rRNA gene revealed the phylogenetic relatedness between the species. The sequence analysis justifies two different strains PT44T and S051T. The PT44T strain was close to Pichia (Ogataea) dorogensis, whereas S051T strain was closely related to Pichia thermomethanolica with some nucleotide substitutions in the phylogenetic tree constructed. The biochemical, molecular, physiological and phenotypic characterisation of the strains proved them novel strains of genus Ogataea and further proposed with the name of Ogataea chonburiensis sp. nov. and Ogataea nakhonphanomensis sp. nov., respectively. Moreover, thermotolerant Pichia siamensis was renamed as Ogataea siamensis and Pichia thermomethanolica was renamed as Ogataea thermomethanolica in this study (Limtong et al. 2005).

3.3  Genetic Regulation in Yeast Methylotrophy A number of methylotrophic yeast strains are reported and characterised by the identification of C1 carbon substrate-inducible gene expression analysis. For the yeast methylotrophy, different essential enzymes such as AOX and DAS and others are required to carry out formaldehyde oxidation metabolic pathway (Nakagawa et  al. 1999; Sakai et  al. 1998; Yurimoto et  al. 2011). Molecular mechanism of methanol-­inducible gene expression is represented during growth on various carbon sources (Fig. 3.2). Genes responsible for carbon substrate metabolism in methylotrophic yeast were investigated in C. boidinii by cloning the genes coding methanolmetabolising enzymes (Yurimoto et al. 2002). Figure 3.2 represents the methanol metabolism in methylotrophic yeasts (Fig. 3.3).

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Fig. 3.2  Molecular mechanism of methanol-inducible gene expression. (a) Relative expression levels of H. polymorpha MOX (encoding AOD), C. boidinii AOD1, and C. boidinii DAS1 during growth on various carbon sources. On glucose-containing media, expression is completely repressed. When glucose is completely consumed or cells are shifted to glycerol medium, a derepressed level of expression of the AOD genes is induced (derepression) and the extent of derepression of the AOD genes differs between H. polymorpha and C. boidinii. When cells are grown on methanol, the maximum level of expression of AOD genes is achieved not only by derepression but also by methanol-specific gene activation. The induction of DAS1 on methanol medium is achieved only by methanol-specific gene activation. (b) During growth on glucose, expression of methanol-­ inducible genes is repressed. When cells are shifted to methanol, initially, a Trm2p-related derepression event occurs followed by a Trm1p-related methanol-specific gene activation. (Adapted from Yurimoto et al. (2011))

For the heterologous protein expression and production, Pichia pastoris strain is generally preferred. In a study, the gene copy number was determined for P. pastoris along with real-time PCR assay for the quantification of integrated expression cassettes (Abad et al. 2010). In yeast methylotrophy, the expression of genes involved in methanol metabolism is regulated by the presence of carbon source. The expression and repression of genes were studied by Ohsawa et al. (2018) in which it was

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Cytosol

NAD+ NADH Peroxisome

1/2 O2 + H2O

CTA

CH2(OH)OCH3 HCOOCH3 ADH (MFS)

CH OH O2 3 H2O2

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NAD+ NADH GS-CH2OH

HCHO RCOOH

RCOOOH

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GS-CH2OH

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DHA

Xu5P

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Xu5P

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1/3 GAP

NADH CO2

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Rearrangements FBP

DAK

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Fig. 3.3  Outline of methanol metabolism in methylotrophic yeasts. Enzymes: ADH (MFS), alcohol dehydrogenase (methyl formate-synthesising enzyme); AOD, alcohol oxidase; CTA, catalase; DAK, dihydroxyacetone kinase; DAS, dihydroxyacetone synthase; FDH, formate dehydrogenase; FGH, S-formylglutathione hydrolase; FLD, formaldehyde dehydrogenase; GLR, glutathione reductase; Pmp20, peroxisome membrane protein which has glutathione peroxidase activity. Abbreviations: DHA, dihydroxyacetone; DHAP, dihydroxyacetone phosphate; F6P, fructose 6-phosphate; FBP, fructose 1,6-bisphosphate; GAP, glyceraldehyde 3-phosphate; GS-CH2OH, S-hydroxymethyl glutathione; GS-CHO, S-formylglutathione; GSH, reduced form of glutathione; GSSG, oxidised form of glutathione; RCOOOH, alkyl hydroperoxide; Xu5P, xylulose 5-­phosphate. (Adapted from Yurimoto et al. (2011))

discussed that a significant and maximum gene expression was observed in the presence of methanol, whereas a low level of gene expression was observed in the absence of methanol (derepression). The characterisation and identification of various transcription factors involved in the expression and regulation of methanol-­ inducible gene expression was done by Ohsawa et al. (2018). Transcription factors KpMit1, leads to the repression of methanol-inducible gene expression or the presence of glucose leads to the repression of methanol-inducible gene expression (Hartner and Glieder 2006; Yamashita et al. 2009). Transcription factors such as KpMit1 in Komagataella phafi (Wang et al. 2016) and KpPrm1 in Candida boidinii (Sahu et al. 2014) along with Hap complex are involved in methanol induction, whereas transcription factors like KpMxr1 and CbTrm2 are involved in derepression (Lin-Cereghino et al. 2006). KpMxr1 and CbTrm2 are homologues to S. cerevisiae Adr1. In a recent investigation, the function of a gene encoding Hac1 transcription factor was characterised in thermotolerant methylotrophic yeast Ogataea thermomethanolica TBRC656 (OtHAC1). Hac1 generally triggers the unfolded protein response pathway in yeasts. Under the characterisation study of this gene, the comparative proteomic analysis was done between OtHAC1 mutant and wild-type Ogataea strain. About 400–500 proteins were detected and gene encoding Hac1 annotated different functions involved in transcription, translation, oxidative stress

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and secretary pathway. Subsequently, two different novel OtHAC1-dependent proteins, viz. Iml1 and Npr2, were also identified which are responsible for the regulation of autophagy. This research on methylotrophic yeast therefore revealed the regulation of different metabolic pathways or processes through OtHAC1 gene expression in thermotolerant Ogataea thermomethanolica TBRC656 (Phithakrotchanakoon et al. 2018). In Pichia pastoris, the regulation of AOX gene expression is done through repression/derepression mechanism along with induction mechanism and mostly this gene regulation resembles the regulation of GAL1 gene expression in Saccharomyces cerevisiae. The rich level of methanol in the media facilitates the high rate of transcription in case of methylotrophic yeast and the repression of gene regulation in the presence of glucose (a repressing carbon substrate) was not seen unlike GAL1 gene in case of Saccharomyces cerevisiae. The rate of gene expression was found directly proportional to the presence of methanol in the medium (Tschopp et al. 1987). The methylotrophic yeast Ogataea thermomethanolica TBRC656 is a well-­known host cell for the heterologous protein expression. In this thermotolerant methylotrophic yeast, maltase gene (mal) promoter-based new gene expression system was developed. The gene expression of xylanase and phytase was found to be enhanced many fold when Ogataea thermomethanolica TBRC656 was supplemented with sucrose in media. The increase in fold expression was due to activation of OtMal promoter gene as compared to constitutive OtGAP promoter gene. The presence of sucrose in the media also activates the more expression of OtMal promoter gene as compared to methanol-inducible OtAOX promoter. This enhances the enzyme activity by increasing higher gene expression of reporter genes coding xylanase and phytase. Therefore, it was suggested that methylotrophic yeast in the presence of sucrose as source of carbon substrate can be utilised for the production of heterologous proteins at large scale (Puseenam et al. 2018).

3.4  Methylotrophic Yeast and Impact to the Environments The methylotrophic communities have been found to be applicable in diverse potential biotechnological applications (Fig. 3.4). The methylotrophic communities have been reported from diverse habitats including plant microbiomes as rhizospheric (Verma et al. 2013, 2014; Yadav 2017), endophytic (Rana et al. 2018; Verma et al. 2015a, 2016b) and epiphytic (Verma et al. 2015b, 2016a) and natural habitats as well as from different extreme environments of acidic, alkaline, drought, low temperature (Yadav 2015; Yadav et al. 2015a, b, 2016, 2019c), salinities and radiations (Yadav and Saxena 2018; Yadav et al. 2015c; Yadav and Yadav 2018). The methylotrophic communities having potential and efficient multifarious plant growth-­ promoting attributes have been used for plant growth promotion, crop yield, and soil health for sustainable agriculture (Biswas et  al. 2018; Yadav et  al. 2017; Yadav 2009). Along with agricultural application, the methylotrophic communities have been reported to use in different processes in medical, industrial and pharmaceutical

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Fig. 3.4  Biotechnological application of methylotrophic yeast. (Adapted with permission from Kuroda and Ueda (2011))

sectors as well as in environment for sustainable future (Rastegari et al. 2019; Yadav et al. 2019a, b). The genetically engineered thermotolerant methylotrophic yeast strains are reported to have properties of bioremediation (Kour et al. 2019; Suman et al. 2016; Yadav et al. 2018). Particularly chromate bioremediation was observed in Hansenula polymorpha which triggers the many fold enhanced gene expression of FCb2 gene coding flavocytochrome b2 enzyme as compared to parental strain (Smutok et al. 2011). The flavocytochrome b2 enzyme is known to be specific for L-lactate. In the presence of L-lactate, the enzyme flavocytochrome b2 leads to the reduction of chromate by living cells. Pichia pastoris is reported to be involved in the degradation of azo dyes and anthraquinone dyes and bioremediation of different xenobiotic compounds (Colao et al. 2006). In the production of fungal laccase, the expression

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of lccI gene was done in Pichia pastoris from cDNA synthesised from the white rot fungus Trametes togi. Methylotrophic yeast isolated from oil-contaminated soil has distinct enzymatic activity and identified as Trichosporon cutaneum. Later, the identification was confirmed by ribosomal DNA-based molecular characterisation. In the isolate, methanol assimilation was found and can use formaldehyde also as a source of carbon substrate along with other carbon substrates like ethanol, glycerol, glucose and other petroleum derivatives. The microorganism was proved as an extremophile. In the isolate, different enzymatic activities were observed such as formaldehyde reductase, unspecific aldehyde dehydrogenase and formaldehyde dehydrogenase activity. Therefore, the biochemical, metabolic and physiological characteristics of methylotrophic isolate explore the new possibilities in the field of environmental biotechnology (Kaszycki et al. 2006). Pichia pastoris is well-known yeast used in the production of animal contaminant-­ free hydroxylated collagen. The enzyme-based and molecular methods are utilised in the production and selection of triple-transformed Pichia pastoris strain, useful in the expression of P4H (prolyl 4-hydroxylase) tetramer obtained from marine sponge Chondrosia reniformis along with a hydroxylated collagen from the same animal (Pozzolini et al. 2015). The environmental pollutants like household, industrial wastes and oil spills cost a lot to the sustainability of environment, and therefore some strains of methylotrophic yeast were found to be a better alternative for the bioremediation of these potent pollutants. The restoration of ecological balance is achieved by diminishing the level of environmental pollution. Methylotrophic yeast are able to degrade a number of xenobiotic compounds as a source of carbon. Methylotrophic strains such as Rhodosporidium, Pichia, Trichosporon, Rhodotorula and Yarrowia are able to degrade xenobiotic compounds like phenol, aromatic compounds and polar compounds. The degradation intensity decreases from n-alkanes to polar and aromatic carbon substrates (Csutak et al. 2010). Genetically engineered methylotrophic yeast Hansenula polymorpha is known for the development of lactate-selective microbial biosensor. This thermotolerant yeast was utilised for the overproduction of lactate:cytochrome c-oxidoreductase enzyme system [FC b(2)] by overexpression of the CYB2 gene encoding FC b(2). The strong alcohol oxidase promoter of H. polymorpha controls the gene expression of HpCYB2 gene, and it was transformed into the host methylotrophic yeast strain H. polymorpha C-105 (gcr1 catX) in the absence of catalase activity and with glucose repression. Using a cathodic electrodeposition polymer, the cells are immobilised over graphite base. The redox mediator phenazine methosulphate used with this reacts with FC b (2) inside the cell in the presence of L-lactate. A higher Km value is observed in a biosensor based on recombinant methylotrophic yeast with a higher linear range towards lactate (Smutok et al. 2011). Now it is very clear that methylotrophic yeast can be exploited as a suitable microbial culture used in the heavy-load wastewater treatment process. The pure monoculture of yeast can be utilised as a biofilter for the treatment of concentrated wastewater. The dilution of contaminants is done through degradation of wastes generated as a result of several technological processes. In earlier reports, it was

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illustrated that up to 10 g/l of formaldehyde can be diluted with the help of these biofilters. Moreover, methylotrophic yeast can be useful in the bioremediation of soil contaminated with oil (Kaszycki et  al. 2006). Since methylotrophic yeast is well known for the production of heterologous proteins, they can change the product or compounds after processing due to genetic changes in their genome. Yeast cells do not contain pyogenes, pathogens, and viral inclusions and therefore can be used in the production of therapeutic administration. Earlier it was assumed that mainly S. cerevisiae was utilised for the production of pharmaceutical proteins but later methylotrophic yeast Hansenula polymorpha was used widely in modern genetics for the production of pharmaceuticals. Hansenula polymorpha possess especial advantageous characteristics as a host cell for the production of pharmaceuticals proteins. The pharmaceutical protein production system based on methylotrophic yeast Hansenula polymorpha has been established for different vaccines, serum proteins and other important therapeutics. In future, different products based on H. Polymorpha will be introduced to market after preclinical and clinical trials (Gellissen and Melber 1996).

3.5  Conclusion and Future Prospects The molecular-level analysis enhances our understanding of methylotrophic yeast structure and function. The gene-level diversity and their study for the phylogenetic relatedness helped us to understand the establishment of methylotrophic yeast with the natural ecosystems. The methylotrophic yeasts are used in the research area and in biotechnological applications, one of the most important being the production of heterologous proteins of a large industrial and medical importance. The biotechnological production of heterologous proteins is reported from methylotrophic yeast Pichia pastoris which is renamed as Komagataella pastoris after recent taxonomy. The increase of SCP (Single Cell Protein) requirements or the remediations of the polluted systems by making use of natural alternatives represent important reasons for the necessity of characterising the methylotrophic yeasts. Moreover, further sophisticated and intensive research in the field of yeast methylotrophy and its molecular basis will explore and reveal new insights of physiological functions along with its importance in the natural ecosystem.

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Kurtzman CP, Robnett CJ (2010) Systematics of methanol assimilating yeasts and neighboring taxa from multigene sequence analysis and the proposal of Peterozyma gen. nov., a new member of the Saccharomycetales. FEMS Yeast Res 10:353–361 Kurtzman CP, Robnett CJ, Basehoar-Powers E (2008) Phylogenetic relationships among species of Pichia, Issatchenkia and Williopsis determined from multigene sequence analysis, and the proposal of Barnettozyma gen. nov., Lindnera gen. nov. and Wickerhamomyces gen. nov. FEMS Yeast Res 8:939–954 Kuroda K, Ueda M (2011) Cell surface engineering of yeast for applications in white biotechnology. Biotechnol Lett 33:1–9 Leão-Helder AN, Krikken AM, Van der Klei IJ, Kiel JA, Veenhuis M (2003) Transcriptional down-­ regulation of peroxisome numbers affects selective peroxisome degradation in Hansenula polymorpha. J Biol Chem 278:40749–40756 Limtong S, Srisuk N, Yongmanitchai W, Yurimoto H, Nakase T, Kato N (2005) Pichia thermomethanolica sp. nov., a novel thermotolerant, methylotrophic yeast isolated in Thailand. Int J Syst Evol Microbiol 55:2225–2229. https://doi.org/10.1099/ijs.0.63712-0 Limtong S, Srisuk N, Yongmanitchai W, Yurimoto H, Nakase T (2008) Ogataea chonburiensis sp. nov. and Ogataea nakhonphanomensis sp. nov., thermotolerant, methylotrophic yeast species isolated in Thailand, and transfer of Pichia siamensis and Pichia thermomethanolica to the genus Ogataea. Int J Syst Evol Microbiol 58:302–307 Lin-Cereghino GP, Godfrey L, Bernard J, Johnson S, Khuongsathiene S, Tolstorukov I, Yan M, Lin-Cereghino J, Veenhuis M, Subramani S (2006) Mxr1p, a key regulator of the methanol utilization pathway and peroxisomal genes in Pichia pastoris. Mol Cell Biol 26:883–897 Lu Y-F, Wang M, Zheng J, Hui F-L (2017) Ogataea neixiangensis sp. nov. and Ogataea paraovalis fa, sp. nov., two methanol-assimilating yeast species isolated from rotting wood. Int J Syst Evol Microbiol 67:3038–3042 Meena KK, Kumar M, Kalyuzhnaya MG, Yandigeri MS, Singh DP, Saxena AK, Arora DK (2012) Epiphytic pink-pigmented methylotrophic bacteria enhance germination and seedling growth of wheat (Triticum aestivum) by producing phytohormone. Antonie Van Leeuwenhoek 101:777–786 Mitsui R, Kusano Y, Yurimoto H, Sakai Y, Kato N, Tanaka M (2003) Formaldehyde fixation contributes to detoxification for growth of a nonmethylotroph, Burkholderia cepacia TM1, on vanillic acid. Appl Environ Microbiol 69:6128–6132 Morais PB, Teixeira LC, Bowles JM, Lachance M-A, Rosa CA (2004) Ogataea falcaomoraisii sp. nov., a sporogenous methylotrophic yeast from tree exudates. FEMS Yeast Res 5:81–85 Nakagawa T, Mukaiyama H, Yurimoto H, Sakai Y, Kato N (1999) Alcohol oxidase hybrid oligomers formed in vivo and in vitro. Yeast 15:1223–1230 Nakagawa T, Miyaji T, Yurimoto H, Sakai Y, Kato N, Tomizuka N (2000) A methylotrophic pathway participates in pectin utilization by Candida boidinii. Appl Environ Microbiol 66:4253–4257 Nakase T, Imanishi Y, Ninomiya S, Takashima M (2010) Candida rishirensis sp. nov., a novel methylotrophic anamorphic yeast species isolated from soil on Rishiri Island in Japan. J Gen Appl Microbiol 56:169–173 Naumov GI, Naumova ES, Lee C-F (2017) Ogataea haglerorum sp. nov., a novel member of the species complex, Ogataea (Hansenula) polymorpha. Int J  Syst Evol Microbiol 67:2465–2469 Naumov G, Shalamitskiy MY, Naumova E, Lee C-F (2018) Phylogenetics, biogeography, and ecology of methylotrophic yeasts of the heterogeneous genus Ogataea: achivements and prospects. Microbiology 87:443–452 Negruţă O, Csutak O, Stoica I, Rusu E, Vassu T (2010) Methylotrophic yeasts: diversity and methanol metabolism. Rom Biotechnol Lett 15:5369–5375 Ohsawa S, Nishida S, Oku M, Sakai Y, Yurimoto H (2018) Ethanol represses the expression of methanol-inducible genes via acetyl-CoA synthesis in the yeast Komagataella phaffii. Sci Rep 8:18051 Okonechnikov K, Golosova O, Fursov M, Team U (2012) Unipro UGENE: a unified bioinformatics toolkit. Bioinformatics 28:1166–1167

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Olah GA (2005) Beyond oil and gas: the methanol economy. Angew Chem Int Ed 44:2636–2639 Péter G, Tornai-Lehoczki J, Dlauchy D (2008) Ogataea nitratoaversa sp. nov., a methylotrophic yeast species from temperate forest habitats. Antonie Van Leeuwenhoek 94:217 Phithakrotchanakoon C, Puseenam A, Phaonakrop N, Roytrakul S, Tanapongpipat S, Roongsawang N (2018) Hac1 function revealed by the protein expression profile of a OtHAC1 mutant of thermotolerant methylotrophic yeast Ogataea thermomethanolica. Mol Biol Rep 45:1311–1319 Pozzolini M, Scarfì S, Mussino F, Salis A, Damonte G, Benatti U, Giovine M (2015) Pichia pastoris production of a prolyl 4-hydroxylase derived from Chondrosia reniformis sponge: a new biotechnological tool for the recombinant production of marine collagen. J Biotechnol 208:28–36 Prasitchoke P, Kaneko Y, Bamba T, Fukusaki E, Kobayashi A, Harashima S (2007) Identification and characterization of a very long-chain fatty acid elongase gene in the methylotrophic yeast, Hansenula polymorpha. Gene 391:16–25 Puseenam A, Kocharin K, Tanapongpipat S, Eurwilaichitr L, Ingsriswang S, Roongsawang N (2018) A novel sucrose-based expression system for heterologous proteins expression in thermotolerant methylotrophic yeast Ogataea thermomethanolica. FEMS Microbiol Lett 365:238 Rana KL, Kour D, Yadav AN (2018) Endophytic microbiomes: biodiversity, ecological significance and biotechnological applications. Res J Biotechnol 14:1–30 Rastegari AA, Yadav AN, Gupta A (2019) Prospects of renewable bioprocessing in future energy systems. Springer International Publishing, Cham Ravin NV, Eldarov MA, Kadnikov VV, Beletsky AV, Schneider J, Mardanova ES, Smekalova EM, Zvereva MI, Dontsova OA, Mardanov AV (2013) Genome sequence and analysis of methylotrophic yeast Hansenula polymorpha DL1. BMC Genomics 14:837 Řezanka T, Lukavský J, Vítová M, Nedbalová L, Sigler K (2018) Lipidomic analysis of Botryococcus (Trebouxiophyceae, Chlorophyta)-identification of lipid classes containing very long chain fatty acids by offline two-dimensional LC-tandem MS. Phytochemistry 148:29–38 Sahu U, Rao KK, Rangarajan PN (2014) Trm1p, a Zn (II) 2Cys6-type transcription factor, is essential for the transcriptional activation of genes of methanol utilization pathway, in Pichia pastoris. Biochem Biophys Res Commun 451:158–164 Sakai Y, Nakagawa T, Shimase M, Kato N (1998) Regulation and physiological role of theDAS1 gene, encoding dihydroxyacetone synthase, in the methylotrophic yeast Candida boidinii. J Bacteriol 180:5885–5890 Smutok O, Broda D, Smutok H, Dmytruk K, Gonchar M (2011) Chromate-reducing activity of Hansenula polymorpha recombinant cells over-producing flavocytochrome b2. Chemosphere 83:449–454 Suh S-O, Zhou JJ (2010) Methylotrophic yeasts near Ogataea (Hansenula) polymorpha: a proposal of Ogataea angusta comb. nov. and Candida parapolymorpha sp. nov. FEMS Yeast Res 10:631–638 Suman A, Yadav AN, Verma P (2016) Endophytic microbes in crops: diversity and beneficial impact for sustainable agriculture. In: Singh D, Abhilash P, Prabha R (eds) Microbial inoculants in sustainable agricultural productivity, research perspectives. Springer-Verlag, New Delhi, pp 117–143. https://doi.org/10.1007/978-81-322-2647-5_7 Tani Y, Yamada K (1987) Diversity in glycerol metabolism of methylotrophic yeasts. FEMS Microbiol Lett 40:151–153 Tschopp JF, Brust PF, Cregg JM, Stillman CA, Gingeras TR (1987) Expression of the lacZ gene from two methanol-regulated promoters in Pichia pastoris. Nucleic Acids Res 15:3859–3876 van der Klei IJ, Yurimoto H, Sakai Y, Veenhuis M (2006) The significance of peroxisomes in methanol metabolism in methylotrophic yeast. Biochim Biophys Acta 1763:1453–1462 Veenhuis M, Van Der Klei I, Titorenko V, Harder W (1992) Hansenula polymorpha: an attractive model organism for molecular studies of peroxisome biogenesis and function. FEMS Microbiol Lett 100:393–403 Verma P, Yadav AN, Kazy SK, Saxena AK, Suman A (2013) Elucidating the diversity and plant growth promoting attributes of wheat (Triticum aestivum) associated acidotolerant bacteria from southern hills zone of India. Natl J Life Sci 10:219–227

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Verma P, Yadav AN, Kazy SK, Saxena AK, Suman A (2014) Evaluating the diversity and phylogeny of plant growth promoting bacteria associated with wheat (Triticum aestivum) growing in central zone of India. Int J Curr Microbiol App Sci 3:432–447 Verma P, Yadav AN, Khannam KS, Panjiar N, Kumar S, Saxena AK, Suman A (2015a) Assessment of genetic diversity and plant growth promoting attributes of psychrotolerant bacteria allied with wheat (Triticum aestivum) from the northern hills zone of India. Ann Microbiol 65:1885–1899 Verma P, Yadav AN, Shukla L, Saxena AK, Suman A (2015b) Alleviation of cold stress in wheat seedlings by Bacillus amyloliquefaciens IARI-HHS2-30, an endophytic psychrotolerant K-solubilizing bacterium from NW Indian Himalayas. Natl J Life Sci 12:105–110 Verma P, Yadav AN, Khannam KS, Kumar S, Saxena AK, Suman A (2016a) Molecular diversity and multifarious plant growth promoting attributes of Bacilli associated with wheat (Triticum aestivum L.) rhizosphere from six diverse agro-ecological zones of India. J Basic Microbiol 56:44–58 Verma P, Yadav AN, Khannam KS, Mishra S, Kumar S, Saxena AK, Suman A (2016b) Appraisal of diversity and functional attributes of thermotolerant wheat associated bacteria from the peninsular zone of India. Saudi J Biol Sci. https://doi.org/10.1016/j.sjbs.2016.01.042 Wang X, Wang Q, Wang J, Bai P, Shi L, Shen W, Zhou M, Zhou X, Zhang Y, Cai M (2016) Mit1 transcription factor mediates methanol signaling and regulates the alcohol oxidase 1 (AOX1) promoter in Pichia pastoris. J Biol Chem 291:6245–6261 Yadav AN (2009) Studies of Methylotrophic Community from the phyllosphere and rhizosphere of tropical crop plants. M.Sc. Thesis, Bundelkhand University, pp  66, https://doi. org/10.13140/2.1.5099.0888 Yadav AN (2015) Bacterial diversity of cold deserts and mining of genes for low temperature tolerance. Ph.D.  Thesis, IARI, New Delhi/BIT, Ranchi pp  234, https://doi.org/10.13140/ RG.2.1.2948.1283/2 Yadav AN (2017) Agriculturally important microbiomes: biodiversity and multifarious pgp attributes for amelioration of diverse abiotic stresses in crops for sustainable agriculture. Biomed J Sci Tech Res 1:1–4 Yadav AN, Saxena AK (2018) Biodiversity and biotechnological applications of halophilic microbes for sustainable agriculture. J Appl Biol Biotechnol 6:1–8 Yadav AN, Yadav N (2018) Stress-adaptive microbes for plant growth promotion and alleviation of drought stress in plants. Acta Sci Agric 2:85–88 Yadav AN, Sachan SG, Verma P, Saxena AK (2015a) Prospecting cold deserts of north western Himalayas for microbial diversity and plant growth promoting attributes. J  Biosci Bioeng 119:683–693 Yadav AN, Sachan SG, Verma P, Tyagi SP, Kaushik R, Saxena AK (2015b) Culturable diversity and functional annotation of psychrotrophic bacteria from cold desert of Leh Ladakh (India). World J Microbiol Biotechnol 31:95–108 Yadav AN, Verma P, Kumar M, Pal KK, Dey R, Gupta A, Padaria JC, Gujar GT, Kumar S, Suman A, Prasanna R, Saxena AK (2015c) Diversity and phylogenetic profiling of niche-specific Bacilli from extreme environments of India. Ann Microbiol 65:611–629 Yadav AN, Sachan SG, Verma P, Saxena AK (2016) Bioprospecting of plant growth promoting psychrotrophic Bacilli from cold desert of north western Indian Himalayas. Indian J Exp Biol 54:142–150 Yadav A, Verma P, Kumar R, Kumar V, Kumar K (2017) Current applications and future prospects of eco-friendly microbes. EU Voice 3:21–22 Yadav AN, Verma P, Kumar V, Sangwan P, Mishra S, Panjiar N, Gupta VK, Saxena AK (2018) Biodiversity of the genus Penicillium in different habitats. In: Gupta VK, Rodriguez-Couto S (eds) New and future developments in microbial biotechnology and bioengineering, Penicillium system properties and applications. Elsevier, Amsterdam, pp  3–18. https://doi.org/10.1016/ B978-0-444-63501-3.00001-6 Yadav AN, Mishra S, Singh S, Gupta A (2019a) Recent advancement in white biotechnology through fungi Volume 1: diversity and enzymes perspectives. Springer International Publishing, Cham

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Yadav AN, Mishra S, Singh S, Gupta A (2019b) Recent advancement in white biotechnology through fungi. Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham Yadav AN, Yadav N, Sachan SG, Saxena AK (2019c) Biodiversity of psychrotrophic microbes and their biotechnological applications. J Appl Biol Biotechnol. Online first Yamada Y, Maeda K, Mikata K (1994) The phylogenetic relationships of the hat-shaped ascospore-­ forming, nitrate-assimilating Pichia species, formerly classified in the genus Hansenula Sydow et Sydow, based on the partial sequences of 18S and 26S ribosomal RNAs (Saccharomycetaceae): the proposals of three new genera, Ogataea, Kuraishia, and Nakazawaea. Biosci Biotechnol Biochem 58:1245–1257 Yamashita S, Yurimoto H, Murakami D, Yoshikawa M, Oku M, Sakai Y (2009) Lag-phase autophagy in the methylotrophic yeast Pichia pastoris. Genes Cells 14:861–870 Yoo SJ, Moon HY, Kang HA (2019) Screening and selection of production strains: secretory protein expression and analysis in Hansenula polymorpha. In: Gasser B, Mattanovich D (eds) Recombinant protein production in yeast. Springer New York, New York, pp 133–151 Young EM, Comer AD, Huang H, Alper HS (2012) A molecular transporter engineering approach to improving xylose catabolism in Saccharomyces cerevisiae. Metab Eng 14:401–411 Yurimoto H, Sakai Y, Kato N (2002) Methanol metabolism. In: Gellissen G (ed) Hansenula polymorpha. https://doi.org/10.1002/3527602356.ch5 Yurimoto H, Oku M, Sakai Y (2011) Yeast methylotrophy: metabolism, gene regulation and peroxisome homeostasis. Int J Microbiol 2011:101298

Chapter 4

White Rot Fungi and Their Enzymes for the Treatment of Industrial Dye Effluents Dhevagi Periasamy, Sudhakarn Mani, and Ramya Ambikapathi

4.1  Introduction Dyes are organic colourants, which are indispensable, as they are widely used in all fields of industry to envisage a colourful world. Among the different classes of dyes, azo dyes are most commonly used in textile industry, and nearly 10–15% of the dye stuffs used do not bind to the fabrics. The unabsorbed dyes were released into sewage treatment systems and in other water resources of the environment (Anliker 1979; Chudgar 1985; Zollinger 1961). It was shown that in environment, an unchanged azo dye was subjected to a reductive transformation of the azo bond. The reductive ring cleavage of the azo linkage is due to unspecific cytoplasmic reductases, resulting in the formation and accumulation of colourless aromatic amines (Chung 2000). The resultant products formed may be toxic, mutagenic and carcinogenic to animals and humans. Biological degradation is a popular, viable and attractive technology that uses the metabolic potential of microorganisms. In recent days, the potential of white rot fungi for the degradation of recalcitrants is gaining importance. White rot fungi (WRF) that come under the division Eumycota are heterogeneous group of fungi having capacity to degrade a wide variety of recalcitrant compounds. It contains the mushrooms, puffballs, conks and crust-like fungi which are used as food source. The xenobiotic degradation capacity of the WRF fungi may be due to extracellular, non-specific enzymes. Recently, white-rot fungi and their ligninolytic enzymes, including laccase, manganese peroxidase (MnP) and lignin peroxidase (LiP), as well as H2O2-producing oxidases, were explored intensively for the degradation of a wide range of xenobiotics. These fungi are having the capacity to break down the lignin in wood without degrading cellulose, and sometimes

D. Periasamy (*) · S. Mani · R. Ambikapathi Department of Environmental Sciences, Tamil Nadu Agricultural University, Coimbatore, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_4

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both lignin and cellulose will be degraded. The non-specific enzymes enable a number of advantages which are not been found in other bioremediation systems. The white rot fungus oyster mushroom (Pleurotus ostreatus) prefers lignin instead of polysaccharides during the degradation. Some of the commonly cultivated Pleurotus ostreatus and other oyster mushrooms will not grow on living trees. They are not parasitic and attack only already dying trees from other causes. During the degradation, the phenyl propane alkyl side chains found abundantly in the lignin was decayed by WRF. One other white rot fungus, Phanerochaete chrysosporium, shows no preference to lignocellulosics. Armillaria spp. is a white rot fungus called as honey mushroom which is notorious for attacking living trees. Turkey tail, artist’s conk and tinder fungus are some of the other white rot fungi involved in degradation.

4.2  Textile Dyes Colour has become an important aesthetic factor in the textile world. Different dyes and pigments are used as colourants, which are based on two major chemistries: azo and anthraquinone. Among these two, azo dyes are largely used in textile industries, and many of these dyes enter wastewater treatment facilities (Chung et al. 1978; Sudhakar et al. 2002). These dyes exhibit high resistance to microbial degradation in wastewater treatment systems and retain their colour, structural integrity upon exposure to sunlight, soil, bacteria and human sweating. Synthetic fabrics like nylon, rayon and polyester necessitate the production of new dyes that can bind with these materials strongly. In addition, updating of azodyes to match the changing social scenario, ideas and styles is also a must. Nearly 1,00,000 commercially available dyes (7 × 105 metric tons) are produced annually (Zollinger 2003). More than 8000 chemicals associated with textile dyeing process are listed in the Colour Index. Brighter, longer-lasting colours with better binding ability are often necessary to satisfy the emerging demand. A multitude of dyes were used in textile industry and different classes of these dyes include azo, acid, reactive, metal complex, disperse, vat, mortant, direct, basic, suphur, etc. (Vijaya et al. 2003). These dyes are usually aromatic and heterocyclic compounds (Vyas and Molitoris 1995) and some are toxic and carcinogenic. Azo dyes contain one to many N=N double bonds, hence many different structures are possible (Cripps et al. 1990; Zollinger 2003). For example, monoazo dyes have only one N=N double bond, while diazo and triazo dyes containing two and three N=N double bonds, respectively. The azo groups are linked to aromatic heterocyclic or enolizable aliphatic groups, but also attached with benzene and naphthalene rings (Zollinger 2003). Aromatic heterocyclic, enolizable aliphatic groups side chains are necessary for imparting the dye colour, with different shades and brightness. Dye molecules are generally described as chromogen, which contains nucleophiles often referred to as auxochromes and the aromatic groups are called as chromophores. Synthesis of most azo dyes involves diazotization of a primary ­aromatic amine, followed by coupling with one or more nucleophiles. Amino- and hydroxyl- groups are the commonly used coupling components. Because of the

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possibilities of the synthesis of diversity of dye components, a large number of structurally different azo dyes exist and are used in industry (McCurdy 1991). Worldwide production of organic dyes is currently estimated at nearly 4,50,000 tons, with 50,000 tons being lost in effluents during application and manufacture (Lewis 1999). Reactive dyes are coloured compounds that contain one or two functional groups capable of forming covalent bonds with the active sites in fibres. Almost 80–95% of the reactive dyes are based on azochromogen (Edwards 2000; Zollinger 2003). The carbon and phosphorus atom found in the dye molecule will bind with hydroxyl groups in cellulose, amino, thiol, and hydroxyl groups in wool, or amino groups in polyamides. Most fibre-reactive azo dyes are used for dyeing cellulosic materials, such as cotton, and are a major source of dye waste in textile effluents. Fibre-reactive azo dyes exhibit a high wet-fastness, due to their ability to covalently bond to substrates. However, dyes that hydrolyse in solution prior to bonding to a substrate are often lost in the washing (Loyd 1992). Schematic diagram of fibre-reactive azo dyes is given in Fig. 4.1. Amino and alkylamino groups are generally used for bridging chromogen and reactive group. The bridging group that binds chromogen and the reactive group must be stable, must be water-soluble and should have flexibility. Mono-, di-, and trichlorotriazinyl are all examples of reactive functional groups, which help to bond the dye molecule with a substrate through nucleophilic substitution, sometimes addition also. Nearly 200 different reactive dye groups are patented with an addition of 25 reactive azo dyes in the list per year from 1998, which was nearly five times as many as from other classes of azo dyes (Freeman and Sokolowska 1999). With such a disproportionate production rate, it is not surprising that a large percentage of dye pollution problems are related to fibre-reactive azo dyes. India, the former USSR, Eastern Europe, China, South Korea and Taiwan consume approximately 600 thousand tons (kt) of dyes per annum (Ishikawa and Leder 2000). The total annual world textile dye production is estimated at about 800 kt (Will et al. 2000; Zollinger 2003). Now Asia is being the largest dyestuff market (about 42%). Classification of dyestuffs into native and synthetic dyes may not be sufficient nowadays, since many natural substances are synthesized easily. According to the chemical structure, the dyes are classified systematically and given as colour index (Table  4.1). This indexing also provides information about the biodegradability of the dyes. A listing of synthetic dyes according to their most predominant chemical structures is given in Table 4.2.

Fig. 4.1  Schematic diagram of fibre-reactive azo dye

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Table 4.1  Classes of synthetic dyes according to colour index Chemical class Nitroso Nitro Monoazo Disazo Trisazo Polyazo Azoic Stilbene Carotenoid Diphenylmethane Triarylmethane Xanthene Acridine Quinoline Methine

Code 10,000 10,300 11,000 20,000 30,000 35,000 37,000 40,000 40,800 41,000 42,000 45,000 46,000 47,000 48,000

Chemical class Thiazole Indamine/Indophenol Azine Oxazine Thiazine Sulphur Lactone Aminoketone Hydroxyketone Anthraquinone Indigoid Phthalocyanine Natural Oxidation base Inorganic

Code 49,000 49,400 50,000 51,000 52,000 53,000 55,000 56,000 57,000 58,000 73,000 74,000 75,000 76,000 77,000

(Source: Wesenberg et al. 2003)

4.3  Dyes Used in India Indian dyestuff industry produces around 60,000 metric tons of dyes, which is 6.6% of total colourants used globally (Teli 2008). Today India exports dyes to USA, Turkey, Bangladesh, China and Germany on which once dependent for imports. The various dyes and dyestuffs used in India are azodyes, disperse dyes, ingrain dyes, naphthols, vat dyes, reactive dyes, pigment emulsion, sulphur dyes and other dyes. The type of dyes and chemicals used in the textile industry are found to differ depending on the fabrics manufactured (Table 4.3). The quantity of wastewater produced from composite industries (cotton and synthetics) was estimated as 840 m3/ day, with synthetic industry alone 180 m3/day. The textile processing and blending industry produces about 150 m3/day and 1500 m3/day, respectively, whereas woollen industry uses 2700 m3/day (Sarayu and Sandhya 2012).

4.4  D  escription of Dyeing Process and Sources of Effluent Generation Among high water-consuming industries, dye industry is also one, which dominates in Coimbatore, Tiruppur and Erode districts of Tamil Nadu. Substantial volume of water and numerous chemicals are used to each kilogram of hosiery. Nearly 200– 300 litres of water is consumed for a kilogram of yarn and 75–90% of which is discharged as effluent containing organic and inorganic pollutants (Banat et  al. 1997). Dyes have strong staining properties and they are visible even at a concentration of 1 ppm. The volume of waste water generated from textile industry is very

Table 4.2  Structure of synthetic dyes used in textile industries Name of dyes Reactive Orange 96 (N=N)

Remazol Brilliant Blue

Bromothymol Blue

Reactive Orange 16

Disperse Violet 93

Amaranth dye

Congo Red

Remazol Black 5

Reactive Orange 16 s

Chemical structure

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Table 4.3  Type of dyes and chemicals used by the Indian textile industries S. No. Dyes 1. Acid, metal–complex and reactive 2. 3.

Reactive, direct, vat, sulphur, indigo, etc. Cellulosics and pigments

4. 5.

Cationic Reactive dyes (Remazol, Procion MX and Cibacron F), direct dyes (Congo Red, Direct Yellow 50 and Direct Brown 116), Naphthol dyes (Fast Yellow GC, Fast Scarlet R and Fast Blue B) and indigo dyes (Indigo White, Tyrian Purple and Indigo Carmine) Acid dyes (azo dyes, triarylmethane dyes and anthraquinone dyes) and Lanaset dyes (Blue 5G and Bordeaux B) Dispersed dyes (Disperse Yellow 218 and Disperse Navy 35), basic dyes (Basic Orange 37 and Basic Red 1) and direct dyes

6. 7.

Material/ Fabrics Wool, Silk Cotton All kinds of materials Acrylics Cellulosic fibres

References Chavan (2001), Ghaly et al. (2014), Teli (2008)

Protein fibres Synthetic fibres

high, and the average concentration of 300 mg/L was also reported in many industries. Until recently, the waste water generated from these industries were discharged into sewage, running or stagnant water bodies and adjoining waste lands without proper treatment. Due to high composition, variety and colour intensity, wastewater from textile industry cannot be treated satisfactorily. However, due to stringent environmental regulations and awareness, the dye industries waste water is treated at Common Effluent Treatment Plants (CETP) or at individual factory premises before recycling. Perhaps, before the introduction of CETP or individual treatment plants, sizeable land area was contaminated already by this industrial waste water. The physicochemical characteristics of untreated textile and dye industries effluent indicate that it will have deleterious effect on soil and soil biological properties. The concentrations of most of the chemical parameters of the waste water were well above the critical limits fixed by Central and State Pollution Control Board of India. Characterizing the untreated textile and dye industry waste water, Gupta (1992) reported a pH range of 10–11.5, electrical conductivity in the range of 8.5–13.9 dS/m and BOD in the range of 400–800 mg/L. Besides high-soluble salts, the effluent also contains toxic trace metals such as lead (1.3 mg/L) and chromium (5–20 mg/L) beyond the critical limits (Kothandaraman et al. 1976). Dye effluent with a pH of 4–12, colour to the level of 500–2000 Pt-Co units, Chromium (VI) 1–4  mg/L and sulphide 0–50  mg/L was reported by Puscas et  al. (2003) (Fig. 4.2). The following steps are involved in the dyeing process: (a) Scouring  – The raw material, mainly the hosiery cloth, is subjected to the scouring operation for eliminating the cotton impurities by boiling the cloth in a vessel at about 80 °C for about three hours. (b) Bleaching – H2O2 and caustic soda of 3% strength is mixed in the vessel thoroughly prior to carrying out the scouring operation.

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Fig. 4.2  Dyeing process in textile industries

(c) Neutralizing with acid and washing  – The contents are washed using the alkaline water followed by the acid-wash for neutralizing and again the washing operation is repeated. After the final washing of the bleached cloth, the dyeing operation is carried out by transferring the material into the dyeing vat. (d) Dyeing  – The dyeing solution is prepared by dissolving the dyes as per the recipe for a particular shade in the winch along with the salts such as sodium chloride (40–60 g/lit strength) and soda ash for exhaustion and fixing the dyes, respectively. The dyeing operation is done at 70 °C and for about three hours to achieve the desired shade.

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4.4.1  Characterization of the Raw Effluents Textile industry wastewater contains dyes to the level of 20–200 mg/L, and about 10–20% of the dyes are present in effluents along with other organic and inorganic accessory chemicals. Reactive dyes, sodium chloride, soda ash, caustic soda, wetting oil, industrial soap powder, hydrochloric acid, acetic acid, softening agent and fixing agents are some of the accessory chemicals used in textile industries. Tiruppur has large sources of bleaching and dyeing industries which generate between 100 and 120 MLD of effluents. Due to poor dyeing process in the textile industry, 10–15% of the dyes are lost in the effluents of textile units, rendering them highly coloured (Boer et al. 2004; Vaidya 1982). It is estimated that 280,000 tons of textile dyes are discharged in such industrial effluents every year worldwide (Maas and Chaudhari 2005). The raw effluent samples were collected based on composite sampling method from Tirupur. The samples were collected on hourly basis and homogenized, and then the representative samples were analysed (Table 4.4.)

4.5  Enzymes of White Rot Fungi With the advancement of biotechnological tools, the use of fungi or bacteria, often in combination with physico-chemical processes were used eco-efficiently for combating this pollution source (Yadav et al. 2016, 2017, 2018, 2019a, b). Among those microorganisms, WRF are most efficient in breaking down synthetic dyes. These established a diverse eco-physiological group comprising mostly basidiomycetous (and, to a lesser extent, litter-decomposing) fungi capable of extensive aerobic lignin mineralization and depolymerization. This property is based on one or more extracellular lignin-modifying enzymes (LMEs) produced by white rot fungi, and these enzymes are also capable of degrading a wide range of xenobiotics (Beydilli et al. 1998; Borchert and Libra 2001; McMullan et  al. 2001; Robinson et  al. 2001; Willmott et al. 1998; Zissi and Lyberatos 2001). In 1896, laccase was identified first time in fungi by both Bertrand and Laborde. Most of the laccases have been widely isolated from fungal origin specifically from white rot fungi belonging to Ascomycetes, Deuteromycetes and Basidiomycete (Gochev and Krastanov 2007). Many other earlier studies also reported laccase production from white rot fungi (Kiiskinen et  al. 2004) such as Phlebia radiata (Niku-­Paavola et  al. 1988), Trametes versicolor (Bourbonnais et  al. 1995) and Pleurotus ostreatus (Palmieri et al. 2000). Some of Trichoderma sp. are also studied as laccase source. For example, T. harzianum (Hölker et  al. 2002), T. atroviride (Hölker et  al. 2002) and T. longibrachiatum produced higher amount of laccase. Velázquez-­Cedeño et al. (2004) observed that the laccase synthesis was found to be higher in mixed cultures than the pure cultures. Liophora terristrus, Phanerochaete chrysosporium, Lenzitis betulina and Stereum ostrea (Viswanath et  al. 2008) are

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some of the important Basidiomycetes which have been reported as the sources of laccases. WRF possess not only laccases, it also possess a wide range of various enzymes such as extracellular ligninolytic enzymes (manganese peroxidase, lignin peroxides) and hydrolytic enzymes (cellulase, xylanase, pectinase) (Teerapatsakul et al. 2007). The expression pattern of enzymes mainly depends on the organism itself. Certain WRF produce manganese peroxidase and lignin peroxidase, but not laccase; meanwhile, others produce laccase and manganese peroxidase, but not lignin peroxidase (Hatakka 1994). Therefore, among various types of WRF, some can decompose all of the lignocellulose components in wooden material, while some can degrade hemicellulose and lignin (Fang et  al. 2008). These enzymes are mostly important in industrial purpose and have a great potential in the processes of degradation of recalcitrant substances (Tortella et al. 2008).

4.5.1  Lignin Peroxidase (LiP) WRF can degrade lignin and a range of diverse environmental pollutants by means of their extracellular ligninolytic systems. Purified forms of LiP have been found to directly oxidize recalcitrant xenobiotic compounds such as polycyclic aromatic hydrocarbons, chlorophenols and azo dyes (Collins et al. 1997). LiPs are proposed to oxidize lignin with free radicals generated through oxidation of various secreted metabolites (e.g. veratryl alcohol). The most effective stimulant was found to be veratryl alcohol, a secondary metabolite produced by ligninolytic cultures of WRF (Sugiura et al. 2003). Veratryl alcohol plays an important role in LiP catalysis. LiP is oxidized by H2O2 to form a two electron intermediates, compound I, which oxidizes substrates by one electron, forming the more reduced enzyme intermediate, compound II. Compound II can then oxidize substrates by one electron, returning the enzyme to the ferric state. However, compound II has a very high reactivity with H2O2; therefore, in the presence of a poor substrate and excess H2O2, it is instead converted to an inactive form of the enzyme, compound III. Veratryl alcohol, when present, is a more favourable substrate for compound II and functions to convert it to the resting enzyme, completing the catalytic cycle (Vasina et al. 2017) (Fig. 4.3).

4.5.2  Manganese Peroxidase (MnP) MnP plays a vital role in lignin degradation, as it is found in all lignin-degrading WRF.  This haem protein belongs to the commonly occurring class II peroxidase group in basidiomycetous fungi and has a highly specific Mn2+ binding site. In the binding site of classical long MnPs, there are three amino acid residues while several fungal Mn2+-oxidizing enzymes with an additional tryptophan residue on the enzyme surface have been found (Hofrichter et al. 2001). These enzymes are called VPs or hybrid MnPs, bearing resemblance to LiPs and able to perform oxidation though

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Table 4.4  Characteristics of combined raw effluent from Tirupur S. No. 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 Sources: Sivasamy (2008)

Parameters pH EC (dS m−1) Colour (NTU) Turbidity (NTU) Total dissolved solids (mg L−1) Total suspended solids (mg L−1) Biochemical oxygen demand (mg L−1) Chemical oxygen demand (mg L−1) Total organic carbon (mg L−1) Total hardness (mg L−1) M-Alkalinity (mg L−1) P-Alkalinity (mg L−1) Calcium (mg L−1) Magnesium (mg L−1) Oil and grease (mg L−1) Chloride (mg L−1) Sulphate (mg L−1) Sodium (mg L−1) Potassium (mg L−1) Carbonate (mg L−1) Bicarbonate (mg L−1) Total phosphate (mg L−1) Fluorides as F (mg L−1) Nitrate as N (mg L−1) Ammonical nitrogen (mg L−1) Total Kjeldal nitrogen (mg L−1) Total iron (mg L−1) Barium (mg L−1) Boron (mg L−1) Aluminium (mg L−1) Zinc (mg L−1) Lead (mg L−1) Manganese (mg L−1) Copper (mg L–1) Chromium (mg L−1) Cobalt (mg L−1) Nickel (mg L−1) Cadmium (mg L−1) Arsenic (mg L−1) Total silica (mg L−1) Strontium (mg L−1)

Value 7.5–10 10–16 300–324 100–770 6250–8205 60–120 75–500 288–1200 100–111 500–900 570–1000 20–80 37–400 250–847 10–20 200–3900 460–3000 2000–3000 40–100 30–100 700–1200 0.9–4.0 1.1–4.0 0.6–6.0 2.0–6.0 10.0–40.0 0.195–2.00 0.15–0.25 0.50–1.00 0.10–1.20 0.017–0.50 0.046–0.25 0.05–0.30 0.041–0.20 0.00–0.05 0.05–0.10 0.031–0.20 0.04–0.06 BDL 20.0–40.0 1.00–2.00

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Fig. 4.3  Lignin peroxidase (LiP) catalytic cycle

long-range electron transfer as well. The evolution of these class II peroxidases appears to be closely related to each other and even consistent with the sharp decline in coal accumulation rate during the Permo-Carboniferous period (Maijala et  al. 2008). Lignin is the main precursor for coal. MnP catalyses the oxidation of Mn2+ ions to highly reactive Mn3+ ions. Chelated Mn3+ in turn acts as low molecular weight mediators that are able to attack phenolic structures. MnP is able to cause substantial depolymerization in in vitro biomass treatments (Floudas et al. 2012). Potential applications for MnP include pulp bleaching, biomechanical pulping, dye decolourization, bioremediation and production of highly valuable chemicals from residual lignin from biorefineries, pulp and paper side-streams (Järvinen et al. 2012) (Fig. 4.4).

4.5.3  Laccase Laccase is a part of broad group of enzymes called polyphenol oxidases containing copper atoms in the catalytic centre and are usually called multicopper oxidases. Laccases contain three types of copper atoms, one of which is responsible for their

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Fig. 4.4  Manganese peroxidase (MnP) catalytic cycle

characteristic blue colour. The enzymes lacking a blue copper atom are called yellow or white laccases (Diamantidis et  al. 2000). Typically laccase-mediated catalysis occurs with reduction of oxygen to water accompanied by the oxidation of substrate. Laccases catalyse the oxidation of a broad range of substrates such as ortho and paradiphenols, methoxy-substituted phenols, aromatic amines, phenolic acids and several other compounds coupled to the reduction of molecular oxygen to water with one-electron oxidation mechanism (Morozova et al. 2007). Laccase is most widely distributed in a wide range of higher plants, fungi and bacteria. Laccases are secreted out in the medium extracellulary by several fungi during the secondary metabolism but not all fungal species produce laccase such as Zygomycetes and Chytridiomycetes. Fungi belonging to Deuteromycetes, Ascomycetes as well as Basidiomycetes are known producers of laccase (Sadhasivam et al. 2008). Laccase is currently the focus of much attention because of its diverse applications such as dye decolourization, waste detoxifications and bioremediation applications (Fig. 4.5).

4.6  F  actors Influencing Enzyme Production and Dye Degradation Secretion and synthesis of these enzymes are often stimulated by limited levels of nutrients such as nitrogen or carbon sources. Production of manganese peroxide and lignin peroxide is generally induced by agitation in submerged white rot fungi liquid culture, while laccase production is frequently enhanced by agitation. These features are considered as important roles in the process design and optimization of fungal treatment of colour-containing effluents (Wesenberg et al. 2003). While several studies were devoted to bio-decolourization of the textile azo dyes, far less

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Fig. 4.5  Laccase catalytic cycle

attention has been paid to synthetic dye bath effluent in which the presence of salts and high dye concentration may be inhibitory to biological agents (Faraco et  al. 2009; Gomaa et al. 2008). Different WRF and their enzymes involving in dye degradation are given in Table 4.5.

4.6.1  Effect of Inducers on Enzyme Production Laccase production of white rot fungi mainly depended on the condition of fungal cultivation and media supporting. Lignolytic systems of WRF were mainly stimulated during the secondary metabolic phase and were prompted by nitrogen concentration or when sulphur or carbon became limiting. Fungi produced lower concentration of laccase, but higher concentrations can be attained with the addition of supplements to media like veratryl alcohol, 2,3xylidine and lignin. Response surface methodology (RSM) was applied to optimize the decolouration of the diazo dye Reactive Black 5 (RB5) by crude laccase from the white rot fungus Trametes pubescens (Roriz et al. 2009). Laccase production of white rot fungus (Pycnoporus sanguineus) and its growth in a bubble column reactor were studied. Different inducers like copper sulphate, ethanol, saw dust and superficial gas velocities are used to increase the growth and laccase production (Karim and Annuar 2009).

4.6.2  Effect of Nitrogen Source The laccase production was influenced by the sources of nitrogen used in the media. Peralta-Zamora et al. (2003) isolated four WRF, and Lentinus edodes displayed the greatest decolourization ability both in terms of extent and rapidity of decolourization. The dyes used were Reactive Red 195 (0.025%), Reactive Blue 19 (0.05%),

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Table 4.5  White rot fugal species and their enzymes involving in dye degradation Organisms Dichomitus squalens

Enzymes Manganese-­ peroxidase

Bjerkandera adusta

Lignin peroxidase

Pleurotus eryngii

Irpex lacteus

Lignin peroxidase Manganese-­ peroxidase versatile peroxidases (VP) Laccase

Phanerochaete chrysosporium

Manganese-­ peroxidase

Irpex lacteus

Lignin peroxidase

Phanerochaete Chrysosporium

Lignin peroxidase

Phlebia (Merulius) Tremellosa

Lignin peroxidase

Bjerkandera adusta Pleurotus and Bjerkandera

Dye Cresol RedTPM, Brilliant Green (TPM), Crystal Violet (TPM), OrangeII (N=N), Congo Red (N=N) Reactive Violet 5 (N=N), Reactive Orange 96 (N=N), Reactive Black 5 (N=N), Reactive Blue 38 (PC) Reactive Blue 15 (PC) and Remazol Brilliant Blue RPAQ, Poly R-478 (PAQ) Reactive Violet 5 (N=N), Reactive Blue 38PC by MnP Reactive Black 5 (N=N) Reactive Black 5 (N=N), Reactive Violet 5 (N=N), Reactive, Blue 38PC Dyes

Naphtol Blue Black (N=N), Methyl Red (N=N), Congo Red (N=N), Remazol Brilliant Blue R (PAQ), Copper (II) phthalocyaninetetrasulphonic acid Tetrasodium salt (MC), Bromophenol Blue (TPM), Poly R-478 (PAQ) Azure Blue, Azo dyes, Cresol Red (TPM), Bromophenol Blue (TPM), Crystal Violet (TPM) Methyl Red (N=N), Naphtol Blue Black (N=N), Congo Red (N=N), Bromophenol Blue (TPM), Remazol Brilliant Blue R (PAQ), Copper (II) phthalocyaninetetra sulphonic acid tetrasodium salt(MC) and Poly R-478(PAQ) Azo dyes, Azure Blue, Cresol Red (TPM), Crystal Violet (TPM), Bromophenol Blue (TPM) Cibacron Red, Remazol Red, Remazol Navy Blue Cibacron Orange, Remazol Golden Yellow, Remazol Blue, Remazol Black B, Remazol Turquoise Blue

Reference Périé and Gold (1991) Kaal et al. (1995)

Heinfling et al. (1998) Heinfling et al. (1998) Mester and Field (1998) Novotný et al. (2000)

Cameron et al. (2000) Novotný et al. (2000)

Cameron et al. (2000) Kirby et al. (2000)

(continued)

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Table 4.5 (continued) Organisms Irpex lacteus

Enzymes Manganese peroxidase

Trametes(Coriolus) Versicolor

Lignin peroxidase

Pleurotus ostreatus, Schizophyllum commune, Sclerotium rolfsii, Neurospora crassa Phellinus gilus, Pleurotus sajor-­ caju, Pycnoporus sanguineus, Phanerochaete chrysosporium Phanerochaete chrysosporium Lentinula (Lentinus) edodes Trametes versicolor

LiP and/or MnP in addition to Lac

T. trogii

Reference Novotný et al. (2000)

Novotný et al. (2000)

Abadulla et al. (2000)

Lignin peroxidase

Vat textile dyes

Balan and Monteiro (2001)

Manganese-­ peroxidase Manganese-­ peroxidase Lignin peroxidase Laccase

Amaranth dye, Orange G dye, New coccine dye, Tartrazine dye Remazole Brilliant Blue R

Chagas and Durrant (2001) Boer et al. (2004) Christian et al. (2005) Zouari-­ Mechichi et al. (2006) Ben Younes et al. (2007) Urek and Pazarlioglu (2007) Asgher et al. (2008) Michniewicz et al. (2008) Moreira-Neto et al. (2013) Rita de Cássia et al. (2013)

Laccase

Perenniporia tephropora Phanerochaete chrysosporium

Manganese-­ peroxidase

Scyzophyllum commune Cerrena unicolor

Manganese-­ peroxidase Laccase

Bjerkandera sp.

Manganese-­ peroxidase Laccase

Trametes versicolor

Dye Congo Red (N=N), Methyl red (N=N), Naphtol Blue Black (N=N), Bromophenol Blue (TPM), Remazol Brilliant Blue R (PAQ), Copper (II) phthalocyaninetetrasulphonic acid Tetrasodium salt (MC), Poly R-478 (PAQ) Remazol Brilliant Blue Everzol Turquoise Blue GPC, R(PAQ), Poly R-478(PAQ), Everzol Red RBN, Everzol Yellow 4GL, Everdirect Supra Yellow (PG), Orange K-GL Triarylmethane, Anthraquinonic, and Indigoid textile dyes

Remazol Brilliant Blue R Azo and triarylmethane dyes

Neolane pink, neolane blue, and remazol brilliant blue R (RBBR) Direct Green 6, Direct Blue 15, Congo Red Solar Golden Yellow R Acid Blue 62, Acid Blue 40, Reactive Blue 81, Direct Black 22, Acid Red 27 Reactive Blue 38PC, Orange IIN = N, Poly R-478PAQ Indigo dye

(continued)

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Table 4.5 (continued) Organisms Lentinus tigrinus Lentinus tigrinus Phanerochaete chrysosporium

Enzymes Manganese-­ peroxidase Laccase Laccase

Dye Reactive Blue 38(PC), Orange II (N=N), Poly R-478(PAQ) Reactive Blue 38(PC), Orange II (N=N), Poly R-478(PAQ) Red 2BAB New coccine, Amaranth, Orang G, Tartrazine Congo Red (N=N)

Pleurotus ostreatus, Laccase Pleurotus sapidus, Pleurotus florida

Coralene Golden Yellow, Coralene Navy Blue and Coralene Dark Red azo dyes

Reference Moreira-Neto et al. (2013) Moreira-Neto et al. (2013) Jain et al. (2000) Chagas and Durrant (2001) Gill et al. (2002) Kunjadia et al. (2016)

Reactive Black 5 (0.05%) and Reactive Yellow 145 (0.05%). The colour removal by fungal hyphae is mainly by the mechanism of degradation by extracellular and intracellular enzymes (Chagas and Durrant 2001). In a nitrogen-deficient mineral salts medium, Robinson et  al. (2001) studied four WRF, Bjerkandera adusta, Phlebia tremellosa, Pleurotus ostreatus and Coriolus versicolor, for testing their ability to produce manganese peroxidase (MnP), lignin peroxidase (LiP) and laccase. B. adusta and P. tremellosa were selected, on the basis of their high enzyme production potential, for the degradation of five dyes in an artificial textile effluent. In N-rich (C:N ratio, 11.6:1 and N-limited, 116:1) conditions, degradation experiments were carried out to examine degradation potential. At a dye concentration of 100  mg/litre, in N-rich media, P. tremellosa degraded 79% in 9  days, B. adusta degraded 85% of the dyes in 7  days, and in N-limited conditions, 86% of the effluent was degraded in 9 days by B. adusta and 74% by P. tremellosa in 11 days. The results revealed that addition of nitrogen had no significant effect on dye degradation percentage by B. adusta, with a slight increase for P. tremellosa and nitrogen supplementation reducing the decolourization time. In nitrogen-limited glucose ammonium media, Phanerochaete sordida decolourized dye mixtures (reactive textile dyes, including azo and anthraquinone dyes 200 mg/L each) within 48 h by 90%. Manganese peroxidase (MnP) was engaged in dye decolourization by P. sordida. Wastewaters from textile industries often contain polyvinyl alcohol (PVA) which inhibited MnP reaction system and decolourization (Reactive Red 120) potential of Phanerochaete sordida. Addition of Tween 80 to the mixtures in the occurrence of PVA improved the decolourization of RR120 which showed that PVA could hinder with lipid peroxidation or consequent attack to the dye (Harazono and Nakamura 2005). D’Souza-Ticlo et al. (2006) reported the effect of KNO3, glycine, glutamic acid, corn steep liquor and beef extract under stationary conditions. The cultures were oxygenated every third day with pure oxygen for 1 min using Pasture pipettes and Tygon tubing under sterile conditions. They obtained when glutamic acid was used as nitrogen source.

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In majority of the cases, nitrogen-limited medium increased the production of lignin peroxide and manganese peroxide, thereby increasing the rate of decolourization (Singhal and Rathore 2001). These enzymes are produced by WRF during their metabolism. Subsequently, lignin oxidation catalyses the degradation/transformation of aromatic dyes either by precipitation or by opening the aromatic complex ring structure, and therefore no energy source is required to the fungus (Husain 2010).

4.6.3  Effect of Carbon Source Blánquez et  al. (2008) showed that T. versicolor was able to continuously decolourize a spent dyeing bath from a textile factory under non-sterilized conditions for 15 days in a 10-L air-pulsed bioreactor, attaining colour reduction levels between 40 and 60%. Nutrients were added at the start-up period (3 days) of the experiment, and thereafter sterilized glucose was added. Therefore, although the author stated that T. versicolor was able to treat successfully real industrial wastewater in continuous mode (HRT 48 h), the operation time was very short (15 days), and, in addition to this, bacterial contamination in the feeding tank was detected from day 10. The ability of C. versicolor to decolourize the textile effluents collected from 5 different textile industries was tested. However, addition of starch (1% w/v) as a carbon source was required to obtain significant decolouration levels (84% in 3 days) (Asgher et al. 2008). Phenol-degrading white rot fungus T. versicolor was isolated from paper mill. 14C synthetic lignin mineralization assay showed that it assimilated 24.3% of total label. During 5  days of incubation period, 71% of p-hydroxybenzoic acid was utilized when glucose was used as a co-substrate and 56% degradation of protocatechoic acid using fructose (Udayasoorian and Prabu 2005).

4.6.4  Influence of pH and Temperature Azo dye (RB5) wastewater is usually neutral to alkaline. However, though fungi grow optimally under weakly acidic conditions, the azo dyes were degraded even in a slightly alkaline pH range (pH 7–8). P. sordida PBU 0057 yielded 100% decolouration in 72 hours over the pH range 6–8, while P. chrysosporium reached the maximum (96%) by 96  hours, though lesser (93%) at pH  8 (Forgacs et  al. 2004). Selvakumar et al. (2013) studied treated textile wastewater in a batch reactor with Ganoderma lucidum. Under optimized conditions (pH  6.6; temperature 26.5  °C; dye wastewater concentration 1:2; agitation speed 200 rpm), a maximum decolouration of 81.4% and a COD reduction of 90.3% were found. Therefore, dilution and decreased pH of the original effluent were necessary for maximum decolouration and COD reduction.

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Hadibarata et al. (2013) assessed the effect of temperature on decolouration at 25, 30, and 35 °C. Complete RB5 decolouration by P. sordida PBU 0057 occurred by 72 hours at 30 and 35 °C but was delayed until 96 hours at 25 °C. Overall, the tropical isolates have the advantage of being active at higher temperatures – a practical attribute in tropical wastewater treatment facilities.

4.6.5  Concentration of Dye Anastasi et  al. (2010) reported the decolouration of wastewater from a dyeing-­ textile factory by the white rot fungi Bjerkandera adusta packed in a fixed-bed bioreactor. The fungus remains effective during 4 cycles of decolouration for a very long period (70 days) under non-sterile conditions and with no nutrient addition. Osorio Echavarría et  al. (2011) reported the decolouration of wastewater from a textile industry by the anamorph R1 of the white rot fungus Bjerkandera sp. Under sterile conditions, the effluent was decolourized by 65% in 8 days, and its toxicity was reduced by 58%, whereas under non-sterile conditions, the decolouration percentage was only 40% for the same time period. The authors stated that this lower decolouration value was likely due to the presence of contaminant microorganisms competing for the substrate. Ma (2014) revealed that the Ganoderma sp. En3 had a strong potential and tolerance to decolourize and detoxify high concentrations of sulphonated azo dye Reactive Orange 16 containing textile wastewater under in  vitro conditions. Maximum decolourization of 98% was achieved in synthetic dye bath effluent on the third day under normal conditions by white rot fungus Phanerochaete chrysosporium. Experiment with different concentrations of effluent found that the increasing in effluent concentration slows down the decolourization percentage and optimized amounts of nutrients were found to be 0.5%, 0.1% and 0.5% of glucose, manganese sulphate and ammonium salts, respectively. Adding inducers such as lignin and starch augmented enzyme activity and the rate of decolourization (Senthilkumar et al. 2014).

4.7  White Rot Fungi and Decolouration Previous studies reported the ability of P. chrysosporium, Bjerkandera sp. and T. versicolor to decolourize Remazol Orange, Reactive Blue, Remazol Brilliant Blue and Tropaeolin O in agar plates. A basidiomycete fungus, P. chrysosporium, was able to degrade the starch, cellulose, pectin, lignin and lignocelluloses complex compounds, which are characteristics of textile dye effluent (Swamy and Ramsay 1999). Consequently, some strains including S. thermophilum and T. trogii were reported to be able to decolourize and detoxify textile effluents. Phlebia tremellosa and B. adusta showed a good efficiency to decolourize textile effluent in N-limited conditions (Robinson et al. 2001). Tekere et al.

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(2001a) reported the ability of T. versicolor, Trametes cingulata, Pycnoporus sanguineus and Datronia concentrica to decolourize Poly R478 (Tekere et al. 2001b). Conneely et al. (2002) found that, Remazol Turquoise Blue G133, Heligon Blue S4, phthalocyanine dyes and Everzol Turquoise Blue are biosorbed by Phanerochaete chrysosporium and also metabolized by its ligninolytic extracellular enzymes resulting in the formation of free copper ions, dye decolourization and organic metabolites with ultimate extensive phthalocyanine ring breakdown. It is assumed that the ligninolytic extracellular enzyme laccase is involved in the initial production of a metabolite M8 which involves break-up of the conjugated phthalocyanine ring structure but which retains multi-negative charge. Another ligninolytic extracellular enzyme, manganese peroxidase, is assumed to be involved in the release of Cu2+ from the phthalocyanine structure to give a non-copper-containing phthalocyanine metabolite M1 with a slightly longer migration time than the parent dye and absorption at 666 nm. The phthalocyanine ring structure is also broken up by metabolic processes that involve oxidation and desulphonation to give phthalimide (M3) and an unidentified electroactive metabolite M2. WRF are most efficient in degrading synthetic azodyes. This property is due to the production of extracellular LMEs, which, because of the lower substrate specificity, are able to degrade a wide range of xenobiotic compounds (Wesenberg et al. 2003). Mohorčič et  al. (2006) found that Bjerkandera adusta was able to decolourize the black-blue dye through red and violet to pale yellow via its extracellular enzyme; MnP was also reported for its ability to decolourize Remazol Black B and amaranth. Ben Younes et al. (2007) and Zouari-Mechichi et al. (2006) reported that the crude enzyme as well as the purified laccase from Perenniporia tephropora was able to decolourize dyes of the textile waste water, including Neolane Blue, Neolane Pink and Remazol Brilliant Blue R (RBBR). The latter was also efficiently decolourized by laccase from T. trogii. The ability of T. trogii laccase to decolourize triarylmethane and azo dyes was approved in the absence of redox mediators, since MG and BCG were completely degraded with crude laccase within 6 h of treatment. This fungus appears to be the best choice, and it has the potential to degrade and can be very useful in the treatment and disposal of textile and related effluents. P. chrysosporium can also be used to decolourize wastewater containing dyes with complex structures directly (Kaushik and Malik 2009). Manavalan et al. (2013) have first-time reported that Ganoderma lucidum laccase enzyme production using medium with 3% (v/v) ethanol improved the enzyme production up to 14.1 folds. Moreira-Neto et  al. (2013) studied on twelve Basidiomycetes strains from the genus Pleurotus, Trametes, Lentinus, Peniophora, Pycnoporus, Rigidoporus, Hygrocybe and Psilocybe on decolourization of the reactive dyes Cibacron Brilliant Blue H-GR and Cibacron Red FN-2BL, both in solid and liquid media. Among the evaluated fungi, seven showed great ability to decolourize the synthetic textile effluent, both in vivo (74–77%) and in vitro (60– 74%), and laccase was the main ligninolytic enzyme involved on dyes decolourization. Pleurotus ostreatus, Trametes villosa and Peniophora cinerea reduced near to 60% of the effluent colour after 1  h of treatment  and  the decolourization results were still improved by establishing the nitrogen source and amount to be used during the fungal strains cultivation in synthetic medium. For example their action on the

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textile effluent, with yeast extract being a better nitrogen source than ammonium tartarate. Rita de Cássia et al. (2013) study reported the LiP, Lac and MnP activities of Curvularia lunata URM 6179 and Phanerochaete chrysosporium URM 6181 at the end of textile effluent treatment in aerated and non-aerated bioreactors. Both fungi showed higher activity of laccase but under different conditions: 2020 U/l for the P. chrysosporium URM 6181  in aerated bioreactor and 2100  U/l for the Curvularia lunata non-aerated bioreactor. Low LiP activity was observed for both fungi strains. The activity of MnP was more evidenced in treatment with C. lunata URM 6179. Activities of 7000  U/l for MnP and 8000  U/l for LiP in treatment employing Lentinus strains, and 2000 U/l for Lac in treatment employing Coriolopsis byrsina were found. The higher production of MnP and Lac was found in culture medium containing wheat T. harzianum bran and glucose (Gomes et al. 2009). The fungus Mucor racemosus CBMAI 847 produces 898.15 U/l of laccase in medium containing 23% salinity and 4.5 mg/ml wheat bran while Cladosporium cladosporioides CBMAI 857 produces 4.63 U/l. El Monssef et al. (2016) study observed that the production of laccase enzyme was higher at 35  °C and pH  5 after 6  days. The highest activity of laccase was achieved at 35  °C and pH  5 during the reaction. FTIR analysis revealed that the structure of extracted fungal pigments has aromatic ring and phenols group. Crude laccase was capable to decolourize different pigment structures. The enzyme showed great decolourization efficiency towards the extracted yellow pigment produced from Asp. terrus and Asp. ochareceous treated by 200 μl of partially purified enzyme. Toxicity evaluation showed a final product detoxification (Ellouze and Sayadi 2016). On the other hand, the fungal decolourization of RBBR has been reported for other strains such as Ischnoderma resinosum, Dichomitus squalens, P. ostreatus and Pleurotus calyptratus. Kunjadia et al. (2016) studied the role of ligninolytic enzymes of Pleurotus spp. grown with azo dyes. The results indicated that, WRF P. sapidus, P. ostreatus and P. florida were tested for ligninolytic enzyme activity and their role in dye degradation. Percentage of decolourization clearly showed higher removal of Coralene Golden Yellow (CGY) by P. ostreatus. Laccase activity in cultures during dye decolourization was significantly higher compared to MnP, suggesting the important role of laccase in dye degradation process. Meanwhile, no LiP activity was found in any of the cultures. It is clear that enzymes such as LiP, MnP and laccase play an important role in dye metabolism by WRF. P. ostreatusis is the best fungal species out of all three studied organisms for ligninolytic activity and degradation of azo dyes.

4.8  W  hite Rot Fungi and Decolourization of Industrial Dye Effluents Reductions in the oxygen demand and carbon content of azo dye wastewaters, subsequently treated with aerobic conditions, are well cited in previous studies (Horning 1977; Loyd 1992; McCurdy 1991). Due to the toxicity and xenobiotic nature,

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degradation of azo dyes is indeed an uphill task. Now it had been well documented that azo dyes can be degraded by microorganisms. Due to their recalcitrant nature, azo dyes often pass through activated sludge facilities with no or little reduction in colour (Cariell et al. 1995); however, some researchers observed slight colour reductions in their findings (Zissi and Lyberatos 2001). Effluents containing dyes are hardly decolourized by conventional and biological wastewater treatments (Shaul et al. 1991; Willmott et al. 1998). In addition to their adverse impact and their visual effect, many synthetic dyes are toxic with high chemical oxygen demand, carcinogenic and mutagenic properties (Chung et al. 1978; Michaels and Lewis 1985). Also, the higher volumetric discharge of industrial effluent in combination with severe legislation makes the searching of suitable treatment technologies an important priority (O'neill et  al. 2000). Reduction of azo and other dyes with chemical and physical methods requires highly expensive reagents and catalysts (Robinson et al. 2001). Usage of dyes and pigments are enormous in the paper, plastic, textile, cosmetics, pharmaceutical and food industries (Levin et al. 2004). Based on toxicity and carcinogenic nature of dyes and pigments, biodegradation of these synthetic dyes may involve (Revankar and Lele 2006) WRF which are better dye-degraders than prokaryotes. Extracellular non-specific LME system present in WRF plays a major role in degrading a wide range of dyes (Christian et al. 2005). Most of the former dye decolourization experiments were based mainly on Phanerochaete chrysosporium and Trametes versicolor (Toh et al. 2003). However, other WRF including Phellinus gilvus, Pleurotus sajor-caju, Pycnoporus sanguineus (Balan and Monteiro 2001), Dichomitus squalens, Irpex flavus, Daedalea flavida, Polyporus sanguineus (Chander 2007; Eichlerova et  al. 2006; Gill et  al. 2002), Funalia trogii ATCC200800 (Özsoy et  al. 2005), Ischnoderma resinosum (Eichlerova et al. 2006) and Ganoderma sp. WR-1 (Revankar and Lele 2006) have been experimented to have higher dye decolourization rates than P. chrysosporium and T. versicolor. At present, much attention has been concentrated on fungal decolourization processes, especially on WRF due to their capacity of production of non-specific enzymes, such as lignin peroxidase, manganese peroxidase and laccase, which act as sorbents and detoxify many toxic aromatic compounds. Several research findings indicate that WRF and their biodegradation/biotransformation capacity could be an excellent candidate for dye removal (Gomaa et al. 2008; Rita de Cássia et al. 2013; Wesenberg et al. 2003). Among the different groups of fungi, Phanerochaete chrysosporium is an effective dye-degrading microorganism. P. chrysosporium has become known as a model system in textile, pulp and paper mill wastewater bioremediation. P. chrysosporium is a basidiomycete fungus able to detoxify complex compounds such as starch, cellulose, pectin, lignin and lignocelluloses in textile dye wastewater (Senthilkumar et al. 2014). This fungus emerges to be the best choice to degrade complex wastewater in the treatment process and disposal of textile and related wastewater.

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4.9  Conclusion and Future Prospects Much research on dye degradation has been reported by many scientists with individual dyes or under simulated condition by WRF, but under dye industry, wastewater treatment was reported only in limited extent. Application of WRF at industrial scale under non-sterilized condition is a big technical challenge. In addition, the chemistry of intermediates formed during the degradation was also in its infant stage. Bioassay studies of treated dye wastewater should also be established. Aquatic fungi and their ability to produce several non-specific enzymes may serve as a new resource to treat textile industry wastewater. Presence of degradative plasmid encoding for such environmentally significant genotype of azo dye degradation opens up the possibilities of genetic transfer of this character into other microorganisms, which can act as a valuable tool in the remediation of azo dye-polluted habitat. Increasing the contribution of small and medium enterprises of textile, chemical and pharmaceutical in total exports of India is vital to India’s future economic growth. Detailed scientific studies with natural dyes have established that in most cases their properties are comparable to those of synthetic dyes. Therefore, if natural dyes have to be commercialized, they need to conform to the same stringent standards of performance that are applied to synthetic dyes. It thus follows that much more research and developmental effort needs to go in this area. Acknowledgement  The authors are grateful to the Department of Environmental Sciences, Tamil Nadu Agricultural University, Coimbatore, for providing laboratory facilities. The authors extend their gratitude to Mrs. P. Divya and Mr. P. Sivasamy for providing textile wastes-related data.

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Tekere M, Zvauya R, Read JS (2001b) Ligninolytic enzyme production in selected sub-tropical white rot fungi under different culture conditions. J Basic Microbiol 41:115–129 Teli M (2008) Textile coloration industry in India. Color Technol 124(1):1–13 Toh YC, Yen JJL, Obbard JP, Ting YP (2003) Decolourisation of azo dyes by white-rot fungi (WRF) isolated in Singapore. Enzym Microb Technol 33:569–575 Tortella GR, Rubilar O, Gianfreda L, Valenzuela E, Diez MC (2008) Enzymatic characterization of Chilean native wood-rotting fungi for potential use in the bioremediation of polluted environments with chlorophenols. World J Microbiol Biotechnol 24:2805 Udayasoorian C, Prabu P (2005) Biodegradation of phenols by ligninolytic fungus Trametes versicolor. J Biol Sci 5:558–561 Urek RO, Pazarlioglu NK (2007) Enhanced production of manganese peroxidase by Phanerochaete chrysosporium. Braz Arch Biol Technol 50:913–920 Vaidya A (1982) Environmental pollution during chemical processing of synthetic fibers. Colourage 14:3–10 Vasina DV, Moiseenko KV, Fedorova TV, Tyazhelova TV (2017) Lignin-degrading peroxidases in white-rot fungus Trametes hirsuta 072. Absolute expression quantification of full multigene family. PLoS One 12:0173813 Velázquez-Cedeño M, Farnet A, Ferré E, Savoie J (2004) Variations of lignocellulosic activities in dual cultures of Pleurotus ostreatus and Trichoderma longibrachiatum on unsterilized wheat straw. Mycologia 96:712–719 Vijaya P, Padmavathy P, Sandhya S (2003) Decolourization and biodegradation of reactive azo dyes by mixed culture. Indian J Biotechnol 2:259–263 Viswanath B, Chandra MS, Pallavi H, Reddy BR (2008) Screening and assessment of laccase producing fungi isolated from different environmental samples. Afr J Biotechnol 7(8):1129–1133 Vyas B, Molitoris H-P (1995) Involvement of an extracellular H2O2-dependent ligninolytic activity of the white rot fungus Pleurotus ostreatus in the decolorization of Remazol brilliant blue R. Appl Environ Microbiol 61:3919–3927 Wesenberg D, Kyriakides I, Agathos SN (2003) White-rot fungi and their enzymes for the treatment of industrial dye effluents. Biotechnol Adv 22:161–187 Will R, Ishikawa Y, Leder A (2000) Synthetic dyes, chemical economics handbook: synthetic dyes. SRI Chemical & Health Business Services, Menlo Park Willmott N, Guthrie J, Nelson G (1998) The biotechnology approach to colour removal from textile effluent. J Soc Dye Colour 114:38–41 Yadav AN, Sachan SG, Verma P, Kaushik R, Saxena AK (2016) Cold active hydrolytic enzymes production by psychrotrophic Bacilli isolated from three sub-glacial lakes of NW Indian Himalayas. J Basic Microbiol 56:294–307 Yadav A, Verma P, Kumar R, Kumar V, Kumar K (2017) Current applications and future prospects of eco-friendly microbes. EU Voice 3:21–22 Yadav AN, Verma P, Kumar V, Sangwan P, Mishra S, Panjiar N, Gupta VK, Saxena AK (2018) Biodiversity of the Genus Penicillium in different habitats. In: Gupta VK, Rodriguez-Couto S (eds) New and future developments in microbial biotechnology and bioengineering, Penicillium system properties and applications. Elsevier, Amsterdam, pp  3–18. https://doi.org/10.1016/ B978-0-444-63501-3.00001-6 Yadav AN, Mishra S, Singh S, Gupta A (2019a) Recent advancement in white biotechnology through fungi volume 1: diversity and enzymes perspectives. Springer International Publishing, Cham Yadav AN, Mishra S, Singh S, Gupta A (2019b) Recent advancement in white biotechnology through fungi. Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham Zissi U, Lyberatos G (2001) Partial degradation of p-aminoazobenzene by a defined mixed culture of Bacillus subtilis and Stenotrophomonas maltophilia. Biotechnol Bioeng 72:49–54

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Zollinger H (1961) Azo and diazo chemistry: aliphatic and aromatic compounds. Interscience Publishers, New York, p 444 Zollinger H (2003) Color chemistry: syntheses, properties, and applications of organic dyes and pigments. John Wiley & Sons Zouari-Mechichi H, Mechichi T, Dhouib A, Sayadi S, Martinez AT, Martinez MJ (2006) Laccase purification and characterization from Trametes trogii isolated in Tunisia: decolorization of textile dyes by the purified enzyme. Enzym Microb Technol 39:141–148

Chapter 5

Pleurotus ostreatus: A Biofactory for Lignin-Degrading Enzymes of Diverse Industrial Applications Hesham El Enshasy, Farid Agouillal, Zarani Mat, Roslinda Abd Malek, Siti Zulaiha Hanapi, Ong Mei Leng, Daniel Joe Dailin, and Dalia Sukmawati

5.1  Introduction Mushrooms have been recognized as important food and medicine in many ancient civilizations (El Enshasy et al. 2013). These are based on their high nutrient contents of carbohydrates, proteins, vitamins, minerals, and many other growth-promoting ingredients (Eleftherios et  al. 2014; Maftoun et  al. 2015). Nowadays, of thousands types of mushroom studied, Pleurotus sp. is considered as one of the top three economically important and widely grown mushrooms beside Agaricus and Lentinula. Pleurotus ostreatus (widely known as oyster mushroom) gained more interest in the recent years not only because of their high nutritional value but also due to the high content of bioactive polysaccharides and other metabolites of high medicinal values. These compounds exhibited immunomodulatory, antitumor, antioxidant, anti-inflammatory, antihyperglycemic, antihypocholesterolemic,

H. El Enshasy (*) Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia School of Chemical and Energy Engineering, Faculty of Engineering, Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia City of Scientific Research and Technology Applications, New Burg Al Arab, Alexandria, Egypt e-mail: [email protected] F. Agouillal Research Unit on Analysis and Technological Development in Environment (URADTE), Centre de Recherche Scientifique et Technique en Analyses Physico-Chimiques (CRAPC), Tipaza, Algeria Z. Mat · R. A. Malek · S. Z. Hanapi Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_5

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antimicrobial, and antithrombotic properties (El Enshasy et al. 2012; El Enshasy and Hatti-Kaul 2013; Mohamed and Farghaly 2014; Correa et al. 2016). The therapeutic effect of oyster mushroom extract and pure bioactive compounds has been studied and proved in many in vivo and in vitro studies (Ryu et al. 2014; Elsayed et al. 2014; Younis et al. 2015; Masri et al. 2017). In addition, P. ostreatus gained more interest than other types of mushrooms based on their ability to grow on large number of substrates and under different environmental conditions (tropical and subtropical region), high capacity to produce a large number of hydrolases, and ease of cultivation both in solid-state fermentation and in submerged cultivation system as well as higher growth rate compared to other types of mushrooms (El Enshasy et  al. 2010; Maftoun et  al. 2013). However, to survive in nature, mushrooms should be able to degrade complex lignocellulosic materials by different hydrolytic enzymes to break the complex structure of wood and utilize the produced sugars as substrate for growth and metabolite production. Therefore, mushrooms are well known for their high capacity to produce and excrete different types of enzymes of wide range of hydrolytic activities. Therefore, this chapter will provide the latest information about the enzyme systems of this type of mushroom and their potential application in different industries.

5.2  Understanding Lignin Structure Lignins constitute with cellulose and hemicelluloses the three major components of lignocellulosic biomass. They are the second most abundant terrestrial polymers and carbon source after cellulose (Evers et al. 1999; Boerjan et al. 2003). In 1813, A. P. de Candolle has firstly named the substance as “lignine” based on the Latin word lignum which means “wood.” He described lignin as a fibrous, tasteless material, insoluble in water and alcohol, but soluble in weak alkaline solutions and which can be precipitated from solution using acid. The typical composition of the dry weight of wood, considered as a whole, is about 50% cellulose, 25% lignin, 20–25% hemicellulose, and 1–4% pectin (Camarero et al. 2014).

O. M. Leng Harita Go Green Sdn. Bhd., Johor Bahru, Johor, Malaysia D. J. Dailin Institute of Bioproduct Development (IBD), Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia School of Chemical and Energy Engineering, Faculty of Engineering, Universiti Teknologi Malaysia (UTM), Johor Bahru, Malaysia D. Sukmawati Faculty of Mathematics and Natural Sciences, Universitas Negeri Jakarta, Jakarta, Indonesia

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Cellulose forms basically a skeleton that is bounded by hemicelluloses and lignin (Sakakibari 1980). Lignin, a polyphenolic amorphous polymer, is the essential natural glue that fills the spaces between cellulose and hemicellulose and acts like a resin that holds the plants’ lignocellulose matrix together (Ritter 2008; Carmen 2009). Through this cross-linking with cellulose, lignin not only confers the strength, rigidity, and flexibility as well as aids in water transport to the plant but represents also the most significant barrier to wood decay by insects/microorganisms attacks and prevents the access of low molecular weight diffusible agents (Zakzeski et al. 2010). In chemical point of view, lignin is a heterogeneous complex class of compounds that changes according to biomass source and isolation technique (Johnson 2002). However, many aspects in the chemistry of lignin still remain unclear due to its high complex structure (Pouteau et al. 2003); However, the lignin classification as grass, hardwood, and softwood is based on the ratios of three major phenylpropene monomers which vary according to the plant species, plant tissue, individual cell types, and cell wall layers (Faix 1991). In general, both native lignin (as present in biomass) and technical lignin (isolated from biomass through various processes) have threedimensional amorphous polymer made up of methoxylated phenylpropane aromatic units structures of p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol (Chakar and Ragauskas 2004). These phenylpropene monomers are randomly cross-linked polymers arising from an enzyme-mediated dehydrogenative polymerization of three phenylpropane monomer precursors (cinnamyl alcohol: coniferyl, synapyl, and p-coumaryl alcohols) (Kleinert and Barth 2008; Wong 2009). These three monolignols (phenylpropene units of lignins) shown in Fig. 5.1 are as follows:

Fig. 5.1  The chemical structure of the primary lignin building blocks and their corresponding lignin polymer monomeric units (Zakzeski et al. 2010; Abdel-Hamid et al. 2013)

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• Guaiacyl (G) units from the precursor trans-coniferyl-alcohol • Syringyl (S) units from trans-sinapyl-alcohol • p-Hydroxyphenyl (H) units from the precursor trans-p-coumaryl alcohol Then, grass lignin is built up by the three monomeric units (G, S, and H), hardwood lignin contains roughly equal amounts of (G) and (S) units, and softwood lignin is composed mainly of (G) units (Faix 1991).

5.2.1  Lignocellulose Processing and Industrial Applications For more economic profitability, lignocellulosic biorefinery produces multiple products, including fuels, and bulk or fine chemicals, from biomass, analogically compared to a petroleum refinery, which produces fuels and chemicals from crude oil (Zhang 2008; Rastegari et al. 2019). In order to achieve a high energy impact and an economically viable biorefinery, the valorization of all components of lignocellulosic biomass should be carried out (Abdel-Hamid et al. 2013). With various aromatic structures, many authors claim that lignin has a special industrial interest offering the feasibility of replacing relevant aromatic polymeric and fine chemicals (Duval and Lawoko 2014; Norgren and Edlund 2014; Laurichesse and Avérous 2014; Thakur and Thakur 2015; Thakur et al. 2014). Lignins are usually considered as waste products of pulp and paper industry and had limited industrial uses. However, in 1998, only 1% of the lignin produced was used in valuable industrial processes. About 50 million tons of lignin were estimated to be extracted annually from woody biomass for the pulp and paper industry and applied as dispersants, adhesives, and surfactants (Cohen et al. 2002; Shah and Nervd 2002; Karam and Nicell 1997). However, new technologies are currently in use to valorization and converting lignin into value-added chemicals; then, a biomass fractionation technology has been developed by PureVision Technology, Inc., to produce value-added low molecular weight lignin as a coproduct to the cellulose stream and also as fuel substrate (Chheda et al. 2007). In energy production process, the lignin fraction acts as a barrier against enzyme or microbial penetration through lignocelluloses decreasing the fermentable sugar yields and affecting negatively the overall biofuel development, making it an uneconomical process (Margeot et al. 2009; Menon and Rao 2012). Essential mechanical, thermomechanical, and thermochemical pretreatment strategies to overcome this limitation have been reported by many authors (Margeot et  al. 2009; Frigon and Guiot 2010; Kumar et al. 2009) and could be summarized as follows: • Mechanical particle size reduction including both of wet, dry, ball, or vibratory ball milling and other forms of biomass grinding (Sarkar et al. 2012; Agbor et al. 2011). • Thermochemical hydrolysis at temperatures between 140 °C and 180 °C, using a dilute solution of sulfuric acid (0.5–2%) for 10–30 minutes as residence times for rendering the carbohydrate fraction (Yang and Lu 2010).

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• Thermomechanical steam explosion heating briefly the biomass to high temperatures (~200 °C) under high pressure followed by a rapid pressure drop that makes the biomass more penetrable for subsequent fermentation (Chandra et al. 2007). • Microwaves heating at short time create localized hotspots and open up the lignocelluloses biomass, facilitating enzyme access for saccharification (Canam et al. 2013). • Organosolv process modifying chemically and removing low molecular weight lignin fractions from biomass using alcohols such as ethanol, methanol, or other solvents at high-temperature extraction. In this process, some dilute acids like hydrochloric and sulfuric acid are used as a catalyst (Agbor et al. 2011). • As natural modification and degradation of the lignin component, biological pretreatments can reduce the severity requirements of obvious pretreatment strategies. The exploitation of the capacity to access and to modify the lignocellulosic biomass by microorganisms was studied by several authors such as Itoh et al. (2003) who used a variety of lignin-degrading white-rot fungi pretreatments followed by extracting lignin by an organosolv method from wood chips. This process leads to saving of electricity up to 15% and increases the ethanol yield obtained from the solid fraction. The enzymatic degradation of lignocellulose by microbial consortia or lignin-­ degrading fungi involves both oxidative and hydrolytic mechanisms based on OH radical reactivities. Attack on the hydroxyl group in lignin subunits by abstracting aliphatic hydrogens and adding to aromatic rings results in benzyl ketone production and hydroxylated cyclohexadienyl radical generation occurring on subsequent C-C bond cleavage and degradative chain reactions (Gierer 1990). This leads to decreasing energy requirements, less producing of fermentation-inhibiting substances. Thus, biological pretreatment of lignocellulosic materials is very useful when incorporated into any economical biorefinery strategies for biofuels and/or metabolite production (Isroi et al. 2011; Chen et al. 2010). In pulp and paper industries, processed lignin is produced traditionally through three distinct processes: kraft lignin, lignosulfonate lignin, and organosolv lignin (Doherty et  al. 2011). Other than renewable bioenergy resources and pulp and paper industry supplement, significant progress is being made in research for the feasibility of a variety of applications of lignin or lignin-related product as a resource for chemicals and materials. For this, two different strategies are followed for lignin conversion to value-added components, either controlled depolymerization into small molecules (Li et  al. 2015; Behling et al. 2016) or building block to synthesize new functional materials (Kai et  al. 2016; Upton and Kasko 2016). Possessing multiple functional groups that can form inter- and intramolecular hydrogen bonding, lignin is considered as an alternative source for the production of more value-added chemicals according to its compatibility with host matrices (Karam and Nicell 1997; Doherty et  al. 2011; Wang et al. 2016). This compatibility which is improved by chemical modification (oxyalkylation or hydroxyalkylation) minimizing lignin molecules’ auto-­association (Zhao et  al. 2016; Feldman et  al. 2001; Maldhure et  al. 2012). Gallezot (2007) ­indicated that three potential strategies for lignin valorization to produce fine chemicals with a high degree of functionality can be adopted.

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The first strategy acts on gasifying lignin allowed to synthesize gas or degraded by pyrolysis to a mixture of small molecules. The second strategy turns, in the first step, the functional groups present into the lignin monomers’ simple aromatic compounds such as phenol, benzene, toluene, and xylene. In a second step, bulk and fine chemicals are produced using catalytic technology developed for petroleum refineries. Finally, using highly selective catalysts in a one-pot approach, the third strategy converses directly the biomass to valuable chemicals (Karam and Nicell 1997; Doherty et al. 2011; Thakur et al. 2014; Chatterjee and Saito 2015; Liu et al. 2015). Then, several value-added chemicals are produced including: 1. Lignin: as a filler or additive in polymers and biopolymers – usually at less than 20–30% of total weight 2. Lignin-derived functional materials such as carbon fibers, activated carbon, adhesives, and foams 3. Lignosulfonates: as dispersants, water reducer in concrete, additive in coal-water slurry, or viscosity reducer In addition, in light of renewed interest in promoting value-added applications of lignin, it is expected recently lignin nanoparticles will play a vital role in promoting lignin valorization in polymer industry (Stark et al. 2015).

5.2.2  Lignin Degradation Lignin is a complex polymer with very low degradation rate in nature. The biodegradation which is carried out by wood-rotting fungi constitutes a key step for carbon cycling in nature, as well as for the industrial use of plant biomass by increasing accessibility to cellulose. It is an oxidative process that has been investigated for decades as a model for biotechnological application in the pulp and paper industries, animal feeding, and bioethanol production. Ligninolytic oxidoreductases (laccases and different types of peroxidases) secreted by wood-rotting fungi are the sole enzymes able to oxidize the phenylpropane lignin units (Schwarze 2007; Yadav et al. 2016, 2018). However, different enzymes have been reported for their role in lignin degradation such as lignin peroxidase (LiP), manganese peroxidase (MnP), aryl alcohol oxidase (AAO), and glyoxal oxidase (GLOX) as shown in Fig. 5.2. The main enzymes associated with lignin degradation are laccases, lignin peroxidases, and manganese peroxidases; while some white-rot fungi produce all the three classes of enzymes, others produce only one or two (Hatakka 1994). Laccases are multicopper enzymes, which catalyze the oxidation of phenolic compounds including a range of dyes with concomitant reduction of oxygen (Eggert et  al. 1996; Chivukula and Renganathan 1995; Munoz et al. 1997). Recent interest in laccase is, in part, a consequence of the findings that the substrate range of laccase can be expanded to include non-phenolic dyes, eventually in the presence of suitable mediators (Bourbonnais and Paice 1990).

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Fig. 5.2  Lignin-degrading enzymes’ actions. LiP lignin peroxidase; MnP manganese peroxidase; AAO aryl alcohol oxidase; GLOX glyoxal oxidase; VP versatile peroxidase; MED mediator. (Modified from Abdel-Hamid et al. 2013)

5.3  Laccase (EC 1.1.2.2) Mushrooms excrete different oxidative, hydrolytic, and non-hydrolytic enzymes that act synergistically to hydrolyze the complex chemical structure which is composed of cellulose, hemicelluloses, and lignin (Guerriero et al. 2015). Laccases and class II peroxidases belong to the oxidative enzymes including lignin peroxidase, manganese peroxidase, and hybrid lignin/manganese versatile peroxidase which catalyze the cleavage of C-C and C-O-C bonds in a wide variety of organic compounds, such as lignin and polyphenolic structures (Ertan et al. 2012; Siddiqui et al. 2014). Laccases (or phenoloxidase) were firstly described and identified in 1883 by Yoshida, from the Japanese lacquer tree Rhus vernicifera. They are phenol-­oxidizing enzymes and listed as ecofriendly catalyzing variety of reactions involving one-­ electron oxidation with water generation (Hao et  al. 2007; Javed et  al. 2017). Laccases are widespread in nature and have been found in non-microbial sources (plants and insects) and in microbial sources (Thurston 1994; Brijwani et al. 2010; Mayer and Staples 2002; Santhanam et al. 2011; Yavuz et al. 2014). As shown in Table  5.1, the genus Pleurotus showed strong laccase activity (Silva et  al. 2012; Alexandrino et al. 2007; Sethuraman et al. 1999; Ardon et al. 1996). P. ostreatus produces many extracellular enzymes with strong ability for lignin degradation and substrate oxidation. Therefore, this mushroom attracted attention because of its high potential for biofactory laccase production for many biotechnology applications (Karas et al. 2011; Faraco et al. 2009; Pezzella et al. 2013).

Manganese peroxidase (EC 1.11.1.13)

Enzyme (EC number) Laccase (EC 1.10.3.2)

P. ostreatus CBS 411.71

P. ostreatus

P. ostreatus P. ostreatus

P. ostreatus 1804 (Immobilized on PUF) P. ostreatus

2.4 UmL−1 803.3 U

Free mycelia: 272.2 U Immobilized: 312.6 U Packed bed reactor:392.9 U 22.50 U mL−1

Basic Broth Media

Wheat bran

Glucose-based chemically defined medium

Submerged culture using CuSO4 (150 μM) as inducer, incubation for 15 days at 28 °C Solid-state fermentation at 28 °C and pH 5.5 Sugarcane bagasse 167 U g−1 Maximal production of Solid-state fermentation, incubation at 20 °C Three agro-industrial 16 μM min kg−1 in coffee for 20 days. Produce enzyme cocktail of wastes (coffee husks, laccases, MnP, and cellulase eucalyptus sawdust, and husks eucalyptus bark) with 20% rice bran Water polluted with 200 U L−1 Submerged cultures incubated for 24 days at wheat straw extracts 25 °C in the dark Semi-defined medium 451 U L−1 Submerged cultivation for 6 days at 26–28 °C with glucose

Coffee husk

Submerged culture incubated at 28 °C agitated at 100 rpm for 14–16 days at pH 6.5 Submerged culture incubated at 28 °C and pH 5.5 Production increased by addition of (2,5-xylidene; 1.0 mM) as inducer after 96 h Submerged culture in packed bed reactor incubating for 192 h at pH 5.6

0.95 U mL−1

Cotton stalk extract

P. ostreatus Florida f6 P. ostreatus strain V-184 P. ostreatus 1804

Cultivation conditions/parameters Solid-state fermentation, cultivation at wide temperature range 20–35 °C for 20 days (laccase production), 30 days (MnP production) Solid-state fermentation for 16 days at 28 °C

Enzyme activity 75.0 U g−1 (laccase) 6.8 U g−1 (MnP)

Substrate/media composition Orange waste

Mushroom type P. ostreatus

Table 5.1  Different enzymes produced by Pleurotus ostreatus

Parenti et al. (2013) Sarkar et al. (1997)

Karp et al. (2012) Rodrigues dal Luz et al. (2012)

Silva et al. (2012)

Krishna et al. (2005b)

Ardon et al. (1998) Mansur et al. (2003) Krishna et al. (2005a)

References Alexandrino et al. (2007)

108 H. El Enshasy et al.

Aryl alcohol oxidase (EC 1.1.3.7)

Versatile peroxidase (EC 1.11.1.16)

Enzyme (EC number)

P. ostreatus (strain Florida)

P. ostreatus (accession number CECT20311)

Mushroom type P. ostreatus ATCC 66376 P. ostreatus dikaryotic strain #261 ATCC 66376 P. ostreatus IBL-02 P. eryngii (accession number GIM5.280) P. ostreatus (accession number CECT20311) P. eryngii ATCC 90787 0.04 U g−1

Sterile wheat straw

Camarero et al. (1999)

Salame et al. (2012)

(continued)

Sannia et al. (1991)

Cultivation using SSF method

0.16 U mL−1

Cotton stalks

Submerged cultivation in dark for 20 days

Submerged cultivation in dark for 10 days at 28 °C

178.9 U mL−1

Cotton stalks

Ashger et al. (2013) Min et al. (2010)

Feldman et al. (2015)

Submerged cultivation at 25 °C

692 U mL−1

Wheat straw

References Kamitsuji et al. (2005) Tsukihara et al. (2006a)

Submerged cultivation for 24 days in dark at 30 °C

Solid-state fermentation for 7 days at 30 °C

7300 U L−1

Semi-synthetic medium 300 U L−1 supplemented with different inducers: tyrosine, VA, and benzyl alcohol Saw dust-based medium 211 U L−1 with addition of inducers VA, olive oil, and tween 80

Cultivation conditions/parameters Submerged cultivation for 25 days in dark at 28 °C and pH 4.5 Submerged cultivation for 28 days in dark at 28 °C

Enzyme activity 900 U L−1

Substrate/media composition Chemically defined glucose-based medium Rice bran 5  Pleurotus ostreatus: A Biofactory for Lignin-Degrading Enzymes of Diverse… 109

Xylanase (EC 3.2.1.8)

Enzyme (EC number)

P. ostreatus (Jacq.:Fr.) Kumm P. ostreatus

P. ostreatus

P. ostreatus PLO6

Pleurotus HK-37

SSF, optimal enzyme recovery was achieved when SMCs were extracted with 50 mM sodium citrate (pH 4.5) buffer at 4 °C SSF cultivation for 20 days at 26–30 °C and humidity 78 ± 2%

SSF cultivation for 45 day at 25 °C

SSF cultivation in dark for 50 days 20 °C. SSF, fruiting bodies formed after 49 days

Submerged cultivation 30 °C for 7 days

91.56 U g−1,

3.73 U g−1

0.51 U mg−1

21.0 U g−1 48 U g−1,

0.863 U ml−1

Spent mushroom compost (SMC)

10% sisal leaf and sisal boles with ration (0:100) and cow dung manure Oxo-biodegradable plastic without physical treatment Saw dust

Cotton wastes (CW) as growth substrate

Modified Czapek medium

Rice bran and saw dust

Submerged cultivation at 26 °C

24.98 U mL−1 Cultivation in submerged culture in dark for 32 days at 22–26 °C Fruiting bodies were produced after incubated for 20 days at 25 °C and 90%

Cultivation conditions/parameters SSF is the best to secrete xylanase enzyme after 4 days of incubation at 30 °C and pH 4

Enzyme activity 11.47 U mL−1

155.8 (nmol/min × mL) 65 μM min−1 kg −1 of substrate

P. ostreatus (PLO 2 and PLO 6) P. ostreatus

P. ostreatus SYJ042 P. ostreatus

Substrate/media composition Wheat straw supplemented with xylan, peptone, and KCl 2.5% (corncob), 2.5% (wheat bran), peptone Pepper extract

Mushroom type Pleurotus sp.

Table 5.1 (continued)

Karthikeyan, (2015)

Sherief et al. (2010) Elisashvili et al. (2003)

Rodrigues da Luz et al. (2013)

Raymond et al. (2015)

Lim et al. (2013)

Qinnghe et al. (2004) Morais et al. (2005) Rodrigues da Luz et al. (2012)

References Getachew et al. (2016)

110 H. El Enshasy et al.

Pectinase (EC 3.2.1.15)

Endoglucanase (EC 3.2.1.4) Exoglucanase (EC 3.2.1.4) CMCase Cellulase

Exoglucanase (EC 3.2.1.4)

Endoglucanase (EC 3.2.1.4)

Enzyme (EC number) Cellulase (EC 3.2.1.4)

P. ostreatus HK-37 P. ostreatus NRRL 366 Pleurotus HK-37 P. ostreatus

P. ostreatus LCJ 183 P. ostreatus

Mushroom type P. ostreatus (Jacquin ex Fr.) Kummer P. ostreatus (Jacq.) P. Kumm. NRRL 366 P. ostreatus (Jacq.) P. Kumm. NRRL 366 P. ostreatus P. ostreatus

13.21 U g−1 1.25 U ml−1

Saw dust What bran

8.28 U g−1 21.42 U g−1

Sawdust

14.2 U g−1

Papain waste and rice straw 30% sisal leaf

SSF for 30 days at 26–30 °C and 87% humidity SSF dark cultivation at 20 °C

Submerged cultivation

(continued)

Raymond et al. (2015) Rashad et al. (2009) Raymond et al. (2015) Sherief et al. (2010)

6.01 U g−1

Rice straw

Sherief et al. (2010)

Daba et al. (2011)

SSF for 30 days at 26–30 °C and 87% humidity SSF, for 7 days cultivation at 28 °C

SSF cultivation for 20 days at 20 °C

4.02 U g−1

Avicel PH101

Spent mushroom 1.67 U g−1 compost Sisal leaf and Sisal bole 3.73 U g−1

Submerged cultivation for 12 days at 27 °C and pH 5.5

1.80 U ml−1

Avicel PH101 Rice straw

Daba et al. (2011)

Radhika et al. (2013) Lim et al. (2013)

Submerged cultivation for 12 days at 27 °C and pH 5.5

2.46 U ml−1

Avicel PH101

References Khalil et al. (2011)

Submerged cultivation system

Cultivation conditions/parameters Cultivation for 14 days at 37 °C and pH 5.5

Enzyme activity 3.51 U ml

Substrate/media composition Rice straw 5  Pleurotus ostreatus: A Biofactory for Lignin-Degrading Enzymes of Diverse… 111

ProA – serine protease (EC 3.4.22.9) ProB – metalloprotease (EC 3.4.99.32) ProC – metalloprotease (EC 3.4.24.4) Fibrinolytic protease (EC 3.4.21.5)

Enzyme (EC number) Pectin lyase (EC 4.2.2.10) Polygalacturonase (PG) (EC 3.2.1.15) Exopolygalacturonase (exo-PGase) (EC 3.2.1.67) Cysteine protease

Shen et al. (2007)

Submerged cultivation for 5 days at 25 °C

8424 U

- Defined medium supplemented with glucose

P. ostreatus

Dohmae et al. (1995)

8.10 U mg−1

Defined medium supplemented with glucose and soymilk –

P. ostreatus, No 4241

Shin and Choi (1998) Liu et al. (2014)

Rashad et al. (2009)

References Rashad et al. (2011) Rashad et al. (2010)

Obtained from the fruiting bodies grown by 110 U SSF 23,600 U Present (quantitative analysis was not given in this work)

Obtained from the fruiting bodies grown by SSF Submerged cultivation for 6 days at 25 °C

14.68 U g−1

Commercial product

P. ostreatus

P. ostreatus (Hiratake)

Submerged fermentation for 4 days

SSF for 4 days at 28–30 °C

Cultivation conditions/parameters SSF for 4 days at 28–30 °C

1750 U

Enzyme activity 81.30 (U mg−1 protein) 158 U mg−1

Lemon peel

Lemon pulp waste

Substrate/media composition Lemon pulp waste

P. ostreatus NRRL 366

Mushroom type P. ostreatus NRRL 366 P. ostreatus NRRL 366

Table 5.1 (continued)

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Lipase (EC 3.1.1.3)

Amylase (EC 3.2.1.x)

Metalloprotease Serine protease (EC 3.4.21)

Enzyme (EC number) Serine protease (EC 3.4.21.14)

P. ostreatus

P. ostreatus (Jacq.) Pleurotus Kumm. (MCC16) P. ostreatus

P. ostreatus

Mushroom type P. ostreatus (Jacq.:Fr.) Kummer (type:Florida) P. ostreatus (PLO 06, GenBank accession number KC782771 P. ostreatus (Jacq.) Pleurotus Kumm. (MCC16) P. ostreatus

Produced in immobilized cell cultivation system Submerged fermentation (SF), 5 days of cultivation

0,93 U mg−1

Using chemical defined medium Medium supplemented with olive mill wastewater

2.97 U g−1

Spent mushroom compost (SMC) Potato peel waste

30 U L−1

6 U L−1

Obtained from the fruiting bodies grown by SSF Obtained from the fruiting bodies grown by SSF SSF for 23 days at 25 °C

SSF for 23 days at 25 °C

4 U L−1

Potato peel waste

220 μmol/min/mg

Submerged cultivation for 7 days at 28 °C

120 U ml−1

Medium containing (glucose 10 g /l; yeast extract, 10 g/l; K2HPO4, 5 g/l; and MgSO4, 0.10 g/l)

Rice bran

Cultivation conditions/parameters Submerged cultivation in dark for 3 days at 27 °C

Enzyme activity 2.5 U mg−1

Substrate/media composition Basal medium (potato dextrose broth, yeast extract

Wijayati et al. (2017) Guarino and Sannia (2013)

Ergun and Urek (2017)

Akinyele et al. (2010) Lim et al. (2013)

Ergun and Urek (2017)

Genier et al. (2015)

References Palmieri et al. (2001)

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5.3.1  Laccases Structure and Mechanism of Reaction Laccase (EC 1.10.3.2), or benzenediol: oxygen oxidoreductases, belongs to a group of phenol oxidizers enzymes called blue multicopper oxidases, which are considered generally as part of the monomeric glycoprotein group. This group of enzyme is mainly exocellular enzyme that can oxidize phenols and aromatic compounds including amines, esters, and ethers (Rathinasamy and Thayumanavan 2010; Solomon et al. 1996). They act by reducing the molecular oxygen to water and catalyzing the oxidation of ortho- and para-aminophenols, diphenols, aryl diamines, polyphenols, polyamines, and lignin (Shleev et al. 2006a, b). Therefore, laccases can involve in the degradation of a wide range of xenoaromatics substrates such as textile dyes, polychlorinated biphenyls, polycyclic aromatic hydrocarbons, pesticides, and synthetic polymers (Mester and Tien 2000; Bezalel et al. 1997; Novotny et al. 2000; Riva 2006; Yadav et al. 2019a, b). Regarding this high nonspecific oxidation capacity, laccases derive interests and attraction from researchers to be used as biocatalysts for many industrial and biotechnological applications, from food processing to environmental technologies through textile and paper industry. Recent research showed also the potential applications of laccases in nanobiotechnology and biomedicine (Mayer and Staples 2002; Jurado et al. 2009; Zhuo et al. 2011; Javed et al. 2017). Based on genetic research, four different genes and their corresponding cDNAs have been identified from P. ostreatus and named pox1 (laccase isoenzyme not yet identified) (Giardina et  al. 1995), pox2 or poxc (Giardina et  al. 1996), poxa1b (Giardina et  al. 1999), and poxa3 (Palmieri et  al. 2003). According to Mot and Silaghi-Dumitrescu (2012), the structure of laccases is held up by monomeric units consisting of three domains which are arranged in a sequence to form a barrel-type structure. The same study reported that fungal laccases lack high-order oligomeric assemblies making the crystal lattice and are found generally as monomers. According to Hoegger et al. (2006), laccases have four catalytic copper atoms in their structure organized in two individual sites that bind the reducing substrate and O2. The substrate oxidation site contains a paramagnetic type 1 copper (T1 or T1Cu) responsible for the characteristic blue color of the enzyme in the reduced resting environment. Another copper (T2 or T2Cu) contributes in catalytic and redox activity site coordinated by three histidines. The two other coppers (T3 or T3Cu) are responsible for the activation and O2 transport and substrate oxygenation and coordinated by three histidines (Hoegger et al. 2006). Both T2Cu and the two T3Cu are clustered at 12 Å from the T1Cu and form a tri-nuclear site as shown in Fig. 5.3, where O2 is reduced to two molecules of water, receiving four consecutive electrons from four independent mono-oxidation reactions at the T1Cu site through a strictly conserved His-Cys-His electron transfer route (Mot and Silaghi-Dumitrescu 2012). Beyond this molecular configuration, other enzymes can include also 2, 3, or 6 copper atoms (Gil et al. 2009; Messerschmidt and Huber 1990). The high potential of copper T1 is of great interest to biotechnologists because of its geometry. In this site, around the T1 copper atom, two histidines and one cysteine which are orga-

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Fig. 5.3  Representation of a native fungus laccase from Trametes hirsute (Left), showing the copper ions organization (Right). http://www.rcsb.org/pdb/explore/jmol.do?structureId=3FPXopt=3

nized three-dimensionally with two other residues are weakly coordinated from the axial position. One of these residues is isoleucine, and the other can be phenylalanine, leucine, or methionine depending on the type of laccase characterized and reduction potential (Rodgers et al. 2010; Rivera-Hoyos et al. 2013). Several authors have reported that more than one laccase isozyme is secreted by fungi (Soden and Dobson 2001; Hoshida et al. 2001; Palmieri et al. 2003; Rodrïguez et al. 2008). At least, eight different laccase isoenzymes are produced by P. ostreatus and six of which have been isolated and fully characterized (Giardina et  al. 1999; Pezzella et al. 2009; Palmieri et al. 1993, 1997, 2003). The most abundant protein present in the P. ostreatus cultures is POXC (59-kDa with pI 2.7). The enzymes POXA1b and POXA1w have a similar molecular weight around 61 kDa, while POXA2, POXB1, and POXB2 isoenzymes are of higher molecular weight around 67 kDa. POXA3a and POXA3b are heterodimers and composed of small (16 or 18 kDa) and large (61 kDa) subunits (Palmieri et al. 1997; Giardina et al. 1999). However, several studies reported that P. ostreatus produces mainly three laccase isoenzymes called POXCs (Giardina et al. 1996): POXA1w, the white laccase isoenzyme with a singular metal content (Palmieri et  al. 1997); POXA1b, the more stable isoenzyme at alkaline pH (Giardina et al. 1999); and the heterodimeric laccase isoenzyme POXA3 (Palmieri et al. 2003; Giardina et al. 2007; Faraco et al. 2008) with two forms, POXA3a and POXA3b (Palmieri et al. 2003; Giardina et al. 2007), that exhibit unusual structural features, being heterodimeric enzymes (Wahleithner et al. 1996; Yaver et al. 1996). These laccase isoenzymes are endowed with quaternary structure, consisting of two subunits of different molecular weights (Faraco et al. 2008). Palmieri et al. (1997) reported that the two laccase isoenzymes

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POXA1 and POXA2 are monomeric glycoproteins containing 3% and 9% carbohydrate with pI values of 6.7 and 4.0, respectively. The sequencing of the N terminus and three tryptic peptides of POXA1 reveals clear homology with laccases from other microorganisms, while POXA2 showed a blocked N terminus. However, POXA1 showed remarkable high stability at relatively wide range of temperature and pH values, whereas POXA2 is less stable at high temperature (Palmieri et al. 1997). The molecular weights of the two laccase isoenzymes were determined with three different methods. Sodium dodecyl sulfate polyacrylamide gel electrophoresis shown that the Mwt of POXA1 and POXA2 are of about 61 and 67 kDa; Gel filtration in native conditions shows a Mwt of about 54 and 59  kDa; Finally, matrix-­ assisted laser desorption ionization mass spectrometry was applied for POXA1 and give 61 kDa (Palmieri et al. 1997). Substrates with specific linkages and structural similarity with lignin, an amorphous polyphenolic polymer, can induce laccase activity. In addition, a wide range of aromatic compounds can be oxidized by laccases with concomitant water generation by reduction of molecular oxygen (Thurston 1994; Ullah et al. 2000). In general, laccase is not able to oxidize substrates with lower redox potential like non-phenolic compounds. However, Call and Mücke (1997) have reported that the oxidation can be catalyzed in the presence of electron transfer mediators. Oxidation of substrates with chemical structure smilar to lignin like amino-phenols, orth- and para-diphenols, polyamines, aryl diamines, lignins and polyphenols is also carried out using the same mechanism. Then, laccases can act on many substances such as agricultural inputs, pesticides and herbicides, and dyes used in many industrial processes which must be degraded before being discarded (Morozova et al. 2007). According to Baldrian (2006), more than 100 fungal laccases are purified and characterized and usually with optimal activity at acidic pH range with a temperatures range from 50 to 70 °C. Table 5.1 summarizes some other characteristics of laccase isoenzymes from Pleurotus ostreatus strains. As reported by Palmieri et al. (1993, 1997), generally, for the P. ostreatus laccase POXC, the optimal pH is already acidic for both ABTS and DMP degradation but around 6 for guaiacol and syringaldazine with a range of optimum temperature from 50 to 60 °C (Palmieri et al. 1993, 1997). Studying the two laccase isoenzymes POXA1 and POXA2 from P. ostreatus, Palmieri et al. (1997) reported that POXA2 acts with an acidic optimum pH of 3 for ABTS as substrate but an optimal pH of around 6 with other substrates like DMP, guaiacol, and syringaldazine, with optimum range of temperature between 25 and 35 °C. Both POXA1and POXA2 isoenzymes oxidize ABTS and syringaldazine, but POXA1 is unable to oxidize guaiacol (Palmieri et al. 1997).

5.3.2  Laccase Production and Bioprocessing P. ostreatus grows naturally on tree stumps and can be easily cultured in the laboratory scale. However, most of mushroom cultivars usually use cereal straw-based substrates (Arjona et  al. 2009). Different parameters influence the laccase

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production which is linked to a complex regulation of nutrients that affects directly the molecular expression. These include the type and concentration of carbon and nitrogen sources used, C/N ratio, and also the presence of some inducers and their concentrations (Terrón et al. 2004). Beside the nutritional factors, the production of laccase by P. ostreatus is influenced by other factors such as pH, the fermentation technique applied, and cultivation conditions like agitation, aeration, cultivation time, and many other factors (Terrón et  al. 2004; Soden and Dobson (2001); Periasamy and Palvannan 2010; Janusz et al. 2007). Different studies were focused on the optimization of the fermentation medium for laccase production by P. ostreatus to facilitate economic design of the full-scale fermentation operation systems. Studies include the investigation of the effect of carbon and nitrogen source (Stajic et al. 2006a; Liu et al. 2009), addition of some metal ions (Soden and Dobson 2001; Collins and Dobson 1997; Galhaup and Haltrich 2001; Galhaup et  al. 2002; Baldrian and Gabriel 2002), and addition of aromatic compounds as inducers (González et al. 2003; De Souza et al. 2004; Terrón et al. 2004; Krishna et al. 2005a). 5.3.2.1  Media Composition and Cultivation Conditions It has been reported that increasing the concentrations of glucose, wheat bran, urea, yeast extract, KH2PO4 and inoculums can increase laccase production (Krishna et al. 2005a; Fernández-Fernández et al. 2013). The works of Ardon et al. (1998) and Krishna et al. (2005b) show that the maximal laccase production by P. ostreatus was obtained within the pH range of 5.0–5.5 in submerged culture. Another study carried out by Terrón et al. (2004) reported that the presence of nine different aromatic compounds, including p-coumaric acid, guaiacol, and ferulic acid, can enhance laccase activity. Zhuo et  al. (2017) studied the effect of aromatic compounds on the transcript levels of laccase genes, and the results have confirmed the previous results of Nyanhongo et al. (2002) who used ferulic acid as an effective laccase inducer and show that ferulic acid and vanillic acid have the most pronounced stimulatory effect on laccase gene transcription (Zhuo et  al. 2017; Nyanhongo et al. 2002). Krishna et al. (2005a) reported on the positive effect of 2,5-xylidene addition to culture media after 96 h on laccase production. However, the inducers’ effects depend on the chemical structure, concentration, and time of its introduction into the production medium (Pointing et al. 2000; Sethuraman et al. 1998). The production of laccases in P. ostreatus is regulated by the presence of copper. Therefore, the two dimeric isoenzymes POXC and POXA1b have been detected only in the presence of copper (Palmieri et al. 2000, 2003). 5.3.2.2  Cultivation Process and Purification Solid-state fermentation is an interesting operational mode for the production of laccases by P. ostreatus. For solid-state fermentation, Karp et  al. (2012) have reported a productivity of 151.6  Ug−1 after 5  days in sugarcane bagasse-based

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medium. Several studies have showed high laccase productivity by P. ostreatus in submerged culture conditions; a productivity of 3500 U L−1 was reported by Lenz and Hölker (2004). Tlecuitl-Beristain et al. (2008) have reported 80 UmL−1 after 12  days in liquid culture without exogenous inducers (Lettera et  al. 2011); also 12.2 UmL−1 has been reached after 18 days according to Tlecuitl-Beristain et al. (2008). Optimizing the laccase production in submerged culture conditions, Krishna et al. (2005a) and Rathinasamy and Thayumanavan (2010) have increased the laccase expression by 32.9%, from 538.8 to 803.3  U, and by 86.8%, from 485.0 to 906.3 U, using P. ostreatus 1804 and P. ostreatus IMI 395545, respectively. Mansur et al. (2003) studied the secretion of laccase isozymes by P. ostreatus V-184 with different substrate specificities. They showed that laccase activity reached 2.4 U mL−1 using submerged culture for 14–16 days, at pH 6.5. Krishna et al. (2005b) have also studied the feasibility of Pleurotus ostreatus 1804 immobilization on polyurethane foam (PUF) cubes in the objective of laccase production in three different cases of submerged culture system. After 192 hours of cultivation at 5.6 pH, the enzyme yields of free mycelia and immobilized and packed bed reactor were 272.2 U, 312.6 U, and 392.9 U, respectively. Rodrigues da Luz et al. (2012) reported also on the potential use of agrowastes such as coffee husks, eucalyptus sawdust, and eucalyptus bark as substrate in combination with rice bran for laccase production by P. ostreatus as substrates in combination with 20% rice bran. Parenti et  al. (2013) have used water polluted with wheat straw extracts as substrate in submerged cultures incubated in the dark at 25 °C for 24 days with orbital shaking (150 rpm) and reported a laccase activity of 200 UL−1. To separate laccase isoenzymes, Palmieri et al. (1997) have fractioned P. ostreatus culture broth after 70 h of growth using ammonium sulfate precipitation followed by anionic exchange chromatography. Five different laccase fractions were separated (POXA1, POXA2, POXB1, POXB2, and POXC). Elution with a saline gradient at around 0.17, 0.18, and 0.32  M NaCl allowed to separate three isoenzymes POXB1, POXB2, and POXC, respectively, whereas the major laccase peak activity, corresponding to POXA1, and a fraction of POXA2 were recovered using the equilibrating buffer (Palmieri et al. 1997).

5.3.3  Applications of Laccase Since the nineteenth century, laccases are considered as the most effective green catalyst by many researchers (Javed et al. 2017). Due to their nonspecific and high oxidative capacity, laccases present a high potential in many industrial and biotechnological applications with a great market demand for commercial applications in detergent, food, cosmetic, paper/pulp, and textile industries. In addition, they have also many applications in soil bioremediation and wastewater treatment (Shah and Nervd 2002). Recent studies showed also the potential use in pharmaceutical industries based on the enzyme ability for transformation of antibiotics and steroids (Cohen et al. 2002; Wu and Nian 2014; Pezzella et al. 2013; Yadav et al. 2017a, b).

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5.3.3.1  Bioremediation of Environmental Contaminants in Soil Soil bioremediation is a process applied to recover pollutant from contaminated soils using bacteria and fungi which degrade these organic compounds, transforming them into compounds that are less or nontoxic. Bioremediation is considered a safer, cheaper, and more efficient method compared to other physicochemical methods (Chibuike 2013). Acting on substrates, which are insoluble and very linked to the soil particles and nonaccessible to bacteria, makes the application of laccases more attractive. These exocellular enzymes are able to degrade some pollutant compounds similar to lignin such as polyaromatic hydrocarbons, chlorophenols, and nitrophenols (Chibuike 2013). Action of laccases on the xenobiotics releases intermediate products with more bioavailability and less toxic, which can be more efficiently removed by physicochemical and mechanical processes (Javed et al. 2017; Viswanath et al. 2014; Zucca et al. 2015). Anthracene, benzopyrene, and organophosphorus compounds, like the nerve agents VX or Russian VX, have been removed by microbial and fungal laccases (Amitai et al. 1998; Zeng et al. 2016). Other studies showed also that effluents from sewage treatment containing estrogenic hormones have been treated using enzymatic systems containing laccases. Fungal laccase has been used to oxidize the estrogens like estrone, 17b-estradiol, and 17a-ethynylestradiol (Tanaka et al. 2009). 5.3.3.2  Wastewater Treatment The appropriate wastewater treatment is important regarding that industrial sewage can contain many carcinogenic, mutagenic, and teratogenic potential substances with toxic effects to human, fish, microorganisms, and plant species (Aljeboree et al. 2014). Using enzymatic processes in wastewater treatment is relatively a new approach decreasing reagents’ consumption and allowing degradation of many persistent and toxic substances such as dyes, solvents, inks, fertilizers, and pesticides (Niebisch et  al. 2014). One of the applications of laccases in combination with peroxidases in wastewater treatment is related to the discoloration of wastewater from textile industry that contains different types of dyes such as azo, triphenylmethane, and anthraquinone (Fig.  5.4). Laccases act on the degradation of the chemical dyes before the final disposal. In this process, immobilized laccases in alginate beads are commonly used (Niebisch et  al. 2014; Peralta-Zamora et  al. 1997). Another study by Zhuo et al. (2017) investigated the ability of laccase from P. ostreatus HAUCC 162 to decolorize six synthetic dyes belonging to different categories (methyl orange, crystal violet, malachite green, bromophenol blue, Reactive Blue 4, and Remazol Brilliant Blue R (RBBR)). The results showed that within 24 hours, and at an initial concentration of 100 mg/L, methyl orange, crystal violet, malachite green, and bromophenol blue could be decolorized by 81.3%, 87.6%, 85.1%, and 98%, respectively. For Reactive Blue 4 and RBBR, the removal of 64.6% and 89.1% was achieved, respectively, when using initial concentration up to 800 mg/L (Rui Zhuo et al. 2017).

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Fig. 5.4  Different synthetic dyes degraded by extracellular laccase from P. ostreatus

Decolorization of crystal violet by laccase from P. ostreatus MTCC142 has also been studied by Kunjadia et al. (2012), and the results show that 92% of dye decolorization can be achieved at an initial concentration of 20  mg/L.  Palmieria et  al. (2005) have reported the decolorization of the recalcitrant anthraquinonic dye Remazol Brilliant Blue R (RBBR) by P. ostreatus on agar plate and also in liquid media supplemented with veratryl alcohol, where RBBR was completely decolorized in 3 days. Two isoenzyme laccases in mixture (POXC and POXA3) were found to be responsible for this RBBR transformation under acidic conditions reducing its toxicity by 95% (Palmieria et al. 2005). Another study by Hongman et al. (2004) found that the anthraquinone dye SN4R (Remazol Brilliant Blue SN4R) decolorization rate was increased by 90% in the presence of ABTS as a mediator of laccase. The crude laccase alone in the concentration of 30 Uml−1 can effectively decolorize the dye by 66%. Reactive Blue HFRL dye decolorization was also achieved by using mushroom laccase after 3 days (Devi et al. 2012). 5.3.3.3  Food Industry Laccases can catalyze homo- and hetero-polymerization reactions needed in fruit juice processing, in sugar beet pectin gelation, and for wine/beer stabilization (Osma et al. 2010). In addition, laccases are currently being utilized in baking to

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cross-link biopolymers (Javed et  al. 2017). It was also used to preserve and to increase juice stability by reducing the oxidation of phenolic compounds, the formation of polyphenol and protein is delayed, and there are many studies which show that laccases can be used for food preservation/stabilization process (Osma et al. 2010; Sammartino et al. 1998). 5.3.3.4  Paper and Textile Industries In paper industry, the pretreatment of wood pulp with ligninolytic oxidoreductases shows special attention to replace the conventional chlorine-based delignification processes. Unlike peroxidase, the benefit of using laccase is that it requires O2 rather than H2O2 (Javed et al. 2017). In addition, use of laccase for phenols grafting to flax fibers in the production process of paper has been recently studied (Fillat et al. 2012; Aracri et al. 2010; Virk et al. 2012). Laccases can be used in textile industry for bleaching processes and they have now replaced the conventional peroxide bleaching to enhancing the whiteness of cotton (Yavuz et al. 2014; Iracheta-Cardenas et al. 2016; Tzanov et  al. 2003). Laccase-based products are also applied for denim bleaching and against fabrics odor (Rodriguez-Couto 2012; Kunamneni et al. 2008). Different textile dyes like phenoxazine and azo dyes have been produced using laccases based on their ability to oxidize the aromatic compounds (Sousa et al. 2013). 5.3.3.5  Biofuels Lignocellulosic materials are the most reliable feedstock for bioethanol and other organic alcohol production in biofuel industries (Kour et al. 2019a). However, based on the complexity of the material, it requires pretreatment of biomass to eliminate lignin and thus expose the cellulose/hemicellulose to hydrolytic enzymes. An interesting research showed that P. ostreatus enzymes have potential application in bioethanol production process (Yu et al. 2009). The results showed that the combination of ultrasonic pretreatment with enzymatic hydrolysis of rice hull showed a maximal yield by using laccases followed by lignin peroxidase and Mn peroxidase (Yu et al. 2009). 5.3.3.6  Biomedical, Pharmaceutical, and Cosmetic Industries Based on the ability of laccases to catalyze reactions by direct electron transfer, developing biosensors based on laccases systems attracts scientists for many biomedicine application like in insulin, morphine, and codeine analysis (Rodriguez-­ Delgado et  al. 2015). In addition, this enzyme can play a potential role in pharmaceutical industries in many processes such as antibiotic transformation, amino acid derivatization, and synthesis of metabolically stable analogues (Piscitelli et  al. 2012). Complex medical products have been synthesized by

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laccases, like immunosuppressors (e.g., cyclosporin A), antibiotics (e.g., penicillin X dimer and cephalosporins), and anticancer drugs (e.g., vinblastine, mitomycin, and actinomycin) (Kunamneni et al. 2008). By exploiting the oxidative potential of laccases, many products have been developed in the field of cosmetic industry and personal care products (Javed et al. 2017). In addition, laccases have been applied for hair bleaching and dying. Therefore, some cosmetics and dermatological formulations containing laccases were patented and used for skin lightening (Morel and Christie 2011).

5.4  Manganese Peroxidase (EC 1.11.1.13) One of the most widely used substrates for P. ostreatus cultivation is wheat straw, especially in European country. As a white-rot fungus, it can be cultivated of agro-­ industrial lignocellulosic wastes (Sanchez 2010). It also has been described as selective and simultaneous lignocellulose degrader (Banfi et  al. 2015). P. ostreatus start to grow at first by easily utilizable and soluble materials from intra- and intercellular spaces from the plant substrate tissues. After nutrient depletion, the fungal biomass starts to degrade the plant cellulose and lignin polymers for further growth (Banfi et al. 2015. Lignin degradation is an oxidative and nonspecific process usually carried out by white-rot fungi. Ligninolytic enzymes, LiP, manganese peroxidase (MnP), and laccase, catalyze the one-electron oxidation of lignin units producing aromatic radicals that lead to non-enzymatic depolymerization. Ligninolytic enzymes, LiP, are the only enzyme able to oxidize directly non-phenolic ones in the presence of certain compounds (Sarkar et al. 1997). However, P. ostreatus produces two manganese peroxidase (MnP) isoenzymes when grown in solid stationary condition on poplar sawdust (Giardina et al. 2000).

5.4.1  Characterization of MnP MnP was first described in P. chrysosporium (Kuwahara et al. 1984). Peroxidases with Mn2-independent activity on phenolic and non-phenolic aromatic compounds were first reported from Pleurotus species (Martínez et al. 1996; Camarero et  al. 1996; Sarkar et  al. 1997; Palma et  al. 2000). These enzymes have been considered as MnP, despite differences in catalytic properties with P. chrysosporium MnP, because they show the highest affinity for Mn and the existence of a characteristic Mn2 interaction site has been shown in the molecular model of P. eryngii Mn2-­oxidizing peroxidase (Ruiz-Dueñas et al. 1999). Manganese peroxidase (MnP) is the most common lignin-modifying peroxidase produced by almost all wood-­colonizing basidiomycetes causing white-rot and various soilcolonizing litter-­decomposing fungi (Hotrichter 2002; Hatakka 1994; Peláez et al. 1995).

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Fig. 5.5  3D Crystal structure of manganese peroxidase IV from Pleurotus ostreatus. (Source: http://www.rcsb. org/pdb/explore. do?structureId=4BM4)

MnP is developed by multiple forms of glycosylated heme protein with molecular weight between 40 and 50 kDa and secreted by ligninolytic fungi (Fig. 5.5). From that point, MnP preferentially oxidizes manganese (II) ions (Mn2+), commonly present in wood and soils, into highly reactive Mn3+. The trivalent Mn3+ is then stabilized by fungal chelators such as oxalic acid. Like other peroxidases, MnP is sensitive to high concentrations of H2O2, but it can be rescued by Mn3+ ions but become quite unstable in aqueous media. To overcome this drawback, they form complexes with organic acids naturally secreted by the fungus, such as malonic or oxalic acid (Hofrichter 2002; Wong 2009). There are three mnp genes, namely mnp1, mnp2, and mnp3, that have been isolated, and their products were characterized from P. ostreatus (Asada et al. 1995; Giardina et al. 2000; Irie et al. 2001). Besides, both MnP2 and MnP3 have been purified (Kamitsuji et al. 2004; Sarkar et al. 1997). However, it is worth to note that MnP2 from P. ostreatus (strain Florida) did not oxidize non-phenolic compounds such as veratryl alcohol (Giardina et al. 2000). Therefore, the difference in enzymatic properties between these two MnP2 isozymes might be due to the differences in posttranslational modification (Kamitsuji et al. 2005). In fact, the homologous expression system developed by Tsukihara et  al. (2006b) reflects the posttranscriptional modifications, secretion, and stability in the p­ hysiological condition during the enzyme production process in P. ostreatus. Irie et al. (2001) pointed out the successful molecular breeding of a P. ostreatus strain with high MnP productivity, using an expression system employing the promoter and terminator sequence of sdi1.

5.4.2  Mechanism of Action of MnP Manganese peroxidase is a heme-containing glycoprotein which requires hydrogen peroxide as an oxidant. Manganese peroxidase basically oxidases Mn2+ to Mn3+ in the presence of H2O2 and organic acid chelators, such as lactic acid. Mn3+, in turn,

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oxidizes a variety of phenolic substrates (Hotrichter 2002). MnP catalyze the oxidation of several monoaromatic phenol and dyes but depends on both divalent manganese and certain types of buffer. This enzyme has been shown to have the ability to oxidize non-phenolic substrates in the presence of mediators. Due to the intense activity in oxidizing a wide variety of aromatic compounds, it exhibited many potential industrial applications. Mushrooms also produce aryl alcohol oxidase (AAO), an enzyme participating in hydrogen peroxide production. MnP production by different Pleurotus species, which were grown in both submerged and solid-state culture, was shown (Stajic et  al. 2006a, b). MnP are considered to be the key enzymes in the lignin degradation system (Irie et al. 2001). The ability to synthesize MnP is common among distinct taxonomical groups of basidiomycetes. In addition, there are fungal MnP producers from rather exotic habitats such as decaying sea grass, cooling tower wood, and brown coal. The molecular weight of MnP is usually within the range between 38 and 62.5 kDa, but most purified enzymes have Mwt around 45 kDa. MnP is often produced in multiple forms, and these isoforms differ in their isoelectric points which are usually rather acidic (pH 3–4).

5.4.3  Applications of MnP Manganese peroxidase (MnP) is one of the two extracellular peroxidases secreted by the lignin-degrading fungus P. ostreatus. Manganese peroxidase has great potential for industrial waste remediation application such as degrading refractory industrial pollutants (Asgher et al. 2013; Hotrichter 2002). As such, direct disposal of olive mills waste to aquatic bodies’ results in environmental deterioration due to the large amount of organic loading discharged (Fountoulakis et al. 2002). MnP was very helpful in bioremediation of olive mills’ waste. In other applications, MnP is also valuable for decolorization of synthetic dyes and significant potential application for textile industry (Susla et al. 2008; Praveen et al. 2012; Saravanakumar et al. 2013). These complexes function as diffusible oxidants that, in turn, can oxidize terminal phenolic substrates and also possibly non-phenolic substituents via a radical mediator (Giardina et al. 2000).

5.5  Versatile Peroxidase (EC 1.11.1.16) The first study reported on the presence of versatile peroxidases (VP) in Pleurotus eryngii was revealed by Martinez et al. (1996), and the complete purification of the enzyme was carried out by Min et al. (2010). To date, little evidence has been found associating versatile peroxidase in Pleurotus sp. except in few researches established for P. ostreatus and P. pulmonarius (Moreira et al. 2007). Versatile peroxidases are glycoprotein with hybrid properties, included in a group of H2O2-dependent ligninolytic heme peroxidases (POXs) also known as lignin manganese peroxidases,

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since it combines together with catalytic properties of lignin peroxidases (LiP, EC 1.11.1.14) and manganese peroxidases (MnP, EC 1.11.1.13) (Busse et al. 2013). It is characterized by the similar properties of MnP in oxidizing phenolic and nonphenolic aromatic compounds as well as its efficient oxidation of Mn2+ to Mn3+ (Tsukihara et  al. 2006a). In another major studies, their capabilities in oxidizing veratryl alcohol, the typical lignin peroxidase (LiP) substrate (Camarero et al. 1999) and simple phenols, which are the substrates of Coprinopsis cinerea peroxidase (CIP) has been reported (Ruiz-Dueñas et al. 2009). These could conceivably then recognized that versatile peroxidase as a new family of ligninolytic peroxidases. In consequence of this, while sharing almost identical heme environment together with lignin peroxidase (LiP) and manganese peroxidase (MnP), VP differentiates in the catalytic sites in their molecular structure. VPs form an attractive ligninolytic enzyme group due to their dual oxidative ability to oxidize Mn(II) and also phenolic and non-phenolic aromatic compounds (Dashtban et al. 2010). This dual substrate specificity makes VP powerful to oxidize a variety of high and low redox potential substrates (Ruiz-Dueñas et al. 2011).

5.5.1  Biochemical Characteristics of VP The biochemical properties of only a few versatile peroxidases of Pleurotus sp. have been analyzed and studied in details especially in P. eryngii, which is superior than VP reported in P. chrysosporium (Camarero et al. 1999; Min et al. 2010). However, the Pleurotus VP isoenzymes were first described as MnP isoenzymes due to their similarity to Mn-oxidizing activity (Martínez et al. 1996). This has shown that VPs have high affinity for manganese and dyes, and strongly oxidized 6-­dimethoxyphenol (DMP) and veratryl alcohol (VA) in a manganese-independent reaction (Moreira et al. 2005). The catalytic cycle of VP consists of three steps as described clearly in Bjerkandera adusta influenced by pH-independent and pH-dependent steps in the conversion process (Ertan et al. 2012). The cleavage of H2O2 by heme Fe3+ to Fe4+ led to the formation and reduction of compound I and II intermediates. With the excessive of H2O2 in the absence of reducing substrate at low pH (3.0-3.5), the conversion of compound II to compound III will be carried out which lead to the inactivation of the enzyme (Fig. 5.6). This supports the theory of involvement of LiP and MnP in the catalytic cycle activities of VP (Wong 2009).

5.5.2  Structural Characteristics of VP Comprehensive and complete genome sequences for P. ostreatus were done in JGI (U.S.  Department of Energy, Office of Science, Joint Genome Institute; http:// genome.jgi-psf.org). This facilitated the identification of nine genes encoding members of short MnP- and VP-encoding gene family in the inventory of heme

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N

N

H2O2

Fe(III) N

H2O

O

N

•+

Fe(IV)

pH independent

N

N

N

Mnp-1 / LiP-1 (compound I )

VP resting + VA•

2H2O VAD + 2VA•

N

Mn+2 VA

Mn+3

VA Mn+3

VA

•+

Mn+2 N

•+ O2

N

N

Fe(III) N

N

VP-III (compound III)

O

N

Fe(IV) 2H2O

excess 2H2O2

N

N

MnP-II / Lip-II (compound II)

Fig. 5.6  Catalytic cycle of VP involving the activities of LiP and MnP; VA veratryl alcohol; VAD veratryl aldehyde (Ertan et al. 2012)

peroxidases of this mushroom (Ruiz-Dueñas et al. 2011). Gene-expression analysis of P. ostreatus has revealed to have five Mn2+-dependent peroxidases encoded with mnp3, 6, 7, 8, and 9, while for versatile peroxidases encoded with MNP1, 2, 4, and 5, VPs all have the related gene and protein structure (Salame et al. 2012). Among these VPs, MNP3 and MNP2 were demonstrated to be unique in their ability to oxidize high molecular weight compounds such as Poly R-478 and RNaseA (Kamitsuji et al. 2005). According to Ruiz-Dueñas et al. (2011), two model types of VPs (137760 and 137766) show genetic variations in only one or two amino acids from the whole sequence of P. ostreatus MnP1 (GenBank AAA84396) (Asada et al. 1995) and MnP2 (GenBank CAB51617) (Giardina et  al. 2000), respectively. A recent review reported that P. ostreatus genome exhibited three versatile peroxidases (VPs) and six manganese peroxidases (MnPs) in which two of these crystal structures, VP1 and MnP4, designated dissimilarities at 1.0 to 1.1 Å (Fernández-­ Fueyo et  al. 2014). The significant differences between these two isoenzymes involved not to their kinetic constants only, however including the activity T50 and residual activity at both acidic and alkaline pH. The findings from the previous studies make several contributions to a growing body of literatures on VP in Pleurotus sp. in which the presence of VP has strongly replaced the role of LiP in lignin degradation for P. ostreatus. However, complete purification of VPs from P. ostreatus is not fully reported yet. Versatile peroxidases from P. eryngii are gaining much attention so far as it is first revealed to have high redox potential compared to other Pleurotus group. Generally, versatile peroxidase from P. eryngii was reported to

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have 40  kDa molecular weight and isoelectric point of 4.1. ABTS (2-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) is the best to be used as a substrate, and the maximal enzyme activity was achieved at 50 °C, pH 3.0 (Min et al. 2010).

5.5.3  Applications of VP Pleurotus sp. has been described as being able to degrade lignin selectively. The limited attack to cellulose makes them very interesting in different physicochemical and biotechnological pretreatment applications related to the use of plant biomass, including the integrated lignocellulose biorefineries for the future production of chemicals, materials, and biofuels. The versatile catalytic of VP in biotechnological and industrial application is due to its high redox potential and unique characteristics in degrading the aromatic compounds without the use of redox mediators and the presence of polyvalent catalytic sites (Ravichandran and Sridhar 2016). The ability to direct degradation of a variety of recalcitrant compounds that other peroxidases are not able to oxidize directly represents VPs as unique enzymes. Their potential applications are not limited to Mn (II) oxidation only but also extend to VA; phenolic, non-phenolic, high molecular weight compounds; as well as dyes in Mn-independent reactions (Wong 2009). In dye treatment, the capacity of P. ostreatus in the degree of decolorization is depending on the type and concentration of dyes, the ligninolytic enzymes produced, and reaction conditions. An example of this is the study carried out by Vishwakarma et al. (2012) in which the treatment of azo dye (direct blue 14) with enzymes of P. ostreatus has been reported. They reported that immobilized enzymes can reduce color up to 99% only in 18 hours. However, in contrast to Kahraman et al. (2012), the color reduction of indigo carmine dye varied from 93% to 64% as concentration was increased from 50 to 500 mg/L when using dead biomass. The ability of P. ostreatus to degrade aromatic pollutants such as 2,4-dichlorophenol (2,4-DCP) and benzo(a)pyrene ([B(a)P] was reported by Rodrıguez et al. (2004). An increasing number of studies have found that versatile peroxidase from Pleurotus represents an important preference with respect to LiP since no mediator such as veratryl alcohol could be used.

5.6  Aryl Alcohol Oxidase (EC 1.1.3.7) Aryl alcohol (AAO) is an extracellular flavoprotein (flavin adenine dinucleotide (FAD)-dependent proteins) providing the H2O2 required by ligninolytic peroxidases for fungal degradation of lignin (Xiao et  al. 2017). Aryl alcohol oxidase (AAO) which is thermodynamically favorable (Hamdane et al. 2015) has other names in common use including veratryl alcohol oxidase and aromatic alcohol oxidase. Aryl alcohol oxidase was first detected in the culture liquid of Polystictus versicolor (a

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synonym of Trametes versicolor) (Farmer et al. 1960) before it is known as best produced in three species of the genus Pleurotus: P. sajor-caju (Bourbonnais and Paice 1988), P. eryngii (Guillen et al. 1992), and P. ostreatus (Sannia et al. 1991). The isolation of the enzyme from basidiomycetes has been first carried out by Janssen et al. (1965) which later designed this enzyme as “alcohol oxidase” included as the prosthetic group of the enzymes (Janssen and Ruelius 1968). Preliminary work in this field was focused mainly on alcohol oxidase in mycelial extracts of a basidiomycete grown in submerged culture (Kerwin and Ruelius 1969). In 2000, Varela et al. published a paper in which they described a gene of aryl alcohol enzymes in the species of Pleurotus and Bjerkandera. Rodrıguez et al. (2004) define evidence that supported the involvement of AAO in lignin degradation as they observed the simultaneous production of AAO and VP in Pleurotus cultures. However, previous studies have not studied AAO in much detail, and the molecular characteristics are not yet clearly elucidated except in P. eryngii (Ferreira et al. 2009). P. eryngii is the first Pleurotus species that has been cloned (Martínez et al. 1994), and a prokaryotic heterologous expression system has been developed in order to obtain fully non-glycosylated AAO for further biochemical and structural studies (Ruiz-Dueñas et al. 2006). However, further study within the species had discovered the best producer of AAO is P. ostreatus compared to other species (Kumar and Rapheal 2011).

5.6.1  Biochemical Characteristics of AAO Aryl alcohol oxidase cooperates with laccase and other peroxidase in the production of hydroxyl radical, which is believed to be involved in the initial attack of lignocellulosic materials. This is to prevent the repolymerization of laccase oxidation products to occur in order to sustain the lignin degradation process (Goswami et  al. 2013). During degradation by Pleurotus species, H2O2 is generated from the expense of benzylic or other p system-containing primary alcohols such as p-anisaldehyde and p-anisyl alcohol into corresponding aldehydes (Ferreira et al. 2015). They point out that in addition to p-anisyl alcohol, the enzyme also oxidizes other polyunsaturated primary alcohols; where flavin reduction by substrate oxidation and re-­ oxidation of the reduce enzymes by oxygen before the release of the aldehyde product. Aromatic radicals that are produced during the reaction then act to catalyze subsequent degradation, which further generated potentially toxic molecules as defensive tool for the cells from the environment (Li et al. 2015).

5.6.2  Structure of AAO Depending on the substrate specificity, group of alcohol oxidase (alcohol: O2 oxidoreductase; EC 1.1.3.x) may contain four different categories: AAO which was categorized as aromatic alcohol oxidase and others which are short-chain alcohol

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oxidase (SCAO), long-chain alcohol oxidase (LCAO), and secondary alcohol oxidase (SAO) (Goswami et  al. 2013). However, SCAO and LCAO are reported as intracellular in nature. The crystalline structure of AAO was revealed in P. eryngii by Fernández et  al. (2009) which confirmed to share similar fold topology with other members of the glucose oxidase from Aspergillus niger (Frederick et al. 1990) and choline oxidase from Arthrobacter globiformis (Quaye et al. 2008). The use of single-wavelength anomalous diffraction of a selemethionine derivative obtained by Escherichia coli expression and in vitro folding had identified that this monomeric enzyme has two additional structural elements existing in the surroundings of its active site that modulate the access of substrates, which is absent in the structure of model GMC oxidoreductase glucose oxidase (Fernández et  al. 2009). These two domains were defined as FAD-binding domains, which interact non-covalently with the proteins, and substrate-binding domain, with a type of funnel-shaped channel that led the connection between the solvent and the flavin cofactor (Hernández-­ Ortega et al. 2012). The unique properties of AAO associated with its active site, which is able to bind over a wide range of aromatic ligands, include competitive inhibitors such as chavicol (4-allylphenol) and p-anisic (4-methoxybenzoic) acid, while 4-methoxybenzylamine was observed to be the best uncompetitive inhibitor (Ferreira et al. 2005). Until recently, there has been little interest in AAO. Aryl alcohol oxidases from P. ostreatus monomeric glycoproteins were reported to have 67 kDa molecular weight, with optimum pH, and the temperature was found to be around 6 and 40  °C, while Km value of AAO for oxidizing veratryl alcohol was determined to be 0.6 mM, respectively (Kumar and Rapheal 2011).

5.6.3  Applications of AAO The advantage characteristic of veratryl alcohol oxidase to oxidize various alcohols irreversibly and selectively without the necessity of cofactors in the catalysis has gained industrial interest for large-scale production (Goswami et al. 2013; Kerwin and Ruelius 1969). It is widely known that cellulose and hemicellulose in lignocellulose are the main carbon sources for P. ostreatus (Xiao et al. 2017). In a study by Kumar and Rapheal (2011), AAO activity was observed to be induced by aromatic amino acids and aryl alcohols up to a level of 289 U L−1. However, another component of lignocelluloses, lignin, was reported to inhibit the growth P. ostreatus (Barakat et al. 2012). The presence of lignin plays restricting roles in the efficiency of enzymatic hydrolysis of cellulose and hemicellulose (Kumar et al. 2012). This is supported by Feldman et al. (2015) who reported on the formation of toxic degradation products such as 5-hydroxymethylfurfural (HMF). However, by the involvement of aryl alcohol oxidase and other yeast dehydrogenases via heterologous gene expression, P. ostreatus was capable to metabolize and detoxify HMF while converting it into 2,5-bis-hydroxymethylfuran (HMF alcohol) and 2,5-furandicarboxylic acid (FDCA) (Feldman et  al. 2015). The involvement of AAO in lignin degradation requires either substrate from lignin-derived compounds; phenolic

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aromatic aldehydes and acids that being reduced to alcohol substrates, and aromatic fungal metabolites, respectively (Hernández-Ortega et al. 2012).

5.7  Heme-Thiolate Peroxidase (HTP) Heme-thiolate (or haem-thiolate) proteins belong to the class of thiolate-containing hemoproteins. Omura (2005) reported that heme-thiolate proteins contain six different classes: cytochromes P450, chloroperoxidase (CPO), nitric oxide synthases, cystathionine β-synthase, protein CoA, and eIF2α. The specialty for the heme-­ thiolate proteins is based on their function as oxidoreductases in the biological systems and the most versatile biocatalyst. The thiolate group usually comes from cysteine residue and is the axial ligand of heme iron, and the cysteine residue in heme-thiolate peroxidases acts as peroximal axial ligand. The only known heme-thiolate peroxidases (HTP) are CPO from ascomycete Leptoxyphium fumago that has been extensively studied (Smith et al. 2015; Ruiz-­ Dueñas et  al. 2011). CPO was first discovered by Hager and team from marine fungus Caldariomyces fumago (Shaw and Hager 1959; Morris and Hager 1966; Hofrichter and Ullrich 2006). CPO is composed of 299 amino acids and uses hydrogen peroxide for catalytic process under acidic environment. Research in genome sequences of ascomycete and basidiomycete has gathered many putative heme-­ thiolate CPO sequences, where many were being included in PeroxiBase, a database that has been created for heme and non-heme peroxidases (Ruiz-Dueñas et al. 2011; Koua et  al. 2009). However, a second heme-thiolate peroxidase has been reported. It is an aromatic peroxygenase (APO) and first reported in Agrocybe aegerita (Anh et al. 2007; Ullrich et al. 2004). APO and CPO values are based on their characteristics such as wide substrate specificity, ability to catalyze halogenation reactions, shared catalases and cytochrome P450 monooxygenases, and ability to oxygenate aromatic substrates. Although they shared the same catalytic properties of peroxidases, these two enzymes differ from typical heme peroxidases in terms of their amino acid sequence, catalytic activity, and structure. Therefore, they were classified under new heme-­ thiolate peroxidase superfamily. Three heme-thiolate peroxidase gene models were also found in fungus P. ostreatus from genus Pleurotus. All genes were found at sequences 2–4 with high amino acid identities that are similar to the unpublished putative CPO of A. bisporus (GenBankCAC03461) (Ruiz-Dueñas et al. 2011).

5.7.1  Biochemical Characteristics of Structure of HTP CPO of L. fumago have been identified with 21 amino acid N-terminal signal peptide and 52 amino acid C-terminal propeptide (have chaperon-like function). Its molecular structure has also been experimentally established and is available in

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Protein Data Bank. Therefore, despite the low amino acid sequence (approximately 20%), its structure has been used as template to get the theoretical models for the three putative P. ostreatus heme-thiolate peroxidases. Ruiz-Dueñas et  al. (2011) studied homology models of the three P. ostreatus heme-thiolate peroxidase and the crystallographic model of CPO from L. fumago in detail. It has been shown that all models have cysteine as axial heme ligand. The surrounding residues which stabilize the cysteine-ligand loop by hydrogen bonding are conserved at the proximal side. The presence of Ala and Cys residues contiguous to the axial Cys in 114464 could reinforce the hydrogen-bonds structure at this site. The models also show that Glu, Ser, and His are responsible for cation coordination in CPO and suggest different catalytic properties (activation rate and mechanism, substrate specificity, etc.) from P. ostreatus heme-thiolate peroxidases (Ruiz-Dueñas et al. 2011).

5.7.2  Applications of HTP Peroxidases and peroxygenases are beneficial biocatalyst and can play big role in the chemical modification of wide range of organic substrates, including regioselective and stereoselective oxygenations, which are difficult to do using conventional chemical reaction (Martínez et al. 2014). As a green technology, enzymes can act as catalyst in chemical syntheses to perform reactions under mild conditions, environmental friendly, and characterized by high specificity and selectivity compared to traditional chemical methods. The advantages of this enzyme are involvement in the production of fine chemical and the possibility to catalyze unspecific chlorination reaction. In addition, it also suitable in bromination and iodation of electrophilic organic substrates via hypohalous acid as actual halogenating agent (Hofrichter and Ullrich 2006).

5.8  Other Enzymes Produced by Pleurotus ostreatus Beside lignin-degrading enzymes, mushrooms have high capacity to produce a wide range of biocatalysts which were not reported in one type of microorganism. Previous studies reported on the production of different types of lipases, proteases, amylases, and polyphenol oxidases (Deepalakshmi and Mirunalini 2014). Nowadays, Pleurotus sp. has become a major cell factory in the enzyme cocktail production based on the ease of cultivation and ability to grow on most of agro-­ industrial wastes considered as sustainable and green substrate (Saini et al. 2014). Pleurotus ostreatus can produce and excrete a large number of enzymes including xylanases (Getachew et  al. 2016; Masutti et  al. 2015; Raymond et  al. 2015; Karthikeyan 2015; Rodrigues da Luz et al. 2013; Lim et al. 2013; Rodrigues da Luz et al. 2012; Morais et al. 2005; Qinnghe et al. 2004; Elisashvili et al. 2003), cellulases (Raymond et al. 2015; Lim et al. 2013; Radhika et al. 2013; Rodrigues da Luz

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et al. 2013; Daba et al. 2011; Sherief et al. 2010), pectinases (Masutti et al. 2015; Raymond et al. 2015; Morales Huerta et al. 2014; Sherief et al. 2010), proteases (Ergun and Urek 2017; Genier et  al. 2015; Lebedeva and Proskuryakov 2009; Palmieri et al. 2001), amylases (Ergun and Urek 2017; Lim et al. 2013; Akinyele et  al. 2010; Rashad et  al. 2009), and lipases (Wijayati et  al. 2017; Guarino and Sannia 2013). This part describes other biocatalyst produced by P. ostreatus under different cultivation strategies as well as the purifications process. The secretion of these cocktail of enzymes is dependent on the medium composition, pH, and temperature as well as the mycelial growth. For many researchers, solid-state fermentation (SSF) is the most appropriate method for mushroom cultivation to scale up and the production of extracellular enzymes such as laccase, manganese peroxidase, xylanase, cellulose, and amylase (Koyani and Rajput 2015). This is supported by the utilization of inexpensive lignocellulosic which can stimulate enzyme synthesis and supporting fungal growth (Koyani and Rajput 2015).

5.8.1  Cellulases Cellulases (EC 3.2.1.4) are known as hydrolase enzymes that cleave cellulose by catalyzing a series of cellulolytic reactions. Cellulases can be classified under different types such as (i) endoglucanases, endo-1,4-β-glucanase, carboxymethyl cellulose (CMCase), β-1,4-glucanase, endo-1,4-β-D-glucanase, β-1-4-endoglucan hydrolase, β-1,4-glucanase, cellobiohydrolases (CBH), and exogluconases, exocellulases, and β-glucosidases. Generally, cellulases were produced by most of the fungi and have the potential to degrade cellulose; at the same time, the same fungus can secrete other enzymes to hydrolyze lignin and hemicelluloses (Khalil et  al. 2011). Different cellulases have been produced and isolated from P. ostreatus as reported by many authors using SSF and submerged cultivation system and summarized in Table 5.1. Several studies reported the potential use of green substrate, which is in the form of agriculture wastes, for the production of cellulases by P. ostreatus (Radhika et  al. 2013; Rodrigues da Luz et  al. 2012). However, growth phase is critical factor governing the enzyme production in SSF. Singh et al. (2003) reported that cellulase production was highest during fruiting phase of strain Pleurotus sp. Furthermore, cellulases are widely used in textile industries, as well as detergent industries. In addition, they have also many applications in paper/pulp industries and pharmaceutical industries.

5.8.2  Xylanases Xylanases are enzymes which are able to break down the linear polysaccharide β-1,4-xylan into xylose monomer and were classified into two major families based on their protein sequence alignment. The two protein sequence alignments

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are Family 10 (F) and Family 11 (G). According to Alvarez-Cervantes et  al. (2016), the xylanases of GH10 family of P. ostreatus, BOFX60 are related to the GH10 and GH11 families, bBased on the smilimarities in amino acids, 233–318 and 180–193, respectively. Xylanases have gained attention because of their high potential applications in many industrial processes (Yadav et  al. 2015). Xylan plays very important role in insect nutrition and ruminant animal since xylan is the major component of hemicelluloses of most of the plants biomass and the polymer can be easily bioconverted into small digestible molecules. Applications for xylanase can be found in the pulp, feed, food, and paper industries (El-Enshasy et al. 2016; Kour et al. 2019b; Rana et al. 2019a, b). For instance, xylanases are beneficial in improving the quality of bread. It enhances the digestibility of ruminant feeds (El-Enshasy et al. 2016) and also applied widely in the pre-bleaching of kraft pulp. In general, the xylanase activities were higher when mushroom cultivated on waste material is compared to medium containing glucose or other low molecular weight fermentable carbohydrates. Different researchers reported that xylanase can be produced by P. ostreatus as summarized in Table 5.1. The study of Hazra et al. (1997) showed that 90% of the xylanase enzyme from the fungus was secreted out by mushroom T. clypeatus. Other research reported that P. ostreatus SYJ042 was able to produce xylanase. This xylanase was characterized by optimum temperature (40 °C) and stability at pH range between 3.0 and 9.0 (Qinnnghe et al. 2004). Getachew et al. (2016) studied xylanase purification by using ammonium sulfate in concentration between 30% and 80% (wt vol−1), and the best yield was obtained at 40%. The produced enzyme exhibited maximal activity at 50 °C and pH 6.0. This enzyme showed high affinity toward substrate used and can further be used for animal feed processing or other industrial applications.

5.8.3  Pectinases Pectinase (EC 3.2.1.5) or pectinolytic enzyme is a heterogeneous group of enzymes which is used to hydrolyze pectin, pectic acid, and oligo-D-galacturonate (Tapre and Jain 2014). In general, pectic substances can be found broadly in the plant cell wall including fruits and vegetables. The pectic substances are usually acidic and characterized as very high molecular weight (about 25–360 kDa) and negatively charged (Lakshminarasimha Reddy and Sreeramulu 2012). Pectinase is comprised of many groups and types. It is classified into three major types which are pectin pectinesterases (PE), protopectinase, and depolymerizing enzymes (Jayani et al. 2005; Garg et al. 2016). Usually, PE is used for pectic acid formation, whereas protopectinase is used for soluble pectin formation. Different research reported on the ability of P. ostreatus to produce and excrete pectinases using different substrates and cultivation systems (Table 5.1). Most of the studies were focused on increasing of the production yield and secretion of pectinase.

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The presence of extractive substances, derived from wheat bran and grape stalks (Massutti et al. 2015), sawdust (Sherief et al. 2010), sisal leaf (Raymond et al. 2015) as well as cell immobilization in polyurethane foam (PUF) (Morales et al. 2014), in culture medium can increase the production of pectinase by P. ostreatus. Sherief et al. (2010) reported on the cultivation of commercial strain of P. ostreatus on lignocellulosic rice straw and sawdust in plastic bags. The highest pectinase obtained was 21.42 U g−1 after 35 days in case of saw dust, and 13.80 U g−1 after 20 days incubation in rice straw culture. However, pectinase production by P. ostreatus is not widely studied like other enzymes in spite of its importance based on its wide use in textile industry, food processing industries, and for wine and juice clarification (Tapre and Jain 2014; Pasha et al. 2013; Jayani et al. 2005). Pectinylase (PL) EC 4.2.2.10 can only be produced by a small number of microorganisms such as Aspergillus, Penicillium, and Fusarium (Rashad et  al. 2011; Buyukkileci and Fernandez-Lahore 2011). Pectinylase is a pectinase which is only able to degrade highly esterified pectin into small molecules via β-elimination without methanol production. It was reported that P. ostreatus has also the capacity to produce PL. The produced enzyme had molecular weight of 23 KDa (Rashad et al. 2011). This pectin lyase showed high level of activity, with biochemical characteristic with alkaline pH (7.5), high optimum temperature (60 °C), and good affinity toward citrus pectin. Higher pectinase production was obtained by substrates from food processing at different varieties which is orange peel, lemon peel, apple pomace, and sugarcane bagasse using submerged fermentation by P. ostreatus NRRL-­ 366. The best substrate is lemon peel, giving 1750  U of exopolygalacturonase (exo-PGase and 750 U of pectin lyase (PL) per 1 g wet substrate after cultivation for only 4 days (Rashad et al. 2010).

5.8.4  Amylases Amylases (EC 3.2.1.x) or amylolytic enzymes are widely used for starch hydrolysis into glucose oligomers and glucose monomer. Amylases can be divided into four types of enzymes which hydrolyze different linkages and form various products. They are (i) α-amylase, β-amylase, glucoamylase, and pullulanase. Amylase can secrete from plants, microorganism and animals and share at least 30% of the world enzyme market (Deljou and Arezi 2016). However, enzymes from mushroom sources have subjected applications in industrial part. Amylase has a great potential application in industrial such as food, textile, paper, fermentation, detergent, and pharmaceutical industries. The production of β-amylase (Akinyele et al. 2010) and α-amylase (Ergun and Urek 2017; Lim et al. 2013: Rashad et al. 2009) has been reported from the species P. ostreatus (Table 5.1). α-Amylase is an enzyme with a potential to breakdown the α-1,4-glucosidic linkage of starch to produce small molecules such as maltose and malto-oligosaccharides (Deljou and Areze 2016). The isolation and characterization of the β-amylase was reported from the edible mushroom, P. ostreatus by Akinyele

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et al. (2010). The purification of amylase enzyme has been achieved by ion exchange chromatography method. The final finding was proven that the purified α-amylase by ion exchange chromatography on DEAE SephadexA 50 and gel filtration is thermostable with high potential for industrial use. The recent study of Ergun and Urek (2017) showed that the maximal amylase activities were obtained in SSF after 7 days of cultivation using dry potato peel waste after pretreatment with potassium hydroxide.

5.8.5  Proteases Proteases refer to a group of enzymes which catalyze hydrolytic reactions responsible for the degradation of protein into peptides and amino acids. Proteases are among widely used enzymes, which share about 65% of the global market based on catalytic properties (Yin et al. 2014). It was reported that Pleurotus ostreatus can secrete different types of proteases, which are metalloprotease (Shen et al. 2007; Liu et al. 2014; Dohmae et al. 1995); serine protease (ProA, EC 3.4.22.9), amino acid residues containing serine, aspartic acid, and histidine in the active center (Lebedeva and proskuryakov 2009; Palmieri et al. 2001; Dohmae et al. 1995); cysteine protease or thiol protease, containing cysteine and histidine residues at active center (Shin and Choi 1998); and aspartic protease (Yin et al. 2014). Researchers isolated different types of proteases from fruiting bodies, mycelia, and fermentation broth of P. ostreatus (Dohmae et al. 1995; Palmieri et al. 2001; Shen et al. 2007). Interestingly, the aspartic protease showed modeling crystal structure containing Po-Asp D110 and D295, with Asp110 and Asp295 in the active site. This study also has clearly shown that the structure had specific N- and C-terminal sequence (Yin et al. 2014). An aspartic protease from P. ostreatus (Pro-Asp) with Mwt of 35.3 KDa and isoelectric point of 4.57 consisted of 1324 nucleotides with an open reading frame (ORF) of 1212 bp encoding 403 amino acid residues (Yin et al. 2014). In addition to that, the crude aspartic protease had milk clotting activity and can be used in cheese making industry. MEROPS database defined that Aps was engaged into 16 different families according to amino acid sequence homology. Cysteine proteases or thiol proteases are enzymes that degrade protein containing a Cys-HisAsn at the active site. These proteases have histidine residue at active site acting as proton donor to enhance the nucleophilicity of the cysteine residue. Shin and Choi (1998) successfully purified cysteine protease and did full characterization of the enzyme. Cysteine protease from P. ostreatus is well known as leucine pNA (LPNA) consisting of ASGLXXAIL amino acid. This enzyme showed thermal stability up to 37 °C and the pH optimum in the range of 5.5–6.5. A serine protease (ProA, EC 3.4.22.9) and two metalloproteases, which are ProB (EC 3.4.99.32) and ProC (EC 3.4.24.4), were isolated from the fruiting bodies of P. ostreatus (Dohmae et  al. 1995). Paminieri et  al. (2001) reported that serine protease, present in the submerged culture of P. ostreatus, is different from other proteases purified from the fruiting bodies done by Dohmae et  al. (1995).

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Interestingly, these enzymes involved in POXA1b degradation and play a key role in the regulation of laccase activity and thus involved in lignin degradation by P. ostreatus. Furthermore, this finding was similar in the regulation of laccase enzyme in cultures of Trametes versicolor and Phanerochaete chrysosporium (Dosoretz et al. 1990). The purified serine protease (PoSI) was reported to have an isoelectric point of 4.5 and Mwt of 75 kDa, which is larger than other fungal proteases (Palmieri et al. 2001; Dohmae et al. 1995; Mellon and Cotty 1996). Metalloprotease is defined as protease enzymes with catalytic mechanism involve metal ions and mostly require zinc or cobalt for activity. PoFE (a fibrinolytic protease) consisting of 19 amino acids with N-terminal sequence of ALRKGGAAALNIYSVGFTS. Enzyme activity was increased by the addition of Ca2+, Zn2+, and Mg2+ ions, which is similar to the metalloprotease, extracted from fruiting body of P. ostreatus (GenBank Accession No. AAU94648.1) (Shen et al. 2007). In another study by Liu et al. (2014), they successfully purified fibrinolytic protease enzyme which exhibited thermostability up to 40 °C and pH optimum of 7.4. This result for optimum pH was similar to other fibrinolytic enzymes from Cordyceps militaris, Armillaria mellea, and Tricholoma saponaceum and application for medical applications, for thrombolytic treatment (Liu et al. 2014). In general, proteases from P. ostreatus have molecular weight between 18.2 and 75 kDA (Liu et al. 2014; Yin et al. 2014; Lebedeva and Proskuryakov 2009; Shen et al. 2007; Palmieri et al. 2001; Shin and Choi 1998; Dohmae et al. 1995). However, different types of protease produced by P. ostreatus are presented in Table 5.2. Table 5.2  Characteristics and potential application of some proteases from Pleurotus ostreatus

Proteases of Pleurotus sp. Metalloprotease

Purification Yield Isoelectric (%) Fold kDa point 6.5 876 32 ND

Metalloprotease

7.54

147

18.2 8.52

Serine protease (PoSi)

13

70

75

4.5

8 8 – 4

262 34.8 – 238

30 19 42.5 ND

9.0 8.35 – ND

ND

35.3 4.57

Metalloproteases ProA ProB ProC Cysteine protease

Aspartic protease ND

ND not determined or data not given in this study

Optimal conditions Potential Temp application 35 °C Thrombolytic therapy Heart disease 7.4 40 °C Fibrinolytic therapy 7–8 ND Acting as regulatory role for laccase activity Maintenance cellular disorder 6.5 5.6 5.6 5.5– 37 °C Cell 6.5 germination, morphogenesis ND ND Cheese making industry pH 6.5

References Shen et al. (2007) Liu et al. (2014) Palmieri et al. (2001)

Dohmae et al. (1995)

Shin and Choi (1998) Yin et al. (2014)

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5.8.6  Lipases Lipases (EC 3.1.1.3) constitute a very important class of hydrolytic enzymes. Lipases were classified into two types: (a) lipases those of enhanced enzyme activity in emulsion form and (b) lipases with the active site full of protein (Karigar and Rao 2011; Sharma et al. 2011). Generally, lipases are varied in size and sequence of amino acids in primary structure. Lipases are produced in all organisms, and natural function of lipases is to hydrolyze the fats into glycerol and fatty acids (Sharma et al. 2011). Therefore, they also play an important role in food, chemical, cosmetic, detergent, and paper making industries. In addition, they were used as indicator to determine hydrocarbon degradation for soil (Karigar and Rao 2011). Currently, P. ostreatus genome was successfully isolated and characterized. Sixty seven putative lipase coding genes have been annotated in this species (Guarino and Sannia 2013). Moreover, the ability to produce lipase enzymes by P. ostreatus was investigated by many authors. The ability P. ostreatus to produce extracellular lipolytic enzymes was studied by Guarino and Sannia (2013). They reported on the successful production of lipase by using 5% olive mill waste as the main carbon source to achieve a final titer of 30 U L−1 after only 5 days of cultivation. Mushroom enzymes were extracted from the SSF cultures, and the extracted enzymes were successfully immobilized using sodium alginate beads for continuous operation (Wijayati et al. 2017).

5.9  Conclusion and Future Prospects As presented in this chapter, P. ostreatus have strong production and excretion capacity for large number of enzymes from different groups. The process can be easily shifted to the targeted enzyme through proper design of the cultivation medium and optimization of the cultivation parameters. In addition, this mushroom has long history as food without any previously reported negative impact on human and animal health. Therefore, it received GRAS status from FDA which makes it very attractive for any food, feed, cosmetic, and pharmaceutical industries. In addition, based on the high capacity for concomitant production of hydrolase enzyme cocktail, P. ostreatus is considered as a very attractive biofactory in biorefinery for the hydrolysis of complex lignocellulosic biomass structure to fermentable sugars which is subsequently used for the economic production of different primary and secondary metabolites. However, further research is required for better understanding of the combined factors effect on the induction, production, and excretion of mushroom enzymes during large-scale cultivation. Acknowledgments  The authors would like to express their sincere acknowledgment for the support of MOE and UTM-RMC (Malaysia) through HICOE grant no. R.J130000.7846.4J262.

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Chapter 6

Extracellular Fungal Peroxidases and Laccases for Waste Treatment: Recent Improvement Shanmugapriya S., Manivannan G., Selvakumar Gopal, and Sivakumar Natesan

6.1  Introduction Fungi are widespread eukaryotic microorganism exploit subsidiary living conditions through their unusual extracellular enzymes capable of utilizing variable sources as substrates. Mostly, these extracellular enzymes degrade complex organic substances including cellulose, hemicellulose, lignin, phenols, pesticides, hydrocarbons, etc. into simple molecules for their carbon, energy, and nutrition (Burns et al. 2013). Among the various organic substances, lignin, hemicelluloses, and phenolic compounds are the major wastes as environmental pollutants. Due to the chemical complexity, lignins take a long time to its natural degradation. In nature, majority of the fungi in the phylum Basidiomycota have the enzymes such as laccases and peroxidases to actively degrade the lignin containing polyphenol waste from the environment, which have potential biotechnological applications. Three phenotypic groups of fungi specifically white-rot, brown-rot, and soft-rot fungi are the predominant groups which degrade lignin compounds variably. Among them, white-rot fungi execute complete lignin degradation with the ability to cleave Cα- Cβ, β-aryl, and C1-Cα bonds, including aromatic C-C bonds degradation c, but brown-rot fungi partially degrade lignin compounds by Fenton/Haber Weiss chemistry (Arantes et  al. 2012). However, white-rot fungi produce a special group of extracellular enzymes like laccases and peroxidases which entirely degrade lignin

Shanmugapriya S. · S. Natesan (*) Department of Molecular Microbiology, School of Biotechnology, Madurai Kamaraj University, Madurai, Tamil Nadu, India e-mail: [email protected] Manivannan G. Department of Microbiology and Biotechnology, SVN College, Madurai, Tamil Nadu, India S. Gopal Department of Microbiology, Alagappa University, Karaikudi, Tamil Nadu, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_6

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compounds. Also, fungal laccases and peroxidases have enormous potentials in environmental detoxification and bioremediation of phenolic compounds. Using these enzymes, the white-rot fungi can convert wood, paints, pesticides, and plastics etc. into nutrients, and it has lots of industrial application. Laccases have been regarded as a “Green Tool,” because they require molecular oxygen (O2) as the only co-substrate for biocatalysis and not hydrogen peroxide (H2O2) (Surwase et  al. 2016). In this chapter, the structure, functional properties, applications, and their recent advancements are being discussed.

6.2  Laccases Laccases (EC 1.10.3.2; 1,4-benzenediol: oxygen oxidoreductases) are extracellular copper-containing monomeric glycoproteins, which come under multicopper oxidase family (Solomon et al. 1996). It is otherwise called as polyphenol oxidase and blue multicopper oxidases. It oxidizes several aromatic and non-aromatic compounds especially phenols as well as diamines and hexacyanoferrate by using molecular oxygen as an electron acceptor. It was first demonstrated in the sap of the Japanese lacquer tree Toxicodendron vernicifluum (formerly Rhus vernicifera); hence, it is named as laccase. Its molecular weight ranges from 50 kDa to 100 kDa (Galhaup and Haltrich 2001).

6.3  Sources of Laccases Laccases have been generally present as extracellular and intracellular enzymes in several organisms ranging from microbes to higher plants. It was first discovered in plants by Yoshida in 1883. It is widely distributed in all plants, but not yet studied properly, because of the existence of several isoenzymes of laccase in lignified plant tissues (Gavnholt et  al. 2002) and difficulties of their detection and purification from crude plant extracts (Ranocha et  al. 1999), but it was well documented in fungi. Fungi are unique important class of eukaryotic microorganisms and synthesize unusual enzymes capable of performing chemically tricky reactions (Viswanath et al. 2008; Shraddha et al. 2011). Many fungal species are of great value and can remove toxic recalcitrant compounds in an environment-friendly manner. Laccases play an important role in bioremediation of toxic phenolic compounds (Singh et al. 2011) and degradation of recalcitrant xenobiotic compounds. The presence of laccase enzyme in fungi was first reported by Laborde in 1897 (Mayer and Harel 1979). Laccases are found in a wide range of fungi generally in white-rot fungi (Brijwani et al. 2010; Mayer and Staples 2002). Generally, paper and pulp industry effluents contain a large amount of chlorinated phenolic compounds that is formed from incomplete breakdown of lignin during pulp bleaching process. Different groups of fungi can remove such lignin

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and phenol from the effluent by producing extracellular enzymes like laccase, ­manganese peroxidase, and lignin peroxidase. Several studies suggested that filamentous fungi are the best choice than bacteria for the removal of soil pollutants because fungi can reach the pollutant efficiently than bacteria (Rubilar et al. 2008; Kour et al. 2019; Yadav et al. 2018). For example, laccase activity was detected in the cultures of fungi belonging to Basidiomycetes, Ascomycetes, and Deuteromycetes family (Table 6.1). The highest amount of laccase is produced by white-rot fungi (Leonowicz et al. 1997). Laccase enzyme has been reported in many fungal species such as Trichoderma reesei (Levasseur et al. 2010), Xylaria polymorpha (Nghi et al. 2012), Lentinus tigrinus (Pozdnyakova et al. 2006), Pleurotus ostreatus (Zhao et al. 2017), Cerrena unicolor (Kim et  al. 2002) T. versicolor (Minussi et  al. 2007; Rogalski et al. 1991), Trametes pubescens (Shleev et al. 2007) Melanocarpus albomyces (Kiiskinen et  al. 2002), Magnaporthe grisea (Iyer and Chattoo 2003), Aspergillus flavus PUF5 (Priyanka and Uma 2017), Trametes hirsuta (Tapia-Tussell et  al. 2011), Trametes ljubarskyi (Goh et  al. 2017), Aspergillus flavus (Kumar et  al. 2016), etc. Further, Abd El Monssef et  al. (2016) reported that the genus Alternaria, Aspergillus, Cladosporium, Penicillium, Rhizopus, and Trichoderma also produce laccases. Table 6.1  Examples of laccase producing fungi Class and division Agaricomycetes Agaricomycetes Agaricomycetes Sordariomycetes (Ascomycota) Agaricomycetes

Agaricomycetes Agaricomycetes Agaricomycetes Agaricomycetes Agaricomycetes Eurotiomycetes (Ascomycota) Sordariomycetes (Ascomycota) Agaricomycetes (Basidiomycota) Sordariomycetes (Ascomycota)

Source Trametes versicolor Phanerochaete chrysosporium BKM-F-1767 Pleurotus pulmonarius Chalara (syn. Thielaviopsis) paradoxa CH 32 Trametes pubescens

References Bourbonnais and Paice (1992) Srinivasan et al. (1995) Marques de Souza et al. (2002) Robles et al. (2002) Galhaup and Haltrich (2001); Rodriguez Couto et al. (2004); Osma et al. (2007) Kapich et al. 2004

Phanerochaete chrysosporium ME-446 Trametes hirsuta Phanerochaete chrysosporium NCIM 1197 Pycnoporus sanguineus Ganoderma lucidum Aspergillus carbonarius

Vikineswary et al. (2006) Murugesan et al. (2007) Sanjay et al. (2007)

Trichoderma harzianum WL1

Sadhasivam et al. (2008)

Pleurotus ostreatus, Trametes pubescens, Cerrena unicolor, and Trametes versicolor Trichoderma spp.

Osma et al. (2011)

Rodríguez Couto et al. (2006) Gnanamania et al. (2006)

Kalra et al. (2013)

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Other than fungi and plants, laccase enzyme has been reported from bacteria (Santhanam et al. 2011) (Yadav et al. 2016, 2019a, b), lichens (Laufer et al. 2009) and sponges (Li et  al. 2015a). Moreover, polyphenol oxidases with laccase-like activity have been found in oysters (Luna-Acosta et al. 2010) and insect cuticles (Lang et al. 2012). Functions of laccase enzymes are based on their source and the stage of life of the organism producing them.

6.3.1  Structure of Laccases Laccases are glycoproteins synthesized as monomer and containing four copper atoms (Fig.  6.1). After synthesis, laccase was modified with mannose which is accountable for about 10–50% of total weight of laccase. Carbohydrate may contribute structural stability of laccases (Mayer and Staples 2002). Glycosylation of laccases confers copper retention, susceptibility to proteolytic degradation, thermal stability, and secretion. The copper atoms of laccase are divided into three types. They are (i) Type 1 (T1) (ii) Type 2 (T2) and (iii) Type 3 (T3). Laccase contains one T1 and T2 and two T3 copper atoms. The catalytic mechanism of the laccase starts with the donation of an electron to the substrate by the T1 copper site, followed by an internal electron transfer from the reduced T1 to the T2 and T3 copper sites. During oxidation of substrate, molecular oxygen is reduced to water (Jones and Solomon 2015). The reduction reaction takes place at trinuclear cluster which is formed by the association of T2 and T3 copper atoms (Fig.  6.1). Type 1 copper confers blue color to the enzyme because of maximum absorbance around at 600 nm which is the result of the covalent copper–cysteine bond (Matera et  al. 2008). However, in fungal laccases, the axial ligand is leucine or phenylalanine, which possibly provides the mechanism for the regulation of enzyme activity (Claus 2004; Kumar et al. 2003; Enguita et al. 2003; Garavaglia et al. 2004). Type 2 is a non-blue copper and showed weak absorption in the visible spectrum (Niku-Paavola et al. 2004). Type 2 copper is coordinated by two histidine residues and is strategically

Fig. 6.1  Catalytic mechanism of laccase

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positioned close to Type 3 copper. Type 3 copper is a binuclear center that showed maximum absorbance at 330 nm in its oxidized form (Matera et al. 2008; Decker and Terwilliger 2000). Laccase molecular weight was determined to be in the range of 60–390 kDa (Kalme et al. 2009). The pH values vary between pH 4.32–6.51 and pH  5.32–6.19 (Cázares-García et  al. 2013; Moreno et  al. 2017). The catalytic domain of laccase is moderately conserved in diverse fungal species, and the rest of the enzyme structure shows high diversity (Gochev and Krastanov 2007; Moreno et al. 2017). However, laccases with variants in the active site are also reported in Pleurotus ostreatus (Palmieri et al. 1997). In this fungus, enzymes lacking the maximum absorption around 600 nm are usually classified as “yellow” or “white” laccases. Difference in the active center might confer these laccases have different functional properties of interest. Similar white laccase has also been reported in Deuteromycete fungus and Myrothecium verrucaria NF-05 (Zhao et  al. 2012). These white laccases contain only one Cu, one Fe and two Zn atoms (Palmieri et al. 1997; Zhao et al. 2012), but laccase enzyme of Phellinus ribis has one manganese atom instead of T1 copper atom (Min et al. 2001). Many fungi have variable number of laccase genes and they are typically inducible.

6.3.2  Mechanism of Laccase Activity Laccases have wide range of substrate-specific activity on ortho- and para-diphenol groups, as well as mono-, di-, and polyphenols, aminophenols, methoxyphenols, ascorbate, and aromatic amines with the linked four-electron reduction of oxygen to water (Bourbonnais and Paice 1990; Bourbonnais et al. 1995; Madhavi and Lele 2009). Oxidation of aromatic compounds occurs with the concurrent reduction of one O2 molecule to H2O. After four cycles of single-electron oxidation of aromatic compounds, it leads to formation of free radicals and reduction of one molecule of oxygen into two molecules of H2O (Fig. 6.1). Initially, the free radical is unstable and converted to a quinone in a second enzyme-catalyzed step. Alternatively, oxidized phenol-containing polymers may be partially degraded by nonenzymatic radical reactions. Partial degradation is due to the breaking of covalent bonds that join the monomer (Strong and Claus 2011). In the presence of small molecules, known as redox mediators, laccases improve their substrate specificity. Redox mediators are low molecular weight and small-sized molecules that are used as enhancer in the real electron transfer steps of enzymatic degradations process. It is a stable and reusable molecule. It increases the capability of an enzyme to react toward uncommon substrates (Majeau et  al. 2010). The following mediators are frequently used for laccase activity. (1) 2,2′-Azino-bis(3-ethylbenzothiazoline-6sulfonic acid) (ABTS), (2) 1-hydroxy-benzotriazole, (HBT) (3) 1-nitroso-2-naphthol-3,6-disulfonic acid (NNDS), (4) syringaldehyde, (5) 4-Acetylamino-TEMPO 4-hydroxy-TEMPO, (6) violuric acid (VIO), and (7) p-coumaric acid (Majeau et al. 2010).

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6.3.3  Applications of Laccases Laccases have several biological functions such as lignification of plant cell walls (O’Malley et  al. 1993), lignin biodegradation, detoxification of lignin (Baldrian 2006), virulence factors (Williamson et al. 1998), and copper and iron homeostasis (Stoj and Kosman 2003). Further, laccases have potential applications in bioremediation, paper pulp bleaching, finishing of textiles, biofuel cells, etc. Laccases exhibit transformation reactions like oxidation of functional groups to the heteromolecular coupling for production of new antibiotics derivatives or the catalysis of key steps in the synthesis of complex natural products (Xenakis et  al. 2016). However, fungal laccases are largely used for removal of phenols which present in wastewater (Pang et  al. 2016). The following are the significant areas of laccase applications. 6.3.3.1  Environmental Applications Extensive use of chemicals in agriculture and industrialization leads to release of different persistent, hazardous, bioactive, and bioaccumulative chemicals to the environment that causes pollution in land and water. These toxic chemicals create adverse effects on both human and other flora and fauna of soil and aquatic environment. Naturally, phenol and its derivatives are ever-present pollutants that arrived as wastewater from the effluents of industrial activities, such as pulp, petrochemicals, coal refineries, pharmaceuticals, production of resins, paints, and textiles (Rastegari et al. 2019; Yadav et al. 2017). They are highly toxic to aquatic organisms, including fish and shellfishes. The toxic effects of phenol are based on its chemical complexity and the range of free radical formation. It causes acute toxicity, with an effect of damaging DNA or enzymes inducing mutagenicity, carcinogenicity, and hematotoxic and hepatotoxic effects toward humans and other living organisms (Michałowicz and Duda 2007). Therefore, removal of phenol is essential to protect the environment and individual. Most of the conventional oxidation method (chemical method) removes the chemicals but has several drawbacks such as (i) use of hazardous chemicals for oxidation (ii) nonspecific, and (iii) undesirable side reactions. In the present scenario, biological treatment methods (enzymatic oxidation) are most suitable and widely used due to specific and biodegradable catalysts and enzyme reactions are carried out in mild conditions (Rodríguez Couto and Toca Herrera 2006). Laccases are capable of oxidizing, polymerizing, or transforming different xenobiotics including phenolic pesticides into less toxic molecules. Hence, it is a more apt enzyme in water (Majeau et al. 2010) and soil bioremediation. Laccase-based bioremediation has been proposed to remove toxins from textile, paper and pulp, food, distillery, pharmaceutical, printing, paint, and cosmetic industrial effluents. For the remediation, laccase could be used as (1) free enzyme, (2) immobilized enzyme, and (3) laccase containing cells to remove the pollutants from water (Mugdha and Usha 2012).

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6.3.3.1.1  Direct Use of Laccase Producing Fungi The growing fungi are used for waste (harmful pollutants) treatment. In this method, the fungal cells can adapt in the pollutant containing environment, utilize the harmful pollutants as carbon and energy requirements by synthesizing the specialized degradative enzyme, and digest or transform the pollutants to harmless. The introduced organisms produced enzyme that co-metabolized the targeted contaminants (Mugdha and Usha 2012). White-rot fungus Trametes versicolor is able to remove humic acids from a real humic-rich industrial-treated wastewater of a food-­ processing plant (Mostafa Zahmatkesh et al. 2017). 6.3.3.1.2  Cell-Free Laccase Enzyme Enzymes extracted from organisms are used to treat toxic pollutants as a pure form or crude extract. This method is advantageous, because of the following: (1) there is no need of acclimatization of source organisms to the toxic environment, (2) additional nutrients are not essential, (3) growth supportive environment is not required, (4) the growth rate of the source organism does not affect the amount of available enzyme to treat the effluent, (5) usage of cell-free enzyme makes it easier to standardize optimum treatment conditions, and (6) it is easy to handle and monitor the process (Karam and Nicell 1997). Crude enzyme extract is the least processed but contains active form of the enzyme. It is used to treat large-scale effluent treatment. Although usage of pure enzyme is highly expensive, crude enzyme preparation at larger volume should be used for industrial effluent treatment. In general, enzyme function is based on their conformation, under extreme conditions such as very high or low pH and temperature, high ionic strength, high concentrations of reactants, and presence of inhibitors; the structure of free enzyme may be modified and the enzyme becomes nonfunctional (Karam and Nicell 1997). Besides, use of free enzyme is hard to be taken from the residual reaction system for reuse (Wang et al. 2008). Therefore, immobilized preparation of enzymes and the whole-cell biomass for repeated long-time usage have been developed. 6.3.3.1.3  Immobilized Laccase Enzyme Immobilization of enzyme provides an increasing availability of enzyme to the substrate with better turnover over a significant period of time. The practice of immobilized enzymes in effluent treatment overcomes the cell-free enzyme because of the following reasons: (1) high stability, (2) easy to handle, (3) reusability, and (4) cost-effectiveness. Immobilization of laccase was done using different materials and used for bioremediation process (Table 6.2). Several immobilization techniques have been developed and adapted in enzyme immobilization for different applications. Theoretically, enzyme immobilizations are done by two basic methods: they are physical (entrapment, encapsulation, and cross-linking) and chemical

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Table 6.2  Immobilization of laccase enzyme and their applications Organisms name Trametes versicolor

Support Porous glass beads

Method of immobilization Entrapment

Trametes versicolor Aspergillus

Microfibers

Encapsulation

Trametes versicolor

Carbon-based mesoporous magnetic composites Gold electrode

Trametes versicolor

Green coconut fiber Adsorption

Coriolopsis gallica

Calcium alginate beads

T. versicolor

Nanostructured bacterial cellulose

Cyathus bulleri Polyvinyl alcohol Cercospora sp. Alginate SPF-6 Cyathus bulleri Polyvinyl alcohol

Adsorption

Applications References Dye decolorization Champagne and Ramsay (2010) Dye decolorization Dai et al. (2010) Dye decolorization Cristovao et al. (2011) Phenol removal Liu et al. (2012)

Covalent bonding

Biosensor for Phenolic compounds in industrial effluents Entrapment Remazol Brilliant Blue R, Reactive Black 5, and Bismarck Brown R Physical adsorption Biosensors and establishment of and cross-linking with glutaraldehyde bioreactors Entrapment Decolorization of azo dye acid red 27 Entrapment Dye decolorization Entrapment

Acid red 27

Trametes versicolor

ZnO/SiO2 nano-composite

Adsorption

Trametes versicolor

Poly(glycidyl methacrylate-co-­ ethylene glycol dimethacrylate) Chitosan macrobeads

Covalent bonding

Remazol Brilliant Blue B and Acid Blue 25 Bisphenol A

Covalent bonding

Anthracene

Magnetic nanoparticles Sol–gel matrix

Polymerization

4-chlorophenol

Entrapment

Dye decorization

Cross-linked enzyme aggregates Functionalized methacrylate– acrylate microspheres

Cross-linking

Malachite green

Covalent bonding

Biosensor for Tartrazine

Trametes versicolor Trametes versicolor Trichoderma harzianum strain HZN10 Cerrena sp

Sarika et al. (2014)

Daassi et al. (2014)

Chen et al. (2015) Chhabra et al. (2015) Vikram et al. (2015) Chhabra et al. (2015) Li et al. (2015b) Melo et al. (2017)

Azzurra Apriceno et al. (2017) Zhang et al. (2017) Zabin et al. (2017) Yang et al. (2017a) Mazlan et al. (2017)

(continued)

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Fig. 6.2  Enzyme immobilization methods

(adsorption, covalent binding) interactions with enzyme-supportive matrix (Matijosyte et al. 2010) (Fig. 6.2). (i) Entrapment is defined as the preservation of enzymes in a porous solid matrix, such as polyacrylamide, collagen, alginate, or gelatin (Dayaram and Dasgupta 2008; Lu et al. 2007; Niladevi and Prema 2008; Phetsom et al. 2009). (ii) In encapsulation, enzymes are protected in a semi-permeable polymer materials such as polyethyleneimine, sol–gel silica matrix, SiO2, and poly(GMA-co-­ nBA) microspheres (Qiu and Huang 2010; Rochefort et al. 2008; Crestini et al. 2010; Mazlan and Hanifah 2017). (iii) In the adsorption method, the enzyme immobilized onto a support is based on ionic and/or other weak forces of attraction. Adsorption is based on the pH and ionic strength of the medium and the hydrophobicity of the support (Xu et al. 2009; Fang et al. 2009; Forde et al. 2010). Adsorbents like Mobil Composition of Matter (MCM), cyano-modified silica (CNS), and Santa Barbara Amorphous (SBA-15) (Forde et al. 2010) and ion-exchange resins such as dextran, agarose, and chitosan (Cordova et al. 2009; Çorman et al. 2010; Ibrahim et al. 2007) are used for laccase immobilization. (iv) Covalent attachment is widely used enzyme immobilization method in which the chemical groups on the support surface are activated and react with nucleophilic groups on the protein (Arroyo 1998; Brady and Jordaan 2009). For example, silica-based supports such as kaolinite or mesoporous silica nanoparticles and GLU-activated silica nanoparticles (Champagne and Ramsay 2007; Liu et al. 2008; Salis et al. 2009), epoxy-activated resins such as Eupergit and Sepabeads (Berrio et  al. 2007; Russo et  al. 2008), Alumina and Granocel (Crestini et  al. 2010), and electrodes based on carbon, glass, gold, silver or graphite (Balland et al. 2008; Rahman et al. 2008; Szamocki et al. 2009) have been frequently used for laccase.

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(v) In cross-linked method, enzyme immobilization is possible with the use of bifunctional cross-linkers (Brady and Jordaan 2009). For example, dialdehydes, diiminoesthers, diisocyanates, and diamines activated by carbodiimide (Arroyo 1998) have been used. 6.3.3.2  Textile Effluent Treatment Colors and dyes are commonly used in textile, paper, food, cosmetics, and pharmaceutical industries. There are above 1,00,000 different human-made synthetic dyes available on the market, and worldwide, its production is around 7,00,000 tons/year (Hao et al. 2000). Wastewater from textile industries carries 10% of the dye stuffs which has been a significant cause of environmental pollution. Most of the synthetic dyes are lethal to living organisms due to their toxic and carcinogenic properties. The removal of dyes from industrial wastewaters could be very important due to their toxicity and carcinogenicity. The structural complexity of dyes makes effluent treatment difficult by conventional physicochemical methods due to their high cost and low effectiveness. Laccases are promising tools for the detoxification of dyes (Table 6.3) because it has shown efficient decolorization of different industrial dyes at low concentrations (Rodriguez et al. 1999; Reyes et al. 1999) without generation of harmful aromatic amines (Chivukula and Renganathan 1995; Wong and Yu 1999). Dye degradation ability of laccase depends on physiochemical parameters such as cell aging, concentration of dye, immobilized cells, etc. (He et al. 2004; Kalyani et  al. 2008). Laccase from Polyporus rubidus showed efficient decolorization of industrially important synthetic textile dyes in broad range of concentration without the use of redox mediators (Bayoumi et  al. 2014). Immobilized laccase of Paraconiothyrium variabile has pH and thermal stability and exhibited efficient decolorization of Acid Blue 25 and Acid Orange 7 (Mirzadeh et al. 2014). Complete decolorization of malachite green was achieved with Cerrena sp. laccase CLEAs (cross-linked enzyme aggregates) at 60 °C (Yang et al. 2017a). Trametes versicolor CBR43 can decolorize different types of dyes such as acid disperse and reactive textile dyes by producing laccase and Mn-dependent peroxidase (Yang et al. 2017b). Laccase enzyme from Cerrena unicolor strain GSM-01has been purified and identified that laccase is a monomeric protein of 63.2 kDa, their optimal pH and temperature is 2.6 and 45  °C, respectively and effectively decolorize bromothymol blue, evans blue, methyl orange, and malachite green (Wang et al. 2017). 6.3.3.3  Paper Industries Large amount of phenolic compounds such as lignin and their derivatives containing effluent has been discharged from the paper industries. Commonly, chemical bleaching method is used to remove lignin. In this process, chlorine is used, but chlorine formed bond with lignin and produce toxic organochloro-complexes like chlorolignins, chlorophenols, chloroguiacols, and chloroaliphatics. Large volume

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Table 6.3  Decolorization of various dyes by laccase Organism name Ganoderma lucidum Trametes versicolor Aspergillus ochraceus NCIM-1146 Paraconiothyrium variabile

Dye name Ramazol Black B and Ramzol Orange 16 Orange G Methyl Orange

References Murugesan et al. (2009) Casas et al. (2010) Telke et al. (2010)

Bromophenol blue

Trametes versicolor

Reactive Black 5

Armillaria sp. F022

Reactive Black 5

Trametes trogii

Acid Orange 51

Coprinopsis cinerea Aspergillus niger Cerrena sp. circulans BWL1061 Ganoderma sp. Pleurotus ostreatus MTCC 142 Paraconiothyrium variabile

Methyl Orange Basic fuchsin Malachite Green Direct Blue E Congo Red Acid Orange 67, Disperse Yellow 79, Basic Yellow 28, Basic Red 18, Direct Yellow 107, and Direct Black 166 Reactive Magenta HB

Vinoth Kumar et al. (2011) Bibi and Bhatti (2012) Hadibarata et al. (2012) Dalel Daassi et al. (2013) Tian et al. (2014) Rani et al. (2014) Yang et al. (2015) Iyer et al. (2016) Das et al. (2016) Forootanfar Hamid et al. (2016)

Talaromyces funiculosum (M2F) Marasmiellus palmivorus Marasmius cladophyllus

Reactive blue 220 (RB)and Acid blue 80 (AB) Remazol Brilliant Blue R, Orange G, and Congo red

Ankita Chatterjee et al. (2017) Cantele Cantele et al. (2017) Ngieng Ngui Sing et al. (2017)

of dark colored wastewater is produced at the end of bleaching process. Dark colored, toxic wastewater of paper industry are highly hazardous and also create environment pollution. Physical and chemical methods such as ultrafiltration, ion exchange, lime precipitation and aerated lagoons, and activated sludge methods are used to treat wastewater, but they are ineffective and expensive. This triggers the use of microbial laccase enzymes which fulfills the whole requirement and delignification, separates wood into its constituent fibers and lessens the toxic wastewater formation. Laccase-mediated delignification was introduced in the 1900 and uses mediators to oxidize the phenolic compound, lignin. The laccase enzyme itself can effectively break phenolic compound due to its high redox potential. The incorporation of mediator along with laccase increases the availability and dimension of the enzyme against non-aromatic ring-containing compounds. Several mediators ABTS, HBT, N-hydroxyacetanilide (NHA), and violuric acid have been used in delignification process. Effective mediators commonly

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possess N-OH functional group, and it should be biodegradable, specific, and economically feasible. Paper recycling reduces usage of source material and also cost. Laccase facilitated bleaching of old newsprint pulp with improved brightness by removing the lignin component (Hakala 2011; Xu et al. 2007, 2009). Combined action of laccase and hemicellulolytic enzyme exhibited efficient deinking and biobleaching of the pulp. The combination of xylanase and laccase is an effective tool for lessening the amount of lignin and related molecules from the pulp (Valls and Roncero 2009; Saxena and Chauhan 2016). Old newsprint is efficiently recycled with high brightness and low effective residual ink concentration (ERIC) content through a combination of the physical methods like sonication and microwaving and enzymatic method (laccase and xylanase) (Virk et  al. 2013). Pure laccase enzyme from T. versicolor decolorized the paper and pulp mills effluent to a clear light-yellow solution (Karimi et al. 2010), and various structurally different industrial dyes (Dhillon et al. 2012). The expression of delignifying enzymes only commenced complete glucose depletion (Girard et  al. 2013). B. adusta and P. chrysosporium have showed 100% delignification of industrial pulp and paper mill wastewater in 8–10 days, independent from pH control, with a significant reduction of total organic carbon (TOC) of the solution (Costa et al. 2017). 6.3.3.4  Bioremediation of PAHs Polycyclic aromatic hydrocarbons (PAHs) are xenobiotic compounds and consist of a benzene ring arranged linearly, angularly or in clusters (Zeng et al. 2011; Li et al. 2010; Yadav et al. 2018). Rapid industrialization and widespread use of pesticides for better agricultural output liberates large amount of PAHs, the main pollutant of soil, air, or aquatic environment. PAHs and their derivatives such as polychlorinated biphenyls (PCBs); benzene, toluene, ethylbenzene, and xylene (BTEX); polycyclic aromatic hydrocarbons (PAHs); trinitrotoluene (TNT); pentachlorophenol (PCP); and 1,1,1-trichloro-2,2-bis (4-chlorophenyl) ethane (DDT) are highly toxic for humans as well as carcinogenic to living beings. PAHs are less soluble in water and are resistant to biodegradation (Ihssen et al. 2015). Laccase enzyme may convert polycyclic aromatic hydrocarbons to their quinines and then carbon dioxide. Laccase converts acenaphthylene to 1, 2- acenapthalenedione and 1,8-napthelic acid when used along with mediator HBT (Madhavi and Lele 2009). Laccase-­ mediated removal of PAHs is an economically feasible, ecofriendly, and efficient bioremediation process. Polychlorinated biphenyls (PCB) are recalcitrant toxic substances, presently banned in most countries but used as pesticides and wood preservatives. T. versicolor degraded PCP efficiently after the initial uptake by the mycelia (Pallerla and Chambers 1998). Laccase-mediated degradation rate of PCBs is inversely proportional to number of chlorine. The 4–6-chlorine substituted hydroxyl-­ PCB is degraded by laccase in the presence of the mediator 2,2,6,6-­tetramethylpipe ridine-­N-oxy radical (Keum and Li 2004). Heterologously expressed Trametes sanguineus laccase in Trichoderma atroviride efficiently removed phenolic compounds present in industrial wastewater, bisphenol A (an endocrine disruptor) from the culture medium, benzo[a]pyrene, and phenanthrene (Balcázar-López et al. 2016).

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Enzyme immobilized on mesoporous nanofibers that were prepared by Vinyl-­ modified poly (acrylic acid)/SiO2 nanofibrous membranes exhibited a better triclosan removal (Xu et  al. 2014). Laccase enzyme from Trametes versicolor and Myceliophthora thermophile could degrade the hormones and endocrine disrupting compounds (EDCs) (Dennis Becker et al. 2017). Phenolic compounds present in industrial wastewater and bisphenol A (an endocrine disruptor) from the culture medium was removed effectively by the heterologously expressed Trametes sanguineus laccase in Trichoderma atroviride (Balcázar-López et al. 2016). Bjerkandera adusta has the ability to degrade aromatic xenobiotics (Sodaneath et al. 2017) and extractives (Kinnunen et al. 2017) have raised its biotechnological importance in wastewater treatments for lignin removal. The immobilization of Trametes versicolor laccase on carbon-based mesoporous magnetic composites was done by an adsorbing laccase into bimodal carbon-based mesoporous magnetic composites. Adsorption effects of the support were responsible for the quick removal rate in the first hour, and up to 78% and 84% of phenol and p-chlorophenol were removed in the end of the reaction, respectively, indicating that the magnetic bimodal mesoporous carbon is a promising carrier for both immobilization of laccase and further application in phenol removal (Liu et al. 2012). 6.3.3.5  Biosensor for Detection of Pollutants Researchers concentrated to device a system for deduction of phenolic compounds in the environment, food, and biomedical components by a user-friendly and cost-­ effective approach. Biosensors are suitable for monitoring contaminated area continuously with high specificity and sensitivity. Among the various biosensors, enzymatic biosensor has increased eventually, due to its substrate-specific catalytic activities. There are more number of biosensors available to detect phenolic compounds, in particular, horseradish peroxidase (Jaafar et  al. 2006), and tyrosinase. However, those enzyme biosensors have some disadvantages due to their lower structural stability and sensitive to reaction products (Rodríguez-Delgadoa et  al. 2015). On the other hand, laccase shows a strong claimant as a biosensor, having selective advantages over other enzymes including stability, catalytic efficacy (electron transfer reaction), and oxidized phenol and related compounds in the presence of O2 without any cofactors (Munteanu et al. 1998). Laccase can react with wide range of phenolic compounds; therefore, it has been used in biosensor technology to detect the presence of various phenolic compounds, oxygen, aromatic amines, morphine, codeine, catecholamines, and plant flavonoids even at low concentration (Leite et al. 2003; Jarosz-Wilkołazka et al. 2004; Ferry and Leech 2005). The smaller and more efficient biosensors are developed through controlled deposition and specific adsorption of laccase on different types of surfaces, at the micro and nanometer scale. There are two types of laccase biosensors: the first type monitors spectrum variation (at an absorbance of 600  nm) of the enzyme, and the second type monitors voltage changes from a modified oxygen electrode (Madhavi and Lele 2009). Immobilized alkali-tolerant laccase on nitrocellulose membrane can react with different substrates (syringaldazine, catechol,

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catechin, and L- DOPA) at their low concentrations (Singh et al. 2010). Oktem et al. (2012) immobilized laccase enzyme on Whatman filter paper No. 1 with coloring agent MBTH (3-methyl-2-benzothiazolinone) that is used for the identification of oxidation products of phenols by developing maroon-green colors.

6.4  Peroxidase Peroxidases (EC 1.11.1.7) are glycoproteins with a hematin compound as cofactor. This heme protein has iron (III) protoporphyrin IX as the prosthetic group. They catalyze hydrogen peroxide (H2O2)-dependent oxidization of the different organic and inorganic compounds. Its molecular weights range between 30 and 150 kDa (Bansal and Kanwar 2013). It is widely distributed in all living organisms like bacteria, fungi, algae, plants, and animals. Peroxidases have been applied to reduce pollution in environment. They have the potential to oxidize phenols, cresols, and chlorinated phenols and synthetic textile azo dyes in water. Phenolic compounds are degraded by lignin peroxides (LiPs) in the presence of H2O2 (co-substrate) and veratryl alcohol (mediator). In this degradation, H2O2 is reduced to H2O by accepting an electron from LiP (which can oxidize itself). The oxidized LiP returns to its native form (reduced) by gaining an electron from veratryl alcohol thereby veratryl aldehyde is formed. Veratryl aldehyde gets reduced back to veratryl alcohol by receiving an electron from the substrate (Karigar and Rao 2011).

6.4.1  Peroxidases Classification Peroxidases are classified into two types based on the presence or absence of heme group. They are (1) heme peroxidases and (2) non-heme peroxidases (Passardi et al. 2007a, b). Most of the known peroxidase are heme-containing peroxidases (>80%). Small proportion of the non-heme peroxidases such as thiol peroxidase, alkylhydroperoxidase, and NADH peroxidase existed. Heme peroxidases have further been classified into two superfamilies. They are (i) peroxidase-cyclooxygenase superfamily (PCOXS) and (ii) peroxidase-catalase superfamily (PCATS) (Passardi et al. 2007a, b; Zamocky and Obinger 2010) (Fig. 6.3). 6.4.1.1  Peroxidase-Cyclooxygenase Superfamily (PCOXS) Animal peroxidases like myeloperoxidase (MPO), eosinophil peroxidase (EPO), lactoperoxidase (LPO), and thyroid peroxidase (TPO) come under this peroxidase-­ cyclooxygenase superfamily. They revealed major role in the innate immunity, defense responses etc. (Dick et al. 2008; Soderhall 1999). In this superfamily peroxidase, the heme (prosthetic) group is covalently joined with the apoprotein (Pandey et al. 2017).

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Fig. 6.3  Classification of peroxidases

6.4.1.2  The Peroxidase–Catalase Superfamily (PCATS) Non-animal (plant, fungal, and bacterial) heme peroxidases come under this superfamily. At first, based on the sources of peroxidase, this superfamily peroxidase was called as the plant, fungal and bacterial heme peroxidase. But, the name of this superfamily was altered as peroxidase–catalase superfamily after identification of new cnidarians peroxidase. The non-animal peroxidases are further divided into three classes. They are Class I, II, and III peroxidases (Pandey et al. 2017). Class-I: They are intracellular peroxidases. It includes cytochrome c peroxidase (CCP1), ascorbate peroxidases and catalase peroxidase. Class-II: They are extracellular fungal peroxidases, like the lignin (LiP) and manganese (MnP) peroxidase. Both are secreted by white-rot fungi and involved in the degradation of lignin. Versatile peroxidases (VP; EC 1.11.1.16) displayed a hybrid molecular structure between LiPs and MnPs (Pérez-Boada et al. 2005). This group of peroxidases plays a major role in lignin biodegradation. Class-III: They are extracellular plant peroxidases. This includes horseradish peroxidases (HRP), peanut peroxidase (PNP), soybean peroxidase (SBP), etc. They play a major role in plant physiological processes such as cell wall metabolism, lignification, suberization, auxins metabolism, wound healing, etc. Class II and Class III peroxidases contain a N-terminal signal peptides, disulfide bridges, glycans, and calcium in their structure (Pandey et al. 2017).

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6.4.1.3  Fungal Peroxidase (Class II Peroxidase) 6.4.1.3.1  Peroxidase Structure Fungal peroxidases have a high-spin protoporphyrin IX (heme b) prosthetic group. It is located in-between the proximal (C-terminal) and distal (N-terminal) domains. The Fe group of this peroxidase is pentacoordinated form which is associated with the four pyrrole nitrogens in the imidazole group of the proximal histidine. The active site containing Fe coordination of peroxidases is highly conserved. At the active site, distal histidine assisted by an asparagine residue participate transfer of electrons from H2O to the heme. Redox potentials of the enzyme are determined by the length of Fe-imidazolic nitrogen (Fe–Ne2) bond. A higher basicity of the imidazole group gives a higher redox potential, except few (Choinowski et al. 1999). In general, the change of basicity is dependent on the electron extraction from the imidazolic nitrogen to the surrounding (proximal histidine) (Sinclair et al. 1992). This may differ in peroxidases such as LiP and MnP. Active site residues such as Ser177 and Asp201 weaken the basicity charges of imidazolic nitrogen bond. The peroxidases enzymes have four disulfide bonds; it was identified in LiP, ARP, and T. versicolor peroxidases (Kunishima et al. 1994; Limongi et al. 1995; Poulos et al. 1993), but MnP have fifth SH linkage extracellular peroxidases containing both N- and O-glycans; however, the glycosylation may be different in ­various peroxidases, which determines its isozymes (Kjalke et al. 1992). In addition, extracellular peroxidases contain two highly conserved, Ca2+-binding sites which have been located at the proximal and distal domains (Kunishima et al. 1994; Poulos et al. 1993). Presence of Ca2+-binding sites gives structural stability of the extracellular form of peroxidases (Banci 1997), and it gives more strength to the active site. Being extracellular enzymes, fungal peroxidases are synthesized with an N-terminal signal peptide. The LiP has eight Cys residues, all forming disulfide bridges. The enzyme molecule consists of eight major and eight minor α-helices and a limited β structure in the proximal domain. 6.4.1.3.2  Mechanisms of Peroxidase Activity Peroxidase catalyzes the oxidation of several of organic and inorganic compounds by using hydrogen peroxide which acts as the electron acceptor. The native form of enzyme (E) is oxidized to an active intermediate enzymatic form termed compound I (EI) with concurrent reduction of hydrogen peroxide (H2O2) to water molecule. Compound I oxidizes a phenol molecule to phenol-free radical and becomes compound II (EII). Compound II oxidizes another one phenol molecule to phenol free radical and returns to its original state (E) (Fig. 6.4). The formed free radical polymerizes and forms insoluble polyaromatic products which are precipitated by solid– liquid operations (Nicell 1994).

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Fig. 6.4  Steps involved in peroxidase catalytic activity

6.4.1.4  Lignin Peroxidase Lignin peroxidase (EC 1.11.1.14) comes under oxidoreductases family (Higuchi 2004; Martínez et al. 2005; Hammel and Cullen 2008). It was first observed in the basidiomycete fungi Phanerochaete chrysosporium by Burdsall in 1983 (Glenn et al. 1983; Tien and Kirk 1988). LiP is an extracellular H2O2-dependent heme protein (Gold and Alic 1993; Haglund 1999; Piontek et al. 2001; Erden et al. 2009). LiP enzyme contains 343–345 amino acids preceded by a 27-or 28-residue leader sequence (Gold and Alic 1993). LiP has less substrate specificity, reacting with different phenolic compounds. LiP is capable of oxidizing a variety of reducing substrates including polymeric substrates. It can oxidize methoxylated aromatic rings without a free phenolic group and produce cation radicals that undergo ring opening, demethylation, and phenol dimerization (Haglund 1999). LiP needs H2O2 to initiate the reaction, but not mediators to decompose high redox potential compounds. It is used for various industrial application and bioremediation process because of their wide substrate specificity and high redox potentials (Erden et al. 2009). Phenolic compounds are degraded by lignin peroxidase (LiP) in the presence of H2O2 (co-substrate) and veratryl alcohol (mediator). In this degradation, H2O2 is reduced to H2O by accepting an electron from the LiP (which can oxidize itself). The oxidized LiP returns to its native form (reduced) by gaining an electron from veratryl alcohol; thus veratryl aldehyde is formed. Veratryl aldehyde gets reduced back to veratryl alcohol by accepting an electron from the substrate (Fig.  6.5). White-rot fungi secreted lignin and manganese peroxidases degrade lignin. Lignin-­ degrading peroxidases are identified in a number of basidiomycetous fungi: Phanerochaete chrysosporium, Trametes versicolor, Pleurotus spp., Phlebia radiata, Coprinus spp., Bjerkandera adusta, Ceriporiopsis subvermispora, Dichomitus

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Fig. 6.5  Catalytic cycle of a LiP-mediator oxidation system. VA-OH veratryl alcohol, VA-CHO veratryl aldehyde

squalens and Arthromyces ramosus, Cylindrobasidium evolvens, and Daedaleopsis septentrionalis (Kimura et al. 1990; Pelaez et al. 1995; Varela et al. 2000; Kinnunen et al. 2016). 6.4.1.5  Manganese Peroxidases Manganese peroxidases (EC 1.11.1.13) also belong to oxidoreductase family (Higuchi 2004; Martínez et al. 2005; Hammel and Cullen 2008). It is a lignin-­degrading enzyme and was discovered in the fungus Phanerochaete chrysosporium following the discovery of LiP (Glenn and Gold 1985). MnP is present in all white-­rot fungi than lignin peroxidase (Hammel and Cullen 2008). MnPs are present mostly in white-rot fungi, such as Phanerochaete chrysosporium, Ganoderma sp., Pleurotus sp., Trametes sp., and Irpex lacteus (Manavalan et al. 2015; Janusz et al. 2013), Phyllosticta, Aspergillus, Fusarium, and Penicillium (Pant and Adholeya 2007), Hyphodontia sp., Pleurotus pulmonarius and Trametes ochracea (Kinnunen et al. 2016). The MnP enzyme is made up of 330–370 amino acids and has a leader peptide that consists of 21–29 amino acids (Li et al. 1999). It is a glycosylated heme protein; molecular weight is ranging from 38 to 62.5  kDa, and averaging at 45  kDa (Hofrichter 2002). Compared to LiP, MnP redox potential is low and oxidizes the substances with the use H2O2 which act as oxidant. Manganese acts as a mediator in the MnP catalytic cycle. Manganese peroxidase (MnP) activity involves the oxidation of Mn2+ ions to Mn3+. The Mn3+ is highly reactive and chelated with organic molecules such as oxalate and malates which are produced by the fungus (Kishi et al. 1994; Galkin et al. 1998; Mäkelä et al. 2002). Chelated Mn3+ oxidizes phenolic structures to phenoxyl radicals (Hofrichter 2002). 6.4.1.6  Versatile Peroxidase Versatile peroxidase (VP) (EC 1.11.1.16) is a heme-containing ligninolytic peroxidase and, as the name suggests, has the catalytic activities of both MnP and LiP and is able to oxidize Mn2+ similar of MnP and high redox potential non-phenolic

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compounds like LiP. It was first identified in the white-rot fungus Pleurotus eryngii (Martinez et  al. 1996). It was first purified from the fungi Bjerkandera (Moreira et al. 2007) and can transform lignin even in the absence of an external mediator. It is found in Physisporinus vitreus (Kong et  al. 2017), Phlebia radiata, P. pulmonarius, and Galerina marginata (Kinnunen et al. 2016). VPs can oxidize wide range of substrates with low and high redox potentials. Generally, VPs have hybrid molecular structures of LiP and MnP and provide multiple binding sites for the substrates (Camarero et al. 1999). VPs are superior than other peroxidases, because VPs efficiently oxidize phenolic compounds without the use of veratryl alcohol or Mn(II) that are needed for LiPs and MnPs activity, respectively (Ruiz-Duenas et al. 2009). Because of the catalytic versatility, VPs have been involved in the different biotechnological applications. VP can oxidize not only Mn (II), but also veratryl alcohol, phenolic, non-phenolic and high molecular weight compounds, including dyes in Mn-independent reactions (Asgher et al. 2008; Wong 2009). Like MnP, commercial applications of VPs are limited, because of their unavailability in large quantities which can be overcome by the use of DNA recombinant technology (Ruiz-Duenas et al. 2009).

6.4.2  Applications of Fungal Peroxidases Ligninolytic extracellular enzymes especially lignin peroxidase and manganese peroxidase have shown capability toward the degradation of various xenobiotics including dyes, chlorophenols, polycyclic aromatic hydrocarbons (PAHs), organophosphorus compounds, and phenols (Wesenberg et al. 2003), improve the digestibility of wood or straw for animal feed (Valmaseda et al. 1991), and reduce costs for the pulp and paper industry (Martinez et al. 1994) (Table 6.4). The other applications of LiP are delignification of feedstock for ethanol production, textile effluent treatment and dye decolorization, coal depolymerization, treatment of hyperpigmentation, and skin-lightening through melanin oxidation. The lignin peroxidase– graphite electrode biosensor systems have been established for recognition of recalcitrant aromatic compounds because of their effective bioelectrocatalysis (Ferapontova et al. 2006). The applications of peroxidases on various industries are given below. 6.4.2.1  Textile Industry Dye is a synthetic colored substance and is used by various industries to color paper, cotton, polyester, nylon, silk, leather, plastics, hair, etc. to which the dye binds and becomes an integral part. When unbound synthetic dyes are released into water, they cause pollution and cause skin allergy, cancer, and chromosomal aberrations for human beings and also affect plants that reduce photosynthetic activity by reflecting sunlight and affect germination rate of plants. Fungi can rapidly become

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Table 6.4  Applications of LiP and MnP enzymes Enzyme type LiP

Organism Phanerochaete chrysosporium Trametes versicolor

MnP

Phanerochaete chrysosporium

MnP & LiP

Phanerochaete chrysosporium

LiP

Fungal strain L-25

MnP

Compound removal Anisyl alcohol (Monomethoxylated Aromatic Compounds) Pulp Bleaching (oxidation of phenolic lignin substructures) Bentazon (3-isopropyl-1H-2,1,3 benzothiadiazin-4(3H)-one 2,3-dioxide) and MCPA (4-chloro-2methylphenoxyacetic acid) Procion Brilliant Blue HGR, Ranocid Fast Blue, Acid Red 119, and Navidol Fast Black MSRL Azo, diazo, and anthraquinone dyes

P. chrysosporium Burds BKM-F-1767 Phanerochaete chrysosporium P. chrysosporium RP78 Anthracophyllum discolor P. floridensis

LiP

Catechol derivative

MnP

Orange II

LiP & MnP MnP

Azo dyes

Phanerochaete chrysosporium Phanerochaete chrysosporium

LiP LiP MnP

Phenanthrene, anthracene, fluoranthene, pyrene and benzo(a)pyrene Coracryl brilliant blue Paper and pulp industry effluent treatment (Color and lignin removal) Congo Red

References Valli et al. (1990) Paice et al. (1993) Castillo (1997)

Verma and Madamwar (2002) Kariminia et al. (2007) Cohen et al. (2009) Sharma et al. (2009) Ghasemi et al. (2010) Acevedo et al. (2011) Chander and Kaur (2015) Singh et al. (2016) Bosco et al. (2017)

accustomed to varying nutritional sources because they can produce a significant number of intra- and extracellular enzymes that are needed to degrade several complex organic pollutants such as dye stuffs, polyaromatic compounds, organic waste, and steroids (Gadd 2001; Humnabadkar et al. 2008). The fungal system can be utilized in the treatment of colored and metallic textile effluents (Ezeronye and Okerentugba 1999) because they can produce nonspecific enzymes such as lignin peroxidase (LiP), manganese peroxidase (MnP) and laccase (Christian et al. 2005) that can mineralize dyes. Direct participation of fungal ligninolytic enzymes is necessary for the mineralization of dyes (Park et al. 2007). Versatile peroxidases (VPs) have shown effective direct oxidation of high redox potential dyes Reactive Black 5. Reactive Black 5 is oxidized by LiP only in the presence of veratryl alcohol, redox mediators (Heinfling et al. 1998). It can oxidize phenols, including hydroquinones (Gomez-Toribio et  al. 2001). Fungi produce enzymes extracellularly that confer decolorization ability of dyes. Lignin peroxidase of P. prosopidis degrades scarlet RR dye (Fernandes et al. 2008). LiP degrades dye by the following steps

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initial asymmetric cleavage, demethylation and denitrification and form N-ethyl-1  l3-chlorinin-2-amine which is further degraded by laccase. Pleurotus ostreatus can decolorize Remazol Brilliant Blue by producing peroxidase extracellularly (Shin et al. 1997). Ganoderma lucidum, a white-rot basidiomycete, could be capable of the decolorization of four dyes (Drimaren Blue CLBR, Drimaren Yellow X-8GN, Drimaren Red K-4B and Disperse Navy Blue HGL) and degradation of phenol with the aid of manganese peroxidase. Manganese peroxidase (MnP) from Ganoderma lucidum was expressed in Pichia pastoris and recombinant MnP can also degrade four textile dyes and phenol (Xu et al. 2017). Similarly, Pleurotus species have been reported for the production of lignin peroxidases, manganese peroxidases, and laccases enzymes, which play a vital role in the biodegradation and bioremediation (Pandey et al. 2012) of textile effluents. White-rot fungi, Pleurotus flabellatus, P. ostreatus, and P. citrinopileatus, are used effectively and efficiently for dye decolorization and bioremediation of recalcitrant substances (Singh and Srivastava 2016). MnP of Pleurotus pulmonarius could be able to decolorize the anthraquinonic dye Remazol Brilliant Blue R and the azo dye Congo Red. The enzyme is strictly dependent on Mn2+ for oxidizing phenolic and non-phenolic compounds. MnP of Pleurotus pulmonarius can be used for textile dye effluent treatment (da Silva et al. 2017). Agrawal et al. (2018) reported that Ganoderma lucidum will be an effective phenanthrene and pyrene degrader by producing ligninolytic enzymes (laccase, lignin peroxidase, and manganese peroxidase). 6.4.2.2  Paper and Pulp Industry Humic substances (HS) are formed from microbial breakdown of dead plant matter, mainly from lignin. HS tend to be polydisperse polymers of aromatic and aliphatic units that have been synthesized from the polymerization of intermediate lignin degradation products (Abdel-Hamid et al. 2013), and the polymer is physically and chemically structurally complex (Niladevi 2009). HS are existing in soil, marine, and groundwater environments and wastewater from industrial and municipal water treatment (Abdel-Hamid et al. 2013). In the pulp and paper industry, HS are produced from the chemical treatment of wood and removed using membrane filters during wastewater treatment, but they form biopolymer and produce blockage of filter leads to decrease of filtration flux rates (Sutzkover-Gutman et al. 2010). The enzymes are applied to remove the HS in ecofriendly method with low cost (Cavicchioli et  al. 2011). Peroxidases catalyze H2O2-dependent oxidation of aromatic polymers, including HS, by generating radicals which can break aromatic rings, ether and carbon–carbon bonds, and by causing demethoxylation (Wong 2009; Abdel-Hamid et al. 2013). Versatile peroxidase oxidizes complex polymeric humic substances (HS) derived from lignin (humic and fulvic acids) and industrial wastes (Siddiqui et al. 2014).

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6.4.2.3  Bioremediation of Toxic Agrochemicals The herbicide atrazine was converted to the less toxic compounds desethyl atrazine and hydroxyatrazine (N-dealkylated and hydroxylated metabolites, respectively), by the fungus Phanerochaete chrysosporium. Atrazine removal corresponded to the production of LiP and MnP from the fungus (Mougin et al. 1994). LiP and MnP of white-rot fungus P. chrysosporium can degrade the herbicide and isoproturon in in vitro and in vivo conditions (Del Pilar et al. 2001). MnPs from P. chrysosporium have the ability to break bentazon in the presence of mediators like Mn(II) and Tween 80. The herbicide glyphosate was oxidized by MnP that is produced by Nematoloma frowardii (Pizzul et al. 2009). This information evidently indicates the prospective application of lignin-degrading enzymes in the treatment of herbicides contaminated soil and water. Polycyclic aromatic hydrocarbons (PAHs) such as anthracene and pyrene are highly hydrophobic, but they are oxidized by MnP and LiP of wood rotting fungus Nematoloma frowardii. In the presence of low molecular mediator substances, the substrate range and the oxidation rate of LiP, MnP is increased (Günther et  al. 1998). When endocrine-disrupting chemicals and trace organic contaminants like pharmaceuticals and personal care products are released into water, it leads to bioaccumulation, acute, and chronic toxicity to aquatic living organisms and also causes severe effect on human health. Podoscypha elegans degrades lignin and organic pollutant by producing nonspecific extracellular ligninolytic enzymes such as laccase, lignin peroxidase (LiP) and manganese peroxidase (MnP). It can be used for the removal of pollutants from the environment (Nikki Agrawal et  al. 2017). MnP from Pleurotus ostreatus could detoxify aflatoxin B1 (AFB1) depending on the enzyme concentration and incubation period (Yehia Ramy Sayed 2014). Non-lignolytic filamentous fungus Penicillium sp. CHY-2 can degrade different aliphatic and aromatic hydrocarbons. Penicillium sp. CHY-2 efficiently degrades decane than octane, dodecane, ethylbenzene, butylbenzene, naphthalene, acenaphthene, and benzo[a]pyrene by producing MnP enzyme. The relative molecular mass of MnP enzyme from Penicillium sp. CHY-2 is estimated to be 36 kDa, and the native form of MnP is a monomer (Govarthanan et al. 2017).

6.5  Conclusion and Future Prospects Fungal laccases and peroxidases are a promising biocatalyst, used as a better alternative for conventional chemical processes in the treatment of lignin degradation, wastewater treatment, decolorization, and detoxification of textile dyes and biosensor preparation to detect the environmental pollutant. Their substrate range is fairly wide and immobilization technology increases enzyme stability and to achieve its reuse. Acknowledgments  The authors acknowledge to DST-PURSE Phase-II for providing computer facilities to prepare this review. There are no conflicts of interest.

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Pang S, Wu Y, Zhang X, Li B, Ouyang J, Ding M (2016) Immobilization of laccase via adsorption onto bimodal mesoporous Zr-MOF. Process Biochem 51(2):229–239 Pant D, Adholeya A (2007) Enhanced production of ligninolytic enzymes and decolorization of molasses distillery wastewater by fungi under solid state fermentation. Biodegradation 18:647–659 Park C, Lee M, Lee B, Kim SW, Chase HA, Lee J  (2007) Biodegradation and biosorption for decolorization of synthetic dyes by Funalia trogii. Biochem Eng J 36:59–65 Passardi F, Theiler G, Zamocky M, Cosio C, Rouhier N (2007a) PeroxiBase: the peroxidase database. Phytochemistry 68:1605–1611 Passardi F, Zamocky M, Favet J, Jakopitsch C, Penel C (2007b) Phylogenetic distribution of catalase-­peroxidases: are these patches of order in chaos? Gene 397:101–113 Pelaez F, Martinez MJ, Martinez AT (1995) Screening of 68 species of basidiomycetes for enzymes involved in lignin degradation. Mycol Res 99:37–42 Pérez-Boada M, Ruiz-Dueñas FJ, Pogni R, Basosi R, Choinowski T, Martínez MJ, Piontek K, Martínez AT (2005) Versatile peroxidase oxidation of high redox potential aromatic compounds: site-directed mutagenesis, spectroscopic and crystallographic investigations of three long-range electron transfer pathways. J Mol Biol 354:385–402 Phetsom J, Khammuang S, Suwannawong P, Sarnthima R (2009) Copper–alginate encapsulation of crude laccase from Lentinus polychrous Lev. and their effectiveness in synthetic dyes decolorizations. J Biol Sci 9:573–583 Piontek K, Smith AT, Blodig W (2001) Lignin peroxidase structure and function. Biochem Soc Trans 29(2):111–116 Pizzul L, Castillo MDP, Stenström J  (2009) Degradation of glyphosate and other pesticides by ligninolytic enzymes. Biodegradation 20(6):751–759 Poulos TL, Edwards SL, Wariishi H, Gol MH (1993) Crystallographic refinement of lignin peroxidase at 2 Å. J Biol Chem 268:4429–4440 Pozdnyakova NN, Turkovskaya OV, Yudina EN, Rodakiewicz-Nowak Y (2006) Yellow laccase from the fungus Pleurotus ostreatus D1: purification and characterization. Appl Biochem Microbiol 42(1):56–61 Priyanka G, Uma G (2017) Statistical optimization of laccase production by Aspergillus flavus PUF5 through submerged fermentation using agro-waste as cheap substrate. Acta Biol Szeged 61(1):25–33 Qiu L, Huang Z (2010) The treatment of chlorophenols with laccase immobilized on sol–gel derived silica. World J Microbiol Biotechnol 26:775–781 Rahman MA, Noh HB, Shim YB (2008) Direct electrochemistry of laccase immobilized on au nanoparticles encapsulated dendrimer bonded conducting polymer: application for a catechin sensor. Anal Chem 80:8020–8027 Rani V, Mohanram S, Tiwari R, Nain L, Arora A (2014) Beta glucosidase: key enzyme in determining efficiency of cellulase and biomass hydrolysis. J Bioproces Biotechniq 5:197 Ranocha P, McDougall G, Hawkins S, Sterjiades R, Borderies G, Stewart D, Cabanes-Macheteau M, Boudet AM, Goffner D (1999) Biochemical characterization, molecular cloning and expression of laccases-a divergent gene family in poplar. Eur J Biochem 259:485–495 Rastegari AA, Yadav AN, Gupta A (2019) Prospects of renewable bioprocessing in future energy systems. Springer, Cham Reyes P, Pickard MA, Vazquez-Duhalt R (1999) Hydroxybenzotriazole increases the range of textile dyes decolorized by immobilized laccase. Biotechnol Lett 21:875–880 Robles A, Lucas R, Martinez-Canamero M, Omar NB, Perez R, Galvez A (2002) Characterization of laccase activity produced by Hyphomycetes Chalara (syn. Thielaviopsis) paradoxa CH 32. Enzyme Microb Technol 31:516 Rochefort D, Kouisni L, Gendron K (2008) Physical immobilization of laccase on an electrode by means of poly(ethyleneimine) microcapsules. J Electroanal Chem 617:53–63 Rodríguez Couto S, Toca Herrera JL (2006) Industrial and biotechnological applications of laccases: a review. Biotechnol Adv 24:500–513

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Chapter 7

Fungal Enzymes for Bioremediation of Contaminated Soil Prem Chandra and Enespa

7.1  Introduction Fungi are varying in size and shape and are known as eukaryotes. Individual cells of fungi vary in its size such as yeasts. The chains of elongated cells stretch for miles (Alberts et al. 2013). The saprophytic fungi grow on dead organic matter, whereas others are parasites. These are absorptive heterotrophs that secrete digestive enzymes and break down the organic matters into substrates, and then absorb easily available molecules of organic matters (Sankaran et al. 2010). The fungal hyphae have large surface areas and small volumes which improve the absorptive capability of the fungi (Wright and Upadhyaya 1998). The bioremediation (mycoremediation) has been progressively renowned by the potential of fungi (Igiehon and Babalola 2017). Other explanation also describes why the fungi are perfect organisms for the mycoremediation (bioremediation), and why they may be superior to other microorganisms such as bacteria, algae, and plants under various situations (Harms et al. 2011). The mycelial network of fungal organizations has capability to penetrate into soils and contact spaces of soil pores (Boswell et al. 2007). The fungal mycelium can performance as a component, and mature surrounding the hurdles, recycled dead hyphae and reallocate properties and development to resource-rich zones of the soil. In comparison to various other eukaryotes, the cells of fungal hyphae display the indeterminate growth, so the division of cells takes place continuously in the hyphae of a mycelium, providing the mineral components accessible in the substrate (Miller and Fitzsimons 2011). The fungal mycelium in drier soils or soils holding the

P. Chandra (*) Department of Environmental Microbiology, School for Environmental Sciences, Babasaheb Bhimrao Ambedkar (A Central) University, Lucknow, Uttar Pradesh, India Enespa Department of Plant Pathology, School of Agriculture, MPDC, University of Lucknow, Lucknow, Uttar Pradesh, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_7

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nutrient patches locally may be benefitted. This is because their mode of growth permits them to associate with soil openings for their growth (Tsing 2012). The fungi have more potential to tolerate high concentrations of toxins compared to bacteria (Gonzalez-Chavez et al. 2004). Furthermore, several secondary metabolites are produced by the fungi for the bioremediation of polluted soils, and the enzymes secreted are particularly significant, and several of them lack substrate specificity and they are released as exoenzymes into the substrate (Cohen and Hadar 2001). Generally, the fungi depend on the degradation of macromolecules extracellularly, and a large quantity of enzymes is produced and secreted into the substrate (Girish and Kemparaju 2007). In the last years, emergent pollutants are of great interest (Bosco and Mollea 2019). Among them, endocrine-disrupting chemicals (EDCs) and pharmaceutical personal care products (PPCPs) generated from human being chemicals are significant due to their biological effects on nontarget organisms; particularly, the endogenous hormonal effects, antagonized or simulated to EDCs, are toxic at very low concentrations to the organisms (Duarte et al. 2018). Bisphenol A, Estrone, 17 β-estradiol, 17 α-ethinylestradiol, and triclosan are detected in soil and studied mostly. Due to the irrigation of contaminated wastewater, EDCs and PPCPs enter mainly into the soil ecosystem (Dodgen et al. 2014; Ying and Kookana 2005; Chen et al. 2010). The ligninolytic fungi have the capability to convert EDCs, permitting a decrease in their ecotoxicity or the endocrine-disrupting activity; furthermore, the capability of degradation by the heterogeneous class of PPCPs is also reported to occur broadly by unspecific enzymatic systems (Cajthaml 2015; Rodarte-Morales et  al. 2011; Yadav et al. 2018).

7.2  Fungal Diversity In any geophysical area, the variation in living organisms taken together abundantly is known as biodiversity (Sogin et al. 2006; Rana et al. 2019b). However, most of the fungi are not observable by the naked eye and must be considered under a microscope. The cultivable fungi can be recognized due to the sporulation (Tibbett and Carter 2008; Yadav et al. 2017). The fungi are superior to bacteria, viruses, and other smaller forms of life such as viroids (Parlevliet 2002). Approximately, 72,000 named species of fungi are recognized, and some new species are being added at the rate of about 700 to 1500 each year (Tortella et  al. 2005; Yadav et  al. 2019a, b). About 63,500 species of fungi have been labeled, in which 13,500 are found to be associated with algae as lichens. Sometimes, these organisms are termed the lower fungi such as Zygomycetes and Chytridiomycetes or higher fungi such as Ascomycetes and Basidiomycets (Blackwell 2011). They are classified as enzymatic machinery. The name white rot is given for rot fungi that decay the lignin and brown rot for those that decay only the cellulose. This difference is reflected in the overall appearance of the rotten wood (Rytioja et al. 2014). The brown rot fungi are known as soft rot fungi, which decompose only the cellulose, but they attack damper wood, and decay

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ordinarily occurs only near the wood surface (Enespa and Chandra 2017). Currently, the fungal potential for bioremediation (mycoremediation) has been gradually renowned. The bioremediation of hydrocarbons is based on biodegradation, and so it is involved in a partial or a complete breakdown of the relevant pollutants (Alvarez et al. 2017; Rastegari et al. 2019). Fundamentally, this mechanism is different from bioaccumulation which designates the capability of some fungi to accumulate or hyperaccumulate various metals inside the cell from the soil (Alvarez et al. 2017).

7.3  Fungal Enzymes Generally, based on catalytic reaction activity, these enzymes are divided into six classes: oxidoreductases (EC 1), transferases (EC 2), hydrolases (EC 3), lyases (EC 4), isomerases (EC 5), and ligases (EC 6) as expressed in (Table 7.1) (Alexander et al. 2017; Kour et al. 2019b; Rana et al. 2019a; Suman et al. 2016; Yadav 2019). The fungal extracellular enzymes mostly belong to two classes of enzymes. They are oxidoreductases and hydrolases. The transfers of hydrogen or oxygen atoms or Table 7.1  Classification of enzymes and types of reactions Classification of enzymes and their number Oxidoreductases (EC1)

Reaction profile The transfer of electrons from one molecule to another involved in oxidation reactions. We usually see the removal of hydrogen from the substrate in biological systems. Dehydrogenase is the typical enzyme in this class such as, alcohol dehydrogenase. R–CH2OH + A R–CHO + H2A (where, A is an acceptor molecule). If A is oxygen, the relevant enzymes are called oxidases or laccases; if A is hydrogen peroxide, than they are known as peroxidases Transferases (EC2) The enzymes catalyze the transfer of groups of atoms from one molecule to another molecule. The transfer of an amino group from an amino acid to an alpha-oxoacid is promoted by aminotransferases or transaminases Hydrolases (EC3) The cleavage of peptide bonds in proteins, glycosidic bonds in carbohydrates, and ester bonds in lipids by hydrolases. Generally, the hydrolases breakdown the larger molecules to smaller fragments Lyases (EC4) The formation of double bonds through the removal of groups by the catalyzation of lyases. The splitting of the glycosidic linkages by beta-elimination in pectate lyases Isomerases (EC5) Isomerases catalyze the transfer of groups from one position to another in the same molecule. Catalyze isomerization Ligases (EC6) Ligase enzymes participate in biosynthetic reactions where new groups of bond are formed by the joining of molecules together with covalent bonds. These reactions require the energy input in the form of cofactors such as ATP

References Pearce et al. (2003)

Clodoveo et al. (2014)

Goodman (2010)

Munoz-­ Munoz et al. (2017) Wu et al. (2011) Nelson et al. (2008)

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electrons from one substrate to another are oxidation and reduction reactions, known as oxidoreduction reactions and are catalyzed by oxidoreductases catalase (Lucas et al. 2008). The cleavages of chemical bonds by the addition of water are hydrolysis reactions and are catalyzed by hydrolases catalase. In the growing phase of fungi, various extracellular of enzymes are produced, and they degrade various polymers (Leonowicz, et al. 2001). In the lignin degradation, the oxidoreductases enzymes are involved, since lignin cannot be utilized as a source of energy or carbon by the fungi. The cellulose and hemicellulose enzymes have the capability for growth after degradation of complex lignocellulose structure in the plant cell wall (Kirk and Farrell 1987). The hydrolases catalyze lignin degradation and further break down to cellulose and hemicellulose (Langston et al. 2011).

7.3.1  Oxidoreductases (EC 1) Lignin is the most complex natural polymer formed from the random polymerization of phenyl propanoid units. Lignin-modifying enzymes (LMEs) are known as extracellular nonspecific oxidative enzymes, which have the capability of degradation to lignin by radical reactions (Joy et al. 2015). Four copper ions per molecule are found in laccases known as phenol oxidases. The oxygen is reduced to water due to catalysis by laccases and supplemented by the oxidation of various phenols (Galhaup and Haltrich 2001). A low redox potential has been seen in the laccases, but other organic compounds are also oxidized in the presence of redox mediators, such as 2, 2′-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS) and 1-hydroxybenzotriazole (HBT) (Camarero et al. 2007). The peroxidase enzymes are MnPs, LiPs, and VPs, which have heme protein (protoporphyrin IX containing Fe2+ion as a central atom) as their prosthetic group (Table 7.2). The Mn2+ ions are oxidized to Mn3+ ions which are catalyzed by MnPs, and stabilized by chelation with the organic acids. The one-electron oxidation of both phenolic and non-phenolic aromatic compounds is catalyzed by LiPs (Ye et al. 2010). VPs have both catalytic activities of MnP and LiP, and are known as hybrid peroxidases. The non-­ligninolytic heme-containing peroxidases are also produced by the Coprinopsis cinerea, which are dye-decolorizing peroxidases (DyP) and peroxidase (CiP). The smaller dye molecules and phenols are oxidized by the CiP, whereas the DyP also have the ability of oxidizing complex dye molecules (Liers et  al. 2013). Moreover, the lignin degradation is also shown by intracellular cytochrome P450 monooxygenases. The lipophilic compounds are also catalyzed by the monooxygenation of P450 enzymes, and they may degraded to lignin and other organic compounds together with peroxidases (Wittich 1998). The catalytic similarities have been shown in extracellular aromatic peroxygenases (APO) and intracellular P450s. In Agrocybe aegerita and Coprinellus radians, the APOs activities have been described (Pecyna et al. 2009). LMEs used in several applications are potential industrial enzymes used in processing industries of paper and pulp, for example, in the functionalization of lignocellulosic materials, biobleaching, biopulping, pitch removal, and alteration of wood

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Table 7.2  Extracellular oxidative enzymes of fungi and their environmental applications Enzyme activity Laccase

Manganese peroxidase (MnP) Dye-­ decolorizing peroxidase (DyP)

Coprinopsis cinerea peroxidase Versatile peroxidase (VP)

Reaction mechanism O2-dependent one-electron oxidation of various phenols. Extended substrate range in the presence of redox mediators. Mostly acidic and rarely in neutral pH range H2O2-dependent one-electron oxidation of Mn2+ to Mn3+, chelated Mn3+ oxidizes phenolic compounds, acidic pH range H2O2-dependent one-electron oxidation of organic compounds, decolorizes Reactive Blue 5, additional hydrolyzing activity, acidic pH range, also very low pH H2O2-dependent one-electron oxidation of various phenols, acidic and alkaline pH range Reaction mechanism of both MnP and LiP, acidic pH range

Occurrence in fungi Basidiomycota and Ascomycota, in most WRF and LDF

References Ray et al. (2014)

Basidiomycota, common in WRF and LDF

Zhang et al. (2014)

Basidiomycota and Ascomycota

Merckx (2013), Salvachúa et al. (2013)

Basidiomycota, only in C. cinerea

Floudas et al. (2012), Millati et al. (2011) Datta et al. 2017, Kinnunen et al. (2017), Pollegioni et al. (2015)

Basidiomycota, only in Pleurotus sp., Bjerkandera sp. and Trametes versicolor

fibers (Ferhan 2016), improving the contaminated soil from effluents and its bioremediation by enhancing the enzymatic hydrolysis of lignocellulosic substrates (Pérez et al. 2002). The laccases and MnPs are the strategic fungal enzymes for the bioremediation (Reddy 1995). All white rot fungi produce laccase and MnP; they are more widespread among LiP produced by WRF (Christian et  al. 2005). Particularly, the activity of LiP has been identified only in a few experiments from the soil. Moreover, Pleurotus sp., Bjerkandera sp., and Trametes versicolor produced only versatile peroxidase (VP) (Hofrichter 2002).

7.3.2  Hydrolases (EC 3) For the degradation of cellulose and hemicellulose from wood or plant litters, the fungi utilize various extracellular hydrolytic enzymes. The chains of single cellulose are connected together with the bonds of hydrogen to form microfibrils (Lynd et al. 2002). Three different types of cellulases are required for the degradation of cellulose namely endoglucanase (EC 3.2.1.4), exoglucanase (cellobiohydrolase, EC 3.2.1.91), and β-glucosidase (EC 3.2.1.21), and they act synergistically (Schwarz 2001). Endoglucanases hydrolyze β-1, 4-glucosidic linkages in noncrystalline sections of the cellulose chain internally, which results in a decrease in the chain length and an

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increase in the number of free ends. Cellulose is degraded by exoglucanases starting from the free ends and splitting off cellobiose, a disaccharide formed of two glucose units (Eriksson et  al. 1990). Lastly, the cellobiose hydrolyzes to glucose by the β-glucosidase. In recent times, cellulose-breaking accessory enzyme has been discovered (Várnai et al. 2010). This enzyme catalyzes the oxidative cleavage of crystalline cellulose and increases its accessibility to classical hydrolytic enzymes (cellulases) catalyzed by the lytic polysaccharide monooxygenase (also known as GH61 enzyme) (Quinlan, et al. 2011). Branched heteropolysaccharides composed of various hexoses (D-glucose, D-mannose, D-galactose), pentoses (D-xylose, L-arabinose, D-arabinose), deoxyhexoses (Lrhamnose, L-fucose), and uronic acids (4-O-methyl-D-glucuronic acid, D-galacturonic acid, D-glucuronic acid) are hemicelluloses. The hemicelluloses composition in softwood, hardwood, and annual plants varies with different plants (Oliveira et al. 2007). Several enzymes are required for the breakdown of hemicelluloses due to its various compositions (Pérez et  al. 2002). The xylan (in hardwood) and glucomannan (in softwood) are the two most common hemicelluloses. The chains of 1, 4-linked β-D-xylopyranose units are situated in xylan and have homopolymeric backbone (Zhou et al. 2017). Xylan contains on average one glucuronic acid besides xylose and the side group is attached to every tenth xylopyranose unit. β-D--mannopyranose and β-D-glucopyranose units are found in the backbone and β-D--galactopyranose units as side groups in glucomannan, which is a branched heteropolysaccharide (Alamgir 2018). In the soft wood (glucomannan), the ratio of galactose: glucose: mannose is 1:2:7 (Lechat et al. 2000).

7.4  Environmental Contamination 7.4.1  Common Source of Pollutants in Soil In soils, the persistent toxic compounds, chemicals, salts, radioactive materials, or disease-causing agents and heavy metals are present due to the industrial movements and pollution (Förstner and Wittmann 2012) and have adversarial effects on the growth of plants and animal health. Soil can become polluted by the various routes, such as seepage from a landfill, industrial waste discharge into the soil, contaminated water that percolated into the soil, rupturing of underground storage tanks, overuse of pesticides, herbicides, or fertilizers. (Quint 1998).

7.4.2  Contaminated Land and Its Issues The removal of pollutants from the soil and water by the fungal microorganisms is known as bioremediation. There are two methods, namely, in situ and ex situ, to remove pollutants from various sites. If the contaminated material is treated onsite, the method is known as in situ method, and if the treatment of contaminated material is done

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elsewhere, the method is known as ex situ method (Mueller et al. 1989). The bioremediation methods are known to degrade toxic organic materials, for example, from oil spills, pesticides, and industrial waste and heavy metals at the molecular level and convert them into less toxic compounds (Boopathy 2000). The bioremediation is the full mineralization of contaminants, that is, their transformation to CO2, H2O, N2, HCl, by the various fungal strains (Rhodes 2014). The heavy metals and radioactive cations, of course, cannot be decayed, but can be reduced into forms of low solubility, for example, by a change in the oxidation state, such as U (IV) (in UO2), so that it may be less harmful in the soil or might be physically unconcerned by mycoremediation, which involves the harvesting of plant or fungus (Gavrilescu et al. 2009; Glasser 2001).

7.4.3  Heavy Metal Contamination in Soil The elements with metallic properties and have atomic number > 20 are known as heavy metals. Certainly, the metal conversion is the normal mechanism in the soil. However, the high concentrations of heavy metals can be toxic for the living cells such as plants, animal, and microbes (Lasat 1999). The most common heavy metal pollutants in the environment are As, Sr, Cs, U, Cd, Cr, Cu, Hg, Pb, and Zn. Some of these metals are micronutrients and are necessary for the growth and development of plants, for example, Zn, Cu, Mn, Ni, and Co, while others have unknown biological function, such as Cd, Pb, and Hg (Alloway 2013). Heavy metals enter into the environment or soil by both natural and anthropogenic sources (Duruibe et al. 2007). The weathering minerals, erosion of rock, and volcanic activity are the most significant natural sources, and the mining, smelting, electroplating, use of pesticides and fertilizers, as well as biosolids in agriculture, sludge dumping, industrial discharge, atmospheric deposition are the anthropogenic sources (Ali et  al. 2013; Barsainya et al. 2016). The anthropogenic and manmade sources of several heavy metals in the environment or soil are presented in Table 7.3.

7.5  Bioremediation of Heavy Metals by Fungi Various species of filamentous fungi such as Trichoderma sp., Penicillium sp., Aspergillus sp., and Mucor sp., have the capability to tolerate and remediate heavy metals from the soil by the various activities (Deng et al. 2011). Due to the presence of negative charge on the various functional groups, the fungal cell walls have excellent metal-binding properties such as carboxylic, amine or sulfhydryl, and phosphate in several components of the cell wall (Wang and Chen 2009). The most prominent soil microorganism is arbuscular mycorrhizal fungus (AMF). It is established by the physical link directly between the soil and plant roots which increases the surface area of the root surface and smoothing nutrient absorption by the plants (Chapin 1980). In the alleviating of metal toxicity to the host plant, the AM fungi

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Table 7.3  Anthropogenic and natural sources of heavy metals in the soil Heavy metals Zinc (Zn)

Sources Mining, oil refinery, plumbing, Brass manufacturing

Effects on microorganisms Inhibits growth, death, decrease in biomass

Effects on human being Kidney and liver failure, lethargy, macular degeneration, metal fume fever, prostate cancer, seizures, vomiting Ataxia, depression, gastrointestinal irritation, hematuria, icterus, impotence Learning deficits, reduced Inhibit enzyme Lead (Pb) Battery fertility, renal system activitiesand manufacture, damage, risk factor for transcription, and herbicides and Alzheimer’s disease, insecticides, aerial denatures nucleic shortened attention span, emission from the acid anorexia, chronic combustion of nephropathy, damage to lead petrol neurons, high blood pressure, hyperactivity, insomnia Growth inhibition, Irritation of the skin, Chromium Fly ash, paints itching of respiratory tract, inhibition of (Cr) and pigments, liver diseases, lung cancer, tannery industry, oxygen uptake, nausea, renal failure, elongation of lag steel industries reproductive toxicity, phase vomiting bronchopneumonia, chronic bronchitis, diarrhea, emphysema, headache Dizziness, dysphasia, Disrupt cell Mercury Release from membrane, inhibits gastrointestinal irritation, (Hg) Au–Ag mining gingivitis, kidney problem, enzyme function, and coal decrease population loss of memory, pulmonary combustion, edema, reduced immunity, size, denature medical waste, sclerosis ataxia, attention protein Power plants deficit, blindness, deafness, emissions decrease rate of fertility, dementia Chest pain, dermatitis, Nickel Nonferrous metal, Inhibit enzyme (Ni) paints, porcelain activities, oxidative dizziness, dry cough and shortness of breath, stress, disrupt cell enameling, headache, kidney diseases, membrane electroplating, lung and nasal cancer, nausea, cardiovascular diseases

References Karwade et al. (2018)

Shawai et al. (2017)

Dhal et al. (2013) Chandra and Singh (2014)

Ali et al. (2013)

Bhatnagar and Minocha (2010)

(continued)

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Table 7.3 (continued) Heavy metals Arsenic (As)

Cadmium (Cd)

Cooper (Cu)

Sources Atmospheric deposition, mining, pesticides, rock sedimentation, smelting, Pesticides and wood preservatives Electroplating of cadmium containing plastics, phosphate fertilizer, paints and pigments, plastic stabilizers Pesticides and fertilizers

Effects on microorganisms Enzyme deactivations

Transcription, inhibits carbon and nitrogen mineralization, damage nucleic acid, denature protein, inhibit cell division Inhibits enzyme activities, Disrupt cellular function

Effects on human being Cardiovascular and respiratory disorder, conjunctivitis, dermatitis, skin cancer and brain damage

References Matschullat (2000)

Itai-Itai, kidney diseases, lung and prostate cancer, lymphocytosis, microcytic hypochromic anemia, testicular atrophy, vomiting, bone disease, coughing, emphysema, headache, hypertension Liver and kidney damage, metabolic disorders, nausea, vomiting Abdominal pain, anemia, diarrhea, headache

Fleischer et al. (1974)

Branham et al. (1995)

are also involved. The arbuscular mycorrhizae showed specific role in the host plant on exposure to heavy metal and depend on various environmental factors, such as the ecotypes and the plant species, the fungal species and the ecotype, the availability of metal, edaphic conditions of soil, such as the fertility of soil, and the growth conditions of plants like intensity of light or density of root (Khan 2005; Hall 2002). Likewise to PGPR, numerous mechanisms have been assumed for the direction of toxic metal and the allocation in plant roots in the presence of AMF, including the binding of heavy metals to the cell wall and their deposition in the vacuoles of AMF (Chandra and Enespa 2016), the sequestration of metals with the help of siderophores in the soil or into the root apoplasm, the binding of metals to metallothioneins or phytochelatins inside the fungal and plant cells, and the transporters of metals at the tonoplast of both plants and fungi catalyze the transport of metals from the cytoplasm (Neagoe et al. 2013; Jan and Parray 2016).

7.5.1  Heavy Metals and Enzyme Regulation For the utilization of complex nutrients, the saprotrophic basidiomycetes utilized a variety of extracellular enzymes such as ligninolytic enzyme (Kaur 2016). The controlling factors for the production enzyme among white rot fungi have been widely researched (Sanchez and Cardona 2008). The nutrients availability, inhibitory

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compounds, temperature, pH, and interrelationships with other fungi are the main factors which influence the production of enzymes (Mata et al. 2010). The ligninolytic and cellulolytic extracellular enzymes are controlled at the transcription level by the heavy metals and during the course of their action (Lee et  al. 2012). The energy flux in the ecosystem influences the effect of heavy metals on the enzyme activities. A positive regulation of laccase and isoenzymes on the copper application has been reported in P. ostreatus study (Baldrian 2003). With increasing Cd concentration, the Mn-peroxidase activity decreased, and the activities highly increased of endo-1, 4-l-glucanase, 1, 4-l-glucosidase and laccase in the presence of metal (Kapahi and Sachdeva 2017). The P. sajor-caju laccase isozyme genes (phenol oxidase A1b (POXA1b), POXA2, and POXC) that are regulated at the transcriptional level in response to copper and manganese have been reported. The energy flux in the ecosystem influences the effect of heavy metals on the enzymes activities (Goudopoulou et al. 2010). A positive regulation of laccase and isoenzymes on the copper application has been reported in P. ostreatus study (Palmieri et al. 2003). With increasing Cd concentration, the Mn-peroxidase activity decreased, and the activities highly increased of endo-1, 4-l-glucanase, 1, 4-l-glucosidase and laccase in the presence of metal (Kapahi and Sachdeva 2017). The P. sajor-caju laccase isozyme genes (phenol oxidase A1b (POXA1b), POXA2, and POXC) that are regulated at the transcriptional level in response to copper and manganese have been reported (Goudopoulou et al. 2010). It has also been reported that the activity of laccase decreases immediately and reduces the stability of the enzyme after the addition of Hg. Stimulatingly, Cu and Hg increased the activity of MnP slightly (Zeng et al. 2012). At low concentrations of Cd, Cu, and Hg, the activity of MnP decreased when it is incubated in the presence of all three metals, due to the synergetic effects of the heavy metals (Aragay et al. 2011; Dwivedi and Enespa 2014). And Mn has also been found to affect MnP gene transcription, and the activity of enzyme in a positive way in some Pleurotus spp. has been reported (Cohen et al. 2002). The progeny of Pleurotus eryngii was incubated in the Zn-, Cu-, Co-, Cd-, and Ni-enriched substrate and assessed for the effect on morphology and physiology. During the incubation stage in Ni and Cu, the concentration of laccase activity decreased, and completely inhibited during the fruiting stage (Baldrian 2003). The effects of inhibition were more noticeable when multi-metal solution exposure takes place. For the consideration of a fungal species as a biosorbent, the desorption of the adsorbed metal ions and their successive reuse and the productivity of the biomass in biosorption need to be taken into account (Volesky and Holan 1995). The desorption in the acidic solution has been described to be more effective than that in the alkaline desorption solution. Under acidic conditions, the protons participate for the sites releasing metal ions in the medium (Mohan and Pittman 2007). A 97% desorption of the adsorbed Hg from the immobilized and heat-treated P. sajor-­ caju resulted when eluted with HCl. P. ostreatus, using HCl for a contact period of 1 h only showed that 99% Pb could be desorbed (Akar et al. 2008). P. florida could be regenerated and reused for the biosorption of Pb for six times using biomass. The 59% generation rate of Cu has been reported for P. mutilus (Kapahi and Sachdeva

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2017). However, they can be upgraded by involving chemical desorption technique with the recovery of copper; the capacity of regenerated biomass for 10 g/l content has a maximum adsorption capacity that is smaller, but still significant 59.75 mg/g (Malyarenko et al. 2005).

7.5.2  Mechanisms of Mycoremediation The fungi are very resourceful and have a wide range of adaptability and quick responsiveness to stress condition, environmental adversities, and exciting climatic conditions. The reduction of complex hydrocarbons and the chains of hazardous substances into simpler, nontoxic, biodegradable form to tidying the environment can be achieved with the help of fungal strain (Dubos 1987). Various fungi also have excellent capability to binding with metallic ions, which comprise the efflux of metal ions outside the cell and buildup, and construct the metal ion complex inside the cell; later, they reduce the toxic metal ions to a nontoxic state (Hajipour et al. 2012). Various mechanisms have progressed by which they can immobilize, mobilize, or renovate the metals and make them inactive or tolerate the uptake of heavy metal ions (Fig. 7.1) (Prado et al. 2015). The mycoremediation mechanism adopted by fungi includes Exclusion: Due to the formation of a permeable barrier, the metal ions are kept away from the target sites. Extrusion: By the active transport, the metal is pushed out of cells.

Fig. 7.1  Bioremediation mechanism of heavy metals by fungi (modified and adopted from Chandra and Enespa 2019)

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Fixation: By the formation of a complex with metal-binding proteins or other cell components like enzymatic detoxification, intra- and extracellular sequestration, dissolution of metal by acid production, chelation, and precipitation through the production of organic bases, extracellular metal precipitation fixes the metals. Biotransformation: The toxic metals are reduced to less toxic forms like methylation, demethylation, volatilization, oxidation, and reduction. Generally, the immobilization, mobilization, biosorption, and biotransformation are considered main approaches used for mycoremediation of hazarders in the agro ecosystem in order to avail good air and water quality for future generations (Sardrood et al. 2013).

7.6  B  iodegradation and Bioremediation of Pesticides by Fungi 7.6.1  Organic Contaminants in the Soil The hazardous organic compounds are released by the industry and other sources to soil in the form of wastewater and the solid waste (Wuana and Okieimen 2011). This hazardous substance may be heavy metal, pesticides, and herbicides and they can accumulate in other organisms through the food chain and have the capability of long-range transport and are classified as Persistent Organic Pollutants (POPs) by the Stockholm Convention, 2001 (World Health Organization 2010). After its 5th meeting, there are 22 organic compounds that are currently classified as POPs; some are directly produced by the chemicals and some are generated as a form of by-products of reactions (Schwarzenbach and Gschwend 2016). The by-products formed in the industrial processes are polychlorinated biphenyls (PCB), polychlorinated dibenzo-p-dioxins (PCDD), polychlorinated dibenzofurans (PCDF), hexachlorobenzene (HCB), carcinogenic polyaromatic hydrocarbons (PAHs), and certain brominated flame-retardants (Roots 2004). The POPs include some pesticides also, such as dichlorodiphenyltrichloroethane (DDT), and some organometallic compounds, for example, tributyltin (TBT). In transformers and capacitors, the PCBs were widely used as coolants and insulating fluids (Darbre 2015). PCBs are common contaminants in many former industrial soils due to their widespread use. PCBs are problematic to be degraded by bacteria because due to the formation of toxic chlorobenzoic acids (CBAs), the PCBs are not easily degraded by bacteria which have a tendency to accumulate in soils polluted with PCBs (Mikszewski 2004). However, the ligninolytic fungi have capability to further reduce CBAs while instantaneously transforming PCBs (Peu 2014). So, the “emerging contaminants” increased in the environments (Petrie et al. 2015). Previously, these chemicals were not detected in the natural water, but discovered currently such as plasticizers (also known as Bisphenol A (BPA), flame-­retardants, pesticides, pharmaceuticals products such as antibiotics and other personal care products (Loraine and Pettigrove 2006). They naturally disrupt the endocrine system or cause other long-term effects to human being or other living organisms in very low

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concentrations (Colborn et al. 1993). For instance, Bisphenol A has estrogenic activity even at pico- to nano-molar concentrations (Wetherill et al. 2007). BPA is known not for the estrogenic activity since 1993, but it is still used to create polycarbonate plastics and epoxy resins, and reactions can be found in many other consumer products such as water bottles and coatings inside food and beverage cans (Vandenberg et al. 2007). During washing with hot water or in alkaline or acidic conditions, BPA leaches out from these materials (Howdeshell et al. 2003). Furthermore, BPA is not completely degraded in the treatment of wastewater, and thus BPA can spread into the aquatic environment through the wastewater effluent. BPA and other emergent contaminants could be degraded in municipal wastewaters with the help of LMEs (Tijani et al. 2013). In the degradation of BPA with the help of MnP, it was observed that the treatment after 90 min. with MnP resulted in 100% reduction of estrogenic activity and 98% degradation of BPA (Kabiersch et al. 2011). In the arctic environment where the natural reduction is extremely slow, the long-­ range transport is the most important source for pollutants. The sources of long-­range transport are previously used chemicals and already-existing environmental pollution due to their long life spans after the restriction and the production of many POPs (Wania 2003). So, it is necessary to treat the soils affected with POPs with special treatment techniques to remediate the contaminants. The fungal enzymes can provide new approaches to treat polluted soils or wastewaters (Gomes et al. 2013). There is a potential to use fungal mycelia to treat POP soils, including PCB, PAH-, and PCDD/ F-contaminated soils, and fungal enzymes to clean up effluents, to degrade emerging contaminants in municipal wastewaters, synthetic dyes in textile industry wastewaters, and lignin in pulp mill wastewaters (Anasonye et al. 2014; Anasonye et al. 2015).

7.6.2  Bioremediation of Pesticides by Fungal Enzymes Based on their morphological, physiological, and genetic features, the fungi are considered unique organisms. They are able to colonize in natural environments (soil, air, water) and are known as omnipresent, and they maintain the ecosystem’s equilibrium (Barea et al. 2005). Fungi develop their unique bioremediation properties due to adaptation to their environments. Based on laboratory studies, it is known that all the natural organic compounds can be degraded by various fungal species due to the production of enzymes such as amylases, lipases, and proteases and that allow them to use substrates as fats, proteins, and starches (Shah et al. 2008). The pectin, cellulose, and hemicelluloses can be used by some other species as a carbon source. The natural complex polymers are degraded by some main fungal strains and may be resistant to microbial attack, such as keratin, chitin, and lignin (Walker and White 2011). Generally, on the basis of reactions, the enzymes are divided into six classes such as catalyze: oxidoreductases (EC 1), transferases (EC 2), hydrolases (EC 3), lyases (EC 4), isomerases (EC 5), and ligases (EC 6) (Andreini et al. 2008). Mostly, the extracellular fungal enzymes are categorized into two enzyme classes: oxidoreductases and hydrolases. The oxidation or reduction reactions, that

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is, transfer of hydrogen or oxygen atoms or electrons from one substrate to another are carried out by oxidoreductases (Madhavi and Lele 2009). The cleavage of chemical bonds by the addition of water is hydrolysis reaction that is done by the hydrolases. In their growth environment, extracellular enzymes are produced by the fungi to degrade various polymers (Lucas et al. 2008). In lignin degradation, oxidoreductases enzymes are involved. The lignin cannot be used as a source of energy or carbon by the fungi. The complex lignocellulose structure of the plant cell wall has capability to use cellulose and hemicellulose for the growth. The breakdown of cellulose and hemicellulose is catalyzed with hydrolases (Pérez et al. 2002).

7.6.3  Biodegradation of PAHs The PAHs are also formed due to the incomplete combustion of carbon-containing fuels, including fossil fuels, wood, or other biomass, and are also present in crude oil and coal (Huber et al. 2006). The PAH contamination can be either long-winded contamination in urban surface soils, which receive a continuous input of pyrogenic PAHs from the emissions to air, or the contamination from a point source like gasworks soil or oil-polluted soil. In sawmill soil, PAH contaminations originate from coal-tar creosote, which completely consists of PAHs, and is commonly used to preserve wood and in the making of waterproof crossties and power line poles (Winquist et  al. 2014). The United States Environmental Protection Agency (US EPA) (Article IV) has listed 16 PAHs of special concerns. The α-benzopyrene is considered as the most carcinogenic PAH-compound associated with five benzene rings (Ramesh et al. 2004). The PAHs are resistant to microbial degradation due to their complex structure with two or more fused benzene rings and low water solubility (Haritash, and Kaushik 2009). The diffuse contamination of PAH in surface soils has been observed and PAHs like phenanthrene, fluoranthene, and pyrene can be degraded by soil microorganism, while the benzo (a) anthracene and benzo (a) pyrene are examples of more recalcitrant high molecular weight (HMW) PAHs that are biodegraded very slowly in the soil (Tian et al. 2008). The ligninolytic fungi have a capability to degrade and mineralize the PAHs and the mechanism is thought to be similar to that of lignin degradation. The oxidation of PAHs is catalyzed by peroxidases or laccases and results in the formation of PAH-quinones that can be further oxidized (Haritash, and Kaushik 2009). LMEs might have an important role in the initial attack on HMWPAHs in soil. Since the LMEs are extracellular, they have capability to diffuse effectively to the highly immobile HMW PAHs. The resulting metabolites are more water-­ soluble, and thus, more bio-accessible (Turja et al. 2013). The formed compounds can be substrates for many bacteria, but they may also be further degraded by fungal intracellular enzymes, such as cytochrome P-450 monooxygenase (Saratale et  al. 2011). When the degradation of several PAHs by Irpex lacteus was determined, the structures of some of the metabolites suggested the involvement of both LMEs and cytochrome P-450 monooxygenase. In addition to mineralization, a

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significant fraction of PAHs are incorporated into humic substances during the bioremediation (Covino et al. 2016).

7.6.4  Biodegradation of PCDD/Fs Persistent Organic Pollutants (POPs) belong to the most toxic compounds such as polychlorinated dibenzo furans (PCDF) and dibenzo-p-dioxins (PCDD). Two aromatic rings with one to eight chlorine atoms are found in PCDD/Fs (Seo et al. 2009). Chlorine groups have toxic congeners in all of the 2,3,7,8 positions (7 PCDDs and 10 PCDFs), and 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin (2, 3, 7, 8-TCDD) is the most toxic congener (Safe 1993). By multiplying their concentration with a Toxic Equivalency Factor (TEF) (Article III), the toxicity of other congeners is compared to that of 2, 3, 7, 8-TCDD. By adding up all the 2, 3, 7, 8-TCDD equivalents of all the individual congeners, the total toxic concentration (expressed as World Health Organization – Toxic Equivalent, WHO-TEQ) can be calculated (Giesy and Kannan 1998). The compounds PCDD/Fs are chemically stable in structure and poor bioavailability, so the degradation rate of these compounds is very slow in nature. PCDD/Fs are tightly adsorbed on soil particles, and absorbed into organic matter in soils and sediment due to its highly hydrophobic nature (Anyasi and Atagana 2011). In industrial processes, the PCDD/Fs are almost exclusively produced as by-­ products, for example, the municipal waste incineration, chlorine bleaching of paper and pulp, manufacturing of pesticides, herbicides, and fungicides (Mezcua et al. 2012). In Finland, the main source of PCDD/F pollution was the production and use of a chlorophenol-containing wood preservative (Ky-5) during 1940–1984 (Salo et  al. 2008). The composition of Ky-5  in chlorophenols is 2, 3, 4, 6-­tetrachlorophenol (55%), 2, 4, 6-trichlorophenol (36%), and pentachlorophenol (7%) (Anasonye et al. 2014). Furthermore, in Ky-5, the PCDD/Fs were found as impurities. Over the past several decades, most of the chlorophenols in these contaminated sawmill soils have volatilized, leached, and biodegraded (Josefsson et al. 2016). In the top soil with hepta and octachloro dibenzo furans (1, 2, 3, 4, 6, 7, 8-HpCDF and OCDF) as the main congeners, still, the PCDD/Fs continue to be the main source for the contaminations (Seike et al. 2007). Additionally, severe PCDD/F contamination is also found in Kymijoki River sediments due to sawmill soils. The Plant situated of Ky-5 along the river and a fire in the plant in 1960 caused a large spill in the environment (Karademir et al. 2013). It is observed that several white rot fungi (WRF) to degrade all congeners of PCDD/Fs have been reported, even the ones with maximum amount of chlorine atoms (Anasonye et  al. 2015). Additionally, to extracellular LMEs, some WRF have also intracellular P450, and the LMEs attack co-metabolic lyon-chlorinated dioxins under aerobic conditions. P450 enzymes of fungi, together with peroxidases, are also involved in the degradation of lignin. Probably, P450 enzymes and extracellular enzymes are involved with later stages of degradation of chlorinated dioxins by WRF (Anasonye et al. 2014). Several fungi used for the biodegradation of pesticides are given in Table 7.4.

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Table 7.4  Bioremediation of pesticides using fungus Name of chemical compounds PAH Diphenyl ether Anthracene Naphthalene Bleached kraft pulp mill effluent Fungicide: Metalaxyl and Folpet Petroleum products crude oil Gasoline Polychlorinated biphenyles, POPs, Polychlorinated dibenzofurans Phenylurea herbicide diuron Dye decolorization of textiles

Endosulfan Pesticide Chlorinated hydrocarbons; Heptaclor Chloropyriphos Leather tanning effluent Heptachlor O Toxaphene O

Name of the fungi White rot fungi: Pleurotus ostreatus Trametes versicolor Armillaria sp. Pleurotus eryngii Rhizopus oryzae or Pleurotus sajor-caju

Purohit et al. (2018)

Gongronella sp. and R. stolonifer

Pandey et al. (2018)

A. niger, Rhizopus sp., Candida sp., Penicillium sp., Mucor sp. Exophiala xenobiotica Doratomycesnanus sp., D. purpureofuscus, D. verrucisporus, Myceliophthora thermophila, Phoma eupyrena, and Thermoascus crustaceus, Phanerochaete sordida, Mortierella isabellina

Babič et al. (2017)

Aspergillus niger, A. foetidus, T. viride, A. sojae, Geotrichum candidum, Penicillium sp., Pycnoporus cinnabarinus, Trichoderma sp. Penicillium sp.

References Mohapatra et al. (2018) Kumar (2017) Saiu et al. (2018)

Mouhamadou et al. (2013), Takada et al. (1996), Papanikolaou et al. (2004)

Jebapriya and Gnanadoss (2013)

P. ostreatus Aspergillus terreus

Romero-Aguilar et al. (2014) Palmieri et al. (2000) Abraham et al. (2013)

Aspergillus flavus, Aspergillus sp. and A. niger, Aspergillus jegita Trichoderma viridae,

Bhadbhade et al. (2002) Roberti et al. (2006)

7.6.5  Biodegradation of Naphthalene The enzymes are categorized into six general groups according to the Commission on Enzymes of the International Union of Biochemistry. The enzymes catalyzing oxidation– reduction reactions are known as oxidoreductases, and those catalyzing a chemical group from one molecule to another are known as transferases; the hydrolytic enzymes are known as hydrolases; those catalyzing the addition of functional groups to double bonds or vice versa are known as lyases; those catalyzing the intramolecular rearrangements are known as isomerases; and those enzymes catalyzing the condensation of two molecules coupled with the cleavage of a pyrophosphate bond of ATP or similar triphosphate are known as ligases (Karigar and Rao 2011; Faber and Faber 1992). Hydroxylation of aromatic rings is catalyzed by oxygenases and is initiated by various enzymes. One oxygen atom is inserted into their substrates from the monooxygenase types, in another process,

7  Fungal Enzymes for Bioremediation of Contaminated Soil H

COOH

OH OH

OXYGENASES

OH

H

ENZYME

NAPHTHALENE

205

CIS-NAPHTHALENE DIHYDRODIOL

SALICYLIC ACID

OH ACETYL-CO ENZYME A CH3 – CO – SCoA

O

OH COOH COOH

ACETIC ACID CH3 –COOH

CO2 + H2O

3 - OXO - ADIPIC ACID

CATECHOL

NAPHTHALENE DEGRADATION BY FUNGAL ENZYMES

Fig. 7.2  The biodegradation process of naphthalene

two oxygen atoms are inserted into their substrates by dioxygenase-type approach (Ullrich and Hofrichter 2007). Consequently, in a series of reactions, they are converted into 2-ketoadipate or another compound and the ring is cleaved. And these compounds can be consumed by the fungi or other microorganisms (Fan and Krishnamurthy 1995). The biodegradation process of naphthalene is displayed in Fig. 7.2.

7.7  Soil Bioremediation and Its Benefits For the bioremediation, the use of microbes is tormented with several rate-limiting factors. For the detoxifications of hazardous substance, various limitations apply to using microbes. To the production of microbial culture, an expensive and long-­approaching process may be used (Hamelinck et al. 2005). Moreover, severe surroundings such as chemical shock, pH and temperature, hazardous toxins, predators, and high dose of the pollutants or their products may permanently damage or metabolically deactivate microbial cells (Burridge et al. 2010), and the difficulty in continuing active cells during the conveyance to the contaminated sites also limits the use of whole-cell purification machineries (Raafat et al. 2008). The other factors that could control the use of microbes include restricted mobility of the cells within the soil, the alternate carbon sources, and the weakness of the inoculated microorganisms in competition with the native population (Nannipieri et al. 2003). The biotransformation is involved in a series of catalyzed enzymes reactions. If the enzyme reactions (preferably immobilized) are

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used instead of the microbes, most of these hostile factors can either be eliminated or mitigated (Kumar et al. 2015; Kour et al. 2019a). The bio-oxidation catalyst causes a host of reactions by the enzymes. Specificity is shown by the enzymes and is characterized as they show an ideal temperature and pH for their actions (Denard et al. 2013). The other catalysts, similarly, accelerate the reaction rate of chemical by lowering the activation energy for a specific reaction.

7.8  Conclusion and Future Prospects Currently, in the world, contamination and its remediation have received considerable attention due to the fact that numerous heavy metals cannot be degraded in the whole environment and these metals lie in the soil. Various approaches have been followed successfully to generate various plants and sites that have the capability to propagate in polluted soils of metals and accumulate or tolerate the stress of metals. Naturally, the microbial approaches for the remediation of heavy metal tolerance are an eco-friendly and economical approach. Subsequently, the heavy metal uptake by plants and their tolerance depend on various environmental factors, and interactions between plant and fungi can play an important role in successful survival and growth of plants in contaminated soils. The plant growth-promoting microbes also assist plant growth by changing the bioavailability of heavy metal due to various enzymatic activities of fungi. These beneficial effects demonstrated by fungus, together with the interrelationship between heavy metal tolerance and plant growth-­promoting capability, indicate that their exploitation in remediating metal-­contaminated soils might have significant potential in the near future. The synergistic approach of plant and fungi and their metal mobilization mechanism, transformation, and detoxification have been observed. Further, observing and handling the fungal and heavy metal remediation involve categorization of the fate and behavior of the compounds of interest in the environment. So, these highlights represent the importance of a consistent link among the research and development for the valuation and management of unindustrialized metal contaminants and the tools, apparatus, and knowhow that contribute toward the contentment of these experiments. Acknowledgments  The authors are grateful to the Department of Environmental Microbiology, Babasaheb Bhimrao Ambedkar University, Lucknow, and Department of Plant Pathology, School for Agriculture, MPDC, University of Lucknow, Lucknow, for providing valuable support to write this chapter. There are no conflicts of interest.

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Seo JS, Keum YS, Li QX (2009) Bacterial degradation of aromatic compounds. Int J Environ Res Public Health 6(1):278–309 Shah AA, Hasan F, Hameed A, Ahmed S (2008) Biological degradation of plastics: a comprehensive review. Biotechnol Adv 26(3):246–265 Shawai SAA, Nahannu MS, Mukhtar HI, Idris A, Abdullahi II (2017) Assessment of heavy metals concentration in ground water from various locations of Gezawa Local Government Area of Kano State. Adv Mater 6(5):73–76 Sogin ML, Morrison HG, Huber JA, Welch DM, Huse SM, Neal PR, Herndl GJ (2006) Microbial diversity in the deep sea and the underexplored “rare biosphere”. Proc Natl Acad Sci 103(32):12115–12120 Suman A, Yadav AN, Verma P (2016) Endophytic microbes in crops: diversity and beneficial impact for sustainable agriculture. In: Singh D, Abhilash P, Prabha R (eds) Microbial inoculants in sustainable agricultural productivity, research perspectives. Springer-Verlag, pp 117– 143. https://doi.org/10.1007/978-81-322-2647-5_7 Takada S, Nakamura M, Matsueda T, Kondo R, Sakai K (1996) Degradation of polychlorinated dibenzo-p-dioxins and polychlorinated dibenzofurans by the white rot fungus Phanerochaete sordida YK-624. Appl Environ Microbiol 62(12):4323–4328 Tian Y, Liu HJ, Zheng TL, Kwon KK, Kim SJ, Yan CL (2008) PAHs contamination and bacterial communities in mangrove surface sediments of the Jiulong River Estuary, China. Mar Pollut Bull 57(6–12):707–715 Tibbett M, Carter DO (2008) Soil analysis in forensic taphonomy: chemical and biological effects of buried human remains. CRC Press, Boca Raton/London/New York, pp 29–52 Tijani JO, Fatoba OO, Petrik LF (2013) A review of pharmaceuticals and endocrine-disrupting compounds: sources, effects, removal, and detections. Water Air Soil Pollut 224(11):1770–1778 Tortella GR, Diez MC, Durán N (2005) Fungal diversity and use in decomposition of environmental pollutants. Crit Rev Microbiol 31(4):197–212 Tsing A (2012) Unruly edges: mushrooms as companion species: for Donna Haraway. Environ Humanit 1(1):141–154 Turja R, Soirinsuo A, Budzinski H, Devier M-H, Lehtonen KK (2013) Biomarker responses and accumulation of hazardous substances in mussels (Mytilus trossulus) transplanted along a pollution gradient close to an oil terminal in the Gulf of Finland (Baltic Sea). Comp Biochem Physiol Part C Toxicol Pharmacol 157:80–92 Ullrich R, Hofrichter M (2007) Enzymatic hydroxylation of aromatic compounds. Cell Mol Life Sci 64(3):271–293 Vandenberg LN, Hauser R, Marcus M, Olea N, Welshons WV (2007) Human exposure to bisphenol A (BPA). Reprod Toxicol 24(2):139–177 Várnai A, Siikaaho M, Viikari L (2010) Restriction of the enzymatic hydrolysis of steam-pretreated spruce by lignin and hemicellulose. Enzym Microb Technol 46(3–4):185–193 Volesky B, Holan ZR (1995) Biosorption of heavy metals. Biotechnol Prog 11(3):235–250 Walker GM, White NA (2011) Introduction to fungal physiology. In: Fungi, Hoboken, pp 1–35 Wang J, Chen C (2009) Biosorbents for heavy metals removal and their future. Biotechnol Adv 27(2):195–226 Wania F (2003) Assessing the potential of persistent organic chemicals for long-range transport and accumulation in Polar Regions. Environ Sci Technol 37(7):344–1351 Wetherill YB, Akingbemi BT, Kanno J, McLachlan JA, Nadal A, Sonnenschein C, Watson CS, Zoeller RT, Belcher SM (2007) In vitro molecular mechanisms of bisphenol A action. Reprod Toxicol 24(2):178–198 Winquist E, Björklöf K, Schultz E, Räsänen M, Salonen K, Anasonye F, Cajthaml T, Steffen T, Jørgensen KS, Tuomela M (2014) Bioremediation of PAH-contaminated soil with fungi–from laboratory to field scale. Int Biodeterior Biodegrad 86:238–247 Wittich RM (1998) Degradation of dioxin-like compounds by microorganisms. Appl Microbiol Biotechnol 49(5):489–499

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Chapter 8

Bioremediation of Polycyclic Aromatic Hydrocarbons (PAHs) Contaminated Soil Through Fungal Communities Ulises Conejo-Saucedo, Darío R. Olicón-Hernández, Tatiana Robledo-Mahón, Haley P. Stein, Concepción Calvo, and Elisabet Aranda

8.1  Introduction Polycyclic aromatic compounds (PAC), including the well-known subgroup of polycyclic aromatic hydrocarbons (PAHs) and the heterocyclic aromatic compounds, include several thousand individual compounds (Achten and Andersson 2015). In particular, polycyclic aromatic hydrocarbons (PAHs) are benzene rings fused pollutants widely distributed in nature. They are considered small-PAHs when they have from two to four rings (such as naphthalene, anthracene, phenanthrene, and pyrene) and large-PAHs with more than four rings (benzopyrene, chrysene, benzoanthracene, etc.) (Table  8.1). PAHs are listed by EPA (Environmental Protection Agency) as priority pollutants and considered to be mutagenic, teratogenic, and carcinogenic compounds (Luch 2009). The physicochemical properties of PAHs depend on the number of rings, as well as the presence of substituent (alkylated). Thus, vapor pressures and water solubility decrease when the number of rings increases; however, the n-octanol-water partition coefficient log KOW increases (Achten and Andersson 2015). PAHs are distributed in all environments worldwide. PAHs present in carbon-­ containing organic materials, such as coal tar and charcoal, are of petrogenic origin, while those released during combustion of fossil fuels or natural combustions (forest fires, volcanic eruption) are of pyrogenic origin. In addition, PAHs can also be synthesized during the degradation of organic matter or biologically by bacteria and plant life, although this fact has not yet been clarified (Parlanti 1990). The largest contributor of PAHs in the environment is from incomplete combustion (both natural and anthropogenic) varying geographically around the Earth U. Conejo-Saucedo · D. R. Olicón-Hernández · T. Robledo-Mahón · H. P. Stein · C. Calvo · E. Aranda (*) Department of Microbiology, Institute of Water Research, University of Granada, Granada, Spain e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_8

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Table 8.1  Polycyclic aromatic hydrocarbons including some heterocyclic and their details health risk, and environmental data Compound Small-PAHs Naphthalene

Chemical structure

Ranking Molecular positionb formula/(Mw) 81

C10H8/128.174

Anthracene

9 (PAHs)

C14H10/178,23

Phenanthrene

252

C14H10/178,23

Acenaphthene

171

C12H10/154.2

Fluorene

9 (PAHs) 38

C13H10/166.223

Pyrene

260

C16H10/202,25

Fluoranthene

138

C16H10/202.256

Benzo(a) anthracene

Health riska

Environmental dataa

Toxic; Very toxic to noncarcinogenic aquatic organisms, it may cause long-term adverse effects in the aquatic environment; hazardous to the environment Very toxic to Toxic; specific toxicity to some aquatic organisms, it organs; noncarcinogenic; may cause long-term non-mutagenic adverse effects in the aquatic environment; hazardous to the environment

C18H12/228,2879 Toxic; non-genotoxic

Very toxic to aquatic organisms, it may cause long-term adverse effects in the aquatic environment; bioaccumulation may occur in crustacea, fish, milk, molluscs, and algae

(continued)

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Table 8.1 (continued) Compound Large-PAHs Benzo[a] Pyrene

Chemical structure

Ranking Molecular positionb formula/(Mw) 230

C20H12/252,31

Benzo(b) fluoranthene

10

C20H12/256.34

Benzo[g,h,i] perylene

9 (PAHs)

C22H12/276.338

Indenol (1,2,3-cd) pyrene

176

C22H12/276.338

Health riska Toxic; carcinogenic; may cause reproductive difficulties; teratogenic

Environmental dataa Very toxic to aquatic organisms, it may cause long-term adverse effects in the aquatic environment; hazardous to the environment

Data obtained from the US Agency for Toxic Substances and Disease Registry (http://www.atsdr. cdc.gov/substances/index.asp). 2017 CERCLA Priority List of Hazardous Substances b Ranking position in the CERCLA Priority List of Hazardous Substances a

(Zhang and Tao 2009). Once in the atmosphere, small PAHs and gaseous PAHs can be volatilized and transported over long distances, influenced by certain meteorological conditions. Small PAHs can then be degraded by different reactions such as photodecompositions and/or chemical oxidation, defining the PAH halflife in the atmosphere, until they are absorbed into particles, where further degradation could be completely inhibited. At this point, the small PAHs are deposited onto terrestrial, lacustrine, and marine surfaces (Fig.  8.1). Heavy PAHs can be transported in settling atmospheric particles which contribute to the depositions in soils and sediments (Wilcke 2000). Thus, PAHs can be dispersed locally and intercontinentally via atmospheric and aquatic transport (Becker et al. 2006). In soils, PAHs result mostly from atmospheric deposition and tend to be absorbent to soils particles. This absorption is strongly influenced by the soil composition, particle size, and organic carbon content. Retention of PAHs in organic matter and soil particles contributes to the aging of soils with PAHs, and as a result, the ability for PAH-degrading microorganisms decreases and they are accumulated (Loehr and Webster 1996).

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Fig. 8.1  Main sources of PAH in the environment: (1) forest fires, (2) volcanic eruptions, (3) combustion of fossil fuels, and (4) accidental spills during storage, transport, and disposal of fossil fuels

8.2  Fungi as Key Drivers in PAHs-Polluted Soils Microorganisms are found in all ecosystems and are essential biological components to their functioning. Microorganisms are involved in the nutrient cycling which contributes to biogeochemical processes in the biosphere. Microbial metabolism which drives degradation and transformation processes depends on the interactions and the relationships between microbes and substrate availability. The combination of microorganisms which comprises the microbial community includes bacteria, archaea, fungi, and microeukaryotes (Fuhrman 2009; Yadav et al. 2019a, b). The first person to observe and discuss the microbial community was Clements (1916), who considered the community as a supraorganism with levels of

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organization, in which the stratified interactions involved allowed different properties to the system. Gleason (1926) defined the terms as an individualist concept: a microbial community exists as a space where an assortment of species cohabit in a balance, due to the level of tolerance to physical and chemical substances that they produce themselves. In this sense, more recently, Konopka (2009) defined microbial community as “multi-species assemblages, in which organisms live together in a contiguous environment and interact with each other.” Thus, the key factor is that the communities involve a group of microorganisms which share a habitat and engage in different interactions and roles in relation to the biological and the psychochemical conditions. Particularly, these interactions contribute to the challenge in defining the limits of an ecosystem. It is likewise a challenge to define specific actions of the microbial interface, due to feedback systems between the various microorganisms and their effect on the environment. Microbial ecology aims to study these interactions of each community in a specific environment. However, to outline the specific bioreactions in the ecosystem is not an easy task. More often, these interplays are studied at the scale of the micron, an exception being the hyphae of fungi, which can be considered on a larger scale. The dynamics, structure, and succession of microbial communities are useful in understanding the complexity of ecosystems, this being the main goal of observational ecology (Garland 1997). Additionally, just as diversity is related to the stability of an ecosystem, a higher ecosystem diversity involves a wide variety of microorganisms with varying functions and requirements. This broad metabolic profile will favor the community more adapted under unfavorable chemical and biological conditions. As such, the cohabitation of communities with different ecological functions act as a buffer when faced with a change in the environmental conditions, thereby regulating the connections between different microbes and maintaining the balance of the ecosystem (Konopka et al. 2015). In this context, polluted environments make for fascinating ecosystems of study, due to the higher potential of the inhabitants to develop biodegradation and biotransformation processes. In observation of the metabolic interactions between microbial cohabitants and the abiotic environmental factors, researchers may gain significant insights into the ecological functions of each diverse microbial community. Fungal communities are ubiquitous, colonizing terrestrial and aquatic habitats, and even extreme ecosystems such as polar zones, deserts or hypersaline environments (Margesin and Schinner 2001; Rana et al. 2019; Sharma et al. 2019; Yadav et al. 2017a, 2018). Fungi are well-known to play a major role in nutrient cycling and biogeochemical cycles, in both aquatic and terrestrial environments (Kubicek and Druzhinina 2007; Suman et al. 2016). In addition, fungi are frequently found in hostile environments, polluted ecosystems, or environments which have otherwise been altered by anthropogenic activity, characterized by the presence of toxic or recalcitrant compounds, such as heavy metals or PAHs. Fungi have several mechanisms of adaptation to these conditions, including the thick cell wall, the production of spores, the production of intracellular and extracellular unspecific enzymes, and/ or a developed enzymatic system to counteract the effect of reactive oxygen species (ROS) (Harms et al. 2011). In addition, in polluted environments, the fungal com-

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munity is involved in complex interactions with other microorganisms. In fact, the close interaction between fungi, plants, and bacteria, called the rhizospheric effect, has been well documented in scientific literature (Kotoky et al. 2018). The rhizospheric effect is mediated by mutual interactions of plant and microbes, mainly during phytoremediation processes, as well as having a role in highways of fungi mycelial tissue for the movement of bacteria through the soil (Kohlmeier et  al. 2005). Despite the representation of fungi in these environments, fungal communities, and their specific roles in these ecosystems remain poorly studied.

8.2.1  Culture-Dependent Techniques Diverse techniques have been used to analyze fungal communities in PAH-­ contaminated soil over the last decades. Traditionally, culture-techniques based on the direct isolation of fungal strains, and conventional microbial techniques (plate counts, community level physiological profiling or Biolog), have been used as the necessary optimal media to growth and conserve the strains. However, it is known that cultured fungi represent only a fraction of the entire fungal community, and favors fast growing species that produce large quantities of spores (O’Brien et al. 2005). Despite these disadvantages, there is no doubt about the great interest in using culture-techniques as a source of fungal cells with ability to remove pollutants for bioremediation purpose and for general studies focused on deeper knowledge of these specific microorganisms.

8.2.2  Culture-Independent Techniques In the last decades, a number of approaches have been developed based on genetic diversity. These include omics technologies, which are useful in understanding the cellular and molecular behavior of the communities, especially during a bioremediation event. Thus, if metagenomics supply the genetic profile of the fungi involved in a polluted soil, metatranscriptomics reveal information related to gene expression profiles in active populations of the mycobiome involved, for example in signaling, or in the degradation and metabolism of PAHs. Metaproteomics then show the protein expression profile as a response to a specific compound and function, integrating intermediates and final products in pathways through metabolomics (Kachienga et al. 2018; Kotoky et al. 2018). In addition, sequencing technologies have allowed for the elucidation and annotation of several genomes at increasing rates. The “1000 Fungal Genome project” represents a strong push to obtain a number of new genomic sequences to include in a large-scale catalogue of fungal species (https://genome.jgi. doe.gov/programs/fungi/1000fungalgenomes.jsf) (Grigoriev et al. 2014). The application of these technologies has become an indispensable tool in bacterial soil ecology, but its use in studies of fungal communities has so far been limited.

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Fig. 8.2  Principal barcode-primer pair combinations used in fungi

Fungal databases display limitations related to the number of sequenced and well-­annotated fungal genomes and have largely failed to apply the sequence data to phylogenetics, as a way of potentially reorganizing fungi according to the new taxonomic classifications (O’Leary et al. 2016). Nevertheless, the collaboration of the different databases marks an advance in the creation of a complete and update database (Huzefa et al. 2017). On the other hand, the absence of a universally accepted DNA barcode is a restriction in ecological and biological studies, which has generated the necessity to use different biomarkers. Since the 1990s, taxonomic studies of fungi have been based on ribosomal genes of 18S or small subunit (SSU), the 5.8S subunit, and the 28S or large subunit (LSU) genes, being the LSU and SSU more efficient at high taxonomic levels (Cuadros-Orellana et  al. 2013). Currently, the intergenic region (ITS) has been considered the universal DNA barcode markers for fungi as suitable barcode. Different fungal studies have shown the efficiency of the ITS region as the barcode marker with the highest probability of correct identification, for a broad analysis of sampled fungi (Dentinger et al. 2011; Kelly et al. 2011; Schoch et al. 2012). Likewise, most diversity studies on environmental samples were using the ITS region to identify fungi (Bellemain et al. 2010; Wang et al. 2011; Rosling et al. 2011; Ihrmark et al. 2012) (Fig. 8.2). However, it is recommendable to use secondary region if there is a low variability of interspecific region, in order to obtain a high genetic precision. It must be considered that the use of a biomarker may lead to biased amplification of different taxonomic groups (Schoch et al. 2012; Blaalid et al. 2013).

8.3  Fungal Mechanisms in PAH Transformation The isolation of fungi-dwelling polluted environments provides important information about metabolic capability, for further application in PAH-polluted soils. Fungi are strict heterotrophic organisms, which involve the transformation of external carbon sources to obtain energy for basal metabolism as well as biomass production. Keeping in mind this degradative capacity and ecological niche, fungi have been supplied with an array of intrinsic bioprocesses to colonize natural matrices, survive

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in extreme conditions, degrade a variety of substrates, and adapt within adverse habitat (Anastasi et al. 2013; Selbmann et al. 2013; Zafra et al. 2015). As a biotechnological tool, the morphological components, type of growth, and enzymatic systems, among others, are important elements for fungi to be considered as excellent models to implement new technologies to, for example, develop and improve bioremediation processes. In the context of the elimination of PAHs, it was demonstrated that fungi can degrade complex aromatic carbon derivatives such as anthracene (Hadibarata et al. 2013; Marco-Urrea et al. 2015), pyrene (Bhattacharya et al. 2014; Mineki et al. 2015), and phenanthrene (Lee et al. 2014; Fu et al. 2018) as well as simple ring compounds and linear hydrocarbons such as benzene and methyl tert-butyl ether (Thomas 2013). In order to adapt the xenobiotic carbon source into their aerobic metabolism pathway and/or eliminate it by nonenzymatic methods, fungi can interact with contaminants by two main mechanisms: intra- and extracellular processes. In terms of the capability to secrete extracellular enzymes or not, as well as possessing elements for biosorption and/or bioaccumulation, fungi differ in the action mechanisms. However, it seems clear that both systems are functional and works together for the elimination of contaminants (Mougin et al. 2013; Deshmukh et al. 2016).

8.3.1  Extracellular Mechanism Extracellular mechanism for elimination of PAHs in fungi involves the use of low-­ specific enzymes as main component (Morelli et al. 2013). White-rot fungi (WRF) are the most important representatives of this kind of fungal elimination systems due to their ability of producing a set of extracellular lignin-modifying enzymes such as laccase, manganese peroxidase (MnP), and lignin peroxidase (Winquist et al. 2014; Yadav and Yadav 2015) and unspecific peroxygenases, versatile peroxidases, and decolorizing-type peroxidases (DyP-type) (Kües 2015). These kinds of enzymes can transform and/or mineralize the phenolic polymer components in wood that have complex and heterogeneous matrices, making them ideal tools for the degradation of a variety of aromatic compounds (Anastasi et al. 2013). From this point of view, WRF have shown promise for application in the bioremediation of PAH contaminated soil and water by employing these enzymes and focusing on improving the removal rates, while continuing to evaluate kinetic parameters, reusability and efficiency (Xuanzhen et al. 2014; Lee et al. 2014; Bautista et al. 2015; Godoy et  al. 2016). However, studies in fungal diversity by culture independent techniques show that in polluted environments, some of the main fungal populations are fungi lacking of these enzymes (D'Annibale et al. 2006). Another element of extracellular systems in the elimination of PAHs is the use of cell wall and membrane components as biosorbent systems (Harms et  al. 2011). This physicobiochemical surface phenomenon considers the electrostatic interaction of biomass with contaminants involved to gauge the removal of the pollutant and could be

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complemented by bioremediation (Ding et al. 2013; Tony et al. 2014). The use and production of fungal biosurfactants to improve extracellular uptake of xenobiotic compounds by biomass is one branch of bioaugmentation, and may play a role as an element that can improve the overall degradation of the xenobiotics in question (Lladó et al. 2013a). In this context, an emulsifying effect was observed in the biodegradation of ring-PAHs by Pleurotus ostreatus D1, due to the production of natural biosurfactants. The author suggested that the presence of this element increases the solubility and bioavailability of the pollutants and/or increase the production of extracellular enzymes for the elimination of the PAHs (Nikiforova et al. 2009).

8.3.2  Intracellular Enzymatic Pathways Intracellular enzymatic pathways for the biotransformation of PAHs show that xenobiotic entry into the cell occurs by several mechanisms and results in the formation of hydroxy, dihydroxy, dihydrodiol, and quinone derivatives. This process (phase I) is mediated by the cytochrome P450 (CYP) system and epoxide hydrolases (EHs). The next step, the conjugation process occurs when sulfate, methyl, glucose, xylose or glucuronic acid groups are linked to oxidized metabolites (Phase II); this process is mediated by transferase enzymes. During phase III, these metabolites are secreted or stored in organelles. The final step for the elimination of oxy-, methoxy-, or sulfate PAHs has not been fully discerned (Aranda 2016) (Fig. 8.3). CYP enzymes are elements present in the degradation of aromatic and aliphatic compounds in WRF and Ascomycetes such as Aspergillus sp. and Penicillium sp. (Aranda et al. 2017; Camacho-Morales et al. 2018; Huarte-Bonnet et al. 2018).

Fig. 8.3  Proposed phenanthrene degradation pathway in fungi (Marco-Urrea et al. 2015)

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In fact, CYP fungal enzymes involved in hydrocarbon assimilation are dependent on phylogeny (Huarte-Bonnet et al. 2018). These enzymes participate in the initial hydroxylation of aromatic and alkane derivatives in order to oxidize the xenobiotic and produce energy and are the main component in intracellular PAHs degradation (Huarte-Bonnet et  al. 2018). In this context, it was described that CYP52 and CYP53 clans play a significant role in fungi detoxification, however, being a complex superfamily of enzymes the participation of these proteins is not fully clear (Godoy et  al. 2016; Olicón-Hernández et  al. 2017; Huarte-Bonnet et  al. 2018). Bioaccumulation of PAHs in cytoplasm or vacuoles, also complemented the intracellular systems of elimination (Verdin et al. 2005; Gu et al. 2016).

8.4  Case Study of Fungal Communities on PAH Degradation In the last two decades, the interest in the use of biotechnologies, destined to recover contaminated environments from hazardous aromatic pollutants such as PAHs, has increased. The analyses of metagenomics in hydrocarbon-polluted areas have provided information about the richness and diversity of the autochthonous community in this type of environment, the genes involved in the degradation, the protein expression, and the final product generated. Several investigations have focused their attention on the study of the fungal diversity in PAH contaminated soils (Cerniglia and Sutherland 2010; Mineki et al. 2015; Sawulski et al. 2015; Siles and Margesin 2018; Yadav et al. 2017b). Fungi, as a part of a community focused on PAH degradation, have an important role as the main degraders or/and complement of communities of bacteria and complex eukaryotes for elimination of hydrocarbons aromatic derivatives (Zafra et al. 2014; Vasudevan et al. 2018). Borowik, et al. (2017) analyzed the effect of diesel oil contamination on the diversity of fungal communities in soils. After 270 days, the degradation of the PAHs with four rings was 64%; in PAHs with five rings was 28%; and in PAHs with six rings was 16%. However, the colony development index for fungi increased significantly, while the diversity index decreased, indicating a directional selection. In recent studies, Biache et al. (2017) analyzed the impact of PAH contamination on the abundance of microbial communities, and the communities’ degradation capacity for the PAHs. The results showed differences in the degradation rates according to the distribution, the availability of PAHs, and the three soils that were evaluated; reaching 98%, 76%, and 34% of degradation, respectively (Biache et al. 2017). In nature, the biodegradation of PAHs occurs among different microbial communities and is not restricted to single microbial species. Thus, cometabolism of PAHs will result in multitudes of high-polarity metabolites more easily degradable by lower species in the taxonomic class. Consequently, in a mixed microbial population, initial cometabolic transformation may pave the way for subsequent attack by another organism, resulting in a synergism of fungi-bacteria/bacteria-fungi for the elimination of PAHs (Vasudevan et  al. 2018). This step is essential for the

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c­ omplete mineralization of xenobiotics, although in a community is not clear if bacteria or fungi works as final biodegradator. The consortiums isolated from PAHs-­ contaminated environments demonstrated high tolerance of concentration of PAHs; however, in the case of fungi, it was observed the same tolerance, even without the fungi being previously exposed to environments contaminated with aromatic compounds (Simarro et al. 2013). This is an advantage for the use of fungi in bioremediation proposes. Bioremediation using fungal-bacterial cocultures generally favored bacteria in increased log phase and lag periods, utilization of PAHs as carbon source, and tolerance to heavy metals, among others. Thus, in the absence of fungal communities, initial oxidation of PAHs are seldom possible by bacteria, and on the other hand, in the absence of bacterial communities, there is a huge accumulation of dead-end fungal metabolites which may be toxic (Vasudevan et al. 2018). Regardless, Zhou et al. (2017), found a significant correlation of PAH contents with fungal community structure, but not with fungal diversity; interestingly, the community structure of bacteria might be more sensitive to soil PAH contamination than those of fungi and archaea (Zhou et al. 2017). Still, fungi have several characteristics that favor their use in consortium (Bacterial-Fungal) for bioremediation processes. Mainly, fungi can spread through soils, allowing bacteria to extended through the soil and access to contaminants (Nazir et al. 2010; Banitz et al. 2011); mycelium can act as dispersion networks for bacterial transport and actively mobilizing PAHs, called “fungal high-ways” (Kohlmeier et al. 2005; Schamfuß et al. 2013). In addition, in recent studies, it has been shown that bacteria adapt to these environments due to the release of nutrients and signaling molecules by fungi (Nazir et al. 2010). On the other hand, it has been found that these mycelia can increase the mobility of a wide range of PAH due to their translocation during cytoplasmic transmission (“fungal pipelines”) (Furuno et al. 2012); in this way, the access of bacteria and plants to soil pollutants and their degradation is favored (Banitz et al. 2013).

8.5  B  ioremediation Performed by Consortia in Contaminated Soils Two of the most used fungal bioremediation technologies for PAH removal in situ are bioaugmentation and biostimulation. While the process of bioaugmentation implies the introduction of a group of microbes that have the ability to survive in a contaminated environment and carry out the bioremediation process, during biostimulation, the autochthonous microorganisms receive nutrients for an adequate growth and metabolic rate in order to increase the efficiency of degradation (Vidali 2001). Contaminated soils are an excellent source of adapted microorganisms (Björklöf et al. 2009; Bourceret et al. 2016). For this reason, several studies have been carried

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out on native fungal communities in contaminated soils and their subsequent isolation and selection based on their ability to grow, efficiency to degrade different PAHs or use them as a source of carbon and energy (Li et al. 2008; Alrumman et al. 2015; Marco-Urrea et  al. 2015; Marchand et  al. 2017) for bioaugmentation purposes. In this context, some recent investigations showed the isolation of nine nonligninolytic fungal strains native from soil contaminated with Maya crude oil. The results showed strains of the genera Fusarium, Neurospora, Aspergillus, Scedosporium, Penicillium, Neosartorya, and Talaromyces. However, Aspergillus terreus, Talaromyces spectabilis, and Fusarium sp. present the best tolerance to 2000 mg kg−1 of a mixture of phenanthrene and pyrene soil in a solid-state microcosm system for 2 weeks. The elimination of PAHs was close to 21% (Reyes-César et  al. 2014). Similar examples were found in native fungi represented mainly by Ascomycota phylum, Mucoromycotina subphylum and Basiodiomycota phylum isolated from a historically pyrogenic PAH-polluted soil in Spain. Out of the 23 isolated species, 12 species were able to oxidize anthracene at different rates (Godoy et al. 2016). In a study done by Anastasi et al. (2009), the fungal consortium (T. versicolor, Bjerkandera adusta, Bjerkandera fumosa, and Lopharia spadicea) were performed in a contaminated soil composting system. These researchers concluded that these consortia eliminated approximately 56 out of every 100  mg  kg−1 dry weight of pyrene in 28 days (Anastasi et al. 2009); and reduced a high concentration of naphthalene (500 mg kg−1) in three weeks. Nevertheless, several studies have shown that a microbial consortium (Fungal, Fungal-Bacterial) are more effective (Jacques et al. 2008; Lladó et al. 2013b; Balaji et al. 2014; Maddela et al. 2015). In the studies carried out by Zafra et al. (2014), the consortium was composed of 12 fungal and bacterial PAH-tolerant isolates; the results shows a removal of 48.18% pyrene, 56.55% benzo[α]pyrene and 87.76% phenanthrene after 14 days. In addition, a study from Sharma et al. (2016), reported a new microbial consortium composed of Serratia marcescens L-11, Streptomyces rochei PAH-13 and Phanerochaete chrysosporium VV-18. Under controlled conditions, the consortium degraded nearly 70% of PAHs in 7 days; also, the degradation rate of PAHs significantly increased between 56% and 98% under natural conditions (in situ) in 7 days, and almost complete degradation was observed on the 30th day. Recently, Zafra et  al. (2017) have constructed a new, tolerant PAH-degrading microbial consortia, composed of six fungal and seven native bacterial strains with the ability to degrade up to 92% of phenanthrene, 64% of pyrene, and 65% of benzo(α)pyrene out of 1000 mg kg−1 after 2 weeks of incubation and used these compounds as a sole carbon source. The evidence shows that the microbial community (bacterial-fungal) plays a key role in the degradation of PAHs in soils. However, in conjunction with plant life (Plant-Fungal-Bacterial), microbes offer a novel type of consortium that makes the degradation more efficient (Fan et al. 2008; Tejeda et al. 2012; Aranda et al. 2013). For example, Storey et al. (2014) found that in the presence of tomato plants, the degradation of fluoranthene was greater, degrading close to 80.2% and 68.1% in concentrations of 500 and 5000 mg kg−1 fluoranthene over 30 days, respectively.

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Interestingly, fluoranthene in combination with the absence or presence of plants had a significant impact on bacterial and fungal community structures. Biostimulation technologies have been studied using compost/organic substrates in contaminated soils, that helps to improve the degradation, the texture of the soil, favors the transfer of oxygen and provides energy to microbial populations (Tyagi et al. 2011; Li et al. 2012; Lladó et al. 2013a; Bastida et al. 2016). Recent studies examine the addition of both fungal inoculum and composted vegetable waste to a PAH-contaminated sawmill soil in treatments at the laboratory and field scales. Several fungi, such as Agrocybe dura, Agrocybe praecox, Gymnopilus luteofolius, Irpex lacteus, Mycena galericulata, Phanerochaete velutina, Physisporinus rivulosus, Stropharia aeruginosa, Stropharia rugosoannulata, and Trametes ochracea, were found to be potential players in the degradation of PAHs; 96% of the four-ring PAHs and 39% of the five- and six-ring PAHs were degraded in 3 months (Winquist et al. 2014). Andreolli et al. (2015) compared bioagmentation and biostimulation in a burned woodland soil contaminated with high molecular weight hydrocarbons. The bioremediation protocol included a mycelial suspension of a Trichoderma sp. strain and a microbial growth promoter formulation. The results showed that the biostimulation approach was the best treatment; about 70% of the initial concentration of high molecular weight hydrocarbons was degraded after 60 days (Andreolli et al. 2015), indicating the important role of native microorganisms in the bioremediation process. However, in this study, the shift in the microbial community has not been assessed. Zafra et al. (2016) show that bioaugmentation with a fungal-bacterial consortium and biostimulation of native microbiota using easy-to-assimilate carbon sources and texturizer produced appreciable changes in the microbial diversity of polluted soils. The combination resulted in a shift in the native microbial communities to favor degrading specific contaminants such as PAHs. Other studies of biostimulation with NPK fertilizer, characterized soil microbial communities (bacterial, archaeal and fungal communities); the results showed high removal rates of total petroleum hydrocarbon obtained with agricultural inorganic NPK fertilizer in treatments at 10 °C and 20 °C during 15 weeks and resulted in changes in bacterial and fungal community structures (Siles and Margesin 2018). The abundance and diversity of bacterial and fungal communities did not play a decisive role on the effectiveness of soil bioremediation; these results were more influenced by changes in temperature and fertilization, indicating the effect of a correct biostimulation of the native microorganisms. Zafra et al. (2016) have reported a new consortium composed of four fungal species (Aspergillus nomius H7, Aspergillus flavus H6, Trichoderma asperellum H15, Rhizomucor vari-abilis H9) and five bacterial species (Bacillus cereus B4, Klebsiella pneumoniae B1, Klebsiella sp. B10, Pseudomonas aeruginosa B6, Stenotrophomonas maltophilia B14). Results showed that phenanthrene (84.29%), pyrene (59.66%), and benzo[α]pyrene (58.98%) degradation was significantly higher in soils inoculated with this consortium than control. These studies concluded that the consortia used produced appreciable changes in the microbial diversity, changing native microbial communities in favor of the degradation of specific populations.

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8.6  Conclusion and Future Prospects With the even growing emphasis on green technologies, the novel field of bioremediation has introduced opportunities for discovery in biological sciences. The inclusion of and focus on fungi in studies of microbial bioremediation can serve to add to understanding not just xenobiotic metabolism, but also aid in the understanding of microbial interactions in both laboratory and natural settings. Importantly, scientists may better be able to observe how fungal communities function and impact their environment on a larger scale. Isolating specific fungi can have its place in discovering the single players in a systemic whole, but ultimately, the consensus shows that fungi, bacteria, and plants work best all together in contaminant purification. Nonetheless, individually and within these systems of microbial communities, through applying surveys using omics techniques, scientists may be able to uncover levels of enzymatic processes behind PAH degradation. Additionally, studying fungi in polluted settings can carry insights that explicitly speak to the ability of species to adapt to stress in their habitats, as fungi have shown to be more resilient and adaptive in polluted settings. Researchers using omics techniques could ultimately pinpoint specific molecular adaptations that operate to augment chances of survival in PAH contaminated environments. Perhaps genetic engineers can even come to apply these mutations to organisms in degradation consortiums, to design whole, synergistic microbial communities for the complete removal of anthropogenic PAHs in contaminated habitats. In this way, fungi serve an important role in the biological and chemical rehabilitation of terrestrial and aquatic environments for a restoration of native ecological equilibrium in contaminated sites. Acknowledgments  The authors gratefully thank the Ministry of Economy and Competitiveness (MINECO) and European Regional Development Fund (ERDF) funds for the Ramón y Cajal contract of EA (RYC-2013-12481). UC and DRO thank CONACyT Mexico for the postdoctoral fellowship (230592/209148/473970; 231581/454815, respectively). HPS acknowledges the Fulbright Program (PS00247479) for the Open Study/Research Grant. We would like to acknowledge the Environmental Microbiology Research Group [RNM-270] of the University of Granada (Spain).

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Chapter 9

Role of Fungal Enzymes for Bioremediation of Hazardous Chemicals Nitika Singh, Abhishek Kumar, and Bechan Sharma

9.1  Introduction Hazardous chemicals may be defined in terms of any waste material that would be a present or future threat to organism or to the environment. It can also be defined as unwanted materials that exhibit hazardous characteristics (Wu et  al. 2008; Gnanasalomi et  al. 2013). Agricultural practices, industrial processes, and the use of variety of chemicals in different extents of our routine life result in the deliberate or accidental release of potent chemicals (i.e., lethal to humans) into the environment. These hazardous chemicals can be transported via atmosphere and water as well and, in many cases, find their way into sediments and soils after their release in environment. Environmental hazardous chemicals of particular interest are mainly pollutants including petroleum hydrocarbons, halogenated solvents from industrial sources, endocrine-disrupting agents and drugs, explosives, chemicals from agricultural activities, heavy metals, metalloids, and radionuclides (Harms et al. 2011; Kumar and Sharma 2018; Kumar et al. 2018). A number of activities including anthropogenic activities (e.g., agricultural practices) and industrial processes (e.g., mining and metal processing, petrochemical and industrial complexes, industry effluents, chemical weapons production, pulp and paper industries, dye industries, and industrial manufacturing) are responsible for the pollution in the environment. These chemicals may affect the health of humans, plant, and animals and also affect the environment for several causes (Rao et al. 2010). Among all environmental hazardous chemicals, some substances are potentially toxic and present in the environment and affect microbial organisms, plants, animals, humans, soil, sediments, water, and air. These toxic substances are very often present not only as mixtures of different organic compounds but also of organic and inorganic ones. Because the number of industries increases day by day, a large amount of waste N. Singh · A. Kumar · B. Sharma (*) Department of Biochemistry, Faculty of Science, University of Allahabad, Allahabad, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_9

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materials has been let out into the environment. These wastes are very toxic and persist in environment for a long time. They can enter into the food chain from soil and water even after their use. These chemicals have also the capability to bioaccumulate within the cells of organism. Thus, biomagnifications occur over the period because of the increasing levels of toxic compounds within the body. These hazardous pollutants are flammable, explosive in nature, corrosive, toxic, and bioaccumulative. Sometimes, eutrophication takes place due to the presence of inorganic pollutants, and acute and chronic diseases occur due to exposure to these pollutants (Gnanasalomi et al. 2013). Bioremediation is a process mediated by microorganisms that transform or degrade contaminants into nonhazardous or less hazardous substances. The efficient capability of bioremediation of pollutants in many organisms like bacteria, fungi, algae, and plants has been reported (Vidali 2001; Leung 2004; Karigar and Rao 2011). The bioremediation process primarily depends on action of microorganisms and which convert hazardous chemical to innocuous products by enzymatically attacking on the hazardous chemical. An effective bioremediation can only exist where environmental conditions favor microbial growth and activity. Often, the applications of bioremediation involve the manipulation in the environmental conditions which allow microbial growth and rapid degradation of chemical. Generally, degradation of pollutants is a very slow process. Only some certain fungal species have been reported as potent pollutant degraders. Some of these strains are working as some effective bioremediation agents only under laboratory conditions. The environmental factors like pH, temperature, oxygen, soil structure, moisture and appropriate level of nutrients, poor bioavailability of contaminants, and presence of other toxic compounds limit the fungal growth. However, the microorganisms can grow in environmental conditions, but most of them prefer optimal ideal condition that is difficult to maintain outside the laboratory (Dua et al. 2002; Dana and Bauder 2011). Most bioremediation systems operate under aerobic conditions, but anaerobic environments may also permit microbial degradation of recalcitrant molecules. Fungi rely on the participation of different intracellular and extracellular enzymes, respectively, for the remediation of recalcitrant and lignin and organopollutants (Hammel 1997; Karigar and Rao 2011) (Table 9.1).

9.2  O  ccurrence and Morphological Features of Fungi Used in Bioremediation Fungus belongs to a large group of eukaryotic organisms like microorganisms including yeasts, mushrooms, and molds. Fungi are heterotrophic, eukaryotic, and absorptive organisms that typically develop mycelium (a branched, tubular body), and reproduction takes place by sporulation. Less than 100,000 of the 1.5 million estimated fungal species have been described in Stajich et al. (2009). Up to 75% of fungal species account for the soil microbial biomass, i.e., total soil microbial

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Table 9.1  Fungal enzymes and their role in bioremediation Enzyme Laccase

Fungal taxa Ascomycota and Basidiomycota

Mechanism O2-dependent one-electron oxidation of organic compounds

Lignin peroxidases

Basidiomycota

H2O2-dependent one-electron oxidation of aromatic compounds

Manganese peroxidase

Basidiomycota

H2O2-dependent one-electron oxidation of Mn2+ to Mn3+, which subsequently oxidizes organic compounds

Tyrosinases

Ascomycota, Basidiomycota, and Mucoromycotina

O2-dependent hydroxylation of monophenols to o-diphenols (cresolase activity) Oxidation of o diphenols to catechols (catecholase activity) H2O2-dependent one-electron oxidation of organic compounds Additional hydrolyzing activity Incorporation of a single atom from O2 into a substrate molecule, with concomitant reduction of the another atom to H2O

Dye-decolorizing Basidiomycota peroxidases

Ascomycota, Cytochrome Basidiomycota, P450 monooxygenases Mucoromycotina, and Chytridiomycota

Applications Food industry, paper and pulp industry, textile industry, nanotechnology, synthetic chemistry, bioremediation, cosmetics, and so forth. Food industry, paper and pulp industry, textile industry, pharmaceutical industry, bioremediation, and so forth. Food industry, paper and pulp industry, textile industry, pharmaceutical industry, bioremediation, and so forth Cosmetic applications, production of dyes, biosensors, biosynthesis and medical applications, and food applications

References Baldrian (2006), Majeau et al. (2010)

Hofrichter (2002), Ruiz-Dueñas et al. (2009), Hammel (1995) Hofrichter (2002), Ruiz-Dueñas et al. (2009), Hammel (1995), Hofrichter (2010) Halaouli et al. (2006), Ullrich and Hofrichter (2007)

Dye decolorization, Oxidation of anthraquinone dyes with high redox potentials

Ullrich and Hofrichter (2007)

Pharmaceutical industry and bioindustrial applications

Kasai et al. (2010), Yadav et al. (2006)

(continued)

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Table 9.1 (continued) Enzyme Nitroreductases

Fungal taxa Ascomycota, Basidiomycota, and Mucoromycotina

Mechanism NAD(P) H-dependent reduction of nitroaromatics to hydroxylamino and amino(nitro) compounds and of nitro functional groups of N-containing heterocycles

Applications Reduction of TNT to hydroxylamino dinitrotoluene and amino-­ dinitrotoluenes Formation of mononitroso derivatives and ring cleavage products from cyclic nitramine explosives,

References Rieble et al. (1994), Esteve-­ Nunez et al. (2001), Bhushan et al. (2002), Crocker et al. (2006), Scheibner et al. (1997)

biomass, and dry weight is 50–1000 μg per g and 2–45 t per ha, respectively. The hyphae length for arable, pasture and forest top soils can be up to 102  m per g, 103 m per g, and 104 m per g, respectively (Ritz and Young 2004). Some classes of fungi tolerate optimal environmental conditions such as temperatures (−5 to +60 °C) and pH (1–9) and grow at a water activity of only 0.65, or with 0.2% oxygen. Morphologically, fungal species vary in shape and size widely from unicellular (microscopic organisms) to multicellular forms (easily seen with the naked eye). Individual cells range from 1 μ to 30 μ. Molds and yeasts are example of microscopic fungi. The kingdom fungi is estimated to contain about 80,000– 100,000 described species. Most of them are pollutant degraders and belong to the phyla Ascomycota and Basidiomycota, followed by the subphylum mucoromycotina (Harms et al. 2011).

9.2.1  Ascomycota Fungi Among all the fungal groups, meiosporic ascomycetes and mitosporic ascomycetes account for about 64% of all described fungal species and also the largest fungal group in terrestrial and aquatic environments (Shearer et al. 2007). Numerous members of the subphylum Pezizomycotina (the most diverse group of the filamentous ascomycetes) attack environmental organic hazardous. For example, species of the genera Cladophialophora and Exophiala belong to order Chaetothyriales and assimilate the toluene (Prenafeta-Boldú et  al. 2006). Aspergillus and Penicillium spp. belong to order Eurotiales and degrade pollutants such as aliphatic hydrocarbons, chlorophenols, polycyclic aromatic hydrocarbons (PAHs), pesticides, synthetic dyes, and 2,4,6-trinitrotoluene (TNT) (Hofrichter et  al. 1994; Pinedo-Rilla et al. 2009; Prince 2010). This diverse group includes microorganisms that can exist in both reproductive states, i.e., anamorph and teleomorph states, making their classification extremely

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difficult (Hibbett and Taylor 2013). This phylum contains parasitic, symbiotic, or saprotrophic lifestyles of microorganisms and morphologies such as unicellular in yeast and multicellular in filamentous, and dimorphic fungi can exist as mold/ hyphal/filamentous form or as yeast. Studies on the fungal diversity in some anthropogenically polluted samples, including activated sludge or wastewater-treatment plants indicate ascomycota as the dominant phylum (Weber et al. 2009; Evans and Seviour 2012; Maza-Márquez et al. 2016). These groups of fungi have been studied for the involvement of the intracellular enzymes such as Cytochromes P450 (CYP), manganese peroxidase, or laccase that have degradative capability of aromatic substances such as PAHs, chlorinated hydrocarbons, and diverse xenobiotic compounds (Marco-Urrea et al. 2015; Aranda 2016; Bovio et al. 2017). However, ascomycetes are able to degrade a large number of compounds.

9.2.2  Basidiomycota Fungi The phylum Basidiomycota constitutes roughly 34% of all described fungal species. Basidiomycetes are mostly inhabiting in terrestrial environments and are rarely in aquatic environments (Lynch and Thorn 2006). The group of macro- and microorganisms having characteristic features such as formation of basidia, a bottle-shaped cell structure containing haploid and sexual basidiospores, is the basidiomycota division of fungi. The most fungi of basidiomycota group pass between a dikaryotic state and diploid growth to asexual reproduction by conidia with the subsequent formation of basidiospores during their life cycle (Alexopoulos et al. 1996). Due to ecosystem balance and their environmental relations, these fungi have established and enhanced different systems such as cycling of carbon and nitrogen sources. The main mechanisms involved in the degradation of hazardous chemicals including PAHs and aromatic compounds by using basidiomycota fungi are groups of extraand intracellular oxidoreductases like laccases, peroxidases, and CYPs. These enzymes break down and modify the bonds of different compounds, mainly by extracellular pathways (Schmidt-Dannert 2016). Several kinds of LME and fungal mediators (Phanerochaete chrysosporium, Phlebia ochraceofulva, Pycnoporus sanguineus, Pleurotus ostreatus, and T. versicolor) have been studied in relation to the biodegradation of PhACs in the white-rot fungi (Yadav et al. 2006; Cajthaml et al. 2009; Díaz-Cruz et al. 2015) (Fig. 9.1).

9.2.3  Mucoromycotina Fungi Mucoromycotina incertae sedis, formerly known as Zygomycota, represents a heterogeneous group. The characteristic feature of these fungi is the formation of zygospores and aplanospores in the sexual phase and asexual phase, respectively (Benny et  al. 2014). Among all the lifestyles present in this group including mutualists,

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Fig. 9.1  The biochemical pathway of polycyclic aromatic hydrocarbon biodegradation by fungi (Harms et al. 2011)

saprophytes, and pathogens, members of Mucorales group (core group of the traditional Zygomycota) are facultative parasites/saprobes in nature. Cunninghamella elegans, member of Mucorales, have the ability to reproduce regioselective and stereoselective transformations in mammal enzymatic systems due to this nature; it has been extensively used as model fungi in different studies on the metabolism of xenobiotics. The subphylum Mucoromycotina part of the basal fungal lineage accounts for less than 1% of all described. In this subphylum, the genera Cunninghamella, Mucor, and Rhizopus (members of the order Mucorales) include degraders of PAHs, pesticides, textile dyes, and TNT (Scheibner et al. 1997; Pinedo-­ Rilla et al. 2009; Prince 2010). The high ability of this fungus group to transform a broad range of compounds has been studied in detail. The mechanism involved in the biotransformation of PAHs by C. elegans includes phase I and phase II reactions. The phase I reaction includes oxidation, reduction, and hydrolysis that generate hydroxylated metabolites such as 2-hydroxycarbamazepine, hydroxyflumequine, hidroxywarfarin, etc. and sulfoxidated products such as chlorpromazine sulfoxide. These reactions are highly regio- and stereoselective. Sometimes, these compounds undergo phase II reactions to form conjugated metabolites such as fluoresomidearyl glucoside. Other representatives of Mucoromycotina such as Umbelopsis ramanniana and Mucor rammanianus have been investigated in microbial transformations of carbamazepine (Kang et al. 2008) and fluoroquinolones, such as sarafloxacin and enrofloxacin, respectively (Parshikov et al. 2000, 2001).

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9.3  Fungal Enzymes Involved in Remediation As a type of bioremediation technology, microbial remediation, including bioaugmentation or biostimulation, biodegrades petroleum hydrocarbons contained in petroleum-contaminated soil by using the growth activities of microorganisms (Wang et al. 2012; Kuppusamy et al. 2016a; Rana et al. 2019; Yadav et al. 2017a, 2018). In remediation studies of the petroleum-contaminated soil, investigators have found several microorganisms which can be used in biodegradation of petroleum hydrocarbons (Kuppusamy et al. 2016b; Liu et al. 2017). White-rot fungi have been reported to degrade petroleum hydrocarbons by secreting some enzymes like lignin peroxidase (LiP), manganese peroxidase (MnP), and laccase (Canet et  al. 2001; McErlean et al. 2006). Fungi play a major role as decomposers and symbionts in all ecosystems (soil and aquatic habitats) owing to their robust morphology and diverse metabolic capacity, and because of that, they are best suited for the purpose of bioremediation. In bioremediation, when fungi are used as degrader to decontaminate contaminated areas, it is termed as mycoremediation (Shearer et al. 2007). There has been increasing interest in the special capacity of fungi to degrade pollutants by employing a number of extracellular and intracellular enzyme systems such as peroxidases and cytochrome P450, respectively, for the biodegradation and detoxification (Jebapriya and Gnanadoss 2013; Morel et  al. 2013; Durairaj et  al. 2015). In contrast to bacteria, white-rot fungi require a higher nutrition level for growth but have a higher-grade oxidase system presenting better degradation ability for complex petroleum hydrocarbons (D'Annibale et al. 2006; Mohan et al. 2006; Liu et al. 2017). Figure 9.2 represents the mechanism of bioremediation of hazardous chemicals used by fungi.

Fig. 9.2  Mechanism of bioremediation of hazardous chemical used by fungi (Deshmukh et al. 2016)

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9.3.1  Laccase Laccases constitute a family of multicopper oxidases belonging to group of blue oxidases which use copper as cofactor and molecular oxygen as cosubstrate. It is produced by certain plants, fungi, insects, and bacteria. Laccases catalyze mainly the oxidation of a wide range of reduced phenolic and aromatic substrates with concomitant reduction of molecular oxygen to water (Mate and Alcalde 2017; Yadav et al. 2016, 2017b). There are multiple isoenzymes of laccases that are known to occur which are encoded by a separate gene (Giardina et al. 1995). The nonspecificity of laccases makes them ideal for a variety of substrate catalysts for a variety of industrial applications of which these enzymes have been extensively explored for their effective potential for bioremediation (Baldrian 2006; Hildén et al. 2009). However, the mechanism of action of laccases under extreme conditions is less explored. The crystal structure of only a few laccases from ascomycetes (Melanocarpus albomyces (MaL) and Thielavia arenaria (TaLcc1) is fully known (Arora and Rampal 2002). In spite of their incredible potential of bioremediation, the utility of laccases is limited by their low shelf life. Among the biological agents, laccases represent an interesting group of ubiquitous, oxidoreductase enzymes that show promise of offering great potential for biotechnological and bioremediation applications (Gianfreda et al. 1999).

9.3.2  Catalase As reported in other biological systems, generation and accumulation of reactive oxygen species (ROS) mainly result in damage to macromolecules (cellular), which is deleterious for cellular integrity. Primary defense mechanism to ROS generation in fungi consists of monofunctional catalases and bifunctional peroxidase/ catalase enzymes. It has been suggested that catalase activity could be used as monitoring tool for monitoring bioremediation efficiency since their study revealed that catalase activity decreased with increasing oil concentration during bioremediation of oil contaminated soil (Deshmukh et al. 2016). Thus, considering the significance of catalases in providing heavy metal tolerance capacity to fungi, fungi producing this enzyme can be promising candidates for bioremediation of metal contaminated sites (Marco et al. 2013).

9.3.3  Tyrosinases Tyrosinases are produced by some fungal species including Ascomycota, Basidiomycota, and Mucoromycotina. It is a natural enzyme and is often purified to only a low degree and it is involved in a number of functions which primarily

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catalyze the o-hydroxylation of monophenols into their corresponding o-diphenols and further the oxidation of o-diphenols into o-quinones using molecular oxygen, which finally polymerizes to form brown or black pigments (Zaidi et  al. 2014). According to Ba and Kumar (2017), tyrosinases has been investigated as “green products” in the removal of several chemical contaminants in waters as well as soils. Different fungal groups have been studied for the isolation of tyrosinase and obtained from Agaricus bisporus, Neurospora crassa, Amanita muscaria, Lentinula edodes, Aspergillus oryzae, portobello mushrooms, Pycnoporus sanguineus, and Lentinula boryana (Zaidi et al. 2014).

9.3.4  Peroxidase Peroxidase catalyzes the oxidation of a wide variety of chemical compounds. This enzyme occurs in plants, animals, and microorganisms. The biological functions and specificity of peroxidases vary with sources of the enzyme. Some fungal groups such as Basidiomycota and Ascomycota secreted extracellular enzymes that play important role in bioremediation (Silva et al. 2016). On the basis of source and activity of peroxidases, it is classified into lignin peroxidase (LiP), manganese peroxidase (MnP), and versatile peroxidase (VP). Among peroxidases, LiP, MnP, and VP have been studied the most because of their high potential to degrade toxic substances in nature. LiP are heme proteins mainly secreted during secondary metabolism by the white-rot fungus. LiP plays a central role in the biodegradation of the plant cell wall constituent lignin. Degradation of lignin and other phenolic compounds is mediated by LiP in the presence of cosubstrate H2O2 and mediator like veratryl alcohol. LiPs have higher redox potential to oxidize aromatic compounds, and the exact redox mechanism is still poorly understood. MnP is an extracellular heme enzyme that oxidizes Mn2+ to the oxidant Mn3+ in a multistep reaction from the lignin-degrading basidiomycetes fungus. It has been reported that white-­rot and basidiomycetes fungi degraded most of toxic compounds. On the other hand, VP enzymes are broad substratespecific enzymes having capability of oxidizing both phenolic and nonphenolic compounds and are highly valued for biotechnological processes such as bioremediation (Hiner et al. 2002; Koua et al. 2009). There are some other peroxidases (Coprinopsis cinereal peroxidase, dye-­ decolorizing peroxidases, and Caldariomyces fumago heme-thiolate chloroperoxidase) secreted extracellularly by Basidiomycota and Ascomycota. Among these, Coprinopsis cinereal peroxidase and dye-decolorizing peroxidases catadependent one-electron oxidation of organic compounds. lyze H2O2-­ Caldariomyces fumago heme-thiolate chloroperoxidase catalyze H2O2-dependent peroxygenation (two-­ electron oxidation), leading to epoxidation of (cyclo) alkenes, hydroxylation of benzylic carbon, and sulphoxidation of S-containing organic compounds (Liers et al. 2011).

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9.3.5  Fungal Unspecific Peroxygenases A secreted heme-thiolate peroxidase with promiscuity for oxygen transfer reactions was discovered in the basidiomycetous fungus, Agrocybe aegerita. The enzyme turned out to be a functional monoperoxygenase that transferred an oxygen atom from hydrogen peroxide to diverse organic substrates like aromatics, heterocycles, linear and cyclic alkanes/alkenes, fatty acids, etc. (Karich et al. 2017). Later, similar enzymes were investigated from other genera of mushroom such as Coprinellus and Marasmius (Hofrichter et al. 2014). This new enzyme type was classified as unspecific peroxygenase (UPO, EC 1.11.2.1) and placed in a separate peroxidase subclass. Furthermore, UPOs and related heme-thiolate peroxidases such as well-studied chloroperoxidase (CPO) represent a separate superfamily of heme proteins on the phylogenetic level. The reactions catalyzed by UPOs include hydroxylation, epoxidation, O- and N-dealkylation, aromatization, sulfoxidation, N-oxygenation, dechlorination, and halide oxidation (Hofrichter et al. 2015; Peter et al. 2013).

9.3.6  Monooxygenases Monooxygenases are classified into two subclasses based on the presence cofactor: flavin-dependent monooxygenases and P450 monooxygenases. Oxygenases are grouped into two categories, the monooxygenases and dioxygenases, on the basis of number of oxygen atoms used for oxygenation. The enzymes of this group are generally cell bound. They play a key role in the metabolism of organic compounds by increasing their reactivity or water solubility or bringing about cleavage of the aromatic ring. Oxygenases have a broad substrate range and are active against a wide range of compounds, including the chlorinated aliphatic. The desulfurization, dehalogenation, denitrification, ammonification, hydroxylation, biotransformation, and biodegradation of various aromatic and aliphatic compounds are catalyzed by monooxygenases. These properties have been explored in recent years for important application in biodegradation and biotransformation of aromatic compounds (Arora et al. 2010).

9.3.7  Phenol 2-Monooxygenases Phenol 2-monooxygenases are studied under flavin-dependent monooxygenases that contain flavin as prosthetic group and require NADP or NADPH as coenzyme. Monooxygenases act as biocatalysts in bioremediation process and synthetic chemistry due to their high region selectivity and stereoselectivity on a wide range of substrates. Majority of monooxygenases studied previously are having a cofactor, but there are certain monooxygenases which function independently of a cofactor. These enzymes require only molecular oxygen for their activities and utilize the substrate as reducing agent (Cirino and Arnold 2002; Arora et al. 2010).

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9.3.8  Cytochrome P450 Monooxygenases Fungi possess complex oxidative and hydrolytic enzymatic systems for detoxifying toxic compounds in the environment. Besides these systems, certain fungi possess intracellular networks which constitute the xenome, consisting of cytochrome (CYP) P450 monooxygenases for dealing with diverse range of pollutants. The fungal cytochrome P450 system can serve as versatile catalyst for region- and stereo-­ specific oxidation of nonactivated hydrocarbons and can be ideal substitutes for chemical catalysts (Urlacher and Girhard 2012). Ichinose (2013) has highlighted the significance of cytochrome P450 systems in metabolism of series of endogenous and exogenous compounds. CYP63A2 P450 monooxygenase from white-rot fungus P. chrysosporium oxidized crude oil aliphatic hydrocarbon n-alkanes, endocrine-­ disrupting long-chain alkylphenols (APs), and mutagenic/carcinogenic fused-ring high molecular weight PAHs (HMW-PAHs) (Syed et al. 2013). Preinduction of the P450 monooxygenase before application in degradation studies could result in enhanced PAH removal. Enhanced removal of pollutants also achieved by molecular tools aimed at rapid and over production of cytochrome P450 monooxygenase (Bhattacharya et al. 2013; Guengerich and Munro 2013; Theron et al. 2014)

9.3.9  Nitroreductases Among the enzymes identified as involved in the degradation of TNT are the nitroreductases. This group of enzymes reduces a wide range of nitroaromatic compounds such as nitrofurazones, nitroarenes, nitrophenols, and nitrobenzenes including explosives such as TNT, RDX, and GTN. TNT (2,4,6-trinitrotoluene) degradation by fungi is initiated by mycelia bound nitroreductases which reduce TNT to hydroxyl amino dinitro toluene and aminodinitrotoluenes. Further degradation of these products and mineralization is achieved through the activity of oxidative enzymes especially lignin degrading enzymes (lignin and manganese peroxidases). Nitroreductases have generated a significant amount of interest recently for application in bioremediation of nitroaromatic compounds (Spain 1995).

9.3.10  Quinone Reductases It has been reported the involvement of quinone reductases in the degradation of several chlorinated aromatics by P. chrysosporium. Occurrence of quinone reductases in white-rot and brown-rot basidiomycetes has been reported. It catalyzes NAD(P)H-dependent reduction of quinones and is a cell-bound enzyme (Akileswaran et al. 1999).

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9.3.11  Reductive Dehalogenases Reductive dehalogenation proceeds via the replacement of a halide ion by a hydrogen atom and requires the transfer of two electrons. The role of electron donor can be fulfilled by a reduced organic substrate or H2 (Habash et al. 2004).

9.3.12  Miscellaneous Transferases Various transferases act on hydroxyl groups of pollutants and their metabolites to catalyze the formation of conjugates. which are typically not subject to further fungal degradation. These are also cell-bound enzymes. Detoxification through the excretion of water-soluble conjugates is well documented for the fungal metabolism of PAHs and other organic pollutants (Hundt et al. 2000; Cerniglia and Sutherland 2010).

9.4  Fungal Degradation of Hazardous Chemicals Certain hazardous chemical compounds, including polycyclic aromatic hydrocarbons (PAHs), penta-chlorophenols (PCPs), polychlorinated biphenyls (PCBs), polychlorinated dibenzo-p-dioxins (PCDDs), polychlorinated dibenzofurans (PCDFs), 1,1,1-trichloro-2,2-bis(4-chlorophenyl) ethane (DDT), benzene, toluene, ethylbenzene, and xylene (BTEX), trinitrotoluene (TNT), and heavy metals, are persistent in the environment and have carcinogenic and/or mutagenic effects. Environment is being loaded with a large quantum of such types of contaminants and recalcitrant compounds. Many conventional physicochemical methods of treatment/removal of these compounds, though effective, are not feasible for application on large scale (Akcil et al. 2015; Rastegari et al. 2019; Yadav et al. 2019a, b). Other toxic materials include polychlorinated biphenyls, dioxins, phenols, chlorophenols, pesticides, effluents from pulp and paper mills, dyestuffs, and heavy metals that have been successfully degraded by using white-rot fungi (Bennett 2007). Bioremediation does not involve only the degradation of pollutants, but also it is sufficient to remove the pollutant from the environment without degrading it (Broda 1992). However, bioremediation by fungal species has been reported to recognize as environment-friendly and economical. In the bioremediation, the efficient conversion of toxic and recalcitrant compounds into nontoxic or less toxic products takes place by applying natural biological processes, especially in the case of contaminated land and water. The technique involved in this process is use of suitable microbes in the polluted area which perform different physical and chemical reactions (as a part of their metabolism) resulting in degradation and removal of pollutants (Gillespie and Philip 2013; Mishra and Malik 2014). Bioremediation of hazardous chemicals can involve either natural attenuation, biostimulation, and bioaugmentation processes or a combination thereof. These processes have been appro-

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priately demonstrated during the bioremediation of atrazine (Sagarkar et al. 2013), petroleum hydrocarbons, and TNT in soil microcosms (No˜lvak et al. 2013; Lien et al. 2015). Bioremediation can be considered as a “green technology” due to its dependency on biological organisms and processes, and it does not require any treatment such as chemical addition or heating (Juwarkar et al. 2010). Bioremediation technology for microbial remediation involves bioaugmentation and/or biostimulation that biodegrades petroleum hydrocarbons contained in petroleum-­contaminated soil. Several biodegrades of petroleum hydrocarbons have been found in remediation of petroleum-contaminated soil. Of these, there are some bacterial species (isolated from petroleum-contaminated soil) that grow with petroleum hydrocarbons by using their carbon source, and white-rot fungi also degrade petroleum hydrocarbons by secreting enzymes such as LiP, MnP, and laccase. In contrast to bacteria, white-rot fungi have a higher grade of oxidase system and degradation ability for complex petroleum hydrocarbons. White-rot fungi also need a higher nutrition level (Liao et al. 2012). A number of researchers reported the widespread use of pesticides in public health and agricultural programs has caused severe environmental pollution and health hazards. The use of fungal species in degradation of pesticides has been reported in the literature (Singh et al. 2017). A large number of fungal species have been identified for their ability to degrade phenylurea herbicides (Khadrani et al. 1999). Sometimes, the fungal species capable of degrading pesticide have been shown to utilize the compound as a source of carbon and nitrogen (Kulshrestha and Kumari 2011). The filamentous fungus belonging to the group of nonligninolytic fungus, i.e., Cunninghamella elegans, a typical soil fungus, has been studied extensively for its ability to transform PAHs that migrate to the sediment in aquatic ecosystems due to their hydrophobic nature and low water solubility (Cerniglia 1997; Moody et al. 2004). The same fungi can rapidly oxidize PAH compounds such as benz-anthracene, 6-nitrochrysene, 6-nitrobenzo-pyrene, 3-nitrofluoranthene, 2-nitrofluorene, dibenzofuran, 1-nitropyrene, fluoranthene, naphthalene, acenaphthene, fluorene, phenanthrene, and nitrochrysene into an array of metabolites. Some other species, such as Cyclothyrium sp., Penicillium simplicissimum, and Psilocybe sp., transformed pyrene, anthracene, phenanthrene, and benzo[a]pyrene (Silva et al. 2003; Silva and Esposito 2004; Hammel et al. 1991). Degradation of the benzene-­ toluene-­ethylbenzene-xylene (BTEX) group of organopollutants by the white-rot fungus P. chrysosporium was studied (Oh et al. 1998; Yadav and Reddy 1993).

9.5  Recent Advances With recent advances in biomolecular engineering, the bioremediation of persistent organic pollutants using genetically modified microorganisms has become a rapidly growing area of research for environmental protection. Biomolecular engineering holds opportunities for the rapid advancement of bioremediation technology and offers the prospect of degrading some of the most recalcitrant. Most of the examples

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focused on the redesign of various features of the enzymes involved in the bioremediation of persistent organic pollutants, including the enzyme expression level, enzymatic activity, and substrate specificity. Now, with the recent advances in biomolecular engineering, the prospect of short circuiting the process of natural evolution to degrade environmental xenobiotic pollutants has been created.

9.6  Conclusion and Future Prospects Environmental hazard is growing largely due to the indiscriminate and frequently deliberate release of hazardous, harmful substances into the atmosphere. Chemicals that are used in domestic, industrial, and agricultural processes may lead to environmental complications when they deteriorate the quality of soil as well as groundwater. Among biological agents, enzymes have a biochemical potential that can be effectively transform and detoxify polluting substances. Simply, bioremediation is the use of biological organisms for cleaning up hazardous chemical by decreasing the toxicity levels of chemical compounds and restoring in its natural conditions. Fungal enzymes are capable of metabolizing, immobilizing, or absorbing toxic compounds from their environment. However, a major advantage of these systems is that they are less harmful to the environment with minimum or no by-products. Thus, evaluation of earlier research work may contribute to understanding that microbes are capable of completely removing pollutants from the environment and also leads to synthesis of useful compounds as by-products. Bioremediation is a cost-effective and eco-friendly biotechnological treatment that is capable of reducing and even eliminating pollution powered by microbial enzymes. Acknowledgments  NS and AK are grateful to the University Grant Commission, New Delhi, for providing financial assistance in the form of a Research Fellowship. The authors acknowledge UGC-SAP and DST-FIST for the support to the Department of Biochemistry, University of Allahabad, Allahabad, India. The authors declare no conflict of interest.

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Shearer CA, Descals E, Kohlmeyer B, Kohlmeyer J, Marvanová L, Padgett D, Porter D, Raja HA, Schmit JP, Thorton HA, Voglymayr H (2007) Fungal biodiversity in aquatic habitats. Biodivers Conserv 16:49–67 Silva M, Esposito E (2004) O papel dos fungos na recupera¸c˜ao ambiental. In Fungos: Uma Introdu¸c˜ao a Biologia, Bioquimica e Biotecnologia. Esposito E and De Azevedo JL (Eds.). EDUCS Ed 2:337–375 Silva M, Cerniglia CE, Pothuluri JV, Canhos VP, Esposito E (2003) Screening filamentous fungi isolated from estuarine sediment for the ability to oxidize polycyclic aromatic hydrocarbons. World J Microbiol Biotechnol 19:399–405 Silva MC, Torres JA, Castro AA, da Cunha EF, Alves de Oliveira LC, Corrêa AD, Ramalho TC (2016) Combined experimental and theoretical study on the removal of pollutant compounds by peroxidases: affinity and reactivity toward a bioremediation catalyst. J Biomol Struct Dyn 34:1839–1848 Singh N, Gupta VK, Kumar A, Sharma B (2017) Synergistic effects of heavy metals and pesticides in living systems. Front Chem 5:1–9 Spain JC (1995) Biodegradation of nitroaromatic compounds. Annu Rev Microbiol 49:523–555 Stajich JE, Berbee ML, Blackwell M, Hibbett DS, James TY, Spatafora JW, Taylor JW (2009) The fungi. Curr Biol 19:R840–R845 Syed K, Porollo A, Lam YW, Grimmet PE, Yadav JS (2013) CYP63A2, a catalytically versatile fungal P450 monooxygenase capable of oxidizing higher-molecular-weight polycyclic aromatic hydrocarbons, alkylphenols, and alkanes. Appl Environ Microbiol 79:2692–2702 Theron CW, Labuschagné M, Gudiminchi R, Albertyn J, Smit MS (2014) A broad-range yeast expression system reveals Arxula adeninivorans expressing a fungal self-sufficient cytochrome P450 monooxygenase as an excellent whole-cell biocatalyst. FEMS Yeast Res 14:556–566 Ullrich R, Hofrichter M (2007) Enzymatic hydroxylation of aromatic compounds. Cell Mol Life Sci 64:271–293 Urlacher VB, Girhard M (2012) Cytochrome P450 monooxygenases: an update on perspectives for synthetic application. Trends Biotechnol 30:26–36 Vidali M (2001) Bioremediation. An overview. Pure Appl Chem 73:1163–1172 Wang X, Cai Z, Zhou Q, Zhang Z, Chen C (2012) Bioelectrochemical stimulation of petroleum hydrocarbon degradation in saline soil using U-tube microbial fuel cells. Biotechnol Bioeng 109(2):426 Weber SD, Hofmann A, Pilhofer M, Wanner G, Agerer R, Ludwig W, Schleifer KH, Fried J (2009) The diversity of fungi in aerobic sewage granules assessed by 18S rRNA gene and ITS sequence analyses. FEMS Microbiol Ecol 68:246–254 Wu Y, Teng Y, Li Z, Liao X, Luo Y (2008) Advances in applied bioremediation. Soil Biol Biochem 40:789–796 Yadav JS, Reddy CA (1993) Degradation of benzene, toluene, ethylbenzene, and xylenes (BTEX) by lignin–degrading basidiomycetes Phanerochaete chrysosporium. Appl Environ Microbiol 59:756–762 Yadav JS, Doddapaneni H, Subramanian V (2006) P450ome of the white rot fungus Phanerochaete chrysosporium: structure, evolution and regulation of expression of genomic P450 clusters. Biochem Soc Trans 34:1165–1169 Yadav AN, Sachan SG, Verma P, Kaushik R, Saxena AK (2016) Cold active hydrolytic enzymes production by psychrotrophic Bacilli isolated from three sub-glacial lakes of NW Indian Himalayas. J Basic Microbiol 56:294–307 Yadav A, Verma P, Kumar R, Kumar V, Kumar K (2017a) Current applications and future prospects of eco-friendly microbes. EU Voice 3:21–22 Yadav AN, Kumar R, Kumar S, Kumar V, Sugitha T, Singh B, Chauhan VS, Dhaliwal HS, Saxena AK (2017b) Beneficial microbiomes: biodiversity and potential biotechnological applications for sustainable agriculture and human health. J Appl Biol Biotechnol 5:1–13 Yadav AN, Verma P, Kumar V, Sangwan P, Mishra S, Panjiar N, Gupta VK, Saxena AK (2018) Biodiversity of the genus Penicillium in different habitats. In: Gupta VK, Rodriguez-Couto S (eds) New and future developments in microbial biotechnology and bioengineering, Penicillium

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Chapter 10

Biotechnological Applications of β-Glucosidases in Biomass Degradation Sushma Mishra, Deepika Goyal, Amit Kumar, and Prem Kumar Dantu

10.1  Introduction Cellulose, the chief chemical constituent of primary cell walls, forms the most abundant group of carbohydrates produced by plants. For example, the cellulose content forms ~90% in cotton fibres, 40–50% in wood and approximately 57% in dried hemp. In contrast to starch, cellulose is an unbranched linear polymer of D-glucose units linked by β(1→4) glycosidic linkages between C#1 of one glucose and C#4 of the next glucose (Shahzadi et al. 2014). In fact, it is these beta linkages present between the monomer units that enable the formation of long, rigid chains of cellulose microfibrils that bear numerous intra- and intermolecular hydrogen bonds. The chains are oriented in parallel and form highly ordered, crystalline domains that are responsible for high tensile strength of plant cell walls (Beguin and Aubert 1994; Shahzadi et al. 2014). In nature, cellulose is degraded mainly by fungi and bacteria. The degradation of cellulose to glucose molecules is catalysed by the synergistic activity of three individual enzymes (Fig.  10.1): endoglucanase (1,4-β-D-glucan hydrolase, EC 3.2.1.4), exoglucanase (1,4-β-D-glucan glucohydrolase EC 3.2.1.74) and β-glucosidase (β-D-glucoside glucohydrolase EC3.2.1.21) (Dashtban et al. 2010; Tiwari et al. 2013; Seo et al. 2013; Lambertz et al. 2014). Exoglucanases, also known as cellobiohydrolase, hydrolyse cellulose polymers from the ends releasing mainly cellobiose, a disaccharide consisting of two β-glucose molecules. Endoglucanases hydrolyse glucosidic bonds at random positions in cellulose chains to generate oligosaccharide chains of different length, also

S. Mishra (*) · D. Goyal · P. K. Dantu Department of Botany, Dayalbagh Educational Institute, Deemed University, Dayalbagh, Agra, India A. Kumar Host Plant Section, Central Muga Eri Research & Training Institute, Central Silk Board, Lahdoigarh, Jorhat, Assam, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_10

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CELLULOSE CHAIN

CELLOBIOSE

GLUCOSE

Exoglucanase Endoglucanase β 1,4-glucosidic bond β-glucosidase Glucose monomer Fig. 10.1  Cellulose degradation by the synergistic action of endoglucanase, exoglucanase and β-glucosidase

producing new sites to be attacked by exoglucanases. Finally, β-glucosidase breaks down cellobiose and short oligosaccharides into glucose units (Kumar et al. 2008; Sukumaran et al. 2005). In other words, in the enzymatic hydrolysis of cellulose, endoglucanases and exoglucanases are responsible for degrading cellulose to cellobiose, after which β-glucosidases hydrolyse cellobiose to free glucose molecules. In this process, the step catalysed by β-glucosidases is generally the ratelimiting step and hence is responsible for the regulation of the entire cellulose degradation process. This inhibition is mainly caused due to the inhibitory effects by cellobiose on both endoglucanase and exoglucanase activities (Bok et al. 1998; Kruus et al. 1995). The most widely accepted system of classification of β-glucosidases is based on their nucleotide sequence identity (NCI) and hydrophobic cluster analysis (HCA). In this system, enzymes with overall amino acid sequence similarity and well-­ conserved sequence motifs are placed in a family (Henrissat 1991; Cairns and Esen 2010). According to the data collected in June 2018, 153 glycoside hydrolase (GH) families are listed in Carbohydrate Active enZYme website (http://www.cazy.org). This is the most widely accepted classification, and β-glucosidases are placed in glycoside hydrolase (GH). Most of the β-glucosidases are reported in GH1, GH3, GH5, GH9, GH30 and GH116 families. HCA system of classification is believed to reflect structure, evolutionary relationship and catalytic mechanisms of this

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enzyme (Cairns and Esen 2010). β-Glucosidases belonging to GH family 1 are mainly reported from archaebacteria, plants and animals, whereas β-glucosidases belonging to GH family 3 are from bacteria, fungi and yeast. β-Glucosidases could also be classified on the basis of substrate specificity into three classes: aryl-β-glucosidases that hydrolyse only aryl-β-glucoside linkage, cellobiases that hydrolyse only disaccharides and broad substrate specificity β-glucosidases that hydrolyse wide range of substrates with different bonds such as β(1→4), β(1→3), β(1→6), α(1→4), α(1→3) and α(1→6) (Singh et al. 2016). Cellulose recycling forms an important part of the carbon cycle in biosphere, as it is the major carbohydrate synthesised by plants. As mentioned above, this degradation process is brought about by the synergistic activity of a series of enzymes, where the terminal steps catalysed by β-glucosidases form the rate-limiting step. Consequently, the entire cellulolytic process is limited by the activity of this enzyme. Hence, an increased understanding of the factors affecting β-glucosidase activity would promote efficient conversion of the otherwise abundant cellulose into the much needed biofuels and other economically important products. With this objective in mind, the authors have tried to present an overview of β-glucosidase enzyme and its biotechnological applications, followed by major rate-limiting components and possible solutions for large-scale conversion of lignocelluloses into ethanol.

10.2  Sources of β-Glucosidases β-Glucosidases are a class of hydrolytic enzymes produced by various organisms, ranging from microorganisms to higher plants and animals. Wood-degrading organisms like termites and wood-decomposing fungi have been generally targeted by the researchers for isolating cellulolytic enzymes (Sanderson 2011). Among plants, β-glucosidases have been well characterised from Arabidopsis thaliana, rice, cherry, wheat, sorghum and maize (Tiwari and Verma 2017; Sue et al. 2006; Dharmawardhana et al. 1995; Seshadri et al. 2009; Kittur et al. 2007). Some of the in-planta functions of β-glucosidases include chemical defence, plant–microbe interactions, cell wall remodelling, alkaloid metabolism and phytohormone regulation (Seshadri et  al. 2009; Singh et al. 2016). However, for industrial production of β-glucosidases, fungi are the best source due to several advantages like high yield, fast growth, cost-effectiveness, etc. (Kour et  al. 2019b; Yadav et  al. 2015, 2016a, b, 2017b, 2019a, b). Many fungi such as Trichoderma reesei (Chen et al. 1992), the filamentous fungus Acremonium persicinum (Pitson et al. 1997), Aspergillus oryzae (Riou et al. 1998), Thermoascus aurantiacus (Parry et  al. 2001), Chaetomium thermophilum (Venturi et  al. 2002), Penicillium purpurogenum (Karnchanatat et  al. 2007), Daldinia eschscholzii (Kaur et al. 2007), Melanocarpus sp. MTCC 3922 (Chen et al. 2012), Neocallimastix patriciarum W5 (Daroit et al. 2008), Monascus purpureus and brown-rot basidiomycete Fomitopsis palustris (Yoon et al. 2008) produce β-glucosidase (Tables 10.1 and 10.2). In addition, this enzyme has recently been produced from Penicillium

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Table 10.1  Large-scale production methods of β-glucosidases from fungi Fungal species Tolypocladium cylindrosporum Syzx4 Penicillium simplicissimum H-11 Aspergillus strain SA 58 Penicillium citrinum YS40-5 Fusarium proliferatum Fusarium solani Aspergillus niger + A. oryzae Fomitopsis palustris Aspergillus niger SOI017 Flammulina velutipes Monascus sanguineus Phoma sp. KCTC11825BP Thermomucorindicae-­ seudaticae N31 Aspergillus niger HDF05 Gongronella butleri Penicillium miczynskii Fusarium oxysporum Aureobasidium pullulans Candida peltata Kluyveromyces marxianus Aureobasidium sp. Saccharomyces cerevisiae

Fermentation method Submerged fermentation (SMF)

References Bai et al. (2013)

Submerged fermentation

Elyas et al. (2010)

Solid-state fermentation (SSF) Solid-state fermentation Submerged fermentation Solid-state fermentation Solid-state fermentation Submerged fermentation Submerged fermentation Submerged fermentation Solid-state fermentation

Ng et al. (2010) Bhatti et al. (2013) Gao et al. (2012) Raza et al. (2011) Vaithanomsat et al. (2011) Yoon et al. (2008) Qian et al. (2012) Mallerman et al. (2015) Dikshit and Tallapragada (2015) Choi et al. (2011) Ling et al. (2011)

Submerged fermentation Solid-state fermentation Solid-state fermentation Solid-state fermentation Submerged fermentation Submerged fermentation Submerged fermentation Submerged fermentation Submerged fermentation Solid-state fermentation + submerged fermentation Submerged fermentation

Cassia et al. (2015) Ling et al. (2011) Beitel and Knob (2013) Santos et al. (2016) Saha et al. (1994) Olajuyigbe et al. (2016) Rajoka et al. (2004) Saha and Bothast (1996) Iembo et al. (2002)

purpurogenum KJS506, Phoma sp. KCTC11825BP (Choi et al. 2011), Aspergillus fumigatus Z5 (Liu et al. 2012), Penicillium italicum (Park et al. 2012), Fusarium proliferatum NBRC109045 (Gao et al. 2012), Aspergillus saccharolyticus (Sorenson et  al. 2014), Aspergillus niger A20 (Abdel-Naby et  al. 1999), Fusarium solani (Bhatti et al. 2013), Flammulina velutipes (Mallerman et al. 2015), Monascus sanguineus, Sporothrix schenckii (Hernández et al. 2016), Gongronella butleri (Santos et al. 2016) and Fusarium oxysporum (Olajuyigbe et al. 2016). The fungal species, Aspergillus niger, is the major source of commercial β-glucosidase under the name of Novazym188 (Sorenson et al. 2013). β-Glucosidase has been identified, purified and characterised from several bacterial species as well, such as Clostridium thermocellum (Ait et  al. 1982), Pyrococcus furiosus (Kengen et al. 1993), Bacillus circulans subsp. Alkalophilus (Paavilainen et al. 1993), Flavobacterium johnsoniae (Okamoto et al. 2000), actinomycete Thermobifida fusca (Spiridonov and Wilson 2001), Lactobacillus brevis (Michlmayr et al. 2010), Caldicellulosiruptor saccharolyticus (Hong et al. 2009)

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Table 10.2  Optimum physical parameters for industrial production of β-glucosidases from microbial sources Preferred carbon Microorganisms source Aspergillus niger Wheat bran

Penicillium purpurogenum Candida peltate

Fusarium proliferatum Chaetomium thermophilum Aspergillus protuberus Penicillium citrinum

Preferred nitrogen source Ammonium sulphate

Preferred Preferred temp. pH References 30 °C 5.5 Raza et al. (2011), Vaithanomsat et al. (2011) Sucrose Sodium nitrate 28 °C 4.0 Jeya et al. (2010), Jeya and Lee (2013) Glucose and – 50 °C 5.0 Saha and Bothast xylose (1996), Rajoka et al. (2004) 25 °C 5.0 Gao et al. (2012) Corn stover Urea and wheat bran Cellulose Peptone, yeast 45 °C 5.5 Venturi et al. (2002) extract Glucose Ammonium 30 °C 3.0 Yadav et al. (2016a, b) sulphate Maltose Ammonium 70 °C 5.0 Ng et al. (2010) sulphate

and Terrabacter ginsenosidimutans (An et  al. 2010). Thermoanaerobacterium thermosaccharolyticum is known to produce a glucose-tolerant β-glucosidase (Pei et al. 2012).

10.3  Biotechnological Applications of β-Glucosidases The use of cellulases in biotechnology began in the early 1980s in animal feed and food industry (Chesson 1987; Thomke et al. 1980). Subsequently, these enzymes were used in textile, laundry as well as in pulp and paper industries (Godfrey et al. 1996; Wong and Saddler 1992). Today, β-glucosidases are also used for production of biofuels, detoxification of cassava cyanogenic glucosides and in the treatment of Gaucher’s disease (Cairns and Esen 2010; Prasad et al. 2012). The catalytic activity of β-glucosidases include hydrolysis of β(1–4), β(1–3), β(1–6) and β(1–2) glucosidic linkages in aryl-, amino-, or alkyl-β-D-glucosides, cyanogenic glucosides, and disaccharides, polysaccharides and glucose-substituted molecules. Apart from hydrolysis of sugars, some mutant β-glucosidases have been reported to catalyse synthetic reactions of sugars by reverse hydrolysis and transglycosylation (Mackenzie et  al. 1998). Due to the diverse types of reactions and substrates of β-glucosidases, they have several industrial applications, some of which have been described below.

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10.3.1  Conversion of Lignocellulose into Biofuels Lignocellulose refers to the woody parts of plants, mainly agricultural residues like wheat stems, corn stalks and leaves, and forestry wastes like wood shavings from logging. The lignocellulose, which is generally considered to be ‘plant waste’ and discarded, could be converted into energy sources. These so-called second-­ generation biofuels have many advantages in comparison to first-generation biofuels that were derived mainly from food crops (Sanderson 2011; Guo et al. 2015; Prasad et  al. 2014; Rastegari et  al. 2019; Prasad et  al. 2019;  Yadav et  al. 2017a, 2018). Apart from being a renewable and sustainable way of energy production, the bioconversion of lignocellulose into ethanol has an additional advantage of solving the problem of waste disposal of agricultural residues  and other biomass (Khan et  al. 2018, 2019). Lignocelluloses, chemically made up of cellulose, hemicellulose and lignin, which when converted into fermentable sugars, could be used to produce liquid fuels like ethanol or oil and gaseous fuels like biogas and electricity (Menon and Rao 2012; Prasad et al. 2009, 2013; Kour et al. 2019a; Rana et  al. 2019a, b). Briefly, the lignocellulose is first subjected to heat and acid or ammonia to separate lignin, thereby exposing cellulose and hemicelluloses. Thereafter, the combined action of exoglucanase, endoglucanase and β-glucosidases converts this polymer into glucose sugar, which could then be fermented to give rise to ethanol or the longer chain alcohol butanol. The pentose and hexose obtained after cellulose degradation could be fermented to ethanol by the action of yeast and certain enzymes. The preferred physical conditions required by some of the commonly used microbial sources of β-glucosidases for lignocelluloses degradation are mentioned in Table 10.2. The bioconversion of lignocellulose involves two steps: hydrolysis of cellulose in lignocellulosic biomass to produce reducing (fermentable) sugars, and fermentation of the sugars to ethanol (Sun and Cheng 2002). An outline of steps involved in biomass degradation is presented in Fig. 10.2. Briefly, plant parts are first cut into small size by either milling or chipping, followed by a pre-treatment step using either physical or chemical agents. The pre-treatment step mainly disrupts the close inter-component association between main constituents (cellulose, hemicellulose, lignin) of the plant cell wall (Jönsson and Martín 2016). Some of the most commonly used pre-treatment methods include acid hydrolysis with mineral acids/ organic acids, steam heating followed by sudden decompression, hydrothermal processing and oxidative methods (Jonsson and Martin 2016). These methods basically aim to remove hemicelluloses and/or lignin from the lignocellulosic matrix, thereby facilitating the subsequent enzymatic degradation of cellulose to D-glucose. Together, endoglucanases, exoglucanases and β-glucosidases make a potent system for cellulose degradation. These three enzymes could be present either as multienzymatic complex called cellulosome or exist as individual enzymes (Bae et al. 2013). Being the terminal enzyme in cellulose degradation pathway, β-glucosidases play a critical role in this process (Bhatia et al. 2002). If β-glucosidases are not present in sufficient amounts, not enough glucose will be produced, and cellobiose will

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Plant debris (lignocellulosic biomass) Cellulose

Hemicellulose

Lignin

PRE-TREATMENT OF LIGNOCELLULOSIC BIOMASS

Physical methods

Chemical methods

Biological methods

Ball milling

Sodium hydroxide

Pleurotus florida

Compression milling

Perchloric acid

Pleurotus cornucopiae

Cryomilling

Acetic acid

Streptomyces viridosporus

Steam treatment

Ammonia freeze explosion

Cyathus sp.

Ultrasound

Organic solvents

Microwave ENZYMATIC HYDROLYSIS OF PRE-TREATED CELLULOSE FERMENTATION OF SUGARS BIOETHANOL PRODUCTION

Fig. 10.2  Various steps involved in degradation of plant biomass. The pre-treatment of lignocellulosic biomass could be done either by physical, chemical or biological methods. Subsequently, the plant biomass is subjected to cellulase degradation to convert cellulose to sugars, which could be fermented to yield ethanol

accumulate. Since cellobiose is an inhibitor of endo- and exoglucanases, this would negatively affect glucose formation, making it the rate-limiting step of the pathway (Dekker 1986). Therefore, the activity of β-glucosidase could be regulated to increase the efficiency of conversion of cellulose to glucose (Dashtban et al. 2010; Lambertz et al. 2014). This aspect has been dealt with in detail in the next section on ‘Challenges in Lignocellulose Bioconversion’.

10.3.2  Production of High-Value Bioproducts Apart from ethanol, which forms the primary product of lignocellulose degradation, the by-products could be used to generate a number of organic chemicals. For example, the fermentation of hexoses and pentoses obtained after cellulose degradation could be used to produce lactic acid by using the bacteria, Bacillus coagulans (Patel et al. 2006). Likewise, the lignocelluloses conversion of palm waste produces

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xylose, which could be further processed to form xylitol, a sweetening agent (Rahman et  al. 2007). Another by-product, furfural, obtained from hydrolysis of hemicelluloses, is reported to be used in plastic, varnishes and herbicide preparation (Montane et al. 2002).

10.3.3  Hydrolysis of Isoflavone Glycosides Phenolic compounds like flavonoids, flavonones, flavones and isoflavones form a class of secondary metabolites in plants that have antioxidant, anticancerous, antiallergic, anti-inflammatory and antihypertensive properties (Kabera et  al. 2014; Karimi et al. 2012; Servili et al. 2013). Majority of these metabolites are present in the form of glycosides, which increase their water solubility and stability, but limit their absorption. The release of non-carbohydrate part requires the action of specific enzymes such as arabinosidases and β-glucosidases. For example, daizdin, genistin and glycitin are some of the glucosidic isoflavones in soybean and soy (a soybean-­ based food), which are generally found in an inactive state. Aglycone forms of these isoflavones, produced by β-glucosidases, exhibit phytoestrogenic properties and hence are useful in the treatment of various diseases like prostate cancer, breast cancer, cardiovascular disease and menopause treatment (Izumi et  al. 2000; Hati et al. 2015). The different microbial sources of β-glucosidases for the hydrolysis of isoflavone or flavonoid compounds are represented in Table 10.3.

10.3.4  Flavour and Nutrition Enhancement In plants, flavour compounds generally occur in the form of glycoconjugates, in order to suppress the flavour and make them non-volatile. The β-glucosidase enzyme releases the glycoconjugate form of flavour compounds, thereby imparting the unique flavour to plants. For example, β-glucosidase isolated from Sporidiobolus pararoseus and Aureobasidium pullulans have been found to hydrolyse terpenyl glycosides and improve aroma of wines (Baffi et al. 2013). Likewise, it has also been reported to improve the organoleptic properties and reduce the bitterness of citrus fruit, which is caused by the glucosidic compound, naringin (Roitner et  al. 1984). β-Glucosidase isolated from Bacillus subitilis is used for improving sugarcane juice quality (Singh et al. 2016) by immobilising it on alginate beads for industrial production. β-Glucosidases could also be used in the release of some nutritionally important components, such as vitamins and antioxidants from their glycoside-conjugated form. For example, vitamin B6 of rice could be released from pyridoxine glucoside form by the application of β-glucosidase (Opassiri et al. 2004).

Daidzein, Genistein, Glycitein Daidzein, Genistein, Glycitein

Daidzin, genistin

Daidzin, genistin, glycitin

Daidzin, genistin, glycitin

Bifidobacterium bifidum

Bacteroides thetaiotaomicron VPI-5482 Aspergillus terreus

Anticancer, osteoporosis, antihypercholesterolaemia

Anticancer, osteoporosis, antihypercholesterolaemia Anticancer, osteoporosis, antihypercholesterolaemia

Antiallergic, antioxidant, anti-­ inflammatory, antihypertensive

Hesperetin, Hesperetin, Haringenin, Naringenin, Quercetin, Rutinose Daidzein, Genistein

Sesaminol

(S)-Rh1 (S)-Rg2

Protopanaxatriol-type ginsenoside mixture (PPTGM) Paenibacillus sp. KB0549 2,6-O-di(β-D-glucopyranosyl)-β-D-­ glucopyranosylsesaminol (STG) Pyrococcus furiosus Hesperidin, neohesperidin, naringin, poncirin, diosmin, neoponcirin, rutin

Mucilaginibacters

Anticancer, osteoporosis Anti-inflammatory, anti-cancer, anti-aging, antioxidant activities Tonic, adaptogenic, immunomodulatory, anti-aging effects Antineoplastic, antistress, antioxidant activities Antioxidants

Yang et al. (2009)

Anticancer, osteoporosis, antihypercholesterolaemia Anticancer, antipostmenopausal syndrome Anticancer, osteoporosis, etc. Anticancer, osteoporosis, etc.

Yan et al. (2016)

Byun et al. (2015)

You et al. (2015)

Shin et al. (2013)

Nair et al. (2013)

Cui et al. (2013)

Yan et al. (2008)

Yang et al. (2004) Kuo et al. (2006), Kuo and Lee (2007) Fang et al. (2014) Choi et al. (2014)

Song et al. (2011)

References Ávila et al. (2009)

Biological activity Antioxidant

Daidzein, Genistein, Ginsenoside Rd, F2 Compound K (CK) Compound K

Genistein, Daidzein Genistein, Daidzein

Product Gallic, Syringe homogentisic acid Genistein, Daidzein, Glycitein Genistein, Daidzein

Gongronella sp. Daidzin and genistin Saccharomyces cerevisiae Ginseng HJ014 Paecilomyces bainier sp. Ginsenoside Rb1 229

Flavonoid glycoside Delphinidin-3-glucoside, malvidin-3-glucoside Paecilomyces thermophila Daidzin, genistin, glycitin J18 Thermoanaerobacter Daidzin, genistin ethanolicus JW200 Pseudomonas ZD-8 Genitin and daidzin Bacillus subtilis 18 Genistin and daidzin

Source of β-glucosidase L. acidophilus LA-5

Table 10.3  β-Glucosidases with the ability to hydrolyse flavonoid compounds

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10.3.5  Detoxification of Cyanogenic Glycosides in Cassava Cassava is a carbohydrate-rich food and a source of food for ~500 million people in the world. Cassava fruit contains cyanogenic glycosides like linamarin and lotaustralin, which have been reported to cause Konzo syndrome, a human central nervous system disorder (Vasconcelos et al. 1990). Although certain glucosidases are naturally present in cassava roots, their insufficient expression leaves a significant part of the cyanogenic glycosides in the processed food. Therefore, an additional treatment of cassava with β-glucosidases has the potential of detoxifying and liberating these cyanogenic glycosides, thereby improving its nutritional quality (Gueguen et al. 1997; Maduagwu 1983).

10.3.6  Deinking of Waste Paper Waste paper causes environmental pollution; its recycling can solve the two-­ dimensional problem of forest wood consumption and waste management. Removal of ink from paper is the most challenging obstacle, which could be overcome by using enzymes. The enzymatic method for waste paper recycling has been reported to be efficient in solving this problem. The enzyme preparations for waste paper recycling contain cellulase, β-glucosidase and hemicellulase (Prasad et  al. 1992; Pathak et al. 2011; Lee et al. 2013).

10.3.7  Applications Based on Synthetic Activity Apart from the hydrolytic activity, β-glucosidases also exhibit synthetic activity (through reverse hydrolysis and transglycosylation), leading to the production of oligosaccharides and alkyl and aryl β-glucosides (Ahmed et al. 2017). Alkyl glucosides like hexyl, heptyl and octyl glucosides are biodegradable and, therefore, find wide applications as emulsifier and antimicrobial agents (Bankova et  al. 2006). Synthetic glucosides are also used in the preparation of therapeutic drugs like heparin and acarbose. They act as probiotic agents and increase the number of useful microorganisms in human gut. Some of the in-planta functions of these oligosaccharides include fertilisation, embryogenesis and cell proliferation (Singh et  al. 2016; Krisch et  al. 2010). Galacto-oligosaccharides and isobutyl-galactosides are synthesised from lactose in water-organic media by the action of β-glucosidase produced by Aspergillus oryzae (Bankova et al. 2006). Arbutin-βglycoside was found to be synthesised via the transglycosylation reaction of β-glucosidase produced by Thermotoga neapolitana to manufacture a skin whitening agent, and these products were checked for their melanogenesis inhibitory activities (Jun et al. 2008).

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10.4  Challenges in Lignocellulose Bioconversion 10.4.1  Complex Structure of Plant Biomass The primary challenge in lignocellulose conversion to fermentable sugars is its complex structure: constituting approximately 33–40% of cellulose, 20–25% of hemicellulose and 15–20% of lignin (Hess et  al. 2011). Although cellulose is a simple homopolysaccharide composed of D-glucose residues, the linear cellulose microfibrils are associated with several hydrogen bonds that make the macromolecule highly crystalline and difficult to hydrolyse (Jørgensen et al. 2007). In addition, cellulose exists in complex with hemicelluloses, the heterogenous polysaccharides composed of variety of sugars, making it difficult to be converted into a single product, ethanol. In addition, the cellulose and hemicellulose are complexed with lignins, which form extensive cross-linking, making it resistant against microbial degradation.

10.4.2  I nhibition of Enzyme Activity Due to Pre-treatment Methods The pre-treatment of lignocellulosic biomass is often required to promote the subsequent step of enzymatic conversion of cellulose to sugars (Jørgensen et al. 2007). This step is basically aimed at removal of hemicelluloses and/or lignin from the lignocellulosic matrix A drawback of the pre-treatment step is formation of by-­ products like carboxylic acids, gluconic acid and glucaric acid, phenolic compounds, furfurals, benzoquinones, etc. that inhibit downstream processes by interfering with microbial activity (Jönsson and Martín 2016). For example, acid hydrolysis of lignocellulose results in corrosion of pre-treatment equipment and release of heavy metal ions like copper, nickel, chromium and iron, which can be inhibitory to fermenting microorganisms (Watson et al. 1984; Garrote et al. 2008).

10.4.3  End-Product Inhibition End-product inhibition is a method of negative feedback regulation, where the final product in a series of reactions inhibits an enzyme from an earlier step in the sequence. The product binds to an allosteric site of the enzyme and temporarily inactivates the enzyme via non-competitive inhibition. This mode of regulation is also seen in enzyme-mediated lignocellulose degradation. The enzymes, cellobiohydrolase and β-glucosidase, are subjected to end-product inhibition by cellobiose and glucose, respectively (Qing et al. 2010; Teugjas and Väljamäe 2013; Kumar and Wyman 2014). This limits the turnover number of these enzymes, leading to

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decline in product formation. This problem gets adverse when lignocellulosic degradation is performed at high solid concentrations in order to reduce the consumption of water and running cost of the process (Kristensen et al. 2009).

10.4.4  Other Challenges In addition to the above limitations, the activity of cellulases could also be inhibited by non-productive binding to lignins and residual hemicelluloses (Rahikainen et al. 2013; Pareek et al. 2013). Other factors limiting plant biomass degradation include relatively low activity of currently available hydrolytic enzymes, high cost of enzyme production and thermal inactivation of enzyme (Sun and Cheng 2002). The optimum fermentation conditions vary with species and other controlling parameters like source of carbon and nitrogen used in media. The carbon source may contain some contaminants in the form of secondary metabolites and chemicals that can interfere with the rate of β-glucosidase enzyme production. Therefore, a thorough screening of these secondary metabolites/inhibitors and their subsequent degradation or inactivation is crucial for the optimum enzyme production. Research should also be focused on the possibility of temperature stress on the yield and activity of β-glucosidase. Thermal inactivation of β-glucosidase is a major roadblock towards achieving high enzyme efficiency. The thermal stability of β-glucosidases could be enhanced by recombinant DNA technology and genetic modification of microbial strains. Precise genome editing using site-specific nucleases like CRISPR/Cas9 is a suitable option to achieve this goal. The other major hurdles in the commercial β-glucosidase production are product inhibition, low product yields and high cost of enzyme production. The search for better alternatives to the currently available enzyme preparations should be highly promoted. Isolation of novel fungal species having higher β-glucosidase activity would contribute towards revolutionising the field of lignocellulose-mediated production of biofuels.

10.5  Approaches for Enhancing β-Glucosidase Activity Researchers are trying to engineer cellulases with high specific activity, high thermal stability, high adsorption capacities, high catalytic efficiency and lower end-­product inhibition. Some of the major limitations in cellulase- or β-glucosidase-mediated biomass degradation have been addressed by using approaches like increasing the production of β-glucosidase through strain improvement by mutagenesis, co-cultivation of microbes in fermentation to increase the quantity of desirable components of cellulase complex, improving the performance of existing lignocellulose-degrading enzymes by genetic engineering, and finally, metagenomics approach that involves identification of novel β-glucosidases by DNA analysis of environmental samples. These approaches have been described below.

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10.5.1  Co-culturing As mentioned above, cellulose degradation requires synergistic action of three enzymes: exoglucanase, endoglucanase and β-glucosidase; however, no native microbial strain produces optimum amounts of all three enzymes under the same condition. For instance, while T. reesei produces exoglucanase and endoglucanase in abundant amounts, it produces β-glucosidase in very low amounts (Peterson and Nevalainen 2012). Likewise, Aspergillus niger produces large quantity of β-glucosidase, but limited amount of exoglucanase and endoglucanase (Stockton et al. 1991; Kumar et al. 2008). Therefore, co-cultivation of T. reesei and A. niger using paper mill sludge as a cellulosic substrate has proven to be a solution for efficient hydrolysis of cellulosic residues (Maheshwari et  al. 1994). Other successful cases include co-culturing Aspergillus ellipticus with A. fumigatus (Gupte and Madamwar 1997) and T. reesei with A. phoenicis using bagasse and corncobs as cellulose substrate in solid-state fermentation (Duenas et al. 1995). Different strains of Trichoderma fungus are used for production of beta glucosidase and are represented in Table 10.4. Table 10.4  Studies done on β-glucosidase isolated from different strains of Trichoderma Trichoderma strain β-Glucosidase T. citrinoviride Extracellular β-glucosidase T. reesei TrBgl2

References Chandra et al. (2013) Lee et al. (2012)

T. reesei QM9414

Dashtban and Qin (2012)

Recombinant T. reesei strain, X3AB1 T. reesei T. reesei CL847 T. reesei T. ressei T. harzianum C-4 T. reesei T. harzianum strain P1 T. reesei QM9414 T. viride

Isolation strategies Protein purification, biochemical and proteomic characterisation Mutational studies involving active site residues of the enzyme bgl1 Overexpression of bgl1 from Periconia sp. in T. reesei QM9414 under T. reesei tef1 promoter bgl1 Construction of T. reesei strain expressing Aspergillus aculeatus bg1 under control of xyn3 promoter bgl I Molecular cloning and expression in Pichia pastoris BGL1 Protein purification and kinetic characterisation β-Glucosidase Molecular cloning and expression in (cel3a) T. reesei β-Glucosidase Molecular cloning, expression in E. BGLII (Cel1A) coli and characterisation Protein purification and biochemical characterisation BGL2 Molecular cloning and expression in Aspergillus oryzae 1,3-β-Glucosidase Protein purification and characterisation Aryl-β-D-­ Protein purification and glucosidase characterisation β-Gluc I Protein purification and biochemical characterisation

Nakazawa et al. (2012) Chen et al. (2011) Chauve et al. (2010) Murray et al. (2004) Saloheimo et al. (2002) Yun et al. (2001) Takashima et al. (1999) Lorito et al. (1994) Chirico and Brown (1987) Beldman et al. (1985)

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10.5.2  Genetic Manipulation Genetic engineering approach involves introduction of specific desirable genes from one species to another species using recombinant DNA technology (Sticklen 2008). This strategy could be used to generate novel β-glucosidases with desirable properties like high efficiency, thermotolerance and high specificity for plant biomass degradation (Blumer-Schuette et al. 2014). T. reesei, the most commonly used source of cellulases, being mesophilic loses its enzyme activity at higher temperatures. Transformation of thermotolerant β-glucosidase genes into T. ressei from thermophilic fungus like T. emersonii was found to confer higher specific activity and temperature tolerance of up to 71.5 °C (Dashtban and Qin 2012; Druzhinina and Kubicek 2017). Similar results were obtained in Paenibacillus polymyxa where single amino acid substitution contributed increased thermal resistance (Garvey et al. 2013). Apart from targeting β-glucosidase, chimeric proteins have been constructed by the fusion of endoglucanase from Acidothermus cellulolyticus and exoglucanase from T. reesei, which resulted in improved saccharification (Chandel et al. 2012). Another attractive option of increasing cellulase production is to express cellulase from heterologous systems (Garvey et al. 2013). This involves codon optimisation, use of strong and inducible promoters and elimination of inhibitory sequences to enable efficient protein expression from heterologous systems. Cellulases were originally produced from anaerobic bacteria isolated from animal digestive systems (Chandel et al. 2012). In addition, recombinant systems like E. coli and Bacillus subtilis are being increasingly used for protein production because of the increased enzyme yields from these systems. Apart from bacterial expression hosts, yeasts like Saccharomyces cerevisiae, Pichia pastoris and Kluyveromyces marxianus have been employed due to its superior post-translational modification of secreted proteins (Tanaka et al. 2012).

10.5.3  Mutagenesis The inherently low β-glucosidase activity of T. reesei has been improved by mutagenesis through the use of chemical mutagens and UV radiations. The T. reesei RUT-C30 mutant was reported to produce 4–5 times higher β-glucosidase than wild T. reesei (Montenecourt and Eveleigh 1979). In another study, T. atroviride were modified through mutagenesis by the use of N-methyl-N′-nitro-N-nitrosoguanidine and UV light, and these mutants were found to have high cellulolytic activity than wild types (Kovacs et al. 2008). Apart from random mutagenesis through the use of physical and chemical mutagens, site-directed mutagenesis has also been used to enhance cellulase activity. In one of the study, Mahadevan et al. (2008) altered the amino acids around the active site of endoglucanase of Thermotoga maritima creating a mutant which displayed 10% higher activity than the wild-type enzyme. Similarly, mutation of the conserved residue F476 to Y476 from Cel9A of

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Thermobifida fusca displayed 40% improved cellulase activity. This was achieved through the integration of computer modelling with site-directed mutagenesis (Escovar-Kousen et al. 2004).

10.5.4  Metagenomics Approach A relatively recent approach involves analysis of DNA collected from environmental samples, enabling identification and quantification of microbial species that inhabit the natural environment. Metagenomics of microbial communities from cow rumen (Hess et al. 2011), termite hindgut (Warnecke et al. 2007) and mangroves (Simões et  al. 2015) have provided detailed insights into the diversity of lignocellulose-­degrading enzymes through the identification of uncultivable bacteria. Bergmann et  al. (2014) have isolated two novel β-glucosidases from soil of Amazon forest. In addition, new genes could be discovered that encode novel lignocellulolytic enzymes.

10.5.5  O  ther Strategies for Enhancing Lignocellulose Degradation One of the simple ways to prevent end-product inhibition of lignocellulose degradation is continuous elimination of end-products through sophisticated reactor designs (Andric et al. 2010). Another method to relieve end-product inhibition is simultaneous saccharification and fermentation, a process in which fermenting microorganism is added along with hydrolytic enzymes (Teugjas and Väljamäe 2013). This prevents accumulation of cellobiose and glucose in the reaction mixture that may interfere or inhibit cellulase activity. This method, however, has a major drawback that different conditions are required for optimal hydrolysis and fermentation. While the optimum temperature for yeast fermentation is approximately 35 °C, the optimum temperature of ~50 °C is optimal for cellulase activity. This issue could be addressed by the use of thermostable enzymes involved in fermentation of sugars, produced after cellulose hydrolysis, into ethanol.

10.6  Conclusion and Future Perspectives The first-generation biofuels, obtained primarily from food crops such as grains, sugar beet and oil seeds, have raised a number of concerns in terms of food security, climate change mitigation, economic growth and sustainability. Most of these concerns could be addressed through the use of second-generation biofuels that involve

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the use of non-food biomass like cereal straw, bagasse, forest residues and other lignocellulosic materials. This would also serve as an attractive alternative for disposal of non-edible portions of plants. However, compared with the production of ethanol from food crops, the use of lignocellulosic biomass is more complicated because the polysaccharides are more stable and the pentose sugars are not readily fermentable by Saccharomyces cerevisiae. Several biotechnology-based approaches are being used to overcome such problems, including the development of microbial strains, use of alternative yeast species that naturally ferment pentose sugars and the engineering of enzymes that are able to break down cellulose and hemicellulose into simple sugars. Many fungal species are reported to produce various isoforms of β-glucosidases. Thus, it is of utmost importance to screen the best yielding isoform for a particular species. In addition, the thermal stability of β-glucosidases could be enhanced by recombinant DNA technology and genetic modification of microbial strains. Precise genome editing using site-specific nucleases like CRISPR/Cas9 is a suitable option to achieve this goal. The other major hurdles in the commercial β-glucosidase production are product inhibition, low product yields and high cost of enzyme production. The search for better alternatives to the currently available enzyme preparations should be highly promoted. Isolation of novel fungal species having higher β-glucosidase activity would contribute towards revolutionising the field of lignocellulose-mediated production of biofuels. To conclude, the lignocellulosic biomass holds a large potential to meet the energy needs of the world without compromising food security. Acknowledgements  The authors would like to thank Director, DEI, for his continuous support and encouragement. SM is grateful to Dayalbagh Educational Institute, Deemed University, Agra, for sanctioning the Research Project, DEI/Minor Project/2017-18 (iv), as a start-up grant.

References Abdel-Naby MA, Osman MY, Abdel-Fattah AF (1999) Purification and properties of three cellobiases from Aspergillus niger A20. Appl Biochem Biotechnol 76(1):33–44 Ahmed A, Fu HN, Batool K, Bibi A (2017) Microbial β-glucosidase: sources, production and applications. J Appl Environ Microbiol 5(1):31–46 Aït N, Creuzet N, Cattaneo J (1982) Properties of β-glucosidase purified from Clostridium thermocellum. Microbiology 128(3):569–577 An DS, Cui CH, Lee HG, Wang L, Kim SC, Lee ST, Jin F, Yu H, Chin YW, Lee HK, Im WT (2010) Identification and characterization of a novel Terrabacterginsenosidimutans sp. nov. β-glucosidase that transforms ginsenoside Rb1 into the rare gypenosides XVII and LXXV. J Appl Environ Microbiol 76(17):5827–5836 Andrić P, Meyer AS, Jensen PA, Dam-Johansen K (2010) Reactor design for minimizing product inhibition during enzymatic lignocellulose hydrolysis: II. Quantification of inhibition and suitability of membrane reactors. Biotechnol Adv 28(3):407–425 Ávila M, Hidalgo M, Sánchez-Moreno C, Pelaez C, Requena T, de Pascual-Teresa S (2009) Bioconversion of anthocyanin glycosides by Bifidobacteria and Lactobacillus. Food Res Int 42(10):1453–1461 Bae J, Morisaka H, Kuroda K, Ueda M (2013) Cellulosome complexes: natural biocatalysts as arming microcompartments of enzymes. J Mol Microbiol Biotechnol 23(4–5):370–378

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Yadav AN, Mishra S, Singh S, Gupta A (2019b) Recent advancement in white biotechnology through fungi. Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham Yan Q, Zhou XW, Zhou W, Li XW, Feng MQ, Zhou P (2008) Purification and properties of a novel beta-glucosidase, hydrolyzing ginsenoside Rb1 to CK, from PaecilomycesBainier. J Microbiol Biotechnol 18(6):1081–1089 Yan FY, Xia W, Zhang XX, Chen S, Nie XZ, Qian LC (2016) Characterization of β-glucosidase from Aspergillus terreus and its application in the hydrolysis of soybean isoflavones. J Zhejiang Univ Sci B 17(6):455–464 Yang L, Ning ZS, Shi CZ, Chang ZY, Huan LY (2004) Purification and characterization of an isoflavone-conjugates-hydrolyzing β-glucosidase from endophytic bacterium. J  Agric Food Chem 52(7):1940–1944 Yang S, Wang L, Yan Q, Jiang Z, Li L (2009) Hydrolysis of soybean isoflavone glycosides by a thermostable β-glucosidase from Paecilomycesthermophila. Food Chem 115(4):1247–1252 Yoon JJ, Kim KY, Cha CJ (2008) Purification and characterization of thermostable β-glucosidase from the brown-rot basidiomycete Fomitopsis palustris grown on microcrystalline cellulose. J Microbiol 46(1):51–55 You HJ, Ahn HJ, Kim JY, Wu QQ, Ji GE (2015) High expression of β-glucosidase in Bifidobacterium bifidum BGN4 and application in conversion of isoflavone glucosides during fermentation of soy milk. J Microbiol Biotechnol 25(4):469–478 Yun SI, Jeong CS, Chung DK, Choi HS (2001) Purification and some properties of a β-glucosidase from Trichoderma harzianum type C-4. Biosci Biotechnol Biochem 65(9):2028–2032

Chapter 11

Role of Fungi in Climate Change Abatement Through Carbon Sequestration Sandeep K. Malyan, Amit Kumar, Shahar Baram, Jagdeesh Kumar, Swati Singh, Smita S. Kumar, and Ajar Nath Yadav

11.1  Introduction The United Nations Environment Programme and the World Metrological Organization established the Intergovernmental Panel on Climate Change (IPCC) in 1988. The aim of setting up IPCC was to evaluate technical, scientific, and socioeconomic information related to climate change, the potential impact of climate change, and its possible mitigation measures. IPCC published a report in the year 2014 which made it clear that climate change is not a myth. To deal with climate change is a challenge for the worldwide scientific, political, and economic community. Greenhouse gases (GHGs), viz., carbon dioxide (76%), methane (16%), nitrous S. K. Malyan (*) · S. Baram Institute of Soil, Water and Environmental Science, The Volcani Research Center, Agricultural Research Organization (ARO), Rishon LeZion, Israel e-mail: [email protected]; [email protected] A. Kumar Host Plant Section, Central Muga Eri Research and Training Institute, Central Silk Board, Lahdoigarh, Jorhat, Assam, India J. Kumar Department of Hydrology, Indian Institute of Technology Roorkee, Roorkee, Uttarakhand, India S. Singh Department of Environmental Science, Chaudhary Charan Singh University, Meerut, Uttar Pradesh, India S. S. Kumar Center for Rural Development and Technology, Indian Institute of Technology Delhi, New Delhi, India A. N. Yadav Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, Sirmour, Himachal Pradesh, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_11

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Fig. 11.1  Trends of CO2 concentration in the atmosphere at Mauna Loa. (Source: NOAA 2019)

oxide (6%), and chlorofluorocarbons (2%), are likely to affect agricultural productivity and food security adversely through climate change (Ranjan and Yadav 2019; Malyan 2018; Fagodiya et al. 2017a; Kumar et al. 2016, 2017b; Pathak et al., 2016; Gupta et al. 2015, 2016; Kumar and Malyan 2016; IPCC 2014; Bhatia et al. 2013a, b). Malyan et al. (2016a, b) quoted that the rise in atmospheric greenhouses gases may result in rising of global mean temperature up to 1.5  °C by the end of the twenty-first century. Among all greenhouse gases, CO2 only accounts for 76%, so it is considered most important. At the beginning of industrial revolution, atmospheric CO2 concentration was 280 ppm (Rastogi et al. 2002), and it has now increased to 410 ppm in March 2019 (Fig. 11.1). The annual mean growth rate (differences in CO2 concentration between the last month of the year [December] and the first month [January] of that year) was monitored by Global Monitoring Division of National Oceanic and Atmospheric Administration (NOAA) at Mauna Loa site. The rate of growth was 0.54 ppm/yr. in 1960 and it increased to 2.10 ppm/yr. in 1983 (Fig. 11.2). The growth rate of CO2 in the atmosphere was 2.98 ppm/yr. in the year 2016, and it might rise over 3 ppm/ yr. in few upcoming years (NOOA 2019). The CO2 growth rate is increasing continuously due to many anthropogenic actions, including fossil fuel combustion, deforestation, forest fire, automobile, etc. Soil, plant, and ocean are the major natural sink for atmospheric CO2. Recently, the scientific community enhanced the work on mitigating climate change and global warming by reducing atmospheric CO2 concentration through carbon sequestration (Bhattacharyya et al. 2018; Mukherjee et al. 2018). Soil acts as both source (soil respiration) and sink (carbon sequestration) for CO2, and soil respiration contributes to 20% of the total CO2 emission to the atmosphere (Kumar et al. 2017a, b; Gupta et al. 2016; Rastogi et al. 2002). The global soils hold 3.3 times (2500 Gt C) as compared to carbon present in the atmosphere (Lal 2014). Out of 2500 Gt, soil

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Fig. 11.2  Growth rate of carbon dioxide at Mauna Loa, Hawaii. (Source: NOAA 2019)

organic matter (SOC) and soil inorganic carbon retain 1550 Gt and 950 Gt, respectively (Lal 2014). The world soil pool of C is 4.5 times the size of C present in biomass (Lal 2014). In the soil carbon sequestration, biological soil crust plays a significant role. Soil fungi especially arbuscular mycorrhizal fungi enhance the soil carbon sequestration in different types of ecosystems. The objective of this study is to assess the carbon sequestration potential of fungi in soils and the factors affecting thereof.

11.2  Fungi and Carbon Sequestration Carbon is the major building block in all living organisms. It exists in many forms such as soil organic matter, plant biomass, and CO2 in the dissolved form in water and gas in the atmosphere. Carbon in soil is more than the total carbon in the atmosphere and plants biomass. Numerous small fungi in the soil, consist of hyphae that use carbon as a building block (Fig. 11.3). When this hypha dies, it is easily decomposed and its carbon is stored as soil organic matter for a long time (Treseder and Holden 2013). The process of long-term storage of carbon in the soil, terrestrial biomass, or the ocean so that the buildup of CO2 in the atmosphere will be slowed down or reduced is known as carbon sequestration. In other words, carbon sequestration is defined as “the capture and secure storage of carbon that would otherwise be emitted to or remains in the atmosphere” (FAO 2000). In carbon sequestration, fungi play a significant role in the northern hemisphere of the Earth which help to combat global warming. In soil, fungi do symbioses association (mycorrhizal fungi) with plant roots and help the plants to utilize the nutrients from the soil. As a result, mycorrhizal fungi stimulate the plant’s growth which results in the faster removal of atmospheric CO2 through its conversion  into plant biomass. There is a different pathway of converting the atmospheric CO2 to plant biomass (Fellbaum et al. 2012).

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Fig. 11.3  Showing mycorrhizal fungi hyphae in soil Fig. 11.4 Simplified method of carbon sequestration in soil organic matter by fungi

Carbon di-oxide

Biological soil crust

Plants

Fungi

Carbon sequestration in organic matter of soil The photosynthates of the host plant are transferred to the intraradical hyphae followed by extraradical hyphae before it releases to the soil (Fig. 11.4). Carbon sequestration in soil depends on the volume and hyphal biomass of fungi (Solaiman 2014; Zhu and Miller 2003). Fungus is a nonchlorophyllous organism and is heterotrophic (requires an organic source of carbon) in nature, i.e., it obtains its food from either dead organic matter (saprophytic) (Chaubey et al. 2019) or from the autotrophic/heterotrophic associates

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(parasitic and symbiotic). Agaricus, Aspergillus, Morchella, Mucor, Penicillium, Rhizopus, Saprolegnia, etc. are the example of the saprophytic mode of nutrition. The parasitic fungi are Peronospora, Puccinia, Fusarium, Pythium, Melampsora, etc. The associate may be animal, plants, and microbes. If an associate is a plant, the autotrophic association can be stated as true symbiosis. The fungi can use a variety (e.g., yeast used the acetate) of the organic matter as a source of energy but the majority of them prefer carbohydrates. Glucose is a preferred carbohydrate for almost all the fungi as compared to fructose. Starch and cellulose are also used by some of the fungi which are capable of synthesizing hydrolytic enzymes. Organic acids are the least preferred sources of energy by most of the fungi. Basidiomycetes are able to utilize lignin as an energy source. Saprolegniaceae and Blastocladiales can grow only with organic nitrogen, i.e., amino acids. The organic matter is first dissolved by the extracellular enzymes, and then it is directly absorbed by diffusion either through the hyphal walls of the hyphae that penetrate the substratum or by the rhizoidal hyphae. Lichens (a single composite thallus plant) are the best-explained association of fungi with the alga where the fungus provides the minerals, water from the substratum, and space for alga which in return supplies food. Mycorrhiza is the association of fungi with the roots of higher plants and can be ectomycorrhiza, endomycorrhiza, and ectoendomycorrhiza based on the positional association. The input of the plant’s organic matter into the soil carbon is both aboveground and belowground litter in the form of leaves, stem, flower, and roots. The plant also contributes some organic compounded inform of exudates to the soil. This contribution is known as rhizo-deposition. The rhizo-deposition is also an important carbon contribution of higher plants which is based on plant root architecture, environment, physiology, biochemistry, and chemical composition of the deposited matter. The litter’s input is varying in quantity and depends on the climax community of the ecosystem. The belowground input mainly contributes to the organic matter and stabilizes their medium to long duration based on the physicochemical properties of soil and microbiota (Dignac et al. 2017; Clemmensen et al. 2015; Kuzyakov and Domanski 2000; Mendez-Millan et al. 2010). Carbon sequestration entirely depends on the photosynthetic rate of the autotrophs and the respiratory losses of the autotrophs and symbionts (mycorrhizal fungi). These fungi utilize 5 to 20% of the net primary productivity of the symbiotic system (Luo and Zhou 2010). The change in the carbon assimilating or degradation rate can entirely change the carbon sequestration potential of the association until the change in both the processes is in the same direction and potential. The degradation of rhizo-deposited carbon is done by microorganisms. These microbes are present or associated with the specific plants and soils. The quantity of the rhizo-associate is also contributed majorly. The quantified contribution in the respiratory losses of these rhizo-associates especially mycorrhizal fungi is not reported in different associations separately. Thus, the exact carbon sequential potential of mycorrhizal fungi still remains to be estimated. However, few studies have confirmed in controlled and field experiments that carbon sequestration depends on plant photosynthesis and respiration through mycorrhizal fungi (Luo and Zhou 2010; Bahn et  al. 2008;

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Friedlingstein, et  al. 2006; Ussiri et  al. 2006). Global forest soils efflux into the atmosphere through belowground respiration has been estimated to be approximately 24 Pg C y−1 in which mycorrhizal fungi contribute about 16% (Zhang et al. 2007; Hu et  al. 2005). Thus, increasing the sequestration potential of the mycorrhizal fungi can reduce the contribution drastically.

11.3  Factors Affecting Fungal Growth in Soil Fungi have high plasticity and they easily adapt to adverse conditions in the soil. A diverse range of fungi is found in almost every type of ecosystem owing to the fact that fungi can adapt to a wide range of temperature and pH (Frac et al. 2018; Frac et al. 2015). Fungi activity in soil is affected by abiotic (physical disturbance, soil temperature, soil pH, soil texture, moisture, and salinity) and biotic (plant and other organisms interaction) factors, and it may influence the carbon sequestration activity of fungi (Kour et al. 2019; Rana et al. 2019; Yadav et al. 2017, 2018, 2019; Farc et al. 2018; Rouphael et al. 2015; Posada et al. 2008; Treseder and Allen 2000) (Table 11.1). Some factors affecting fungi activity in the soil are discussed below in brief.

11.3.1  Temperature The Northern hemisphere consists of 67.3% of total Earth’s landmass; Moritz et al. (2002) reported that global warming has led to a rise in temperature by ~1.5 °C. The IPCC (2007) predicted that by the end of 2100, the temperature may elevate to Table 11.1  Factor affecting the fungi in soils Factor Temperature

P-Fertilizer

Pesticides Physical disturbances

Remarks Soil warming results in rapid fungal respiration which results in carbon losses to the atmosphere No effect on fungal carbon sequestration under soil warming On addition of N fertilizer fungal biomass increase which helps in carbon sequestration To an optimum level (20 mg kg−1) shows a positive response while at a higher dose (40 mg kg−1) shows a negative impact on fungal root growth Application of phosphorous fertilizer has a negative effect on fungal diversity in soil Have a direct and indirect negative impact on fungi diversity and carbon sequestration in soil Biocrust removal drastically affects inoculum potential of AM (Arbuscular Mycorrhiza) fungal mycelial due to web fragmentation

References Hawkes et al. (2008) Rosenstock et al. (2018) Aliasgharzad et al. (2018) Aliasgharzad et al. (2018) Smith et al. (2011) Willis et al. (2013) Jasper et al. (1989)

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additional 4–7 °C which will affect all the ecosystem services. There are contradictory reports on the effect of soil temperature on the fungal diversity and activity (Rosenstock et al. 2018; Solley et al. 2017; Allison and Tresseder 2008; Fujimura et al. 2008; Zogg et al. 1997; Yadav et al. 2018). On one hand, some studies have reported significant changes in fungal activity; to the contrary, some others could not assess any change related to temperature. This suggests that the effect of temperature is species-specific. In Tundra, Fujimura et al. (2008) observed no change in fungal diversity even after soil warming. Warming of soil samples taken from the environment resulted in a significant change in fungal community of soil (Zogg et al. 1997). Rosenstock et al. (2018) observed that the soil warming from a range of 0–5.5 °C above the control has no or limited effect on the growth of ectomycorrhizal and no effect on community composition and fungal carbon sequestration in Picea sitchensis forest soils (Rosenstock et al. 2018).

11.3.2  Physical Disturbance Disturbance in several ecosystems is a universal process and it affects all levels of biological organisms prevailing in that ecosystem. The causes of disturbance may be natural or anthropogenic. Anthropogenic physical disturbance in top soil (20– 30 cm) results in fragmentation of fungal mycelia (Jasper et al. 1989) which results in a lower rate of soil carbon sequestration. Korb et al. (2003) reported that in the soil, fungi diversity recovered rapidly after a forest fire. On the other hand, in an agricultural field, tillage practices disturb the fungal diversity and they recover at a slow rate. However, in the Gangetic Plains of India, the fungal diversity was observed to recover at a rapid rate. The high rate of fungal recovery in Indian plains was due to a large diversity of spores (Oehl et al. 2005).

11.3.3  Soil pH Soil pH is the primary factor affecting fungal activity in the soil and provides an environment that is essential for carbon sequestration in soil. Minor increases in pH are associated with higher root colonization by fungi in acidic soils with less phosphorus availability (Ge et al. 2017; Oehl et al. 2005).

11.3.4  Fertilizer Phosphorus(P)-based fertilizers have environmental concerns related to eutrophication of fresh waterbodies. Generally, soil has an abundant amount of both inorganic and organic P in soil, but its bioavailability is very low for the crop and plants

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(Sarabia et al. 2017). Mycorrhiza increased the P availability for host plant, and in return, fungi get the photosynthetic product and result in more development of hyphae which increases the soil carbon sequestration. Aliasgharzad et  al. (2018) reported that application of 20 mg-P kg−1 soil showed positive fungal root growth while increasing the P dose to 40  mg  kg−1negatively affected the growth rate of fungi (Table 11.1).

11.3.5  Pesticide Fungi in agricultural soils are exposed to different type of pesticides. Application of pesticide directly or indirectly affects the host plant and thus indirectly suppresses the rate of carbon sequestration by fungi in soils (Table 11.1). Zocco et al. (2011) reported that fungicides like fenpropimorph inhibit the growth and development of fungi in soil. Nevertheless, pesticides like dimethoate, fenamiphos, and aldicarb were not found to inhibit the fungal diversity (Karpouzas et al. 2014; Schweiger and Jakobsen 1998; Nemec 1985). Spokes et al. (1981) studied the impact of eight chemicals on the development of fungi. It was observed that aldicarb had negligible impact on fungi development, while fungicide chloroneb acted as development stimulator for fungal population in the soil. Then again, triadimefon and benomyl fungicides application had an inhibitory impact on fungal development (Spokes et al. 1981).

11.3.6  Moisture Soil moisture is an important constituent for the growth of the soil fungi. The soil fungi and degradation of the rhizo-deposition material is highly influenced through the moisture percentage. The decrease in the moisture percentage can directly limit the availability of organic matter to the fungi, so that the rate of degradation can decrease and ultimately the growth of soil fungi can also get limited. Therefore, moisture also directly controls the soil temperature and soil heat flux. Moreover, the higher the soil temperatures, the lower is the soil moisture, especially in the topsoil horizon, and ultimately the lower will be the soil fungi due to the scarcity of the food material.

11.4  Research Gap and Future Recommendation At certain places and in some of the ecosystems, fungi can play an important role in the sequestration of carbon dioxide present in the atmosphere by playing principal roles in fixation of organic matter in the soil and degradation. However, the mechanism

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of sequestration is not yet fully understood. There are certain avenues that need to be explored in order to have a deeper understanding of this phenomenon. Some of the gaps that have been observed from literature to direct the future research have been summarized as follows: (i) The quantified contribution of soil and mycorrhizal fungi in different ecosystems and various associations has not yet been evaluated and reported. This quantification can serve a major role to define the direction of the soil and mycorrhizal fungi as a carbon sequestrate. (ii) The minimizing of the respiratory losses of soil and mycorrhizal fungi can also enhance the carbon sequestration potential. Therefore, future research should be conducted in this direction. (iii) The role and potential of different fungal communities along with different types of soil and mycorrhizal fungi can also serve to enhance the carbon sequestration in different ecosystems. (iv) The factors influencing growth of fungi such as soil moisture, soil temperature, CO2 enrichment, quality of rhizo-deposition, and precipitation changes control the heterotrophic respiration. Thus, the impact of these factors with respect to the soil and mycorrhizal fungi can also lend a hand to understand their behavior in changing the environment. (v) Relevant studies in different interactions, i.e., moisture vs mycorrhizal diversity, soil temperature vs mycorrhizal composition and diversity, and soil temperature vs moisture, are very limited in respect of the soil fungal respiration and also poorly understood. (vi) Contribution of mycorrhizal association on carbon storage in contaminated/ mineral/nutrient-rich soil is not available. Therefore, future research in this direction can help in improved understanding of the mechanism of fungal carbon sequestration.

11.5  Conclusions Based on this study, the following conclusions can be drawn. The global soil carbon is more than the carbon present in the atmosphere, but the rising temperature due to global warming has the potential to affect the fungal diversity and fungal activity. The rate of soil carbon sequestration by fungi depend upon direct (quality and quantity of hyphae) and indirect (soil pH, texture, moisture, soil management, physical disturbance, pesticide application, etc.) factors of the ecosystem. There are three important strategies to combat climate change: development of less or zero carbon fuel, limiting of fossil energy use, and carbon dioxide sequestration from atmosphere or emission point source through abiotic (engineered) or biotic methods (photosynthesis). Soil carbon sequestration by fungi in soil is the environmentally sound method to reduce the carbon dioxide levels in the atmosphere.

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Acknowledgments  The financial support to the first author, Sandeep Kumar Malyan, provided by Ministry of Agriculture and Rural Development, Israel, under ARO-Postdoctoral ­Fellowship-­India and China, is highly acknowledged. The authors are also very thankful to the Central MugaEri Research and Training Institute, Central Silk Board, Lahdoigarh, Jorhat-785700, India, for having provided the necessary support.

References Aliasgharzad N, Afshari Z, Najafi N (2018) Carbon sequestration by glomerular fungi in soil is influenced by phosphorus and nitrogen fertilization. Int J Adv Sci Eng Inf Technol 6:1–5 Allison SD, Tresseder KK (2008) Warming and drying suppress microbial activity and carbon cycling in boreal forest soils. Glob Chang Biol 14:2898–2909 Bahn M, Rodeghiero M, Anderson-Dunn M, Dore S, Gimeno C, Drösler M, Williams M, Ammann C, Berninger F, Flechard C, Jones S (2008) Soil respiration in European grasslands in relation to climate and assimilate supply. Ecosystems 11:1352–1367 Bhatia A, Kumar A, Kumar V, Jain N (2013a) Low carbon option for sustainable agriculture. Indian Farm 63:18–22 Bhatia A, Kumar A, Das TK, Singh J, Jain N, Pathak H (2013b) Methane and nitrous oxide emissions from soils under direct seeded rice. Int J Agric Stat Sci 9(2):729–736 Bhattacharyya R, Bhatia A, Das TK, Lata S, Kumar A, Tomer R, Singh G, Kumar S, Biswas AK (2018) Aggregate-associated N and global warming potential of conservation agriculture-based cropping of maize-wheat system in the north-western Indo-Gangetic Plains. Soil Tillage Res 182:66–77 Clemmensen KE, Finlay RD, Dahlberg A, Stenlid J, Wardle DA, Lindahl BD (2015) Carbon sequestration is related to mycorrhizal fungal community shifts during long-term succession in boreal forests. New Phytol 205:1525–1536 Chaubey R, Singh J, Baig MM, Kumar A (2019) Recent advancement and the way forward for Cordyceps. In: Yadav A, Singh S, Mishra S, Gupta A (eds) Recent advancement in white biotechnology through fungi, Fungal biology. Springer, Cham Dignac M-F, Derrien D, Barré P, Barot S, Cécillon L, Chenu C, Chevallier T, Freschet GT, Garnier P, Guenet B, Hedde M, Klumpp K, Lashermes G, Maron P-A, Nunan N, Roumet C, Basile-­ Doelsch I (2017) Increasing soil carbon storage: mechanisms, effects of agricultural practices and proxies. A review. Agron Sustain Dev 37:14 Fagodiya RK, Pathak H, Bhatia A, Kumar A, Singh SD, Jain N (2017a) Simulation of Maize (Zea Mays L.) yield under alternative nitrogen fertilization using infocrop-maize model. Biochem Cell Arch 17:65–71 Fagodiya RK, Pathak H, Kumar A, Bhatia A, Jain N (2017b) Global temperature change potential of nitrogen use in agriculture: a 50-year assessment. Sci Rep 7:44928 FAO (2000) Carbon sequestration options under the clean development mechanism to address land degradation (World Soil Resources Reports), by Food and Agricultural Organization of the United Nations. ISBN-10:9251045151 Fellbaum CR, Mensah JA, Pfeffer PE, Kiers ET, Bucking H (2012) The role of carbon in fungal nutrient uptake and transport: implications for resource exchange in the arbuscular mycorrhizal symbiosis. Plant Signal Behav 7:1509–1512 Frąc M, Jezierska-Tys S, Takashi Y (2015) Occurrence, detection, and molecular and metabolic characterization of heat-resistant fungi in soils and plants and their risk to human health. Adv Agron 132:161–204 Frac M, Hannula SE, Jedryczka M (2018) Fungal biodiversity and their role in soil health. Front Microbiol 9:707 Friedlingstein P, Cox P, Betts R, Bopp L, von Bloh W, Brovkin V, Cadule P, Doney S, Eby M, Fung I, Bala G (2006) Climate-carbon cycle feedback analysis: results from the C4MIP model intercomparison. J Clim 19:3337–3353

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Fujimura KE, Egger KN, Henry GH (2008) The effect of experimental warming on the root-­ associated fungal community of salixarctica. ISME J 2:105 Ge ZW, Brenneman T, Bonito G, Smith ME (2017) Soil pH and mineral nutrients strongly influence truffles and other ectomycorrhizal fungi associated with commercial pecans (Caryaillinoinensis). Plant Soil 418:493–505 Gupta DK, Bhatia A, Kumar A, Chakrabarti B, Jain N, Pathak H (2015) Global warming potential of rice (Oryza sativa)-wheat (Triticumaestivum) cropping system of the Indo-Gangetic Plains. Indian J Agric Sci 85:807–816 Gupta DK, Bhatia A, Kumar A, Das TK, Jain N, Tomer R, Malyan SK, Fagodiya RK, Dubey R, Pathak H (2016) Mitigation of greenhouse gas emission from the rice-wheat system of the Indo-Gangetic Plains: through tillage, irrigation and fertilizer management. Agric Ecosyst Environ 230:1–9 Hawkes CV, Hartley IP, Ineson P, Fitter AH (2008) Soil temperature affects carbon allocation within arbuscular mycorrhizal networks and carbon transport from plant to fungus. Glob Chang Biol 14:1181–1190 Hu ZQ, Wei ZY, Qin P (2005) Concept and methods for soil reconstruction in mined land reclamation. Soil 37:8–12 IPCC (2007) Climate change report. Cambridge University Press, Cambridge, p 73 IPCC (2014) Fifth assessment report on climate change. Cambridge University Press, Cambridge Jasper DA, Abbott LK, Robson AD (1989) Soil disturbance reduces the infectivity of external hyphae of arbuscular mycorrhizal fungi. New Phytol 112:93–99 Karpouzas DG, Papadopoulou E, Ipsilantis I, Friedel I, Petric N, Udikovic-Kolic N, Kandeler E, Menkissoglu-Spiroudi U, Martin-Laurent F (2014) Effects of nicosulfuron on the abundance and diversity of arbuscular mycorrhizal fungi used as indicators of pesticide soil microbial toxicity. Ecol Indic 39:44–53 Korb JE, Johnson NC, Covington WW (2003) Arbuscular mycorrhizal propagule densities respond rapidly to ponderosa pine restoration treatments. J Appl Ecol 40:101–110 Kour D, Rana KL, Yadav N, Yadav AN, Singh J, Rastegari AA, Saxena AK (2019) Agriculturally and industrially important fungi: current developments and potential biotechnological applications. In: Yadav AN, Singh S, Mishra S, Gupta A (eds) Recent advancement in white biotechnology through fungi, volume 2: perspective for value-added products and environments. Springer International Publishing, Cham, pp 1–64. https://doi.org/10.1007/978-3-030-14846-1_1 Kumar SS, Malyan SK (2016) Nitrification inhibitors: a perspective tool to mitigate greenhouse gas emission from rice soils. Curr World Environ 11:423–428 Kumar A, Tomer R, Bhatia A, Jain N, Pathak H (2016) Greenhouse gas mitigation in Indian agriculture. In: Pathak H, Chakrabarti B (eds) Climate Change and Agriculture Technologies for Enhancing Resilience. ICAR-IARI, New Delhi, pp 137–149 Kumar A, Bhatia A, Fagodiya RK, Malyan SK, Meena BL (2017a) Eddy covariance flux tower: a promising technique for greenhouse gases measurement. Adv Plants Agric Res 7(4):337–340. https://doi.org/10.15406/apar.2017.07.00263 Kumar SS, Kadier A, Malyan SK, Ahmad A, N Bishnoi NR (2017b) Phytoremediation and rhizoremediation: uptake, mobilization and sequestration of heavy metals by plants. In: Singh D, Singh H, Prabha R (eds) Plant-microbe interactions in agro-ecological perspectives. Springer, Singapore, pp 367–394. https://doi.org/10.1007/978-981-10-6593-4_15 Kuzyakov Y, Domanski G (2000) Carbon input by plants into the soil. Review. J Plant Nutr Soil Sci 163:421–431 Lal R (2014) Soil carbon sequestration impacts on global climate change and food security. Science 304:1623–1627 Luo YQ, Zhou XH (2010) Soil respiration, and the environment. Academic Press, San Diego Malyan SK (2018) Reducing methane emission from rice soil through microbial interventions. Ph.D. Thesis, ICAR-Indian Agricultural Research Institute, New Delhi-110012 Malyan SK, Bhatia A, Kumar A, Gupta DK, Singh R, Kumar SS, Tomer R, Kumar O, Jain N (2016a) Methane production, oxidation, and mitigation: a mechanistic understanding and comprehensive evaluation of influencing factors. Sci Total Environ 572:874–896

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Malyan SK, Kumar SS, Kumar A, Kumar J  (2016b) Water management tool in rice to combat two major environmental issues: global warming and water scarcity. In: Kumar S, Beg MA (eds) Environmental concerns of 21st century: Indian and global context, pp 43–58. (ISBN: 978-93-83281-65-7) Mendez-Millan M, Dignac M-F, Rumpel C, Rasse DP, Derenne S (2010) Molecular dynamics of shoot vs. root biomarkers in an agricultural soil estimated by natural abundance 13Clabeling. Soil Biol Biochem 42:169–177 Moritz RE, Bitz CM, Steig EJ (2002) Dynamics of recent climate change in the Arctic. Science 297:1497–1502 Mukherjee J, Mridha N, Mondal S, Chakraborty D, Kumar A (2018) Identifying suitable soil health indicators under variable climate scenarios: a ready reckoner for soil management. In: Bal S, Mukherjee J, Choudhury B, Dhawan A (eds) Advances in crop environment interaction. Springer, Singapore Nemec S (1985) Influence of selected pesticides onGlomus species and their infection in citrus. Plant Soil 84(1):133–137 NOAA (2019) Earth System Research Laboratory Global Monitoring Division, Online link (https://www.esrl.noaa.gov/gmd/ccgg/trends/full.html) Oehl F, Sieverding E, Ineichen K, Ris EA, Boller T, Wiemken A (2005) Community structure of arbuscular mycorrhizal fungi at different soil depths in extensively and intensively managed agroecosystems. New Phytol 165:273–283 Pathak H, Jain N, Bhatia A, Kumar A, Chatterjee D (2016) Improved nitrogen management: a key to climate change adaptation and mitigation. Indian J Fertil 12(11):151–162 Posada RH, Franco LA, Ramos C, Plaza LS, Sua JC, Lvarez FA (2008) Effect of physical, chemical and environmental characteristics on arbuscular mycorrhizal fungi in Brachiaria decumbens (Stapf) pastures. J Appl Microbiol 104:132–140 Rana KL, Kour D, Sheikh I, Yadav N, Yadav AN, Kumar V, Singh BP, Dhaliwal HS, Saxena AK (2019) Biodiversity of endophytic fungi from diverse niches and their biotechnological applications. In: Singh BP (ed) Advances in endophytic fungal research: present status and future challenges. Springer International Publishing, Cham, pp  105–144. https://doi. org/10.1007/978-3-030-03589-1_6 Ranjan R, Yadav R (2019) Targeting nitrogen use efficiency for sustained production of cereal crops. J Plant Nutr. https://doi.org/10.1080/01904167.2019.1589497 Rastogi M, Singh S, Pathak H (2002) Emission of carbon dioxide from the soil. Curr Sci 82: 510–517 Rosenstock N, Ellstrom M, Oddsdottir E, Singurdsson BD, Wallander H (2018) Carbon sequestration and community composition of ectomycorrhizal fungi across a geothermal warming gradient in an Icelandic spruce forest. Fungal Ecol. https://doi.org/10.1016/j. funeco.2018.05.010 Rouphael Y, Franken P, Schneider C, Schwarz D, Giovannetti M, Agnolucci M (2015) Arbuscular mycorrhizal fungi act as biostimulants in horticultural crops. Sci Hortic 196:91–108 Sarabia M, Cornejo P, Azconc R, Carreon-Adudd Y, Larsen J (2017) Mineral phosphorus fertilization modulates interactions between maize, rhizosphere yeasts, and arbuscular mycorrhizal fungi. Rhizosphere 4:89–93 Schweiger PF, Jakobsen I (1998) Dose-response relationships between four pesticides and phosphorus uptake by hyphae of arbuscular mycorrhizas. Soil Biol Biochem 30:1415–1422 Smith SE, Jakobsen I, Gronlund M, Smith FA (2011) Roles of arbuscular mycorrhizas in plant phosphorus nutrition: interactions between pathways of phosphorus uptake in arbuscular mycorrhizal roots have important implications for understanding and manipulating plant phosphorus acquisition. Plant Physiol 156:1050–1057 Solaiman ZM (2014) Mycorrhizal fungi: use in sustainable agriculture and land restoration. Soil Biol 41. https://doi.org/10.1007/978-3-662-45370-4_18 Solley EF, Lindahal BD, Dawes MA, Peter M, Souza RC, Rixen C, Hagedorn F (2017) Experimental soil warming shifts the fungal community composition at the alpine tree line. New Phytol 215:766–778

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Chapter 12

Microbial Enzymes and Their Application in Pulp and Paper Industry Abdulhadi Yakubu, Upasana Saikia, and Ashish Vyas

12.1  Introduction Industrial utilization of wastepapers in the production of new one is increasing globally. Currently, pulp and paper industry is one of the largest consumer of wood. Based on their demands due to global economic growth, more trees will be harvested and waste will be consumed and disposed in the environment (Pathak et al. 2011). Chemical agents such as sodium hydroxide, hydrogen peroxide, sodium carbonate, diethylenetriaminepentaacetic acid, sodium silicate, and surfactants are used in large quantities by paper industries as conventional methods of deinking wastepaper which lead to expensive wastewater treatment to meet environmental regulations (Pathak et  al. 2011; Saxena and Chauhan 2016). Enzymes such as lipase, xylanase, pectinase, cellulase, hemicellulase, amylase, and esterase are used as substitute to chemical conventional methods of deinking wastepapers (Yadav et  al. 2015, 2016). These enzymes are reported to be environmentally friendly as compared to conventional method. It was realized several decades ago that microbial enzymes might be useful in processing of papers since it is composed of natural polymers such as cellulose, hemicellulose, and lignin. Microbial enzymes have been commercially used in pulp and paper industry only in the previous decade, while microorganisms are presently used in other industrial processing steps, though long been used in the treatment of wastewater. This is due to the fact that wood and pulps which act as substrates are difficult to degrade. In addition, most research now focuses on lignin biodegradation since it is lignin that is A. Yakubu Department of Science Laboratory Technology, College of Science and Technology, Jigawa State Polytechnic, Dutse, Nigeria U. Saikia · A. Vyas (*) Department of Microbiology, School of Bioscience and Bioengineering, Lovely Professional University, Phagwara, Punjab, India e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_12

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removed from wood in chemical pulping and pulp in bleaching. Lignin likely evolved in part as a deterrent to microbial degradation, and it continues to be an impediment to biotechnological processing of wood and pulps. During the last decade, the number of possible applications of enzymes in paper and pulp industries in which many are of commercial quantity has grown rapidly. These include xylanase for enzymatic bleaching, lipase for pitch removal, as well as cellulase and hemicellulase for freshness enhancement (Kirk et al. 1996). Globally, our lives are somewhere governed by little inscriptions laid on paper in one way or the other. The major domains of public sector are reliant on paper and paper products directly or indirectly. The history of paper dates back almost 2000  years, while the manufacturing process happened to be reported in China when inventors imprint their writings on crafted cloth sheets. The knowledge of papermaking then became popularized through westward and eventually reached India around 605  AD.  A wide variety of raw materials used in the papermaking process include cellulose originated from agriculture waste, forests, and wastepaper, noncellulosic coal, talcum powder, etc. However, raw materials like wood logs and its waste, bamboo, wastepaper, bagasse, and agricultural residues like bran of wheat, rice straw, grasses, and seaweed are majorly used in the process. The basic product coming out during paper processing is the cellulosic pulp which is used in papermaking and as animal feedstock. Pulp being a lignocellulosic fibrous material separates the cellulosic fibers from wood logs, fibers, wastepaper, and rags. Two steps are involved in pulp production: wood pulping and pulp bleaching (Fig. 12.1). Both processes require high-energy input and reagents which leads to the production of significant amount of gaseous, liquid, and solid wastes that need to be treated. Biotechnological approaches provide substantial solution of the aforementioned problem. Lignin degradation by white-rot fungi has been intensively studied for biotechnical applications such as biopulping, biobleaching, and treatment of pulp mill effluents. Also, lignolytic enzymes and hemicellulases provide a prospective way to decrease the usage of hazardous bleaching reagents and to make an improvement in the quality of bleached pulp (Tavares et  al. 2014). The application of

Fig. 12.1  The stages of enzymatic deinking process

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­ icrobial enzymes in paper and pulp industries has increased rapidly which lead to m the decrease of adverse effect on ecosystem due to the increased awareness of sustainability issues (Yadav et  al. 2019a, b). Utilization of these microbial enzymes also leads to the decrease in energy consumption, processing time, and amount of chemicals in deinking wastepaper. They are equally used to aid deinking and bleach in paper and pulp industry as well as waste treatment by increasing biological oxygen demand (BOD) and chemical oxygen demand (COD). It is difficult to deink laser-printed office papers by employing the conventional method of deinking. Due to higher demand of laser printers and copy machines every year, there has been an increase in the amount of nonimpact printed papers entering the recycled papers. It is challenging for the inventors to remove ink from these papers. The reason is primarily due to the strong adherence of the toner particles to the paper surface. The photocopier printers are indulged in using thermosetting toners made up of synthetic polymers as ink. Due to applying high amount of heat during inking, the ink particles become physically bonded to the paper making it difficult as well as expensive to remove by conventional method. A biological process using enzymes had been evaluated which implied positive results in deinking process of different types of wastepaper. One of the major advantages of using enzymes is production of minimum treatment effluent, and it is also less harmful to the environment. The most important step involved in recycling process is removal of ink from the paper (Table 12.1). Several enzymes like cellulase, hemicellulase, α-amylase, lipase, xylanase, and other lignolytic enzymes are involved in the biological process of deinking. The enzymatic treatment is favorable because enzymes are eco-friendly in nature and during its processing ink detachment occurs without any discharge of harmful chemicals, thus rendering our environment green. Most of the cited studies reported deinking of mixed office waste consisting of photocopier paper using commercially available enzymes (Roushdy 2015). India generates approximately 36.5 million tons of municipal solid waste annually, of which 14.6 million tons consist of paper wastes. The Indian Agro and Recycled Paper Mills Association (IARPMA) estimated that India is among the countries with low recycling of waste of wastepapers (26%) as compared to countries like China (38%), Thailand (45%), and Germany (80%). Based on the shortage of raw materials, resources, and high demands being imposed on green plants, Indian paper industries are facing many challenges on daily basis. One of the attractive solution to these problems in India is the recycling of municipal office waste (MOW) papers but still very difficult to remove nonimpact ink. Deinking process which involves the removal of printed ink from used paper involves dislodgement of ink particles from fiber surface and the separation of dispersed ink from the fiber suspensions by washing and flotation (Tavares et al. 2014). In chemical deinking process, industries used high quantity of chemicals such as chlorine, chlorine-based derivatives, sodium hydroxide, sodium carbonate, sodium silicate, hydrogen peroxide, hypochlorite, and chelating agents, which lead to hazardous effluent disposal problems (Vega et al. 2012; Pala et  al. 2004). For these reasons, biological deinking by using microbial

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Table 12.1  Microbial enzyme and their applications in pulp and paper industry Microorganism Enzymes Commercial enzymes Cellulase Trichoderma harzianum

Cellulase and xylanase

Commercial enzymes Cellulase Commercial enzymes Cellulase Aspergillus niger Streptomyces sp. L22001 Bacillus altitudinis

Cellulase and hemicellulase Xylanase Xylanase

Bacillus sp. CKBxID

Xylanase

Alkalothermotolerant Aspergillus niger

Xylanase and pectinase Xylanase

Aspergillus nidulans

Xylanase

Commercial enzymes Laccase and hemicellulase Commercial enzymes Cutinase and amylase Commercial enzymes Amylase and cellulase

Application Ink removal, freshness, and reduction of drainage time Improved drainage, high deinking efficiency, brightness, and reduction of drainage time Not good if specks surface of the deink paper are used Detach significant amount of ink from ONP/OMG Enhanced optimum deinking efficiency Biobleaching effect Potential for biodeinking and biobleaching Deinking agent of recycled wastepaper Commercially viable with better paper quality Deink old newspaper with improved brightness, removal of surface ink particles from ONP pulp Reduction of ink and increased brightness of recycled paper Deink old newspaper Increase brightness and ink removal Improve brightness

References Pathak et al. (2011) Pathak et al. (2014) Tsatsis et al. (2017) Zhang et al. (2008) Lee et al. (2007) Li et al. (2010) Adhyaru et al. (2017) Maity et al. (2012) Singh et al. (2012) Desai and Iyer (2016) Taneja et al. (2002) Xu et al. (2011) Wang et al. (2018) Gil et al. (2013)

enzymes which act directly either on the fiber or on the ink film becomes more attractive. For example, hydrolysis of cellulose and hemicellulose brings detachment at fiber/ink interbonding regions and finally releases ink particles into the suspension when treated with cellulase/hemicellulase enzymes (Lee et al. 2007). Microbial enzymes can also detach small fibrils from the surface of the ink particles; hence, they can change the usual hydrophobicity of the particles, which brings about their separation in the flotation/washing step. These enzymatic technologies have been described as especially advantageous in deinking high-quality wastepapers, whose reuse is usually limited as the nonimpact inks (toners) polymerize onto the paper surface using thermoplastic binders during the high-temperature printing process. In the chemical deinking process, the toner particles usually remain large, flat, and rigid and separate very poorly from papers during the fiber/ink separation stages (Jiang and Ma 2000).

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12.2  Enzyme Producing Microbes Twelve bacterial culture has been isolated from alkaline sediments from Lonar Lake in Maharastra India. Based on 16S rRNA sequencing, biochemical characterization, and phylogenetic analysis, all the isolates were identified. For the first time in Lonar lake, bacteria such as haloalkaliphilic Marinobacter excellens, Alkalimonas delamerensis, Roseinatronobacter monicus, and Rhodobaca bogoriensis were identified (Borgave et  al. 2012). In another research conducted by Kladwang et  al. (2003), about 490 alkaline-tolerant fungi from a natural environment using petri dishes containing potato dextrose agar medium buffered at pH 11.0 were identified. Many of the alkaline tolerant fungi has been isolated from fifty one samples from different habitats of Thailand. This research indicated that a good source of alkaline enzyme production can be found from alkaline-­tolerant fungi isolated from tree holes in alkaline and acidic environments. A bacterium Bacillus pumilus SV-205 produces xylanase in an optimized fermentation condition. A medium containing a mixture of yeast extract and peptone yields high xylanase as compared to other combinations. High pH stability over a range of 6–11 in 24 h is the major characteristic feature of this enzyme which can also maintain 65% activity after 2  h incubation at 60  °C (Nagar et al. 2012). In Barabanki district of Uttar Pradesh, India, George et al. (2001) isolate a novel alkalothermophilic actinomycete with optimum growth at pH 9 and 50 °C from self-­ heating compost. Thermomonospora sp. was able to produce 23  IU/ml carboxymethyl cellulase (CMCase) enzyme purified under fractional ammonium sulfate precipitation followed by cellulose affinity chromatography and Sephacryl S-200 gel filtration. The enzyme retained 100% activity at 50 °C for 72 h and had halflives of 7 and 3 h at 60 and 70 °C, respectively. The enzyme activity was tested as an additive to laundry detergents based on its stability in the presence of commercial detergents viz. Ariel, Henko, and Surf Excel. During an investigation exploring possible sources of novel thermophilic species in natural products, a novel thermophilic and alkaliphilic actinomycete capable of producing alkaline cellulase from soil of a tropical rain forest in Yunnan province, China, was isolated and identified (Wu et al. 2018). The whole-­cell hydrolysates were found to contain glucose and ribose. The organism was identified as Genus Streptomyces based on 16S rRNA gene sequence analysis. Organism formed a distinct phyletic line together with closely related type strain called Streptomyces burgazadensis ZIR7 T. Strain named Streptomyces thermoalkaliphilus represent a novel species in the genus Streptomyces based on its phenotypic, chemotaxonomic, and phylogenetic characteristics. Bilanenko et al. (2005) reported an isolate representing the group Ascomycete from saline soda soils of Central Asia and Africa. The bacterium described as Heleococcum alkalinum sp. novel was isolated on alkaline agar with carboxymethyl cellulose (CMC) and was a dominant species in samples of soda soils with pH >10 and relatively high salinity. It shows an alkali-­tolerant adaption by growing within the pH range of 6.7–10.8. This cellulolytic activity of an alkaliphilic obligate anaerobic bacterium, Z-7026, which was isolated from the microbial community of soda-lake

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sediments belonging to the cluster III of Clostridia with low G  +  C content was investigated by Zvereva et al. (2006). The bacterium has the ability of growing in media with cellulose or cellobiose as the sole energy source. The maximum growth rate on cellobiose was found at a pH range of 8.5–9.0, while that of cellulase synthesis, assayed by using a novel fluorimetric approach, was observed at pH 8–8.5. In the laboratory, bacterium Penicillium citrinum (MTCC 6489) was previously isolated from soil producing an alkali-tolerant and thermostable cellulase (Dutta et al. 2008). The study reports the presence of alkali-stable cellulase from alkali-tolerant fungus Penicillium citrinum for the first time, which may have potential effectiveness as additives to laundry detergents. According to Vyas and Lachke (2003), two extracellular alkali-stable 1,4-β-d-­ glucan-4 glucanohydrolase (EC3.2.1.4) fractions, namely, EndoA and EndoB, were separated from the culture filtrate of an alkalotolerant Fusarium strain. These enzymes are found to be suitable for deinking mixed office wastepapers leading to the increase in brightness with decrease in ink counts of the recycled paper. Probable mechanism of enzymatic deinking process was schematically presented based on the distinct properties of endoglucanases. Picart et al. (2007) reported one fungal strain from subtropical soils on the medium supplemented with rice straw exhibiting high cellulase activity. Using isoelectric focusing, zymography, and sodium dodecyl sulfate polyacrylamide gel electrophoresis, these new strains were identified as Penicillium sp. CR-316 and Penicillium sp. CR-313 which indicated that the strains secreted multiple enzymes that hydrolyze cellulose. Crude cellulose produced by Penicillium sp. CR-316 has potentials in industrial applications since it showed activity and stability at high temperature and produced a thermostable cellulase. Kalpana and Rajeswari (2015) has isolated Streptomyces from agricultural waste capable of producing enzymes for degrading xylan. Streptomyces sp are vital source of enzymes involved in lignocellulosic degradation. Isolate was reported to grow on different types of feedstuffs such as oat spelt xylan, sugarcane molasses, tomato pomace, rice bran, wheat bran, and sawdust under submerged fermentation conditions. The xylanase activity in each production medium was confirmed by measuring the amount of reducing sugars liberated from the medium by the DNS method using crude extract which was found to have an application in deinking of newsprint. Nadagouda et al. (2016) isolate cellulase enzyme from Trichoderma viride GSG12 under solid-state fermentation technique using cheap and readily available agricultural waste material called rice bran. This indicates the possibility of using rice bran to produce cellulase using Trichoderma viride under solid-state fermentation. The finding shows that cellulase production can be influence by optimal pH, initial moisture level of the medium, incubation temperature, inoculum size, and incubation time. The optimum pH, initial moisture level, incubation temperature, and inoculums size were 5.5, 70%, 32 and 2  ×  108 spores/flask, 120  h, respectively. Increased enzyme production was favored by supplement of lactose and corn-steep solid to the rice bran. From various mangrove sites in the Philippines, conventional as well as analytical profile index (API) were used to characterize and phenotypically identify five

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promising cellulase producing bacterial strains. The finding provide data regarding species of Bacillus producing cellulase enzyme and additional knowledge regarding the bacterial diversity of mangrove forests in the Philippines (Tabao and Monsalud 2014). Makky and Abdel-Ghany (2009) described old newspaper (ONP) waste as a carbon source for growing Bacillus subtilis where avicelase and carboxymethylcellulase (CMCase) enzymes are estimated in the culture filtrate. Bacillus subtilis CMCase has more activity at optimal temperature and pH than Avicelase. Deinking with these enzymes brings about an increase in brightness of the sheet effective removal of ink particles and also prevents redeposition onto the fiber surfaces. These findings indicate that enzymatic deinking can perform better than the conventional chemical method. Bacillus halodurans was purified to homogeneity to produce an extracellular haloalkaline cellulase by bioconversion of lignocellulosic waste by (Annamalai et al. 2013). The enzyme has retained up to 80% activity at 80 °C, 12% and 35% temperature, pH and NaCl with optimum of 60 °C, 9.0 and 30% respectively. When detergents and organic solvents such as n-hexane, acetone and acetonitrile are present, the enzyme was found to be stable. This indicates that a purified cellulase produce from Bacillus halodurans utilizing ligncellulosic biomass could be of great potentials in industrial process. Maitan-Alfenas et al. (2016) used Aspergillus nidulans to isolate and characterize xylanase in Pichia pastoris. At 60 °C and 7.5 as well as 50 °C and 6.0 temperature and pH, respectively, this enzyme has its optimum activity which is completely inhibited by SDS and HgCl2. Another important bacterium capable of producing an extracellular and thermostable xylanase enzyme is Bacillus pumilus ASH when grown on solid-state fermentation (SSF). When wheat bran is moistened with deionized water at a substrate to moisture ratio 1:2.5 (w/v), higher xylanase activity is obtained with optimum production temperature of 37 °C. The enzyme activity was slightly lower in solid-state fermentation (SSF) than in submerged fermentation technique, but the ability of the organism to produce such a high level of xylanase at room temperature, with deionized water and with no addition of any mineral salts in SSF, could lead to substantial reduction in the overall cost of enzyme production.

12.3  Cellulase and Its Applications Microbial cellulases have been focused as the important biocatalysts being multiplex in nature and bearing extensive applications. Cellulase and hemicellulase enzymes are both synthesized by fungi and bacteria as seen in Table 12.2. These microorganisms can be aerobic, anaerobic, mesophilic and thermophilic. Among them, the most commonly studied fungal genera are Thermomonospora, Trichoderma, and Aspergillus. Fungal and bacterial cellulases are structurally similar (Fig. 12.2). Fungal cellulase having two domains: a catalytic domain and a cellulose binding module. Commercially, cellulase enzymes have been accessible for 30 years or more, and these enzymes have been used for study purpose as well as

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Table 12.2  Microbial enzymes and their sources Sample Soil

Soil

Soil

Wastepaper

Agricultural waste

Waste photocopy paper Wild herbivore, rain deer Soil, compost, animal waste slurry Wastepaper Water

Soil

Location Pulp and paper industries, India Macuya rain forest, Pucallpa, Peru

Organism Bacillus subtilis

Enzyme Cellulase

pH 4.0

Temp (°C) References 60 Pala et al. (2004)

Cellulase Aspergillus sp. LM-HP32 and Penicillium sp. LM-HP33 and 37 Cellulase Penicillium sp. Iguazú CR-313 and rainfalls, CR-316 Argentina USM Campus, Aspergillus niger Cellulase, hemicellulose Penang, Malaysia Cellulase, Cairo, Egypt Bacillus xylanase thuringiensis MAM-29 and MAM-38 Medellin, NA Cellulase, Colombia amylase

4.8– 9.4

28

Vega et al. (2012)

4.5

65

Picart et al. (2007)

6.0

50

Lee et al. (2013)

Wayanad, Kerala, India

Escherichia coli SD5

3–7.6 60– 80

Abo-State et al. (2013)

7.0

40

Gil et al. (2013)

Cellulase, xylanase

NA

37– 39

Kumar et al. (2018)

Jeju Island, South Korea

Bacillus subtilis CMCase, C5–16 and S52–2 avicelase, xylanase

5.0

50

Kim et al. (2012)

NA

NA

9–11 50

Lonar Lake, Buldhana, Maharashtra, India Vellore, Tamil Nadu, India

Many haloalkalihpilic bacteria

Cutinase, amylase Lipase, amylase, caseinase, cellulose Xylanase

Bacillus halodurans FNP135

Xylanase

8–9.5 65

Bacillus pumilus

Xylanase

6–11 60

Streptomyces rameus L2001

Xylanase

5–8

Chandigarh, Old Punjab, India newsprint, magazine, inkjet, Xerox Soil Ambala Cantt, Haryana, India Soil Tianshan Xinjiang, China

Streptomyces sp.

10.5

23

7.5

37

70

Wang et al. (2018) Kanekar et al. (2008)

Kalpana and Rajeswari (2015) Virk et al. (2013)

Nagar et al. (2012) Li et al. (2010) (continued)

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Table 12.2 (continued) Sample Industrial effluents

Compost pit Wastepaper Soil

Location Shreyans Paper Industry, Ahmedgarh, Punjab, India BREC Sadra, Gujarat, India Chandigarh, Punjab, India Effluents of paper industries, India

Organism Aspergillus nidulans KK-99

Enzyme Xylanase

Temp pH (°C) References 8–8.5 55 Taneja et al. (2002)

Bacillus altitudinis DHN8 Bacillus halodurans Bacillus pumilus AJK10414

Xylanase

8.0

Xylanase and laccase Xylanase, pectinase

45– 55 8–9.5 65 8.5

55

Adhyaru et al. (2017) Virk et al. (2013) Singh et al. (2012)

Fig. 12.2  A simplified schematic representation of the enzymatic action of cellulase, involving exoglucanase, endoglucanase, and β-glucosidase

industrial researchers. The different applications of cellulase enzyme in this industry have reached a considerable increase in the last decade. The conventional method of the woody raw material using refining and grinding happens to pulp fines with high content, bulks, and stiff. However, the enzymatic pulping process using cellulase leads to decrease in the utilization in the energy during refinement and improving the strength content of handsheet. In Peru, a soil from an undisturbed forest was investigated for fungi capable of producing alkaline cellulase. At different PH value, plate clearing assay and carboxymethyl

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cellulase as substrate, 11 of 50 morphological colonies were selected. These 11 fungal strains produced cellulase of high alkaline PH values in a liquid culture media. The best producers of cellulase in highest productivities are the Penicillium sp. LM-HP33, Penicillium sp.LM-HP37 as well as Aspergillius sp. LM-HP32. These fugal strains are found to be suitable production of alkaline cellulase (Vega et al. 2012). Pathak et al. (2011), conducted a study on deinking photocopier papers using chemicals and commercial cellulase enzyme where they optimized all the parameters of deinking experiment for hydro pulping. Ink removal efficiency and freshness were improved by 24.6% and 12.6%, respectively, along with reduction of drainage time of 11.5% as compared to chemical deinking. As compared to fungi, bacteria have a high rate of cellulase enzyme production rate due to its advantage of high bacterial growth rate. The most important parameters for successful production of cellulase enzyme are the screening of the organism, optimization of fermentation conditions and selection of substrates. ­ Using carboxymethyl cellulase as substrate, (Ariffin et al. 2006) produced enzyme cellulase from local isolate of Bacillus pumilus EB3.This enzyme screened from this bacterium was purified using ion exchange chromatography using anion exchanger for cellulase characterization. Rawat and Tewari (2012) isolated and identified microorganism which hydrolyzed carboxymethyl cellulose (CMC) as Bacillus subtilis strain LFS3. Gel filtration chromatography, ion exchange, and sodium sulfate precipitation are the methods used to isolate and screen enzyme carboxymethylcellulase with overall recovery of 15%. Optimum temperature and pH for active profile of this enzyme was 60 °C and 4.0 respectively. A fungi called Coprinopsis cinerea was found to have the ability of producing a crude cellulase and xylanase enzymes with potentials of deinking photocopier wastepaper deink photocopier wastepapers as reported by Pathak et al. (2014). In their view to achieve maximun and possible efficiency without affecting paper and its strength propertis, enzyme dose, point of enzyme addition, pulp consistency, and reaction time were investigated which also confirmed the potential of crude enzyme of C. cinerea for deinking of photocopier wastepapers. Effects on the use of cellulase for deinking of office wastepaper were investigated by Tsatsis et al. (2017). Better results were achieved by the use of enzyme in deinking experiment as compared to those in which enzymes are inactive. It was discovered that enzyme application has disadvantage if specks surface of the deink paper sheets was uses as compared conventional deinking. Based on their finding, more research is needed in formulations of enzyme with better performance under alkaline conditions as well as the types of paper printed in different photocopier and laser printers. Abo-State et al. (2013) isolated Bacillus strains from agricultural waste and identified as Bacillus thuringenesis which have the ability to produce cellulase and xylanase based on their pH and temperature. Stability at different temperatures (60–80 °C) at separate duration was also investigated. Zhang et al. (2008) evaluate three commercial cellulase enzymes for their application on deinking artificially aged old newspaper (ONP) mixed with fresh old magazine (OMG) in a ratio of 7:3. At the start of repulping, these enzymes were added followed by incubation for 3 h.

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Despite the fact that cellulase enzymes were able to remove a significant amount of ink from ONP/PMG, they have low efficiency than using conventional methods of either sulfite or alkaline deinking chemistry. None of the three cellulase enzymes tested were able to separately deink aged ONP/OMG, and a poor deinkability was also observed by using either sulfite or alkaline chemistry. However, the research indicates a significant increase of deinking when a combination of enzyme and sulfite is applied which provide a potential strategy of achieving effective deinking of old newspapers at neutral pH. Enzymatic deinking has added advantages over conventional deinking viz. Reduced alkali usage, improving fiber brightness, and greater strength property. Moreover, enzymatic deinking process also prevents alkaline yellowing and reduces environmental pollution. However, excessive use of enzymes can be degradable as it might lead to significant hydrolysis. With the aim of increasing the rate of production, cellulase has been pursued by several mills in improving the drainage. These enzymes are also used in the production of easily biodegradable stationary objects including paper towels and sanitary paper (Kuhad et al. 2011). Laccase mediator system was used in a study conducted to identify the similarity on the application of cellulase/hemicellulase for deinking printed fibers from newspapers and magazines. In this regard, commercially available endoglucanase and endoxylanase activities and a commercial laccase were evaluated in the presence of synthetic or natural mediators. They concluded that factors to be considered for the application of enzymatic deinking processes in addition to enzymes include ink types, printing m ­ ethods, and fiber/ink separation process (Ibarra et al. 2012). Lee et al. (2007) also developed a laboratory procedure for enzymatic deinking of wastepapers using cellulase and hemicellulase enzymes produced from Aspergillus niger. Using an optimized floatation system of 6.0 and 45 °C pH and temperature, respectively, an optimum deinking efficiency with these enzymes was enhanced to 95%. The deinked papers are found to have similar properties with commercial papers indicating the effectiveness of the developed enzymatic process.

12.4  Xylanase and Its Applications Xylanase comprises the hydrolytic enzymes which are capable of breaking the β-1,4 backbone of the multiplex plant cell wall polysaccharide. Xylan is the second largest polysaccharide after cellulose (Yadav et  al. 2017, 2018). A diverse array of microorganisms like bacteria, actinomycetes, yeast and fungi are involved in the hydrolysis of hemicellulose as indicated in Table  12.2. Wood is processed for debarking, chipping and screening. Then a chip undergoes steaming process so that the microorganisms become lesser in number. After this, the chips are allowed to cool down and inoculated with biopulping fungus. The biopulping process is cost effective and technologically feasible. The main advantage is the decrease in consumption of energy as well as the increase in mill consumption. The processes also lead to enhancement in the properties such as strength of the paper, and reduced

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environmental impacts (Khonzue et  al. 2011). From the past studies, it has been concluded that pre bleaching method with xylanase enzyme is cheaper and eco-­ friendly. It also decreases the significant amount of chemicals which are indulged in order to get brightness in chemically bleaching process. In conventional method of paper making process, the manufacturers use hazardous chemicals which impart negative impact to the environment. A high amount of xylanase enzyme was produced from Bacillus pumilus SV-205 using optimized fermentation conditions. The bacterium secretes maximum amount of cellulase free xylanase in combination with yeast and peptone which also enhanced highest xylanase production that differ from other combinations. The enzyme maintained a thermal stability of 65% activity after incubation at 60 °C for 2 h (Nagar et al. 2012). Li et al. (2010) isolate xylanase with biobleaching potentials on wheat straw pulp from Streptomyces L2001 with stable optimum temperature of 70 °C and pH of 5.3. High production of xylanase from another bacterium called Bacillus altitudinis DHN8 followed by its purification and application was presented by Adhyaru et al. (2017). Using response surface technology, enzyme yield was improved by optimizing submerged fermentation conditions which includes incubation time, temperature, agitation speed, sorghum straw, inoculum size, and gelanin. This leads to twofold increase in activity based on the abovementioned optimized conditions as compared to activity in one factor at a time optimization. The research indicates a potential use of Bacillus altitudinis for biodeinking and biobleaching. For pollution free environment, the recently employed technique is the recycling of civic paper waste by enzyme based technology. In these regards, a newly isolated bacterium for recycling of laser jet paper waste was isolated for its potential ability to purify xylanase enzyme by Maity et al. (2012).This potent xylanase producing bacterium from microbial consortia of termite gut was identified as Bacillus sp CKBx1D based on 16S rRNA sequence. Response surface methodology was the technique used to optimize all operational parameters, while purified enzyme mixture was used for laser printed paper waste deinking potentials. This deinking potential was detected from the enzyme at a pH of 6.8 with 72 h continuous shaking at constant temperature of 35 °C. Hence, the bacterial isolate and its xylanase enzymatic system could efficiently be used in recycling paper waste as deinking agent. Using cheap agricultural residue, pectinase and xylanase enzymes were isolated for the first time from alkalo thermotolerant bacterial strain with potentials of deinking capabilities. The enzymes may also play important role in making enzymatic deinking an eco-friendly having 50% decrease of chemicals, commercially viable with better paper quality (Singh et al. 2012). According to (Gessesse and Mamo 1999), overall cost of xylanase enzyme production from Bacillus sp. AR-009 can be greatly reduce using solid-state fermentation technique. This bacterium has the ability to produce dry bacterial bran xylanase activity when grown in solid-state fermentation and produced a high bacterial bran xylanase activity with wheat bran as substrate. The ability of the organism to produce high xylanase activity at alkaline pH and lower wheat bran to moisture ratio could have a potential advantage in minimizing the risk of contamination. In addition, the cost of downstream processing during

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product upgrade and that of waste treatment steps can be greatly reduce since the enzyme can be produce with a least quantity of liquid. Dutta et  al. (2007) studied the culture filter of Penicillium citrinum grown on wheat bran bed in solid-state fermentation to purify an extracellular xylanase enzyme. Moderately thermostable xylanase showed optimum activity at 50 °C at pH 8.5. Purification and characterization of this novel endoglucanase free alkaliphilic xylanase from the alkali tolerant fungus P. citrinum were discovered for the first time which may have potential implications in paper and pulp industries. Desai and Iyer (2016) isolate cellulase free xylanase enzyme from fungi for deinking of Old News Paper (ONP) pulp. Aspergillus niger strain DX-23 from the total 16 fungal producing enzyme isolates had a maximum xylanase of 48.9  ±  0.02  Uml−1. The enzyme deinked ONP pulp efficiently with improved brightness after optimization and subsequent H2O2 treatment as compared to untreated pulp. According to (Taneja et al. 2002), an alkaline thermostable xylanase was produced by alkalophilic fungi called Aspergillus nidulans KK-99 in an alkaline medium consisting of wheat bran, KNO3, pH 10.0 and temperature of 37 °C. this partially purified enzyme was stable at a pH range of 4.0–9.5 and temperature of 55 °C and reach optimum activity at a pH  8.0 and same temperature of 55  °C where decrease of ink and brightness of recycled paper was achieved by this enzyme treatment. An investigation in to xylano pectinolytic enzymes for deinking of school wastepaper was conducted by Shatalov and Pereira (2008). This biodeinking in combination with conventional deinking approach leads to an increase in BOD and COD values of effluent as ­compare to use of only conventional deinking method. The usage of xylanase enzymes in deinking process has some limitations depending on different parameters. These comprise different factors like narrow pH ranges, thermal instability, no proper end product, and cost of production of enzyme. Biological treatment using xylanase enzyme has proved to be helpful in both reducing the costs and also improved the quality of the fiber. Xylans are highly available to hydrolytic enzymes as they are not having a complex structure. Therefore, its specific activity becomes two to three times higher as compared to crystalline cellulose form (Shatalov and Pereira 2008). In order to obtain whiter and brighter pulpy matter to process better quality paper, it becomes necessary to separate the residual lignin with the use of bleaching method (Azeri et al. 2010). The most important advantages of biobleaching include: (a) Reduction in the use of the bleaching chemicals, (b) Enhanced quality of the paper and pulp, (c) Improving whiteness and brightness of the pulp, and (d) Decrease effluent toxicity and pollution load.

12.5  Laccase and Its Applications Laccase is the oldest and widely studied enzymatic system discovered for the first time in 1883 by Yoshida from the exudates of Japanese lacquer tree called Rhus vernicifera. In 1985, Bertrand discovered it as a metal containing oxidase, while in

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1896, it was demonstrated by both Bertrand and Laborde to be present in fungi. The redox potential is found to be higher in fungal laccases rather than bacterial or plant laccases up to 800 mV. Therefore, these are found to be beneficial in lignin degradation or in the eradication of phenolic toxins arising during lignin degradation. Laccase is found to be present in higher plants and fungi such as basidiomycetes, white root fungi and ascomycetes. White rot basidiomycetes are responsible for efficient degradation of lignin and the enzymes involved in lignin degradation includes lignin peroxidases, manganese dependent peroxidases and laccases. Also, fungal laccases have been involved in cellular processes in many ways including sporulation, plant pathogenesis and other specificity. Woody chips when pretreated with lignolytic fungus leads to increase in the strength of the pulp while decrement in the requirement of energy for the mechanical pulping process. The most widely studied application of laccase in this industry is bleaching of kraft pulp. It was studied that when SL4, a lignolytic fungal strain, was applied, it reduced 25% usage of chlorine during bleaching of kraft pulp and produced 1.8 unit brightness (Kaur and Nigam 2014). C. albidus responsible for producing laccase helps in the reduction of lignin content found in the eucalyptus wood and happen to be useful in the process of biopulping. Fungal laccases also efficiently remove toxic effluents coming from the pulp mills which contain a significant number of phenolic compounds and chlorolignins. Laccase mediated biobleaching process is a friendly way to improve pulp and paper production. The ink removal capability of laccase and xylanase enzymes from an alkalophilic bacteria on recycled old newspaper (ONP) in combination with physical deinking method of sonication and microwaving were investigated. Parameters such as PH, enzyme dose as well as treatment time of these enzymes were optimized statistically using Response Surface Methodology in which any mediator supplementation for deinking was not required. Optimization of these deinking parameters were conducted statistically using response surface methodology in which for the first time laccase did not need any mediator supplementation (Virk et al. 2013). An enzyme showing alkaliphilic laccase activity was purified from the culture supernatant of Myrothecium verrucaria 24G-4 by (Sulistyaningdyah et  al. 2004). The enzyme was very stable in alkaline conditions with an optimum pH of 9.0 and was able to remove synthetic dyes under the same conditions which confirm the function of this enzyme in alkaline environment.

12.6  Amylase and Their Applications Amylases hydrolyze starch molecules and give diverse array of products including dextrin and glucose units (Fig. 17.2). Microbial amylase enzymes are available in market in large number and they are replacing chemicals involving hydrolysis of starch. The first commercially produced microbial enzyme was amylase which was of fungal origin in 1894 and used as therapeutic aid to cure digestive disorder. Nearly 25% of the global enzyme comprises amylase. Enzymatic pretreatment of

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wastepaper pulp was conducted for deinking process at neutral conditions using amylase and cellulase enzymes and ethoxylated fatty acids as surfactant. At a temperature of 40 °C, this enzymatic deinking process was conducted by flotation consistency of 0.8% within 6 min (Gil et al. 2013). In a future work, effectiveness of combine enzymes in removal of waste photocopy paper and other wastepapers mixtures is expected to be discussed. Amylases are distributed in plants, animals and microbes. However, amylases produced by microbes are replacing the chemical processing methodology indulged in paper industries due to cost effectiveness and technical advantages. Amylases classified into two types on the basis of catalysis, endoamylase and exoamylase. Endoamylases carry out hydrolysis in a random manner while exoamylases hydrolyze from the nonreducing end. Endoamylases results in oligosaccharides with branching of different chain length and exoamylases forms short end products. Main fungi involve in the production of industrially important enzymes are Aspergillus niger, A. oryzae, and A. flavus. Also these fungi are capable of producing large quantities of amylases which can be used commercially. Aspergillus niger having significant amount of hydrolytic capacities in the production of α-amylase (Sahni and Goel 2015) Sakthivel et al. (2010) isolated and screened some bacteria that inhabit decaying vegetables for the production of amylase enzyme. Enterococcus pseudoavium is the only specie identified to have a relatively higher amylase activity of the bacterial species tested. Four days after growth, the organism deinked pulp completely when grown with paper pulp. When cultured with paper pulp, the bacterial culture immobilized in sodium alginate beads can decolorize this paper within 4 days. This shows the ability of extracellular amylase produced from Enterococcus pseudoavium to effectively deink and decolorize paper pulp within 4 days of incubation. The role of α-amylases in the paper industry as shown in Table 17.4 is to modify starch of coated mixed paper which comprises production of lower viscous, starch with high molecular weight, deinking, drainage and cleaning of paper. The coating tends to make the paper smoother and stronger, to improve the quality of writing. Since the viscosity of natural starch remains high enough for paper sizing and this can be changed by degradation of the polymer with the use of α-amylases in the batch or continuous culture. Starch being good sizing agent helps in giving fine finish of the paper, improvement of the paper quality, and reusability, besides behaving as a perfect paper coating (Singh et al. 2012).

12.7  Lipase and Its Applications Lipases are the enzymes which hydrolyze long chain triglyceride and constitute as one of the most focal biocatalysts for biotechnological applications. It was first found to be present in pancreas by J. Eberle in 1834 and during 1856 by C.I Bernard. Under aqueous conditions, they are able to release fatty acids and glycerol by acting on carboxyl ester bonds present in triacylglycerol (Gupta 2004). These are serine

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hydolases and are able to work independently without the use of any cofactor. Microbial lipases play an important part in the field of biotechnology due to its versatility and ease of mass production. It has got wide enzymatic properties. Both bacterial lipase and fungal lipase are widely used in different industries. Lipase producers include bacteria, fungi, yeast, and actinomycetes. Examples of fungi producing lipase are Acinetobacter radioresistens, Aeromonas hydrophila, Aspergillus oryzae (Andualema and Gessesse 2012). Wood is the cheap source of paper pulp and pitch describes the hydrophobic contents of wood (triglycerides and waxes). These are appearing to create serious problem in the paper processing. The problems might be sticky deposits and holes or spots in the finish product. However, lipases remove the pitch from the pulpy matter during the process of paper making process. Around 90% of triglycerides present in the pitch get hydrolyzed into glycerol, monoglycerides, and fatty acids by lipase enzymes which are having less stickiness and more hydrophilic character (Jaeger and Reetz 1998). In Japan, a paper industry called Nippon Paper industries developed a method to control pitch that utilizes the Candida rugosa fungal lipases to hydrolyze up to 90% of the woody triglycerides present. Lipases have many advantages in general as its utility increase the rate of pulping, enhancing whiteness and intensity power, decrement in the usage of chemical, equipment with prolong life, reduction in the level of pollution in the wastewater, saving energy utility, and time and reduction in the composite cost. Pseudomonas sp. (KWI-56) has an added advantage as addition of its lipase to the deinking process leads to increase in the whiteness of the paper and reduce usage of residual inking. A thermophilic isolate of Bacillus coagulans BTS-3 can be used to screen alkaline lipase. The bacterium can be enhanced substantially when parameters like nitrogen source, carbon source, and initial pH of culture medium were consecutively optimized. Enzyme activity of culture medium was obtained in 48 h at 55 °C and pH 8.5 with refined mustard oil as carbon source and a combination of peptone and yeast extract as nitrogen sources. Maximum activity of the enzyme was achieved at 55 °C temperature and 8.5 pH and was also stable between the pH range of 8.0 and 10.5 at temperatures up to 70 °C. This purified lipase enzyme indicates a variable hydrolytic activity toward various 4-nitrophenyl esters (Kumar et al. 2005).

12.8  Other Microbial Enzymes and Their Applications Cutinase and other microbial enzymes have also play a vital role in recycling of wastepaper (Table 17.2). For the first time, Wang et al. (2018) reported the effect of cutinase enzyme on the deinking of mixed office wastepaper (MOW). Combination of cutinase, amylase, and some complicated surfactants can be used to replace the conventional chemical deinking methods at neutral deinking process. When these enzymes are treated in combination of surfactants mixed with pulp and office wastepaper at a temperature of 50 °C for 30 min, the brightness, ink removal rate, tensile index, and tear index of the deink paper will increase significantly. Hemicellulase

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was combined with laccase mediator system (LMS) by Xu et al. (2011) to deink old newsprint (ONP). The result indicates the effective residual ink concentration was lower as compared to pulp deink with hemicellulase or LMS individually. According to this study, an environmentally friendly and effective deinking method can be achieved when cutinase and amylase enzymes combine with cardanol polyoxyethylene ether and other surfactants.

12.9  Conclusion and Future Prospects Industrial utilization of wastepapers in the production of new one is increasing globally. Currently, pulp and paper industry is one of the largest consumers of wood. Based on their demands due to global economic growth, more trees will be harvested, and waste will be consumed and disposed in the environment. With increasing demand of paper globally and high cost of conventional methods for deinking and recycling of wastepapers, attentions are now focused on the use of enzymes for eco-friendly deinking of wastepapers especially extremozymes such as cellulase, hemicellulase, xylanase, amylase, and laccase that are able to deink wastepapers at optimum environmental conditions. Combination of enzymatic and chemical methods can greatly increase the brightness of deink wastepaper, ink removal, and quality of recycled wastepapers. However, biological deinking method of wastepaper with the help of varied microbial enzymes comprises an eco-friendly way that can produce high-quality paper without relying upon the traditional usage of harmful chemicals. Thus, more efforts should be efficiently directed to still tap the wide underlying potential of these fungal lignolytic enzymes.

References Abo-State MA, Fadel M, Abdellah EM, Ghaly MF (2013) Studying the stability of cellulases and xylanase produced by Thermophilic and alkaliphilic bacterial strains isolated from agricultural wastes. Am Eurasian J  Agric Environ Sci 13(11):1568–1575. https://doi.org/10.5829/idosi. aejaes.2013.13.11.11262 Adhyaru DN, Bhatt NS, Modi HA, Divecha J (2017) Cellulase-free-thermo-alkali-solvent-stable xylanase from Bacillus altitudinis DHN8: over-production through statistical approach, purification and bio-deinking/bio-bleaching potential. Biocat Agric Biotechnol 12:220–227. https:// doi.org/10.1016/j.bcab.2017.10.010 Andualema B, Gessesse A (2012) Microbial lipases and their industrial application: review. Biotechnology 11(3):100–118. https://doi.org/10.3923/biotech.2012.100.118 Annamalai N, Veeramuthu M, Elayaraja S (2013) Thermostable haloalkaline cellulase from Bacillus halodurans CAS 1 by conversion of lignocellulosic wastes. Carbohydr Polym 94(1):409–415. https://doi.org/10.1016/j.carbpol.2013.01.066 Ariffin H, Abdullah N, Kalsom MSU, Shirai Y, Hassan MA (2006) Production and characterisation of cellulase by Bacillus pumilus EB3. Int J Eng Technol 3(1):47–53 Azeri C, Tamer AU, Oskay M (2010) Thermoactive cellulase-free xylanase production from alkaliphilic Bacillus strains using various agro-residues and their potential in biobleaching of kraft pulp. Afr J Biotechnol 9(1):63–72

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Bilanenko EN, Sorokin DY, Kozlova MV (2005) Heleococcum alkalinum, a new alkali-tolerant ascomycete from saline soda soils. Mycotoxin 91(1):497–507 Borgave SB, Joshi AA, Kelkar AS, Kanekar PP (2012) Screening of alkaliphilic, haloalkaliphilic bacteria and alkalithermophilic actinomycetes isolated from alkaline soda lake of Lonar, India for antimicrobial activity. Int J Pharma Biosci 3(4):258–274 Desai DI, Iyer BD (2016) Biodeinking of old newspaper pulp using a cellulase-free xylanase preparation of Aspergillus niger DX-23. Biocatal Agric Biotechnol 5:78–85. https://doi. org/10.1016/j.bcab.2015.11.001 Dutta T, Sengupta R, Sahoo R, Ray SS, Bhattacharjee A, Ghosh S (2007) A novel cellulase free alkaliphilic xylanase from alkali tolerant Penicillium citrinum: production, purification and characterization. Lett Appl Microbiol 44:206–211. https://doi.org/10.1111/j.1472-765X.2006.02042.x Dutta T, Sahoo R, Sengupta R, Ray SS, Bhattacharjee A, Ghosh S (2008) Novel cellulases from an extremophilic filamentous fungi Penicillium citrinum: production and characterization. J Ind Microbiol Biotechnol 35:275–282. https://doi.org/10.1007/s10295-008-0304-2 George SP, Ahmad A, Rao MB (2001) Studies on carboxymethyl cellulase produced by an alkalothermophilic actinomycete. Bioresour Technol 77(2):171–175. https://doi.org/10.1016/ S0960-8524(00)00150-4 Gessesse A, Mamo G (1999) High-level xylanase production by an alkaliphilic Bacillus sp. by using solid-state fermentation. Enzym Microb Technol 25:68–72 Gil HHA, Dovale SAM, VirneyHadely CL, Munoz OA, Casas BAE, Quintana MGC, Velasquez JJA (2013) Study of the enzymatic/neutral deinking process of waste photocopy paper. O Papel 74(8):61–65 Gupta MR (2004) Bacterial lipases: an overview of production, purification and biochemical properties. Appl Microbiol Biotechnol 64:763–781. https://doi.org/10.1007/s00253-004-1568-8 Ibarra D, Monte MC, Blanco A, Martínez AT, Martínez MJ (2012) Enzymatic deinking of secondary fibers: cellulases/hemicellulases versus laccase-mediator system. J  Ind Microbiol Biotechnol 39(1):1–9. https://doi.org/10.1007/s10295-011-0991-y Jaeger KE, Reetz MT (1998) Microbial lipases form versatile tools for biotechnology. Trends Biotechnol 16(9):396–403. https://doi.org/10.1016/S0167-7799(98)01195-0 Jiang C, Ma J (2000) Deinking of waste paper: flotation. Academic Press, Norcross, pp 2537–2544 Kalpana VN, Rajeswari VD (2015) Production of xylanase from various lignocellulosic waste materials by Streptomyces sp. and its potential role in deinking of newsprint. Asian J Biochem 10(5):222–229. https://doi.org/10.3923/ajb.2015 Kanekar PP, Joshi AA, Kelkar AS, Borgave SB, Sarnaik SS (2008) Alkaline Lonar lake, India – a treasure of alkaliphilic and halophilic bacteria. In: Sengupta M, Dalwani R (eds) Proceedings of Taal2007: The 12th World Lake Conference. The Ministry of Environment and Forests of the Government of India, Jaipur, pp 1765–1774 Kaur S, Nigam V (2014) Production and application of laccase enzyme in pulp and paper industry. Int J Res Appl Nat Soc Sci 2(4):153–158 Khonzue P, Laothanachareon T, Rattanaphan N, Tinnasulanon P, Apawasin S, Paemanee A, Eurwilaichitr L (2011) Optimization of xylanase production from Aspergillus niger for biobleaching of eucalyptus pulp. Biosci Biotechnol Biochem 75(6):1129–1134. https://doi. org/10.1271/bbb.110032 Kim YK, Lee SC, Cho YY, Oh HJ, Ko YH (2012) Isolation of cellulolytic Bacillus subtilis strains from agricultural environments. ISRN Microbiol 2012:1–9. https://doi.org/10.5402/2012/650563 Kirk TK, Jeffries TW, Gifford O, Drive P (1996) Roles for microbial enzymes in pulp and paper processing. American Chemical Society, Washington, DC Kladwang W, Bhumirattana A, Hywel-jones N (2003) Alkaline-tolerant fungi from Thailand. Fungal Divers 13:69–84 Kuhad RC, Gupta R, Singh A (2011) Microbial cellulases and their industrial applications. Enzyme Res 2011(1):1–10. https://doi.org/10.4061/2011/280696 Kumar S, Kikon K, Upadhyay A, Kanwar SS, Gupta R (2005) Production, purification, and characterization of lipase from thermophilic and alkaliphilic Bacillus coagulans BTS-3. Protein Expr Purif 41:38–44. https://doi.org/10.1016/j.pep.2004.12.010

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Kumar NV, Rani ME, Gunaseeli R, Kannan ND (2018) Paper pulp modification and deinking efficiency of cellulase-xylanase complex from Escherichia coli SD5. Int J  Biol Macromol 111:289–295. https://doi.org/10.1016/j.ijbiomac.2017.12.126 Lee CK, Darah I, Ibrahim CO (2007) Enzymatic deinking of laser printed office waste papers: some governing parameters on deinking efficiency. Bioresour Technol 98(8):1684–1689. https://doi.org/10.1016/j.biortech.2006.05.052 Lee CK, Darah I, Ibrahim CO (2013) Enzymatic deinking of various types of waste paper: efficiency and characteristics. Process Biochem 48(2):299–305. https://doi.org/10.1016/j. procbio.2012.12.015 Li X, She Y, Sun B, Song H, Zhu Y, Lv Y, Song H (2010) Purification and characterization of a cellulase-­free, thermostable xylanase from Streptomyces rameus L2001 and its biobleaching effect on wheat straw pulp. Biochem Eng J  52(1):71–78. https://doi.org/10.1016/j. bej.2010.07.006 Maitan-Alfenas GP, Oliveira MB, Nagem RAP, de Vries RP, Guimarães VM (2016) Characterization and biotechnological application of recombinant xylanases from Aspergillus nidulans. Int J Biol Macromol 91:60–67. Elsevier B.V. https://doi.org/10.1016/j.ijbiomac.2016.05.065 Maity C, Ghosh K, Halder SK, Jana A, Adak A, Mohapatra PKD, Mondal KC (2012) Xylanase isozymes from the newly isolated Bacillus sp. CKBx1D and optimization of its deinking potentiality. Appl Biochem Biotechnol 167(5):1208–1219. https://doi.org/10.1007/s12010-012-9556-4 Makky EA, Abdel-Ghany TM (2009) Cellulases applications in biological de-inking of old newspaper wastes as carbon source produced by Bacillus sp. Egypt J Exp Biol (Bot) 89(5):85–89 Nadagouda MG, Lingappa K, Bheemareddy VS, Malipatil SG (2016) Optimization of solid state fermentation conditions for the production of cellulase by using Trichoderma viride GSG12. Biosci Discov 7(1):1–6 Nagar S, Mittal A, Kumar D, Gupta VK (2012) Production of alkali tolerant cellulase free xylanase in high levels by Bacillus pumilus SV-205. Int J Biol Macromol 50(2):414–420. https://doi. org/10.1016/j.ijbiomac.2011.12.026 Pala H, Mota M, Gama FM (2004) Enzymatic versus chemical deinking of non-impact ink printed paper. J Biotechnol 108(1):79–89. https://doi.org/10.1016/j.jbiotec.2003.10.016 Pathak P, Bhardwaj NK, Singh AK (2011) Optimization of chemical and enzymatic deinking of photocopier waste paper. Bioresources 6(1):447–463 Pathak P, Bhardwaj NK, Singh AK (2014) Production of crude cellulase and xylanase from Trichoderma harzianum PPDDN10 NFCCI-2925 and its application in photocopier waste paper recycling. Appl Biochem Biotechnol 172:3776–3797. https://doi.org/10.1007/ s12010-014-0758-9 Picart P, Diaz P, Pastor FIJ (2007) Cellulases from two Penicillium sp. strains isolated from subtropical forest soil: production and characterization. Lett Appl Microbiol 45(1):108–113. https://doi.org/10.1111/j.1472-765X.2007.02148.x Rawat R, Tewari L (2012) Purification and characterization of an acidothermophilic cellulase enzyme produced by Bacillus subtilis strain LFS3. Extremophiles 16:637–644. https://doi. org/10.1007/s00792-012-0463-y Roushdy M (2015) Biodeinking of photocopier waste paper effluent by fungal cellulase under solid state fermentation. J  Adv Biol Biotechnol 2(3):190–199. https://doi.org/10.9734/ JABB/2015/15378 Sahni TK, Goel A (2015) Microbial enzymes with special reference to α-amylase. BioEvolution 2(1):19–25 Sakthivel M, Karthikeyan N, Meenakshi J, Palani P (2010) Optimization of culture conditions for the production of extracellular cellulase from Enterococcus pseudoavium. J  Exp Sci 1(11):25–29 Saxena A, Chauhan PS (2016) Role of various enzymes for deinking paper: a review. Crit Rev Biotechnol 8551(7):1–15. https://doi.org/10.1080/07388551.2016.1207594 Shatalov AA, Pereira H (2008) Effect of xylanases on peroxide bleachability of eucalypt (E. globulus) kraft pulp. Biochem Eng J 40(1):19–26. https://doi.org/10.1016/j.bej.2007.11.012

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Singh A, Yadav RD, Kaur A, Mahajan R (2012) An ecofriendly cost effective enzymatic methodology for deinking of school waste paper. Bioresour Technol 120:322–327. https://doi. org/10.1016/j.biortech.2012.06.050 Sulistyaningdyah WT, Ogawa J, Tanaka H, Maeda C (2004) Characterization of alkaliphilic laccase activity in the culture supernatant of Myrothecium verrucaria 24G-4  in c­ omparison with bilirubin oxidase. FEMS Microbiol Lett 230(2):209–214. https://doi.org/10.1016/ S0378-1097(03)00892-9 Tabao NS, Monsalud RG (2014) Characterization and identification of high cellulase-­producing bacterial strains from Philippine mangroves. Philipp J  Syst Biol 4(6):13–20. https://doi. org/10.3860/pjsb.v4i0.1562 Taneja K, Gupta S, Kuhad RC (2002) Properties and application of a partially purified alkaline xylanase from an alkalophilic fungus Aspergillus nidulans KK-99. Bioresour Technol 85(1):39– 42. https://doi.org/10.1016/S0960-8524(02)00064-0 Tavares APM, Xavier MRB, Evtuguin DV (2014) Biotechnology applications in pulp and paper industry. In: Biotechnology Vol. 12: Bioprocess engineering. Studium Press LLC, Houston, pp 561–581 Tsatsis DE, Papachristos DK, Valta KA, Vlyssides AG, Economides DG (2017) Enzymatic deinking for recycling of office waste paper. J  Environ Chem Eng 5(2):1744–1753. https://doi. org/10.1016/j.jece.2017.03.007 Vega K, Villena GK, Sarmiento VH, Ludeña Y, Vera N, Gutiérrez-Correa M (2012) Production of alkaline cellulase by fungi isolated from an undisturbed rain forest of Peru. Biotechnol Res Int 2012:1–7. https://doi.org/10.1155/2012/934325 Virk AP, Puri M, Gupta V, Capalash N, Sharma P (2013) Combined enzymatic and physical deinking methodology for efficient eco-friendly recycling of old newsprint. PLoS One 8(8):1–8. https://doi.org/10.1371/journal.pone.0072346 Vyas S, Lachke A (2003) Biodeinking of mixed office waste paper by alkaline active cellulases from alkalotolerant Fusarium sp. Enzym Microb Technol 32(2):236–245. https://doi. org/10.1016/S0141-0229(02)00273-9 Wang F, Zhang X, Zhang G, Chen J, Sang M, Long Z, Wang B (2018) Studies on the environmentally friendly deinking process employing biological enzymes and composite surfactant. Cellulose 25:1–11. https://doi.org/10.1007/s10570-018-1778-3 Wu H, Liu B, Ou X, Pan S, Shao Y (2018) Streptomyces thermoalkaliphilus sp. nov., an alkaline cellulase producing thermophilic actinomycete isolated from tropical rainforest soil. Antonie van Leeuwenhoek 111(3):413–422. https://doi.org/10.1007/s10482-017-0964-x Xu QH, Wang YP, Qin MH, Fu YJ, Li ZQ, Zhang FS, Li JH (2011) Fiber surface characterization of old newsprint pulp deinked by combining hemicellulase with laccase-mediator system. Bioresour Technol 102(11):6536–6540. https://doi.org/10.1016/j.biortech.2011.03.051 Yadav AN, Sachan SG, Verma P, Saxena AK (2015) Prospecting cold deserts of north western Himalayas for microbial diversity and plant growth promoting attributes. J  Biosci Bioeng 119:683–693 Yadav AN, Sachan SG, Verma P, Kaushik R, Saxena AK (2016) Cold active hydrolytic enzymes production by psychrotrophic Bacilli isolated from three sub-glacial lakes of NW Indian Himalayas. J Basic Microbiol 56:294–307 Yadav A, Verma P, Kumar R, Kumar V, Kumar K (2017) Current applications and future prospects of eco-friendly microbes. EU Voice 3:21–22 Yadav AN, Verma P, Kumar V, Sangwan P, Mishra S, Panjiar N, Gupta VK, Saxena AK (2018) Biodiversity of the genus Penicillium in different habitats. In: Gupta VK, Rodriguez-Couto S (eds) New and future developments in microbial biotechnology and bioengineering, Penicillium system properties and applications. Elsevier, Amsterdam, pp  3–18. https://doi.org/10.1016/ B978-0-444-63501-3.00001-6 Yadav AN, Mishra S, Singh S, Gupta A (2019a) Recent advancement in white biotechnology through fungi. Volume 1: Diversity and enzymes perspectives. Springer International Publishing, Cham

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Yadav AN, Mishra S, Singh S, Gupta A (2019b) Recent advancement in white biotechnology through fungi. Volume 2: Perspective for value-added products and environments. Springer International Publishing, Cham Zhang X, Renaud S, Paice M (2008) Cellulase deinking of fresh and aged recycled newsprint/ magazines (ONP/OMG). Enzym Microb Technol 43:103–108. https://doi.org/10.1016/j. enzmictec.2007.11.005 Zvereva EA, Fedorova TV, Kevbrin VV, Zhilina TN, Rabinovich ML (2006) Cellulase activity of a haloalkaliphilic anaerobic bacterium strain Z-7026. Extremophiles 10:53–60

Chapter 13

Arbuscular Mycorrhizal Fungi-Mediated Mycoremediation of Saline Soil: Current Knowledge and Future Prospects Dileep Kumar, Priyanka Priyanka, Pramendra Yadav, Anurag Yadav, and Kusum Yadav

13.1  Introduction The explosive rise of global population led to the increased exploitation of natural resources in response to the high demands for food, energy, and other things. Soil is the ultimate source of nutrients for the plant kingdom. The weathering of rocks present in the earth’s crust forms soil. The rocks are composed of minerals and the process of weathering occurs continuously. The soil has different concentrations of these minerals. If any specific type of minerals has a greater concentration, then it changes the quality of soils. For example, the saline soil holds a greater salt concentration; the acidic soil has low pH and sodic or alkaline soil possess high pH. The soil salinization is a serious problem for arable lands, which decreases the productivity of agricultural products. According to some reports, the soil salinization is gradually increasing in many parts of the world, mostly in arid and semiarid regions (Giri et al. 2003; Al-Karaki 2006). The amount of evaporation causes increased salt concentration or precipitation amount leading to a decrease in salt concentration in a given land area (Mahajan and Tuteja 2005). Salinization of land is a global problem, which is rapidly increasing. The United Nations Environment Program (UNEP) estimated that approximately 20% of agricultural land and 50% of cropland in the world is facing salt stress (Flowers and Yeo 1995). The extent of salt-affected soils is highest in the Asia Pacific region including Australia. The countries like Argentina, Australia, China, Egypt, India, Iran, Iraq, Pakistan, Thailand, former Soviet Union, and the United States of America (USA) are predominantly affected by soil salinization. Salt-affected soils are occupied about 7% of the earth’s land surface

D. Kumar · P. Priyanka · P. Yadav · K. Yadav (*) Department of Biochemistry, University of Lucknow, Lucknow, Uttar Pradesh, India A. Yadav Department of Microbiology, College of Basic Science and Humanities, Sardarkrushinagar Dantiwada Agricultural University, Banaskantha, Gujarat, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_13

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(Ruiz-Lozano and Azcon 2000) and about 5% of the total cultivated land around the world, i.e., 1.5 billion hectares (Sheng et al. 2008). In India, about 30 million hectares of coastal land is barren and uncultivable due to salinity. Soil salinization in arid and semi-arid regions is one of the major problems, which impose a challenge for sustainable agriculture in irrigated production systems. Plant growth is directly affected by high levels of sodium chloride and other salts in the soil  (Yadav et  al. 2019a; Yadav and Saxena 2018; Yadav et  al. 2015). The area of saline soil is rapidly increasing because of climate change, saline irrigation, and high evaporation, which induce salt accumulation in the soil, thus causing a significant decrease in irrigation practices. According to Kohler et  al. (2009), saline soils and saline irrigation constitute a serious production problem for vegetable crops, as saline conditions are known to suppress plant growth particularly in arid and semiarid areas. The high salt concentration is deposited in the soil to generate a low water potential zone in the soil, making it difficult for the plant to absorb water and nutrients. The basic physiology of salt and drought stress overlaps, therefore salt stress essentially results in a water-deficit condition in the plant that takes the form of a physiological drought (Mahajan and Tuteja 2005; Yadav 2017; Yadav and Yadav 2018). Wang et al. (2003) anticipated that the increased salinization of arable land would result in 50% land loss by the middle of the twenty-first century. The significance of soil salinity in agricultural yield is crucial (Tester and Davenport 2003) which negatively affect the establishment, growth, and development of plants, leading to huge losses in productivity (Mathur et al. 2007). Irrigated land is only 15% of the total cultivated land but has at least twice the productivity of rainfed and accounts for one-third of the world’s food supply (Munns 2005). The three main direct effects of salt on plant growth are (a) reduced osmotic potential of soil, causing a decrease in the amount of water availability to plant, leading to physiological drought; (b) toxicity of excessive Na+ and Cl− ions in the cell, causing disruption of enzyme structures and other macromolecules, damage to cell organelles and plasma membrane, as well as disruption of photosynthesis, respiration, and protein synthesis (Feng et al. 2002); and (c) nutrient imbalance in the plant caused by nutrient uptake leading to ion deficiencies (Adiku et  al. 2001). The plants must maintain lower internal osmotic potentials to prevent water movement from roots into the soil (Feng et al. 2002). The general response of plants to high soil salinity is reduced growth (Ghoulam and Foursy 2002). At low or moderate salt concentration, plants adjust osmotically and maintain the potential for water influx (Gunes et al. 1996). High salt concentrations disturb membrane integrity and interfere with internal solute balance and nutrient uptake, causing nutritional deficiency symptoms similar to drought (Grattan and Grieve 1999). Grattan and Grieve (1999) reported that high NaCl uptake competes with the uptake of other nutrient ions, such as K, Ca, N, and P, resulting in nutritional disorders and eventual reduction in yield and quality. In most cases, salinity decreases the concentration of P in plant tissues (Kaya et al. 2001). Osmotic stress, ion imbalances, and direct toxic effects of Na+ and Cl− ions on the metabolic processes are most important and widely studied physiological impairments caused

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by salt stress (Munns 2005). Cusido et al. (1987) revealed that salinity inhibits the plant growth by affecting water absorption, protein biosynthesis, and by N as well as CO2 assimilation. Certain halophytes are also known to remediate soil salinity. Gallagher (1985) and Glenn and O’Leary (1985) searched for new salt-tolerant crop plants that could deal with saline soils and minimize the crop loss. Development of salt-tolerant crop varieties through breeding programs is underway. To challenge the detrimental effects of salinity, a salt-tolerant perennial rye-grass (Lolium perenne) was obtained by transforming rice vacuolar membrane Na+/H+ antiporter gene via Agrobacterium-­ mediated transformation. The salt tolerance of perennial ryegrass was improved by overexpression of the Na+/H+ antiporter gene (Wu et al. 2005). Most of such successful methods are costly and beyond the economy of several developing nations (Cantrell and Linderman 2001). Hence, an alternative attempt is needed  to challenge the deleterious effects of saline soils by inoculating salt-tolerant AMF in the agricultural crops. External and internal microorganisms in their natural environment colonize plants. Under stress environment, plant performance could be improved by the deliberate introduction of some microorganisms in rhizosphere. The AMF association with plants is one such association that could mitigate several types of plant stress. The AMF is associated with the roots of over 80% terrestrial plant species including halophytes, hydrophytes, and xerophytes (Mathur et al. 1999; Yadav et al. 2019b, c; Hejiden et al. 1998). The AMF form a symbiotic relationship with the plants in which the plant supplies carbohydrate to the fungi while the mycorrhizal fungi extend the surface area of roots, thus increasing their ability to absorb nutrients and water from the soil. These fungi constitute an integral component of the natural ecosystem and are known to exist in saline environments (Giri et al. 2003). The AMF holds the potential to reduce the impact of salinity-induced stress on plants. The AMF associations often result in greater yield of crop plants even under saline conditions. The AM fungi widely exist in salt-affected soils (Juniper and Abbott 1993). To some extent, AM fungi are considered as bio-ameliorators of saline soils (Azcon-Aguilar and Barea 1997; Rao 1998). The AMF is increasingly being considered for mycoremediation of soil salinity. According to Kohler et al. (2009), the use of plant symbiotic microorganisms, especially AM fungi, is useful in developing strategies to facilitate plant growth in saline soils. The AMF maintains physiological and biochemical processes of the host plant. In salt-stressed soil, PO43− ions usually precipitate along with Ca2+, Mg2+, and Zn2+ and remain  less available to plants (Azcon-Aguilar et  al. 1979). However, AMF symbiosis in plants enhance the uptake of less mobile phosphorus by extending their external hyphal network beyond nutrient depletion zones. Apart from salt stress mitigation, AMF also improves plant growth and hormonal status, increases nutrient acquisition, maintains osmotic balance and reduces ion toxicity (Juniper and Abbott 1993). It also stabilizes soil for plant growth by producing glomalin, a substance that binds soil aggregates (Wright and Upadhyaya 1998). The purpose of this chapter is to outline the current state of knowledge on soil salinity mycoremediation. This chapter highlights interaction mechanisms between AMF and

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­ MF-­colonized plants in which AM fungi ameliorate the deleterious effects of A salinity. It includes a brief discussion on how this knowledge is currently being used for mycoremediation of soil salinity with future prospects.

13.2  M  orphological Features of AMF and Development of Fungal Symbiosis The AMF are soil fungi that form a mutualistic symbiosis with the plant roots. It is reported that AMF occurs in most of the plant species and taxonomically the symbiotic associations of AMF with the plants are widespread in nature (Schwab et al. 1991). The AMF occurrence is too diverse from geography and plant taxonomy point of view. However, it has become possible to define two morphological types, the Arum-type and the Paris-type. These types were named after the plants in which they were first described for Arum maculatum and Paris quadrifolia (Cavagnaro et al. 2001). Smith and Smith (1997) found that the morphology of the AMF symbiosis with the angiosperms depends on plant taxa (plant species). It was concluded that 30 families belong to Arum-type, 41 to Paris-type and 21 families form either an intermediate morphology or had members with both Arum and Paris-types. The plant largely controls AMF morphology through the presence or absence of extensive air-spaces in roots (Brundrett and Kendrick 1990). The general morphology of AMF can be differentiated into six different types: 1. The intracellular hyphae forming coils often found in the outer layers of cortical parenchyma 2. The intercellular hyphae 3. The intracellular hyphae with numerous ramifications, i.e., the arbuscules 4. The inter- or intracellular hypertrophied hyphae, i.e., the vesicles 5. The extracellular ramified hyphae, i.e., branched-absorbing structures (BAS) 6. The resistance propagules, i.e., the spores The AMF spores are capable of independent germination. However, AMF development is hindered for their inability to complete the life cycle in the absence of host and the fungus start producing vigorous mycelia. This is because the AM fungi coevolved along with host plants and require each other for mutual development (Remy et al. 1994; Redecker et al. 2000). The AMF has different survival strategies like regulation of infection structure (Giovannetti et al. 1994), the ability of multiple germinations (Koske 1981), induction of energy-saving mechanism of spores germinate in the absence of the host (Logi et al. 1998), etc. In addition, the ability to form wide hyphal networks by pre-symbiotic and symbiotic mycelia represents a fundamental mechanism for increasing the chances of AM symbionts to contact host roots (Giovannetti et al. 1999). The hyphae of AM fungal soil propagules and asexual spores or mycorrhizal roots begin the establishment of AMF symbiosis by colonizing on compatible plant

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roots. Even dead roots from annual plants might be a good source of AMF growth because roots protect the fungus from environmental hazards until new hyphae grow out of the roots and colonize plants (Requena et al. 1996). The hyphae of fungus penetrate the plant cortex through an aspersorium to form distinct morphologically special structures like inter- and intracellular hyphae, coils, and arbuscules. Out of these structures “arbuscules” are specialized hyphae similar to haustoria of plant pathogenic fungi. They are presumed to be the main site of nutrient exchange between the plant and the fungus (He and Nara 2007). After host colonization, the fungal mycelium grows out of the root, exploring the soil in search of mineral nutrients to colonize other susceptible roots (Breuninger and Requena 2004). The fungal life cycle completes after the formation of asexual chlamydospores on the external mycelium. The life cycle and different morphological parts of AMF are shown in Fig. 13.1.

Fig. 13.1  Showing the life cycle of AM fungi and different morphological parts of AMF. (From Porcel et al. 2012)

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13.3  Ecophysiology Mechanisms of AMF The interactions between plants and AMF affect the plant ecophysiology and have the following consequences on nutrition, growth, competition, stress tolerance, fitness, and soil structuring: 1 . Enhanced uptake of low mobile ions 2. Improved quality of soil structure 3. Enhanced plant community diversity 4. Improved rooting and plant establishment 5. Improved soil nutrient cycling 6. Enhanced plant tolerance to biotic and abiotic stress

13.3.1  Development of Extraradical AMF Networks A network of AM fungal hyphae is present in almost every terrestrial ecosystem but our knowledge lacks the proper understanding of morphology and development of AM fungal network. The reason for this knowledge gap lies in methodological difficulties to extract and observe AM mycelia from the soil. The common mycelia extraction method includes blending, wet sieving, and decanting of soil samples (Jakobsen et al. 1992). The soils rich in organic or porous particles may not yield the complete mycelium. Nevertheless, artificial growth media like sand or glass bead supplied with nutrient solution are frequently used as substrates for AMF growth (Chen et al. 2001). These nutrient solutions allow the extraction and analysis of AM mycelium samples. However, but it cannot be neglected that morphology and activities of AM hyphae in nutrient solution differ considerably from those in the soil. Studies of extraradical AMF development on employed axenic cultures of AM fungi with Ri T-DNA-transformed roots have been done (de Souza and Declerck 2003). A substrate consisting of 40  μm wet sieved soil and glass beads, which allows for the complete extraction of intact AM mycelium, was proposed as a growth substrate for AM mycelia (Neumann and George 2005). This substrate may reproduce soil conditions better than previously used artificial growth substrates. The AM mycelia are made of morphologically different hyphal types. The coarse and thick-walled hyphae with a diameter between 5 and 20 μm that appear to function mainly in nutrient transport and extension of the fungal colony at rates of 1–2 mm per day (Olsson and Wilhelmsson 2000) are called “runner-hyphae” that elongate and form hyphopodia parallel to the plant root axis (Friese and Allen 1991). The runner-hyphae extend radially around the root and can reach distance up to 24  cm (Drew et  al. 2006). They may rapidly spread AM infection over non-­ colonized neighboring roots of the same or different host plants (Drew et al. 2006; Voets et al. 2009).

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The AM hyphae bridges between neighboring roots and may be formed by hyphal anastomosis of the same species (Croll et al. 2009). The coarse extraradical AM hyphae form clusters of fine branched, thin-walled structures with a diameter below 4 μm (Bago 2000). When AMF infection is watched on Petri plates, it resembled in their morphology with 5–7 days old intraradical arbuscules (Cano 2005), which show high metabolic activity (Bago et al. 2002). Since intra-radical arbuscules are presumably the major sites of AMF nutrient uptake from the soil, therefore called branched absorbing structures (BAS).

13.4  E  ffect of Soil Salinity on Plants and Their Self-Defense Mechanisms Salinity represents the concentration of dissolved mineral salts present in the soil and water. The dissolved mineral salts consist of the electrolytes of several types of cations and anions. The establishment, growth, and development of plants are affected by soil salinity that causes a huge loss in the productivity of the plants (Evelin et al. 2009; Yadav et al. 2017a, b). Due to the soil salinity, plants undergo some major physiological stresses that affect the plant system very badly. The toxic effect of ions, mostly Na+ and Cl−, damage the structure of enzymes and other macromolecules as well as cell organelles, disrupt photosynthesis and respiration, inhibit protein synthesis and induce ion deficiencies (Juniper and Abbott 1993). The high salt concentration induces physiological drought condition in plants. The plant root cells are osmotically sensitive and under normal conditions they must maintain lower internal osmotic potentials to prevent water from moving into the soil. In addition, soil salinity creates a nutrient imbalance in the plant caused by decreased nutrient transport to the shoot (Adiku et al. 2001). Consequently, it affects all the systemic process of plants such as growth, photosynthesis, protein synthesis, energy, and lipid metabolism (Ramoliya et al. 2004). The plant cell membranes have receptors for perceiving stress signal to activate production of few secondary signal molecules such as Ca2+, inositol phosphates, reactive oxygen species (ROS), and abscisic acid (ABA). The stress signals are transduced inside the nucleus where they induce multiple stress-responsive genes and the products of genes to induce  plant adaptation to salinity. Plants start expressing early genes within minutes of stress perception, and their products (e.g., various transcription factors) can activate the expression of delayed genes (e.g., RD gene [responsive to dehydration], KIN gene [cold induced], COR gene [cold responsive]). Such gene products are either directly involved in cellular protection against the stress (e.g., late embryogenesis abundant proteins, antifreeze proteins, antioxidants, chaperons, and detoxification enzymes) or indirectly (e.g., transcription factor and enzymes of phosphatidylinositol metabolism) (Tuteja 2007) protect the plant.

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13.5  Effect of Soil Salinity on AMF Soil salinity negatively impacts the host plant and associated AMF by inhibiting colonization capacity, spore germination, and growth of fungal hyphae. It was reported that AMF colonization on plant roots is reduced in the presence of NaCl (Ojala et al. 1983; Poss et al. 1985; Duke et al. 1986; Rozema et al. 1986; Menconi et al. 1995; Juniper and Abbott 2006; Giri et al. 2007; Sheng et al. 2008). The higher NaCl concentrations directly affect fungal growth and metabolism (Juniper and Abbott 2006) and could suppress the formation of AM (Tian et al. 2004; Sheng et al. 2008). The variable behavior of AM fungi in a similar ecosystem could be associated with soil salinity (Kilironomos et al. 1993) and some other environmental factors (Carvalho et al. 2001). In the presence of higher concentrations NaCl, the germination of spores is delayed (Cantrell and Linderman 2001). The rate of germination and maximum germination of AMF spores may also depend on the salt type. According to Juniper and Abbott (1993), the sodium salts like NaNO3 and Na2SO4 with osmotic potentials similar to NaCl impart differential effects on the rate and germination of spores. Jahromi et al. (2008) studied in vitro effects of salinity on the AM fungus, Glomus intraradices and observed that there was no significant difference in hyphal length and BAS between control (no salt) and 50 mM NaCl, but a significant decrease in hyphal length and the number of BAS at 100 mM NaCl was observed. All these results indicate that soil salinity directly affects fungal development by reducing fungal mycelia formation and host root colonization. However, contrary to the mentioned reports, increased AMF sporulation and colonization under salt stress were reported by Aliasgharzadeh et al. (2001). Yamato et al. (2008) reported that AM colonization rates were not reduced in coastal vegetations of Okinawa Island, Japan even when treated with salinity as high as 200 mM of NaCl. The variation in the results invites researchers to look out for salt-tolerant AMF species and to test mycorrhizal  isolates that could  maintain colonization capacity and symbiosis efficiency under salinity.

13.6  Effect of AMF on Plant Biomass and Nutrient Uptake For remediating the harmful effect of salinity, the AMF colonization plays a very crucial role in plants. The AMF protects the symbiotic plants against salt stress. It was demonstrated that the symbiosis results in increased uptake of nutrient, accumulation of osmoregulator compounds, increase in photosynthetic rate, and water use efficiency (Andrea et al. 2016). It was suggested that salt stress remediation by AMF results from a combination of nutritional, biochemical, physiological, and molecular effects (Atakan et al. 2018). However, remediation effect on plant development depends on the AMF species (Marulanda et al. 2003, 2007; Wu et al. 2007). Several workers reported that AMF-inoculated plants grow better than

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non-inoculated ones under salt stress (Al-Karaki 2000; Giri et al. 2007; Sannazzaro et al. 2007; Zuccarini and Okurowska 2008). Hajiboland et al. (2010) reported that although high salinity reduces dry matter production of tomato cultivars; however, in all treatments, mycorrhizal plants grow better by improving plant nutrient uptake, especially P (Smith and Read 1997). Many other workers related improved growth of mycorrhizal plants in saline conditions with the mycorrhiza-mediated enhancement of P nutrition in plants (Hirrel and Gerdemann 1980; Ojala et al. 1983; Pond et  al. 1984; Poss et  al. 1985; Al-Karaki 2000; Verma et  al. 2017; Yadav et  al. 2018; Kour et al. 2019; Rana et al. 2019a, b).

13.6.1  I ncreased Nutrient Uptake and Transport by AM Hyphae The enhanced growth of AM plants with improved P uptake is a common observation in greenhouse studies, particularly when plants are grown on soil with low nutrient availability. Investigations on the uptake, transport, and transfer of mineral elements by AMF mycelia to host plants were done under in vivo or in vitro microcosms. Wang et al. (2002) studied the uptake, transport, and delivery of P and N of plant-AM fungal combinations under varied environmental conditions. In many studies, uptake and transport of Zn (Jansa et al. 2003), S (Allen and Shachar-Hill 2009), Fe (Caris et al. 1998), and Cu (Lee and George 2005) were verified via AM pathway. However, the transport of nutrients other than P, N, Zn, Fe, and S through AM mycelium has not been unequivocally verified. The AM symbiosis shows increased tolerance to biotic and abiotic stress. Several studies of plant-AM symbiosis confer host tolerance to drought (Ruiz-Lozano 2003; Miransari 2010), heat (Compant et al. 2010), salinity (Evelin et al. 2009; Miransari 2010), or osmotic stress (Ruiz-Lozano 2003). Nevertheless, the early works on biotic stress of mycorrhizae were mostly descriptive (Linderman 2000). The AM symbiosis occurs in almost all the habitats including disturbed soils contaminated with heavy metals and plays an important role in improving plant metal tolerance. Several AM-mediated mechanisms of plant protection include dilution of plant tissue toxins, sequestration of the toxic metals in the fungus, and tolerance development by the fungi (Hildebrandt et al. 2007; Gamalero et al. 2009). Thus, it can be concluded that some plants are unable to endure habitat-imposed abiotic and biotic stresses in the absence of fungal endophytes. The AM colonization of various plants improve salt tolerance and growth. However, relatively few mechanisms demonstrate the increased tolerance of AM plants to salt stress (Cho et  al. 2006). The improved growth of AMF-colonized plants was shown to enhance nutrient uptake, particularly N and P (Jeffries et al. 2003). In one study, salt tolerance was not found correlated with P concentration (Ruiz-Lozano and Azcon 2000). According to Auge (2001), the contribution of AM symbiosis to the salinity remediation of host plants can explain the salt tolerance

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mechanisms such as enhanced osmotic adjustment, leaf hydration, increased water use efficiency, reduced oxidative stress, and improved nutritional status. Due to soil salinity, the water uptake ability of plants is inhibited leading to the slower growth, causing osmotic or water-deficit effect (Munns 2005). This effect was reported in wild and cultivated Phaseolus sp. where leaf water, osmotic and turgor potentials of the plants decreased due to the salinity (Jimenez et  al. 2003). Rosendahl and Rosendahl (1991) reported improved water uptake by AM plants under salinity. Kumar et al. (2009) reported that relative water content in the leaves of Jatropha sp. was significantly higher in AM-inoculated plants growing under salinity. Under salinity stress, the beneficial effects of AMF on plant growth may occur due to improved water absorption, nutrient uptake and enhanced photosynthesis (Miransari et al. 2008). Decreased K+ concentration and K+/Na+ ratios at high salinity is another harmful effect of salinity that disturbs the K+-specific plant functions (Tabatabaei 2006). Generally, K+ is a preferred ion of cytoplasm as it provides a reactive environment for cellular enzymes. In addition, K+ is also an osmotic equivalent under water stress and salinity. However, under high salinity, the Na+ spontaneously enters plant cells via nonselective K+ channels (Hammer et al. 2010). The spontaneous accumulation of Na+ can be prevented and K+ uptake could be improved by AMF inoculation on plants. Many studies show decreased Na+ accumulation under salt stress in Sesbania spp. (Giri and Mukerji 2004) and Lotus glaber (Sannazzaro et al. 2006) due to AM association. In addition, K+ uptake is also enhanced by the influence of mycorrhiza (Rabie and Almadini 2005). Interactions of Na+, Cl−, and many mineral nutrients affect the plant growth, causing an imbalance in nutrient availability, uptake, or distribution within plants, thus increasing plant’s requirement for essential elements (Grattan and Grieve 1992). Many reports show lowered Na+ concentrations in AMF plants growing under salinity (Ashraf et al. 2004; Ghazi and Al-Karaki 2006; Sharifi et al. 2007; Kaya et al. 2009; Porras-Soriano et al. 2009). The growth and metabolism of plants are affected by soil salinity, N availability, and other environmental factors (Shenker et al. 2003). The P availability and activity also reduces in saline soil (Grattan and Grieve 1999). However, plants colonized by Glomus sp. show higher N and P contents in shoots of lettuce at the highest salt level than non-colonized ones at the lowest salt level (RuizLozano and Azcon 2000). It was reported that the efficiency of P uptake by an AMF is strongly affected by the spatial distribution of its hyphae in the soil and possibly also by the differences in the capacity for uptake per unit length of hyphae (Jakobsen et al. 1992). Mycorrhizal inoculation improves P nutrition of plants under salinity stress and reduces the negative effects of Na+ by maintaining vacuolar membrane integrity, which prevents this ion from interfering in growth metabolic pathways (Rinaldelli and Mancuso 1996). It has been observed that the chief mechanism for enhanced salinity tolerance in the mycorrhizal plant is  actioned by improving P availability (Copeman et al. 1996). The AMF plants increase P uptake in compensation of salt stress (Tian et al. 2004). A symbiotic association of specific mycorrhizal fungi with plant species, and also the changes made by the AMF in plants for the amelioration of soil salinity are shown in Table 13.1.

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Table 13.1  Showing the effect of different AMF on enhancement tolerance of several plants growing under salt stress Plant species Glycine max

AMF species Glomus etunicatum

Gossypium arboretum Lycoperssicon Esculentum

Glomus mosseae Glomus mosseae

Acacia nilotica

Glomus fasciculatum

Acacia auriculiformis

Glomus fasciculatum and Glomus macrocarpum Mixed inoculum of Glomus sp. and Gigaspora sp. Glomus mosseae

Effect of AMF on plant Increase the uptake of P, K, Ca, and Zn Increase the uptake of P, Na, and Cl Increase the uptake of P and Na and decreases the uptake of Cu and Zn Increase the uptake of Cu, Na, and P Increase the uptake of P and Na

References Sharifi et al. (2007) Tian et al. (2004) Al-Karaki (2000)

Giri et al. (2007) Giri et al. (2003)

Increase the uptake of N and P

Murkute et al. (2006) Feng et al. (2002)

Glomus macrocarpum

Increase the uptake of P and increases soluble sugar and electrolyte concentrations Increase the uptake of N and Mg

Glomus macrocarpum

Increase the uptake of N and Mg

Musa sp.

Glomus intraradices

Increases the uptake of K and Cl

Lactuca sativa

Glomus clarum

Increase the uptake of K

Lactuca sativa

Glomus sp.

Increases transpiration, stomatal conductance, and water use efficiency

Citrus karma

Zea mays

Sesbania aegyptiaca Ocimum basilicum

Giri and Mukerji (2004) Zuccarini and Okurowska (2008) Yano-Melo et al. (1999) Ruız-Lozano et al. (1996) Ruız-Lozano et al. (1996)

13.7  Mechanism of Remediation of Salinity Stress by AMF 13.7.1  Biochemical Change The strategies adopted by the plant to remediate soil salinity include (1) synthesis and accumulation of compatible solutes, (2) control of ion uptake by roots and transport to plant tissues for ion homeostasis, (3) fine regulation of water uptake and distribution to plant tissues by the action of aquaporins, and (4) reduction of oxidative damage through improved antioxidant capacity. Additional plant responses include selective build-up or exclusion of salt ions, maintenance of photosynthesis at values adequate for plant growth, changes in membrane structure and synthesis of phytohormones (Turkan and Demiral 2009). Many studies were done to understand

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mechanisms for enhanced salt tolerance of AMF plants (Al-Garni 2006). The AMF facilitate plants by enhancing nutrient mobility, improving water uptake through extended fungal hyphae that act as root hairs, boosting root hydraulic conductivity, and by maintaining osmotic balance by enhancing K+/Na+ ratios and lowering accumulation of Na+ in the host plant shoots. Among the mechanisms, the best characterized biochemical response of plant cells is osmotic stress in which accumulated inorganic ions like Na+ and compatible organic solutes like proline, glycine betaine, and soluble sugars were studied (Flowers and Colmer 2008). The compatible solutes can accumulate to high levels in the cells without disturbing intracellular biochemistry (Bohnert and Jensen 1996) and, consequently, protect subcellular structures, mitigate oxidative damage caused by free radicals, and maintain the enzyme activities under salt stress (Yokoi et al. 2002). 13.7.1.1  O  smotic Adjustment with the Help of AMF for Remediation of Soil Salinity The salt-accumulated soil has negative water potential, which implies less availability of soil water to the plant leading to the dehydration of the plant cells. Therefore, plants must respond accordingly by decreasing their water potential to maintain a favorable concentration gradient. Osmotic adjustment or osmoregulation is the most important mechanism adopted by plants to reduce osmotic potential in which some inorganic ions like Na+ and K+ and compatible organic solutes accumulate in the cells (Morgan 1984; Hoekstra et al. 2001). Among them, proline, betaines, sugars (fructose, sucrose, and glucose), and complex sugars (trehalose, raffinose, and fructans) are some compatible solutes that  chiefly accomplish this function in halophytes. The two major osmoprotectant osmolytes include proline and glycine betaine (N, N, N-trimethylglycine betaine) which are synthesized by plants in response to stress. These osmoprotectants help in maintaining the osmotic status of the cell to ameliorate abiotic stress. Proline plays a role in scavenging free radicals, stabilizing subcellular structures, and buffering cellular redox potential under stresses. The salinity stress-responsive genes whose promoters contain proline-­responsive elements (ACTCAT) are also induced by proline (Chinnusamy et al. 2005). Proline is an important osmoprotectant that plays a vital role in stabilizing subcellular structures, buffering cellular redox potential under stress and in scavenging free radicals (Chen and Dickman 2005). In higher plants, proline is synthesized from glutamic acid by the actions of two enzymes, pyrroline-5-­ carboxylate synthetase (P5CS) and pyrroline-5-carboxylate reductase (P5CR). Overexpression of P5CS gene in transgenic tobacco increases the production of proline that helps in salinity tolerance (Kishor et al. 1995). Proline concentration also increases during AMF colonization on plants. Several authors reported higher proline concentration in AMF plants than in non-AMF plants at different salinity levels (Sharifi et al. 2007; Talaat and Shawky 2011). In addition, another osmoregulator named betaines also stabilize the structures and activities of enzymes, protein complexes and maintain the integrity of membranes against the damaging effects of

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excessive salt (Gorham 1995). The AMF plants increase the accumulation of betaines under salt stress (Al-Garni 2006). The increase in sugar content is found to be positively correlated with the mycorrhization of the host plant as reported by Thomson et al. (1990). Porcel and Ruiz-­ Lozano (2004) reported increased sugar concentrations in soybean roots colonized with Glomus intraradices subjected to drought stress. The positive correlation between sugar content and mycorrhization is due to the sink effect of the fungus demanding sugars from the shoot tissues (Augé 2000). The increased sugar accumulation in plants may also be due to AMF-facilitated starch hydrolysis. However, some authors reported negative correlations between AMF colonization and sugar accumulation in host plants. Pearson and Schweiger (1993) reported a reduction in carbohydrate content with an increase in the percentage of root colonization. Sharifi et  al. (2007) observed no role of soluble carbohydrates in the responses of AM (colonized by Glomus etunicatum) soybean plants to salinity. 13.7.1.1.1  Polyamines The polyamines are small organic cations necessary for eukaryotic cell growth. The three main polyamine types existing in plants are putrescine (Put), spermidine (Spd), and spermine (Spm). The plant-secreted polyamines play a major role against a wide array of environmental stresses like salinity (Delauney and Verma 1993), high osmolarity (Besford et al. 1993), and antioxidative stress (Kurepa et al. 1998). They are also candidates for the regulation of root development under saline situations (Couee et al. 2004). Under salinity stress plants reduce the concentration of free polyamine. The polyamine increases membrane stability and show a significant effect on H+/ATPase and Ca2+/ATPase transporters during salinity stress (Pottosin and Shabala 2014). 13.7.1.2  P  hytohormone Content Regulation by AMF for Remediation of Salinity Many phytohormones regulate cell division, plant growth, fruit ripening, stem elongation, and other biochemical function. Out of these, ABA responds against abiotic stress, and its concentration rise when plants experience stress. The ABA promotes stomatal closure to reduce water loss. It also induces the expression of stress-related genes and diminishes the environmental damage (Evelin et  al. 2009). A report showed that AMF plants change the ABA concentration under the saline condition (Estrada-Luna and Davies 2003). It was observed that ABA concentration in AMF plants decreased in lettuce plants colonized by G. intraradices (Jahromi et al. 2008). The AMF-infected plants experience less environmental stress and, as a result, accumulate less ABA. Evelin et al. (2009) observed the effect of AMF species on ABA content varies with the host plants. As AM fungi help in remediating the salinity effect, therefore could be used as agents for salinity-induced remediation.

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13.7.1.3  Antioxidant System A number of biotic, abiotic, and xenobiotic stresses can be linked to some degenerative reactions that produce the ROS such as superoxide (O2−), hydroxyl radicals (OH−), hydrogen peroxide (H2O2), and singlet oxygen in plants. These are produced as reaction by-products of electron transport chains of chloroplast and mitochondria where oxygen and electrons react to form ROS (Scandalios 1993). The plant ROS production must be minimized because hydroxyl radical and singlet oxygen are highly reactive and damage life-giving biomolecules (Jakob and Heber 1996). The O2− and H2O2 are especially toxic ROS compounds that initiate cascade reactions, resulting in the production of the hydroxyl radicals. According to Bowler et  al. (1992), free radicals and their derivatives are dangerous and reactive and could damage biomolecules through lipid peroxidation, denaturation of proteins, and DNA mutation. Plant cells contain an array of protective and repair mechanisms that minimize oxidative damage. According to Smirnoff (1993), the repair mechanisms can be divided into two categories: (a) systems that scavenge activated oxygen species which include carotenoids, glutathione, tocopherols, ascorbic acid, flavonoids, and anthocyanins (Wu and Xia 2006) and (b) systems which possess ROS-scavenging antioxidative enzymes like superoxide dismutase (SOD), catalase (CAT), ascorbate peroxidase (APOX), glutathione reductase, dehydroascorbate reductase, monodehydroascorbate reductase, guaiacol peroxidase, oxidized glutathione, glutathione peroxidase, and the enzymes involved in the ascorbate-glutathione cycle (Alguacil et al. 2003). Like other abiotic stresses, salinity also induces oxidative stress in plants (Hajiboland and Joudmand 2009). A correlation between antioxidant capacity and salinity tolerance was reported in several plant species (Nunez et al. 2003; Turkan and Demiral 2009). The reports suggest that AMF symbiosis helps plants to remediate salt stress by enhancing the activities of antioxidant enzymes like superoxide dismutase (SOD), catalase (CAT), ascorbate peroxidase (APOX), glutathione reductase, dehydroascorbate reductase, monodehydroascorbate reductase, and guaiacol peroxidase (Porcel et al. 2003; Garg and Manchanda 2009; Talaat and Shawky 2011).

13.7.2  Role of AMF in Physiological Changes Salt stress disrupts plant physiological mechanisms by decreasing photosynthetic efficiency, gas exchangeability, water status, and membrane integrity (Aroca et al. 2006; Porcel et al. 2006). Several pieces of evidence demonstrate that AMF symbiosis can alleviate salt-­ stress effects by employing several mechanisms that are discussed below.

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13.7.2.1  Chlorophyll Content Salt stress inhibits photosynthetic ability and induces physiological drought in plants, causing decreased crop production. The soil salinity was reported to reduce plant chlorophyll content (Sheng et al. 2008). The salinity-induced reduction in chlorophyll content could be due to suppression of enzymes responsible for the synthesis of photosynthetic pigments (Murkute et al., 2006). Under salinity the plant uptake of Mg2+ decreases, which is needed for chlorophyll biosynthesis. The reduction in Mg2+ concentration lowers leaf chlorophyll concentration (El-Desouky and Atawia 1998). However, many studies report higher chlorophyll content in leaves of AM plants facing saline stress (Sannazzaro et al. 2006; Zuccarini 2007; Colla et al. 2008; Sheng et al. 2008). This suggests lower salt interference in chlorophyll biosynthesis in mycorrhizal than in non-mycorrhizal plants (Giri and Mukerji 2004). According to Giri et al. (2003), the antagonistic effect of Na+ on Mg2+ uptake is counterbalanced and suppressed by the presence of mycorrhiza. By chlorophyll content measurement, it was estimated that the photosynthetic capacity of salt-stress AMF plant is superior to the non-stressed plants, which implies that mycorrhization of the plants can fully counterbalance the salt stress (Zuccarini 2007). 13.7.2.2  Relative Permeability The AMF inoculation of host plants enable them to maintain higher electrolyte concentration that improves the integrity and stability of the cell membrane. Consequently, the electrical conductivity of mycorrhizal roots was found to be higher than the non-mycorrhizal roots (Garg and Manchanda 2008). The mycorrhizal roots of Cajanus cajan showed higher relative permeability than non-­ mycorrhizal plants at different levels of soil salinity (Garg and Manchanda 2008). Kaya et al. (2009) reported the electrolyte leakage of 31.66 and 42.45, respectively, in leaves of Capsicum annum treated with 50 mM and 100 mM NaCl, while the AMF-inoculated plants had a relatively lower electrolyte leakage of 26.87 and 30.98, respectively. This suggests that AM plants have lower root plasma membrane electrolyte permeability than the non-mycorrhizal plants. The increased membrane stability has been attributed to mycorrhiza-mediated enhanced P uptake and increased antioxidant production (Feng et al. 2002). 13.7.2.3  Water Status The water is essentially needed to mobilize plant nutrients. In saline soils, Na+ and Cl− ions bind with water and create a physiological drought-like condition for plants (Fuzy et al. 2008). Several studies indicate that AMF colonization can help plants to remediate the physiological drought. It has been reported that plants inoculated with AMF maintain relatively higher water content

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(Colla et al. 2008; Jahromi et al. 2008; Sheng et al. 2008). This is facilitated by the improved hydraulic conductivity of the root at low water potential (Kapoor et al. 2008). The improved root conductance is associated with a longer root and an altered root system morphology induced by AMF (Kothari et al. 1990). The AMF plants exhibit a higher stomatal conductance to increased leaf transpiration (Jahromi et  al. 2008; Sheng et  al. 2008). Mycorrhizal plants hold lower osmotic potential that is maintained by fungal accumulating solutes, resulting in improved plant osmotic adjustment. Lower water saturation deficit and higher turgor potential of AMF plants also improve their water status (Al-Garni 2006). All the improved parameters facilitated by mycorrhizal colonization enable the host to utilize water more efficiently and allow them to maintain a lower intercellular carbon dioxide concentration. As a consequence, the gas exchange capacity increases in the mycorrhizal plants. 13.7.2.4  Nodulation and Nitrogen Fixation Nitrogen-fixing bacteria-mediated nodule formation in plants is considered a target for salt stress, and its occurrence decreases due to salt  accumulation (Harisnaut et al. 2003). This is likely due to premature nodule senescence triggered by salt stress (Gonzalez et al. 1998), which causes accelerated lytic activities, formation of green pigments from leghemoglobin, and loss of nitrogen fixation (Delgado et al. 1994). Application of AMF in legumes could counteract the harmful effects of salinity on nodulation and nitrogen fixation. The AMF symbiosis could alleviate drought stress-induced premature nodule senescence (Ruiz-Lozano et al. 2001). Giri and Mukerji (2004) reported a positive effect of mycorrhizal inoculation on nodule formation under salt stress. The legume colonization by AMF could increase the number of nodules (Garg and Manchanda 2008). This indicates a positive influence of AMF on legume–nitrogen-fixing bacteria symbiosis due to enhanced leghemoglobin content, which is determined by observing the nodule color change from pink to brownish pink due to the synthesis of green pigments by leghemoglobin. The greening of the nodule was observed much earlier in AMF-colonized legume plants (Garg and Manchanda 2008). In addition, mycorrhizal plants hold higher nitrogenase activity. All these parameters contribute to the higher nitrogen-fixing ability of AMF plants. The increased nitrogenase activity and nitrogen fixation in AMFcolonized legume plants is beneficial for nitrogenase enzyme functioning possibly through enhanced uptake of essential micronutrients, resulting in improved plant growth (Founoune et al. 2002). Therefore, it may be concluded that mycorrhizal and nodule symbioses often act synergistically on mineral nutrition, and plant growth to fulfill the need of N and P and to increase tolerance of plants to salinity (Rabie and Almadini 2005).

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13.7.3  Molecular Changes in Plants The beneficial effects of the AMF symbiosis on plant salinity tolerance are assessed by measuring plant growth, water status, or the expression of plant-specific stress-­ related genes. The effects of AMF on plant responses to salinity stress are difficult to measure due to the complex nature of the association. The heterokaryotic and obligate nature of AMF also creates difficulty in understanding plant molecular changes. However, molecular studies on salt tolerance by AMF are speeding up for holistically understanding the mechanism of salt stress remediation by AM symbiosis. The expression and overexpression studies of proteins like Δ1-pyrroline-5-­ carboxylate synthetase (P5CS), aquaporins, cation channels and transporters, late embryogenesis abundant protein (LEA), and ABA synthesis in AMF plants treated with salinity stress are underway (Hanin et al. 2011). 13.7.3.1  1-Pyrroline-5-Carboxylate Synthetase (P5CS) The PC5S enzyme catalyzes the rate-limiting step in the biosynthesis of proline. Proline is the widespread osmoprotectant in plants. The PC5S has two catalytic sites: (1) γ-glutamyl-kinase and (2) glutamic-γ-semialdehyde-dehydrogenase. The P5CS catalyzes the first two steps of proline biosynthesis from glutamate using γ-glutamyl-kinase and glutamic-γ-semialdehyde-dehydrogenase activities. The last step of proline biosynthesis is catalyzed by the Δ1-pyrroline-5-carboxylate reductase (P5CR) that reduces the Δ1-pyrroline-5-carboxylate to proline (Hu et al. 1992). Yoshiba et al. (1995) observed that the P5CS-encoding gene is induced by drought stress, salinity, and by ABA treatment in Arabidopsis thaliana. The P5CS-encoding gene is of key importance for the biosynthesis of proline in plants (Abraham et al. 2003). The overexpression of the P5CS-encoding gene in transgenic tobacco plants increase proline production, which confers plant tolerance to osmotic stress (Kishor et al. 1995). Jahromi et al. (2008) reported the higher expression of Lactuca sativa P5CS gene in non-AMF plants at 50 mM NaCl and 100 mM. 13.7.3.2  Aquaporins Aquaporins belong to the major intrinsic protein (MIP) family of transmembrane channels that facilitate and regulate the passive movement of water molecules following a water potential gradient (Hill et al. 2004). In plants, aquaporins are subdivided into five evolutionarily distinct subfamilies: (1) the plasma membrane intrinsic proteins (PIPs), (2) the tonoplast intrinsic proteins (TIPs), (3) the small basic intrinsic proteins (SIPs), (4) the nodulin-like intrinsic proteins (NIPs), and (5) the uncharacterized X intrinsic proteins (XIPs) (Gupta and Sankararamakrishnan 2009), which were shown to transport a variety of uncharged substrates (Bienert et al. 2011). The role of aquaporin in water uptake was confirmed by inhibiting root water transport by the

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general aquaporin blocker mercury ions (Maggio and Joly 1995). In general, AMF differentially exerts control over the expression of different members of the large family of aquaporins (Ouziad et al. 2006). The AM-plant symbiosis enhances the expression of PIP genes, and its product contributes in regulating root water permeability to tolerate salinity-induced osmotic stress (Aroca et al. 2007). Such differences could be a consequence of the mode of salt stress in plants. Sarda et al. (1999) observed complex and differential expression pattern members of the large family of aquaporins in tested plant species. This highlights the complex regulation of aquaporin genes in response to the AMF symbiosis under abiotic stress with osmotic component. 13.7.3.3  Late Embryogenesis Abundant Proteins Close (1996) reported the accumulation of proteins groups called late embryogenesis abundant (LEA) proteins in mature plant seeds, responsible for desiccation tolerance. During cellular dehydration, the LEA proteins maintain the structure of other proteins, vesicles, or endomembrane constituents required for the sequestration. In addition, the LEA proteins help in absorbing water and also function as molecular chaperones (Koag et al. 2003). The reports show that overexpression of LEA proteins in plants and yeasts confer tolerance to osmotic stress (Imai et al. 1996; Babu et al. 2004). Dehydrins belong to LEA group 2 and represent the most noticeable soluble proteins induced by dehydration stress. Biochemically, the activities of dehydrins are diverse because they have multiple existences on the target sites (euchromatin, cytosol, and cytoskeleton). Studies show increased gene expression of the LEA genes during AMF symbiosis under salt stress. However, LEA gene expression and its effect on AMF symbiosis under salt stress is not fully understood. Porcel et  al. (2005) cloned two dehydrin encoding genes from Glycine max (gmlea8 and gmlea10) and one from L. sativa (lslea1) to analyze their response against drought stress of AMFinfected soybean and lettuce plants. Results demonstrate that the levels of LEA transcript accumulation in soybean and lettuce plants colonized by G. mosseae or G. intraradices were considerably lower than corresponding non-AMF plants. It is suggested that the accumulation of LEA proteins is not a mechanism by which the AMF symbiosis protects their host plant. The results suggest that mycorrhizal plants were less strained by drought stress through primary drought-­avoidance mechanisms. Jahromi et al. (2008) reported that LEA is expressed under salt stress and is induced at lower rates in AMF plants.

13.7.4  Cation Channels and Transporters 13.7.4.1  Na+/H+ Antiporters In soil the sodium uptake and distribution within the plant is responsible for their sensitiveness to salts. Plants survive high salinity by restricting root entry of Na+ and by allocating Na+ within the leaf to sequester into the vacuole. The Na+/H+

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antiporters mediate Na+ transfer out of cytoplasm into vacuole or apoplast. Transgenic plants were reported to be more salt tolerant than the controls as shown for Arabidopsis sp. (Sottosanto et al. 2004) and rice (Fukuda et al. 1999) due to overexpression of Na+/H+ antiporters. In Arabidopsis sp., the Na+/H+ antiporters are found at various locations. The transporters contributing to Na+ homeostasis include plasma membrane (SOS1) antiporter, vacuolar Na+/H+ antiporters (e.g., NHX1), and the plasma membrane uniporter HKT1 (Zhu 2003). The loss-of-function mutations in AtHKT1 render plants Na+ hypersensitive and disturb the distribution of Na+ between roots and shoot. Ouziad et  al. (2006) analyzed the expression of two Na+/H+ antiporter genes in relation to salt and mycorrhizal colonization. They observed no significant mycorrhization alterations through the expression of LeNHX1 and LeNHX2, the latter of which had previously been shown as K+/H+ antiporter (Venema et al. 2003). It is clear from the presented studies that AMF colonization does not assist Na+/H+ antiporters for balancing Na+ concentration. 13.7.4.2  Cyclic Nucleotide-Gated Channels Ion influx is essential for signal transduction. Cyclic nucleotide-gated ion channels (CNGCs) are one of the potential pathways for ion uptake (Talke et  al. 2003). Cyclic nucleotides monophosphate (CNMP) has only recently been accepted as an important secondary messenger that might be involved in plant response to biotic and abiotic stress. Donaldson et al. (2004) demonstrated that rapid increase in cyclic guanosine monophosphate (cGMP) levels in Arabidopsis thaliana are related to salt and osmotic stress, and these findings are consistent with another finding according to which cAMP and cGMP improve tolerance to salt stress (Maathuis and Sanders 2001). Interestingly, improved salt tolerance is correlated with the cNMP-­dependent decrease of channel opening probability and the reduced influx of Na+ (Rubio et al. 2003). Members of the CNGC family belong to the group of nonselective cation channels which enable the uptake of Na+, K+, and Ca2+ (Kaplan et al. 2007). Plants hold a structure where CNGC and cyclic nucleotide-binding domains occur in the cytosolic C-terminus of overlapping regions. The Arabidopsis CNGC gene family is made of 20 members (Maser et al. 2001). The ability of CNGCs to transport cations play a role in mediating various environmental stresses, plant defense responses, and development (Borsics et al. 2007). Kugler et al. (2009) reported upregulation of AtCNGC19 and AtCNGC20  in Arabidopsis thaliana shoots toward elevated NaCl. Salt induction of CNGC20 was also observed in the shoot. No differences in K+ and Na+ contents of the shoots were measured in homozygous T-DNA insertion lines for CNGC19 and CNGC20, respectively, which developed a growth phenotype in the presence of up to 75 mM NaCl similar to that of the wild type. All these results suggest that CNGC19 and CNGC20 channels are involved in salinity response of different cell types in the shoot. Both the channels assist the plant to survive toxic effects caused by salt stress probably by real-

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locating sodium within the plant. For obtaining a wider picture these studies should be accomplished in combination with measurements of sodium and potassium content in different plant tissues. This will allow us to understand the effect of AMF symbiosis on Na+ and K+ uptake, distribution, and compartmentalization within the plant cell and may shed light on the mechanisms involved.

13.8  Conclusion and Perspectives for Future Studies It is well-recognized fact that halophytes conveniently grow in saline soil. Now it is known that AM-inoculated plants can also grow under salinity. The AM fungi do not completely remove the salt from the local environment. However, it has the potential to alleviate the effect of soil salinity on plants. Several counter-balancing processes of AMF symbiosis help in alleviating the effect of soil salinity. In the saline soil, AMF-mediated mycoremediation of the soil salinity causes accumulation of different compatible solutes and higher uptake of water as well as nutrients. The symbiotic association of AMF regulates the plant physiological and biochemical processes like water potential, ionic balance, stomatal conductance, maintenance of photosynthesis, reduction of oxidative damage through antioxidant production, and hormonemediated signal transduction. Molecular basis of regulation of ionic homeostasis, cation to proton antiporter and cyclic nucleotide-gated channels under AMF symbiosis are less known and therefore should be the thrust area for future research. In Arabidopsis, the 20-member CNGC-encoding genes have overall sequence similarities between 55% and 83% (Maser et al. 2001). However, in mammalian genomes only six CNGC-encoding genes are present. The reason for the large number and diversity of CNGCs in plants is because of important and specialized physiological role there. For AM symbiosis no major CNGCs specific studies were done. Only a few investigations were carried out on cation and proton antiporters, and therefore partial knowledge about the participation of cation/proton antiporters and cyclic nucleotide-gated channels is available. In future research, the efforts should be made to understand the effect of AM symbiosis on cation/proton antiporters, cyclic nucleotide-gated channels, and AMF-mediated changes on plants. Mycoremediation-based research is a promising field to shed light on mechanisms involved in the enhanced tolerance of AM plants to salt stress. Further, transcriptomic analysis of AMF is a promising route to furnish information about fungal genes participating in AMF-plant symbiosis. Overall, these investigations may open new research lines aimed at harnessing maximum benefit from the AM symbiosis under salinity or other osmotic stress.

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Chapter 14

Fungal Enzymes for Bioconversion of Lignocellulosic Biomass Subhadeep Mondal, Suman Kumar Halder, and Keshab Chandra Mondal

14.1  Introduction Plant’s lignocellulosic material is the principal renewable biopolymer of our planet. Globally the estimated production of lignocellulosic materials is around 1 × 1010 ton annually (Hekkert and Negro 2009). Lignocellulosic biomass obtained from plant cell wall (PCW) is chiefly composed of cellulose, lignin, hemicellulose, and pectin (Guerriero et al. 2016). Lignin is the copious biopolymer on earth ranked after cellulose. A. P. de Candolle, a Swiss botanist, in 1813 coined the term “lignine” originated from the Latin word “lignum,” which means wood (Sjostrom 1993). In 1838, French Chemist Anselme Payen discovered that the fibrous component of all higher plant cells had an identical chemical structure, which he named “cellulose” (Payen 1938). Schulze first ever coined the term “hemicellulose” to represent the remnants extracted from plant materials using a dilute alkali (Schulze 1891). Henri Braconnot in 1825 first extracted and narrated pectin (Braconnot 1825). Despite the huge morphological diversity of PCW, it is basically divided into three strata: primary cell wall, secondary cell wall, and middle lamella (Buchanan et al. 2016). Primary cell wall (PCW) (CWp) is defined as wall synthesized during cell growth. The essential components of CWp are pectin (35–45%), cellulose (25– 45%), and hemicellulose (20–30%). The cellulosic microfibrils are associated with the hemicellulosic chain to constitute the cellulose-hemicellulose network. The principal hemicellulose in the CWp is xyloglucan (Fry 1989). When the plant cell enlargement stops, secondary cell wall (SCW) (CWs) synthesis started. The components of CWs are deposited between plasma membrane (PM) and the CWp. The chief components of CWs are cellulose (35–50%), hemicellulose (20–35%), and lignin (10–25%). Lignin is located by excluding water and provides rigidity to the cell wall. The middle lamella layer is flourishing with pectin. S. Mondal · S. K. Halder (*) · K. C. Mondal Department of Microbiology, Vidyasagar University, Midnapore, West Bengal, India e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_14

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Cellulose is a polysaccharide of β (1–4)-linked d-glucose and the chain length varied as minimum of hundreds to few thousands (Klemm et al. 2005). At the PM of vascular plants, the cellulose biosynthesis is done by the hexameric terminal, rosette terminal complex (RTC). Cellulose microfibrils are synthesized by cellulose synthase which form individual glucans, contained within each unit of the RTC (Kimura et  al. 1999). Hemicellulose is a heteropolymer that consists of mainly xylan, arabinoxylan, glucuronoxylan, xyloglucan, and glucomannan. In majority of cases, xylose is present in large amount as a sugar monomer, but in the case of softwoods, mannose is present as the principal sugar. Along with regular sugars, the acidified form of hemicellulose, for example, galacturonic acid and glucuronic acid, can be found (Chen 2014). Proteins synthesized on the ribosome of the endoplasmic reticulum of the plant cell can be transferred to the Golgi complex and form glycosides; the hemicellulose produced is contained in the Golgi vesicles and moved to the cell membrane. In the cell membrane, the Golgi vesicles inosculate to the continuous plasma membrane, further causing the hemicellulose to be stuck to the cell wall. The Golgi apparatus can produce hemicellulose because it can produce the enzymes needed for its synthesis. Lignin is extremely recalcitrant to degradation, synthesized in plants from the phenylpropanoid precursors such as coniferyl, synapyl, and p-coumaryl alcohols (Niladevi 2009). The phenylpropanoid units analogous to lignin polymer are designated as p-hydroxyphenyl (H), syringyl (S), and guaiacyl (G) units, respectively, depending on the methoxy group substitution on the aromatic rings. Usually softwood (gymnosperms, conifers such as hemlock, spruce, and cedar) lignin consists of principally G units and very minute level of H units. Hardwood (leafy deciduous trees, viz. willow, poplar, alder, and birch, angiosperms) lignin is rich in G and S units with minute level of H units. Grasses (monocots) have all three types of units. Monolignol biosynthesis is conducted through the phenylpropanoid pathway. In the shikimate pathway, glucose generated from the carbon dioxide by photosynthesis first is converted to the final intermediate: shikimic acid. Then, through the prephenic acid, the shikimic acid is converted into the final products of the shikimate pathway: phenylalanine and tyrosine. These two amino acids are widely present in plants and serve as starting materials for the cinnamic acid pathway. Under the effects of various enzymes, three monomers of lignin are finally synthesized after a set of reactions, such as deamination, hydroxylation, methylation, reduction, and so on (Geng et  al. 2003). Pectin principally contains d-galacturonic acid units (Sriamornsak 2003), tethered in chains via α-(1,4)-glycosidic linkage. Mostly four kinds of pectic substances are found in PCW: pectic acid, pectinic acids, protopectin, and pectin (Alkorta et  al. 1998). Highest amount of pectin is found in the middle lamella of PCW, with the continuous lessening from the primary wall end route to the PM. Glycosyltransferase (GT) accountable for pectin biosynthesis required specific nucleotide-sugar substrates and acceptors for their activities. The recently accepted model for pectin biosynthesis depends on an active site of Golgi luminal GT transmembrane protein with a nucleotide-sugar ­substrate which is thought to reach into the Golgi lumen through membrane traversing protein transporters or otherwise be synthesized within the Golgi lumen.

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14.2  Enzymes Attacking Lignocellulosic Biomass In the PCW, lignin surrounds cellulose along with mainly hemicellulose and pectin forming a matrix, its degradation leading to the major loss of utilizable wood and concern for plant pathogenesis (Bholay et al. 2012). In Fig. 14.1, the structure and composition of primary and secondary cell wall of plant was depicted with the enzymes responsible for the degradation of the constituents. In the subsequent section, a brief synopsis of those enzymes is discussed.

14.2.1  Ligninase Ligninase is an array of oxidoreductive class of enzyme that mainly consists of lignin peroxidase (LiP; EC 1.11.1.14), manganese peroxidase (MnP; EC 1.11.1.13), and laccase (EC 1.10.3.2). Apart from these, the degradation of highly intractable form of lignin requires the synergistic activities of several accessory enzymes, which may include pyranose 2-oxidase/glucose-1-oxidase (EC 1.1.3.4), aryl

Fig. 14.1  Organization and composition of primary and secondary cell wall of plant with the arrangement of cellulose, hemicellulose, pectin, and lignin moieties, along with respective degrading enzymes

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alcohol oxidase/veratryl alcohol oxidase (EC 1.1.3.7), cellobiose dehydrogenase (EC 1.1.99.18), glyoxal oxidase (EC 1.2.3.5), and cellobiose quinone oxidoreductase (EC 1.1.5.1) (Wong 2009). Ligninase production is regulated by nutrient starvation, especially in nitrogen- and carbon-limited cultures (Ntwampe et al. 2010). MnP and LiP are pertaining to the heme proteins since they contain protoporphyrin IX prosthetic group. Related to diverse heme peroxidases, such as horseradish peroxidase and cytochrome c peroxidase, the enzymatic reactions catalyzed by MnP and LiP have three consecutive steps (Plácido and Capareda 2015). Hydrogen peroxide causes the enzyme oxidation to produce compound I and water:

Reduced peroxidase + H 2 O 2 → Compound I + H 2 O

The first altered enzyme (compound I) catalyzes the production of free radical (S•) and the second altered form of the enzyme (compound II) with an electron shift from the substrate (SH: reduced substrate):

Compound I + SH → Compound II + S •

Ultimately, compound II reacts with a second substrate molecule to synthesize an additional free radical and water. In the meantime, the enzyme reduced to its previous actual form.

Compound II + SH → Reduced peroxidase + S • + H 2 O

Ligninin Peroxidase  This enzyme was first isolated from fungi Phanerochaete chrysosporium in 1983 (Tien and Kirk 1988). LiPs are chemically oligomannose-­ type glycoprotein with a MW that varies from 38 kDa to 43 kDa (Schmidt et al. 1990). LiP is having low optimum pH  3–4.5 and comparatively elevated redox potential. This particular character of LiP makes it an important part of ligninolytic system. LiP oxidizes both phenolics and nonphenolics. The catalytic reactions carried out by LiP are involving cleavage of the propyl side chains, oxidation of benzyl alcohols, hydroxylation of benzylic methylene groups, phenol oxidation, as well as cleavage of aromatic ring in nonphenolic lignin compounds (Renganathan et  al. 1985; Umezawa and Higuchi 1987). Manganese Peroxidase  Kuwahara first discovered manganese peroxidase in 1984 from Phanerochaete chrysosporium (Plácido and Capareda 2015). They are glycoproteins with a MW ranging from 38 to 62.5 kDa. MnP especially favored Mn(II) as its reducing substrate. MnP oxidizes Mn2+ to Mn3+, with the help of organic acid chelators, viz., oxalate, malonate, glyoxylate, etc., and in turn oxidizes several compounds randomly via hydrogen and electron abstraction (Glenn and Gold 1985). The organic acids also assist the relief of Mn3+ from the catalytic site. The one-­electron oxidation of Mn2+ to Mn3+ is a multistep reaction as follows (Niladevi 2009):

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MnP + H 2 O2 → MnP compound I + H 2 O



MnP compound I + Mn ( II ) → MnP compound II + Mn ( III )



MnP compound II + Mn ( II ) → MnP + Mn ( III ) + H 2 O





Laccase  Although laccases originated in plants, bacteria, and insects, the leading producers of these enzymes are fungi. In 1983, laccase was first time isolated from Rhus vernicifer (Japanese lacquer tree). Later, both Bertrand and Labrode in 1896 individually isolated laccase from fungi. In a holoenzymatic monomeric form of fungal laccase (MW 60–85  kDa), generally it has four copper atoms (Coll et  al. 1993) arranged in three groups: T1, T2, and T3 (Leontievsky et al. 1997). Laccase catalysis involves (i) reduction of the T1 copper by reducing substrate; (ii) internal electron shifting from T1 to T2 and T3 copper; and (iii) reduction of oxygen to water at T2 and T3 (Archibald et al. 1997). Laccase certainly attacks the phenolics of lignin, followed by Cα oxidation, aryl-alkyl cleavage, and Cα–Cβ cleavage. The spectrum of substrate accessibility of laccase expanded toward the nonphenolic constituents of lignin with the help of mediator such as ABTS and HBT. ABTS-­ mediated oxidation of nonphenolic substrate is associated with an electron transfer mechanism (Fabbrini et al. 2002; Baiocco et al. 2003). ABTS is first oxidized to the radical cation (ABTS.+) and subsequently to dication (ABTS2+) with redox potentials of 4.72  V (ABTS/ABTS.+) and 8.85  V (ABTS.+/ABTS2+), respectively (Bourbonnais et al. 1998). The dicationic active intermediate is predominantly liable for the oxidation of the nonphenolic compounds. Fungal laccases are monomeric or homodimeric glycosylated protein having fewer monosaccharides (10–25%), such as mannose, arabinose, fucose, galactose, and glucose. Glycosylation is mainly subjected to secretion, proteolytic susceptibility, copper retention, activity, and thermostability (Xu 1997).

14.2.2  Cellulase Chemically, cellulase comprised a family of at least three groups of enzymes: endo(1, 4)-β-d-glucanase (EC 3.2.1.4), exo-(1, 4)-β-d-glucanase (EC 3.2.1.91), and β-glucosidase (EC 3.2.1.21), which act cooperatively to break down cellulose. Endoglucanase  Endoglucanase or endo-(1, 4)-β-d-glucanase randomly digests the internal O-glycosidic bond, resulting in glucan chains of various lengths. Exoglucanase  Exoglucanase or exo-(1, 4)-β-d-glucanase or cellobiohydrolase (CBH) releases β-cellobiose from the reducing (CBH I) and nonreducing (CBH II) ends of cellulose chain. Cellobiohydrolase is inhibited by its hydrolysis product, the glucose.

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β-Glucosidase  It causes the degradation of cellobiose to form glucose and ingests glucose from oligosaccharides by randomly hydrolyzing the β-1, 4 glycosidic bonds in cellulose. β-Glucosidase is feedback inhibited by its end product glucose (Singh 1999; de Castro et al. 2010; Yadav et al. 2016, 2017a) The cellulase complex from Trichoderma reesei comprised an endoglucanase of about 52  kDa, an exoglucanase of about 61  kDa, and a β-glucosidase of about 76 kDa. Two catalytic models have been hypothesized for the reactions catalyzed by cellulases: The Inverting Mechanism  In this mode of catalytic mechanism, the hydrolysis of a glycosidic bond with anomeric configuration inversion is usually achieved in a single-step, single-displacement reaction mechanism including transition states. The catalytic reaction is typically performed with general acid or base support from two specific amino acid side chains, usually glutamate or aspartate that are generally positioned 6–11 Å apart (McCarter and Withers 1994). The Retaining Mechanism  Hydrolysis with configuration retention is accomplished by two steps, double-displacement reaction mechanism including a covalent glycosyl-enzyme transition state. Reaction is carried out with acid/base and nucleophilic support assisted by side chain of glutamate or aspartate, situated 5.5 Å apart. In the initial step, one residue acts as a nucleophile, which attacks the anomeric center to make a glycosyl-enzyme intermediate by displacing the aglycone. At this situation, the other residue acts as acid catalyst and donates protons to the glycosidic oxygen upon breakdown of the bond. In the next (deglycosylation) step, water molecules cause the hydrolysis of glycosyl enzyme with the help of other residue which being a base catalyst deprotonates the water molecules (Koshland Jr 1953).

14.2.3  Pectinase According to their mode of action, the pectinase enzyme is basically divided into three categories: pectin esterase, hydrolases, and lyases (Sharma et al. 2013; Garg et al. 2016) (Kour et al. 2019b). Pectin Esterase  Pectin esterase (PE, EC 3.1.1.11) carried out de-esterification reaction of the methoxy group of pectin resulting in pectic acid. The MW of majority of microbial PEs ranges between 30 and 50 kDa, and the optimum pH for enzyme activity ranges between 4.0 and 7.0 (Hadj-Taieb et al. 2002). Hydrolases  Hydrolases, namely, polygalacturonases and polymethylgalacturonases, hydrolyze the α-(1,4)-glycosidic bonds in pectic acid and pectin, respectively. There are three types of polygalacturonase: endopolygalacturonase (endo-PG, EC 3.2.1.15) cleaves pectic acid randomly and produces oligogalacturonates, exopolygalacturonase 1 (exo-PG1, EC 3.2.1.67) catalyzes the terminal cleavage of polygalacturonic acid from the nonreducing termini and produces

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monogalacturonates, and exopolygalacturonase 2 (exo-PG2, EC 3.2.1.82) catalyzes the penultimate cleavage of pectic acid and produces digalacturonates. The MW of endo-PG lies between 30 and 80  kDa, and in the case of exo-PGs, the MW lies between 30 and 50  kDa. There are two types of polymethylgalacturonase: endopolymethylgalacturonase (endo-PMG) that cleaves pectin in random manner, producing oligo-methyl-­galacturonates, and exopolymethylgalacturonase (exo-PMG) that catalyzes the terminal cleavage of pectin from the nonreducing termini, producing methyl monogalacturonates. Lyases  Lyases (polygalacturonate lyase and polymethylgalacturonate lyase) cleave the α-(1,4)-glycosidic bond in pectin and pectic acid, respectively, through trans-­ elimination reaction, thereby forming unsaturated methyl galacturonates and galacturonates, respectively. The MW of polygalacturonate lyase lies between 30 and 50 kDa, and the optimum pH of enzymatic activity is 8.0 and 10.0. There are three types of polygalacturonate lyase: (i) endopolygalacturonate lyase (endo-PGL, EC 4.2.2.2) that randomly cleaves pectic acid to generate unsaturated oligogalacturonates, (ii) exopolygalacturonate lyase (exo-PGL, EC 4.2.2.9) that cleaves pectic acid from the nonreducing termini at the penultimate site to generate unsaturated digalacturonates, and (iii) oligopolygalacturonate lyase (oligo-PGL, EC 4.2.2.6) that cleaves the oligogalacturonate to generate unsaturated monogalacturonates. The MW of PL lies between 30 and 40 kDa, and its optimum pH for enzymatic activity ranges between 4.0 and 7.0. There are two types of polymethylgalacturonate lyase: endopolymethylgalacturonate lyase (endo-PMGL EC 4.2.2.10) that cleaves pectin randomly, producing unsaturated methyl oligogalacturonates, and exopolymethylgalacturonate (exo-PMGL) that catalyzes the terminal cleavage of pectin, producing unsaturated methyl monogalacturonates.

14.2.4  Hemicellulases Hemicellulase is an array of enzymes mainly comprising xylanase, galactanse, xyloglucanse, and mannanase (Bhat 2000). In an industrial scale, xylanase is mainly beneficial because greater fraction of agricultural biomass utilized is composed of mainly xylan as a hemicellulosic material. Xylanase  The substrate for xylanase, xylan, is the third most plentiful biopolymer on earth (Mellerowicz and Gorshkova 2011), considering up to one third of the renewable organic carbon on this planet (Collins et al. 2005). Xylans are polysaccharides that consist of β-1,4-linked xylose (pentose sugar) residues with side chains of α-arabinofuranose and α-glucuronic acids and assists in cross-linking of lignin and cellulose microfibrils by ferulic acid residues (Balakshin et al. 2011). According to substituted groups, xylan is of three types: (i) glucuronoxylan, (ii) neutral arabinoxylan, and (iii) glucuronoarabinoxylan (Faik 2010). Generally, the amount of xylan in hardwoods and softwoods of hemicelluloses is 10–35% and 10–15%, respectively. The major xylan ingredients in hardwoods and softwoods are O-acetyl-­

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4-O-methylglucuronoxylan and arabino-4-O-methylglucuronoxylans, respectively. Basically, xylans of softwood varies from hardwood by the absence of acetyl groups as well as the presence of arabinose units tethered by α-(1,3)-glycosidic linkages to the backbone (Sixta 2006). The xylanolytic enzyme system that performed xylan hydrolysis normally comprised a reservoir of hydrolytic enzymes, involving endoxylanase (endo-1,4-β-xylanase, E.C.3.2.1.8), β-xylosidase (xylan-1,4-β-xylosidase, E.C.3.2.1.37), acetylxylan esterase (E.C.3.1.1.72), α-arabinofuranosidase (α-l-­ arabinofuranosidase, E.C.3.2.1.55), and α-glucuronidase (α-glucosiduronase, E.C.3.2.1.139) (Juturu and Wu 2012). These enzymes act synergistically to transform xylan into its organizing sugars (Belancic et  al. 1995). In xylanase family, endoxylanases are the most efficient as they are directly involved in the cleavage of glycosidic bonds, excluding short xylooligosaccharides (Verma and Satyanarayana 2012). Xylanase carried out the formation of xylan to xylooligosaccharides by random hydrolysis, whereas β-xylosidase liberates xylose subunits from the nonreducing terminus of xylooligosaccharides. However, the entire breakdown necessitates the cooperation of acetyl esterase to liberate the acetyl units from the β-1,4-linked d-xylose backbone of xylan (Wong and Saddler 1992; Coughlan and Hazlewood 1993).

14.3  Biodiversity of Fungi-Producing Enzymes Microorganisms are considered as a better source of valuable enzymes because of its higher rate of multiplication and synthesizing biologically active products. From the end of the twentieth century, remarkable level of increment in the utilization of enzymes for industrial purposes was realized. These enzymes overcome the disadvantages of traditional chemical agents for various reasons, such as high catalytic activity and substrate specificity, high production rate, biodegradability, economical, and environment friendly (Motta et al. 2013; Yadav et al. 2016, 2017b, 2018). Fungi are the best lignin-degrading enzyme producer, and thereby, industrial-­ scale production of the same is usually carried out with mainly fungal sources. Within fungi, the white rots are the most potent producers of these enzymes, succeeded by the brown rots and the soft rots. White-rot fungi cause decaying of wood that leads to pale color because of oxidative bleaching and loss of lignin and often preserves a fibrous texture. Preferentially some white-rot fungi attack lignin more easily than cellulose and hemicellulose of the woody tissue (Blanchette 1984; Mester et al. 2004). This selective delignification procedure leaves enriched cellulose that is detectable in the white portions of a mottled rot and in the pockets of a white pocket rot. The lignin degradation system of Phanerochaete chrysosporium was mostly studied. Lignin breakdown by P. chrysosporium is an orthodox secondary metabolic activity initiated particularly during nitrogen starvation. This organism secretes LiP, MnP, and laccase for lignin degradation (Tien and Kirk 1983; Glenn and Gold 1985; Srinivasan et al. 1995). Different genera of white-rot fungi, Phlebia ostreatus, P. radiata, Trametes hirsuta, T. versicolor, and T. ochracea, have

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been found to be secreting LiP and MnP along with laccase. In disparity to white-rot fungi, which are mainly found on deciduous (angiosperms) wood, brown-rot fungi primarily grow on conifers (gymnosperms) (Gilbertson 1980). Brown-rot fungi cause the breakdown of wood carbohydrates leading to brown-colored rot due to the remaining oxidized lignin (Cowling 1961). Due to the cellulose breakdown, the decayed wood represents a cross-checking pattern with cracks and clefts; this phenomenon is known as “cubical” brown rot. These include Gloeophyllum trabeum, Postia placenta, Fomitopsis lilacinogilva, Laetiporus portentosus, Piptoporus betulinus, and Serpula lacrymans. The roles of ligninolytic enzymes of fungal origin include lignin degradation, breakdown of plant’s toxic products, and spore/fruiting body formation (Dos Santos et al. 2007). Fungi contributes lion’s share in the worldwide cellulose breakdown. This is particularly important in forest ecosystem, where fungi are the chief decomposers of cellulose (Alexopoulos et al. 1996). Many fungal strains synthesized cellulases as inducible enzymes at the time of their growth in the presence of cellulosic components. For cellulase production, fungi are more preferable than bacteria because (a) bacterial cellulase usually lacks one of the three cellulolytic activities, i.e., FPase activity, (b) downstream processing of fungal cellulase is much easier than bacterial cellulases, and (c) the activity of fungal cellulase is far greater than that of the bacterial cellulase (Sajith et al. 2016; Tomme et al. 1995). Most extensively studied cellulolytic fungal genera are classified as soft-rot fungi like Aspergillus niger (Ong et al. 2004), A. nidulans (Kwon et al. 1992), A. oryzae (Takashima et al. 1998), A. fumigatus (Ximenes et  al. 1996), Fusarium solani (Wood and McCrae 1977), Humicola grisea (Takashima et  al. 1996), Melanocarpus albomyces (Miettinen-­ Oinonen et  al. 2004), Penicillium brasilianum (Jørgensen and Olsson 2006), Trichoderma reesei (Saloheimo et al. 1988), T. viride (Khokhar et al. 2012), T. harzianum (Galante et  al. 1998), Chaetomium cellulyticum (Fähnrich and Irrgang 1981), Neurospora crassa (Romero et al. 1999), C. thermophilum (Li et al. 2009), Thermoascus aurantiacus (Kalogeris et  al. 2003), Mucor circinelloides (Saha 2004), and Paecilomyces inflatus (Kluczek-Turpeinen et al. 2007); brown-rot fungi like Coniophora puteana (Schmidhalter and Canevascini 1992), Poria placenta (Highley 1977), Tyromyces palustris (Yoon and Kim 2005), and Fomitopsis sp. (Deswal et al. 2011); and white-rot fungi, viz., Phanerochaete chrysosporium (Jäger et al. 1985), Sporotrichum thermophile (Kaur et al. 2004), Agaricus arvensis (Jeya et al. 2010), Pleurotus ostreatus (Khalil et al. 2011), and Phlebia gigantea (Niranjane et al. 2007). Microbial sources of pectinolytic enzymes include filamentous and nonfilamentous bacteria, yeasts, and filamentous fungi. Most extensively studied cellulolytic fungal genera are Aspergillus niger, A. fumigatus, A. flavus, A. oryzae, A. sojae, Penicillium viridicatum, P. chrysogenum, P. atrovenetum, Rhizomucor pusillus, Trichoderma viride, Phytophthora infestans, etc. (Couri et al. 2000; Phutela et al. 2005; Demir et al. 2012; Silva et al. 2002; Banu et al. 2010; Siddiqui et al. 2013; Irshad et al. 2014). The xylanolytic enzymes are widely distributed among fungi. Industrially important xylanases are preferentially obtained from thermophilic organism. Some fungal

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genera show elevated thermal stability such as Thermoascus aurantiacus and Paecilomyces themophila have been found to be stable at high temperature (Yang et al. 2006). Thermostable Thermomyces lanuginosus exhibited optimum activity at 65–75°C, at pH 5–6.5 (Cesar and Mrša 1996). Except these, other important xylanase producers of fungal origin are Aspergillus awamori, A. foetidus, Fusarium oxysporum, Geotrichum candidum 3C, and T. reesei (Christakopoulos et al. 1996; Rodionova et al. 2000).

14.4  Production of Fungal Enzymes Solid-state fermentation (SSF) is the current trend for industrial-scale production of most of the enzymes in comparison to the submerged fermentation (SmF). The benefits of SSF over SmF are economically lower fermentation cost, minor risk of contamination (Beg et al. 2001), simple recovery of the enzymes, compatible with the natural environment of the fungus, production of a protein-augmented by-product, production of enzymes with intensified enzyme activities, and higher specific activities (Coughlan 1989). SSF is especially compatible for the fungal growth because their growth sustained at comparatively low-water activities at which the growth of bacteria and yeast is not favored (Corral and Villaseñor-Ortega 2006). The selection of fermentable substrate(s) is gaining special attention for the production of lignocellulosic biomass-degrading enzymes to reach the expected level. The lignocellulosic waste from agricultural-based industries has shown potentiality to act as substrate for the high titers of laccase production. Laccase biosynthesis by Pleurotus florida PF05 was reported by utilizing corncob oil seed cakes as substrate. Marques De Souza et al. (2002) reported laccase production by utilizing both wheat bran and straw as substrates in 1:1 ratio by Penicillium pulmonarius. Shinners-Carnelley et  al. (2002) studied the production of laccase from sugarcane bagasse by Penicillium chrysosporium. Rosales et al. (2007) performed SSF using orange peel by Trametes hirsuta. Trametes versicolor efficiently utilizes barley bran for maximum laccase production. Karim and Annuar (2009) used coconut husk for laccase synthesis by Pycnoporus sanguineus. During SSF, Coriolus versicolor produced laccase when rice bran was used as substrate (Chawachart et al. 2004). Agricultural wastes such as brans and straws of wheat and rice, sugarcane bagasse, corn stover, sawdust, etc. are principal substrates used for the synthesis of cellulase. For instance, Liu et al. (2011) investigated the cellulose preparation on different lignocellulosic substrates such as straws of rice, wheat, and cotton, corn stover, and corncob using Aspergillus fumigatus; of them, the corncob supported the maximum synthesis of endoglucanase. Penicillium echinulatum showed the maximum biosynthesis of cellulolytic enzymes on the medium containing a mixture of pretreated wheat bran and sugarcane bagasse (Camassola and Dillon 2007). Dutta et al. (2008) studied the production of cellulases from Penicillium citrinum using brans of wheat and rice and rice straw as substrate; all these substrates supported the

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cellulases production at high magnitude. Aspergillus niger efficiently utilized the lignocellulosic substrates like grass, bagasse, and corncob with variable cellulase activities (Sohail et al. 2009). Aspergillus flavus competently utilized different lignocellulosic substrates and supported higher levels of cellulase activity (Sajith et al. 2014). Aspergillus flavus produces cellulose when grown on various natural substrates like bagasse, sawdust, and corncob (Ojumu et al. 2003). Manimaran et  al. (2009) reported higher-level synthesis of xylanase by Thermoascus aurantiacus during growth on bagasse pulp under optimized solid-­ state fermentation. Kang et al. (2004) reported xylanase biosynthesis by A. niger KK2 using rice straw as substrate. Mohana et al. (2008) studied the production of xylanase by Burkholderia sp. by employing distillery spent wash pretreated anaerobically. Yang et al. (2006) synthesized higher amount of xylanase under optimized conditions by Paecilomyces thermophila J18 by utilizing wheat straw as substrate. Thangaratham and Manimegalai (2014) carried out a comparative study by involving Aspergillus oryzae, Aspergillus flavus, and Rhizopus oryzae for pectinase production by applying different agro-industrial wastes, including sawdust, pineapple peel, cassava waste, lemon peel, and wheat bran. Jahan et al. (2017) reported higher production of polygalacturonase using Bacillus licheniformis using wheat bran as SSF substrate. A combination of wheat bran and orange bagasse as substrates in the ratio 1:1 allows the higher production of pectinase using the filamentous fungus Penicillium viridicatum RFC3 (Silva et  al. 2005). Biz et  al. (2016) construct an identical process to synthesize pectinases using sugarcane bagasse and citrus peel as SSF substrates.

14.5  Biotechnological Applications The aforementioned enzymes have versatile fields of application (Fig. 14.2) in different sectors either in purified or in cocktail form. In the following section, the prime applications are stated.

14.5.1  In Biofuel Industry The combustion of fossil fuel has a serious impact on atmospheric CO2 levels and is an important agent to man-made global climate change (Topakas et  al. 2013). Production of biofuels overcomes this issue, but the major problem in producing biofuels from lignocellulosic biomass is the processing cost rather than availability and cost of feedstock (Lynd et al. 2008). To reach the sufficient level of biofuels production to compete with fossil fuels, the ability of transforming lignocellulosic biomass to monomeric sugars must be enhanced by producing enzymes of cost-­ effective and greater specific activities (Galbe et  al. 2007; Kour et  al. 2019a; Rastegari et al. 2019)

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Fig. 14.2  Different fields of application of enzymes associated with lignocellulosic biomass degradation

Laccases assist the pretreatment of lignocellulosic materials by excluding their phenolic compounds for the subsequent yield of bioethanol (Fang et  al. 2015). White-rot fungal genus Phlebia sp. MG-60 selectively converts lignin under SSF and from this delignified biomass ethanol is synthesized under semi-aerobic submerged fermentation (Kamei et al. 2012a). Phlebia sp. MG-60 is unique in that it degrades lignin and subsequently produces ethanol from cellulose (Kamei et  al. 2012b). After removing lignin and hemicellulose contents from lignocellulosic biomass, cellulase is treated to hydrolyze cellulose residue to generate fermentable sugars, and finally a fermentable microbe is applied for biofuel production by utilizing those sugars (Sukumaran et al. 2005). Xylanase along with ligninase and cellulase facilitates the biofuel production from lignocellulosic residues (Dominguez 1998; Olsson and Hahn-Hägerdal 1996). The plants which are rich in pectin source like apple pomace, citrus, and sugar beet have been recommended as possible sources of hydrocarbon for bioethanol production (Edwards and Doran-Peterson 2012).

14.5.2  In Paper and Pulp Industry The mechanical pulping process involving refining and grinding of the woody raw material assists pulps with fine particles of greater amount of bulkiness and stiffness (Bhat 2000). Wood fiber has a multilayered structure consisting primarily of cellulose, hemicellulose, and lignin. In pulp and paper industry, enzymes related to lignocellulosic biomass breakdown aid biomechanical pulping for prospering the rough mechanical pulp (Bedford et  al. 1997), hand-sheet strength characteristics (Akhtar 1994), deinking of reprocessed papers (Pere 1995), and elevating drainage waste effluent from paper mills (Prasad et al. 1992). Lignin is an insoluble complex biopolymer of phenolic compounds. Conventional methods for delignifying paper pulp include either chlorinated or oxygenated agents (Bajpai and Bajpai 1992). Albeit effective, these methods have severe disadvantages such as environmental pollution, weakening the cellulose fiber strength, etc. Enzymatic delignification procedure overcomes these problems and has the

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potential as an alternative to the conventional methods. During delignification, laccases are found to act on small phenolic lignin parts that subsequently reacted with the lignin polymer, leading to its degradation (Madhavi and Lele 2009; Yadav et al. 2019a, b). Application of cellulase for pulping enhances the energy efficiency of the process and also improves the physical properties, such as interfiber bonding and mechanical strength of the ultimate paper product (Chen et al. 2012). Cellulase was also used for deinking of waste papers. During deinking, the ink attached to the surface of recycled cellulose fibers was released by the enzymatic hydrolysis of carbohydrates, leading to the peeling of individual fibers or bundles (Kuhad et al. 1997). Recycling of used papers reduces the solid wastes and also decreases the burden of deforestation for wood fibers (Lee et  al. 2011; Singh et  al. 2012). Enzymatic deinking of waste papers, especially employing the mixtures of cellulase and hemicellulase, enhances the quality and brightness of the recycled paper (Lee et al. 2007; Ibarra et al. 2012). Cellulases are also used to elevate the drainage of several paper mills by dissolving clogged fiber residues (Kuhad et  al. 2011). Moreover, the cellulase preparations are also used to make easily biodegradable cardboards, tissue papers, and sanitary papers (Buchert et al. 1998; Bajpai 1999). During the Kraft process, i.e., the chemical transformation of wood into wood pulp (lignocellulosic fibrous components), the lignin-carbohydrate complex is hydrolyzed. Thus, endo-1,4-β-xylanase can be used in this process to increase the extraction of lignin. In that way it becomes more accessible for bleaching. By this approach the consumption of chlorine chemicals for bleaching is reduced. In addition to being environment friendly, enzymatically treated pulp is brighter and has improved fiber quality (Kalim et al. 2015). It was also reported that pulps and papers treated with hemicellulases lead to the reduction in necessity of chlorine bleaching (Viikari et al. 2001). Pectins found in papers impair dewatering during sheet production because of their high cationic requirement and give yellowness of paper. Depolymerization of polygalacturonic acids by pectinases lowers the cationic requirement in the hydrogen peroxide-bleached thermomechanical pulp filtrate (Viikari et  al. 2001). Pectinases exclusively or with other enzymes synthesized by same or other microorganisms have been effectively utilized for biobleaching of bamboo kraft pulps and mixed hardwood (Dhiman et al. 2009).

14.5.3  In Food Processing Along with progressive health-conscious civilization, fruit and vegetable juice preparations gain huge impact on both the human health and industrial sector. Fruit and vegetable juice production at industrial level needs detailed knowledge about the methods of extraction, clarification, and stabilization. During the early 1930s, when fruit industries were just blooming to produce juice, many difficulties were encountered, such as the yields were low and clarity of fruit juices was not to a receivable

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grade (Uhlig 1998). At present, a mixture of pectinases, laccase, cellulases, and hemicellulases, collectively acquainted as macerating enzymes, are utilized in the juice extraction and clarification from fruits and vegetables (Galante et al. 1998). Stutz (1993) prepared a clear and stable apple juice with the help of laccase and ultrafiltration. Laccase treatment at suitable pH and consequent ultrafiltration ameliorated the juice quality by means of flavor and durability in contrast to the conventional treatments with ascorbic acid and sulfites. Cellulases performed a leading role in macerating enzymes complex (mainly cellulases, pectinase, and xylanases); they are used to ameliorate texture and cloud stability and reduce viscosity of the purees and nectars of the juice obtained from tropical fruits like, mango, papaya, peach, apricot, pear, and plum (Sukumaran et al. 2005; Bhat 2000; De Carvalho et al. 2008). Improving sensory qualities of fruits and vegetables juices like texture, aroma, and flavor can be achieved by reducing the extra bitterness of citrus fruits through the addition of enzymes such as β-glucosidases and pectinases (Baker and Wicker 1996; Youn et al. 2004; Rai et al. 2007). The same enzyme mixture is also helpful for improving the extraction procedure of olive oil. Cellulase along with pectinase is utilized in the extraction of pigments (e.g., carotenoid) which are used as food coloring agents (Choudhari and Ananthanarayan 2007). A mixture of pectin methylesterase with calcium is used as softening agent in pickle industries (Baker and Wicker 1996). Pectinase treatment improved tea fermentation by degrading pectin that is present in the tea leaves and improves the quality of instant tea powder. A characteristic aroma of tea is associated with alteration in tea color during the fermentation (Praveen and Suneetha 2015). Marimuthu et al. (1997) documented that a mixture of cellulase, pectinase, and xylanase upgrades the quality of tea by increasing ­thearubigin (TR), theaflavin (TF), high polymerized substances (HPS), and total soluble solids (TSS)  content. The robusta coffee obtained from the bean is  surrounded by the gelatinous and viscous mucilage layer that comprised sugars (4.0%), pectin (2.8%), moisture (84.0%) with protein (8.9%), and ash (0.9%). Pectinases eliminate the layer of pectic substances from the coffee bean. Pectinase treatment minimizes demucilization time associated with sugar content increment and lowering of pH (Murthy and Naidu 2011).

14.5.4  In Wine and Brewery Industry One of the vital applications of laccase in alcoholic beverage industry is wine stabilization. Wine aroma depends on the alcohol and organic compounds, whereas phenolic compounds determine the color and taste of the wine. Availability of metal ions (like copper and iron), enzymes, amino acids, proteins, aldehydes and polyphenols in wine causes oxidative reactions, resulting in turbidity, aroma, color intensification, and flavor changes. This oxidative phenomenon is called maderization. In comparison to the conventional chemical treatment of Riesling wine with sulfur dioxide and polyvinylpyrrolidone (PVP), the polyphenol content, sensorial quality, color, and haze stability were uplifted with laccase treatment (Madhavi and Lele 2009).

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In wine production, a combination of enzymes such as glucanases, pectinases, and hemicellulases in well-balanced conditions performs a major role by ameliorating color extraction, must clarification, skin maceration, filtration, and lastly the wine stability and quality (Singh et al. 2007; Galante et al. 1998). β-Glucosidases are responsible for developing aroma in wines by modifying glycosylated precursors. Macerating enzymes also maintain the pressability, settling, and juice production from grapes utilized in wine fermentation. Oksanen et al. (1985) reported that enzymatic preparations of exoglucanase II and endoglucanase II from Trichoderma cause maximum amount of reduction in polymerization and wort viscosity level. Galante et al. (1998) conducted an experiment for wine production by treating three qualities of white grapes (namely, Soave, Chardonnay, and Sauvignon) by a cocktail of cellulase, pectinase, and xylanase, commercially known as Cytolase 219, and found that improvement in wine must extraction, wine must filtration rate, and stability was realized with minimization of pressing time and must viscosity.

14.5.5  In Textile Industry Laccases play an essential role in preventing back staining of printed or dyed textiles. Laccases in washing solution thoroughly bleach released dyestuff, leading to the reduction in energy, processing time, and water needed to obtain textile up to satisfactory level. Laccase-based bleaching system replaced the traditional chemical oxidation step and showed its potentiality to bleach the indigo dye in denim and acquire various bleached outlooks on the fabric (Madhavi and Lele 2009). Conventional stonewashing of jeans included amylase-mediated dispel of starch coating followed by pumice stone-mediated machine washing of jeans. In commercial sector, cellulases find its role by biopolishing the cotton and various other cellulose-­based fabrics and biostoning of jeans. During the biostoning procedure, cellulases attack the cotton fabrics and split the tiny fiber endings on the yarn surface which leads to dye slacking causing easy removal during washing by mechanical friction. The benefits associated with cellulose-based treatment include less harm to the cotton fibers, less intensive work, and improved production efficiency of the machines in an environmentally protected manner (Sukumaran et al. 2005; Uhlig 1998). Acidic cellulases are particularly helpful in improving softness and water-absorbing properties of fibers, sharply reduce the propensity for pill formation, and give a bright and cleaner surface feature with reduced fuzz (Sreenath et al. 1996). Endoglucanases-rich cellulase preparations are efficient in biopolishing by improving look, color, and feel without any chemical anointment of fibers (Galante et al. 1998). In regular washing, the cotton clothes became fluffy and dull which is chiefly due to the partially loosed microfibrils on the clothes surface. Application of cellulase causes removal of those microfibrils, thereby retaining a smooth surface, original color, and appearance to the clothes (Hebeish and Ibrahim 2007; Ibrahim et al. 2011). Infliction of cellulase also helps in softening the clothes by removing dirt particles entrapped within microfibril network.

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Pectinases in combination with amylases, cellulases, and hemicellulases are used to dispel sizing agents from cotton in such a safe and ecofriendly way that displaces traditional process of using toxic sodium hydroxide. Pectinases are employed for bioscouring (noncellulosic impurities removal from the fiber by applying specific enzymes) (Jayani et al. 2005).

14.5.6  In Bakery Industry The application of enzyme preparations in bread making to produce fluffier bread has been reported long ago (Blagoveschenski and Yurgenson 1935). It was reported that the incorporation of laccase in baked products in bakery industries causes oxidation of dough materials and strengthens gluten structures of wheat as well as increased dough volume, smoothness of final baked product, a better crumb structure, increased strength, reduced stickiness, and stability, thus overall ameliorating the machinability of the dough. Xylans play a major role in bread making because of their interaction with gluten and high water absorption potentiality. Endoxylanase attacks arabinoxylan backbone to reduce the degree of polymerization and consequently its structure and function (Courtin and Delcour 2002). Xylanase heightens dough stability, dough machinability, lump volume, crumb structure, oven spring, and shelf life when applied in optimum level (Hamer 1995; Poutanen 1997).

14.5.7  In Biomedical Sector Laccase covalently conjugated to bioadhesive molecules (antibody, antigen, DNA, RNA, biotin, and streptavidin) can be employed as a marker enzyme in the immunological, histological, cytological, or molecular biological assay. In such protocols, the binding moiety attached to the target, and subsequently the laccase signals the binding incident (Schmid and Urlacher 2007). Laccases have shown its ability to cure aceruloplasminemia. The disease is associated with the lack of ceruloplasmin, a multicopper serum oxidase whose ferroxidase activity controls iron homeostasis (Harris et  al. 2004). Poison ivy dermatitis is caused mainly by urushiol (catechol-derived toxin). The application of laccase causes oxidation, polymerization, and detoxification of urushiol (Madhavi and Lele 2009). The first significant pharmaceutical chemical that has been made by employing laccase is actinocin, produced from 4-methyl-3-hydroxyanthranilic acid. Actinocin is an anticancer compound which restricts the tumorous growth by blocking DNA transcription (Burton 2003). Another example of the anticancer drug is vinblastine which is useful in curing leukemia. The plant Catharanthus roseus naturally produces vinblastine in small amount. This compound is synthesized in plant from the precursors—katarantine and vindoline. Laccases find its role by 40% transformation of

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these precursors to the end product (Yaropolov et al. 1994). Catechins found in tea, vegetables, and herbs consist of small amount of tannins. The antioxidative property of catechin makes them beneficial for healing several diseases including cancer, cardiovascular diseases, and inflammatory diseases. The antioxidative property of catechin can be upgraded by the oxidation of laccase and producing catechins in numerous goods with amplified antioxidative capability (Kurisawa et  al. 2003). Erb-Downward et al. (2008) reported that laccase was involved in the creation of immunomodulatory prostaglandins. Laccase of oyster mushroom can obstruct hepatitis C virus penetration into hepatoma and peripheral blood cells (El-Fakharany et  al. 2010). Laccase of edible mushroom Agrocybe cylindracea was reported to inhibit HIV-1 reverse transcription (Hu et al. 2011). Cellulases are applied to synthesize chitosan with antibacterial, immunomodulatory, and antitumor activities (Qin et al. 2004; Wu and Tsai 2004). Cellulases are directly applied to human for the treatment of phytobezoars. A phytobezoar is a calculus formed in the gastrointestinal tract that consists of swallowed foods of plant origin, such as indigestible vegetables and/or fruit fibers. Few medical complications are healed by using cellulases alone (Pinos et al. 2015). Cellulases cause breakdown of biofilm by attacking cellulose which is one of the vital constituents in the matrix of biofilm (Flemming and Wingender 2010; McCrate et  al. 2013). Pathogenic microbes construct biofilms, allows spreading and protecting themselves. It is a matter of concern because maximum drugs are incapable to penetrate the biofilm structure. Several studies and patents suggested that the direct insertion of cellulases acts as antibiofilm agent for medical implants (Loiselle and Anderson 2003), diverse prosthetic materials (Rajasekharan and Ramesh 2013), treatment of cystic fibrosis (Ma et al. 2009; Rajasekharan and Ramesh 2013), treatment of nosocomial infections (Huertas et al. 2014), among others. Endoxylanase-catalyzed xylan hydrolysis results in xylooligosaccharides (XOs). Xylan results in the formation of xylose, arabinose, and methyl-d-glucuronic acid-­ bearing XOs (Goswami and Rawat 2015). XOs have numerous impacts in various fields like agricultural, food pharmaceuticals purposes, and feed formulations (Vazquez et al. 2000). XOs have prebiotic action as food additives by improving the intestinal health by flourishing the growth of health-beneficial Bifidobacteria (Fooks and Gibson 2002). XOs are applied as dietary supplements due to their instrumental effect on gastrointestinal tract and may decrease the threat of colon cancer (Whitehead and Cotta 2001). Acceptance of XOs is due to the allowable odor and noncarcinogenic nature (Kazumitsu et al. 1987; Kazumitsu et al. 1997). XOs have low calorific value and therefore are applied as antiobesity diet products (Taeko et al. 1998; Toshio et al. 1990). Non-digestibility of pectin in the gut of human makes it as a dietary fiber by increasing the viscosity in the intestinal tract. Enhancement of the viscosity of the intestinal tract results in reduced cholesterol absorption by increasing the excretion of neutral sterols and fecal bile acids, by intervening with the formation of micelles, and/or by reducing the diffusion rate of bile acid and cholesterol-carrying micelles through the bolus, therefore curtailing the incorporation of cholesterol and bile acids. High amount of negative charges on demethylated pectin makes them as a

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weak cation exchanger, and therefore, at favorable pH, they chelate toxic ions. Intestinal bacteria easily ferment demethylated pectins, subsequently producing short-chain fatty acids, propionate, butyrate, and acetate. These products protect the bowel against inflammatory diseases, and also regulate the insulin release and appetite by modulating the release of gut hormones (Tolhurst et al. 2012).

14.5.8  In Animal Feed Sector Zearalenone (produced by Fusarium sp.) is globally present in an array of cereal crops, such as maize, barley, oats, wheat, rye, rice, millet, and sorghum. Zearalenone is the main toxin responsible for infertility, abortion, and breeding-associated problems, especially in pigs. The symptoms are particularly complicated in prepubertal gilts involving enlarged breast, atrophy of the ovaries, and swelling of uterus and vulva. Humans come in contact with zearalenone by consuming animal meat, by swallowing contaminated grain, and also from bread baked with contaminated wheat. Vikso-Nielsen and Sorensen (2015) reported that laccase treatment causes the conversion of zearalenone into nontoxic substances. Treatment of cereal straw with enzyme alone or in conjunction with other treatments improved their degradability by the rumen microbes (Rodrigues et  al. 2008). Feeding the sheep with 24  hours lignolytic enzymes-treated straw (Eleusine coracana) causes not only higher lignin degradation but also higher digestibility in comparison to consuming the immediate enzyme-treated straw (Sridhar et al. 2014). The cereal-centered food of poultry and pigs are far less complicated than forage food of ruminants comprising cellulose, pectin, hemicellulose, and lignin. An enzymatic complex having higher amounts of cellulase, hemicellulase, and pectinase increases the nutritive value of forages (Lewis et al. 1996). Together, cellulases and hemicellulases improve the nutritional status of animal feed by semihydrolysis of lignocelluloses, hydrolysis of β-glucans, better emulsification and pliability, and dehulling of cereal grains (Galante et  al. 1998; Cowan 1996). Furthermore, these enzymes are involved in incomplete hydrolysis of PCW at the time of silage or fodder preservation. Propionic acid, a bacteriostatic agent produced during cellulases-­based cereal fermentation, has a positive impact on ruminant animals by decreasing the colonization of pathogenic bacteria (Fortun-Lamothe et al. 2001). Supplementation of fibrolytic enzymes like xylanases and cellulases significantly raised the average milk production per day in Murrah buffaloes (Shekhar et al. 2010). Similar results were also noticed for goats (Bala et al. 2009). Wheat, triticale, and rye are rich in arabinoxylans (Bonnin et al. 1998). Endo-(1,4)-d-xylanase ease the mobility of the gut materials leads to the improved nutrient absorption and diffusion of the enzymes secreted from pancreas. Consumption of xylanase-treated rye-based feed to broiler chickens leads to the reduction in intestinal viscosity, improved feed conversion efficiency, and weight gain (Bedford and Classen 1993). Incorporation of pectinases to the animal feed lessens feed viscosity which in turn improves nutrients absorption and decreases the amount of fecal matter (Jayani et al. 2005).

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14.5.9  In Agricultural Sector Mixture of cellulases, hemicellulases, and pectinases produced by fungal strains of Trichoderma and Penicillium are utilized for the synthesis of protoplast of plant and fungi which are fused to yield mutant or hybrid strains with anticipated characteristics (Beguin and Aubert 1994; Bhat and Bhat 1997; Yadav et  al. 2015, 2017b). The pathogenic fungal genera Fusarium sp. and Rhizopus solani are liable for leakage of cytoplasm, hyphal tip swelling, formation of many septae, etc. Such pathogen-­inducing plant diseases are limited by β-1,3-glucanase isolated from T. harzianum CECT 2413 (Kuhad and Singh 2013; Kubicek and Harman 1998). Cucumber seedlings infected by the plant pathogen Pythium sp. can be limited by the inhibition of its growth by hypercellulolytic mutant of T. longibrachiatum, which synthesize high amount of β-1,4-endoglucanase compared to the wild type (Kubicek and Harman 1998). Thus, cellulase performed beneficial role as biocontrol agents to defend the plants and its seeds from plant pathogens (Bhat 2000). Application of exogenous cellulase leads to improvement in soil fertility by decomposition of cellulose (straw) in soil and thus minimizes reliance on mineral fertilizers (Han and He 2010; Fontaine et al. 2004). Paenibacillus ehimensis KWN38 synthesizes antifungal compounds along with the lytic enzymes, mainly cellulases and β-glucanases, which defend plants against pathogenic oomycetes, such as strains Phytophthora (Naing et  al. 2014). Moreover, cellulases synthesized by this Paenibacillus species may be responsible for rhizospheric soil decomposition; thus more nutrients are now accessible to the plant (Han and He 2010; Kour et al. 2019b; Yadav et al. 2017a; Yadav 2018). Xylanase treatment of plant cells can induce glycosylation and fatty acylation of phytosterols. Treatment with purified endoxylanase of Trichoderma viride of tobacco suspension cells leads to a 13-fold rise in the intensities of acylated sterol glycosides which in turn causes large amount of phytoalexins synthesis (Moreau et  al. 1994). Xylanases from the germinating plant seed during ripening convert reserve food to the suitable end product. Xylanases are also reported to take active part in softening of fruits, e.g., papaya. During ripening when the fruits become soften, endoxylanases modify the polysaccharide content in the matrix of cell wall (Manenoi and Paull 2007). The fiber which is obtained from fiber crop holds gum. Ramie fiber is an excellent natural textile, but it contains 20–35% ramie gum that mostly contains pectin and hemicellulose and is therefore detached by further treatment for textile processing. Conventionally degumming can be done by chemical treatment, 12–20% NaOH in solution form is used to remove gum of decorticated fibers (Cao et al. 1992). But chemical treatment is not much efficient, and this type of degumming is toxic, contaminating, and nonbiodegradable. Pectinases with xylanase can be employed for degumming in a sustainable manner (Jayani et al. 2005). A mixture of xylanase with pectinase can be employed for the degumming of bast fibers such as flax, ramie hamp, and jute (Sreenath et al. 1996). The same enzymatic mixture can also be involved in the debarking process, the early

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step in wood handling (Bajpai 1999). Alkaline pectinases and cellulases are employed for the extraction of viruses in pure form when the viral particles are present in phloem (Jayani et al. 2005).

14.5.10  In Waste Management A leading problem of our present world is contamination of air, soil, and water by harmful chemicals which may have catastrophic influence not only on human health but also their surrounding environment. Unrelenting legislation have been executed on industries for treating their waste effluents before ultimate discharge into the ecosystem. Fungal laccases are employed for the detoxification and decolorization of industrial effluents as well as wastewater treatment (Chandra and Chowdhary 2015). Major environmental role of laccases is the remediation of toxic soils involving the oxidation of toxic organic chemicals, such as polycyclic aromatic hydrocarbons, chlorophenols, organophosphorus compounds, lignin-related structures, phenols, and azo dyes in free or immobilized state (Saratale et  al. 2011; Khan et  al. 2013). Gaitan et  al. (2011) found that laccase of Trametes pubescens can degrade mixture of pentachlorophenol, 2,4-dichlorophenol, and 2,4,6-trichlorophenol. Saparrat et  al. (2010) reported that the water-soluble fraction of “alpeorujo,” a by-product pulled out during the olive oil extraction, can be detoxified by laccase. Zhao et al. (2010) studied that the application of laccase in soil causes the degradation of dichlorodiphenyltrichloroethane. Along with the biodegradation of 2,4-dichlorophenol by laccase, treatment of the same can weaken the bisphenol-A-mediated cancer cell propagation (Bolli et  al. 2008). Pozdnyakova et al. (2006) reported the biodegradation of polycyclic aromatic hydrocarbons like fluoranthene, anthracene, fluorene, phenanthrene, pyrene, and perylene using synthetic mediator-based laccase production by Pleurotus ostreatus. The wastes from agricultural fields, agroindustries, and forests contain a huge mass of unprocessed or underprocessed cellulose, resulting in environmental pollution (Milala et al. 2005). At present these biowastes are judiciously applied to produce valued products, viz., enzymes, biofuels, sugars, nutrients, biochemicals, as well as cheap substrate for fermentation and improved animal feeds (Kuhad et al. 2010; Kuhad et al. 1997; Karmakar and Ray 2011; Humpf and Schreier 1991; Gupta et al. 2009). Hemicelluloses (xylan)-rich agrowastes are treated with xylanase to convert xylan into xylose which are subsequently employed for the production of biogas, enzyme (xylanase), and other value-added products (Stalin et  al. 2012; Fang et al. 2010). The wastewater discharged from the citrus fruit processing industries contains large amount of pectinaceous substances which are not fully decomposed during the activated sludge treatment process (Tanabe et al. 1986). Vegetable food processing

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industries also discharged pectin substances as waste effluent. Treatment of this waste effluent with pectinases makes it suitable for decomposition by activated sludge treatment (Jayani et al. 2005).

14.6  Conclusion and Future Perspectives Irresistible demand for environment-friendly products elevated the value of industrial enzymes, among which lignocellulose-degrading enzymes gained pivotal role. Ample supply of lignocellulosic biomass makes it a potential source in agricultural, biofuel, textile, biomedical, food processing, and paper and pulp industries, as well as in research and development. Most significant lignocellulose biomass attacking enzymes such as laccase, cellulase, xylanase, and pectinase are produced by fungi belonging to diverse genera. These enzymes also availed worldwide attention in an expected level regarding the waste management, and their utilization overcome the drawbacks in various conventional chemical-based treatments in different industries. Present scenario for the fermentative production of these enzymes from lignocellulosic biomass in an industrial scale is still not reached to the expectation. The major aims for future research regarding the synthesis of these enzymes would be cost-effective production and increment in the potentiality of the enzymes. Attaining such goals required tremendous level of research for searching new microbes of better enzyme production both quantitatively and qualitatively, their growth optimization, and genetic engineering to develop the properties of these enzymes. Acknowledgement  The authors are grateful to the Department of Science and Technology and Biotechnology, Govt. of West Bengal, India for financial assistance (Memo No: 532/(Sanc.)\ST/P/ S&T/2G-48/2018 dated: 27/03/2019).

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Tolhurst G, Heffron H, Lam YS, Parker HE, Habib AM, Diakogiannaki E, Cameron J, Grosse J, Reimann F, Gribble FM (2012) Short-chain fatty acids stimulate glucagon-like peptide-1 secretion via the G-protein-coupled receptor FFA2. Diabetes 61:2364–2371 Tomme P, Warren RAJ, Gilkes NR (1995) Cellulose hydrolysis by bacteria and fungi. Adv Microb Physiol 37: 1–81 Topakas E, Panagiotou G, Christakopoulos P (2013) Xylanases: characteristics, sources, production, and applications. In: Bioprocessing technologies in biorefinery for sustainable production of fuels, chemicals, and polymers. Wiley, New York, pp 147–166 Toshio I, Noriyoshi I, Toshiaki K, Toshiyuki N, Kunimasa K (1990). Production of Xylobiose. Japan Patent JP 2119790 Uhlig H (1998) Industrial enzymes and their applications. John Wiley Sons, New York Umezawa T, Higuchi T (1987) Mechanism of aromatic ring cleavage of β-O-4 lignin substructure models by lignin peroxidase. FEBS Lett 218(2):255–260 Vazquez MJ, Alonso JL, Dominguez H, Parajo JC (2000) Xylooligosaccharides: manufacture and applications. Trends Food Sci Technol 11:387–393 Verma D, Satyanarayana T (2012) Molecular approaches for ameliorating microbial xylanases. Bioresour Technol 117:360–367 Viikari L, Tenkanen M, Suurnäkki A (2001) Biotechnology in the pulp and paper industry. In: Biotechnology: Special processes, vol 10. Wiley, Hoboken, pp 523–546 Vikso-Nielsen A, Sorensen BH (2015) U.S. Patent No. 9,040,275. Washington, DC: U.S. Patent and Trademark Office Whitehead TR, Cotta MA (2001) Identification of a broad-specificity xylosidase/arabinosidase important for xylooligosaccharide fermentation by the ruminal anaerobe Selenomonas ruminantium GA 192. Curr Microbiol 43:293–298 Wong KK, Saddler JN (1992) Trichoderma xylanases, their properties and application. Crit Rev Biotechnol 12(5–6):413–435 Wong DW (2009) Structure and action mechanism of ligninolytic enzymes. Appl Biochem Biotechnol 157(2):174–209 Wood TM, McCrae SI (1977) Cellulase from Fusarium solani: purification and properties of the C1 component. Carbohydr Res 57:117–133 Wu GJ, Tsai GJ (2004) Cellulase degradation of shrimp chitosan for the preparation of a water-­ soluble hydrolysate with immunoactivity. Fish Sci 70:1113–1120 Ximenes EA, Felix CR, Ulhoa CJ (1996) Production of cellulases by Aspergillus fumigatus and characterization of one β-glucosidase. Curr Microbiol 32(3):119–123 Xu F (1997) Effects of redox potential and hydroxide inhibition on the pH activity profile of fungal laccases. J Biol Chem 272(2):924–928 Yadav A, Verma P, Kumar R, Kumar V, Kumar K (2017a) Current applications and future prospects of eco-friendly microbes. EU Voice 3:21–22 Yadav AN (2018) Biodiversity and biotechnological applications of host-specific endophytic fungi for sustainable agriculture and allied sectors. Acta Sci Microbiol 1:01–05 Yadav AN, Kumar R, Kumar S, Kumar V, Sugitha T, Singh B, Chauhan VS, Dhaliwal HS, Saxena AK (2017b) Beneficial microbiomes: biodiversity and potential biotechnological applications for sustainable agriculture and human health. J Appl Biol Biotechnol 5:1–13 Yadav AN, Mishra S, Singh S, Gupta A (2019a) Recent advancement in white biotechnology through Fungi. Volume 1: Diversity and enzymes perspectives. Springer, Cham Yadav AN, Mishra S, Singh S, Gupta A (2019b) Recent advancement in white biotechnology through Fungi. Volume 2: Perspective for value-added products and environments. Springer, Cham Yadav AN, Sachan SG, Verma P, Kaushik R, Saxena AK (2016) Cold active hydrolytic enzymes production by psychrotrophic bacilli isolated from three sub-glacial lakes of NW Indian Himalayas. J Basic Microbiol 56:294–307 Yadav AN, Sachan SG, Verma P, Saxena AK (2015) Prospecting cold deserts of north western Himalayas for microbial diversity and plant growth promoting attributes. J  Biosci Bioeng 119:683–693

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Chapter 15

Bioconversion of Biomass to Biofuel Using Fungal Consortium Pavana Jyothi Cherukuri and Rajani Chowdary Akkina

15.1  Introduction The global economy is highly influenced by fuel energy. Global pollution and energy consumption expanded tremendously during the last two decades which led to exhaustion of fossil fuels, resulting in emerging of energy crisis. Therefore, there is a need to search for alternative new energy sources and technologies which have increased logistically in recent years. The entire globe depends on petroleum as a sole energy source. The high usage of petroleum has led to adverse impact on environmental issues such as catastrophic emission of greenhouse gases (Hill et al. 2006). In India, the required petroleum is imported from Middle East; these high imports of petroleum significantly influence the Indian economy. Many attempts are being done to search for alternative fuel sources in transportation sector, such as diesel, gasoline and natural gas (Tabassum Ansari and Choube 2012). But no fuel exhibits unique feature like petroleum such as high energy density, compatibility with vehicles, and being in liquid state. Developed countries like the United States, Europe, Brazil, and China invented new technologies such as solar, hydro, and wind energy usage as an alternative to petroleum or alternative liquid fuels such as butanol, ethanol, methane, and CNG. Among all the liquid fuels, ethanol occupies the first place by its unique properties.

P. J. Cherukuri (*) · R. C. Akkina Department of Microbiology & Food Science and Technology, Institute of Science, GITAM (Deed to be University), Visakhapatnam, Andhra Pradesh, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_15

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15.2  Ethanol Production Ethanol exhibits almost similar properties with petroleum. It can mix with gasoline and reduce emission of smog by unburned hydrocarbons and CO (carbon monoxide) from vehicles. Ethanol blending with gasoline enhances octane number and oxygen content of fuel (Limayem and Ricke 2012). The specific properties of ethanol like high octane number, water free, low flame temperature, high gas volume change, high heat of vaporization, and total combustibility make it convenient to use as automotive fuel (Balan et al. 2013). Ethanol is a transparent liquid, with mild odor boiling temperature at 78  °C and freezing temperature at −112  °C.  Energy Policy Act of 2005 in the United States, demands the blending of 7.5 billion gallons of alternative fuels by 2012. Global rapid depletion of fossil fuels has increased the demand for alternate fuels. The usage and production of ethanol have to be increased markedly (Haghighi et al. 2013). The production of ethanol is not sufficient to meet today’s demand. Industrial ethanol production is obtained from coarse grains (56%), cane (32%), molasses (4%), wheat (3%), nonagricultural substrates (3%), and sugar beet (2%) (Tabassum Ansari and Choube 2012). Economically feasible ethanol production from cellulosic wastes is one of the best ways (Rastegari et al. 2019). Nowadays, integrated biomass system is being used for production of biofuel through biotechnological process. Lignocellulosic biomass provides competitive renewable resource for generation of ethanol in an eco-friendly manner. This technology ensures increased energy security and economic progress and eliminates problems of solid waste management (Das and Singh 2004). According to the studies by Sanchez and Cardona (2008), 73.9 Tg of dry crop waste and 1 × 1010 MT weed biomass is produced annually. Numerous edible crops like cane, corn, beet, and sorghum are used in ethanol commercial production. In this technology, the raw material cost is very expensive. This leads to food insecurity and very high cost of food products. Therefore, plant biomass (lignocelluloses waste) has become an inexpensive carbon source for the production of ethanol. The lignocellulosic wastes such as agricultural residues like corn stover, corncobs, rice straw, wheat straw, sugarcane bagasse, rice husk, wheat husk, and weed biomass like Lantana camara, Prosopis juliflora, Saccharum spontaneum, Eichhornia crassipes, etc. are promising substrates for the production of ethanol.

15.2.1  Demand for Ethanol Production It was estimated that there was 90% increase from the existing levels on the India’s dependency on oil imports by 2020 AD. In order to reduce the oil imports, India proposed an EBP (ethanol blending program report 2014–2015) during the year 2002 with 5% blending of ethanol in petrol which is mandatory in nine major sugarproducing states (Tamil Nadu, Gujarat, Andhra Pradesh, Karnataka, Uttar Pradesh,

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Maharashtra, and Union Territories). In the year 2008, Indian Government introduced a “National Biofuel Policy” for blending of petrol with 20% ethanol (Tabassum Ansari and Choube 2012). To fulfill the supply of bioethanol, it is highly essential to search for new cost-effective raw materials like agricultural residues and weeds. Global ethanol production is around 30 billion liters per year. Global ethnaol production is around 30 billion litters per year, in this, the United States occupies a major share (53%), followed by Brazil (21%), Europe (6%), China (7%) and India (3%). In India, the main source of ethanol production is by molasses (80%) and grains (20%). The production of bioethanol in India is around 2.4 billion liters per annum. The industrial production of bioethanol by biotechnological process using microorganisms and gross productivity ranges from 75% to 90%, and the remaining small fraction of production is from chemical technology by ethylene hydration reactions (Mc. Millan 1997). Currently, ethanol production is by sugarcane and corn in developed countries like the United States and Brazil (Limayem and Ricke 2012). The first-generation biofuels are the results of bioconversion of edible food crops (sugarcane, corn), the second-generation biofuels are from nonedible sources like agricultural and nonagricultural residues, and third-generation biofuels are from algal biomass (Bacovsky et al. 2010; Yewale et al. 2016; Kim and Dale 2004; Chen et al. 2010).

15.3  Biomass for Biofuels Hard wood angiosperms such as a poplar, eucalyptus, and beech wood are rich in cellulose, hemicellulose, and lignin. Efficient conversion of ethanol production from hard wood spent sulfite liquor (HSSL) using Pichia stipitis, Candida shehatae, and Pachysolen tannophilus was reported (Jeffries et al. 2007). HSSL contains high amount of microbial inhibitors. Biofuel production from renewable plant sources is an attractive and alternative process in many countries. Mainly, three substrates like sugars, starches, and cellulose are used for ethanol production in a cost-effective manner. Among these, cellulose materials are renewable and plenty. The agro-­ residues such as sugarcane bagasse, corncobs, corn fiber, corn stover, wheat straw, rice straw, forestry, paper pulp, weed plants, sawdust, sorghum straw, cotton seeds, sunflower seed coats, kitchen waste, and fruit and vegetable waste are collectively known as organic biomass (Lin and Tanaka 2006). The plant biomass is an attractive feedstock for ethanol production consisting of rich carbohydrate composition in various polymeric complex forms (lignin, cellulose, and hemicellulose). Pretreatment process is necessary to use available carbohydrates in biomass. The current technologies are not cost-effective process for commercialization of bioethanol. Elaborate investigations have been done from the past two decades by many researchers for value addition of lignocellulosic biomass (Zhang et al. 2014; Yewale et al. 2016). Weed, starch, and by-products of paper industry were used for ethanol production which includes spent sulfite (Pereira et al. 2013).

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15.4  Composition of Biomass In the present chapter, focus is on biofuel production from lignocellulosic biomass. According to the plant taxonomy, lignocelluloses are classified into soft wood (belongs to gymnosperms), hard wood (belongs to woody angiosperms), and annual plants (herbaceous angiosperms, crops). Biomass of lignocelluloses is a heterogenous mixture consisting of hemicellulose, cellulose, and lignin, and these compositions vary from plant to plant (Yadav et al. 2019a, b). Generally, cellulosic fractions of biomass comprise 40–60% by weight, and long-chain polymers of glucose are bonded together which appear as fiber bundles. Hemicellulose comprises 20–40% by weight and contains short-chain polymers of heterogenous sugars (glucose, galactose, mannose, arabinose, and xylose) and co-jointly binds the cellulose fibers (Bobleter 1994). Lignin consists of 10–30% by weight and contains three-­ dimensional propyl-phenol polymers and contributes rigidity to the entire structure. Lignin is derived from the dehydrogenated products of lignin monomers (p-­coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol). These main aromatic phenol rings of monomer residues of p-hydroxy phenyl, guaiacyl, and syringyl and their composition vary from plant to plant (Campbell and Laherrere 1998). Lignocellulosic biomass shows resistance toward degradation and hydrolysis because of structural firmness. The presence of cross-linkages exists between the cellulose, hemicellulose, and lignin components in biomass. Hemicellulose and lignin are compactly packed cellulosic fraction. For hydrolysis of cellulose, initially digest the hemicellulose and lignin fractions. Cellulosic fraction is protected and occupies the central position in the three-dimensional structure of lignin and hemicellulosic fraction (Balan et al. 2013). For exposure of cellulose by hydrolysis, it is essential to remove the lignin seal. The degradation of cellulosic and hemicellulosic fractions of plant biomass are carried by acid hydrolysis or enzymatic hydrolysis or enzyme producing microrganisms. From these methods, acid hydrolysis requires high temperatures and energy, and this process releases various non-eco-friendly inhibitors (furfurals, hydroxyl furfurals, acids) during the hydrolysis process (Sreenivas et al. 2006). Secondly, enzymes used for the hydrolysis of biomass are eco-friendly and expensive. Among these, bioconversion of biomass using fungal consortium is cost-effective and eco-friendly process (Dirk et al. 2003). The bioconversion of biomass to ethanol requires more processing steps, i.e., pretreatment of lignocellulosic biomass, hydrolysis, and conversion (fermentation/ transformation) of sugars into (biofuel) ethanol. An extensive work has been carried out by utilization of microorganisms (yeast, bacteria, and fungi) for ethanol production. Commercial production of ethanol by fermentation process is highly influenced by fungal and yeast strains. In pretreatment process, initially digest the hemicellulose and lignin crystalline structure and then expose it to cellulose. In the pretreatment process of lignocellulosic matrix, major sugar released from hemicellulosic fraction is xylose, and small amounts of arabinose, galactose, and glucose are also released by the laccase and xylanase enzymes using fungal consortium. Then, cellulose is broken down to monomeric sugars by cellulase-producing fungal

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strains and release of hexose (glucose) sugars. These sugars are converted to ethanol by fermentation process using yeast strains such as Saccharomyces and Candida sp. The yeast Saccharomyces cerevisiae is used for ethanol production from glucose as carbon source. This strain is unable to utilize xylose as carbon source. The Candida and Pichia sp. of yeast are used to convert pentose sugars to ethanol by biotechnological process (Steven and Lee 1990).

15.5  Pretreatment of Lignocellulose Materials The lignocellulosic biomass derived from plant biomass is initially treated by chemical or enzymatic or by both methods. By this degradation, polymeric forms of biomass such as lignin, cellulose, and hemicelluloses are digested to release monomeric sugars. These monomeric sugars are further fermented by microorganisms to produce bioethanol (Claudio et al. 2011). The pretreatment methods are classified into three types: 1. Physical process: Physical process such as steam explosion or auto-hydrolysis, carbon dioxide explosion, ammonium fiber/freeze explosion, and liquid hot water. 2. Chemical process: Chemical process consists of wet oxidation ozonolysis, acid pretreatment, alkaline pretreatment, and organosolv. 3. Biological process employed by microorganism or their enzymes (Buruiana et al. 2013). The pretreatment of starch substrates includes digestion by gasification of lignocellulosic materials by acid or enzymatic hydrolysis for solubilization of cellulose. Novel pretreatment methods such as ultra-sonication, nano-technological methods, and microwave digestion processes are used for pretreatment of lignocellulosic biomass in order to improve the bioethanol production and reduce inhibitors. After this treatment process, the obtained slurry contains two different types of fractions: one is liquid fraction consisting xylose and small amounts of glucose, galactose, and arabinose, and another is solid fraction containing lignin and cellulose material. The potential utilization of these two (hemicellulosic and cellulosic) fractions of biomass is one of the cost-effective method for ethanol production (Agbogbo and Wenger 2007).

15.5.1  Parameters Affecting Pretreatment of Biomass Optimization by screening the efficient strains and maintaining the culture conditions can make this process more effective and reduce the treatment time. The governing process parameters include biomass type, nature, and composition, and physical parameters include incubation temperatures, pH, incubation time, moisture

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content, and aeration rate. Biological parameters such as type of microorganism, culture conditions, growth characteristics, optimum conditions for growth, and enzyme production are the key factors for this process. Lignocellulosic biomass is available abundantly and consists of various proportions of lignin, cellulose, and hemicelluloses. The selection of microorganism for biological treatment is based on the nature and composition of the biomass. The optimum temperature varies with fungal strain such as ascomycetes with optimum temperature at 39 °C while basidiomycetes at 25–30 °C. Incubation time also varies depending on strain and nature of biomass. Long incubation time is required for pretreatment and delignification process. Moisture content plays a key role in solid-­ state fermentations and is essential for growth establishment of microorganisms. White-rot fungi or brown-rot fungi are used for efficient enzymatic saccharification. Pretreatment with fungal strains could enhance enzymatic hydrolysis of lignin. Several studies have revealed that fungal consortium has faster degradation ability of lignocellulosic biomass when compared with single strain. Lignin degradation is an oxidative process, and aeration is a critical factor for production of lignolytic enzymes such as lignin peroxidase and manganese peroxidase (Millati et al. 2011). pH is another important factor for microbial growth; majority of fungal strains grow tremendously in acidic pH range between 4.0 and 5.0. Inoculum level and cell biomass are also critical factors and influence the treatment time of biomass. In SSF (solid-state fermentation), particle size plays a crucial role in biological treatment. Penetration limitations are large with particle size, when compared to small particle size (Kuijk et al. 2015).

15.6  Limitations by Chemical Pretreatment Inhibitors were produced during the digestion of hemicellulosic fraction. These inhibitors are classified into three types: (a) organic acids (acetic acid, formic acid, levulinic acid, ferulic acid, and p-coumaric acid), (b) furan derivatives (furfural and 5-hydroxy furfural), and (c) phenolic compounds (4-hydroxybenzoic acid and ferulic acid) (Jonsson and Martin 2016). All these inhibitors influence the growth and ethanol production efficiency of microorganism (Cragg et  al. 2015). Advanced treatment methods (ultrasonication, microwave digestion) are used to reduce the inhibitor formation and increase the fermentation efficiency of organism. Therefore, detoxification steps are required for removal of inhibitory compounds from hydrolysate before the fermentation process. Detoxification strategies include active charcoal treatment, ion exchange resins, alkali treatment, overliming using calcium hydroxide, change in the fermentation methodologies, and treatment with soft-rot fungi Trichoderma reesei to degrade inhibitors (Yu et al. 2011). The need for efficient pretreatment or hydrolysis process for the recovery of maximum amount of fermentable sugars with minimum toxic chemicals is a major challenge. WRF (white-rot fungi) is one of the promising organisms to convert biomass to bioethanol production with eco-friendly and cost-effective manner. This

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process has ample advantages over chemical process such as simpler technique, less energy utilization, less wastage, and absence of inhibitors. In this process, various strains of WRF that include Pleurotus ostreatus, Cyathus stercoreus, Ceriporiopsis subvermispora, Trametes versicolor, and Phanerochaete chrysosporium are used for conversion of biomass to ethanol production (Nigam and Pandey 2009).

15.7  F  ugal Consortium Used for Biofuel Production from Plant Biomass Several studies have proved that co-culture studies are promising processing methods for production of ethanol from lignocellulosic biomass. Development of fungal consortium has played a significant role for the conversion of polymeric fractions of lignin, cellulose, and hemicellulose into sugars. This consortium includes lignin-, cellulose-, hemicellulose-degrading white-rot fungi and ethanol producing efficient strains. Examples including P. chrysosporium, Pleurotus ostreatus, Pycnoporus cinnabarinus and Cyathus stercoreus are able to produce lignin-degrading enzymes. Laccases are involved in degradation of lignin and show activity with lignin peroxidase and manganese peroxidase (Binod et al. 2010). Cellulose-degrading enzymes endoglucanases, cellobiohydrolase, and β-glucosidease are produced by a number of fungal species. Hemicellulose-degrading enzymes are xylanases and β-xylosidases and produced by Aspergillus niger, Trichoderma reesei fungal strains (Zhang et al. 2012; Kour et al. 2019; Rana et al. 2019a, b). Currently, genetic engineering techniques are widely used for bioconversion of value-added products from lignocelluloses biomass. Transfer of genes encodes xylose reductase and xylitol dehydrogenase from Pichia stipitis to S. cerevisiae (wild strain) for utilization of xylose for enhanced production of ethanol (Agbogbo and Wenger 2007). In industrial scale, it is expensive to maintain the biochemical and fermentative characters of recombinant strains. Due to this disadvantage, a coculture process (both glucose- and xylose-utilizing strains) is cost-effective method for ethanol production along with enzyme-producing white-rot fungi (Cheng et al. 2010). Various factors influencing the production efficiency of the strain include oxygen, aeration, agitation, pH, temperature, concentration of carbon source, inhibitor presence in hydrolysate, and medium components (Sreenivas Rao et al. 2006).

15.8  Enzymes Produced by White-Rot Fungi White-rot fungi produce various extracellular oxidases such as laccase, Mn peroxidase, and lignin peroxidase (LiP) including lignin-modifying enzymes (LME). These enzymes effectively degrade the lignin content in lignocellulosic biomass. Lignin degradation is a key process for biofuel production from lignocellulosic biomass. White-rot fungi (WRF) Phanerochaete chrysosporium produce multiple

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isoenzymes (LiP, MnP) but not laccase. Other WRF were able to produce laccases. Based on the enzyme production, WRF are categorized into three main groups: (1) lignin-manganese peroxidase (P. chrysosporium, Phlebia radiata), (2) manganese peroxidase (Dichomitus squalens, Rigidoporus lignosus), and (3) lignin peroxidase (Phlebia ochraceofulva and Junghuhnia separabilima). These enzymes are able to degrade various types of plant polymers (Heinzkill 1998; Yadav et  al. 2016) (Table 15.1). Table 15.1  Enzymes released by WRF Biomass Corn stover Cotton stalks Milled tree leaves, banana peel, apple peel, mandarin peel Sugarcane trash Sorghum husk Young plant leaves (from Aster genus), lamella from oats and maize plants Sugarcane bagasse

Microorganism Gloeophyllum trabeum KU-41 Phanerochaete chrysosporium Pleurotus spp., Lentinus edodes Aspergillus terreus Phanerochaete chrysosporium Fusarium oxysporum

Aspergillus niger Clavel leaves, young Fusarium merismoides plant leaves (from Aster genus), lamella from oats and maize plants Sugi wood Strobilurus ohshimae Bagasse of cane Pleurotus maize straw ostreatus Grape seeds, barley bran, and wood shavings

Phanerochaete chrysosporium

Clavel leaves, young Clonostachys rosea plant leaves (from Aster genus), lamella from oats and maize plants

Enzyme Cellulase

Reference Gao et al. (2012)

Cellulase

Jian et al. (2008)

Laccase

Songulashvili et al. (2005)

Cellulases

Singh et al. (2008)

Lignin peroxidase and manganese peroxidase Endopolygalacturonases galactosidase

Pankajkumar et al. (2018) Mikan and Castellanos (2004)

Xylanases, cellulases

Park et al. (2002)

Endo-xylanase, cellulases, arabinofuranosidase, acetylesterase

Fernández-Martín et al. (2007)

Lignin peroxidase and manganese peroxidase Xylanases, cellulases, laccase, manganese peroxidase Lignin peroxidase and manganese peroxidase

Homma et al. (2007)

Endopolygalacturonases galactosidase endo-xylanase, cellulases, arabinofuranosidase, acetyl

Márquez et al. (2007), Okamoto et al. (2002) Rodríguez et al. (1997), Srinivasan et al. (1995), Kersten and Cullen (2007), Quintero et al. (2006) Mikan and Castellanos (2004), Rezácová et al. (2006)

(continued)

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Table 15.1 (continued) Biomass Coffee pulp, used nappy, grass residues, cleaned coffee (substrates analyzed separately and in mixture), wheat straw, industrial cotton fiber Clavel leaves, young plant leaves (from Aster genus), lamella from oats and maize plants Wood shaving, carozo maize, and compost of gardening wheat straw

Oat husk Saw dust

Microorganism P. ostreatus, P. pulmonarius

Enzyme Endoglucanase, cellobiohydrolase, laccases, manganese peroxidase

Reference Marnyye et al. (2002), Delfin and Duran de bazúa (2003), Okamoto et al. (2002)

Streptomyces

Cellulases, xylanases, arabinofuranosidase xylosidase, acetylesterase

Mikan and Castellanos (2004), Benimelia et al. (2007)

Trametes versicolor

Laccases

Cerrena unicolor Coriolopsis gallica

Laccases, manganese peroxidase Laccases

Moredo et al. (2003), Márquez et al. (2007), Dumonceaux et al. (2001), Villagran and Renan (1991), Cabuk et al. (2006), Tong et al. (2007) Moilanel et al. (2015) Daassi et al. (2016)

15.9  Ligninolytic Enzymes from White-Rot Fungi Ligninolytic enzymes from WRF are used in industrial biotechnological process. Free enzyme applications were limited in industrial scale due to instability and lack of reusability. Immobilization techniques were used to improve stability and can be reusable. Voberkova et  al. (2018) have reported that immobilization methods are desirable, operational stability and cost-effective process. The delignification is a key task for proper utilization of lignocelluloses biomass. The first demonstration of commercial plant for the ethanol production from lignocellulosic biomass is in operation in Canada since 2004 (Tampier et  al. 2004). White-rot fungi is a filamentous fungi, which can produce ligninase, cellulase enzymes for degradation of lignocellulosic plant material. Pleurotus cystidiosus, P. ostreatus, Phlebia, Ganoderma lucidum, and Flammulina velutipes are able to produce ethanol from biomass. Various WRF are used for production of enzymes, which degrade the lignocellulosic material. Integrated production of ethanol by WRF is influenced by factors like moisture content, temperature, pH, and chemical nature. Various reports of biological pretreatment of lignin are presented in Table 15.2.

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Table 15.2  Lignin degradation by fungal strains Microorganism Merulius tremellosus Phanerochaete chrysosporium Trichoderma viride Trichoderma reesei Ceriporiopsis subvermispora Irpex lacteus Phanerochaete chrysosporium

Pleurotus ostreatus Pleurotus ostreatus Ceriporiopsis subvermispora Fusarium spp.

Biomass Effect Aspen wood 0.26–0.37 mg/ml as compared to 0.15–0.16 mg/ml of control Cotton stalks 0.027 g/g

Reference Bradley et al. (1989) Jian et al. (2008)

Rice straw

Ghorbani et al. (2015) Barakat and Rouau (2014) Wan and Li (2011) Du et al. (2011) Liu et al. (2014)

56% of lignin reduction

Wheat straw Sugar Yield -270 Reduction (mg/g dry substrate) Corn stove 2- to 3-fold increase in reducing sugar yield Corn stalks 82% of hydrolysis yield Corn stover Improved degradation silage of substrate cell wall components 39% lignin removal of initial substrate Wheat straw 35% of lignin reduction Rice straw 33% lignin removal Corn stover

31.59% lignin loss

Paddy straw

17.1% decrease in lignin content, 10.8% decrease in silica content compared with controls

(1983) Mustafa et al. (2016) Wan and Li (2010) Phutela and Sahni (2012)

15.10  Biochemical Pathway of Ethanol by Fungi The yeast cells have the sense to identify the sugar-rich environment. This intensity by the yeast can affect the enzyme activity during biochemical processes, change of translation by mRNA, stability of protein degradation, and concentration of metabolites (Yadav et  al. 2017, 2018). After glucose uptake then enters the glycolytic pathway and is converted to pyruvate and produces ATP and then coupled to intermediate products and reducing power through NADH for biosynthetic pathway. Pyruvate in glycolysis enters TCA cycle or fermentative pathway. In alcoholic fermentation, decarboxylation of the pyruvate gives acetaldehyde by pyruvate decarboxylase enzyme. This enzyme converts acetaldehyde to ethanol by reduction of NADH to NAD+, from one molecule of glucose to two CO2 molecules and ethanol is formed. The second abundant sugar, from hemicellulosic fraction of biomass, is xylose. Xylose is a five-carbon sugar molecule, utilized through pentose phosphate pathway in fungi. The xylose transportation in fungi is followed by two different mechanisms. Pichia stipitis and Candida spp. follow proton symport (PS) mechanism, whereas Saccharomyces cerevisiae follows facilitated diffusion system (FDS) for transport of xylose. The PS transport is for pentose sugars and FDS transport for both hexose and pentose sugars. The medium consisting of low quantities of hex-

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Lignocellulosic biomass Pretreatment Cellulose

Hemicellulose

Cellulases

Cellobiose Cellodextrin transporter Cellobiose β-glucosidase

Glucose

Xylose Xylose XR Xylitol XDH Xylulose XK PPP Glycolysis

Acetate Acetate ACS Acetyl-CoA AADH Acetaldehyde ADH Ethanol

Ethanol

Fig. 15.1  Biochemical pathway of ethanol production by fungi

oses will inhibit xylose transport by FDS mechanism. In fungi xylose metabolism, xylose is converted to xylulose through xylitol by xylitol dehydrogenase, and then xylulose is phosphorylated and enters to pentose phosphate cycle. Conversion of xylose to xylitol in the presence of xylose reductase utilizes NADH or NADPH as a cofactor. In anaerobic or microaerophilic conditions, yeast utilizes NADH for conversion. High expression of xylose reductase and xylitol dehydrogenase will tend to enhance the ethanol production. Co-current isomerization of xylose and co-fermentation of xylose and glucose increase the production of ethanol (Fig. 15.1).

15.11  Conclusion and Future Prospects Overcoming the challenges of fossil fuels through the biotechnological route by fungal consortium is one of the promising approaches to reach global demand. The lignocellulosic biomass is a significant substrate for bioethanol production using fungal consortium for commercial production of ethanol throughout the year in a cost-effective process. This biotechnological approach of ethanol production is an eco-friendly process through enhancing the yield and reduction of the greenhouse gas emission. Saccharifications of lignocellulosic biomass by white-rot fungi have overcome the challenges with chemical digestive methods such as gasification and acidification. The chemical process using high temperatures, acids, and high-energy

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input lead to release of inhibitors and pollutants. The development of fungal consortium consists of lignin-, cellulose-, and hemicellulose-degrading strains in combination with ethanol-producing strains (from hexoses and pentoses) which can achieve the global demand for ethanol production. The enzymes like laccases, cellulases, and xylanases play a significant role for the digestion of lignocellulosic plant biomass by eco-friendly manner. Several advantages were reported by fungal consortia which include high adaptability, productivity, and efficiency of the production. This process is also considered as inexpensive and eco-friendly. The future prospect for ethanol production using fungal consortium is a need to development of unique fungal consortia with noncompetitive synergistic fungal strains for production of lignocellulosic digestive enzymes along with efficient ethanol-producing strains. Standardization of co-culture studies for optimization of digestive enzymes and ethanol production, optimized conditions for microbial growth, and metabolite production (enzymes) were varying from one strain to another. In co-culture studies (development of fungal consortium), physical factors (temperature, pH, agitation, moisture levels, surface area, and SSF) and chemical factors such as carbon source, nitrogen sources, minerals, salts, and their concentrations need to standardize for commercial production of ethanol. Simultaneous saccharification and fermentation have enhanced the yield. Acknowledgments Authors sincerely acknowledge University Grant Commission (UGC), Government of India, for providing financial support for major research project entitled “Overcoming fossil fuel challenges: Co-culture fermentations for biofuel production using agro-­ industrial waste materials.”

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Chapter 16

Role of Fungi in the Removal of Heavy Metals and Dyes from Wastewater by Biosorption Processes Ajay Kumar, Vineet Kumar, and Joginder Singh

16.1  Introduction World population is increasing day by day; hence, to meet the demand of the growing population, clean water is the major concern. Water is being polluted by human activates and industrial discharges. These pollutants are categorized into three major groups: organic, inorganic, and biological particles. Heavy metals and dyes as waste from various industries including textile, pharmaceutical, leather, etc. are the major pollutant present in water (Burakov et al. 2018). Heavy metal ions are elements from the fourth period of the periodic table, mostly chromium (Cr), cobalt (Co), nickel (Ni), copper (Cu), zinc (Zn), arsenic (As), lead (Pb), and mercury (Hg). Removing heavy metals is necessary because they are toxic substances with carcinogenic nature that should not to be discharged directly into the environment. Conventional techniques like membrane separation, precipitation, coagulation, and flocculation are widely used for removal of heavy metals (Azimi et al. 2017; Marzougui et al. 2017). Biosorption is preferred over these conventional techniques due to high affinity, capacity, and selectivity of the materials from the solution. There are different mechanisms involved in biosorption phenomenon (Fig. 16.1). There are different kinds of adsorbent available for wastewater treatment. Adsorbents are broadly classified into conventional and nonconventional. Biosorbents are nonconventional adsorbents and have several advantages over other conventional and nonconventional methods (Fig. 16.2). Microorganism is one type of biosorbents that has been used for wastewater treatment. Many living or dead microorganisms such as bacteria, fungus, and

A. Kumar (*) · V. Kumar School of Bioengineering and Biosciences, Lovely Professional University, Phagwara, Punjab, India J. Singh Department of Biotechnology, School of Bioengineering and Biosciences, Lovely Professional University, Jalandhar, Phagwara, Punjab, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_16

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Fig. 16.1  Different mechanisms involved in biosorption phenomenon (Asgher 2012)

algae are widely used for heavy metal removal because of high adsorption capacity, low cost, and its availability in large quantities (Kour et al. 2019a; Rastegari et al. 2019). However, it suffers from some of the drawbacks like waste may be converted into more potential toxic compounds. Most of the microorganism-based methods deal with discoloration of dye instead of removal of dye or other wastes. Dyes are mostly used in textile, leather, and pharmaceutical industries. Discharge of dyes directly into the water bodies causes threats to aquatic fauna and flora as it interferes with gas solubility. Higher concentration of dyes resulted in carcinogenicity and toxicity. Use of fungi for wastewater treatment is one of the promising alterative to physical and chemical process (Couto 2009; Yadav et al. 2016, 2019b). Classification of dyes according to the chemical structure is depicted in Fig. 16.3. However, nanomaterial-based approaches of wastewater treatment are considered more useful as they allow comparatively better removal of wastewater (Masoudi et al. 2018). With advances in nanotechnology, more and more useful nanomaterials such as graphene, carbon nanotubes, and fullerenes are being produced for wastewater treatment. Along with wastewater treatment, these nanomaterials are also useful for various other environmental applications (Hegab et al. 2018; Nyairo et al. 2018; Park et al. 2017). Figure 16.4 shows nanomaterials used for heavy metal treatment of water. Synthesis of bionanocomposite is now in practice for wastewater treatment. Wang et al. (2018) reported the removal of methylene blue by adsorption on yeast composite assisted with Fe2O3 nanoparticles. Figure 16.5 shows the scheme diagram for the synthesis of bio-nanocomposites.

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Fig. 16.2  Various methods for wastewater treatment methods (Crini et al. 2018)

16.2  Fungi as Biosorbent Fungi are osmo-heterotrophic eukaryotes placed in the kingdom Fungi. The fungal cell wall is composed of acid polysaccharides such as chitin (a polymer of acetylglucosamine unit), and chitosan, which is characterized by phosphate, amine, and

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Fig. 16.3  Classification of dyes according to the chemical structure (Yagub et al. 2014)

Fig. 16.4  Nanomaterials for heavy metal treatment of water (Lu and Astruc 2018)

hydroxyl groups, is involved in biosorption of heavy metals, dyes, and phenolic compounds (Zhu et al. 2019). They lack chlorophyll and their vegetative structure may be filamentous or unicellular. They reproduce through spore formation (Raghukumar 2017). Figure 16.6 shows the structure of chitin (Fig 16.6a) and chitosan (Fig 16.6b) and their binding with metal ions. Table 16.1 shows the characteristics of major fungal divisions.

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Fig. 16.5  The scheme diagram for the synthesis of bio-nanocomposites

Fig. 16.6 (a) The units of a chitin polymer molecule. (b) Chitosan is deacetylated chitin. (c) Binding of metal anions on chitin or chitosan (Kotrba 2011)

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Table 16.1  Characteristics of major fungal divisions (Stajich et al. 2009) Division Chytridiomycota Zygomycota

Ascomycota

Basidiomycota

Deuteromycota

Characteristics The fungi produce zoospores capable of moving on their own through a liquid medium by simple flagella The hyphae do not have one nucleus per cell but rather have long multinucleate, haploid hyphae that comprise their mycelia. Asexual reproduction is by spores produced in stalked sporangia They contain more than 30,000 species of unicellular (yeasts) to multicellular fungi. Yeasts reproduce asexually by budding and sexually by forming a sac/ascus Mushrooms, toadstools, and puffballs are commonly encountered basidiomycetes. These conspicuous features of the fungi are the reproductive structures. Sexual reproduction involves the formation of basidiospores on club-shaped cells known as basidia A group of fungi that either lack the perfect stage (i.e., sexual reproduction) or whose perfect stage is as yet undiscovered. They reproduce most frequently by conidia or conidia-like spores. Many forms of deuteromycota are pathogenic, affecting man, animals, or plants

For removal of dyes from wastewater, different forms of fungal sorbents are used such as fungal pellets, mycelium, or dead fungus by many researchers (Yagub et al. 2014). Many molds and filamentous microorganisms such as Aspergillus niger, Penicillium simplicissimum, Aspergillus fumigatus, Termitomyces clypeatus, Penicillium brevicompactum, Saccharomyces cerevisiae, Trichoderma, etc. are used for removal of heavy metals and dyes (Rana et al. 2019a, b; Yadav et al. 2019a, b). Fungus can survive in the presence of high metal concentration. So it can be used for heavy metal removal from wastewater. Heavy metal adsorption by fungi through ion exchange and coordination is due to the presence of chitin–chitosan, glucuronic acid, phosphate, and polysaccharides present in/on the cells of fungi. Different kinds of functional groups such as amine, carboxyl, hydroxyl, phosphate, and sulfhydryl pay a vital role in the adsorption of heavy metals and dyes by fungal stains (Yin et al. 2018). Fungi especially white-rot fungi and their enzymes (laccase, lignin peroxidase, and Mn peroxidase) can be used to bioremediate various xenobiotics and wastewaters (Kour et al. 2019b; Yadav et al. 2017a, b; 2018). Figure 16.7 shows the schematic diagram of adsorption mechanism model.

16.3  Growth Models for Filamentous Organisms Fungal biomass can be easily cultivated, or it can be available as industrial waste product such as Aspergillus niger (waste from citric acid production) and Saccharomyces cerevisiae (brewery industry waste) (Dhankhar and Hooda 2011). At high cell density, filamentous organisms such as molds often form microbial pellets in suspension culture. During the growth process, filamentous organism

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Fig. 16.7 (a) Schematic diagram of adsorption mechanism model. (b) SEM images of fungus mycelia and (c) dyes adsorbed onto fungus mycelia (Li et al. 2019)

increases in their size and mass. Thus, in the absence of mass transfer limitations, the radius of the microbial pellet increases linearly with time.



dR = k= cons tan t p dt

(16.1)

The growth rate of mold colony can be expressed as follows:



dM dR = ρ 4π R 2 = kp 4π R 2 ρ dT dt

(16.2)



dM = γ M 2/3 dT

(16.3)

where γ = kp(36πρ)1/3 The mass of spherical pellet as a function of time is given as follows: 3



3

γt  γt   M =  M 01/ 3 +  ≈   3 3 

(16.4)

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where M0 is the initial mass which is very small as compared with the M and therefore M varies with cubic power with time.

16.4  Surface Modification of Fungal Biomass Surface modification of biomass is one of the strategies used for adsorption of heavy metals and dyes. Various pretreatment methods such as acid, base, and thermal treatment are used for surface modification of biomass to enhance the adsorption capacity of biomass (Yin et al. 2018) as shown in Fig. 16.8.

16.4.1  Acid Pretreatment Biomass treated with acid improved the positive charge density on the surface which provide strong electrostatic attraction for negatively charged heavy metal ions.

16.4.2  Base Pretreatment Biomass treated with alkali may increase negative charge on the surface of biomass to enhance the electrostatic attraction for positively charged heavy metal ions.

Fig. 16.8  Enhancing heavy metal removing efficiency of biomass through surface modification (Yin et al. 2018)

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16.4.3  Thermal Treatment The porosity and surface area of biomass can be enhanced by thermal treatment for adsorption capacity of biomass. The thermal treatment can also increase the surface functional groups by adding metal-binding groups.

16.5  Biosorption Models and Isotherms Biosorption is defined as the removal of substance from biological materials whether it is living or dead which involves the phenomenon of the mass transfer. Biosorption involves both the adsorption and absorption processes. Sorption mechanism can be divided into four consecutive steps: (i) Transport of solute in the bulk solution (ii) Diffusion of solute through the liquid film surrounding the adsorbent particles (iii) Diffusion of solute in the pores of the sorbent (intraparticle diffusion) (iv) Chemical reaction as adsorption and desorption on the solid surface

16.5.1  Adsorption Thermodynamics Thermodynamic behavior of heavy metals and dyes on biosorbent is cited as exothermic or endothermic sorption processes. Free energy gives the information about the physical sorption or chemical sorption (Yao et al. 2010; Madala et al. 2017).

∆G  = − RT ln K

(16.5)



∆G  = ∆H  − T ∆S 

(16.6)



ln K =





∆S ∆H − R RT

(16.7)

where ∆G0 (J/mol) is Gibb’s energy, R is the ideal gas constant (8.314 J/mol K), T is temperature in Kalvin (K), ∆S0 (J/mol K) is adsorption entropy, and ∆H (J/mol) is adsorption enthalpy. K can be obtained from qe/Ce, while the values of ΔH0 and ΔS0 were determined from the slope and intercept of the van’t Hoff plot of ln K versus 1/T. The negative values of ΔGo suggested the spontaneous behavior of adsorption process. The positive values of ΔHo indicate the endothermic process for the adsorption of metals and dye. The positive value of ΔSo suggested the increasing randomness between the solid and solution interface during the adsorption process.

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16.5.2  Adsorption Isotherm Several mathematical models have been developed by the researchers to validate the process of adsorption (Lei et al. 2018). Kumari and Abraham (2007) describe biosorption of anionic textile dyes by nonviable biomass of fungi and yeast. The Freundlich model assumes that the adsorbent surface is heterogeneous and sorption on its surface is multilayer.

qe = K f c1/e n

(16.8)



1 ln qe = ln K f + ln ce n

(16.9)

where Ce (mg/L) is the equilibrium concentration in solution, qe (mg/g) is the lead adsorbed at equilibrium, n is Freundlich constant related to adsorption intensity, and Kf is adsorption constant for Freundlich model. Thermodynamic parameters of adsorption of silver onto biochar were calculated from Langmuir isotherm as documented by Antunes et al. (2017). qe =

bqm ce 1 + bce

(16.10)

The linear form of Langmuir isotherm model equation



Ce Ce 1 = + qe qm ( qm .b )

(16.11)

where ce (mg/L) is the equilibrium concentration of Cu(II), qe (mg/g) is the adsorption capacity, qm (mg/g) is the theoretical maximum sorption capacity, and b (L/mg) is the Langmuir constant related to adsorption energy. The plot of Ce/qe against Ce gives a straight line with a slope and intercept of 1/qm and 1/qmb, respectively. The separation factor, RL, can be determined from Langmuir plot as per the following relation: RL =

1 (1 + bC0 )

(16.12)

where RL values indicate the type of adsorption to be irreversible (RL = 0), favorable (0 20), metals are termed as heavy metals. Some heavy metals are essential nutrients (Fe, Co, Zn), relatively harmless (Ru, Ag, and Id), but potentially can be toxic in larger amounts or certain forms. Conversely, heavy metals, such as Cd, Hg, and Pb, are highly poisonous. The common source of heavy metals is antiseptics, fertilizers, sedimentation, cars, golf clubs, mobile phones, plastics, selfcleaning ovens, solar panels, and particle accelerators (Gupta et al. 2018; Singh et  al. 2013;  Hübner et  al. 2010; Singh et  al. 2017; Yadav et  al. 2018b, c) (Table  18.1). The potential sources are atmospheric deposition; automobile exhausts, metal industries, mine spoils, river dredging and urban refuse disposal, pyrometallurgical industries, and fossil fuel combustion are also the main sources of heavy metals (Lottermoser 2010a, b; Matta et  al. 2018; Prasad 2001) (Table 18.1). Industries such as microelectronics, plastics, refinery textiles, wood preservatives, agrochemicals (fertilizers and pesticides), sugar-based industries and waste disposal sewage sludge, landfill leachate, and fly ash disposal are also some of the chief sources of the heavy metals (Bhatia et  al. 2015; Gupta et  al. 2018; Singh et al. 2013a; Kumar et al. 2016; Singh and Kumar 2006; Yadav et al. 2018b, c) (Fig. 18.1).

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Table 18.1  Sources of heavy metals and respective anthropogenic activities (adapted and modified from Yadav et al. 2017) Heavy metals Antimony (Sb) Arsenic (As) Beryllium (Be) Cadmium (Cd) Chromium (Cr) Cobalt (Co) Copper (Cu) Iron (Fe)

Anthropogenic activities Alloys, Britannia metal, electrical applications, flame-proof pigments and glass, pewter, medicines for parasitic diseases, queen’s metal, semiconductors Geogenic processes, fuel, smelting operations, thermal power plants Alloy, electrical insulators in power transistors, moderator, nuclear power plants e-waste, incinerations and fuel combustion, paint sludge, waste batteries, Zn smelting Mining, industrial coolants, chromium salt manufacturing, leather tanning

Ceramics, glass industry, metallurgy (in super alloys), paints Mining, electroplating, smelting Alloys, cast iron, construction, machine manufacturing, steel, transportation, wrought iron Lead (Pb) Alloys, antiknock agents, cable sheathings, ceramics, glassware, lead-acid batteries, plastic, ordinance, pigments, solder, tetramethyl lead, pipes Manganese Alloys, antiknock agents, batteries, catalysts, coating welding rods, (Mn) ferromanganese steels production, fungicides, pigments, dryers, wood preservatives Mercury(Hg) Catalysts, dental fillings, fungicides, electrodes, electrical and thermal measuring apparatus, metals extraction by amalgamation, mobile cathode production, mercury vapor lamps, pharmaceuticals, oscillators, scientific instruments, solders, rectifiers, X-ray tubes Molybdenum Alloys, cast irons, catalysts, corrosion inhibitors, dyes, electroplating, flame (Mo) retardants, lubricants, nonferrous metals, smoke Nickel (Ni) Alloys, arc-welding, catalysts, computer components, electroplating, Ni/Cd batteries, paint pigments and ceramics, rods, surgical and dental instruments, ceramic molds, and glass containers Selenium (Se) Dandruff treatment glass industry, inorganic pigments, lubricants, photoelectric and photo cells, rubber production, semiconductors, stainless steel, thermo-­ elements, and xerographic materials Stannum (Sn) Brasses, bronzes, catalysts, dental amalgam, pesticides, pewter, stabilizers, tin-plated steel Titanium (Ti) Ti as alloy in aeronautics nucleation, catalyst, deep temperature thermometers, electronics industry, glass ceramics, infrared optical systems, low melting glasses, semiconductors, supraconductors, UV-filtering agents, white pigments Vanadium (V) Alloys, catalyst, steel production Zinc (Zn) Electroplating, smelting

18.2.2  Water Resources Water contamination due to heavy metals is a known threat and has been attributed to anthropogenic sources involving untreated domestic and industrial wastewater discharges, chemical spills, and agricultural residues (Malyan et  al. 2014;

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Fig. 18.1  Overview of sources of heavy metal pollution and its agroecological consequences. (Source: Srivastava et al. (2017))

Tchounwou et al. 2012). The outcome is poor water quality, degradation, and water borne-human health risks even at lower doses of heavy metals (Kumar et al. 2014; Micó et al. 2006; Wongsasuluk et al. 2014). Major heavy metals such as lead, mercury, chromium, cadmium, copper, and aluminum for water contaminations are originated through anthropogenic activates and natural incidents like seepage from rocks, volcanoes, and forest fires. Over a time period, heavy metals enter in the food chain through water, and there chronic effects could be manifested for many years and may exert several threats such as mental disorders, pain in joints, gastric disorders, and even cancer. Human population living near industries are more susceptible to heavy metal toxicity. Along with that, pregnant women and malnutritioned children are more vulnerable to heavy metal toxicity. Freshwater bodies are heavily affected by pathogens from untreated wastewater and heavy metals from mining and industrial release (Caravanos et  al. 2016). It has been reported that more than 80% of the world’s wastewater is released to the environment without treatment, which is the major cause of nearly 58% diarrheal disease (major cause of child mortality) (Connor et al. 2017). Hence, it is of utmost importance in the coming future to mitigate this global threat of water toxicity with proper remediation measure, and techniques are required for the treatment of water. In that context, fungal phytoremediation serves as an environment-friendly, pocket-friendly, and reliable technique.

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18.3  Role of Heavy Metals in Living Beings Heavy metals such as chromium (glucose metabolism), cobalt (metabolism), copper and iron (oxygen and electron transport), zinc (hydroxylation reactions) (Nieboer and Richardson 1978), manganese and vanadium (enzyme regulation), nickel (cell growth), and selenium (antioxidant and hormone production) (Emsley 2011) are important for certain biological processes. Molybdenum (catalysis of redox reactions), cadmium (in marine diatoms), tin (growth in a few species), and tungsten (metabolic processes of archaea and bacteria) may be required for growth of different species (Emsley 2011). A deficiency and excess of any of these above-discussed heavy metals may impart heavy metal poisoning of living beings (Venugopal and Luckey 1978). Hence, excess amount of heavy metals could dysfunction various physiological and biological effects in the human beings which have been elaborated in next sections.

18.4  Possible Impacts of Heavy Metal Contaminations 18.4.1  On Humans Non-essential metals can escape control mechanisms such as binding to specified cell constituents, cellular processes malfunctioning, compartmentalization, homeostasis, oxidative deterioration, and transport, and therefore, they have toxic and further lethal effects (Gupta et  al. 2018). The important health symptoms of heavy metal toxicity in human are central nervous system disorders, dementia in adults, emotional instability, insomnia, intellectual disability in children, kidney diseases, liver diseases, depression, vision disturbances, and increased morbidity and mortality rate (Jain et al. 2015; Yadav et al. 2018b, c). The metal toxicity depends on the generation of oxidative stress (increased reactive oxygen species (ROS) and reactive nitrogen species (RNS) production; depletion of intracellular antioxidant stores and free radical scavengers) (Jan et al. 2015). Heavy metals toxicity due to occupational exposure mainly responsible for multiple organ systems and toxicity levels mainly depends on the form and type of the heavy element, on route and duration of the exposure, and, to a greater extent, on a person’s individual susceptibility (Jan et al. 2015) (Fig. 18.2).

18.4.2  On Plants Heavy metal contamination in soil and water resources affects growth and yield performance as well as nutritional quality of plants to a great extent. For the plants which are grown in close vicinity to the contaminated soil and water or at the contaminated site, metals cause physiological dysfunctioning and biochemical

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Fig. 18.2  Trophic transfer of toxic HMs from soil to plants to humans and organism’s food to humans and their toxicity. (Adapted with permission from Saxena et al. (2019))

alterations (Sharma et al. 2012a; 2012b). In case of vegetables requiring high moisture percentage, the use of heavy metal-contaminated irrigation water is one of the major causes for high metal toxicity in plants. Some of the heavy metals at a lower concentration are required for optimum performance of plants; however, excess amount may cause toxicity, e.g., chromium (Yadav et al. 2018b, c). Common features pertaining to metal toxicity are reduced biomass reduction, leaf chlorosis, and root growth and seed germination inhibition (Ghani 2011). Cr toxicity considerably affects the physio-biochemical processes in barley, cauliflower, citrullus maize, wheat, and vegetables (Ghani 2011). ROS signalling and oxidative damage affect enzymes like catalase; cytochrome oxidase and peroxidase with iron as their component are affected by chromium toxicity. The catalase activity stimulated with an excess supply of chromium-inducing toxicity has been studied, concerning nitrate reductase activity, photosynthesis, photosynthetic pigments, and protein content in algae (Nath et al. 2008). Pb and Cd also affect the gas exchange attributes, ROS system, cause chlorophyll deterioration, and ultimately the overall performance of major agricultural crop worldwide (Anjum et al. 2015; Mobin and Khan 2007; Pinho and Ladeiro 2012; Zhu et al. 2007). The microbes are ubiquitous in nature and have been reported from diverse sources including extreme habitats (Yadav et al. 2015a, b, c, 2017b) and as plant microbiomes (Kour et al. 2019b; Yadav 2018; Yadav et al. 2016). These microbes have potential applications in agriculture, industry, pharmaceutical, and environment (Kour et al. 2019a; Yadav et al. 2017a, 2018a, 2019a, b).

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18.5  P  hytoremediation of Contaminated Soils/Water Resources In general, phytoremediation is the process of bioremediation using plant species called hyperaccumulators to reduce the toxic contaminants in the environment. This is a novel advanced technology, considered as eco-friendly having lesser investment cost. Current scenario explains the feasibility and accountability of this technique. Many plant species are being used as hyperaccumulators, and new species are being explored (Ali et  al. 2013). Eventually, phytoremediation is an interdisciplinary branch that requires knowledge for soil composition, soil microbiology and environment engineering, plant physiological processes, and in recent development use of lower plant groups as a sustainable system for the bioremediations of toxic heavy metals (Pisani et al. 2011). Some of the species of the plants used in phytoremediation are Robinia pseudoacacia and Sesbania drummondii for Pb, Stanleya pinnata for Se, etc. (Yang et al. 2016).

18.6  Fungal Phytoremediation As its name explains, fungal phytoremediation or mycoremediation is a form of bioremediation where the degradative abilities of fungi are utilized to remove or neutralize the harmful contaminants present in soil and water. It is a relatively new form of bioremediation where its use only spans a few decades, beginning as early as 1966 (Matsumura and Boush 1966), but it is known or being practiced to a lesser extent. Malathion (an insecticide and neurotoxin) breakdown was successfully done using Trichoderma viride and Pseudomonas (Matsumura and Boush 1966). There are several mushroom species identified till date to remove the heavy metals from the contaminated resources. The important species are Galerina vittiformis (Cu, Cd, Cr, Pb, and Zn), Hypholoma capnoides (Ti, Sr, and Mn), and Marasmius oreades (bismuth and titanium). The other important fungal species which are having high fungal phytoremediation potentials are Agaricus bisporus, Lentinus squarrosulus, Phanerochaete chrysosporium, Pleurotus ostreatus, Pleurotus tuber-regium, P. ostreatus, P. pulmonarius, and Trametes versicolor (Adenipekun and Lawal 2012; D’Annibale et al. 2005). In this chapter, the sources of different heavy metals (HMs) with adverse effects in major countries on human health along with the permissible limits of HMs has been highlighted to have the understanding on the current scenario of fungal ­phytoremediation works (Table 18.2). Similarly, the different groups of fungus having remediation potential for the most potent heavy metals have been highlighted in Table  18.3. Further, the categorical classification of different fungus and their importance in particular metal have been worked out with extensive literature survey in order to target potential fungal phytoremediation techniques for the metal

Pulmonary, skin

Ni

Industrial effluents, kitchen appliances, surgical instruments, steel alloys, automobile batteries

Release from Au-Ag mining Nervous system, and coal combustion, medical renal waste

Hg

Liver, kidney, blood

Pesticides, fertilizers

Cu

Cr

Paints and pigments, plastic stabilizers, electroplating, incineration of Cd-containing plastics, phosphate fertilizers Tanneries, steel industries, fly Pulmonary ash

Cd

Target organs Pulmonary, nervous system, skin Renal, skeletal pulmonary

Sources Pesticides and wood preservatives

HMs As

Major world mine Harmful effects countries China, Chile, Morocco, As (especially as arsenate) is a phosphate analog which interferes with oxidative phosphorylation and Russian Federation ATP synthesis China, Korea, Japan, Cd is carcinogenic, mutagenic, teratogenic, and Mexico, Canada endocrine disruptor; Cd interferes with calcium regulation in livings; renal failure and chronic anemia Cr causes hair loss, nephritis, cancer, and ulceration South Africa, in humans Kazakhstan, India, Turkey, Russian Federation Chile, China, Peru, Elevated Cu levels may cause brain and kidney damage, liver cirrhosis and chronic anemia, stomach Australia, the United States and intestinal irritation China, Kyrgyzstan, Anxiety, autoimmune diseases, depression, Chile, Russian balancing difficulty, drowsiness, fatigue, hair loss, Federation insomnia, irritability, memory loss, recurrent infections, restlessness, vision disturbances, tremors, temper outbursts, ulcers and damage to brain, kidney and lungs Allergic dermatitis known as nickel itch; inhalation Philippines, Russia, Brazil, Indonesia, can cause cancer of the lungs, nose, and sinuses; Canada, Russia cancers of the throat and stomach have also been attributed to its inhalation; hematotoxic, immunotoxic, neurotoxic, genotoxic, reproductive toxic, pulmonary toxic, nephrotoxic, and hepatotoxic; causes hair loss 0.02

2.0

–

0.2

–

1.3

2.0

–

(continued)

0.05

5.0

HMs limitation in (ppm) EPA WHO 0.10 –

Table 18.2  Sources of heavy metals (HMs) and their target organs in human with adverse effects in major countries along with the permissible limits of HMs

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Fertilizers

Industrial dust and fumes

Zn

Mn

Major world mine Target organs Harmful effects countries China, Australia, the Nervous system, Impaired development in children, reduced United States, Peru, intelligence, loss of short-term memory, learning hema-­topoietic disabilities and coordination problems; causes renal Mexico system, renal failure; increased risk for development of cardiovascular disease Brain, respiratory Over dosage can cause dizziness and fatigue China, Australia, Peru, tract India, the United States Nervous system Central and peripheral neuropathies South Africa, Australia, China

Sources: Ali et al. (2013); Mahurpawar (2015); Yadav et al., 2017; Gupta et al. 2018; Rajendran et al., 2003; USGS, 2012 

Sources Aerial emission from combustion of leaded petrol, battery manufacture, herbicides and insecticides

HMs Pb

Table 18.2 (continued)

–

0.5

–

–

HMs limitation in (ppm) EPA WHO 15 0.01

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Table 18.3  Categorical classification of fungal species targeting metal remediates for fungal phytoremediation Species Agaricus bisporus

Metals remediate References Ni, Cu, Pb, Mn, Cd, Nagy et al. (2014) Zn, Hg, Fe Agaricus bitorquis Cu, Zn, Fe, Cd, Pb, Ni, Lamrood and Ralegankar (2013) Alternaria alternata Cd, Cr, Cu, Ni Seshikala and Charya (2012) Armillaria mellea Ni, Cu, Pb, Mn, Cd, Zn Ita et al. (2008) Ascochyta betae Cr Seshikala and Charya (2012) Aspergillus fumigatus Cu, Cd, Ni, Co, Pb Rao et al. (2005) Aspergillus flavus Zn, Cu, Ni, Pb Thippeswamy et al. (2012a) Aspergillus foetidus Cr Prasenjit and Sumathi (2005) Aspergillus fumigates Pb Kumar Ramasamy et al. (2011) Aspergillus niger Cd, Pb, Zn, Cu, Ni, Cr, Pal et al. (2010) Aspergillus ochraceus Cr Seshikala and Charya (2012) Aspergillus oryzae Cr Nasseri et al. (2002) Aspergillus terreus Pb, Cu, Ni, Cr Seshikala and Charya (2012) Aspergillus versicolor Cr, Ni, Cu Taştan et al. (2010) Aspergillus versicolor Pb Çabuk et al. (2005) Candida tropicalis Zn Akhtar et al. (2008) Candida utilis Cr Pattanapipitpaisal et al. (2001) Circinella sp. Ni Alpat et al. (2010) Cladonia rangiformis (lichen) Pb Ekmekyapar et al. (2012) Cladosporium resinae Cu Gadd and de Rome (1988) Cunninghamella echinulata Pb, Ni, Zn Shouaib et al. (2011) Curvularia lunata Cu, Cr, Cd Seshikala and Charya (2012) Drechslera rostrata Cr Seshikala and Charya (2012) Fusarium oxysporum Cr Amatussalam et al. (2011) Fusarium solani Cr, Zn, Ni Sen and Dastidar (2011) Ganoderma lucidum Cu Muraleedharan et al. (1995) Ganoderma lucidum, Penicillium sp. Ar Loukidou et al. (2003) Gliocladium sp. Cu Tahir (2012) Lactarius piperatus Cd Nagy et al. (2014) Lentinus edodes Cd, Pb, Cr Tu and Huang (2005) Metarhizium anisopliae Pb Çabuk et al. (2005) Mucor hiemalis Cd, Cu Srivastava and Hasan (2011) Mucor rouxii Pb, Cd, Ni, Zn Majumdar et al. (2010) Mucor sp. Cu Tahir (2012) Neurospora crassa Pb, Cu Kiran et al. (2005) Penicillium canescens Cr Say et al. (2003) Penicillium canescens As, Pb, Cd, Hg Say et al. (2003) Penicillium chrysogenum Cu, Ni, U, Cr, Th, Zn, Tan and Cheng (2003) Cd, Pb (continued)

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Table 18.3 (continued) Species Penicillium cyclopium Penicillium decumbens Penicillium digitatum Penicillium notatum Penicillium purpurogenum Penicillium verrucosum Phanerochaete chrysosporium Pleurotus florida Pleurotus floridianus Pleurotus ostreatus Pleurotus sajorcaju Pleurotus sapidus Polyporus frondosus Polyporus sulphureus Pyrenochaeta cajani Rhizoctonia solani Rhizopus arrhizus Rhizopus arrhizus Rhizopus cohnii Rhizopus nigricans Rhizopus sp. Saccharomyces cerevisiae Serpula himantioides Species of Aspergillus, Mucor, Penicillium, and Rhizopus Trichosporon cutaneum Volvariella diplasia Volvariella volvacea

Metals remediate Cu Cd, Ni, Cr Cd, Cu, Pb Cr Cr Pb Cu, Ni, Cd, Pb, Zn, Mn, Fe Cu, Zn, Ni, Pb Cu, Zn, Pb, Cd, Fe, Ni

References Ianis et al. (2006) Levinskaite (2001) Galun et al. (1987) Seshikala and Charya (2012) Say et al. (2003) Çabuk et al. (2005) Mamun et al. (2011)

Cr Cu, Cd, Pb, Ni

Bajgai et al. (2012) Lamrood and Ralegankar (2013) Lamrood and Ralegankar (2013)

Prasad et al. (2013) Lamrood and Ralegankar (2013) Pb, Ni, Cu, Zn, Cu, Cr, Arbanah et al. (2012) Mn Pb, Cd, Cu, Hg, Zn, Fe Arıca et al. (2003) Ni, Cu, Pb, Cd, Mn, Zn Ita et al. (2008) Ni, Cu, Pb, Cd, Mn, Zn Ita et al. (2008) Ni, Cu, Pb, Cd, Mn, Zn Ita et al. (2008) Cr Seshikala and Charya (2012) Cr Seshikala and Charya (2012) Ni, Zn, Cd, Pb Fourest and Roux (1992) Pb, Cr, Cd, Cu, Zn, Ni Prakasham et al. (1999) Cd Luo and Xiao (2010) Cr, Pb, Zn Bai and Abraham (2001) Cu, Cd Tahir (2012) Cd, Ni, Pb, Cr, Zn, Cu Thippeswamy et al. (2012b) As Adeyemi (2009) Cd, Cu, Fe Fulekar et al. (2012)

Cu, Zn, Pb, Cd, Ni, Fe

Sources: Adapted and modified from Archana and Jaitly (2015)

contamination in soil. In addition to that for the mechanistic understanding on growth conditions, enzyme production, type of compound degradation has been explored (Table 18.4). The bioconversion efficiency of wastes by some fungal species has been reported worldwide (Table 18.5).

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Table 18.4  Some commonly used fungal species, for mechanistic understanding on growth conditions, enzyme production, type of compound degradation, and key references Fungal species Phanerochaete chrysosporium Aspergillus flavus Bjerkandera adusta Fusarium oxysporum

Rhizopus arrhizus

Growth condition required Enzymes produced Lignin peroxidases and manganese peroxidases Laccase Grows best in cereals nuts legumes Commonly grows Lignin peroxidases on dead wood Grows in desert, Endoglucanase temperate, and tropical, soils of tundra Lipases Arises from nodes where rhizoids are borne

Compound degraded Reference Xenobiotic Paszczynski compounds and Crawford (1995) Removing Ghosh and surfactants and dyes Ghosh (2018) Xenobiotic compounds Degrades silver

Rhodes (2014) Danesh et al. (2013)

Fourest and Heavy metals like Roux (1992) Ni, Zn, Cd, Pb Also remediated uranium- and thorium-affected soil

Table 18.5  Bioconversion of waste by fungal species Fungal species Pleurotus citrinopileatus

Aspergillus niger

Bioconversion of waste Handmade paper and cardboard industrial waste

Remarks Successfully cultivated. Basidiocarps possessed good nutrient content and no genotoxicity Waste office paper to Used turbine blade reactor gluconic acid and production increased to four times in the presence of oxygen than air Biodecoloration Decoloring reactive dye

References Kulshreshtha et al. (2013)

Ikeda et al. (2006)

Vinciguerra et al. (1995) Beauveria bassiana Production of Utilized prawn chitinous Suresh and chitinase enzyme waste Chandrasekaran (1998) Mäkinen et al. Phlebia radiata Production of ethanol Successfully produced biocompound and biofuels at (2018) low cost Ghosh and Ghosh Aspergillus flavus Production of laccase Potential candidate for the enzyme production of lactase, used in (2018) bioremediation and bleaching of dyes, etc. Kantifedaki et al. Successfully produced by Monascus purpureus Production of (2018) biobased pigments semisolid fermentation and and Penicillium submerged fermentation purpurogenum technique

Lentinus edodes

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18.6.1  Mechanistic Approach of Fungal Phytoremediation In fungal phytoremediation, mechanism of fungal partner is very important to understand. Fungal phytoremediation has got several mechanistic pathways for bioremediation process. In general, fungus increases the ability of roots to absorb more heavy metals. Its mechanism could be devised as (i) avoidance and (ii) sequestration mechanisms. Avoidance ameliorates the metal toxicity though decreasing the concentration of metal by biosorption, precipitation, and uptake or efflux. Conversely, sequestration involves the formation of compounds for intracellular chelation (−) and further dilution in plant tissues due to plant growth, exclusion from uptake through precipitation, and chelation in the rhizosphere (Danesh et al. 2013). Both of these mechanisms may play part or even could counteract. Overall, the reduction in absorption owing to retention and immobilization takes part in fungal structures or mycorrhizal roots. The activation of specific/nonspecific transporters and pores play the part in the plasma membrane in plants and fungi, chelation in the cytosol and the sequestration into the vacuoles of plants as well as in fungal cells. Further, transportation and exportation occur through the fungal hyphae, involving both active and passive transportation into the mycorrhizae (Fig. 18.3). Fungal phytoremediation is proven to be efficient, where the abilities of hyperaccumulators diminished. One of the limitations of hyperaccumulators is to accumulate

Fig. 18.3  Mechanisms involved in remediation of HM-contaminated soil by HMT-PGP microbes-­ plant interaction. (Sources: Mishra et al. (2017))

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less concentration of contaminants due to their small biomass while fungi can accumulate more due to their some molecular mechanisms. Hence, intervening the interaction of hyperaccumulator plant with fungi and other legume plant and herbs could help us to use it as a potent strategy for phytoremediation (Yang et al. 2016). Therefore, further exercise is required for explaining the molecular mechanisms underlying.

18.6.2  Factors Influencing the Fungal Phytoremediation Several factors influencing the fungal phytoremediation include species of plant and fungi, their association strength, plant-soil interaction, physical and chemical properties of soil, and biophysical aspects such as temperature, pH, salinity, soil microbes, and metal characteristics (Fig. 18.4). 18.6.2.1  Temperature The fungi are having their different temperature range for growth based on different habitat, such as mesophilic (5–35  °C), psychrophilic (below °C), thermophilic (above 40 °C), etc. With the change in the temperature, the bioavailability of the

Plant characteristics

Plant type, species and varieties

Climate characteristics

Humidity

Rootzone characteristics

Root depth

Medium characteristics

Soil type

Chelator characteristics

Pollutant characteristics

Chelation

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Temperature

Exudation

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Air

pH

Resistance to disease and pests

Rainfall

Sunlight

Microbial activity

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Time of application

Concentration of metal pollutants

Phytoremediation

Growth rate and biomass

Salinity

Toxicity

Organic matter

Cation exchange capacity

Bioavailability

Speciation and toxicity

Chemical structure

Fig. 18.4  Relationships among the factors affecting phytoremediation efficiency. (Adapted with permission from Saxena et al. (2019))

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heavy metals is also changed. An increase in soil temperature tends to speed up the concentration of metals in the soil due to increase rate of organic matter degradation. It was observed that high temperature is favorable for the absorption of heavy metals. However, the temperature also affects the growth of fungi. So, fungi with high temperature tolerance will be beneficial for the bioremediation process (Yadav et  al. 2018b, c). Fe and Mn are mobile in alternating in dry and wet conditions (Boisselet 2012). 18.6.2.2  pH pH is an important parameter which controls the availability of heavy metals to get remediated. Heavy metals are present in a dissolved state if the pH of the solution is at 2–3. However, the bioavailability, dissolution, and precipitation of each metal have its own intrinsic capacity along with the pH range. 18.6.2.3  Redox Potential The redox potential affects the state of oxidation of the metals, as different forms show different behaviors in solubility. Anaerobic conditions in deeper parts of the soil for oxidoreductive reactions of microorganisms can accelerate the heavy metal degradation. Redox potential along with pH affects the fungal-phyto interactions with the soil components by altering the sorption capacity and influencing stability of complexes. 18.6.2.4  Heavy Metals Bound with Hydrocarbon Some of the heavy metals are present in the bound form of the other compounds such as polycyclic aromatic hydrocarbons (PAH). The remediation of such metals can be achieved only after degradation of the host compound. Some fungal species such as Agaricus bisporus, Pleurotus ostreatus, and Ganoderma lucidum are observed to degrade the hydrocarbons in petroleum. Pleurotus ostreatus is beneficial in degrading the PAH (García-Delgado et al. 2015). 18.6.2.5  Other Growth Requirements Apart from the temperature, other factors such as moisture percentage, sugar and other organic materials, oxygen, amino acids, vitamins, fatty acids, etc. are also important for fungal growth. The change in these requirements can also enhance/ limit the fungal phytoremediation potential.

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18.6.2.6  Fungal Species Different fungal species are having different capacity to remediate the heavy metals from the soil and water based on their internal genetic constitutes and external growth and environmental factors. To check the any new/existing species remediation potential, the arsenic test (preliminary assessment) will serve as good choice. Later on, the heavy metals-based potential check can be made and compared with the existing data. However, some of the fungal species can serve as a bioindicator of particular heavy metals. In this case, these species serve as the reference species for the remediation potential. For example, Lycoperdon perlatum may be employed as a bioindicator of heavy metals and selenium in soil pollution (Quinche 1990). Filamentous fungi are known to possess higher adsorption capacities for heavy metal removal (Singh and Gauba 2014). Trichoderma and Mortierella species isolated from the soil and Aspergillus and Penicillium species isolated from marine and terrestrial environments, respectively, have the high ability to remediate contaminated environment (Thenmozhi et al. 2013). Arbuscular mycorrhizal fungus Glomus mosseae formed a symbiotic associate of P. vittata L. and possessed substantial resistance to arsenic toxicity by increasing the plant biomass, and this mycorrhiza can enhance the arsenic sink. Mycorrhiza can be a potential tool for fungal phytoremediation by choosing the native species of fungi/host and alteration in the a­ ssociation by changing any of the fungi/host or controlling factors or inoculation of the new fungal strains. This can be achieved through re-vegetation on the contaminated sites such as mine areas.

18.7  Precaution Prerequisite Some prerequisite precautions are needed for successful achievement of fungal phytoremediation which involves selection of correct fungal species for targeted metal contamination for developing a screening protocol (Matsubara et al. 2006). Among these precautions, major points have been prescribed in general which should be considered. These involve as follows: • The catabolic activity and capacity of organisms involved to transform the target compound(s) and bring the concentrations to levels that meet regulatory standards • The rate of bioremediation • The possible production of toxic by-products at dangerous levels during the remediation process • Adaptability of the process to site conditions (environmental and anthropogenic) • Economic viability of the process

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18.8  Conclusion and Future Prospects As explained above, fungal phytoremediation is a very potent technology for sustainable bioremediation of contaminated soils and water. In general, it is still in infancy in laboratory conditions and greenhouse, which limits the outcome to the actual field condition pertaining to multiple factors. Hence, the assessment of the efficiency rate of fungal phytoremediation must be tested in field condition in order to commercialize this green technology by evaluating the different plant for targeted heavy metal. Similarly, there are few lags in this eco-friendly remediation technology such as to increase the growth rate of plants, increase the biomass of such plants for maximum absorption of heavy metals, and take a look on possible hazards on food chain. Field experiments should be devised to explore the hyperaccumulators from where these metals can be harvested easily and feasible techniques to harvest these metals without exerting a negative impact on environment. Also, the key to mycoremediation is determining the right fungal species to target a specific pollutant. Desirable traits should be identified from the hyperaccumulator and fungi genome. Such gene can be selected by the conventional techniques or new technologies of hybridization such as protoplast fusion. Identification of genes coding for different toxicants from different hyperaccumulators and their transformation in same plant can develop SUPERBUG plant for phytoremediation. Besides the several constraints and limitations, fungal phytoremediation appears to be the most potent, eco-friendly, economical, and environmentally attractive option of bioremediation in heavy metal-contaminated soils and water resources. Many fungal species can grow under various contaminated conditions, thus enabling remediation in the contaminated environment that may not be suitable for other organisms. Based on our review of the subject and key questions raised on the concerned topic, we do not conclude that it could solve the issues of metal or hydrocarbon contamination completely. Conversely, a synergistic approach involving proactive policy designing in the field of fungal phytoremediation ranging from lab-based desirable trait based targeted metal contaminations with a particular fungal species, after testing it in green houses it has a potential to be replicated in the field environment for the future safety of soil, plant, water resources and rising human population prone to future heavy metal contaminations. Acknowledgments  The authors are grateful to Prof. Harcharan Singh Dhaliwal, Vice Chancellor, Eternal University, Baru Sahib, Himachal Pradesh, India, for providing infrastructural facilities and constant encouragement.

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Chapter 19

Fungal Enzymes for Bioremediation of Xenobiotic Compounds Peter Baker, Araven Tiroumalechetty, and Rajinikanth Mohan

19.1  Introduction Xenobiotics are natural or synthetic organic compounds foreign to an organism that are potentially toxic and entail negative ecological or physiological consequences, in the form of pollution and disease, respectively (Olicon-Hernandez et al. 2017). Industry and agriculture are two major sources of environmentally prevalent xenobiotics which include fertilizers, insecticides, pesticides, dyes, plastics, and hydrocarbon derivatives (Sharma et  al. 2018). Chemically, the vast majority of these compounds are aromatic with one or more frequently substituted phenyl functional groups. Many of these xenobiotic compounds are of major health and ecological concern as they are frequently carcinogenic and teratogenic, thereby disrupting development and reproductive capabilities in humans, fish, fish-eating birds, and other animals. A serious environmental and health issue is the accumulation of persistent, toxic chemical pollutants requiring new cost-effective and efficient ways to tackle the growing threat of environmental toxicity in the modernized world. The majority of xenobiotic compounds can be decomposed or modified by microbes (Cameron et al. 2000). Bioremediation utilizes the metabolic potential of biological organisms to degrade or transform hazardous compounds in the environment into less toxic or nontoxic forms (Watanabe 2001; Yadav et al. 2019a, b). In particular, the use of fungi, referred to as mycoremediation, has attained widespread Peter Baker and Araven Tiroumalechetty contributed equally with all other contributors. P. Baker · A. Tiroumalechetty Department of Biology, Colgate University, Hamilton, NY, USA R. Mohan (*) Department of Biology, Colgate University, Hamilton, NY, USA Department of Biology, Mercyhurst University, Erie, PA, USA e-mail: [email protected] © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_19

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attention. Although bacteria are also versatile decomposers of xenobiotic pollutants, certain fungal strains are able to tolerate higher levels of pollutants. In some cases, such as the degradation of polyaromatic hydrocarbons (PAHs), bacteria can metabolize them and utilize them as carbon and energy sources, but they cannot mineralize them completely the way fungi can (Mougin et al. 2009). Fungi are capable of metabolizing a wider range of pollutants than their prokaryotic counterparts due to both extra- and intracellular degradation mechanisms and the powerful, nonspecific nature of the enzymes involved in both processes (Christian et  al. 2005a). Additionally, the enzymes in question are tolerant of an active under diverse conditions including broad pH ranges, making fungi and their enzymes desirable for bioremediation (Verma and Madamwar 2002) (Rastegari et al. 2019; Yadav et al. 2017, 2018). Filamentous fungi, particularly from the group called white rot fungi and mainly from Basidiomycetes, demonstrate a striking ability for oxidative decomposition of lignin, a recalcitrant component of wood composed of polyphenols (Mougin et al. 2009). These enzymes were evolved by such fungi to assist in the breakdown and detoxification of potentially hazardous by-products resulting from the decomposition of wood and other organic matter (Morel et al. 2013). The presence of these enzymes in many species of white rot fungi has been confirmed using comparative genomics to identify the so-called xenome, which consists of genes involved in xenobiotic detoxification. In the last two decades, these enzymes have been extensively adapted for bioremediation of pollutants in a low-cost, eco-friendly manner. Bioremediation involves environments toxic to the survival of biological systems and despite fungal robustness and innate degradation infrastructure, there is a limit to the organisms’ tolerance of xenobiotic-induced environmental toxicity. Frequently, the polluted areas are too nutrient-poor to support microbial growth. Moreover, the fungi capable of detoxifying a pollutant may be sensitive to other pollutants in the environment (Mougin et al. 2009). To circumvent these problems, one option is to utilize isolated fungal enzymes or enzyme mixtures. The use of fungal enzymes rather than complete fungal populations is also a more eco-friendly approach in contrast to using live microbes as it allows for the mitigation of any adverse environmental impact resulting from the introduction of novel species (Sharma et  al. 2018; Kour et  al. 2019; Rana et  al. 2019a, b). This is especially desirable as release of genetically modified fungi or other organisms is contingent on both acceptance by regulatory bodies like EPA and the general public (Ang et al. 2005). The most prominent groups of enzymes utilized in xenobiotic bioremediation transformations are oxidoreductases: peroxidases, laccases, and oxygenases (Sharma et al. 2018). Oxidoreductases can detoxify compounds by catalyzing oxidative coupling reactions using oxidizing agents to support the reactions. Laccases (LACs) and CYP monooxygenases (P450s) use molecular oxygen as the electron acceptor, while peroxidases use hydrogen peroxide to oxidize the substrates; both reactions result in the formation of water as a by-product. Peroxidases are heme-­ containing proteins, and the major types of peroxidases involved in detoxification processes in fungi are manganese peroxidase (MnP), lignin peroxidase (LiP), and versatile peroxidases (VP) (Doddapaneni et  al. 2005). Xenobiotic detoxification

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enzymes are produced by a great diversity of fungi, but many are often produced by a large physiological group called the white rot fungi. These fungi actively degrade lignin  – a large, complex, aromatic-containing networked polymer, creating a bleached appearance in the host, hence the name white rot fungi (Pointing 2001). Given the complexity of lignin and the structural motifs common to both lignin and various xenobiotics, these enzymes are well suited for xenobiotic detoxification (Fig. 19.1). A few genera of fungi are used extensively in bioremediation processes; these include Trametes, Pleurotus, Phanerochaete spp., etc. In this chapter, we will review the utility of fungal enzymes in bioremediation processes. In particular, we focus on the detoxification of organic xenobiotics by oxidoreductase enzymes including the extracellular peroxidases and laccases as well as the intracellular CYP450.

19.2  Sources of Xenobiotic Pollutants Synthetic dyes released from paper, textile, plastic, cosmetic, food, and drug industries can be toxic and carcinogenic (Asgher et al. 2008; Levin et al. 2005). Polycyclic aromatic hydrocarbons (PAHs) like anthracene, pyrene, benzopyrene, and naphthalene are toxic, potentially carcinogenic xenobiotics that are produced from fuel combustion, gas plants, and industrial applications. Endocrine disrupting chemicals (EDCs) include alkylphenols such as nonylphenol and octylphenol as well as biphenyls such as bisphenol A (BPA), stilbene, and genistein estrogens. These are pharmaceutically active compounds that disrupt endocrine homeostasis in animals and are of grave concern to human health (Asgher et al. 2008). Other sources of xenobiotics include bleach plant effluents from paper and pulp industry, which contain toxic polychlorinated phenols (PCPs) and other organic compounds used for bleaching, and include dyes like azo dye and crystal violet (D'Souza et  al. 2006). Chlorinated aromatic compounds like dichloro- and trichlorophenol and derivatives such as DDT, chlordane, and lindane are used in pesticides.

19.3  Metabolism of Xenobiotic Pollutants by Fungi Bioremediation through fungi may be achieved by direct metabolism, whereby a fungus may completely degrade a xenobiotic compound to innocuous end products like carbon dioxide and other simple inorganic compounds (Mougin et al. 2009). This is particularly likely under nutrient-limiting conditions when the substrate compound could serve the carbon and/or energy needs of the organism. While this metabolic pathway is preferred as the toxic compound is eliminated or nearly so, the more common method of detoxification is co-metabolism. In this process, a cosubstrate is used to transform a xenobiotic compound without utilizing the compound for growth or energy needs. Generally, this only results in minor changes in the structure of the pollutant. For instance, substrate-free radicals may be generated

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Fig. 19.1  The relationship and commonalities between organically produced lignin and varied xenobiotic chemical structures. (a) The structure of lignin according to Laurichesse and Averous (Laurichesse and Avérous 2014) and the monomeric alcohols which polymerize to form the cross-­ linked complex. (b) Common xenobiotic classes and associated structures as lignin model compounds targeted by lignin-degrading enzymes. Alkylated phenolic structures are common to both EDCs and lignin, as is the geminal phenyl motif of BPA. Nitrogen is not commonly present in lignin structures, nor are fused polycyclic aromatics nor polychlorinated phenols making lignin-­ degrading enzymatic degradation of these compounds less obvious

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which subsequently cross-link the substrates with themselves or with environmental structures such as soil components, thereby forming adducts with decreased bioavailability and toxicity (Bollag 1992). Alternatively, these modified compounds could be further metabolized by other organisms leading to further detoxification. A third detoxification pathway involves conjugation or oligomerization of a pollutant. In this process, the resulting products of enzymatic action have larger and more complex structures than the parent compounds, but the structural modification or aggregation reduces the bioavailability of the compound, thereby negating its biological potency. In conjugation, typically, the compound may be methylated, acetylated, alkylated, or conjugated with sugars and amino acids and subsequently excreted or sequestered into storage structures. In oligomerization, a xenobiotic is coupled with a second molecule of the same or different xenobiotic following oxidation, aggregating into structures that become less bioactive.

19.4  Fungal Enzymes in Xenobiotic Remediation The most commonly studied enzymes involved in mycoremediation all catalyze oxidoreduction reactions in the transformation of xenobiotics (Sharma et al. 2018). Among these enzymes are some active extracellularly and other which are intracellularly involved in the transformation and subsequent detoxification of xenobiotics. Within these oxidoreduction-catalyzing enzymes, some such as laccases (LACs) and CYP monooxygenases (P450s) use molecular oxygen as an eventual electron acceptor, while peroxidases use hydrogen peroxide as electron acceptors in their respective reaction pathways. One specific class of enzyme common to many xenobiotic degradation mechanisms are the heme peroxidases of which the most extensively involved and studied are manganese peroxidase (MnP), lignin peroxidase (LiP), and versatile peroxidases (VP) (Doddapaneni et  al. 2005). The relatively lower substrate specificity of the extracellular enzymes in oxidative degradation allows them to target a wider variety of xenobiotic compounds and is as such of great interest in bioremediation processes (Harms et  al. 2011). As with many secreted proteins, these peroxidases and laccases are glycosylated, at times heavily; for instance, certain laccases show 16% sugar content (Salony et al. 2006). Many fungal metabolites bearing diverse functional groups can serve as redox mediators for the oxidative function of these enzymes and can enhance their activity (Asgher et al. 2008). These include veratryl alcohol, N-hydroxyacetanilide, and acetosyringone to name a few. Mediators are particularly important when the substrates are too large to be accommodated into the active site of the enzyme (Mougin et  al. 2009). The toxic xenobiotics or the products formed following their metabolism by extracellular enzymes can be further detoxified by intracellular enzymes including CYPs that are ubiquitously present in all organisms (Morel et  al. 2013). These enzymes can catalyze various reactions including dealkylation, hydroxylation, and sulfoxidation and frequently function by inserting molecular oxygen into various substrates.

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19.4.1  Laccase LACs, originally identified as ligninolytic enzymes, are present in ascomycetes, basidiomycetes, and deuteromycetes and are commonly found in gene families (Harms et al. 2011). Laccases are secreted blue multicopper oxidases that catalyze single-electron oxidation of phenolic and other aromatic substrates to create free radicals with the concomitant reduction of molecular oxygen to water. The radical products cross-link by self-coupling or cross-coupling to form less toxic polymers; during this process they may undergo further decomposition reactions such as decarboxylation, dechlorination, and demethoxylation. Following LAC-dependent oxidation events, metabolites commonly exhibit oxidative coupling or radical polymerization resulting in compounds of greater molecular masses than the parent compound (Junghanns et al. 2005) which potentially may further negate the biological effects of such foreign compounds (Tsutsumi et al. 2001). The use of abundantly available oxygen as an oxidizing agent with the formation of water as a harmless by-product makes laccases a desirable bioremediation enzyme. Compared to peroxidases, different laccases can tolerate a wider pH range (2–10) and have broader substrate specificities (Xu 1996). The broad substrate range of laccases encompasses various xenobiotic compounds contaminating soil and water, i.e., PAHs, organophosphorus insecticides, and toxic dyes which include aromatic compounds like phenols, trichlorophenols, aminophenols (anilines), phenylenediamine derivatives, and benzenethiols (Amitai et al. 1998; Kues 2015; Sharma et al. 2018; Xu 1996; Yadav et al. 2016). The potential of LACs in xenobiotic remediation has long since been a point of interest and investigation. Laccases from Trametes versicolor and Pleurotus ostreatus have been used in the detoxification of PCBs (polychlorinated biphenyls) which were used as insulators in electrical equipment until they were banned in 1979 for their high toxicity (Keum and Li 2004). Laccases facilitate substrate oligomerization and dechlorination, leading to detoxification of these compounds. LACs catalyze the initial oxidation of polycyclic aromatic hydrocarbons (PAHs), beginning their extracellular degradation (Pozdnyakova et al. 2018b). A LAC isolated from Coriolopsis gallica oxidizes the PAHs carbazole, N-ethylcarbazole, and dibenzothiophene in concert with proper mediators such as 1-hydroxybenzotriazole and 2.2′-azino-bis-(3-ethylbenzothiazoline)-6-sulfonic acid (Viswanath et  al. 2014). When free laccase was applied to PAH-contaminated soil, 15 PAHs including anthracene and benzopyrene were degraded, demonstrating the potential of direct enzyme application for bioremediation (Wu et al. 2008). Covalently immobilized laccases from Trametes versicolor were shown to efficiently degrade the PAHs, anthracene, naphthalene, and phenanthrene (Bautista et al. 2015). Multiple studies revealed the ability of laccases from Pleurotus ostreatus, Trametes versicolor, and other strains to effectively degrade BPA and other endocrine disruptors like alkylphenols. The detoxification of these compounds by laccases may be brought about by oligomerization or mineralization to carbon dioxide, and this action was enhanced by supplementation with mediators (Macellaro and Pezzella 2014;

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Margot et  al. 2013; Zhang et  al. 2015). Laccases from Pycnoporus sanguineus, Myceliophthora thermophila, Trametes trogii, and Ceriporiopsis subvermispora effectively metabolized the toxic dyes like bromophenol blue, methyl violet, and malachite green and other dyes used in the textile industry (Antosova et al. 2018; Chmelova and Ondrejovic 2016; Mougin et al. 2009). Synthetic laccases are also used in the paper and pulp industries to bleach paper and clothes (Sharma et  al. 2018). The addition of mediator compounds can dramatically enhance the detoxification of substrates as witnessed in the transformation of halogenated pesticides by laccases from the fungus, Coriolopsis gallica (Torres-Duarte et al. 2009); in this study, acetosyringone and syringaldehyde proved to be the most effective mediators. Finally, laccases from Trametes versicolor have been used to oxidize the pharmaceutical drugs, diclofenac and mefenamic acid in municipal wastewater (Margot et al. 2013). Beyond naturally occurring LACs, some researchers have explored the possibility of engineering LACs with the intention of improving transformational efficiency and demonstrating transient expression by means of directed evolution (Camarero et al. 2012; Gu et al. 2014; Mate and Alcalde 2015; Theerachat et al. 2012; Wong et al. 2013b).

19.4.2  Peroxidase The ligninolytic peroxidases of interest in bioremediation carry high redox potentials (>1.4 V) and catalyze the oxidative breakdown of lignin and other compounds with aromatic ring structures using hydrogen peroxide as a cosubstrate and with the help of certain mediator compounds like veratryl alcohol (Piontek et  al. 2001; Sharma et al. 2018). These peroxidases are glycosylated secreted proteins carrying an iron protoporphyrin (heme) ring at the catalytic center. Peroxidases can oxidize and generate phenolic radicals which may aggregate and precipitate. There are several groups of secreted peroxidases in fungi: 19.4.2.1  Lignin Peroxidase LiPs are heme-containing enzymes which can metabolize various aromatic compounds many of which are generally refractory to breakdown, with pH optima in the acidic range (2–5) (Shrivastava et  al. 2005). Since the 1983 discovery of P. chrysosporium, LiP in extracellular media, isozyme forms among various basidiomycetes have been isolated ranging in weight from 38 kDa to 43 kDa (Falade et al. 2017). LiPs have been shown to completely oxidize both methylated lignin model and non-­methylated lignin model compounds as well as PAHs (Kadri et al. 2017). Through powerful, nonspecific catalytic transformative activity, the aptly named LiP is capable of direct transformation of up to 90% of lignin structural components (Falade et al. 2017).

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LiPs are commonly produced by Phanerochaete and Trametes spp. As demonstrated by LiP isolated from P. chrysosporium which transforms the PAHs benzo[a]pyrene, anthracene, 1-methylanthracene, 2-methylanthracene, 9-methylanthracene, fluoranthene, acenaphthene, and dibenzothiophene, these enzymes possess broad substrate ranges (Pozdnyakova 2012). LiPs also demonstrate expanded substrate ranges in the presence of veratryl alcohol which increases oxidation of weak or terminal LiP substrates. With the advantageous kinetics conferred by mediators like veratryl alcohol, LiP transforms most aromatic compounds with an ionization potential less than 8 eV. LiP also has the unique ability to cleave esters in non-­phenolic aromatics, thereby further demonstrating the significance of LiPs to bioremediation efforts of diverse, aromatic-containing xenobiotics (Pozdnyakova 2012). 19.4.2.2  Manganese Peroxidase MnPs are also heme-containing secreted enzymes which function under relatively less acidic conditions (pH 4–7) than those of LiP (Asgher et al. 2008). As a heme peroxidase, MnPs share much of their characteristics and mechanism with LiPs (Deshmukh et al. 2016). MnP appears not to occur in the same large gene families characteristic of other ligninolytic enzymes such as LAC (Torres-Farrada et  al. 2017) and is only found in Basidiomycota (Harms et al. 2011). MnP is capable of aromatic ring cleavage within monoamino-dinitrotoluene and chlorophenol derivatives (Harms et al. 2011). The oxidative potential of MnP relies on the oxidation of Mn2+ to Mn3+ by the enzyme followed by the indirect, single-electron oxidation of subsequent substrates as Mn3+ is reduced, thereby reverting to Mn2+. Due to the indirect mechanism, the substrate range is broad, and the extent of transformation of resulting metabolites is near complete. MnP oxidizes phenols, aromatic amines, and dye compounds as well as mineralizes CO2 from various quinones – common products of PAH radical polymerization – by ring fission transformations. MnPs isolated from Anthracophyllum discolor clearly demonstrate this PAH-substrate promiscuity as they are able to oxidize pyrene, anthracene, fluoranthene, and phenanthrene as well as various derivatives of these compounds (Pozdnyakova 2012). A MnP from the white rot fungus, Trametes, displayed a strong ability to degrade azo and indigo dyes as well as PAHs (Zhang et al. 2016). Another MnP from Peniophora incarnata not only displayed the potential to break down the PAH anthracene, but this ability was transferable by heterologous expression in yeast, signifying a bioremediatory potential. MnP from Ganoderma lucidum used as cross-linked enzyme aggregates efficiently degraded the endocrine disrupting nonylphenol and triclosan (Bilal et al. 2017a). MnP is of great interest in bioremediation efforts as it has been shown to be stable under adverse conditions; however, a complicating factor of its use in remediation effort and applications is the mechanistic need for a suitable chelator (Bogan et al. 1996). Such chelators are commonly organic acids such as oxalic or malonic

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acid derivatives (Kadri et al. 2017). These compounds complex with Mn3+, enabling the oxidation of substrate lignin model compounds. In addition to a mechanistic dependence on chelators, MnP activity significantly increases in the presence of redox mediators such as Tween 80. With the added effect of Tween 80, MnP has been shown to transform compounds with ionization potentials of 8.2 eV. Despite the powerful oxidative potential and resistance to adverse conditions of MnP, the application of these enzymes in biotechnical pursuits is complicated by not only chelator requirements and redox mediator reliance for elevated efficiency but also by LAC-dependent initiation of Mn3+ complexing (Schlosser and Hofer 2002). No Mn3+ complexing was observed in in vitro mixtures of semi-purified MnP, Mn2+, and oxalate or malonate when H2O2 sources were excluded. However, the addition of LAC stimulated Mn3+ complexing and ultimate MnP-stimulated substrate oxidation. In response to PAH-polluted media, MnP secretion by Fomes is very high, reaching concentrations of approximately 1299  U/L after 21  days of xenobiotic exposure (Godoy et al. 2016). 19.4.2.3  Versatile Peroxidase Versatile peroxidases (VP) are a hybrid between LiPs and MnPs as they contain a heme group and oxidize Mn2+to Mn3+, inducing indirect oxidations, as well as oxidize phenolic and non-phenolic substrates; VPs conjoin mechanisms and substrate ranges between these two enzymes (Kues 2015; Pozdnyakova 2012). VPs have thus far only been identified in Basidiomycetes (Harms et al. 2011). In the initial characterization of the first identified VP – specifically, PS1 isolated from Pleurotus eryngii  – it was shown to possess both the Mn oxidation domain of MnPs and the aromatic substrate oxidation center (AS) of LiPs (Camarero et al. 1999). Furthermore, PS1 and subsequently characterized VPs have been shown to actually retain LiP- or MnP-like enzymatic activity in conditions that would inactivate LiPs or MnPs, respectively. Little is understood about the role of VPs in the mediation of xenobiotic transformation; however, it is known that VP production is induced by the presence of PAH pollutants (Pozdnyakova et al. 2018b). In addition to PAHs, VPs can also degrade polyhalogenated aromatic pesticides containing diverse functional groups as demonstrated by VP isolated from Bjerkandera adusta which successfully transformed dichlorophen (an antimicrobial polycyclic), bromoxynil (a nitrile herbicide), and pentachlorophenol (PCP) – a potent pesticide (Davila-Vazquez et al. 2005). Given the hybridization of LiP and MnP substrate ranges, they have the potential to be a major component of the subsequent remediation of many chemically diverse xenobiotics. Due to their broad substrate range and various transformation mechanisms, VPs warrant further characterization and investigation regarding their potential application in bioremediation and biotechnical efforts. Examples of fungal laccases and peroxidases employed for xenobiotic detoxification are presented in Table 19.1.

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Table 19.1  Details of microbes producing laccase and peroxidase in the bioremediation of xenobiotics Enzyme source Laccases Trametes versicolor T. versicolor, Pleurotus ostreatus P. ostreatus P. pulmonarius T. sanguineus

Echinodontium taxodii Clavariopsis aquatic Peroxidases Trametes spp.

Peniophora incarnata Irpex lacteus Phanerochaete chrysosporium Penicillium chrochloron

Xenobiotic

Compounds

Mediator

Reference

PAH

Anthracene, benzopyrene

ABTS

PCB

Hydroxy PCBs

TEMPO

Dodor et al. (2004) Keum and Li (2004)

Insecticides, VX, Russian VX, nerve agents diisopropyl-Amiton Toxins Aflatoxin B1 Endocrine disruptor, PAH Azo dyes Endocrine disruptor

Bisphenol A, benzopyrene, phenanthrene

ABTS ABTS, AS, SA ABTS

Brilliant Violet 5R, Direct Lignin Red 5B, Direct Black 38 derivatives Nonylphenol ABTS

Amitai et al. 1998) Loi et al. (2016) Balcazar-­Lopez et al. (2016) Han et al. (2014) Junghanns et al. (2005)

–

Zhang et al. (2016)

PAH

Indigo, anthraquinone, azo, triphenylmethane, fluorene, fluoranthene, pyrene, phenanthrene, anthracene Anthracene

–

Lee et al. (2016)

Dyes PCB

Azo, indigo dyes 2,4-dichlorophenol

– –

Dyes

Cotton blue

–

Qin et al. (2014) Chen et al. (2011) Shedbalkar et al. (2008)

Dyes, PAH

ABTS 2,2′-azino-bis(3-ethylbenzothiazoline)-6-sulfonic acid, AS acetosyringone, PAH polyaromatic hydrocarbons, PCB polychlorinated biphenyls, TEMPO 2,2,6,6-tetramethylpiperidineN-oxyl radical, SA syringaldehyde

19.4.3  Cytochrome P450 Monooxygenase Many fungi also possess intracellular transformation enzymatic machinery capable of further degradation of xenobiotics (Syed and Yadav 2012). They belong to the larger group of oxygenases that are the principal intracellular enzymes involved in the aerobic degradation of aromatic xenobiotics using oxygen (Sharma et al. 2018). These are also heme-containing enzymes that add one or more oxygens to destabilize aromatic rings to break down and even solubilize the compound. Oxygenases could catalyze the addition of one oxygen molecule (monooxygenase) or two molecules (dioxygenase) to the substrate. Many halogenated herbicides, fungicides, and pesticides are detoxified using oxygenases, and the best-studied enzyme belongs to the cytochrome P450 enzyme family, which utilized NADPH as a cofactor to catalyze redox reactions (Doddapaneni et al. 2005).

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P450s are very distinct from other xenobiotic-degrading fungal enzymes as it is active intracellularly (Deshmukh et al. 2016). Furthermore, CYP450 are found in massive gene families with multiple subfamilies as discovered in the genome of P. chrysosporium with at least 149 isozymes (Doddapaneni et  al. 2005; Olicon-­ Hernandez et al. 2017; Syed et al. 2011). Such gene families have been identified in ascomycetes, basidiomycetes, mucoromycetes, and chytridiomycetes (Harms et al. 2011). As monooxygenases, P450 incorporates a single atom from molecular oxygen into the substrate while reducing the remaining oxygen to water. Compared to CYPs involved in primary metabolism, CYPs mediating detoxification processes are generally less specific (Cresnar and Petric 2011). P450 monooxygenases are a major component in the degradation of various xenobiotics as they are quite nonspecific and demonstrate very powerful enzymatic activity in the transformations of such diverse chemical motifs. However, as these are intracellularly active proteins, their application in fungal enzyme isolation and immobilization is further complicated. Within each class of P450s, multiple subfamilies exist sharing general function and some sequence identity. Many class-II subfamilies are involved in biosynthetic pathways and homeostatic processes. The subfamily CYP51 has been indicated in maintaining cell wall integrity while CYP61 (a fungi-specific subfamily) is responsible for spore outer wall formation. Various Class II P450s transform PAHs, alkyl phenols (APs), alkanes, and polychlorinated dibenzo-p-dioxins (PCDDs) (Harms et al. 2011; Kues 2015). CYPs from the basidiomycete Phanerochaete chrysosporium have been shown to catabolize PAHs including anthracene and the endocrine disrupting alkylphenols (Hirosue et  al. 2011; Syed et  al. 2011). Among Class II P450 subfamilies, CYP57 has been shown to detoxify pisatin, a heterocyclic, aromatic-­containing fungus growth inhibitor produced in pea plants upon microbial attack. Furthermore, CYP53 has been shown to degrade and detoxify benzoate derivatives within multiple species from both the Ascomycota and Basidiomycota phyla. Additionally, CYP504 is known to degrade phenylacetate derivatives. The significance of Class II P450 activity is further emphasized in observed biodegradation by non-ligninolytic fungi such as Scopulariopsis brevicaulis which was shown to completely transform anthracene (a tricyclic PAH), producing the same major metabolite – 9,10-anthraquinone – as ligninolytic fungi such as the basidiomycete Fomes, despite the absence of extracellular enzymes (LAC, LiP, and MnP) demonstrating the versatility and potency of Class II P450-dependent intracellular transformation pathways (Godoy et  al. 2016). CYPs have been tapped to detoxify a variety of pharmaceutical compounds, including antibiotics, anti-inflammatories, and β-blockers (Olicon-Hernandez et  al. 2017). Naproxen is an environmentally prevalent pharmaceutical pollutant and has even invaded drinking water systems due to overuse in treating human and animal diseases. CYPs were shown to detoxify this drug through demethylation and hydroxylation (Aracagok et al. 2017). A fundamental limitation in utilizing CYP enzymes in cell-free systems is the fact that CYP functions in intracellular networks that also involve other enzymes (Haroune et al. 2017). Shotgun proteomics of proteins in response to the PAH, anthracene in Penicillium oxalicum, revealed regulation of hundreds of proteins, and intracellular metabolism of such xenobiotics typically follows two distinct phases with the CYPs working in conjunction with other enzymes like epoxide hydrolases and transferases (Lucero Camacho-Morales et al. 2018).

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19.4.4  Unspecific Peroxygenase UPOs are secreted hybrid enzymes which combine the functionalities of heme peroxidases and P450 monooxygenases (Karich et al. 2017). Among this class of peroxygenases, two common structural motifs have been observed: short UPOs are approximately 29 kDa, while long UPOs are approximately 44 kDa. Short UPOs are found across all fungal phyla, but long UPOs are exclusive to ascomycetes and basidiomycetes. Both short and long UPOs have immense substrate ranges, acting in association with hydrogen peroxide as a cofactor, a marketed improvement in simplicity from the nuanced requirements of MnP. Furthermore, 41 of EPA-listed priority pollutants have been shown to be transformed by UPOs, and more than 300 other aromatic, poly- and heterocyclic, and aliphatic substrates have been identified thus far. Despite the recent findings regarding xenobiotic substrates, the physiological role of UPOs remains to be identified (Olicon-Hernandez et al. 2017); however, Karich et  al. theorized that UPOs and P450s may work in harmony with UPOs crudely transforming xenobiotics extracellularly to reduce negative biological effects, while P450s “fine-tune” the resulting metabolites so that they may be further transformed and rendered inert within the cell or even consumed as carbon sources due to the incredibly diverse functionality of these enzymes. Thus far, present understanding explains that UPOs’ substrate transformations are limited by steric hindrance, bioavailability and potential substrate solubility, or strong inactivation of an aromatic ring by electron withdrawing groups (Karich et al. 2017). Despite these regulatory factors in UPOs’ substrate specificity, UPO isolated from Agrocybe aegerita demonstrated significant transformation of naphthalene, phenol, anisole, toluene, ethylbenzene, acenaphthylene, acenaphthene, fluorene, phenanthrene, anthracene, pyrene, benzo[a]anthracene, 1,2-diphenylhydrazine, benzidine, and 2,4-dimethylphenol as well as various phthalate esters, nitroarenes, and polychlorinated benzenes (Karich et al. 2017). Among these known substrates exist several previously only thought to be transformed by P450s or MnPs and LiPs, again demonstrating the remarkable potential of these enzymes. Such versatile enzymes in extracellular degradation processes carry great implications for biotechnical development of xenobiotic mitigation strategies.

19.5  M  echanisms of Xenobiotic Detoxification by Fungal Enzymes Fungal extracellular xenobiotic degradation occurs in two conserved steps (Rao et al. 2010). Firstly, the hydrolytic system targets macromolecules for degradation by hydrolase activity. The hydrolyzed substrates are then further transformed by the ligninolytic system comprised of nonspecific oxidative enzymes. The enzymatic mechanisms we focus on in this section are the oxidoreductases whose mechanisms are well researched (Fig. 19.2).

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Fig. 19.2  Simplified mechanism of fungal enzymes used in bioremediation. (a) Laccase enzymatic cycle. Mediators include veratry alcohol (VA), Tween 80, and other small organic molecules. (b) Lignin peroxidase enzymatic cycle. Both oxidized states of LiP are able to oxidize substrates and mediators. Mediator molecules are used when the substrates cannot access enzyme active site. (c) Manganese peroxidase enzymatic cycle. MnP preferentially oxidizes Mn2+ in the first redox reaction but can also oxidize other mediators. Mn2+ oxidation is highly specific for the second redox reaction which restores the base state. (d) CytP450 enzymatic cycle. The reduction of heme-­ bound molecular oxygen is either directly catalyzed by the reductase coenzyme or indirectly via the use of cytochrome B5 as a redox mediator. (Adapted from Kues (2015) and Guengerich (2001b))

19.5.1  Laccases Laccases are multicopper oxidases with low substrate specificity. Within basidiomycetes these families range from 5 to 17 isozymes (Yang et al. 2017). Within LAC isozymes, enzymatic copper involvement is conserved with four copper atoms at +2 oxidation states in the resting enzyme. The four copper atoms are characterized as T1, T2, and T3 – which is binuclear. The T1 copper facilitates substrate oxidation, while T2 and T3 copper atoms store the resulting electrons which then convert diatomic oxygen to water. They catalyze coupled redox reaction between a substrate and molecular oxygen resulting in the formation of a radical cation and water, respectively. The type I copper is found in a wide cavity of the enzyme surface, which allows it to bind to many different types of substrates (Su et al. 2018). The significance of LACs in the remediation of xenobiotics is only amplified by laccase-mediator systems (LMSs) which act as electron transfer chains and thereby

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further broaden the substrate range as well as increase kinetic favorability of transformation (Wong 2013a). LMSs share the common mechanism of facilitating the indirect oxidation of a substrate following the primary oxidation event of the mediator by LAC. The enzymatic cycle of laccases can be simplified as a four-electron abstraction from the substrate by the oxidized enzyme followed by the reduction of molecular oxygen to water to regenerate the active enzyme (Fig. 19.2a). The enzyme primarily exists in the resting oxidized state (RO) with each of the copper atoms oxidized. The catalytic reaction is initiated by the donation to the type I copper of four electrons from suitable substrates (reductants). Three electrons are transferred through a conserved His-Cys-His sequence of amino acids to the trinuclear cluster (TNC) consisting of the type II copper and the two type III copper atoms, resulting in the fully reduced enzyme. The electrons are used to reduce molecular oxygen which binds to the TNC, forming a peroxide intermediate (PI). The reduction then proceeds by donation of a hydrogen by a nearby glutamic acid residue resulting in the cleavage of the oxygen-oxygen bond and the native intermediate (NI) state of the enzyme. This state can directly proceed to the fully reduced state in the presence of large amounts of suitable substrates, producing two water molecules or proceed to the fully oxidized state with loss of a single water molecule in the absence of significant levels of reductants (Jones and Solomon 2015). Laccases tend to accommodate substrates with relatively low redox potentials such as phenolic compounds which are oxidized to phenoxyl radicals which may undergo coupling reactions or isomerization to form quinones. Laccases are generally unable to directly oxidize compounds with high redox potentials or steric hindrance. Instead, such compounds are oxidized via the laccase-mediator system which involves oxidation of micromolecular organics which can then oxidize target compounds through redox reactions leading to cleavage of bonds (Su et al. 2018).

19.5.2  Peroxidases Of the various peroxidases discussed herein, only the general mechanism of lignin peroxidase and manganese-dependent peroxidases has been elucidated. The shared mechanism of these two types of peroxidases involves the oxidation of the enzyme by two electron abstractions by the H2O2 cosubstrate followed by two one-electron transfer steps where the oxidized enzyme abstracts an electron from the substrate (Mougin et al. 2009). 19.5.2.1  Lignin Peroxidases In LiPs, Fe(III) in the heme ring is coordinated with four heme tetrapyrrole nitrogens and to a histidine residue. LiP has been shown to fold into a globular profile measuring about 50 × 40 × 40 ¯ ; this form is subdivided into proximal and distal

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domains relative to the heme group. Two small molecular channels allow for heme accessibility despite its fixed position within the protein’s tertiary structure. LiPs are capable of cleaving Cα-Cβ bonds as well as bonds between aryl Cα (Kadri et  al. 2017). LiP is further differentiated from other peroxidases by the optimal pH of approximately 3.0 (Falade et al. 2017). Lignin peroxidases are activated by a two-­ electron oxidation of the native enzyme by the cosubstrate H2O2 (Mougin et  al. 2009). This results in compound I which is reduced by the substrate as part of a one-electron redox reaction to compound II. Compound II then abstracts a second electron from the substrate resulting in regeneration of the native enzyme. The activated forms of the enzyme have a high redox potential allowing it to oxidize compounds such as lignin that other peroxidases are unable to transform. LiPs act through three different mechanisms based on the availability of its heme cofactor toward the substrate. First, LiPs act directly on certain phenolic and non-phenolic compounds which can access the heme group. This can lead to breakage of carbon-­ carbon bonds in substrates leading to conformational changes (Fig. 19.2b). The second lignin peroxidase mechanism is indirect, acting through a redox mediator to oxidize compounds that cannot access the heme group (Fig. 19.2b). Lignin peroxidase oxidizes mediators such as veratryl alcohol (VA) to a cation radical (VA+) which then oxidizes compounds through a redox reaction. The third mechanism occurs through further reactions of VA. The cation radical oxidizes organic acids into anion radicals which act as reductant. These radicals can also reduce molecular oxygen to a dioxygen anion which acts as a reductant. Through the reduction of ferrous ions, this dioxygen anion can also reduce hydrogen peroxide to a hydroxyl radical which can oxidize compounds through the non-enzymatic Fenton’s reaction (Christian et al. 2005b). 19.5.2.2  Manganese Peroxidase Manganese peroxidases, also known as manganese-dependent peroxidases, are oxidized to a two-electron-deficient state (compound I) and restore its basal oxidation state by two single-electron abstracting steps (Christian et al. 2005b). The first step which reduces the compound I to a single-electron-deficient state (compound II) is not very specific and can use either Mn2+ or small compounds such as VA as reductants. The second step which restores the basal oxidation state of the enzyme is highly specific to Mn2+ (Fig.  19.2c). Both of the activated enzyme states oxidize Mn2+ to Mn3+ which acts as a nonspecific small, diffusible redox mediator. Mn3+ are chelated to carboxylic acids such as oxalic acid and are thus able to cause one-­ electron abstraction oxidation of various compounds (Fig.  19.2c). They can also react with carboxylic acids to produce radicals such as VA+ and superoxide. Manganese peroxidases are also able to act indirectly as reductors. This is done through oxidation of hydroquinone to semiquinone radicals which reduce highly oxidized compounds. This generates a quinone which is then reduced back to hydroquinone by quinone reductase enzyme.

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19.5.3  Cytochrome P450 Cytochrome P450 enzymes are part of a superfamily with remarkable variation. Due to this variation within the CYP superfamily, the reactions associated with P450 enzymes are classified based on reactionary motifs and various protein involvements into ten distinct classes, three of which have been observed in fungi: class II, class VIII, and class IX (Cresnar and Petric 2011). Most commonly, CYP450 enzymes act as monooxygenases that incorporate an atom of oxygen into substrates (van den Brink et al. 1998). P450 enzymes therefore serve as versatile catalysts for region-specific and stereospecific oxidation resulting in hydroxylation, heteroatom oxygenation, dealkylation, and epoxidation of C=C bonds (Deshmukh et  al. 2016; OliconHernandez et al. 2017). However, the basis of all these reactions can be summarized as being XH + NAD ( P ) H + H + + O2 → XOH + NAD ( P ) + H 2 O +





with R-H as the substrate of cytochrome P450. Among these transformations, NADH or NADPH frequently acts as an electron donor to the second oxygen as it is reduced to water (Kues 2015). The reaction does not usually produce a phenol but instead an oxide which isomerizes to the more favorable phenolic conformation (Christian et  al. 2005b). The phenolic form is activated toward further enzyme-­ catalyzed reactions, resulting in trans-dihydrodiols (Tongpim and Pickard 1999). Much like the peroxidases, P450 enzymes are heme-based, with their catalytic active site containing a heme-bound iron atom (Guengerich 2001a). The substrate first binds to the heme iron Fe3+ which is reduced by an accessory enzyme, NADPH reductase (Fig. 19.2d). This reduced iron Fe2+ then binds with molecular oxygen. The accessory enzyme then reduces molecular oxygen which is then protonated. This leads to cleavage of the O-O bond, causing the protonated oxygen to be released as H2O. The resulting FeO3+complex radicalizes the substrate by proton (or electron abstraction). The radical then accepts the hydroxyl group (or oxygen atom in case of electron abstraction) before being released, simultaneously returning the iron to its base state. There exists a shunt pathway in which a peroxide is used as ­cosubstrate instead of molecular oxygen, skipping the need for the reductase coenzyme (Fig. 19.2d). The monooxygenation mechanism of P450 is not limited to carbons but can also apply to heteroatoms such as nitrogen, sulfur, phosphorus, and iodine. In these cases, the oxygen transfer is more complicated, with the oxygen being added to the substrate after two successive electron transfer steps.

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19.6  S  trategies and Considerations in Xenobiotic Bioremediation Either fungi or enzymes isolated from fungi could be used for the detoxification of xenobiotic compounds. Some fungal phyla possess more versatile degradation systems relative to bacterial strains (Pozdnyakova et al. 2018a). The fundamental limitations in using isolated fungal enzymes include productivity as well as stability and retention of activity (Sharma et al. 2018). Laccases, for example, have an acidic pH optimum that may not be available in effluent soils and waters, presenting a major limitation to their use. Furthermore, cost of application, intolerance of enzymes to high levels of cosubstrate hydrogen peroxide, and issues with enzyme reusability have hampered the application of these enzymes in the field (Nicell and Wright 1997). Once applied to sites for remediation, the enzymes are also under threat of denaturation and destruction by physical and chemical forces and by the action of microbes and their enzymes. Besides, excessive exposure of heme-containing proteins to oxidative species can lead to inactivation and subsequent degradation of the protein (Valderrama et al. 2002). Indeed, these limitations are so critical that they have delayed or limited large-scale application of these enzymes in bioremediation processes (Ayala et al. 2008). One approach to the identification of potential fungal strains with significant remediation potential and biotechnical application is to isolate populations from xenobiotic-polluted environments (Godoy et al. 2016). Doing so results, in part, in fungi with innate tolerance to the environmentally present pollutants. The pollutant-­ resistant detoxifying fungi often produce biodegradative enzymes that are too limited in amount to isolate and utilize. To scale up production of these enzymes, genetic engineering could be used to overexpress fungal enzymes in fungi, plants, or other organisms that could colonize the polluted substratum. Alternatively, enzymes could be mass-extracted from such organisms by heterologous expression and utilized for bioremediation. This is a cost-effective strategy that not only allows purification of large amounts of enzyme with stability and activity, the purification process of recombinant proteins is also simpler as they can be isolated using cleavable tags (Alcalde et al. 2006). Furthermore, the stability, activity, and other features of an enzyme can also be enhanced using genetic and enzyme engineering approaches. By introducing mutations using strategies like DNA shuffling, error-­ prone PCR, and site-directed mutagenesis (Dua et al. 2002), enzyme engineering could be accomplished, where a change in protein sequence from mutations results in a possible change in enzyme structure or regulation with improved traits including increased stability and activity; increased xenobiotic substrate specificity or wider substrate range; tolerance to a wider range of pH, temperature, and stress conditions; and decreased susceptibility to proteases in the natural environment (Rayu et al. 2012). One such mutated laccase protein from Pleurotus ostreatus identified in a screen gained greater enzymatic activity and higher stability at acidic pH, widening its pH tolerance in detoxifying toxic industrial dyes (Miele et al. 2010).

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Similarly, directed evolution of a laccase resulted in 3.5-fold increase in acetonitrile catabolism while tolerating relatively high concentrations of it (Alcalde et al. 2005). As enzymes are sensitive to physical and chemical environmental changes, various strategies have been evolved to fortify and preserve their structure and function while exposed to adverse conditions. Enzyme immobilization in carbohydrate-­ based matrices has been shown to be effective in prolonging the half-life and enhancing the stability and catalytic function of the remediative enzymes. Enzymes can be immobilized to solid support systems for more effective use in bioremediation. Enzyme immobilization can be done by covalent linkage or adsorption of enzymes onto solid surfaces like glass, affinity tag-bearing beads or nylon membranes, suspension in polymeric gels like chitosan, gelatin and encapsulation in solid matrices such as sodium alginate and could also accompany enzyme crosslinking using glutaraldehyde (Diano et al. 2007; Koyani and Vazquez-Duhalt 2016; Sirisha et al. 2016). Immobilization has been shown to improve the stability, catalytic function, and longevity of enzymatic function of the fungal enzymes and additionally protects the enzymes from proteolytic degradation (Asgher et  al. 2007; Cheng et al. 2007; Jing and Kjellerup 2018). This is brought about in part by the increased resistance to physical, chemical, and biological denaturing agents afforded by the immobilization. Additionally, immobilized enzymes can be recovered and reused, thus economizing the process. Laccases immobilized on glass beads from Trametes retained 90% of their activity and showed greater resistance to proteases (Bilal et al. 2017b; Dodor et al. 2004). Similar benefits have been observed for other fungal enzymes like MnPs (Bilal and Asgher 2016). Enzymatic nanoreactors with dendritic copolymers bearing glycosidic groups anchoring laccases not only showed improved enzymatic activity but also increased thermostability (Gitsov et al. 2008). Similarly, vault nanoparticles with hollow cores covalently anchoring multiple MnP enzymes showed enhanced stability while displaying a threefold increase in phenol degradation (Wang et al. 2015). The addition of cofactors, cosubstrates, and mediators is important to drive the action of the enzymes in bioremediation scenarios; this is particularly important for recalcitrant pollutants. Additionally, the presence of mediator compounds can enhance enzyme activity. The fungal metabolites, veratryl alcohol and 2-chloro-­1,4dimethoxybenzene, can stimulate LiP enzyme activity, and this is particularly useful when treating recalcitrant substrates (Asgher et al. 2008), while acetohydroxamic acid serves as an excellent mediator for laccases (Minussi et  al. 2007). In some cases, the addition of glucose as a carbon source to fungal cultures also enhanced the detoxification process (Asgher et al. 2008). Marine fungi are adapted to much harsher conditions than terrestrial fungi. Their enzymes are able to withstand high salinity and concentration of phenolic compounds. Certain marine fungi have shown bioremediation potential for water-­soluble crude oil fractions between 0.01 and 0.25 mg/mL (Deshmukh et al. 2016). Other extremophilic fungi such as Pestalotiopsis palmarum are able to survive in high concentration of extra-heavy crude oil and salt while producing oxidative exoenzymes (laccases and lignin peroxidases) which degrade the maltene and asphaltene fractions of oil for use as a carbon and energy source (Naranjo-Briceno et al. 2013).

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The isolation of enzymes from marine organisms can select for traits such as heat and cold tolerance and salt and pressure tolerance (Lima and Porto 2016). Recently, non-protein enzyme mimics or next-generation artificial enzymes in nanoparticles or nanocomposites have been used to replace enzymes in bioremediation processes (Gao and Yan 2016). Desired for their low cost and higher stability, these structures display enzyme-like properties under physiological conditions. Nanozymes, although lacking an active site, bind substrates specifically and catalyze their transformation. The Fe3O4-based magnetic nanoparticles mimic peroxidases and can degrade toxic dye compounds like methylene blue (Wu et al. 2015). Carbon-based nanomaterials made of graphene oxide (GO) also display peroxidase-­ like activity (Ma et al. 2017). Similarly, a guanosine monophosphate (GMP) coordinated copper nanocomposite mimicked laccases in being able to degrade phenolic compounds including hydroquinone and naphthol (Liang et al. 2017). Although it is desirable to identify and isolate detoxification enzymes in polluted environments, a basic handicap is the ability to culture these microbial species. Only a handful of microbes are culturable, and these may not include the microbes that make the enzyme of interest. This problem is circumvented by recent developments in metagenomics which can identify potential detoxification enzyme-coding genes. These genes could be expressed in the heterologous system to mass-produce the enzymes of interest for bioremediation. Clues to the functionalities of these genes could be revealed by metatranscriptomic and metaproteomic approaches, which could also reveal biochemical pathways and synthetic pathways for xenobiotic-transforming enzyme production. In silico approaches can be employed to understand the evolution of xenobiotic detoxification by phylogenetic analyses. More recently, bioinformatic analysis in the form of molecular docking tools has been developed to predict pollutant substrates of the detoxification enzymes. This approach has been particularly useful for laccases which have broad substrate specificity; in one study, laccase enzyme structures were screened against a database of toxic compounds to identify putative substrates (Suresh et al. 2008). The study found that 30% of the studied compounds that were recognized as environmental pollutants could be potentially metabolized by fungal laccases. Similar approaches could be employed for other enzymes with known structures to test if they could be used to detoxify pollutants of interest.

19.7  Conclusions and Future Perspectives Increased global industrialization has presented many challenges including the production of ecotoxic industrial wastes that also present health threats as xenobiotic compounds. Fungi have specialized enzymes that are highly efficient in detoxifying these pollutants. Harnessing the power of these enzymes is proving to be an effective strategy for the bioremediation of the xenobiotic compounds. In particular, oxidoreductase enzymes including laccase, peroxidase, and oxygenases like CYP from various fungal species are employed to detoxify a wide range of pollutants.

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In addition to bioremediation by detoxification, enzymes can also be employed for the detection and quantification of xenobiotics in the environment. White rot fungi (WRF) are a versatile group of microbes that are capable of oxidative detoxification of a variety of chemical pollutants and xenobiotic compounds that are environmentally harmful. These organisms and their enzymes have the ability to not only tolerate the xenobiotics but in many cases also metabolize or help sequester them. The limitation in biodegradation of xenobiotic compounds is frequently the initial stages of degradation. The oxidoreductase enzymes are all extracellular and specialized for less specific activity in initial stages of xenobiotic metabolism and, as such, as ideal candidates for detoxification of these compounds. Development of heterologous expression systems and industrial scale expression is still a limitation. Limitations of enzyme quantity and stability are being overcome by the adoption of genetic engineering technology and enzyme immobilization approaches. Fungal enzymes can also be modified for enhanced catalytic activity in addition to thermostability to maximize the efficiency of the detoxification process. Creation of enzymes with increased redox potentials, especially in those such as laccases which have a lower redox potential, could widen the substrate range. The utility of laccases and CYPs may be limited by the availability of molecular oxygen. Development of peroxidases to target those substrates could be one possible approach. Present bioengineering tactics are focused on developing superior enzymes for the bioremediation of known pollutants, but such approaches may be extended to target novel pollutants, not known to be biodegradable. Alternatives to fungal enzymes in the form of nanozymes are also being explored. An inherent limitation in the use of single enzymes in bioremediation systems is the fact that detoxification processes are often multi-step pathways, requiring the action of multiple enzymes in a sequence. The ability to use fungal enzymes in synthetic pathways for biotransformation of toxic pollutants into economically valuable compounds would be a great future direction. Since fungal-fungal and fungal-bacterial consortia have been employed for remediation of xenobiotics like PAHs, a similar strategy with enzymes could follow suit. Structural studies of enzymes as well as directed evolution can facilitate changes in structure that could substantially enhance activity and xenobiotic metabolism. Better structural understanding of laccases as well as other enzymes could lead to minimization of the use of redox mediators, especially in in situ bioremediation scenarios in environmental settings, where the massive release of these ­eco-­unfriendly compounds may be undesirable. The solving of crystal structures of fungal detoxification enzymes enables molecular docking analyses to predict the ability to metabolize various xenobiotic substrates. Similarly, the application of enzymes also requires a fine understanding of soil structure and soil-water interactions as these can significantly affect enzyme activity. Recent advances in this area are likely to benefit soil modeling studies and assessment of enzyme function for future bioremediation projects. Adopting a bioprospecting-like approach to find new strains of fungi with superior ligninolytic enzymes in environments like rainforests could be a promising way forward in continued efforts to combat environmental pollutants of industrial origin in cost-effective and environmentally sustainable tactics.

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Chapter 20

Fungal White Biotechnology: Conclusion and Future Prospects Ajar Nath Yadav

This book contains current knowledge about fungi; their biodiversity from diverse sources including natural as well as extreme, associated with plants (epiphytic, endophytic, and rhizospheric); and potential biotechnological applications of fungi and fungal products for different processes in agriculture, industry, pharmaceutical, food and feed processing, and environments for sustainable developments. Fungi are prominent sources of pharmaceuticals and are used in many industrial fermentative processes such as the production of enzymes, vitamins, pigments, lipids, glycolipids, polysaccharides, and polyhydric alcohols. In the past 50 years, several major advancements in medicine came from lower organisms such as molds, yeasts, and the other diverse fungi. Fungi are extremely useful in making high-value products like mycoproteins and act as plant growth promoters and disease suppressors. Fungal secondary metabolites are important to our health and nutrition and have tremendous economic impact. In addition to this, fungi are extremely useful in carrying out biotransformation processes. Recombinant DNA technology, which includes yeasts and other fungi as hosts, has markedly increased the market for microbial enzymes (Fig. 20.1). Today, fungal white biotechnology is a major participant in the global industry due to its mind-blowing potential in medical with pharmaceutical importance. The secondary metabolites with pharmaceutical importance of Aspergillus nidulans and other fungi could be used as drug worldwide (Keller et al. 2005). Ergot alkaloids (ergometrine and Ergotamine) and lovastatin, a popular cholesterol-lowering drug, are the secondary metabolites (Beekman and Barrow 2014). Fungal metabolites have antitumour, antiviral, antibacterial, and immunosuppressant activities. Fungi are used as high-cost food because of its high protein and low calorific value (Siso 1996). Some of the edible fungi (mushrooms) are used such as Agaricus bisporus (white button mushroom), Agaricus campestris, Morchella (temperate zone A. N. Yadav (*) Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, Sirmour, Himachal Pradesh, India © Springer Nature Switzerland AG 2019 A. N. Yadav et al. (eds.), Recent Advancement in White Biotechnology Through Fungi, Fungal Biology, https://doi.org/10.1007/978-3-030-25506-0_20

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Fig. 20.1  Biotechnological applications of fungi and their value-added products in agriculture, health, industry, and environments. (Adapted with permission from Kour et al. (2019a))

­mushroom), Pleurotus sp. (oyster mushroom), and Volvariella (paddy straw mushroom) (Mathur et al. 2011; Reddy 2015). Fungi are used as the rich sources of single-cell proteins. Some of the fungi for SCP are given as Aspergillus niger, Fusarium avenaceum, Neurospora sitophila, Penicillium chrysogenum, and Saccharomyces cerevisiae (Kuhad et al. 1997; Ravindra 2000) (Fig. 20.2). Fungi are widely used in fermentative industries for the production of ethanol, organic acids, antibiotics, and enzymes like fungal cellulases, gluconase, and glycosidase. Certain fungi like Penicillium notatum, P. chrysogenum, and Cenococcum sp. are used in antibiotic production (Broadbent 1966; Mishra et al. 2019), whereas Saccharomyces cerevisiae and Monilia sp. are used in ethanol production (Chiang et al. 1982; Molaverdi et al. 2019). Fungi are also useful in ripening of cheese and processing of other products. With the expanding population, environment is changing greatly, and agriculture is one of the most exposed sectors to these changes and faces a number of challenges like pollution, pathogenic attack, salinity, drought, high temperature, low temperature, and so on. All these challenges ultimately affect the productivity (Fig. 20.3). To overcome such issues, eco-friendly approaches are very vital. The use of fungi as biofertilizers is one emerging area which is getting a greater attention, as it is proving its importance by enhancing the plant growth and productivity by diverse plant growth-promoting traits including the production of phytohormones, siderophores, and hydrolytic enzymes, making the availability of different nutrients and protecting plants against pathogens (Kour et  al. 2019b; Yadav et  al. 2019b). Fungi are very important for the soil ecosystem and play a considerable role in the daily life of human beings; additionally, they are important for agriculture, bioremediation, natural cycling, food industry, and as biofertilizers (Karthikeyan et  al. 2014). Plant

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Fig. 20.2  White fungal biotechnology applications. (Adapted with permission from Challa et al. (2019))

growth-promoting fungi are gaining a significant interest to be used as bioinoculants, as they possess manifold benefits on quantity as well as the quality of the plants as well as the positive relation they exhibit with the ecological environment. Vesiculararbuscular mycorrhizae are the mutualistic symbiosis between the roots of higher plants and certain fungi. The mycorrhizae help in the phosphate nutrition of plants and protect the roots by forming the mantle (Mathur et  al. 2011; Yadav et  al. 2019a). Furthermore, fungi have been demonstrated to produce phytohormones including indole-3-acetic acid (IAA), gibberellins (Maor et  al. 2004; Tudzynski and Sharon 2002; Kumar et  al. 2018), and siderophores (Kumar et  al. 2018; Milagres et al. 1999). Fungi are known for its quite specific and effective action and have low residual effects in comparison with synthetic pesticides. Fungi are used as bioherbicides such as Cercospora ageratinae (Pamakani weed), Colletotrichum gloeosporioides (mistletoes), Leptosphaerulina trifolii (Passiflora), Phyllosticta (Glycosmis), Puccinia chondrillina (rush weed), Septagloeum gillis (mistletoes), and Wallrothiella arecuthobii (mistletoes) (Charudattan and Dinoor 2000; Poudel et al. 2019). Employing live fungi or fungal enzymes for industrial applications is known as fungal white biotechnology. However, fungal biotechnology, an essential ­technology,

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Fig. 20.3 A schematic representation of plant-associated rhizospheric microbiota. The rhizosphere-­associated fungi in relation with plants exhibit widespread applications in the field of agriculture, pharmaceutical and biomedical industries, and industrial sectors. (Adapted with permission from Pattnaik and Busi (2019))

uses renewable sources for sustainable growth of population. Fungi or fungal enzymes play a role in food and feed industries. Fungal white biotechnology brings down greenhouse emissions and is eco-friendly in nature. The applications of fungi as food (edible fungi) and fodder and using fungi in processing food (bread, cheese, and other bakery products) and fermenting food (alcohols, beverages) are indispensable. Fungal white biotechnology enhances flavor in cheese, bread, and beverages, protein quality and yield in SCPs, and stability and shelf-life of the products with much efficacy. Though there are many advantages with white fungal biotechnology, tolerance to the extreme conditions during processing and enrichment of products are the major challenges observed with white fungal biotechnology (Challa et al. 2019) (Fig. 20.4). Enzymes from microbes have gained great appreciation worldwide for their extensive uses in a variety of sectors such as food, agriculture, chemical, and medicine. In the field of medicine, these are used to treat health disorders associated with deficiency of human enzymes caused by genetic problems (Anbu et al. 2015; Singh et al. 2016). The processes mediated by enzymes are speeding up in food, pharmaceutical, textile, paper, and leather industries and are gaining interest because of certain advantages such as reduced process time, intake of low energy input,

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Fig. 20.4  Classification and biological activity of fungal natural products. (Adapted with permission from Gholami-Shabani et al. (2019))

c­ost-­ effectiveness, greater efficiency, nontoxicity, higher-quality products, and eco-­friendly characteristics (Gurung et al. 2013; Kamini et al. 1999; Singh et al. 2016). The fungi and fungal enzymes are used for biodegradation of azo dye and hydrocarbons for sustainable environments. Peroxidase enzymes of Penicillium chrysosporium and Streptomyces sp. have potential biodegradable activities that degrade amaranth dye, Orange G, heterocyclic dyes like Azure B and Lip dye (Fig. 20.5). The filamentous fungi are also playing a role in the degradation of toxic hydrocarbons (Hasan 2018; Leahy and Colwell 1990). Another potential application of fungal communities for sustainable environments is remediation of explosive contaminated soil by its lignin-degrading enzymes (Maqbool et  al. 2016). The fungi have eminent role in the removal and recovery of heavy metals from wastewater and industrial effluents. Hg, Cu, Ni, Pb, and Cd are extracted at pH  2–5 by myceliar beads of Penicillium.

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Fig. 20.5  Fungal enzymes and their biotechnological applications in diverse sectors. (Adapted with permission from Kour et al. (2019a))

White biotechnology or industrial biotechnology uses enzymes and microorganisms to make bio-based products in sectors such as chemicals, food and feed, detergents, paper and pulp, textiles, and bioenergy. White biotechnology also employs living organisms as cell factories preferably utilizing renewable natural resources such as lignocellulose for production of a variety of materials and biocompounds with energy efficiency, increased productivity, and environmentally sustainable characteristics. Fungi are the organisms that play a potential role in degradation of explosives. It is observed by repeated laboratory studies involving pure cultures of white rot fungi. It also helps in degradation of hydrocarbons in the environment. Fungi attract considerable attention due to their possible involvement in the

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diverse applications. So far, large numbers of enzymes have been purified from fungal cultures and characterized in terms of their biochemical and catalytic properties. It possesses antimicrobial activities and is used in bio-mineralization, as a food for its high protein contents and as a biofertilizer. Acknowledgments  The authors are grateful to the Department of Biotechnology, Akal College of Agriculture, Eternal University, Baru Sahib, and Department of Environment, Science and Technology, Shimla, HP-funded project “Development of microbial consortium as bio-inoculants for drought and low temperature growing crops for organic farming in Himachal Pradesh,” for providing the facilities and financial support to undertake the investigations. There are no conflicts of interest.

References Anbu P, Gopinath SC, Chaulagain BP, Tang T-H, Citartan M (2015) Microbial enzymes and their applications in industries and medicine 2014. Biomed Res Int 2015:816419 Beekman AM, Barrow RA (2014) Fungal metabolites as pharmaceuticals. Aust J Chem 67:827–843 Broadbent D (1966) Antibiotics produced by fungi. Bot Rev 32:219–242 Challa S, Dutta T, Neelapu NRR (2019) Fungal white biotechnology applications for food security: opportunities and challenges. In: Yadav AN, Singh S, Mishra S, Gupta A (eds) Recent advancement in white biotechnology through fungi: Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham, pp 119–148. https://doi. org/10.1007/978-3-030-14846-1_4 Charudattan R, Dinoor A (2000) Biological control of weeds using plant pathogens: accomplishments and limitations. Crop Protect 19:691–695 Chiang L, Hsiao H, Flickinger M, Chen L, Tsao G (1982) Ethanol production from pentoses by immobilized microorganisms. Enzym Microb Technol 4:93–95 Gholami-Shabani M, Shams-Ghahfarokhi M, Razzaghi-Abyaneh M (2019) Natural product synthesis by fungi: recent trends and future prospects. In: Yadav AN, Singh S, Mishra S, Gupta A (eds) Recent advancement in white biotechnology through fungi: Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham, pp 195–228. https://doi.org/10.1007/978-3-030-14846-1_7 Gurung N, Ray S, Bose S, Rai V (2013) A broader view: microbial enzymes and their relevance in industries, medicine, and beyond. Biomed Res Int. https://doi.org/10.1155/2013/329121 Hasan IF, AI-Jawhari (2018) Role of filamentous fungi to remove petroleum hydrocarbons from the environment. In: Kumar V, Kumar M, Prasad R (eds) Microbial action on hydrocarbons. Springer Singapore, Singapore, pp 567–580. https://doi.org/10.1007/978-981-13-1840-5_23 Kamini N, Hemachander C, Mala JGS, Puvanakrishnan R (1999) Microbial enzyme technology as an alternative to conventional chemicals in leather industry. Curr Sci 77:80–86 Karthikeyan P, Kanimozhi K, Senthilkumar G, Panneerselvam A, Ashok G (2014) Optimization of enzyme production in Trichoderma viride using carbon and nitrogen source. Int J  Curr Microbiol App Sci 3:88–95 Keller NP, Turner G, Bennett JW (2005) Fungal secondary metabolism—from biochemistry to genomics. Nat Rev Microbiol 3:937 Kour D, Rana KL, Yadav N, Yadav AN, Singh J, Rastegari AA, Saxena AK (2019a) Agriculturally and industrially important fungi: current developments and potential biotechnological applications. In: Yadav AN, Singh S, Mishra S, Gupta A (eds) Recent advancement in white biotechnology through fungi: Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham, pp 1–64. https://doi.org/10.1007/978-3-030-14846-1_1

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Kour D, Rana KL, Yadav N, Yadav AN, Singh J, Rastegari AA, Saxena AK (2019b) Agriculturally and industrially important fungi: current developments and potential biotechnological applications. In: Yadav AN, Singh S, Mishra S, Gupta A (eds) Recent advancement in white biotechnology through fungi, Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham, pp 1–64. https://doi.org/10.1007/978-3-030-14846-1_1 Kuhad RC, Singh A, Tripathi K, Saxena R, Eriksson K-EL (1997) Microorganisms as an alternative source of protein. Nutr Rev 55:65–75 Kumar CS, Jacob T, Devasahayam S, Thomas S, Geethu C (2018) Multifarious plant growth promotion by an entomopathogenic fungus Lecanicillium psalliotae. Microbiol Res 207:153–160 Leahy JG, Colwell RR (1990) Microbial degradation of hydrocarbons in the environment. Microbiol Mol Biol Rev 54:305–315 Maor R, Haskin S, Levi-Kedmi H, Sharon A (2004) In planta production of indole-3-acetic acid by Colletotrichum gloeosporioides f. sp. aeschynomene. Appli Environ Microbiol 70:1852–1854 Maqbool Z, Hussain S, Imran M, Mahmood F, Shahzad T, Ahmed Z, Azeem F, Muzammil S (2016) Perspectives of using fungi as bioresource for bioremediation of pesticides in the environment: a critical review. Environ Sci Pollut Res 23:16904–16925 Mathur N, Vyas P, Joshi N, Choudhary K, Purohit DK (2011) Mycorrhiza: a potent bioinoculant for sustainable agriculture. In: Pathak H, Sharma A (eds) Microbial technology “the emerging era” lap lambert. Academic Publishing Ag & Co. Kg, Dudweiller Landstr, pp 230–245 Milagres AM, Machuca A, Napoleao D (1999) Detection of siderophore production from several fungi and bacteria by a modification of chrome azurol S (CAS) agar plate assay. J Microbiol Meth 37:1–6 Mishra R, Kushveer JS, Revanthbabu P, Sarma VV (2019) Endophytic fungi and their enzymatic potential. In: Singh BP (ed) Advances in endophytic fungal research: present status and future challenges. Springer International Publishing, Cham, pp  283–337. https://doi. org/10.1007/978-3-030-03589-1_14 Molaverdi M, Karimi K, Mirmohamadsadeghi S, Galbe M (2019) High titer ethanol production from rice straw via solid-state simultaneous saccharification and fermentation by Mucor indicus at low enzyme loading. Energy Convers Manag 182:520–529 Pattnaik SS, Busi S (2019) Rhizospheric fungi: diversity and potential biotechnological applications. In: Yadav AN, Mishra S, Singh S, Gupta A (eds) Recent advancement in white biotechnology through fungi: Volume 1: diversity and enzymes perspectives. Springer International Publishing, Cham, pp 63–84. https://doi.org/10.1007/978-3-030-10480-1_2 Poudel A, Jha P, Shrestha B, Muniappan R (2019) Biology and management of the invasive weed Ageratina adenophora (Asteraceae): current state of knowledge and future research needs. Weed Res 59:79–92 Ravindra P (2000) Value-added food:: single cell protein. Biotechnol Adv 18:459–479 Reddy SM (2015) Diversity and applications of mushrooms. In: Bahadur B, Venkat Rajam M, Sahijram L, Krishnamurthy KV (eds) Plant biology and biotechnology: Volume I: plant diversity, organization, function and improvement. Springer India, New Delhi, pp 231–261. https:// doi.org/10.1007/978-81-322-2286-6_9 Singh R, Kumar M, Mittal A, Mehta PK (2016) Microbial enzymes: industrial progress in 21st century. 3 Biotech 6:174 Siso MG (1996) The biotechnological utilization of cheese whey: a review. Bioresour Technol 57:1–11 Tudzynski B, Sharon A (2002) Biosynthesis, biological role and application of fungal phytohormones. In: Osiewacz HD (ed) Industrial applications. Springer, Berlin/Heidelberg, pp  183– 211. https://doi.org/10.1007/978-3-662-10378-4_9 Yadav AN, Mishra S, Singh S, Gupta A (2019a) Recent advancement in white biotechnology through fungi Volume 1: diversity and enzymes perspectives. Springer International Publishing, Cham Yadav AN, Mishra S, Singh S, Gupta A (2019b) Recent advancement in white biotechnology through fungi. Volume 2: perspective for value-added products and environments. Springer International Publishing, Cham

Index

A Abiotic stress, 324, 327, 330–332, 336, 337 Actinomycin, 122 Adsorption metal ions, 397 wastewater treatment, 397 Adsorption activation energy, 408 Adsorption effects, 165 Adsorption mechanism model, 402, 403 Adsorption method, 161 Adsorption process, 408 Adsorption studies batch adsorption study, 409–410 breakthrough analysis, 411 capacity, 409 column adsorption study, 410–411 equilibrium adsorption capacity, 410 packed bed column, 411 Agricultural practices, 420 Agrowastes, 118 Airlift bioreactor, 414 Alcohol oxidase (AOX), 58, 65 Aliphatic hydrocarbons, 240 Alkyl glucosides, 266 AMF symbiosis, 327, 334 Amino and alkylamino groups, 75 Amylases, 134, 135, 310, 311 Anaerobic conditions, 452 Anaerobic rumen fungi (ARF) brown rot fungi, 6 plant biomass degradation, 3, 4 secretome, 12, 13 wood-decaying fungi, 8 wood-degrading fungi, 13 Analytical profile index (API), 302 Anthracene, 119, 217, 224, 228

Anthraquinone, 119, 120 Anthropogenic activities, 17 Anthropogenic and natural sources, 196–197 Antihyperglycemic, 101 Antihypocholesterolemic, 101 Antioxidant system, 332 Antioxidative property, 365 AOX1 gene expression, 58 Aquaporins, 335, 336 Arabidopsis, 337, 338 Arabinosidases, 264 Arbuscular mycorrhizal (AM), 438 Arbuscular mycorrhizal fungi (AMF), 30, 195, 425 biochemical processes, 321 biodiversity, 423–427 biodiversity studies, 422 cation antiporter (see Cation channels and transporters) colonization, 420 community composition, 420 counter-balancing processes, 338 ecophysiology mechanisms extraradical AMF networks, 324, 325 and plants, 324 enhancement tolerance, plants, 329 extraction and propagation, 420–423 fungal symbiosis, 322, 323 germination, 427 life cycle, 323 microbiota, 419 mobilization and transportation, 427 morphology, 322, 323 mycelia, 324 mycoremediation, soil salinity, 321 mycoremediation-based research, 338

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500 Arbuscular mycorrhizal fungi (AMF) (cont.) nutrient uptake, 326, 327 AM colonization, plants, 327 interactions, Na+ and Cl-, 328 K+, 328 Na+, 328 P availability and activity, 328 P uptake, 327 salinity stress, 328 physiological processes, 321 plant biomass, 326, 327 plant growth-promoting bacteria, 420 plants, 321 pot-culture technique, 421 propagation, 421, 422 roots, 321 saline soils, 321 salinity-induced stress, 321 salinity stress remediation (see Salinity stress remediation) soil salinity (see Soil salinity) species, 420, 421 trap cultures, 422 Arbuscular mycorrhizas, 427 Arbuscules, 323 Arctic environment, 201 Aromatic peroxygenase (APO), 130, 192 Arrhenius equation, 408 Aryl alcohol oxidase (AAO), 106, 127, 128 applications, 129, 130 biochemical characteristics, 128 structure, 128, 129 Ascomycota, 240, 241, 244, 245 2-Azino-bis-(3-ethylbenzothiazoline-6-­ sulfonic acid (ABTS), 127 Azo dye (RB5) wastewater, 74, 89 B Basidiomycota, 240, 241, 244, 245 Batch adsorption study, 409–410 Batch biosorption equilibrium, 410 Benzene, toluene, ethylbenzene and xylene (BTEX), 248, 249 Benzoanthracene, 217 Benzopyrene, 119, 217 β-glucosidases, 7, 10, 354, 363 approaches, 268 aryl-β-glucosidases, 259 biotechnological applications, 261 biotechnology-based approaches, 272 cassava, 266 cellobiose, 258 cellulose degradation, 258

Index classification, 258 co-culturing, 269 CRISPR/Cas9, 272 deinking of waste paper, 266 flavour and nutrition enhancement, 264 fungi, 260 genetic manipulation, 270 high-value bioproducts, 263 hydrolyse flavonoid compounds, 265 isoflavone glycosides, 264 lignocellulose, 262 lignocellulose bioconversion (see Lignocellulose bioconversion) lignocellulose degradation, 271 metagenomics approach, 271 microbial sources, 261 mutagenesis, 270, 271 plant biomass, 263 sources, 259, 260 synthetic activity, 266 Betaines, 330 Bioaugmentation process, 225, 227–229, 243, 248, 249 advantages, 24 degradation properties, 24 DNA manipulation, 23 GEMs advantage, 24 biodegradation process, 24 biotic and abiotic factors, 24 genetic engineering microbes strategies, 24 rate-limiting steps, 24 recombinant DNA technology, 24 molecular biological techniques, 24 recombinant DNA, 24 Bioconversion edible food crops, 383 fungal consortium, 384 value-added products, 387 Bio-decolourization, 84 Biodegradation process, 205 Biofertilizers, 492, 497 Biofuel bioconversion, 383 biomass, 383 lignocellulose, 262, 263, 387 production, 261, 272, 382, 384, 387 Biofuels, 121 Bioherbicides, 493 Biological oxygen demand (BOD), 299 Biomass algal, 383 biofuels, 383

Index composition, 384, 385 ethanol production, 387 hemicellulosic fraction, 390 lignocellulosic, 382, 385, 387, 389, 391 plant, 387 pretreatment, 385, 386 Biomedical industries, 121 Bio-mineralization, 497 Bio-nanocomposites, 398, 401 Biopulping process, 307 Bioremediation, 64, 204, 444 advantages, 18 aerobic conditions, 21 agents, 18 application, 238, 244 atrazine, 249 biological processes, 19 chemical pollutants, 22 classifications, 20 definition, 18 extracellular enzymes, 18, 19 fungal enzymes, 19, 31–34, 239–240 (see also Enzymes) fungus, 19 hazardous chemical by fungi, 243 limitations, 25, 26 metabolic potential, 21, 22 microbes, 20, 21 microbial bioremediation, 19, 20 microorganisms, 20 monooxygenases act, 246 nitroaromatic compounds, 247 oil contaminated soil, 244 organic pollutants, 18 persistent organic pollutants, 249 phytoremediation, 19 pollutants, 238 types, 20, 22, 243 bioattenuation, 23 bioaugmentation, 23 biopiles, 25 biostimulation, 22 bioventing, 25 mycoremediation/fungal remediation, 26 Bioremediation mechanism, 199 Bioremediation processes, 467 environmental toxicity, 464 fungal enzymes, 465 metabolic potential, 463 oxidoreductases, 464 pH ranges, 464 white rot fungi, 464 xenobiotic pollutants, 464, 465, 467 xenome, 464

501 Biosensors, 165 Biosorbent, 397 chitin, 399 chlorophyll, 400 Biosorption, 397, 405 heavy metals and dyes, 405 thermodynamic behavior, 405 Biosorption phenomenon, 398 Biostimulation, 227, 229, 243, 248, 249 Biostimulation/bioaugmentation, 23 Biotechnological applications, 259 in agricultural sector, 367, 368 in animal feed sector, 366 in bakery industry, 364 β-glucosidases, 261 in biofuel industry, 359, 360 in biomedical sector, 364–366 in food processing, 361, 362 paper industry, 360, 361 pulping process, 360, 361 in textile industry, 363, 364 in waste management, 368, 369 in wine and brewery industry, 362, 363 Biotechnology, 312 Bisphenol A, 201 Branched absorbing structures (BAS), 325 Breakthrough curve (BTC) behavior, 411 Brown rot fungi (BRF), 11, 12, 357, 386 Brunauer–Emmett–Teller (BET), 408 Bubble column reactor, 414 C C1-compounds, 53 Carbohydrate esterases (CEs), 10, 11 Carbohydrate-active enzymes (CAZymes), 4, 6, 9, 10, 12, 13 Carbon sequestration, 286, 287 Carboxymethyl cellulase (CMCase), 132, 301, 303 Catalase, 244 Catalase activity, 443 Catalytic mechanism, 156 Cation channels and transporters CNGCs, 337, 338 Na+/H+ antiporters, 336, 337 Cellobiohydrolase, 7, 10, 11, 132, 257, 267, 387 Cellobiose, 257, 258, 263, 267, 271 Cellulase alkaline, 305 application, 361 biocatalysts, 303 deinking photocopier papers, 306

502 Cellulase (cont.) endoglucanase, 353 enzymatic deinking, 307 enzymatic pulping process, 305 exoglucanase, 353 fungal, 303 fungi vs. bacteria, 306 and hemicellulase, 303 inverting mechanism, 354 isolated Bacillus strains, 306 laccase mediator system, 307 methods, 306 office wastepaper, deinking, 306 OMG, 306, 307 ONP, 306, 307 retaining mechanism, 354 Cellulose in cotton fibres, 257 enzymatic hydrolysis, 258 microfibrils, 257 Cellulosome, 262 Cephalosporins, 122 Chemical oxygen demand (COD), 299 Chitin, 400 Chitin polymer molecule, 401 Chlorophenols, 119, 240, 248 Chlorophyll, 333 Chromium-inducing toxicity, 443 Chrysene, 217 Column adsorption study, 410–411 Common Effluent Treatment Plants (CETP), 78 Coniferyl alcohol, 384 Contaminants HMs (see Heavy metals (HMs)) Conventional techniques, 397 Coralene Golden Yellow (CGY), 92 Cosmetic industries, 121 Crude laccase, 92 Cutinase, 312 Cyclic guanosine monophosphate (cGMP), 337 Cyclic nucleotide-gated ion channels (CNGCs), 337, 338 Cyclic nucleotides monophosphate (CNMP), 337 Cyclosporin A, 122 CYP monooxygenases (P450s), 464 Cytochrome P450 (CYP), 225, 241, 247 D Decolouration, 84, 85, 88, 90–92, 120 Dehalogenases, 248 Dehydrins, 336

Index Deinking process biological, 299 cellulase, 306 chemical, 299 cutinase, 312 enzymatic technologies, 300 enzymatic treatment, 299 laccase, 310 lipase, 312 microbial enzymes, 300 xylanase, 309 Depolymerization, 2, 3, 6–9, 12 Desorption, 413 Deuteromycete fungus, 157 Deuteromycetes family, 155 Differential scanning calorimetry (DSC), 408 6-Dimethoxyphenol (DMP), 125 Dubinin–Radushkevich (D-R) equation, 407 Dye-decolorizing peroxidases (DyPs), 39 Dye degradation, 162 Dye industry, 76 Dyeing process, 79 CETP, 78 chemicals, 80 effluents, 80 Dyes, 400 Dyestuffs, 75 E Ecosystem’s equilibrium, 201 Ecosystems, 290 Effluents containing dyes, 93 Elovich equations, 407, 408 Endoamylases, 311 Endocrine disrupting chemicals (EDCs), 465 Endocrine disrupting compounds (EDCs), 165 Endoglucanase, 257, 258, 262, 269, 270, 353, 387 Energy-and cost-efficient passive phytoremediation methods, 438 Enterococcus pseudoavium, 311 Environmental contamination metallic properties, 195 soil and water, 194 toxic compounds, 194 Enzymatic deinking, 307 Enzymatic system, 221, 224 Enzyme commission (E.C.) number, 35 Enzyme immobilization methods, 161 Enzymes active sites, 31 advantages, 40, 41 apoenzyme, 35

Index biological moieties, 31 classification and types, 191 disadvantages, 41 E.C. number, 35 holoenzyme, 35 scope, 42 types, 35 catalase, 38 laccases, 36, 37 oxidoreductase, 36 peroxidases, 38–40 Epoxide hydrolases (EHs), 225 Ergot alkaloids, 491 Ethanol production, 382, 383, 391 Ethanol blending program (EBP), 382 Ex situ bioremediation technique, 20, 21 Exoglucanases, 257, 258, 262, 269, 270, 353 Extracellular oxidative enzymes, 193 F Facilitated diffusion system (FDS), 390 Ferulic acid, 386 Fibre-reactive azo dye, 75 Filamentous fungi, 453 Filamentous organisms growth process, 402 industrial waste product, 402 Fixed-bed experimental setup, 411 Fluidized bed reactor, 414 Fluoresomidearyl glucoside, 242 Fluoroquinolones, 242 Food industry, 120 Free radicals, 468 Freundlich model, 406 Fungal biomass acid pretreatment, 404 biomass, 404 thermal treatment, 405 Fungal bioreactor, 414 Fungal biosorbent, 408 Fungal biosorption, 413 Fungal biotechnology, see White biotechnology Fungal cellulase, 303 Fungal colonization, 26 Fungal communities bacterial and archaeal, 229 contaminated soils, 228 indispensable tool, 222 PAH-contaminated soil, 222 PAH degradation, 226, 227

503 polluted environments, 221 terrestrial and aquatic habitats, 221 Fungal consortium bioethanol production, 391 biomass, 384 development, 392 ethanol production, 392 laccase and xylanase enzymes, 384 Fungal decolourization processes, 93 Fungal divisions, 402 Fungal enzymes agricultural wastes, 358 agro-industrial wastes, 359 application, 479 biodegradation, 495 bioremediation, 239–240, 465, 475 catalytic reaction activity, 191 EDCs, 190 food and feed industries, 494 fungal extracellular enzymes, 191 geophysical area, 190 high-cost food, 491 hydrocarbons, 191 immobilization, 480 laccase biosynthesis, 358 limitations, 479, 482 marine fungi, 480, 481 mediators, 480 medicine, 494 multicellular fungal mycelium, 189 non-protein enzyme mimics, 481 oxidoreduction reactions, 192 remediation, 243, 244, 250 secondary metabolites, 190, 491 size and shape, 189 SSF, 358 synthesis of xylanase, 359 value-added products, 492 xenobiotic detoxification (see Xenobiotic detoxification) xenobiotic pollutants, 465, 467 xenobiotic remediation (see Xenobiotic remediation) Fungal metabolites, 491 Fungal mycelium, 26 Fungal pellet reactor, 415 Fungal peroxidase distal histidine, 168 mechanisms, 168–169 Fungal phytoremediation bioremediation, 444, 454 categorical classification, 444 eco-friendly, 454 factors, 451

504 Fungal phytoremediation (cont.) fungal species, 453 growth requirements, 452 HMs (see Heavy metals (HMs)) hydrocarbon, 452 mechanisms, 450, 451 metal contamination, soil, 448 pH, 452 precautions, 453 prerequisite precautions, 453 redox potential, 452 species, 444 temperature, 451, 452 Fungal species AMF, 30 dye decolorization, 27 ectomycorrhizal fungi, 30 electrostatic interactions with metals, 27 extremophilic fungi, 29, 30 laccases, 26, 27 marine fungi, 28, 29 white-rot fungi, 26–28 wood degraders, 27 yeasts, 27 Fungal strains, 390 Fungal unspecific peroxygenases, 246 Fungi AM, 438 bioherbicides, 493 biotransformation processes, 491 brown rots, 357 and carbon, 285 detoxification, environmental chemicals, 438 ecological and biochemical capacity, 438 ecosystems, 438 factor, soil, 288 fermentative industries, 492 (see also Fungal enzymes) growth ecosystem services, 289 fertilizer, 289–290 pesticides, 290 physical disturbance, 289 soil moisture, 290 soil pH, 289 temperature, 288 high-value products, 491 lignin-degrading enzyme producer, 356 literature, 291 and metals, 438 pectinolytic enzymes, 357 pharmaceuticals, 491 plant growth promoters, 493

Index play, 290 secondary metabolites, 491 soft rots, 357 sustainable fungal phytoremediation, 438 usage, 438 wastewater treatment, 398 white rots, 356 xylanolytic enzymes, 357, 358 Fungus, 286, 402 2,5-Furandicarboxylic acid (FDCA), 129 G Galacto-oligosaccharides, 266 Generally Regarded As Safe (GRAS), 137 Genetic diversity, methylotrophic yeast evolutionary analysis, 58 expression system, 58 gene regulation study, 54 H. polymorpha, 57, 58 LSU rRNA, 59 methanol utilisation, 59 O. Polymorpha, 59 phylogenetic analysis, 58 Pichia pastoris, 58 Pichia strains, 57 SSU rRNA, 59 strains, 60 woody materials, 56 Genetically engineered microbes (GEMs), 23 Global forest soils efflux, 288 Global sustainable environments, 429 Glucanases, 10 Glucomannans, 1 Glucosidic isoflavones, 264 Glycoside hydrolases (GHs), 9–11 Glycosyltransferase (GT), 4, 350 Glyoxal oxidase (GLOX), 106 Golgi apparatus, 350 Greenhouse gases (GHGs), 283 Guaiacyl, 104 H Haloalkaliphilic bacteria, 301 Hansenula polymorpha, 65, 66 Hard wood spent sulfite liquor (HSSL), 383 Hazardous chemical bioremediation, 243 definition, 237 degradation, 241 environmental, 237 fungal degradation, 248, 249 Heavy metal removing efficiency, 404

Index Heavy metals (HMs) alkaline desorption solution, 198 AM fungi, 438 anthropogenic essential nutrients, 439 sources, 439 water resources, 440, 441 binding, 197 bioaccumulation, 439 living beings, 442 MEUF, 439 on humans, 442 Ni and Cu, 198 nutrients availability, 197 on plants, 442, 443 phytoremediation, 444 Heme-thiolate peroxidase (HTP) applications, 131 biochemical characteristics, 130, 131 catalytic properties, 130 classification, 130 CPO, 130 Hemicellulase, 266, 312 xylan, 355 xylanase, 355 xylanolytic enzyme system, 356 xylooligosaccharides, 356 Hemicelluloses, 1, 2, 7, 9, 10, 262, 264, 267, 268 Hemicellulosic fraction, 386 Hidroxywarfarin, 242 High molecular weight (HMW), 202 High molecular weight PAHs (HMW-PAHs), 247 Histidines, 114 Homopolysaccharide, 267 Humic substances (HS), 173 Hydrolases, 193–194 Hydrolytic enzymes cellulose/hemicellulose, 121 class, 137 complex lignocellulosic materials, 102 and non-hydrolytic, 107 Hydrophobic cluster analysis (HCA), 258 Hydroxyalkylation, 105 4-Hydroxybenzoic acid, 386 2-Hydroxycarbamazepine, 242 Hydroxyflumequine, 242 Hydroxylation, 204 I In situ bioremediation technique, 20, 21 Indian Agro and Recycled Paper Mills Association (IARPMA), 299

505 Indian dyestuff industry, 76 Industrial applications lignocellulose process, 104–106 in VP, 127 Industrial biotechnology, 496 Industrial dye effluents, 92–93 Industrial Wasteland Soil, 425–427 Inorganically Managed Agricultural Soil, 424–425 Intergovernmental Panel on Climate Change (IPCC), 283 Intra-radical arbuscules, 325 Ion influx, 337 Isobutyl-galactosides, 266 Isoflavone glycosides, 264 K Konzo syndrome, 266 Kraft lignin, 105 Kraft process, 361 L Laccase-based bioremediation, 158 Laccase catalytic cycle, 85 Laccase enzyme, 160 Laccase-mediator systems (LMSs), 307, 313, 475 Laccase producing fungi, 155 Laccases (LACs), 83, 84, 107, 114, 116, 244, 309, 310, 467 agriculture and industrialization, 158 applications, 118 biological functions, 158 bioprocessing, 116, 117 bioremediation, 154 catalytic domain, 157 covalent attachment, 161 cross-linked method, 162 detoxification copper involvement, 475 LMSs, 475 multicopper oxidases, 475 redox potentials, 476 TNC, 476 type I copper, 475 type II copper, 476 type III copper, 476 encapsulation, 161 enzyme-supportive matrix, 161 extracellular and intracellular enzymes, 154 functions, 156 fungal laccases, 156

506 Laccases (LACs) (cont.) fungi and plants, 156 glycosylation, 156 growing fungi, 159 immobilization, 159 mechanism, 157 paper and pulp industry effluents, 154 polyphenol oxidase, 154 remediation applications, 468, 469 aromatic compounds, 468 detoxification of PCBs, 468 free radicals, 468 ligninolytic enzymes, 468 PAHs, 468 pH range, 468 toxic dyes, 469 structure, 156–157 toxic environment, 159 types, 156 Langmuir isotherm model equation, 406 Late embryogenesis abundant (LEA) proteins, 336 Leghemoglobin, 334 Legume colonization, 334 Leucine pNA (LPNA), 135 Lichens, 287 Lignin, 298 polymer monomeric units, 103 Lignin degradation, 390 Lignin-degrading enzymes actions, 107 Lignin-modifying enzymes (LME), 192, 387 Lignin peroxidase–graphite electrode biosensor systems, 171 Lignin peroxidase (LiP), 39, 40, 81, 83, 106, 125, 169–170, 243, 245, 249, 387, 464 Lignin vs. xenobiotic chemical structures, 466 Ligninase enzymatic reactions, 352 heme peroxidases, 352 laccase, 353 LiPs, 352 MnP, 352 oxidoreductive class, 351, 352 Ligninolytic extracellular enzymes, 171 Ligninolytic fungi, 123 Ligninolytic oxidoreductases, 106 Lignins, 102 Lignocellulose, 382, 384, 387, 389 biorefineries, 127 cellulose and hemicellulose, 129 and industrial applications, 104–106

Index Lignocellulose bioconversion challenges, 268 end-product inhibition, 267 plant biomass, 267 pre-treatment methods, 267 Lignocellulose degradation, 263, 267, 271 Lignocellulosic biomass applications (see Biotechnological applications) cellulase, 353, 354 fungi, 356 hemicellulase, 355, 356 hemicellulose, 350, 351 lignin polymer, 350 ligninase, 351–353 monolignol biosynthesis, 350 morphological diversity of PCW, 349 PCW, 349 pectinase enzyme, 354, 355 RTC, 350 Lignolytic fungi, 26 Lignosulfonate lignin, 105 Lignosulfonates, 106 LiP and MnP enzymes, 172 Lipases, 137, 311, 312 Lovastatin, 491 Lytic polysaccharide monooxygenases (LPMOs), 7, 10 M Manganese peroxidase (MnP), 40, 81–84, 88, 106, 122, 170, 243, 245, 249, 464 application, 124 characterization, 122, 123 mechanism of action, 123, 124 Manganese peroxidases, 40, 170 Mannans, 1 Mediators, 467 Meiosporic ascomycetes, 240 MEROPS database, 135 Metagenomics, 271 Metal pollutants, 438 Metal tolerance, 438 Metal toxicity, 442, 443 Metaproteomics, 222 Methane cycle, 54 Methanol, 54 Methanol oxidising genes, 54 Methylotrophic yeast biotechnological application, 64 environmental impact bioremediation, 64 bioremediation, soil, 66

Index biotechnological applications, 63 heavy-load wastewater treatment process, 65 lactate-selective microbial biosensor, 65 oil-contaminated soil, 65 Pichia pastoris, 65 plant growth promotion, 63 pollutants, 65 genetic diversity, 55–56 (see also Genetic diversity, methylotrophic yeast) genetic regulation, 60, 62, 63 glycerol metabolism, 54 metabolism and physiology, 57 methane cycle, 54 methanol inducible gene expression, 61 metabolism, 62 oxidising genes, 54 methanol utilisation, 53 recycling, carbon, 54 Micellar-enhanced ultrafiltration (MEUF), 439 Microbial enzymes alkaline-tolerant fungi, 301 amylases, 310, 311 API, 302 avicelase, 303 biological deinking, 299–300 biological process, 299 biotechnical applications, 298 cellulases (see Cellulases) CMCase, 303 Crude cellulose, 302 cutinase, 312 deinking process, 299 deinking wastepapers, 297 EndoA, 302 EndoB, 302 enzymatic technologies, 300 enzymatic treatment, 299 haloalkaliphilic bacteria, 301 Heleococcum alkalinum sp., 301 hemicellulase, 312 IARPMA, 299 laccase alkaliphilic laccase activity, 310 bleaching, kraft pulp, 310 ink removal capability, 310 plants and fungi, 310 lignin, 298 lipases, 311, 312 MOW, 299 natural polymers, 297 novel alkalothermophilic actinomycete, 301 papermaking, 298

507 Penicillium citrinum, 302 photocopier printers, 299 pulp, 298 pulp and paper industry, 297 SSF, 303 sustainability issues, 299 thermophilic, novel, 301 wastepaper (see Wastepaper) whole-cell hydrolysates, 301 xylanase, 303 (see also Xylanase) Microeukaryotes, 220 Microorganism, 238, 240, 241, 243, 245, 249 autochthonous, 227 PAH-degrading, 219 Microwaves, 105 Middle lamella layer, 349 Mitomycin, 122 Mixed office wastepaper (MOW), 312 Molecular biological techniques, 24 Monolignol biosynthesis, 350 Monooxygenases, 246 Mucoromycotina, 241, 242, 244 Multicopper oxidases, 83 Multifarious techniques, 409 Municipal office waste (MOW), 299 Mushroom enzymes, 137 Mutagenesis, 270 Mycoremediation, 189, 195, 444, 454 biotransformation, 200 exclusion, 199 extrusion, 199 fixation, 200 stress condition, 199 Mycoremediation/fungal remediation, 26 Mycorrhiza, 429, 453 Mycorrhizal-based technology, 426 Mycorrhizal fungi hyphae, 286 Mycorrhization, 429 Mycorrhizosphere, 419 Mycosorption, 429 N Na+/H+ antiporters, 321, 337 Nanomaterial-based approaches, 398 Naphthalene, 204–205, 217, 228 National Oceanic and Atmospheric Administration (NOAA), 284 Natural Biomass Utilization Systems (NBUS), 3 Nitrogen fixation, 334 Nitrophenols, 119 Nitroreductases, 247 Nodulation, 334

508 Nucleotide sequence identity (NCI), 258 Nutrient uptake, 320, 325–328 O Ogataea polymorpha, 59 Ogataea thermomethanolica TBRC656, 62, 63 Old magazine (OMG), 306 Old newspaper (ONP), 306 Organic biomass, 383 Organic compounds, 200 Organically Managed Agricultural Soil, 425 Organosolv lignin, 105 Organosolv process, 105 Osmoregulation, 330 Osmotic stress, 336 Oxidoreductases, 192–193, 464 Oxyalkylation, 105 Oyster mushroom, 101, 102 P P5CS-encoding gene, 335 Packed Bed Column Model, 411–412 BDST model, 412 kinetic data, 413 Thomas model, 411 Paper and pulp industry, 173 Paper industries, 121, 162–164 P-coumaryl alcohol, 384 Pectin esterase, 354 Pectinase, 133, 134 hydrolases, 354, 355 lyases, 355 pectin esterase, 354 Pectinolytic enzymes, 357 Pectinylase (PL), 134 Peroxidase, 245 Peroxidase catalytic activity, 169 Peroxidase–catalase superfamily (PCATS), 167 Peroxidase-cyclooxygenase superfamily (PCOXS), 166 Peroxidases, 166, 241, 243, 245–247 detoxification cytochrome P450, 478 LiPs, 476, 477 MnPs, 477 heme peroxidases, 38, 39 lignin, 39, 40 LiP, 166 manganese, 40 non-heme peroxidases, 39 PCOXS, 166

Index remediations LiP, 469, 470 MnPs, 470, 471 P450s, 472, 473 UPOs, 474 VP, 471 types, 166 versatile, 40 Persistent organic pollutants (POPs), 203 Pesticides, 194, 195, 200–205, 240, 242, 248, 249, 290 pH, 452 Phanerochaete chrysosporium, 9, 10 Pharmaceutical industries, 121, 398 Pharmaceutical personal care products (PPCPs), 190 Pharmaceutical protein production system, 66 Phenanthrene, 217, 224, 225, 228, 229 Phenol 2-monooxygenases, 246 Phenol oxidases, 192 Phenylpropane lignin, 106 Phenylpropene monomers, 103 Phosphorus(P)-based fertilizers, 289 Photocopier printers, 299 Photocopier wastepaper, 306 Photosynthates, 286 p-Hydroxyphenyl, 104 Phylogeny, 55, 59 Phytoremediation, 19, 437, 444 Pichia pastoris, 58, 61, 63 Pichia strains, 57 Plant cell wall (PCW), 349 Plant growth promoters, 491–493 Pleurotus ostreatus amylases, 134, 135 cellulases, 132 different enzymes, 108–113 laccase isoenzymes, 116 lipases, 137 manganese peroxidase IV, 123 pectinase, 133, 134 proteases, 135, 136 PUF cubes, 118 xylanases, 131–133 Polluted soils metatranscriptomics, 222 microbial diversity, 229 PAH, 220–223, 228 Polyamines, 331 Polyaromatic hydrocarbons (PAHs), 18, 119, 464, 465, 468 biodegradation, 202–203 contamination, 202 oxidation, 202

Index Polychlorinated biphenyls (PCB), 164, 248 laccase-mediated degradation rate, 164 Polychlorinated dibenzofurans (PCDFs), 248 Polychlorinated dibenzo-p-dioxins (PCDDs), 248 Polycyclic aromatic compounds (PAC), 217 Polycyclic aromatic hydrocarbon (PAH), 164, 174, 240–242, 247–249, 452 atmospheric and aquatic transport, 219 barcode-primer pair, 223 biological sciences, 230 contaminated soils, 227–229 culture-dependent techniques, 222 culture-independent techniques, 222, 223 environments, 217, 220 extracellular mechanism, 224, 225 fungal communities, 226, 227 fungal mechanisms, 223, 224 genetic engineers, 230 heterocyclic, 218–219 intracellular enzymatic pathways, 225, 226 microbial communities, 230 phenanthrene degradation pathway, 225 photodecompositions, 219 physicochemical properties, 217 polluted soils, 220–222 Polysaccharide lyases (PL), 4 Polyurethane foam (PUF), 134 Polyvinylpyrrolidone (PVP), 362 Pot-culture technique, 421 Powdered activated carbon (PAC), 18 Prerequisite precautions, 453 Primary cell wall (PCW), 349 Proline, 330, 335 Proteases, 135, 136 Proton symport (PS) mechanism, 390 Proxidases classification, 167 Pulping process, 360 Pyrroline-5-carboxylate reductase (P5CR), 330 Pyrroline-5-carboxylate synthetase (P5CS), 330, 335 Q Quinone reductases, 247 R Reactive Black 5 (RB5), 85 Reactive dyes, 75 Reactive nitrogen species (RNS), 442 Reactive oxygen species (ROS), 38, 221, 244, 325, 442

509 Reactors, 414 Recalcitrant, 221, 238, 248, 249 Recombinant DNA technology, 24, 491 Redox mediators, 157 Redox potentials, 168, 452 Remazol Brilliant Blue R (RBBR), 119, 120 Remazol Brilliant Blue SN4R, 120 Response surface methodology (RSM), 85 Resting oxidized state (RO), 476 Rhizo-deposited carbon, 287 Rhizo-deposition, 287 Rhizo-deposition material, 290 Rhizospheric soil, 423 Rosette terminal complex (RTC), 350 S Saline soil AM fungi, 321 and irrigation, 320 salt concentration, 319 Salinity stress remediation biochemical change antioxidant system, 332 osmotic adjustment, 330, 331 osmotic stress, 330 phytohormones regulation, 331 polyamines, 331 molecular changes, plants aquaporins, 335, 336 heterokaryotic and obligate nature, 335 LEA proteins, 336 P5CS, 335 physiological changes chlorophyll content, 333 nitrogen fixation, 334 nodulation, 334 relative permeability, 333 water, 333, 334 Salt-affected soils, 319 Salt-stressed soil, 321 Secondary cell wall (SCW), 349 Secretome fungal, 4, 7 wood-degrading fungi, 2 Sexual basidiospores, 241 Sinapyl alcohols, 2, 384 Single-spore culture, 422–423 Soil, 319 Soil bioremediation, 119, 205–206 Soil fungi, 285 Soil moisture, 290 Soil organic matter (SOC), 284–285

510 Soil salinity AMF, 326 development, salt-tolerant crop varieties, 321 dissolved mineral salts, 325 nutrient imbalance, 325 nutritional disorders, 320 physiological stresses, 325 plant cell membranes, 325 reduced growth, 320 salt concentrations, 320 significance, 320 stress signals, 325 Soil salinization, 319, 320 Solid–liquid operations, 168 Solid-state fermentation (SSF), 132, 303, 358, 386 Sorption mechanism, 405 Starch and cellulose, 287 Stirred tank reactor (STR), 414 Stress tolerance, 324 Subphylum mucoromycotina, 240 Subtropical agricultural soil, 423–424 Symbiosis, 427–429 Syringaldazine, 116 Syringyl, 104 T Temkin isotherm, 407 Temperature, 451, 452 Textile dyes azo groups, 74 colourants, 74 fabrics, 74 Textile effluent treatment, 162 Textile industries, 73, 76, 121 dye, 171 Theaflavin (TF), 362 Thearubigin (TR), 362 Thermochemical hydrolysis, 104 Thermodynamic behavior, 405 Thermotolerance, 270 Thermotolerant yeast, 57 Thomas model, 411 Thyroid peroxidase (TPO), 166 Toxic agrochemicals, 174 Trametes hirsute fungus laccase, 115 Transcription factors, 62 Transcriptome, 7–9, 13 Transglycosylation, 261, 266 Trap culturing, 422 Trichoderma viride, 302

Index 1,1,1-Trichloro-2,2-bis(4-chlorophenyl) ethane (DDT), 248 Trinitrotoluene (TNT), 18, 240, 242, 247–249 Trinuclear cluster (TNC), 476 Triphenylmethane, 119 2,4-Dichlorophenol (2,4-DCP), 127 Tyrosinases, 244 U Ultrasonication, 386 United Nations Environment Program (UNEP), 319 Unspecific peroxygenase (UPOs), 39, 474 V Veratryl alcohol, 245 Versatile peroxidases (VP), 40, 170, 172, 193, 245, 471 applications, 127 biochemical characteristics, 125 CIP, 125 lignin manganese peroxidases, 124 ligninolytic enzyme, 125 in Pleurotus sp., 124 structural characteristics, 125, 127 Vesicular-arbuscular mycorrhizae, 493 Vinblastine, 122 Volcanic eruption, 217 W Wastepapers cutinase, 312 deinking biological process, 299 chemical process, 299, 300 conventional methods, 297 enzymatic pretreatment, 311 laser-printed office papers, 299 microbial enzymes, 299 industrial utilization, 297, 313 raw materials, 298 Wastewater treatment, 119, 120, 398, 399 Water, 333, 334 Water toxication, 440, 441 White biotechnology advantages, 494 bio-based products, 496 eco-friendly, 494 greenhouse emissions, 494

Index in medical, 491 renewable natural resources, 496 White rot fungi (WRF), 3, 6, 9, 10, 73, 153, 173, 203, 224, 225, 356, 386–389, 391, 464, 482 colour reduction levels, 89 decolouration, 90 dye degradation, 86–88 En3, 90 enzymes, 89 laccases, 80, 81, 83 ligninolytic enzymes, 73 lignolytic systems, 85 LiP, 81 manganese peroxide and lignin peroxide, 84 MnP, 81 nitrogen, 85 nitrogen-limited glucose ammonium media, 88 oyster mushroom, 74 recalcitrant nature, 93 toxicity and xenobiotic nature, 92 types, 81 xenobiotic degradation capacity, 73 Wood biopulping, 310 cheap source, paper pulp, 312 chipping, 307, 310 debarking, 307 lignin biodegradation, 297 pulp and paper industry, 297, 313 screening, 307 triglycerides, 312 Wood-degrading fungi, 2, 4, 7–9 Wood-rotting fungi, 106 Woollen industry, 76

511 X Xenobiotic compounds, 463 Xenobiotic detoxification LACs, 475, 476 peroxidases, 476–478 silico approaches, 481 Xenobiotic pollutants, 465 Xenobiotic remediation bioremediation processes, 467 fungal strains, 479 heme peroxidases, 467, 479 heterologous expression, 479, 482 LACs, 468, 469 mediators, 467 oxidoreduction-catalyzing enzymes, 467 peroxidases (see Peroxidaes) Xenobiotics, 242 Xenome, 464 Xylan, 307 Xylanase, 9, 12, 132, 133, 307–309 Bacillus pumilus SV-205, 308 biological treatment, 309 biopulping process, 307 deinked ONP pulp, 309 hydrolysis, hemicellulose, 307 optimizing submerged fermentation conditions, 308 pollution free environment, 308 SSF, 308, 309 xylano pectinolytic, 309 Xylanolytic enzyme system, 356–358 Xyloglucans, 1 Xylulose, 391 Z Zygospores, 241