Fungi in Fuel Biotechnology [1st ed.] 9783030444877, 9783030444884

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Fungal Biology

Gholamreza Salehi Jouzani Meisam Tabatabaei Mortaza Aghbashlo   Editors

Fungi in Fuel Biotechnology

Fungal Biology Series Editors Vijai Kumar Gupta AgroBioSciences (AgBS) and Chemical & Biochemical Sciences (CBS) Department Mohammed VI Polytechnic University (UM6P) Benguerir, Morocco 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

Gholamreza Salehi Jouzani Meisam Tabatabaei • Mortaza Aghbashlo Editors

Fungi in Fuel Biotechnology

Editors Gholamreza Salehi Jouzani Microbial Biotechnology Department Agricultural Biotechnology Research Institute of Iran (ABRII) Agricultural Research, Education and Extension Organization (AREEO) Karaj, Iran Mortaza Aghbashlo Department of Mechanical Engineering of Agricultural Machinery Faculty of Agriculture Engineering and Technology College of Agriculture and Natural Resources University of Tehran Karaj, Iran

Meisam Tabatabaei Institute of Tropical Aquaculture and Fisheries (AKUATROP) Universiti Malaysia Terengganu Terengganu, Malaysia Microbial Biotechnology Department Agricultural Biotechnology Research Institute of Iran (ABRII) Agricultural Research, Education and Extension Organization (AREEO) Karaj, Iran

ISSN 2198-7777     ISSN 2198-7785 (electronic) Fungal Biology ISBN 978-3-030-44487-7    ISBN 978-3-030-44488-4 (eBook) https://doi.org/10.1007/978-3-030-44488-4 © Springer Nature Switzerland AG 2020 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Preface

Second-generation biofuels, including lignocellulosic-based bioethanol and waste oil-oriented biodiesel, have been considered promising in mitigating greenhouse gas emissions resulting from wide use of petro-fuels and in combating climate change. However, since these kinds of biofuels require additional pretreatment and hydrolysis steps, development of more economically viable technologies is of importance. The present book offers a comprehensive review on the obstacles faced in the production of waste-based liquid biofuels with a focus on the potentials of different fungi and yeasts in bioethanol and biodiesel production. Overall, fungi could be effectively used in biofuels production in three different ways: application as the source of cellulase for hydrolyzing lignocellulosic materials for bioethanol production, as the source of lipids for biodiesel production, or as the source of lipase for catalyzing the transesterification of lipids into biodiesel production. In this context, different types of fungi, including plant-pathogenic fungi, fungal biocontrol agents, endophytic fungi, wood-rotting fungi, and anaerobic rumen fungi as well as their potentials for bioethanol and biodiesel production have been reviewed and thoroughly discussed in Chaps. 2, 3, 4, 5, 6, and 7 of the present book. Moreover, basic considerations in designing bioprocesses for fungal-based biofuel production systems are presented in Chap. 8. One of the most promising third-generation feedstocks for biodiesel production is the microbial oil obtained from oleaginous fungi. Accordingly, Chap. 9 is aimed at briefly reviewing oleaginous fungi and their application for biodiesel production while comprehensively presenting the principles and implementation of life cycle assessment (LCA) for measuring the environmental impacts associated with fungal biodiesel production systems. It is expected that the present book would assist both the scientific and industrial communities in further developing fungal-based biofuel production systems. We are thankful to authors of all the chapters for their efficient cooperation and also for their readiness in revising the manuscripts. We also would like to extend our appreciation to the reviewers who in spite of their busy schedule assisted us by evaluating the manuscripts and provided their critical comments to improve the manuscripts.

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We sincerely thank the team of Springer Nature, in particular Professor Vijai Kumar Gupta, Dr. Eric Stannard, Mr. Rahul Sharma, and Ms. Savita Rockey Samuel, for their cooperation and invaluable efforts in producing this book. Karaj, Iran Terengganu, Malaysia  Tehran, Iran

Gholamreza Salehi Jouzani Meisam Tabatabaei Mortaza Aghbashlo

Contents

1 Biofuels: Types, Promises, Challenges, and Role of Fungi������������������    1 Gholamreza Salehi Jouzani, Mortaza Aghbashlo, and Meisam Tabatabaei 2 Bioethanol Production by Using Plant-­Pathogenic Fungi��������������������   15 Amin Alidadi, Hamed Kazemi Shariat Panahi, Mona Dehhaghi, Reeta Rani Singhania, Hossein Ghanavati, Reza Sharafi, Mortaza Aghbashlo, Meisam Tabatabaei, and Gholamreza Salehi Jouzani 3 Fungi as Bioreactors for Biodiesel Production��������������������������������������   39 Meisam Tabatabaei, Amin Alidadi, Mona Dehhaghi, Hamed Kazemi Shariat Panahi, Su Shiung Lam, Abdul-Sattar Nizami, Mortaza Aghbashlo, and Gholamreza Salehi Jouzani 4 Fungal Biocontrol Agents as a New Source for Bioethanol Production������������������������������������������������������������������������������������������������   69 Hamed Kazemi Shariat Panahi, Mona Dehhaghi, Gholamreza Salehi Jouzani, Rasoul Zare, Mortaza Aghbashlo, and Meisam Tabatabaei 5 Endophytic Fungi for Biodiesel Production������������������������������������������  105 Cristiano E. Rodrigues Reis, Heitor B. S. Bento, Ana K. F. Carvalho, Yan Yang, Heizir F. de Castro, and Bo Hu 6 Wood-Rotting Fungi for Biofuel Production ����������������������������������������  123 Ichiro Kamei 7 Anaerobic Rumen Fungi for Biofuel Production����������������������������������  149 Mona Dehhaghi, Hamed Kazemi Shariat Panahi, Gholamreza Salehi Jouzani, Sivakumar Nallusamy, Vijai Kumar Gupta, Mortaza Aghbashlo, and Meisam Tabatabaei

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8 Process Design in Fungal-Based Biofuel Production Systems ������������  177 Behzad Satari and Keikhosro Karimi 9 Life Cycle Analysis for Biodiesel Production from Oleaginous Fungi����������������������������������������������������������������������������  199 Homa Hosseinzadeh-Bandbafha, Meisam Tabatabaei, Mortaza Aghbashlo, Anh Tuan Hoang, Yi Yang, and Gholamreza Salehi Jouzani Index������������������������������������������������������������������������������������������������������������������  227

Contributors

Mortaza  Aghbashlo  Department of Mechanical Engineering of Agricultural Machinery, Faculty of Agricultural Engineering and Technology, College of Agriculture and Natural Resources, University of Tehran, Karaj, Iran Amin Alidadi  Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran Heitor  B.  S.  Bento  Chemical Engineering Department, Engineering School of Lorena, University of São Paulo, São Paulo, Brazil Ana  K.  F.  Carvalho  Chemical Engineering Department, Engineering School of Lorena, University of São Paulo, São Paulo, Brazil Heizir  F.  de  Castro  Chemical Engineering Department, Engineering School of Lorena, University of São Paulo, São Paulo, Brazil Mona Dehhaghi  Department of Microbial Biotechnology, School of Biology and Centre of Excellence in Phylogeny of Living Organisms, College of Science, University of Tehran, Tehran, Iran Faculty of Medicine and Health Sciences, Macquarie University, Sydney, NSW, Australia Biofuel Research Team (BRTeam), Karaj, Iran Hossein  Ghanavati  Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran Vijai  Kumar  Gupta  AgroBioSciences (AgBS) and Chemical & Biochemical Sciences (CBS) Department, Mohammed VI Polytechnic University (UM6P), Benguerir, Morocco Anh  Tuan  Hoang  Faculty of Mechanical Engineering, Ho Chi Minh City University of Transport, Ho Chi Minh city, Vietnam ix

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Engineering Institute, Ho Chi Minh city University of Technology (HUTECH), Ho Chi Minh city, Vietnam Homa  Hosseinzadeh-Bandbafha  Department of Mechanical Engineering of Agricultural Machinery, Faculty of Agricultural Engineering and Technology, College of Agriculture and Natural Resources, University of Tehran, Karaj, Iran Bo  Hu  Department of Bioproducts and Biosystems Engineering, University of Minnesota, Saint Paul, MN, USA Ichiro Kamei  Faculty of Agriculture, University of Miyazaki, Miyazaki, Japan Keikhosro  Karimi  Department of Chemical Engineering, Isfahan University of Technology, Isfahan, Iran Industrial Biotechnology Group, Research Institute for Biotechnology and Bioengineering, Isfahan University of Technology, Isfahan, Iran Hamed Kazemi Shariat Panahi  Department of Microbial Biotechnology, School of Biology and Centre of Excellence in Phylogeny of Living Organisms, College of Science, University of Tehran, Tehran, Iran Faculty of Medicine and Health Sciences, Macquarie University, Sydney, NSW, Australia Biofuel Research Team (BRTeam), Karaj, Iran Su  Shiung  Lam  Pyrolysis Technology Research Group, Institute of Tropical Aquaculture and Fisheries (AKUATROP) & Institute of Tropical Biodiversity and Sustainable Development (Bio-D Tropika), Universiti Malaysia Terengganu, Kuala Nerus, Terengganu, Malaysia Sivakumar Nallusamy  Department of Biology, College of Science, Sultan Qaboos University, Muscat, Oman Abdul-Sattar  Nizami  Sustainable Development Study Center, Government College University, Lahore, Pakistan Cristiano  E.  Rodrigues  Reis  Chemical Engineering Department, Engineering School of Lorena, University of São Paulo, São Paulo, Brazil Gholamreza  Salehi  Jouzani  Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran Behzad Satari  Department of Food Technology, College of Abouraihan, University of Tehran, Tehran, Iran Reza  Sharafi  Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran

Contributors

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Reeta  Rani  Singhania  Centre for Energy and Environmental Sustainability, Lucknow, India Meisam Tabatabaei  Institute of Tropical Aquaculture and Fisheries (AKUATROP), Universiti Malaysia Terengganu, Terengganu, Malaysia Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran Yan Yang  Department of Bioproducts and Biosystems Engineering, University of Minnesota, Saint Paul, MN, USA Yi Yang  Department of Bioproducts and Biosystems, University of Minnesota, St. Paul, MN, USA Key Lab of Urban Environment and Health, Institute of Urban Environment, Chinese Academy of Sciences, Xiamen, China Institute on the Environment, University of Minnesota, St Paul, MN, USA Rasoul Zare  Iranian Research Institute of Plant Protection, Agricultural Research, Education and Extension Organization (AREEO), Tehran, Iran

Chapter 1

Biofuels: Types, Promises, Challenges, and Role of Fungi Gholamreza Salehi Jouzani, Mortaza Aghbashlo, and Meisam Tabatabaei

1.1  Introduction The world energy demand is predicted to increase to 6.6  ×  1020  J in 2020 and 8.6 × 1020 J in 2040 (Guo et al. 2015), leading to an increase in the utilization of fossil-oriented energy carriers as the main energy sources, particularly by the industrial and transportation sectors. However, these kinds of fuels are nonrenewable, their reserves are limited, and, more importantly, their exploitation is linked with serious environmental and health concerns (Hosseinzadeh-Bandbafha et al. 2018). Among those concerns is the critical increase in the amounts of atmospheric

G. Salehi Jouzani (*) Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran e-mail: [email protected] M. Aghbashlo (*) Department of Mechanical Engineering of Agricultural Machinery, Faculty of Agricultural Engineering and Technology, College of Agriculture and Natural Resources, University of Tehran, Karaj, Iran e-mail: [email protected] M. Tabatabaei (*) Institute of Tropical Aquaculture and Fisheries (AKUATROP), Universiti Malaysia Terengganu, Terengganu, Malaysia Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran e-mail: [email protected]

© Springer Nature Switzerland AG 2020 G. Salehi Jouzani et al. (eds.), Fungi in Fuel Biotechnology, Fungal Biology, https://doi.org/10.1007/978-3-030-44488-4_1

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greenhouse gases (GHG) resulting in global warming and climate change (Strobel 2015; Salehi Jouzani and Taherzadeh 2015). The International Energy Agency (IEA) has suggested to limit the global temperature increase to 2 °C to address these challenges, based on which, by the year 2060, the global CO2 emissions should be reduced by up to 70% compared to the 2014 baseline. These have been pushing policy makers and scientists to seek and develop alternatives sources of energy to partially or completely replace fossil fuels (Soltanian et al. 2019). For instance, the transport sector alone generates approximately 23% of total CO2 emissions (Oh et  al. 2018), highlighting the importance of its decarbonization. To achieve such targets, the application of renewable energy carriers such as biofuels is instrumental (Khalife et al. 2017). It has been confirmed that the wide global application of various generations of biofuels could have deep positive impacts on the environment (soil, water, land, and wild life) as well as on the human and animal health while also exerting positive socioeconomic effects on civil development (Ahlgren et al. 2017). To achieve the target set forth by the 2 °C scenario, biofuels production must be increased by a factor of 10 (Oh et al. 2018). However, the major challenge would be to supply a sufficient deal of feedstock. This becomes more critical given the fact that the majority of the biofuels produced currently relies on edible resources such as sugar and vegetable oil (i.e., first-generation biofuels). On the contrary, biomass could serve as a promising feedstock for biofuels production (i.e., second-generation biofuels), posing no threat to food security (Aghbashlo et al. 2018). It is estimated that the annual terrestrial dry biomass production stands at about 120 × 1015 g, which is capable of storing more than 2.2 × 1021 J of energy. This amount of energy, which is captured annually by plants, is three to four times greater than the human energy need (Guo et al. 2015; Haberl et al. 2013). Biofuels exist in different forms, including gaseous, liquid, and solid forms, which are discussed in the subsequent sections.

1.2  Different Types of Biofuels Based on the feedstock used, biofuels are classified into three generations. First-­ generation biofuels are those produced using edible resources, such as starch (corn, potato, and wheat) and oilseed crops (rapeseed, soybean, and sunflower). The second-­generation biofuels include those derived from inedible crops or agricultural wastes (lignocellulosic materials). Application of different microorganisms such as microalgae for production of different types of biofuels, such as biodiesel and bioethanol, results in the production of third-generation biofuels (Raven et al. 2019). Based on the physical state of biofuels, they are also classified into three forms as follows (Fig. 1.1) (Guo et al. 2015): 1 . Solid biofuels: wood pellets, wood chips, wood charcoal, and firewood 2. Liquid biofuels: biodiesel, bioethanol, drop in biofuels, and pyrolysis bio-oil 3. Gaseous biofuels: biogas and syngas

1  Biofuels: Types, Promises, Challenges, and Role of Fungi Fig. 1.1 General classification of the globally used biofuels

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Biofuels

Solid biofuels

Liquid biofuels

Gaseous biofuels

Fire woods

Bioethanol

Biogas

Wood chips

Biodiesel

Syngas

Wood pellets

Pyrolysis biooil

Charcoal

Drop in biofuels

1.2.1  Solid Biofuels Biomass feedstocks used for production of different solid biofuels are commonly agricultural, forestry, and waste-oriented resources. Based on the European standards, the classification of solid biofuels is based on their origin and source, and accordingly, solid biofuels are divided into four subcategories: (1) woody biomass, (2) herbaceous biomass, (3) fruit biomass, and (4) blends and mixtures (Guo et al. 2015). Plant materials, such as wood, straw, and wood chips, have been directly used for burning since thousands of years ago. In fact, prior to the application of fossil fuels, firewood was the most important energy source for domestic purposes. Direct combustion of firewood is among the most common strategies to generate bioenergy. However, this application has some disadvantages; for instance, firewood is bulky and not suitable for small and automated heating systems. To address this challenge, wood chips, i.e., small pieces of wood, have been introduced which could be easily used as energy source for heating and electricity generation (Shepherd 2000). Wood pellets are a more processed form of solid biofuels with their volumetric energy content reaching 18 MJ.m−3. Different types of plant biomass, such as tree trunks, grasses, crop residues, and nutshells, can be efficiently used for pellet production. It should be noted that the energy contents of wood fuels (firewood, woodchips, and wood pellets) are generally lower than those of fossil fuels. To overcome this shortcoming, charcoal, a carbon-enriched, porous, and black solid, was produced by the ancient populations through using different types of pyrolysis techniques. More specifically, to produce charcoal, wood materials are heated in a kiln or a retort at 400  °C in the absence air. The energy content of the resultant charcoal is about

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28–33 MJ.kg−1, which is significantly higher than that of its fossil-oriented counterparts, i.e., coal.

1.2.2  Liquid Biofuels The most common types of liquid biofuels include bioethanol, biodiesel, and biobutanol. –– Bioethanol Bioethanol is an alcohol produced through the fermentation of organic matters mostly by the yeast Saccharomyces cerevisiae. Bioethanol contains 34.7% oxygen and, therefore, compared to gasoline, which does not contain oxygen, is an eco-­ friendly oxygenated fuel (Zabed et al. 2017). The most widely commercially practiced technology for bioethanol production is by fermentation of simple sugars (monosaccharides) derived from various plant carbohydrates, i.e., corn, sugarcane, barley, sugar beet, potato, sweet sorghum, wheat, etc. The global trend of bioethanol production has been continuously on the rise during the last three decades with 4.0 billion gallons in 1990, 4.5 billion gallons in 2000, and 23.3 billion gallons in 2010, while it is estimated that the production volume will reach 40 billion gallons in 2020, corresponding to approximately 75% of the biofuel produced in the world (Zabed et al. 2017; Branco et al. 2019). First-generation bioethanol has caused a competition between the food industry and biofuel industry over arable land, irrigation water, and other raw materials, resulting in increases in food and feed prices (Salehi Jouzani and Taherzadeh 2015; Manochio et al. 2017; Dutta et al. 2014). Production of bioethanol from nonedible plants and from globally available lignocellulosic residues (second-generation bioethanol) has been proposed to minimize the negative impacts of first-generation biofuels. The former, such as miscanthus (Miscanthus spp., C4), switchgrass (Panicum virgatum, C4), reed canary grass (Phalaris arundinacea, C3), giant reed (Arundo donax, C3), shrub willow, and hybrid poplar, can be cultivated on marginal lands which are not suitable for crop production (Zabed et al. 2017). Lignocellulosic residues include forest slashes, crop residues, wood chips, yard trimmings, food processing waste, organic part of municipal solid waste, etc. The lignocellulosic parts of plants contain cellulose, hemicellulose, and lignin (Aghbashlo et  al. 2019a). Cellulose and hemicellulose are polysaccharides and could be converted into their building blocks, i.e., simple sugars, through hydrolysis. These sugars can be readily used by yeasts for bioethanol production. Lignin is another complex material present in the lignocellulosic biomass, serving as a cementing substance in the whole structure. Lignin is in fact the main reason to the recalcitrant nature of lignocellulosic biomass reducing the efficiency of the saccharification process. This necessitates the implementation of a pretreatment stage (physical, chemical, biological, etc.) to enhance the accessibility of c­ ellulose/hemicellulose to hydrolyzing enzymes during the saccharification stage (Binod et  al.

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2019). Accordingly, a huge number of research works has been carried out to develop more efficient pretreatment methods for enhancing the production of simple sugars from lignocellulosic materials (Mood et al. 2013). Commonly, first-generation bioethanol production from food crops includes milling, liquefaction and cooking, saccharification by hydrolyzing enzymes (glucoamylase and α-amylase), fermentation of simple sugars at 32  °C for 50  h by S. cerevisiae, distillation, drying, and denaturing. Through this process, 2.5–2.9 gallon of bioethanol is produced from 25  kg corn grains (Mosier and Ileleji 2015). There are additional processes involved in second-generation bioethanol production, in particular biomass collection and pretreatment, leading to increased production costs. Among these, the major bottleneck is the cost of pretreatment and saccharification for conversion of cellulose into simple fermentable sugars. Therefore, achieving cost-effective pretreatment and saccharification technologies for efficient production of simple sugars (hexoses and pentoses) from lignocellulosic biomass is of importance. During the last two decades, different feedstock pretreatment technologies, such as acidic, alkali, thermal, enzymatic, and integrated hydrolysis techniques, have been tested and developed (Salehi Jouzani and Taherzadeh 2015; Mood et al. 2013). However, the most widely used methodology for lignocellulosic materials pretreatment is acidic pretreatment. In this process, commonly, a dilute acid solution (e.g., 1–10% sulfuric acid) at high temperatures (e.g., 237 °C) and high pressures (e.g., 13 atm) is used for a short time (up to 15 s) to facilitate the enzymatic hydrolysis and sugar release (Badger 2002). To facilitate hemicellulose degradation and lignin solubilization, it is also possible to use different physical treatments, such as steam explosion, freezing, or radiation (Guo et  al. 2015). Different genera of fungi, such as Trichoderma, Aspergillus, Myceliophthora, Fusatium, Tribulus, Talaromyces, Chaetomium, and Thermoascus, have also been studied for exploring their efficient cellulases, which could efficiently be used for pretreatment purposes (Salehi Jouzani and Taherzadeh 2015; Kumar et al. 2009). Despite the development of different pretreatment and hydrolysis technologies for bioethanol production, these technologies are not cost-­ effective yet. The first commercial-scale cellulosic ethanol plant was constructed and operated in 2013 in Italy (The Crescentino bio-refinery). This company uses wheat straw as substrate and has the production capacity of more than 20 million gallons of bioethanol. Recently, some other cellulosic ethanol plants have been constructed or are under construction in the USA and other countries (Guo et al. 2015). –– Biodiesel The mono-alkyl esters of vegetable oils and animal facts as well as algal and microbial oils are referred to as biodiesel (Tabatabaei et al. 2019a). This biofuel, known as an attractive alternative to conventional petro-diesel in the transportation sector, is typically produced through the “transesterification” process (Aghbashlo et al. 2017a). Transesterification is the production of one ester from another ester, in which a light alcohol, mostly methanol, replaces the heavy alcohol found in the oil/ lipid structure, i.e., glycerol, and this reaction is mostly catalyzed by an alkaline catalyst, e.g., NaOH or KOH (Aghbashlo et al. 2017b; Salehi Jouzani et al. 2018;

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Sharafi and Jouzani 2019). The global annual biodiesel production stands at more than 35 million tons. Biodiesel production in the European Union and the USA in 2018 was recorded at about 13–7 million tones, respectively (Guo et al. 2015). The quality of biodiesel is commonly defined by its fatty acid alkyl ester composition, which is in turn defined by the oil feedstock used. In better words, biodiesel properties vary depending on the used oil feedstock (Hosseinpour et  al. 2016). Nevertheless, typically the average specific gravity of biodiesel is about 0.88, its kinematic viscosity ranges from 3.8 to 4.8 mm.s−2, and its cetane number ranges from 50 to 62. In addition, biodiesel energy density is between 38 and 45 MJ.kg−1 (Hoekman et al. 2012). Since the physicochemical properties of biodiesel are very similar to those of petro-diesel (Aghbashlo et al. 2019b), various blends of biodiesel and petro-diesel could be used in diesel engines (up to 20%, i.e., B20, without any modifications). It should be noted that biodiesel has a cleaner emission profile, higher cetane number, flash point, and lubricity, and its production and use are simple and easier compared to petro-diesel. In addition, biodiesel does not contain sulfur and aromatics (Guo et al. 2015; Lee and Lavoie 2013; Tabatabaei et al. 2011). Biodiesel is commonly produced from edible oils, i.e., soybean, canola, rapeseed, palm, safflower, almond, barley, coconut, cotton seed, etc. (Hegde et al. 2015; Taher and Al-Zuhair 2017). However, the application of these crops as source of biodiesel production has contributed to increased food price and has negatively impacted global food security, especially in the third world and some developing countries suffering from vulnerable economy and agriculture. This production scenario is also not quite economically feasible due to the high cost of these oil feedstocks. Thus, other sources, such as nonedible oils, waste animal fats, and used vegetable oils, have been proposed for biodiesel production. The most important nonedible oil plants with high potentials as feedstock for biodiesel production include physic nut (Jatropha curcas), camelina (Camelina sativa), castor beans (Ricinus communis L.), neem (Azadirachta indica A.), karanja (Pongamia pinnata L.), mahua (Madhuca spp.), simarouba (Simarouba glauca DC.), and cheura (Diploknema butyracea). However, the yield efficiency of such oil producing plants is too low to meet the global bioenergy demands. In addition, the application of such plant oils is faced with some disadvantages, such as the presence of trace salts, water, and high free fatty acid contents. In light of that, efforts have been put into their genetic improvements to enhance oil yield and quality (Sharafi and Jouzani 2019; Alaba et al. 2016). –– Pyrolysis bio-oil Pyrolysis is the process of fast or slow heating of lignocellulosic materials at 300–900  °C in the absence of air. As a result of such processes, typically, three products, including biochar, bio-oil, and syngas, are produced (Vamvuka 2011). Typically, slow pyrolysis by heating biomass at 300–600 °C for a number of days, with the absence of air, primarily results in the production of biochar, residual tar, and syngas. Fast pyrolysis (fast increasing of the temperature up to 500 °C within 2  s) converts biomass to mainly bio-oil and other chemicals (Kabir and Hameed 2017; Laird et al. 2009; Carpenter et al. 2014). Through the fast pyrolysis of plant

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biomass, about 10–30% biochar, 50–70% bio-oil, and 15–20% syngas are obtained (Laird et al. 2009). This method is also cost effective for bio-oil production from lignocellulosic materials. Pyrolysis bio-oil contains more than 300 different compounds, such as acids, alcohols, ketones, phenols, sugars, aldehydes, guaiacol, syringol, furans, alkenes, esters, aromatics, sugars, etc. However, the main components of bio-oil are typically phenolic and alkylated poly-phenols, along with relatively small amounts of phenol, eugenol, cresols, and xylenols (Aho et al. 2011). All lignocellulosic materials, such as wood chips, straw, bagasse, and hulls, can be used for bio-oil production. The lignocellulosic materials with low contents of nitrogen and ash are known as the most efficient for this purpose. Wood is the most preferred biomass used for production of qualified bio-oil (Vamvuka 2011). Crude pyrolysis bio-oil may be straightly used in industrial scale combustion systems; nevertheless, significant improvements are needed before crude bio-oil could be used as a petrol distillate fuel alternative. This includes reduction of moisture and acidity as well as improvement of heating value and storage stability. During the last 20 years, a wide range of improvement and upgrading technologies, such as esterification, supercritical fluidization, hydrogenation, catalytic cracking, steam reforming, hydrodeoxygenation, emulsification, and molecular distillation, have been developed (Guo et al. 2015; Kabir and Hameed 2017; Zhang et al. 2013; Ruddy et al. 2014). –– Drop-in biofuels There are two major concerns about bioethanol and biodiesel. One is that the energy density of bioethanol and biodiesel is about 67% and 90% of those of their petroleum-oriented counterparts, i.e., gasoline and petro-diesel, respectively. Second concern is related to the higher dissolution capability of these types of biofuels compared with their petroleum-oriented counterparts, therefore causing corrosion to engines, storage tanks, and distribution facilities at high blending rates (Bohre et al. 2015). Drop-in biofuels are those liquid biofuels derived from biomass with similar fuel specifications to petroleum-oriented fuels, which are ready to “drop in” to the existing fuel systems without requiring any changes in the infrastructure (Guo et al. 2015; Karatzos et al. 2014). Different biofuels, such as syngas complexes, sugar hydrocarbons, liquefied biomass, and butanol, have been proposed as drop-in biofuels. Production and application of drop-in biofuels are not cost-efficient yet, and more highly focused studies are needed to develop more efficient technologies for production of these biofuels.

1.2.3  Gaseous Biofuels –– Biogas Natural gas, as a gaseous fossil fuel, has formed from buried organisms under high levels of heat and pressure during thousands of years. This fuel contains about 95% methane and ethane. It also contains a partial share of propane, butane,

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nitrogen, and carbon dioxide. While natural gas as a fossil fuel is associated with environmental pollution, its renewable alternative, biogas generated through the anaerobic digestion (AD) process, has attracted a great deal of attention during the last decades (Aghbashlo et al. 2019c). In addition to economic advantages of biogas generation, there are also other environmental benefits associated with AD, such as efficient waste management and mitigation of water, soil, and air sources of pollution. AD helps reduce unfavorable odors such as that of manure while it is an efficient process to eliminate plant, animal, and human pathogenic populations. In addition, the digestate produced during the biogas production process can be directly used as fertilizer and could therefore significantly contribute to decreased application of synthetic fertilizers (Barati et  al. 2017). This could not only improve the economy of the agricultural practices but also could improve their eco-benignity by reducing nutrient runoffs and avoiding methane emissions (Thompson et al. 2013). Biogas could be produced through the AD of different organic wastes including livestock manure, food processing waste, lignocellulosic materials (straw, branches, bagasse, etc.), municipal solid waste, yard trimmings, sewage sludge, etc. Based on the type of organic waste and the technology of AD, biogas traditionally consists of 60–65% of methane, 30–35% of carbon dioxide, and a partial share of water vapor, hydrogen, and hydrogen sulfide (Aghbashlo et al. 2019d). However, before commercial use of raw biogas, different impurities, such as hydrogen, hydrogen sulfide, and water vapor, should be removed (Weiland 2010). The process of anaerobic digestion for biogas production includes four steps (Tabatabaei et al. 2019b): The first step is hydrolysis, in which some anaerobic bacteria start hydrolyzing large organic molecules (i.e., complex carbohydrates, proteins, and lipids) into their building blocks or monomers (i.e., simple sugars, amino acids, and fatty acids). In the second step, acidogenesis occurs in which acidogenic bacteria transform simple sugars, amino acids, and fatty acids into CO2, H2, NH3, and organic acids. The third step is acetogenesis in which organic acids are converted into acetic acid, NH3, CO2 +, and H2 through the function of acetogenic bacteria. Finally, during the fourth step, i.e., methanogenesis, methane is generated by methanogenic archaea. Commonly, one-ton dry weight of organic waste yields about 120 cubic meters of biogas, which can produce about 200 kWh of electricity (Guo et al. 2015). The C/N ratio, total solids, temperature, microbial flora in the process, and the technology used for the process are the most important factors influencing on the AD and biogas production efficiency (Tabatabaei et al. 2020). –– Syngas Syngas, a gaseous biofuel, is a mixture of gases dominated by carbon monoxide (30–60%), hydrogen (25–30%), and carbon dioxide (5–15%). Other materials present in syngas are CH4, H2S, COS, NH3, and water vapor. Typically, gasification or pyrolysis of lignocellulosic materials or other renewable materials leads to syngas production. The term “bio-syngas” is usually used for syngas produced from renewable sources (Abatzoglou and Fauteux-Lefebvre 2016). For production of bio-­ syngas, it is necessary to heat dry plant biomass at high temperatures (e.g., 1200 °C) within a gasifier in the presence of controlled airflow. In such combustion chambers,

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three phases, including dehydration (drying at temperatures up to 200 °C), pyrolysis (by increasing temperature, biomass is transformed to char and vapor), and partial oxidation (production of CO, CO2, and H2O), occur. Finally, through the reaction of CO with H2O, CO2 and H2 are produced. The carbon conversion efficiency of this process for wood is about 92% (Guo et al. 2015; Abdoulmoumine et al. 2015). The energy density of syngas is about 5.3 MJ.Nm−3. Syngas can be directly burned for electricity generation. It could also be purified and applied for production of transportation fuels, such as methanol, ethanol, methane, and dimethyl ether. Syngas production from biomass could be costly owing to the presence of moisture in the biomass necessitating drying and purification of crude syngas. Therefore, more comprehensive and in-depth research and development are needed to develop more efficient and cost-effective feedstock preparation and bio-syngas production protocols. Currently, hundreds of gasification plants have been constructed through the world.

1.2.4  Challenges in Biofuel Production As mentioned earlier, biofuels, in particular second-generation biofuels, have been regarded as one of the most promising options to mitigate GHGs and combat climate change. However, due to the limitations discussed above and the consequent poor cost competitiveness [higher production cost of lignocellulosic biofuels vs. fossil fuels or first-generation biofuels (Rastogi et al. 2018)], the share of conventional biomass-based biofuels in the transportation sector is only about 5% (Mood et al. 2013). The poor economy of lignocellulosic biofuels is attributed to the complexity and crystalline structure of cellulose and the presence of lignin, requiring additional steps (Mood et al. 2013; Kumagai et al. 2014). More specifically, the pretreatment and enzymatic hydrolysis are known as the costliest processes during bioalcohol production from lignocellulosic biomass. Implementation of biorefineries aimed at generating a wide range of products could be regarded as a strategy to address the abovementioned challenges and enhance the economical features of the whole lignocellulosic biofuel production process (Khounani et  al. 2019). Fungi, owing to their wide range of lignocellulolytic enzymes, could be an important part of these lignocelluloses-based biorefineries.

1.3  Fungi Biorefineries Fungi with a huge diversity and a diverse biochemical processes present in them have been widely used in the biotechnology as biocatalyst. Commonly, fungal biotechnology is often referred to modern biotechnological processes involving fungi and yeasts. In spite of that, only 5% of the fungi are cultivable, which have been

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used to produce a diverse range of enzymes, organic acids, pigments, sweeteners, biopesticides, bio-herbicides, biofertilizers, vitamins, antibiotics, and other pharmaceutical products. During the last decades, more than 9000 different bioactive compounds have been produced by fungi (Brakhage and Schroeckh 2011). By the development of new omics technologies, such as genomics, transcriptomics, proteomics, metabolomics, metagenomics, next-generation sequencing technologies, genetic engineering, etc., more new biological compounds have been detected and purified from filamentous fungi. Typically, fungal enzyme genes are present in clusters which code multi-domain and multi-modular enzymes, such as polyketide synthases, prenyltransferases, non-ribosomal peptide synthases, and terpene cyclases (www.scribd.com).

1.3.1  Promises of Fungi in Biofuel Production Fungi have shown tremendous potentials for the production of biofuels from a variety of crop plants and other lignocellulosic agricultural wastes. Different studies have confirmed that a diverse range of fungi could be used for biofuel production or to be used as a part of biofuel production process. A wide range of fungal species have been evaluated for their capability and efficiency in the production of different biofuels, such as biodiesel, biohydrogen, and cellulosic ethanol. Different protocols have been developed for production of second-generation (degradation of lignocellulosic biomass) and third-generation (straight production of biofuels by fungi) biofuels using filamentous fungi (Raven et  al. 2019). Recently, Salehi Jouzani and Taherzadeh (2015) reviewed the possible applications of different fungi, such as Trichoderma, Aspergillus, and Fusarium species through the consolidated bioprospecting (CBP) for biofuel production. Bio-delignification of lignocelluloses has been carried out by ascomycetes including T. reesei, basidiomycetes such as the white rot fungi, Trametes, Pleurotus ostreatus, and Phanerochaete chrysosporium (Chandel et  al. 2015). Exo-β-1,4-­ glucanases and endo-β-1,4-glucanases, which are capable of cellulose degradation, are commonly produced by different T. reesei strains in large concentrations. More than 200 glycoside hydrolase genes controlling the production of at least 10 different lignocellulose-hydrolyzing enzymes have been detected and characterized in this fungus. In addition, thermostable cellulases with high cellulose-degrading activity at high temperatures have been identified from several fungal species, such as Myceliophthora thermophile, Talaromyces emersonii, Tribulus terrestris, Chaetomium thermophilum, and Thermoascus aurantiacus (Guo et al. 2015). Filamentous fungi are capable of effective production of different types of cellulases (about 100  g/ L). Currently, some big companies, including Novozymes, Genencor, and Iogen Corporation, commercially use Trichoderma for cellulase production, while some others, such as Dyadic International, Inc., use Chrysosporium lucknowense. Production of 100 g/L is much higher than what obtained in bacteria.

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Recently, through the application of genetic engineering technologies, the beta-­ glucosidase gene of Periconia sp. was expressed in T. reesei, which could enhance the level of beta-glucosidase produced and improve the overall cellulase activity on biomass residues (Dashtban and Qin 2012). Therefore, the widest application of filamentous fungi for biofuels production is their application in the processes of pretreatment and enzymatic hydrolysis of lignocellulosic materials to produce sugar monomers. Another application of fungi for biofuels production is their application in biodiesel production. Fungi have been used for biodiesel production in two different ways. The first one concerns fungi with the capability to produce high concentrations of lipids. The fungal lipids can be directly used for biodiesel production. The second way concerns the application of fungi as biocatalyst for the transesterification process. Application of lipases as catalyst is more favorable than chemical acidic or basic catalysts, as it does not pose any environmental and health risks. In addition, lipases derived from fungi have a wide substrate specificity, are highly stable in different organic solvents, and do not need cofactors in the process. Yields of enzymatic transesterification can reach to 100%. Genetic engineering has been used to convert S. cerevisiae into a biodiesel producer, i.e., one that is oleaginous, producing fatty acids and alcohols, and could be directly converted into biodiesel (Sanchez and Demain 2017).

1.4  Conclusions Increasing global demand for fuels and the consequent increases in environmental pollution and human health risks have collectively driven research toward finding sustainable and economically viable alternatives. The second- and third-generation biofuels have been considered as promising strategies for meeting this goal. However, as these kinds of biofuels are produced from lignocellulosic materials or waste oils, their production is not commonly cost-effective due to additional stages required. In light of that, on one hand, researchers have been striving to develop more cost-effective pretreatment and hydrolyzing technologies, while, on the other hand, they have been focused on more sophisticated production pathways, i.e., biorefineries. Fungi owing to their huge potentials for biofuels production have been at the center of attention throughout the implementation of these strategies. More specifically, fungi could be used as the source of cellulase for hydrolyzing lignocellulosic materials in the bioethanol production process. Moreover, they could be used as source for lipid production for biodiesel production. Finally, fungi could also serve as the source of lipase for catalyzing transesterification of lipids into biodiesel. In spite of all these attractive features, however, more research and development will be required in the future to further explore the applications of fungi for second- and third-generation biofuel production.

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Chapter 2

Bioethanol Production by Using Plant-­Pathogenic Fungi Amin Alidadi, Hamed Kazemi Shariat Panahi, Mona Dehhaghi, Reeta Rani Singhania, Hossein Ghanavati, Reza Sharafi, Mortaza Aghbashlo, Meisam Tabatabaei, and Gholamreza Salehi Jouzani

2.1  Introduction Currently, one of the greatest challenges facing the modern world is the secure provision of transportation fuels, preferably by renewable energy. Some of the negative environmental and health impacts of fossil-derived fuels application  could be neglected through biofuel consumption (Dehhaghi et  al. 2019b; Kazemi Shariat Panahi et al. 2019b; Tabatabaei et al. 2019a). Conventional biofuels (e.g., bioethanol, biobutanol, biocrude oil, and biodiesel) and unconventional ones (such as glycerol and bioammonia) have extensively been studied for climate change and greenhouse gas mitigation, environment protection, and health risk reduction that

A. Alidadi · H. Ghanavati · R. Sharafi · G. Salehi Jouzani (*) Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran e-mail: [email protected] H. Kazemi Shariat Panahi · M. Dehhaghi Department of Microbial Biotechnology, School of Biology and Centre of Excellence in Phylogeny of Living Organisms, College of Science, University of Tehran, Tehran, Iran Faculty of Medicine and Health Sciences, Macquarie University, Sydney, NSW, Australia Biofuel Research Team (BRTeam), Karaj, Iran R. R. Singhania Centre for Energy and Environmental Sustainability, Lucknow, India M. Aghbashlo (*) Department of Mechanical Engineering of Agricultural Machinery, Faculty of Agricultural Engineering and Technology, College of Agriculture and Natural Resources, University of Tehran, Karaj, Iran e-mail: [email protected]

© Springer Nature Switzerland AG 2020 G. Salehi Jouzani et al. (eds.), Fungi in Fuel Biotechnology, Fungal Biology, https://doi.org/10.1007/978-3-030-44488-4_2

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are associated with liquid energy production and consumption (Dehhaghi et  al. 2019b; Kazemi Shariat Panahi et al. 2019c, d; Rahimzadeh et al. 2018; Shirzad et al. 2019; Tabatabaei et al. 2019a). Among them, fuel bioethanol accounts for more than 70% of the global biofuel production, of which 80% comes from fermentation and the remaining comes from chemical synthesis (Kazemi Shariat Panahi et al. 2019c). In 2018, ~108.14 billion L of this biofuel was produced (mainly through ethanolic fermentation of corn and sugarcane in Brazil and the United States, respectively) (Kazemi Shariat Panahi et al. 2019c). On this context, bioethanol could be used as a gasoline or fuel extender, which usually blends in a ratio of 5–10% v/v with gasoline but the ratio could be up to 85% v/v for flexible fuel vehicles (Kazemi Shariat Panahi et al. 2019c). Despite the high versatility, one of the main constraints of the bioethanol production is food vs. fuel concern (Kazemi Shariat Panahi et al. 2019a, b). To address this issue, plant biomass (i.e., lignocelluloses) could be exploited for the production of second-generation biofuels (including bioethanol, biobutanol, and biogas) (Dehhaghi et al. 2019b; Kazemi Shariat Panahi et al. 2019c; Shirzad et al. 2019; Tabatabaei et al. 2019b, c). More specifically, lignocelluloses are the most abundant and biorenewable biomass on the earth which can be potentially used as substrates for microbial production of diverse metabolites, ranging from therapeutic compounds (Dehhaghi et  al. 2018, 2019c; Mohammadipanah et  al. 2016) to enzymes (Hamedi et al. 2015; Mohammadipanah et al. 2015) and biofuels (Dehhaghi et al. 2019a; Kazemi Shariat Panahi et al. 2019c). Lignocellulose generally consists of three main polymers, including cellulose, hemicelluloses, and lignin, and also minor concentrations of acetyl groups, minerals, and phenolic substituents (Kazemi Shariat Panahi et al. 2019a). The cellulosic portion composes of long linear chain of β-(1 → 4)-glucose monomers along with micro-fibril bundles (Kazemi Shariat Panahi et al. 2019b), whereas hemicellulose is made up of xyloglucans or xylans linked to micro-fibrils through hydrogen bonds. On the other hand, lignin is a phenolic compound that consists of p-coumaryl, coniferyl, and sinapyl alcohol monomers and confers mechanical strength to the cell wall (Kang et al. 2014). The bioethanol production from lignocelluloses involves four steps, including (1) biomass delignification/pretreatment, (2) enzymatic saccharification or chemical hydrolysis, (3) ethanolic fermentation, and (4) distillation (Kazemi Shariat Panahi 2019a) (Fig. 2.1).

M. Tabatabaei (*) Institute of Tropical Aquaculture and Fisheries (AKUATROP), Universiti Malaysia Terengganu, Terengganu, Malaysia Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran e-mail: [email protected]

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Fig. 2.1  Major steps in bioethanol production

The pretreatment step involves the application of biological, physical, or chemical method (or their combination thereof) to degrade the lignin complex structure and to decrease cellulose crystallinity of lignocelluloses for facilitation of the subsequent saccharification and fermentation steps. A detailed explanation of different pretreatment techniques for preparation and delignification of various types of biomass as well as their subsequent saccharification methods has been provided in our previous report (Kazemi Shariat Panahi et  al. 2019a). The enzymatic hydrolysis, also known as saccharification, is the application of different microbial hydrolases to release monosaccharides (i.e., fermentable sugars) from cellulose and hemicellulose polysaccharides (Kazemi Shariat Panahi et al. 2019b). Prior to distillation, the fermentable sugars coming from saccharification are converted into ethanol by suitable ethanologens (Kazemi Shariat Panahi et al. 2019b,c). There are different ways to integrate hydrolysis and fermentation bioprocesses, including separate hydrolysis and fermentation, simultaneous saccharification and fermentation, and consolidated bioprocessing (Kazemi Shariat Panahi et al. 2019b) (Fig. 2.2). Enzymes used in saccharification step can significantly be produced by the lignocellulose-­degrading microbes, especially different filamentous fungi (Kazemi Shariat Panahi et al. 2019a, b; Tabatabaei et al. 2019b). Among them, many phytopathogenic fungi could overcome plant cell wall barrier through the secretion of copious amounts of hydrolytic enzymes. More specifically, these plant pathogens could be categorized into three groups according to their feeding strategies: (1) biotrophic, (2) nectrotrophic, and (3) hemibiotrophic (i.e., intermediate between the two previously mentioned groups). Typically, necrotrophic pathogens produce overwhelming amounts of hydrolytic enzymes that can degrade wide range of hosts, compared to biotrophics (Meinhardt et al. 2014; Oliver and Ipcho 2004). Therefore, plant pathogenic fungi can potentially be used in biofuel production, improving conversion efficiency of cell wall polysaccharides into glucose and other

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Fig. 2.2  The major steps of bioethanol production from lignocelluloses

fermentable sugars. In this chapter, the promising applications of plant pathogenic fungi in bioethanol industry in respect to hydrolytic enzyme production have comprehensively been discussed.

2.2  P  hytopathogenic Fungi’s Potential in Bioethanol Industry Plant pathogenic fungi are one of the most important groups of fungi that, together with fungal-like organisms, are responsible for ~85% of plant diseases. More specifically, over 20,000 fungi species are plant pathogens, causing more than 10,000 different plant diseases. The pathogenesis involves the hydrolytic enzyme secretion that degrades the plant cell wall into carbohydrates (i.e., energy) for fungal growth and reproduction. In other hand, various plants (or plants residues) could also be used as substrates for the production of different biofuel types (especially biogas, ethanol, and butanol) (Kazemi Shariat Panahi et al. 2019b, 2019c; Tabatabaei et al. 2019a). In better word, pathogenic fungi or their enzymes could be utilized in delignification process (via enzymatic or biodelignification method) and/or subsequent saccharification step, prior to anaerobic digestion (i.e., biogas production) or fermentation (i.e., butanol and ethanol production) step (Kazemi Shariat Panahi et al. 2019b, c; Tabatabaei et al. 2019c). Alternatively, these fast-growing microorganisms could ferment mixed pentose-­ hexose sugars in saccharified lignocellulosic biomass into ethanol, a fermentation process commonly referred to as simultaneous saccharification and co-fermentation (Christakopoulos et al. 1997; Kazemi Shariat Panahi et al. 2019a; Tabatabaei et al. 2019a). For example, the shortcomings of wild Saccharomyces cerevisiae in pentose (e.g., xylose and arabinose) conversion into bioethanol could be addressed by some plant pathogenic filamentous fungi (Demeke et  al. 2013; Kazemi Shariat Panahi et al. 2019a; La Grange et al. 2010). This section discusses some important plant pathogenic fungal genera as well as species that have been investigated for their application in bioethanol production process (Fig. 2.3).

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Fig. 2.3  Some plant pathogenic fungi genera that could be used in bioethanol production industry

2.3  Genus Fusarium Fusarium is a large cosmopolitan genus and widely distributed genus of filamentous fungi. It belongs to family Nectriaceae, order Hypocreales, class Sordariomycetes (subclass Hypocreomycetidae), phylum Ascomycota (subdivision Pezizomycotina), and kingdom Fungi (subkingdom Dikarya). With few exceptions (see Chap. 4), Fusarium spp. are commonly known as ubiquitous soil-borne fungal plant pathogens due to their devastating effects on different agricultural crops, for example, Fusarium wilt of banana. According to the American Phytopathological Society’s plant disease list (www.apsnet.org), most of economically important plants have at least one disease associated with Fusarium spp. More specifically, Fusarium spp. are highly potent plant pathogens affecting different plants parts (e.g., root, stem, leaf, fruit, and seed) and causing serious diseases (e.g., rots, cankers, wilts, and spots) with wide range of symptoms (Leslie and Summerell 2008).

2.3.1  Fusarium verticillioides Fusarium verticillioides (F. verticillioides), also known as Gibberella moniliformis, is one of the most common pathogens in the world, particularly causing ear rot, stalk rot, and cob rot in maize (Covarelli et al. 2012; Duan et al. 2016). This pathogen can produce various phytotoxins and mycotoxins (i.e., fumonisins A, B, C, and P) which could further complicate the plant infection, following their accumulation in maize kernel, by posing human and livestock health at risk (e.g., cytotoxic and carcinogenic issues)  (Jimenez-Garcia et  al. 2018; Leyva-Madrigal et  al. 2017;

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Szécsi 2014). However, from ethanol industry viewpoint, F. verticillioides could be a potent cellulase-producing filamentous fungus. More specifically, the cellulases produced by this species, including ß-glycosidase, endoglucanase, cellobiohydrolase, endocellulase, and exocellulase, catalyze the conversion of cellulose into sugars. Some of these hydrolytic enzymes, together with F. verticillioides hemicellulase cocktails, have potentials for bioethanol production through lignocellulose saccharification. For example, a high hydrolysis efficiency could be expected from its endoglucanase and glycosidase multi-enzyme complex (de Almeida et al. 2011, 2013a). In another study (de Almeida et al. 2013a), a free endoglucanase and glycosidase multi-enzyme complex consisting of one xylanase, one cellobiohydrolase, and two endoglucanases were characterized from F. verticillioides. The characterized enzymes were highly thermophilic with the optimum activity at 80 °C. Saha (2001) partially purified a xylanase from F. verticillioides with capabilities to digest various xylan substrates into higher short-chain xylooligosaccharides and xylobiose. Moreover, the enzyme showed high temperature and pH stability ranges of up to 50 °C (optimum, 50 °C) and 4–9.5 (optimum, 5.5), while it produces no xylose and requires no metal ion for stability and activity. Recently, two enzyme cocktails (i.e., cellobiase-rich and endoglucanase-rich cocktails) were prepared from F. verticillioides cultivation on the medium containing Andropogon gayanus (Gamba grass) as the carbon source (de Almeida et  al. 2019). A blend of these two cocktails (cellobiase-­to-endoglucanase ratio, 1:12) at the concentration of 40 mg/g was used for sugarcane bagasse saccharification (solid content, 6  wt.%), converting 73.1% xylan and 43.4% glucan into sugars. The mentioned hydrolytic enzyme production capacities, therefore, could be exploited for bioethanol production either by consolidated bioprocessing or separate hydrolysis and fermentation process. The former approach, also known as direct microbial conversion, simultaneously involves cellulase production, pretreated lignocellulose hydrolysis, and ethanolic fermentation in a single vessel (Kazemi Shariat Panahi et al. 2019a). In addition to cellulase production, F. verticillioides could co-ferment glucose and xylose at high yields. Thus, it can be used for direct ethanol production from pretreated lignocellulosic biomass in a single process via consolidated bioprocessing system. de Almeida et al. (2013b) used F. verticillioides for direct ethanol production (concentration, 4.6 g/L, and yield, 0.31 g/g) from delignified sugarcane bagasse (40 g/L) in a consolidated bioprocessing system. Interestingly, the fungus could ferment glucose, xylose, and their mixture (i.e., mixed sugar co-fermentation) into ethanol with yields of 0.47, 0.46, and 0.50 g/g, respectively. According to their results, F. verticillioides could be a suitable candidate for the direct ethanol production from lignocelluloses via consolidated bioprocessing system (de Almeida et al. 2013b; Salehi Jouzani and Taherzadeh 2015). As already mentioned, bioethanol could also be produced through separate hydrolysis and fermentation process, in which the delignified lignocellulose enzymatically is hydrolyzed into reducing sugars prior to ethanolic fermentation (Kazemi Shariat Panahi et al. 2019a). This approach takes the advantage of optimum enzyme activities through flexibility in process temperature (i.e., due to the absence of microorganisms) (Kazemi Shariat Panahi et  al. 2019d). Trichoderma reesei

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(T. reesei) is the most common microorganism for the production of commercial cellulase (i.e., endoglucanases, cellobiohydrolases, and ß-glucosidases) (see Chap. 4). However, wild T. reesei cannot produce some enzymes such as hemicellulase and pectinase, compared to other filamentous fungi (Martinez et al. 2008), which are crucial for biodegradation of lignocellulose and the subsequent monosaccharide liberation. This drawback could be addressed by enriching the T. reesei enzyme cocktails by hemicellulase and pectinase produced by other microorganisms such as F. verticillioides. On this basis, Ravalason et al. (2012) supplemented T. reesei commercial cellulases with F. verticillioides enzymes. The resulting enzyme cocktail enhanced wheat straw hydrolysis and the subsequent release of glucose (24%), xylose (88%), and arabinose (68%) sugars, compared to the un-supplemented control. Some studies have already sequenced the genome of F. verticillioides which could be used in performance improvements (in respect to hydrolytic enzyme and ethanol productions), mycotoxin-producing gene knockout, etc., via recombinant technology (de Almeida et al. 2019; Gardiner 2018).

2.3.2  Fusarium graminearum Fusarium graminearum (F. graminearum) is another pathogenic Fusarium sp., i.e., a major cereal pathogen, all over the world. This pathogen causes head blight disease on various commercial plants, particularly cereal crops, such as wheat (Triticum aestivum), barley (Hordeum vulgare), oat (Avena sativa), and also ear/stalk rot of corn (Zea mays) (McMullen et al. 1997; Sutton 1982). In addition, F. graminearum could affect some oil crops, especially soybean, leading to damping-off, and crown and root rots (Martinelli et al. 2004; Pioli et al. 2004). The F. graminearum virulence factors are a number of hydrolytic enzymes with cell wall-degrading ability. In respect to phytopathogenesis, the most important of these enzymes are pectinases, arabinoxylanases, and endo-polygalacturonases (Kikot et al. 2009; Paccanaro et al. 2017). It is worth mentioning that xylan is one of the most abundant hemicellulose polymers in the plants. Three important enzymes, i.e., endo-1,4-ß-xylanase, 1,4-ß-­xylosidase, and α-L-arabinofuranosidase are associated with hemicellulose hydrolysis. Therefore, F. graminearum’s ability in xylanase production could be exploited in bioethanol industry. In addition to hemicellulases and pectinases, this fungal species could secrete various cellulases for digestion of many oligo- and polysaccharide substrates (Kikot et  al. 2009; Phalip et  al. 2005). Thus, F. graminearum can improve lignocellulose saccharification by the production of important hydrolytic enzymes, especially hemicellulases, which is a crucial step for bioethanol production from lignocelluloses. It is also possible for transferring these genes into a potent microbial host, for example, xylanase genes (XylD, XyloA, and Xylo/ArabA) have been expressed in Escherichia coli for a production of high amounts (10–38 mg/L) of endo-1,4-ß-xylanase, 1,4-ß-xylosidase, and bifunctional xylosidase/arabinofuranosidase (Carapito et  al. 2009). Accordingly, the produced

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enzymes could efficiently be used for hydrolyzing hemicellulose into reducing sugars during bioethanol production via separate hydrolysis and fermentation process.

2.3.3  Fusarium oxysporum The Fusarium oxysporum species complex (FOSC) comprises both plant pathogenic and nonpathogenic Fusarium oxysporum (F. oxysporum) strains. More specifically, a total of 143 formae speciales and races (106 well-documented ff. ssp. and 37 insufficiently documented ff. ssp.) of F. oxysporum have been described that mostly are worldwide pathogens on 45 plant families (more predominantly on Asteraceae, Cucurbitaceae, Fabaceae, and Solanaceae), including various economically important plants (Edel-Hermann and Lecomte 2019; Leslie and Summerell 2008). For example, F. oxysporum ff. ssp. radicis-lupini, radicis-cucumerinum, and radicis-lycopersici are the causative agents of lupine, cucumber, and tomato root rotting. It should be noted that when the term radicis has been mentioned in the formae speciales, it always means the strain can develop root rotting symptoms. However, the term is not present in all formae speciales such as F. oxysporum ff. ssp. opuntarium, lilii, and cepae. Moreover, there are 58 additional host plants with still no characterized formae speciales (Edel-Hermann and Lecomte 2019). Generally, Fusarium disease symptoms are vascular wilting and/or, to a lesser extent, rotting. Briefly, the common pathogenesis involves the fungal penetration into host plant roots. Upon reaching the xylem vessels, this fungal species colonizes upward causing progressive plant yellowing and wilting (Edel-Hermann and Lecomte 2019). The plant diseases associated by the mentioned symptoms and pathogenesis are Fusarium wilt, Fusarium blight, and Fusarium yellows. However, F. oxysporum can cause Fusarium stem rot, or crown and root rot, or basal rot through developing black to brown necrotic spots and the subsequent progressive infection of roots and hypocotyl cortical tissues without colonizing the vascular system. Intriguingly, short-stem plants (e.g., ginger, potato, and lily) with storage organs (e.g., rhizomes, tubers, and bulbs) are more susceptible to rot diseases, possibly due to the fungus adaptability to the plant anatomical peculiarity (Edel-­ Hermann and Lecomte 2019). The hydrolytic enzyme production capabilities of plant pathogenic F. oxysporum could be exploited for bioethanol production in the same manner as non-pathogenic ones (See Chap. 4). Briefly, simultaneous saccharification and fermentation, simultaneous saccharification and co-fermentation, and consolidated bioprocessing may be used for ethanol production by F. oxysporum. Accordingly, F. oxysporum is a filamentous fungus that is able to generate many cellulase and hemicellulase enzymes to hydrolyze the wide range of delignified lignocellulosic biomass under submerged or solid-state conditions (Xiros et al. 2008, 2009). Moreover, it can ferment glucose, xylose, cellulose, and hemicellulose into bioethanol (Panagiotou et al. 2005). Thus, this species is one of the unique filamentous fungi that can be used for bioethanol

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production from various delignified lignocellulosic biomasses via consolidated bioprocessing system. Interestingly, the ability of F. oxysporum for pentose (e.g., xylose) fermentation into bioethanol could be exploited under simultaneous saccharification and co-­ fermentation process. Through this strategy, which is a derivative of simultaneous saccharification and fermentation process, mixed sugars (pentoses and hexoses) are fermented into bioethanol at the same time (Kazemi Shariat Panahi et al. 2019a). Therefore, F. oxysporum could be used along with other potent ethanologens such as wild S. cerevisiae which could ferment hexoses (e.g., glucose and mannose) to bioethanol but is unable to ferment xylose into bioethanol. This co-cultivation could be translated into higher bioethanol yield through xylose conversion (See Chap. 4). Alternatively, high-solid bioethanol production process could be used to reduce the bioethanol production costs while at the same time minimizing water consumption. The lower water input requires lower energy for distillation, thus improving the economic feasibility of the process (Kawa-Rygielska and Pietrzak 2014; Koppram et al. 2014). In contrast, poor mass transfer and high microbial inhibitor concentration (formed during the lignocellulose pretreatment) would be encountered during high-solid content fermentation (Kazemi Shariat Panahi et al. 2019a, d; Koppram et al. 2014). These problems could partially be overcome by the inoculation of resistant ethanologens such as F. oxysporum that harbor powerful enzymatic system for the conversion of delignified lignocellulose into bioethanol even in the presence of considerable amounts of inhibitory compounds (Paschos et al. 2015; Kazemi Shariat Panahi et al. 2019c; Xiros et al. 2011). In this context, it is worth mentioning that ionic liquid pretreatment is gaining more attention as a more economically feasible method for lignocellulose delignification, compared to other chemical pretreatments, such as acid treatment, alkali treatment, ozonolysis, etc. (Wang et  al. 2011). Other advantages of ionic liquid include raw material invariance, high monomeric sugar production yield, short saccharification process time, and efficient delignification activity (Klein-Marcuschamer et al. 2011). However, a main disadvantage of this method is its biological toxicity (ionic liquid concentration, 0.2–5% w/v), decreasing the growth of ethanologens during fermentation process (Ouellet et al. 2011; Ruegg et al. 2014). Interestingly, F. oxysporum enzymatic system shows good tolerance against ionic liquids as well as other inhibitory compounds generated during lignocellulose delignification process (Xiros et al. 2011). For example, F. oxysporum (strain BN) produced a cellulase that is capable of growing at high concentrations of 1-ethyl-3-methylimidazolium phosphinate (10 wt.% w/v). Overall, F. oxysporum could be a promising ethanologen for the conversion of delignified lignocellulose into bioethanol. However, this fungus suffers from several major disadvantages including low growth rate, low conversion rate, and large amounts of acetic-acid production, compared to yeasts (see Chap. 4). Therefore, genetic engineering techniques are required to construct a high efficient enzyme- or ethanol-producing F. oxysporum (see Chap. 4).

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2.4  Genus Aspergillus Aspergillus is another diverse filamentous fungal genus with ubiquitous occurrence. This genus belongs to family Aspergillaceae, order Eurotiales, class Eurotiomycetes (subclass Eurotiomycetidae), phylum Ascomycota (subdivision Pezizomycotina), and kingdom Fungi (subkingdom Dikarya). Aspergillus spp. have high adaptability to various habitats (e.g., soil, salterns, agroecosystems, polar, living plants, animals and lichens, stones, water-related, fossil records, and human) and climatic zones and can infect a wide range of plants. More specifically, many Aspergillus spp. are weak and opportunistic plant pathogen, commonly associated with post-harvest diseases. On the other hand, some Aspergillus spp. have commercial biotechnological exploitation for the production of different metabolites ranging from antibiotics to organic acids and enzymes (especially cellulolytic enzymes), e.g., citric acid production by Aspergillus niger (A. niger) (Samson et al. 2014).

2.4.1  Aspergillus niger A. niger prefers warm and humid places and is commonly present in mesophilic environment (e.g., decaying vegetables) but can grow anywhere at 6–47  °C and pH 1.4–9.8 with a growth water activity limit of as low as 0.88 (Palacios-Cabrera et  al. 2005; Schuster et  al. 2002). This species harbors a 35.5–38.5  Mb genome, assembled into four separate chromosomal bands (ranging 3.5–6.5 Mb) and composed of ~14,000 open reading frames (with potential ability for protein encoding) and ~13,000 genes (including 8000–8500 genes with functional assignments) (Debets et al. 1990; Gautam et al. 2011). This species is responsible for post-harvest damage of fresh and dried fruits (e.g., apple, citrus, grape, and fig), vegetables (e.g., tomato, breadfruit, onion, and garlic), and other crops (i.e., corn, barley, soybean, rapeseed, sorghum, and chickpea) (Pitt and Hocking 2009; Plascencia-Jatomea et al. 2014; Thamizharasi and Narasimham 1993; Vitale et al. 2008). Moreover, it is the causative agent of black molds on plant surfaces and is responsible for seed deterioration (Plascencia-Jatomea et al. 2014; Ziani et al. 2009). Compared to other filamentous fungi, A. niger is less destructive and has been labeled as generally recognized as safe by the US Food and Drug Administration. Unlike other Aspergillus spp., A. niger produces spores ranging from very dark brown to carbon black in color from biseriate phialides (Gautam et al. 2011). During the pathogenesis, a wide range of hydrolytic and oxidative enzymes are secreted by this species, degrading the lignocellulosic plant cell wall. Consequently, various diseases targeting different plant parts are resulted, including back rots of onion, crown rot of peanut, tuber rot of yam, stem rot of Dracaena, black mold rot of cherry, kernel rot

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of maize, fruit rots of grapes and banana, and rots of tomato and mango (Abbasi and Aliabadi 2008; Adebesin et al. 2009; Gautam et al. 2011; Palencia et al. 2010). In addition to citric acid, A. niger is commercially exploited in various fermentation techniques for production of useful enzymes such as pectinase, invertase, lactase, glucoamylase (also known as amyloglucosidase), cellulase, and amylase (Gautam et al. 2011; sheng Xue et al. 2018; Tiwari et al. 2015). A. niger SH3 can also produce a holocellulase cocktail (i.e., xylanase, 1951.6 IU/g; FPase, 31.5 IU/g; ß-glucosidase, 540.6  IU/g; and endoglucanase, 655.9  IU/g) for liberating up to 375 mg/g dried substrate (mg/gds, pretreated paddy straw) in an optimized saccharification process (15 FUP/gds, 40 °C) (Tiwari et al. 2015). Moreover, some A. niger spp. (such as strain van Tieghem) can also secrete thermostable ß-xylosidase (optimum activity, 70–75 °C and pH 4.5) (Boyce and Walsh 2018). The enzyme showed up to 91% activity, following 72  h incubation at 60–65  °C and could digest p-­ nitrophenyl α-L-arabinofuranoside, p-nitrophenyl β-D-glucopyranoside, and p-nitrophenyl-β-D-xylopyranoside. A 19-time increase in xylose amount was observed during the saccharification (70 °C, 6 h) of pretreated straw, following the supplementation of a commercial lignocellulosic enzyme cocktail by the purified A. niger thermostable ß-xylosidase (Boyce and Walsh 2018). Recently, an acidic, thermostable, and ethanol-tolerant endoglucanase was reported from A. niger that required up to 8% ethanol for maximum activity (60 °C, pH 5) (sheng Xue et al. 2018). Therefore, A. niger may be a good candidate for hydrolysis of delignified lignocellulose prior to bioethanol production process. On this context, Rocha et al. (2013) produced an enzyme cocktail (40 U/gds) by A. niger ATCC 16404 via solid state fermentation (40  °C, 500  mL rotating drum bioreactor) of rice by-product (86 wt.%), CaCl2 (2 wt.%), and whey (12 wt.%). The enzyme cocktail was used in simultaneous saccharification and fermentation of same substrate in the presence of S. cerevisiae Y904 as the ethanologen. With a 12-h pre-saccharification step, this fermentation setup delivered up to 11.7  g/L of bioethanol at 35  °C and pH  4.5. Compared to that of T. reesei, saccharification of the substrate by this A. niger enzyme cocktail allowed up to 2.25-time higher bioethanol production under the same simultaneous saccharification and fermentation conditions (Rocha et  al. 2013). A solid-state fermentation was also used by Rehman et  al. (2014) for the production of various plant cell degrading enzymes by A. niger in the presence of banana peels as the substrate. In contrast, Nitesh et al. (2013) applied submerged fermentation for the production of cellulase enzyme from A. niger for the subsequent saccharification of weed plant (Cyperus rotundus). A. niger has also been exploited for bioethanol production using simultaneous saccharification and fermentation. Through this strategy, Hatami et al. (2015) converted rice cooker wastewater into bioethanol using co-culture of A. niger (as hydrolyzing microorganism) and S. cerevisiae (as ethanologen). Under optimum condition (35  °C, pH  5, and 36  h), bioethanol concentration and yield of 16.97  g/L and 0.36 g/g, respectively, were achieved. More recently, a similar approach has been

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used for potato waste conversion into bioethanol in biofilm reactors (Izmirlioglu and Demirci 2017). Respectively, maximum bioethanol yield and production of 0.41  g/g starch and ~37.9  g/L were delivered at 35  °C and pH  5.8 after 72  h fermentation.

2.4.2  Aspergillus aculeatus Aspergillus aculeatus (A. aculeatus) causes post-harvest disease in fresh or dried fruits, vegetables, oil seeds, and nuts in association with other Aspergillus spp. (e.g., A. niger and Aspergillus carbonarius) (Perrone et  al. 2007). More significantly, A. aculeatus as well as Aspergillus japonicas are grape pathogens (Perrone et al. 2007; Rosa et al. 2002). A. aculeatus could produce all three cellulase groups for complete cellulose degradation. Among them, β-glucosidase, the most important enzyme for cellulose hydrolysis, is secreted in a greater amount by A. aculeatus (Du et al. 2010; Molina et al.2016). This is a highly desirable trait when it is considered that T. reesei, one of the most important cellulolytic microorganisms, secretes a less amount of β-glucosidase in its enzyme cocktail, compared to those of endo-β-1,4-­ glucanases and cellobiohydrolases. In better word, insufficient β-glucosidase activity in commercial cellulolytic cocktails could be addressed through A. aculeatus β-glucosidase supplementation. Moreover, an acidothermotolerant ß-glucosidase was characterized from A. aculeatus, exhibiting maximum activity at 70  °C and pH  5–6 (Li et  al. 2018). Recently, an enzyme cocktail showing stability against ionic liquid was prepared from A. aculeatus and successfully used in the saccharification of 1-ethyl-3-methylimidazolium methane sulfonate pretreated Parthenium hysterophorus biomass (Nargotra et al. 2019). Alternatively, recombinant technology could be employed for more prominent secretory expression of β-glucosidase in T. reesei from A. aculeatus. On this basis, a recombinant T. reesei strain X3AB1 was constructed, expressing A. aculeatus β-glucosidase 1 (Nakazawa et al. 2012). A significantly higher β-glucosidase activity (63 times) against cellobiose was observed from culture supernatant of T. reesei strain X3AB1, compared to that of the parent strain PC-3-7 (Nakazawa et al. 2012). Similarly, a recombinant T. reesei expressing A. aculeatus β-glucosidase 1 was constructed (Treebupachatsakul et al. 2015), producing an enzyme cocktail with a comparative saccharification performance to that of T. reesei PC-3-7 supplemented with Novozyme 188. In another study (Treebupachatsakul et  al. 2016), a recombinant S. cerevisiae expressing A. aculeatus β-glucosidase gene was used for bioethanol production (i.e., simultaneous saccharification and fermentation) in the presence of T. reesei. Overall, A. aculeatus produces high amount of stable β-glucosidase that could be used in the saccharification of wide range of cellulosic biomasses in bioethanol industry. The β-glucosidase gene could also be expressed in either potent hydrolytic-enzyme-producing microorganisms (e.g., T. reesei) or potent ethanologens (e.g., S. cerevisiae). The mentioned approaches could improve the bioethanol production feasibility from lignocelluloses.

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2.4.3  Aspergillus flavus Aspergillus flavus (A. flavus) is a saprophytic soil and opportunistic pathogenic fungus that infects important agricultural crops. It is the causative agent of pre- or post-­ harvest diseases such as ear rot (on maize) and yellow mold (on peanut) (Amaike and Keller 2011; Yu et  al. 2005). This species produces a secondary metabolite aflatoxin (toxicant carcinogens) in the infected seeds of a number of crops, further complicating the pathogenesis by posing risks to humans and animals health. A. flavus has a rich genome with high capabilities for encoding a wide range of cell wall-­ degrading enzymes that have important role in the fungal virulence. Efforts have been done to exploit the A. flavus enzyme capabilities in bioethanol industry. Extremozymes such as an alkaline halophilic cellulase (optimum activity; 60 °C, pH 10, and 200 g/L NaCl) and a halotolerant (65% activity in 2 M NaCl solution after 1 d incubation) and acidic (62% activity in pH 4–7 after 1 h incubation) thermostable 63-kDa-glucoamylase (optimum activity, 50 °C and pH 5.5) have already been characterized from various A. flavus strains (Ayodeji et al. 2017; Bano et al. 2019). A. flavus strain FPDN1 and ARC-12 could also produce xylanase (1530 and ~2220  IU/gds/min, respectively) under solid-state fermentation with pearl millet bran or stover as the substrate (Archana et al. 2017; Nikhil et al. 2012). It was also reported that co-cultivation of A. flavus and Trichoderma viride could ecofriendly and efficiently produce an enzyme cocktail exhibiting xylanase (180 IU/g) and cellulase (FPase, 11 IU/g) activities (Narsale et al. 2018; Singh et al. 2018). In 2017, rice husk was enzymatically hydrolyzed into simple sugars in the presence of A. flavus. The subsequent ethanolic fermentation using S. cerevisiae yielded ~14.9 vol.% bioethanol after distillation (Onwuakor et al. 2017). More recently, A. flavus was also successfully applied for the saccharification of pretreated millet straw (cellulose, 30%; hemicellulose, 10%; total organic carbon, ~9%; and total nitrogen, ~1%) under solid-state fermentation (70% moisture content, 28 °C, pH 4.5, and 120 h) (Narsale et al. 2018). Alternatively, A. flavus hydrolytic enzyme-expressing genes could be exploited to improve S. cerevisiae performance in consolidated bioprocessing systems for bioethanol production. On this basis, endoglucanase gene (i.e., endo753) from A. flavus NRRL3357 was successfully cloned and efficiently expressed on the surface of S. cerevisiae EBY100 (Gao et al. 2017). The resulting whole-cell enzyme showed a high pH stability (7–10; optimum, 8) and moderate thermostability (≥70 °C; optimum, 50 °C).

2.5  Phoma exigua Genus Phoma belongs to family Didymellaceae, order Pleosporales, class Dothideomycetes (subclass Pleosporomycetidae), phylum Ascomycota (subdivision Pezizomycotina), and kingdom Fungi. This diverse filamentous fungal genus

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contains many plant pathogenic species, which are generally found in soil, organic matter, plants, and water sources (Boerema 2004). Among them, Phoma exigua (P. exigua), also known as Boeremia exigua, is one of the important plant pathogenic fungi that can infect injured parts of many plant genera, including Cucumis, Acroptilon, Glycine, Lupinus, Pisum, Humulus, Lactuca, Raphanus, etc. (Koike et al. 2006; Machowicz-Stefaniak et al. 2008; Tunali et al. 2003). In 2013, P. exigua ITCC 2049, a potato pathogen, was successfully used for the production of cellulases (i.e., ß-glucosidase, 2.67 IU/mL; FPase, 1.13 IU/mL; and endoglucanase, 37 IU/mL) and hemicellulase (i.e., xylanase, 24.92 IU/mL) in an 8-d submerged fermentation (Tiwari et al. 2013). The subsequent saccharification of paddy straw and Parthenium biomass with this enzyme cocktail delivered up to 651–698 mg/gds sugars in the presence of commercial ß-glucosidase. In 2014, the hydrolytic system of P. exigua was unwrapped for the first time (Tiwari et al. 2014). Accordingly, it was found that the secretome consists of 33 proteins when this species is cultured on α-cellulose as the sole carbon source. The enzyme system consisted of hemicellulases (i.e., endo-1,4-ß-xylanase and 1,4-ß-xylosidase), cellulases (i.e., ß-glucosidase, exoglucanase, cellobiohydrolase I, and endo-1,4-ß-glucanase), and some hypothetical proteins (such as GH5–GH7, GH20, and GH54) (Tiwari et al. 2014). Following secretome decipher, the enzymatic exploitation on P. exigua in bioethanol industry received more attentions. However, despite the rich enzyme cocktail, the ß-glucosidase activity was very limited. The supplementation of the enzyme cocktail with commercial ß-glucosidase is therefore required for a successful saccharification of delignified lignocellulose. Through this strategy, high yields of arabinose (25.2  mg/gds), xylose (209.2  mg/gds), and glucose (177.2  mg/gds) could be delivered from the saccharification (55 °C, 72 h) of alkali-treated wheat straw (1% w/v) by P. exigua enzyme cocktail (15.23 FPase units) supplemented with Novozyme 188 (31.95 units) (Tiwari et al. 2014).

2.6  Chrysoporthe cubensis Genus Chrysoporthe belongs to family Cryphonectriaceae, order Diaporthales, class Sordariomycetes (subclass Diaporthomycetidae), phylum Ascomycota (subdivision Pezizomycotina), and kingdom Fungi. Chrysoporthe cubensis (C. cubensis), previously known as Cryphonectria cubensis, is a serious and often deadly canker pathogen that primarily affects Syzygium spp., Tibouchina spp., and eucalypts (i.e., Corymbia and Eucalyptus spp.). The symptoms may range from foliage wilt and dieback (due to stem girdling) to bark swelling, cracking, and splitting at the trees bases and sunken canker formation on stems. The formation of cankers on the trunks of trees reduces the plant growth and could lead to stem breakage or tree death (Gryzenhout et al. 2004). In contrast, C. cubensis could be a novel source for hydrolyzing enzymes (i.e., cellulases and hemicellulases) with potential applications in delignified lignocellulose saccharification step during bioethanol production. Various cellulases and

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hemicellulases could be produced in the presence of different lignocellulosic biomass as the carbon source using C. cubensis via solid-state fermentation. For example, wheat bran as the carbon source could deliver 33.84, 2.52, 21.55, and 362.38 U/g of endoglucanase, FPase, β-glucosidase, and xylanase activities, respectively (Falkoski et al. 2013). The enzymes showed maximal activity at 50–60 °C and pH 4, delivering up to 321 mg/g glucose and 288.7 mg/g xylose in the subsequent saccharification of the alkaline-pretreated sugarcane bagasse (Falkoski et  al. 2013). Intriguingly, C. cubensis enzyme cocktail showed higher specific activities (U/mg of protein) in terms of endoglucanase (~331.8), pectinase (~127.5), ß-glucosidase (~29.5), ß-xylosidase (~3.0), and laccase (~2.5), compared to three commercial enzyme cocktails (i.e., Accellerase®, Multifect® CL, and Multifect® XL) (Maitan-­ Alfenas et al. 2015). A 80% enzyme synergy, especially form endoglucanase and FPase activities (48% and 76% higher than theoretical activities, respectively) could be obtained by blending C. cubensis enzyme extract with that of Penicillium pinophilum (ratio, 1:1 v/v) while the blend could resist up to 35 mg phenols/mg protein (~116.5- and 23.3-time higher phenol concentration, compared to enzymes from A. niger and T. reesei, respectively) (Ázar et al. 2018; Visser et al. 2013). An enzyme blend quantity of 20 FPU/g could, respectively, deliver 93.3% and 63.8% xylose and glucose conversions during saccharification of alkali-treated sugarcane bagasse. More specifically, >30  g/L glucose and ~17  g/L xylose were liberated from the substrate (solid content, 8%) at 45 °C within 120 h incubation (Visser et al. 2013). Further research showed that C. cubensis could produce (via solid-state fermentation of wheat bran) a group of three ß-glucosidases with specific activity for hydrolysis of glucose residues in ß position (de Andrade et al. 2017). Accordingly, the purified ß-glucosidases could withstand temperatures up to 70  °C while showed maximum activity at 60 °C and pH 4 during sugarcane bagasse saccharification. In natura sugarcane bagasse could also be used as a substrate for cellulases and hemicellulase production using C. cubensis. Through this strategy, an enzyme cocktail showing respective carboxymethylcellulase, xylanase, β-xylosidase, α-galactosidase, and mannanase activities of 33.2 U/g, 602 U/g, 2.0 U/g, 2.4 U/g, and 7.1 U/g could be produced (Dutra et al. 2017). The purification and characterization of xylanases from C. cubensis showed three acidic endo-ß-1,4-xylanases groups, including P1, P2, and P3, with maximum activities at 60 °C and pH 4, 55 °C and pH 3, and 80 °C and pH 3, respectively (de Sousa Gomes et al. 2017). Using oat spelt xylan as the substrate, the KM values of 2.65 mg/mL, 1.81 mg/mL, and 1.18 mg/mL were, respectively, recorded for the mentioned xylanases while delivering xylotriose and xylobiose as the main xylooligosaccharides (de Sousa Gomes et al. 2017).

2.7  Ustilago maydis Genus Ustilago belongs to family Ustilaginaceae, order Ustilaginales, class Ustilaginomycetes (subclass Ustilaginomycetidae), phylum Basidiomycota (subdivision Ustilaginomycotina), and kingdom Fungi. Ustilago, the largest genus of the

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family Ustilaginaceae, consists of about 300 species, most of which are parasites on monocotyledonous hosts. Among them, Ustilago maydis (U. maydis), also known as Ustilago zeae, is one of the major pathogenic species on corn, causing common corn smut disease (Kämper et  al. 2006). This phytopathogenic fungus can easily penetrate into plant through plant cell wall hydrolysis by its unique enzymatic system (Couturier et  al. 2016). More specifically, this species could secrete a wide range of lignocellulose-degrading enzymes (e.g., hemicellulases and oxidoreductases) during corn pathogenicity. Post-genomic analyses of 20 filamentous fungi were performed and compared with each others using enzyme profiling method (Couturier et  al. 2012). Among them, the most promising result was observed by wheat straw co-saccharification using U. maydis and industrial T. reesei CL8 47 enzyme cocktails. Compared to industrial T. reesei CL8 47 enzyme cocktail alone, the enzyme blend could enhance glucose and the total sugar liberations by 22% and 57%, respectively. The improvement effect could be attributed to U. maydis ability for an abundant secretion of carbohydrate-acting enzymes that were only expressed in limited amounts by T. reesei. In a later study, a new lignin-targeting aryl-alcohol oxidase enzyme (i.e., glucose-methanol-choline) was characterized from U. maydis and expressed in Pichia pastoris (Couturier et al. 2016). Therefore, U. maydis, its enzyme cocktail, or its gene expression on other microbial hosts could be used during delignification process, i.e., a crucial step during lignocellulose valorization into bioethanol.

2.8  Colletotrichum graminicola Genus Colletotrichum belongs to family Glomerellaceae, order Glomerellales, class Sordariomycetes (subclass Hypocreomycetidae), phylum Ascomycota (subdivision Pezizomycotina), and kingdom Fungi. Anthracnose, caused by different Colletotrichum sp. (such as C. graminicola), is one of the most important diseases that affects approximately 2200 plant species including grass, vegetables, fruits, and trees (Farr et al., 2016). Some of the main symptoms of this disease include dark lesions or blight on different part (i.e., leaves, stems, flowers, and fruits) of the infected hosts (Hyde et al. 2009). C. graminicola is a hemibiotrophic fugal pathogen that causes anthracnose disease of maize following a mechanical penetration into the plant epidermis (via melanized appressorium) (Sukno et al. 2008; Torres et al. 2016). The disease is associated with leaf blight, stem rot, and root infections. In fact, a period of biotrophy (intracellular colonization of living host cells) is observed prior to necrotrophic phase (symptom development). During these periods, plant immune system is evaded with minor responses through stealthy methods, and plant cells are rapidly killed and major molecular responses from plants are elicited, respectively. C. graminicola showed a good potential for the production of cellulolytic enzymes (β-glucosidase, 109.7  U/g; β-xylosidase, 57.9  U/g; and xylanase, 189.3 U/g) under solid-state fermentation of wheat bran (Zimbardi et al. 2013). The

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substrate supplementation with sugarcane trash, peanut hulls, and corncob could improve the concentration of ß-glucosidase (~45.2%), β-xylosidase (~121.2%), and xylanase (~99.7%), delivering 159.3, 128.1, and 378.1 U/g enzymes, respectively (Zimbardi et al. 2013). Accordingly, these enzymes showed highest activity at 65 °C and pH 5 and were fully active at 60–65 °C for 2 h. A glucose yield of 33.1% was obtained, following a 48-h saccharification process of sugarcane trash by these enzymes blended with T. reesei cellulases. In 2016, the first fungal thermostable, halotolerant, and solvotolerant endo-xylanase was purified from C. graminicola (Excg1) that could be applied for lignocellulose hydrolysis under harsh condition (Carli et al. 2016). Excg1 (monomeric, molecular weight of 17.3–20 kDa) was fully stable at 50 °C (half-life, 48 h), pH 3–10, and against various organic solvents (5% v/v). While salt presence had no effect on thermotolerancy, the pH stability range slightly dropped to 4–10 in the solutions containing 2.9–14.6 wt.% NaCl. The optimum activity was observed at 65 °C and pH 5.5 in the presence of 0–2.9 wt.% NaCl, whereas the optimum pH rose to 6 when salt concentration elevated to 14.6 wt.% NaCl (Carli et al. 2016). With or without saline water (up to 2.9 wt.% NaCl), Excg1 exhibited good maximal velocity (481.3 U/mg) and affinity constant (3.7 mg/mL) during beechwood xylan conversion into xylotriose and xylobiose (Carli et al. 2016).

2.9  Conclusions Biofuels can be considered as a promising replacement for nonrenewable fossil-fuel to compensate its devastating environmental and health impacts. A contribution more than 70% of the total biofuel production made the bioethanol the major alternative for phasing out the gasoline. Bioethanol production reached ~108.14 billion L in 2018, with fermentation technology contribution of about 80%. However, the feedstocks mainly consumed were food commodity such as corn and sugarcane. Application of these crops as substrates has great environmental burdens while imposing food insecurity on nations. Changing these substrates with lignocelluloses could provide reliefs in respect to biofuel production, waste and forestry management, social benefits, and environment conservations. In contrast, the bioethanol production cost is elevated due to several additional pretreatments steps (i.e., delignification and hydrolysis) that are required prior to fermentation step. On the other hand, high requirement for inorganic acid during hydrolysis step has some negative environmental consequences, further limiting the commercial production of lignocellulose-­derived bioethanol. Microorganism-aided lignocellulose degradation could significantly boost both environmental and economic feasibilities of bioethanol production process. Among them, pathogenic filamentous fungi are known for their devastating crop deterioration due to their strong hydrolytic enzyme machinery. Mainly, Basidiomycota could participate in delignification process (e.g., U. maydis) as whole fungal cell or their enzyme cocktails. Compared to delignification process, plant pathogenic fungi (both Ascomycota and Basidiomycota) could provide substantial contributions on

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cellulase and hemicellulase saccharification step. On this context, Fusarium spp. and Aspergillus spp. are the top two most studied pathogenic fungal genera, followed by C. cubensis, C. graminicola, U. maydis, and P. exigua. These fungi could be used in bioethanol industry via simultaneous saccharification and fermentation, simultaneous saccharification and co-fermentation, and/or separate hydrolysis and fermentation methods. It is worth mentioning that some plant pathogenic fungi have even the capacity to fermented pretreated lignocellulose into bioethanol. The potential of F. verticillioides and F. oxysporum for direct ethanol production in consolidated bioprocessing system has already been investigated. However, some challenges including large amounts of by-products (i.e., reducing bioethanol yields and production efficiency), low productivity, and low growth rates must be addressed (e.g., through genetic technology) for commercial bioethanol production using pathogenic filamentous fungi. Moreover, precautionary measures must be taken to avoid accidental release of these pathogenic fungi into the environment. Alternatively, valuable hydrolyses genes of pythopatogenic fungi could be expressed in either superb hydrolytic-­ enzyme-­producing microorganisms (e.g., T. reesei) or potent ethanologens (e.g., S. cerevisiae). Through this strategy, various recombinant microorganisms such as S. cerevisiae EBY100, Pichia pastoris, Escherichia coli, and T. reesei strain X3AB1S and S. cerevisiae have already been constructed from A. flavus NRRL3357 (endoglucanase gene, endo753), U. maydis (lignin-targeting aryl-alcohol oxidase enzyme, glucose-methanol-choline), F. graminearum (xylanase genes, XylD, XyloA, and Xylo/ArabA), and A. aculeatus (β-glucosidase).

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Nakazawa H, Kawai T, Ida N, Shida Y, Kobayashi Y, Okada H, Tani S, Sumitani Ji, Kawaguchi T, Morikawa Y (2012) Construction of a recombinant Trichoderma reesei strain expressing Aspergillus aculeatus β-glucosidase 1 for efficient biomass conversion. Biotechnol Bioeng 109:92–99 Nargotra P, Sharma V, Bajaj BK (2019) Consolidated bioprocessing of surfactant-assisted ionic liquid-pretreated Parthenium hysterophorus L. biomass for bioethanol production. Bioresour Technol 289:121611 Narsale PV, Patel SR, Acharya P (2018) Role of Aspergillus flavus on biodegradation of lignocellulosic waste millet straw and optimization parameters for enzyme hydrolysis and ethanol production under solid state fermentation. Int J Curr Microbiol App Sci 7:429–445 Nikhil B, Adhyaru D, Thakor P (2012) Production of xylanase by Aspergillus flavus FPDN1 on pearl millet bran: optimization of culture conditions and application in bioethanol production. Int J Res Chem Environ 2:204–210 Nitesh K, Singh J, Ravi R, Subathradevi C, Srinivasan V (2013) Bioethanol production from weed plant (Cyperus rotundus) by enzymatic hydrolysis. Adv Appl Sci Res 4:299–302 Oliver RP, Ipcho SV (2004) Arabidopsis pathology breathes new life into the necrotrophs-vs.biotrophs classification of fungal pathogens. Mol Plant Pathol 5:347–352 Onwuakor CE, Hans-Anukam U, Uzokwe MJ (2017) Production of ethanol and biomass from rice husk using cultures of Aspergillus flavus, Aspergillus eamarii and Saccharomyces cerevisiae. Am J Microbiol Res 5:86–90 Ouellet M, Datta S, Dibble DC, Tamrakar PR, Benke PI, Li C, Singh S, Sale KL, Adams PD, Keasling JD (2011) Impact of ionic liquid pretreated plant biomass on Saccharomyces cerevisiae growth and biofuel production. Green Chem 13:2743–2749 Paccanaro MC, Sella L, Castiglioni C, Giacomello F, Martínez-Rocha AL, D’Ovidio R, Schäfer W, Favaron F (2017) Synergistic effect of different plant cell wall–degrading enzymes is important for virulence of Fusarium graminearum. Mol Plant-Microbe Interact 30:886–895 Palacios-Cabrera H, Taniwaki MH, Hashimoto JM, Menezes HCd (2005) Growth of Aspergillus ochraceus, A carbonarius and A niger on culture media at different water activities and temperatures. Braz J Microbiol 36:24–28 Palencia ER, Hinton DM, Bacon CW (2010) The black Aspergillus species of maize and peanuts and their potential for mycotoxin production. Toxins 2:399–416 Panagiotou G, Christakopoulos P, Villas-Bôas SG, Olsson L (2005) Fermentation performance and intracellular metabolite profiling of Fusarium oxysporum cultivated on a glucose–xylose mixture. Enzym Microb Technol 36:100–106 Paschos T, Xiros C, Christakopoulos P (2015) Simultaneous saccharification and fermentation by co-cultures of Fusarium oxysporum and Saccharomyces cerevisiae enhances ethanol production from liquefied wheat straw at high solid content. Ind Crop Prod 76:793–802 Perrone G, Susca A, Cozzi G, Ehrlich K, Varga J, Frisvad JC, Meijer M, Noonim P, Mahakarnchanakul W, Samson RA (2007) Biodiversity of Aspergillus species in some important agricultural products. Stud Mycol 59:53–66 Phalip V, Delalande F, Carapito C, Goubet F, Hatsch D, Leize-Wagner E, Dupree P, Van Dorsselaer A, Jeltsch J-M (2005) Diversity of the exoproteome of Fusarium graminearum grown on plant cell wall. Curr Genet 48:366–379 Pioli R, Mozzoni L, Morandi E, Menard M (2004) Disease notes. Plant Dis 88:220 Pitt JI, Hocking AD (2009) Fungi and food spoilage. Springer, Dordrecht/Heidelberg/London/ New York Plascencia-Jatomea M, Susana M, Gómez Y, Velez-Haro JM (2014) Aspergillus spp. (black mold). In: Bautista-Baños S (ed) Postharvest decay–control strategies. Elsevier, Cambridge, MA, pp 267–286 Rahimzadeh H, Tabatabaei M, Aghbashlo M, Panahi H, Rashidi A, Goli S, Mostafaei M, Ardjmand M, Nizami A (2018) Potential of acid-activated bentonite and SO3H-functionalized MWCNTs for biodiesel production from residual olive oil under biorefinery scheme. Front Energy Res 6:137

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Ravalason H, Grisel S, Chevret D, Favel A, Berrin J-G, Sigoillot J-C, Herpoël-Gimbert I (2012) Fusarium verticillioides secretome as a source of auxiliary enzymes to enhance saccharification of wheat straw. Bioresour Technol 114:589–596 Rehman S, Aslam H, Ahmad A, Khan SA, Sohail M (2014) Production of plant cell wall degrading enzymes by monoculture and co-culture of Aspergillus niger and Aspergillus terreus under SSF of banana peels. Braz J Microbiol 45:1485–1492 Rocha NRdAF, Barros MA, Fischer J, Coutinho Filho U, Cardoso VL (2013) Ethanol production from agroindustrial biomass using a crude enzyme complex produced by Aspergillus niger. Renew Energy 57:432–435 Rosa CDR, Palacios V, Combina M, Fraga M, Rekson ADO, Magnoli C, Dalcero AM (2002) Potential ochratoxin a producers from wine grapes in Argentina and Brazil. Food Addit Contam 19:408–414 Ruegg TL, Kim E-M, Simmons BA, Keasling JD, Singer SW, Lee TS, Thelen MP (2014) An auto-inducible mechanism for ionic liquid resistance in microbial biofuel production. Nat Commun 5:3490 Saha B (2001) Xylanase from a newly isolated Fusarium verticillioides capable of utilizing corn fiber xylan. Appl Microbiol Biotechnol 56:762–766 Salehi Jouzani G, Taherzadeh MJ (2015) Advances in consolidated bioprocessing systems for bioethanol and butanol production from biomass: a comprehensive review. Biofuel Res J 2:152–195 Samson RA, Visagie CM, Houbraken J, Hong S-B, Hubka V, Klaassen CH, Perrone G, Seifert KA, Susca A, Tanney JB (2014) Phylogeny, identification and nomenclature of the genus Aspergillus. Stud Mycol 78:141–173 Schuster E, Dunn-Coleman N, Frisvad J, Van Dijck P (2002) On the safety of Aspergillus niger–a review. Appl Microbiol Biotechnol 59:426–435 sheng Xue D, Zeng X, Lin D, Yao S (2018) Ethanol tolerant endoglucanase from Aspergillus niger isolated from wine fermentation cellar. Biocatal Agric Biotechnol 15:19–24 Shirzad M, Kazemi Shariat Panahi H, Dashtic BB, Rajaeifard MA, Aghbashlo M, Tabatabaei M (2019) A comprehensive review on electricity generation and GHG emission reduction potentials through anaerobic digestion of agricultural and livestock/slaughterhouse wastes in Iran. Renew Sust Energ Rev 111:571–594 Singh N, Devi A, Jaryal R, Rani K (2018) An ecofriendly and efficient strategy for cost effective production of lignocellulotic enzymes. Waste Biomass Valoriz 9:891–898 Sukno SA, García VM, Shaw BD, Thon MR (2008) Root infection and systemic colonization of maize by Colletotrichum graminicola. Appl Environ Microbiol 74:823–832 Sutton J (1982) Epidemiology of wheat head blight and maize ear rot caused by Fusarium graminearum. Can J Plant Pathol 4:195–209 Szécsi Á (2014) Screening of Fusarium verticillioides strains for cellulase activity. Acta Phytopathol Entomol Hung 49:1–10 Tabatabaei M, Aghbashlo M, Dehhaghi M, Kazemi Shariat Panahi H, Mollahosseini A, Hosseini M (2019a) Reactor technologies for biodiesel production and processing: a review. Prog Energy Combust Sci 74:239–303 Tabatabaei M, Aghbashlo M, Valijanian E, Kazemi Shariat Panahi H, Nizami A-S, Ghanavati H, Sulaiman A, Mirmohamadsadeghi S, Karimi K (2019b) A comprehensive review on recent biological innovations to improve biogas production, part 1: upstream strategies. Renew Energy Tabatabaei M, Aghbashlo M, Valijanian E, Kazemi Shariat Panahi H, Nizami A-S, Ghanavati H, Sulaiman A, Mirmohamadsadeghi S, Karimi K (2019c) A comprehensive review on recent biological innovations to improve biogas production, part 2: mainstream and downstream strategies. Renew Energy 146:1392–1407 Thamizharasi V, Narasimham P (1993) Growth of Aspergillus niger on onion bulbs and its control by heat and sulphur dioxide treatments. Trop Sci 33:1 Tiwari R, Singh S, Nain PK, Rana S, Sharma A, Pranaw K, Nain L (2013) Harnessing the hydrolytic potential of phytopathogenic fungus Phoma exigua ITCC 2049 for saccharification of lignocellulosic biomass. Bioresour Technol 150:228–234

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Tiwari R, Singh S, Singh N, Adak A, Rana S, Sharma A, Arora A, Nain L (2014) Unwrapping the hydrolytic system of the phytopathogenic fungus Phoma exigua by secretome analysis. Process Biochem 49:1630–1636 Tiwari R, Nain PK, Singh S, Adak A, Saritha M, Rana S, Sharma A, Nain L (2015) Cold active holocellulase cocktail from Aspergillus niger SH3: process optimization for production and biomass hydrolysis. J Taiwan Inst Chem Eng 56:57–66 Torres MF, Ghaffari N, Buiate EA, Moore N, Schwartz S, Johnson CD, Vaillancourt LJ (2016) A Colletotrichum graminicola mutant deficient in the establishment of biotrophy reveals early transcriptional events in the maize anthracnose disease interaction. BMC Genomics 17:202 Treebupachatsakul T, Shioya K, Nakazawa H, Kawaguchi T, Morikawa Y, Shida Y, Ogasawara W, Okada H (2015) Utilization of recombinant Trichoderma reesei expressing Aspergillus aculeatus β-glucosidase I (JN11) for a more economical production of ethanol from lignocellulosic biomass. J Biosci Bioeng 120:657–665 Treebupachatsakul T, Nakazawa H, Shinbo H, Fujikawa H, Nagaiwa A, Ochiai N, Kawaguchi T, Nikaido M, Totani K, Shioya K (2016) Heterologously expressed Aspergillus aculeatus β-glucosidase in Saccharomyces cerevisiae is a cost-effective alternative to commercial supplementation of β-glucosidase in industrial ethanol production using Trichoderma reesei cellulases. J Biosci Bioeng 121:27–35 Tunali B, Eskandari F, Berner D, Farr D, Castlebury L (2003) First report of leaf blight caused by Phoma exigua on Acroptilon repens in Turkey. Plant Dis 87:1540–1540 Visser EM, Falkoski DL, de Almeida MN, Maitan-Alfenas GP, Guimarães VM (2013) Production and application of an enzyme blend from Chrysoporthe cubensis and Penicillium pinophilum with potential for hydrolysis of sugarcane bagasse. Bioresour Technol 144:587–594 Vitale A, Castello I, Polizzi G (2008) First report of Aspergillus vine canker on table grapes caused by Aspergillus niger in Europe. Plant Dis 92:1471–1471 Wang Y, Radosevich M, Hayes D, Labbé N (2011) Compatible ionic liquid-cellulases system for hydrolysis of lignocellulosic biomass. Biotechnol Bioeng 108:1042–1048 Xiros C, Topakas E, Katapodis P, Christakopoulos P (2008) Evaluation of Fusarium oxysporum as an enzyme factory for the hydrolysis of brewer's spent grain with improved biodegradability for ethanol production. Ind Crop Prod 28:213–224 Xiros C, Katapodis P, Christakopoulos P (2009) Evaluation of Fusarium oxysporum cellulolytic system for an efficient hydrolysis of hydrothermally treated wheat straw. Bioresour Technol 100:5362–5365 Xiros C, Vafiadi C, Paschos T, Christakopoulos P (2011) Toxicity tolerance of Fusarium oxysporum towards inhibitory compounds formed during pretreatment of lignocellulosic materials. J Chem Technol Biotechnol 86:223–230 Yu J, Cleveland TE, Nierman WC, Bennett JW (2005) Aspergillus flavus genomics: gateway to human and animal health, food safety, and crop resistance to diseases. Rev Iberoam Micol 22:194–202 Ziani K, Fernández-Pan I, Royo M, Maté JI (2009) Antifungal activity of films and solutions based on chitosan against typical seed fungi. Food Hydrocoll 23:2309–2314 Zimbardi A, Sehn C, Meleiro L, Souza F, Masui D, Nozawa M, Guimarães L, Jorge J, Furriel R (2013) Optimization of β-glucosidase, β-xylosidase and xylanase production by Colletotrichum graminicola under solid-state fermentation and application in raw sugarcane trash saccharification. Int J Mol Sci 14:2875–2902

Chapter 3

Fungi as Bioreactors for Biodiesel Production Meisam Tabatabaei, Amin Alidadi, Mona Dehhaghi, Hamed Kazemi Shariat Panahi, Su Shiung Lam, Abdul-Sattar Nizami, Mortaza Aghbashlo, and Gholamreza Salehi Jouzani

3.1  Introduction The world population will increase up to 9 billion by 2050, significantly increasing the energy demand. This, in turn, increases the emissions of greenhouse gases (GHG), polluting gases, and particulate matters, causing environmental pollution and climate change and global warming (Kazemi Shariat Panahi et al. 2019b; Shirzad et al. 2019). Therefore, to ensure environment protection and to achieve sustainable economic growth, application of different types of biofuels as renewable energy carriers have been considered in some regions of the world (Dehhaghi et al. 2019b; Kazemi Shariat Panahi

M. Tabatabaei (*) Institute of Tropical Aquaculture and Fisheries (AKUATROP), Universiti Malaysia Terengganu, Terengganu, Malaysia Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran e-mail: [email protected] A. Alidadi · G. Salehi Jouzani (*) Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran e-mail: [email protected] M. Dehhaghi · H. Kazemi Shariat Panahi Biofuel Research Team (BRTeam), Karaj, Iran Department of Microbial Biotechnology, School of Biology and Centre of Excellence in Phylogeny of Living Organisms, College of Science, University of Tehran, Tehran, Iran Faculty of Medicine and Health Sciences, Macquarie University, Sydney, NSW, Australia

© Springer Nature Switzerland AG 2020 G. Salehi Jouzani et al. (eds.), Fungi in Fuel Biotechnology, Fungal Biology, https://doi.org/10.1007/978-3-030-44488-4_3

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et al. 2019a; Shirzad et al. 2019; Tabatabaei et al. 2019b, c). Liquid biofuels (e.g., bioethanol, biobutanol, biocrude oil, and biodiesel) are well-known alternatives to fossil fuels and are used (or have been used in many field trials) in the transportation sector (Hosseinzadeh-Bandbafha et al. 2018; Kazemi Shariat Panahi et al. 2019b, c; Salehi Jouzani et al. 2018; Shirzad et al. 2019; Tabatabaei et al. 2019a). Among them, bioethanol (see Chaps. 2, 4, and 7) and biodiesel are the most promising alternatives (Kazemi Shariat Panahi et al. 2019a, c; Tabatabaei et al. 2019a). The former biofuel is a mixture of fatty acid alkyl esters, which are produced by transesterification or, to a lesser extent, esterification method (Rahimzadeh et al. 2018; Tabatabaei et al. 2019a). Biodiesel contains monoalkyl esters with long-chain fatty acids that could be derived from plant oils or animal fats (Tabatabaei et al. 2019a). It is one of the most attractive biofuels, as it is environmentally friendly, a low CO2 and particulate matter emitter, renewable, highly degradable, compatible with current commercial diesel engines, and a non-­hydrocarbon emitter and has zero toxicity (Tabatabaei et al. 2019a). Compared to petroleum diesel, some advantages of biodiesel include high cetane number, high flash point, excellent lubricity, no sulfur content, and lower aromatic content (Tabatabaei et al. 2019a; Salehi Jouzani et al. 2018; Tabatabaei et al. 2011). This type of biofuel could basically synthesize from lipids as the substrate. These molecules act as storage materials in different organisms and are present in the form of lipid bodies in the cells (i.e., triglycerides (TGs) and membranes lipids). TGs are long-­chain fatty acids, which are chemically bound to a glycerol backbone (Hegde et al. 2015; Salehi Jouzani et al. 2018). Plant oils (e.g., soybean, canola, rapeseed, palm, safflower, almond, barley, coconut, cotton seed, and groundnut) and animal fats could be rich sources of TGs for biodiesel production (Tabatabaei et al. 2019a). However, the production of biodiesel from plant oils has some negative impacts on world food security, and in fact, considerable area of cultivable lands must be used for sufficient production of oil feedstock (i.e., oil crops) to meet the growing biodiesel demands (Tabatabaei et al. 2019a). Alternatively, novel oil feedstock sources (e.g., single-cell oils, SCOs) that are cheaper and do not interfere with food security are appreciable (Salehi Jouzani et al. 2018; Vicente et al. 2009). On this basis, SCOs could be obtained from microorganisms (e.g., algae and fungi) which can produce and accumulate high concentrations of lipids. It is worth mentioning that microorganisms are potent producers of various industrial important metabolites ranging from S. S. Lam Henan Province Engineering Research Center for Forest Biomass Value-added Products, School of Forestry, Henan Agricultural University, Zhengzhou, China Pyrolysis Technology Research Group, Institute of Tropical Aquaculture and Fisheries (AKUATROP) & Institute of Tropical Biodiversity and Sustainable Development (Bio-D Tropika), Universiti Malaysia Terengganu, Kuala Nerus, Terengganu, Malaysia A.-S. Nizami Sustainable Development Study Center, Government College University, Lahore, Pakistan M. Aghbashlo (*) Department of Mechanical Engineering of Agricultural Machinery, Faculty of Agricultural Engineering and Technology, College of Agriculture and Natural Resources, University of Tehran, Karaj, Iran e-mail: [email protected]

3  Fungi as Bioreactors for Biodiesel Production

41

therapeutic agents (Dehhaghi et al. 2018; Mohammadipanah et al. 2016; Sajedi et al. 2018) to enzymes (Dehhaghi et al. 2018, 2019a; Hamedi et al. 2015; Mohammadipanah et al. 2015, 2016; Sajedi et al. 2018). From the biodiesel industry point of view, microbial enzymes (lipases), preferably those produced by oleaginous microorganisms, could be exploited during enzymatic transesterification reaction. In this chapter, the potentials of fungi in biodiesel industry have been scrutinized with respect to their capabilities as oil feedstock (lipids) and sources of enzyme (lipase).

3.2  Oleaginous Microorganisms and Lipid Production SCOs could conventionally be produced by oleaginous microorganisms, i.e., microbes with lipid contents of more than 20% dry cell weight (DCW). During this process, different organic carbon sources (e.g., glucose and other hexoses, disaccharides, glycerol, and polysaccharides) could be assimilated. Compared to oil crops, two main advantages of the oleaginous microorganism application as the promising oil-rich feedstock include (1) year-round culturing and oil production and (2) higher biomass productivities with a greater lipid-to-biomass ratio than food oil crops. Moreover, it is possible to convert cheap agro-industrial wastes into SCOs that structurally and compositionally are similar to several high-value fats using these microorganisms (Kazemi Shariat Panahi et al. 2019d; Shuba and Kifle 2018; Sitepu et al. 2019). To achieve this goal, the first step should be a careful screening and identification of the microorganisms with significant lipid production capabilities. Different types of microorganisms, such as microalgae (e.g., Botryococcus braunii, Cylindrotheca sp., Nitzschia sp., and Schizochytrium sp.), bacteria (e.g., Arthrobacter sp., Acinetobacter calcoaceticus, Rhodococcus opacus, and Bacillus alcalophilus), fungi (e.g., Aspergillus oryzae, Aspergillus niger, Thamnidium elegans, Mucor sp., Colletotrichum sp., Alternaria sp., Mortierella ramanianna, Epicoccum purpurascens, Chaetomium globosum, Cunninghamella echinulate, Cunninghamella bainieri, Zygorhynchus moelleri, Mortierella alpine, Mortierella isabellina, and Mortierella vinacea), and yeasts (e.g., Candida curvata, Yarrowia lipolytica, Cryptococcus albidus, Humicola lanuginosa, Trichosporon fermentans, Trichosporon dermatis, Trichosporon cutaneum, Kodamaea ohmeri, Metschnikowia pulcherrima, Lipomyces starkeyi, Rhodosporidium toruloides, Rhodosporidium kratochvilovae, and Rhodotorula glutinis) have been identified as the potential oleaginous microorganisms for biodiesel production. Respectively, Tables 3.1 and 3.2 represent different oleaginous fungi and yeasts, their substrates (i.e., carbon sources), and their oil production efficiency. Typically, unlike non-oleaginous microorganisms, the lipid biosynthesis pathways in oleaginous microorganisms have been equipped with certain key enzymes (Papanikolaou and Aggelis 2011a). The energy sources could be obtained from organic carbon sources (e.g., sugars, fatty materials, glycerol, etc.) or CO2 sequestration (in the presence of sunlight) by heterotrophic or autotrophic photosynthetic oleaginous microorganisms, respectively (Bellou et  al. 2014; Kazemi Shariat Panahi et  al. 2019a; Ratledge 1988;

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M. Tabatabaei et al.

Table 3.1  Some oleaginous fungi, their energy source (i.e., hydrophilic and hydrophobic carbon substrates), and their oil production efficiency

Fungi Carbon source Aspergillus niger Glycerol (crude)

8.2

41.4

Experiment level References Flasks André et al. (2010) Flasks André et al. (2010) Flasks Papanikolaou et al. (2011) Flasks Papanikolaou et al. (2011) Flasks Van der Merwe et al. (2005)

49.5

Waste cooking olive oil 17.9

22

12.4

15

8.7

54

Flasks

Waste cooking olive oil 16.2

27.2

Flasks

Crude glycerol

16.3

71.1

Flasks

Glucose and xylose (50–50% w/w) Fructose (commercial)

12.6

45.9

Flasks

12.5

70.4

Flasks

Glucose (commercial)

31.9

47.1

Flasks

30.1

46.1

13.3

70.0

Batch bioreactor Flasks

14.9

34.5

Flasks

Sunflower oil and glucose Blackstrap molasses

18

53

Flasks

12.1

31.4

Flasks

Fructose (commercial)

16.7

21.5

Flasks

Glucose (commercial)

13.6

69.5

Flasks

Glucose/tomato waste hydrolysate

32.6

24.9

Flasks

20.3

42.1

Flasks

7.8

25.6

Flasks

Edible-oil-containing waste (sunflower oil, ~80% w/w) Aspergillus flavus Soybean oil

Saccharose (commercial) Xylose (pure)

Cunninghamella echinulata

Oil contents (wt.%) 57.4

Waste cooking olive oil 14.4

Aspergillus fumigatus

Penicillium expansum Thamnidium elegans

Dry cell weight (g/L) 5.4

Glycerol (crude)

Koritala et al. (1987) Papanikolaou et al. (2011) Chatzifragkou et al. (2011) Zikou et al. (2013) Papanikolaou et al. (2010) Zikou et al. (2013) Zikou et al. (2013) André et al. (2010) Zikou et al. (2013) Čertík et al. (1997) Chatzifragkou et al. (2010) Chatzifragkou et al. (2010) Chatzifragkou et al. (2010) Fakas et al. (2007) Fakas et al. (2008) Fakas et al. (2009b) (continued)

3  Fungi as Bioreactors for Biodiesel Production

43

Table 3.1 (continued)

Fungi

Cunninghamella bainieri

Epicoccum purpurascens Mortierella ramanniana

Mortierella isabellina

Carbon source Glycerol (pure)

Dry cell weight (g/L) 6.9

Oil contents (wt.%) 25.1

4.2

15.4

Experiment level References Flasks Bellou et al. (2012) Batch Bellou et al. bioreactor (2012) Flasks Fakas et al. (2008) Flasks Chen and Liu (1997) Flasks Fakas et al. (2009a) Flasks Van der Merwe et al. (2005)

Glycerol/tomato waste hydrolysate Starch

18.1

27.4

26.1

29.3

Xylose (pure)

12.5

53.6

Edible-oil-containing waste (sunflower oil, ~80% w/w) Sunflower oil and glucose Glucose (pure)

24.9

24

18.3

58

Flasks

12.4

31.9

11.9

31.9

Molasses

33.5

80.0

Batch bioreactor Batch bioreactor Flasks

Glycerol (pure)

7.0

53.1

Flasks

9.7

32.7

Blackstrap molasses

9.5

53.7

Batch bioreactor Flasks

Corn fiber hydrolysate

39.2

54.6

Flasks

Corn stover acid or alkali hydrolysate

14.1

34.5

Flasks

10.9

29.5

Flasks

12.8

24.8

Flasks

17.2

29.7

Flasks

18.7

37.0

13.8

21.2

Batch bioreactor Flasks

Giant reed hydrolysate

Čertík et al. (1997) Saad et al. (2014) Saad et al. (2014) Koutb and Morsy (2011) Bellou et al. (2012) Bellou et al. (2012) Chatzifragkou et al. (2010) Xing et al. (2012) Ruan et al. (2012) Ruan et al. (2012) Ruan et al. (2013) Ruan et al. (2014) Ruan et al. (2014) Ruan et al. (2013) (continued)

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M. Tabatabaei et al.

Table 3.1 (continued)

Fungi

Carbon source Glucose (commercial)

Dry cell weight (g/L) 13.2

Oil contents (wt.%) 74.9

22.3

45.0

17.8

72 25.3

Experiment level References Flasks Chatzifragkou et al. (2010) Flasks Ruan et al. (2012) Batch Chatzifragkou bioreactor et al. (2010) Flasks Vamvakaki et al. (2010) Flasks Ruan et al. (2013) Flasks Fakas et al. (2009b) Flasks Papanikolaou et al. (2008) Flasks Economou et al. (2011) Flasks Harde et al. (2016) Flasks Harde et al. (2016) Flasks Papanikolaou et al. (2017) Flasks Ruan et al. (2013) Flasks Ruan et al. (2012) Flasks Ruan et al. (2012) Flasks Papanikolaou et al. (2004) Flasks Ruan et al. (2013) Flasks Ruan et al. (2015)

Lactose-enriched 32.0 cheese whey Miscanthus hydrolysate 12.3

32.2

Glycerol (crude)

6.2

53.2

8.5

51.7

Rice hull hydrolysate

3.6

64.3

Glucose and xylose (50–50% w/w) Xylose

18.8

50.5

15.3

66.0

Glucose (commercial)

10.2

83.3

Switch grass hydrolysate Synthetic acid hydrolysate (glucose and xylose)

12.6

35.6

12.6

38.4

12.24

30.2

Glucose (pure)

35.9

50.4

Synthetic hydrolysate (glucose and xylose) Synthetic lignocellulosic hydrolysate Xylose(commercial)

10.4

30.7

11.0

37.5

11.4

43.0

Flasks

Xylose (pure)

8.7

65.5

Flasks

Glycerol (crude)

8.1

66.7

Flasks

Ruan et al. (2012) Fakas et al. (2009a) Papanikolaou et al. (2017) (continued)

3  Fungi as Bioreactors for Biodiesel Production

45

Table 3.1 (continued)

Fungi Mortierella ramanniana

Mortierella alpina

Mucor circinelloides

Mucor sp.

Zygorhynchus moelleri Chaetomium globosum

Colletotrichum sp.

Carbon source Xylose

Dry cell weight (g/L) 7.8

Oil contents (wt.%) 15.4

Glycerol (crude)

9.3

42.0

Glycerol (crude)

15.6

33.3

Glycerol (crude)

20.5

31.9

Glycerol (pure)

28.6

33.4

Glycerol (pure)

26.7

35.4

Linseed oil

n.d.

56.0

Olive oil and glucose

39.9

65

Sesame oil

12.9

40

Molasses

n.d.

19.8

Canola oil

9.6

36.6

Starch

28.0

17.8

Olive oil

24.1

54

Sunflower oil

5.4

36.8

Sunflower oil and glucose Sunflower oil and glucose Glycerol (crude)

19.4

62

19.8

60

3.7

42.4

Glucose

18.6

25.0

Glucose and xylose (50–50% w/w) Xylose

16.0

54.4

7.4

29.7

Experiment level References Flasks Hansson and Dostálek (1986) Flasks Papanikolaou et al. (2017) Flasks Dedyukhina et al. (2014) Flasks Dedyukhina et al. (2014) Flasks Dedyukhina et al. (2012) Flasks Dedyukhina et al. (2012) Flasks Shimizu et al. (1989) Flasks Szczęsna-­ Antczak et al. (2006) Flasks Tauk-Tornisielo et al. (2009) Flasks Carvalho et al. (2018) Flasks Tauk-Tornisielo et al. (2009) Flasks Ahmed et al. (2006) Flasks Szczęsna-­ Antczak et al. (2006) Flasks Aggelis et al. (1995) Flasks Čertík et al. (1997) Flasks Čertík et al. (1997) Flasks Chatzifragkou et al. (2011) Flasks Harde et al. (2016) Flasks Harde et al. (2016) Flasks Dey et al. (2011) (continued)

46

M. Tabatabaei et al.

Table 3.1 (continued) Dry cell weight (g/L) 10.4

Oil contents (wt.%) 41.3

Experiment level References Flasks Dey et al. (2011) Flasks Van der Merwe et al. (2005)

Fungi Alternaria sp.

Carbon source Xylose

Curvularia lunata

Edible-oil-containing waste (sunflower oil ~80% w/w) Edible-oil-containing waste (sunflower oil ~80% w/w) Evening primrose oil

14.4

33

23.8

45.0

Flasks

Van der Merwe et al. (2005)

10.6

40.0

Flasks

Sunflower oil and glucose Sunflower oil and glucose Fried soybean oil

18.2

43

Flasks

18.9

63

Flasks

10.1

28.4

Flasks

Canola oil

10

40.6

Flasks

Palm oil

11.9

54.3

Flasks

Aggelis et al. (1997) Čertík et al. (1997) Čertík et al. (1997) Tauk-Tornisielo et al. (2009) Tauk-Tornisielo et al. (2009) Tauk-Tornisielo et al. (2009)

Emericella nidulans Langermannia gigantea Rhizopus microscopus Rhizopus stolonifer Rhizopus sp.

Updated from Athenaki et al. (2018)

Sharma et al. 2001). The organic carbon sources that are assimilated by heterotrophic oleaginous microorganisms could be hydrophilic (e.g., glucose, other hexoses, disaccharides, glycerol, polysaccharides, acetic acid, butyric acid, and ethanol), hydrophobic (e.g., free fatty acids (FFAs), TGs, and n-alkanes), or their mixtures (e.g., blends of glucose or glycerol with FFAs or TGs). Techno-economic outcomes for biodiesel production could significantly be improved by avoiding feedstock drying and oil extraction procedures. On this context, second-generation biodiesel (i.e., advanced biodiesel) could be produced from oleaginous microorganisms. This approach was used for biodiesel production through one-step acid-catalyzed transesterification of oil extracted from Yarrowia lipolytica biomass grown on waste cooking oil (lipid productivity, 0.042  g/L/h) (Katre et al. 2018). Under optimized conditions (catalyst, 0.2 M H2SO4 1.0 mL/g; methanol-to-chloroform-to-biomass ratio, 10:1:4 v/v/w; 50 °C; and 8 h), a biodiesel yield of 22 g/g biomass was achieved. The fatty acid methyl ester profile displayed desirable amounts of saturated (32.81%), monounsaturated (36.41%), and polyunsaturated (30.59%) methyl esters (Katre et al. 2018). One of the newly developed biodiesel production processes is direct transesterification of wet biomass feedstock, which can reduce the costs. Direct transesterification of oleaginous microorganisms significantly reduces both production costs (shorter processing time and wet biomass conversion) and environmental burdens (no requirement for toxic

3  Fungi as Bioreactors for Biodiesel Production

47

Table 3.2  Some oleaginous yeasts, their energy sources (i.e., hydrophilic and hydrophobic carbon sources), and their oil production efficiency

Yeasts Aureobasidium melanogenum Rhodotorula colostri Sporidiobolus ruineniae Sporobolomyces carnicolor Trichosporon fermentans

Trichosporon sp.

Lipomyces starkeyi

Lipomyces tetrasporus Trichosporon dermatis Trichosporon cutaneum

Yarrowia lipolytica

Carbon source Glucose

Dry cell weight (g/L) n.d.a

Oil contents (wt.%) 66.3

Experiment level Flasks

Glucose

n.d.

62.9

Flasks

Glucose

n.d.

66.0

Flasks

Glucose

n.d.

61.4

Flasks

Bagasse acid hydrolysate Glucose and xylose blend Molasses

29.8

39.9

Flasks

38.9

61.7

Flasks

36.4

35.3

Flasks

28.8

40

Flasks

25.4

61

Batch bioreactor

25.8

52

Flasks

Gong et al. (2012)

153

54

9.4

68

Fed-batch bioreactor Flasks

Rice straw acid hydrolysate Methyl-stearate and methyl-palmitate Cellobiose, xylose, and glucose Glucose

References Wang et al. (2014) Garay et al. (2016) Garay et al. (2016) Garay et al. (2016) Huang et al. (2012b) Huang et al. (2012c) Zhu et al. (2008) Huang et al. (2009) Matsuo et al. (1981)

Glucose and sewage sludge Glucose and xylose blend Glycerol (crude)

20.5

61.5

Flasks

18.2

37

Flasks

Glucose

n.d.

66

Flasks

Corncob hydrolysate Corn stover hydrolysate Glucose and xylose blend Xylose Borage oil

60.9

40.1

Flasks

49.2

39.2

Flasks

Yamauchi et al. (1983) Angerbauer et al. (2008) Zhao et al. (2008) Tchakouteu et al. (2015) Sitepu et al. (2014) Huang et al. (2012a) Hu et al. (2011)

47.9

49.7

Flasks

Hu et al. (2011)

45.6 22.6

46.5 51.6

Flasks Flasks

Echium oil

26.1

56.9

Flasks

Hu et al. (2011) Saygün et al. (2014) Saygün et al. (2014) (continued)

48

M. Tabatabaei et al.

Table 3.2 (continued)

Yeasts

Dry cell weight (g/L) 8.7

Oil contents (wt.%) 44

Experiment level Flasks

Carbon source Industrial fats blend Defatted rice bran hydrolysate Glucose

22.5

48

Flasks

9.2

25

Continuous bioreactor

Stearin

15.2

52

Flasks

Stearin

12.5

54

Flasks

Stearin

15.1

48

Stearin and hydrolyzed rapeseed oil Stearin and hydrolyzed rapeseed oil Stearin and glycerol (crude) Trout oil

8

36

Batch bioreactor Flasks

4.2

46

Batch bioreactor

11.4

30

Flasks

20.4

53.8

Flasks

9

37.7

Flasks

8.4

45.9

Flasks

4

88

Flasks

7.6

57.9

Flasks

8

34

Flasks

3.2

30.7

Flasks

3.9

22.1

Flasks

11.6

31

Flasks

24.7

23.5

Repeated-­ batch bioreactor

Chicken fat and glucose Waste oil from frying fish and glucose Waste cooking oil Waste oil from frying vegetables and glucose Meat fat and glucose Glycerol (crude)

References Papanikolaou et al. (2001) Tsigie et al. (2012) Aggelis and Komaitis (1999) Papanikolaou et al. (2007) Papanikolaou et al. (2001) Papanikolaou et al. (2002) Papanikolaou et al. (2001) Papanikolaou and Aggelis (2003) Papanikolaou et al. (2003) Saygün et al. (2014) El Bialy et al. (2011) El Bialy et al. (2011) Katre et al. (2018) El Bialy et al. (2011) El Bialy et al. (2011) Papanikolaou et al. (2013) Poli et al. (2014) Sestric et al. (2014) Bellou et al. (2016b) (continued)

3  Fungi as Bioreactors for Biodiesel Production

49

Table 3.2 (continued)

Yeasts

Carbon source

Glycerol (pure)

Candida sp.

Dry cell weight (g/L) 49.1

Oil contents (wt.%) 46

6.3

26.4

6.5

31

8.1

43

4.7

23.1

42.2

38.2

42.1

30.9

5.5

50.8

41

34.6

Fed-batch bioreactor

Fontanille et al. (2012) Tsigie et al. (2011) Najjar et al. (2011) Saygün et al. (2014) Xiong et al. (2015) Saygün et al. (2014) Saygün et al. (2014) Gill et al. (1977) Duarte et al. (2013)

Experiment level Fed-batch bioreactor Repeated-­ batch bioreactor Flasks Single stage continuous bioreactor Repeated-­ batch bioreactor Fed-batch bioreactor Fed-batch bioreactor Batch bioreactor

Glycerol (crude) and decanter effluent from palm oil blend Glycerol (pure) and volatile fatty acids blend Sugarcane bagasse hydrolysate Olive oil

19.5

58.5

Flasks

9.7

37.8

Flasks

Linseed oil

26.7

61.7

Flasks

Mutton fat

15

33.1

Flasks

Rapeseed oil

25.1

48

Flasks

Sesame oil

23.9

47.3

Flasks

Glucose

18.1

37.1

Glycerol (crude)

19.7

50.2

Continuous bioreactor Flasks

References Rakicka et al. (2015) Mirończuk et al. (2014) André et al. (2009) Papanikolaou and Aggelis (2002) Makri et al. (2010) Fontanille et al. (2012) Celińska and Grajek (2013) Louhasakul and Cheirsilp (2013)

(continued)

50

M. Tabatabaei et al.

Table 3.2 (continued)

Yeasts Candida lipolytica

Carbon source Corn oil Evening primrose oil Soap-stocks

Candida guilliermondii Rhodosporidium toruloides

Dry cell weight (g/L) 29.2

Oil contents (wt.%) 62

7.8

21

Experiment level Batch bioreactor Flasks

n.d.

33.5

Flasks

Methyl-stearate and methyl-palmitate Glucose

24.8

35

Batch bioreactor

106.5

67.5

Glycerol (crude)

16.2

54.3

Fed-batch bioreactor Flasks

19.3 26.6

47.7 69.5

35.3

46

24.9

48.9

15.1

57

27.9

29

26.5 8.7

37.8 61.8

12.5

42.9

37.2

64.5

9.8 5.8 11.7

Glucose

Glucose and pure stearic acid blend Glycerol (pure) Pure stearic acid

Flasks Batch bioreactor Batch bioreactor Flasks Batch bioreactor Flasks

References Bati et al. (1984) Aggelis et al. (1997) Montet et al. (1985) Matsuo et al. (1981) Li et al. (2007) Tchakouteu et al. (2015) Xu et al. (2012) Xu et al. (2012) Kiran et al. (2013) Yang et al. (2014) Bommareddy et al. (2015) Leiva-Candia et al. (2015) Xu et al. (2017) Shen et al. (2013) Moreton (1988)

47

Flasks Continuous bioreactor Batch bioreactor Fed-batch bioreactor Flasks

Tchakouteu et al. (2017) Gierhart (1984)

34.6 35

Flasks Flasks

Moreton (1988) Gierhart (1984) (continued)

3  Fungi as Bioreactors for Biodiesel Production

51

Table 3.2 (continued)

Yeasts Rhodotorula glutinis

Rhodotorula mucilaginosa Rhodosporidium kratochvilovae

Rhodosporidium fluviale

Rhodosporidium sphaerocarpum Rhodosporidium babjevae Cryptococcus curvatus

Dry cell weight (g/L) 4.9

Oil contents (wt.%) 39

Experiment level Flasks

5.5

35.2

Flasks

44.8

62.1

30.8

53

16.8

34.6

25

20

Fed-batch bioreactor Fed-batch bioreactor Fed-batch bioreactor Flasks

n.d. 13.9

47.9– 52.9 61.7

Batch or fed-batch Flasks

8.9

53.2

Flasks

15.1

55.6

Flasks

14.1

63.8

23

69.5

Batch bioreactor Flasks

Glucose

n.d.

74.1

Flasks

Glucose

n.d.

62.4

Flasks

Glycerol (crude)

118

25

32.9

52.9

22

49

50.4

37.7

Fed-batch bioreactor Fed-batch bioreactor Fed-batch bioreactor Flasks

18.0

31

Continuous bioreactor

63.3

Flasks

Carbon source Glucose and xylose blend Glycerol (crude)

Monosodium glutamate wastewater Cassava starch hydrolysate Pulp and paper industry effluent Aqueous extract of Cassia fistula Aqueous extract of Cannabis sativa Glycerol (crude)

Glycerol (pure) and spent yeast lysate blend Lactose

Cryptococcus aerius

Glucose

References Zhang et al. (2011) Saenge et al. (2011) Yen et al. (2015) Karamerou et al. (2017) Karamerou et al. (2016) Xue et al. (2008) Li et al. (2010) Patel et al. (2017) Patel et al. (2015) Patel et al. (2014) Polburee et al. (2016) Polburee et al. (2016) Garay et al. (2016) Garay et al. (2016) Meesters et al. (1996) Liang et al. (2010) Cui et al. (2012) Ryu et al. (2013) Evans and Ratledge (1983) Sitepu et al. (2014) (continued)

52

M. Tabatabaei et al.

Table 3.2 (continued)

Yeasts Cryptococcus freyschussii Cryptococcus albidus Cryptococcus psychrotolerans Metschnikowia pulcherrima Kodamaea ohmeri

Trichosporanoides spathulata

Apiotrichum curvatum

Carbon source Glycerol (crude)

Dry cell weight (g/L) 30.5

Oil contents (wt.%) 29.8

Glycerol (pure)

1.4

43.8

Experiment level Fed-batch bioreactor Flasks

Groundnut shell acid hydrolysate Glycerol

n.d.

46

Flasks

7.4

40

Flasks

Glycerol (crude)

10.3

53.3

Flasks

Glycerol (crude)

10.5 17.1

30.4 43.4

Flasks Flasks

13.8

56.4

Fed-batch bioreactor

20

36.1

Continuous bioreactor

Whey

References Raimondi et al. (2014) Hansson and Dostálek (1986) Deeba et al. (2017) Santamauro et al. (2014) Kitcha and Cheirsilp (2013) Kitcha (2012) Kitcha and Cheirsilp (2013) Kitcha and Cheirsilp (2013) Ykema et al. (1988)

Updated from Athenaki et al. (2018) a Not determined

solvents) (Kumar 2017; Sitepu et al. 2019; Yousuf et al. 2017). For instance, compared to the conventional ex situ multistep transesterification method, direct wet Pichia guilliermondii biomass transesterification could reduce the processing time up to 7 h (Chopra et al. 2016). Moreover, microwave and ultrasonication processes could economically be used for in situ extraction of oil from biomass or intensified (higher mass transfer) the transesterification reaction (Tabatabaei et al. 2019a). For example, through this strategy, a 4-min process delivered conversion efficiencies of 92% and 94.3% from direct transesterification of wet Cryptococcus curvatus and Yarrowia lipolytica biomasses, respectively (Yellapu et al. 2017). It should be noted that a vast majority of direct transesterification of fungal biomass have been performed at temperatures of 60–100  °C, consuming significant energy. Therefore, some technologies such as vortex microfluidic platform or high shear turbo thin film device must be coupled to direct transesterification of wet oleaginous fungal biomass to improve its economic feasibility in terms of residence time and reaction temperature. For example, the latter technology has been used for continuous conversion of Mucor plumbeus biomass into biodiesel (conversion efficiency, >90%; catalyst, 3  wt.% NaOH; room temperature; water content, 50%;

3  Fungi as Bioreactors for Biodiesel Production

53

atmospheric pressure; and residence time, ~2  min) and saved up to 94% energy consumption, compared to conventional direct transesterification system (Sitepu et al. 2019). Overall, some advantages of biodiesel feedstock (i.e., SCOs) production using fungal and yeast systems include fast growth, high lipid production efficiency, reduced land requirement, no threat to food security (i.e., no food vs. fuel issue), short oil production period, and their oil similarity (i.e., in terms of fatty acid composition) to vegetable oils (i.e., containing 16–18 carbons) (Amara et  al. 2016; Athenaki et al. 2018; Cho and Park 2018; Xu et al. 2015). However, novel biodiesel production technologies must be developed to further improve the economic and environmental feasibilities of biodiesel production through direct transesterification of wet oleaginous microorganisms. Figure 3.1 shows the simplified process of biodiesel production from lipids produced by fungi and yeasts.

3.3  Fungi as Bioreactors for Lipid Production During past 30 years, more than 64 different fungal species have been identified that could accumulate high lipid content (>25% w/DCW) in their complex cellular organizations (Table 3.1). Commonly, fungal lipids are less saturated than those accumulated by yeasts. More specifically, fungi can produce different types of polyunsaturated fatty acids (PUFAs), such as arachidonic acid, c-linolenic acid, dihomo-γ-linolenic acid, docosahexaenoic acid, and eicosapentanoıc acid (Athenaki et  al. 2018; Bellou et  al. 2016a; Bharathiraja et  al. 2017). Some fungi (e.g., Zygomycetes), especially those related to Mucoromycota, are potential producers of c-linolenic-acid-containing SCOs, which have crucial dietary and significant pharmaceutical applications. For instance, Mortierella isabellina, one of the most important oleaginous Zygomycetes, could produce and accumulate lipids up to 84% w/DCW, following its cultivation on several agro-industrial wastes consisting of glucose, molasses, fructose, starch, cheese whey, glycerol, and lignocelluloses (e.g., corn stover, corn cob, etc.) (Table 3.1). Mortierella alpina is another important oleaginous Zygomycetes (subdivision Mucoromycota) that could be used in biodiesel industry due to its ability for arachidonic acid production. Kosa et al. (2018) used a reproducible and high-throughput method (i.e., microplate combined with FTIR spectroscopy) for screening the potent (i.e., quality and yield) omega-6-PUFA-­ producing fungi for biodiesel feedstock production. Accordingly, Mortierella humilis, Cunninghamella blakesleeana, and Umbelopsis vinacea showed the highest lipid production concentration (i.e., 7.0–8.3 g/L), whereas Absidia glauca accumulated the highest total lipid content (47.2% of biomass). Lipid containing high concentrations of saturated (Cn: 0) and monounsaturated (Cn: 1) fatty acids could be produced from different agro-industrial residues (e.g., sugarcane molasses, sugarcane bagasse hydrolysate, corn milling residues, and glycerol) using Mucor circinelloides URM 4182 (Carvalho et al. 2018). The lipid

Fig. 3.1  The simplified processes for microbial lipids and subsequent biodiesel productions

54 M. Tabatabaei et al.

3  Fungi as Bioreactors for Biodiesel Production

55

concentrations could be improved from 19.8 to 33.2% by replacing sucrose-based substrate with glucose. The produced SCOs (wet biomass) could be directly transesterified (solid catalyst, 10  wt.% 12-molybdophosphoric acid supported on alumina; ethanol-to-oil ratio, 120:1; 200 °C; and 4 h) into biodiesel (conversion yield, 98.5%) in a high-pressure reactor.

3.4  Yeasts as Bioreactors for Lipid Production Some yeast species can also produce and accumulate lipids up to 50–75% w/DCW (Table 3.2). More specifically, these oleaginous yeasts accumulate lipids up to 50% w/DCW under normal environmental conditions, whereas some environmental stresses (e.g., nutrition limitation) could improve their lipid content. For instance, the oleaginous yeasts Rhodosporidium spp. (i.e., R. sphaerocarpum, R. toruloides, and R. fluvialis) produce lipids up to 70% w/DCW in nitrogen-limited medium. The biosynthesized lipids consist of 80–90% TGs and a small fraction of sterol esters (SEs) (Athenaki et al. 2018; Xue et al. 2018). Additionally, both cheap hydrophilic (e.g., molasses, bagasse, straw, corn stover, glycerol, corncob, and xylose) and hydrophobic carbon sources could be efficiently converted into lipids by oleaginous yeast. Some examples of the hydrophobic carbon sources (edible, non-edible, and waste oils) that have already been transformed into OSCs by yeasts include fried soybean oil, canola oil, sunflower oil, edible-oil-containing waste, methyl-stearate and methyl-palmitate, stearin, rapeseed oil, industrial fats, chicken fat, waste oil from frying fish, waste oil from frying vegetables, meat fat, palm oil, olive oil, sesame oil, linseed oil, trout oil, borage oil, echium oil, mutton fat, corn oil, and evening primrose oil (Table 3.1). The most common oleaginous yeast genera are Yarrowia, Candida, Rhodotorula, Rhodosporidium, Aureobasidium, Cryptococcus, Metschnikowia, Kodamaea, Trichosporon, Sporidiobolus, Sporobolomyces, Occultifur, Starmerella, Pichia, and Lipomyces (Maina et  al. 2017; Papanikolaou and Aggelis 2011a, b; Santamauro et al. 2014; Saygün et al. 2014). Many red yeast species such as Rhodotorula mucilaginosa and Rhodosporidium toruloides synthesize and accumulate high lipid concentrations in their cells (47.9% and 70% w/DWC, respectively). However, both of them only grow on glucose, sucrose, maltose, cellobiose, D-xylose, and glycerol and do not grow on lactose and soluble starch (Kurtzman et al. 2011; Li et al. 2007, 2010; Xue et al. 2018). TGs and SEs are known as the main lipid classes produced by oleaginous yeasts. Yeasts have a short duplication time (20 g/L), and high lipid content (>50% w/DWC). In addition, simple genetic improvement and convenient large-scale fermentation further increase the feasibility of biodiesel production form yeast lipids as an efficient alternative for oil plants (Fakas et al. 2009b; Papanikolaou and Aggelis 2011b; Sitepu et al. 2014; Xue et al. 2018). For example, in 2015, the lipid production capabilities in some oleaginous yeasts (e.g., Yarrowia lipolytica) were improved by genetic engineering

56

M. Tabatabaei et al.

techniques (Zhu and Jackson 2015). More specifically, the key genes involved in the lipid biosynthesis pathways (e.g., inulinase gene, PYC gene encoding a pyruvate carboxylase, and endogenous ACL1 gene encoding ATP citrate lyase) were overexpressed. Alternatively, the genes (e.g., MIG gene encoding the main glucose repressor Mig1) that inhibit lipid biosynthesis could be silenced. Overall, compared to fungi, oleaginous yeasts could more easily be cultured in large-scale bioreactors due to their unicellular morphology, allowing transformation of different cheap carbon sources to SCOs (Table 3.2).

3.5  B  iodiesel Production Catalyzed by Fungal and Yeast Lipases Currently, the alkali-catalyzed transesterification technologies are used for commercial-­scale biodiesel production (Tabatabaei et al. 2019a). For this purpose, plant oils or animal fat must meet some quality standards including low water and FFA contents (15 wt.%) (Tabatabaei et al. 2019a; Rahimzadeh et al. 2018). The incorporation of this FFA pretreatment step improves the quality of cheap oil feedstocks by reducing the acidity (i.e., free fatty acid content) prior to alkaline transesterification (Tabatabaei et  al. 2019a; Rahimzadeh et  al. 2018). However, it is obvious that this reduction in the price of feedstock is in the expense of an increase in conversion cost. A more promising alternative could be the application of microbial lipases (i.e., as biocatalyst) in biodiesel production process to overcome the abovementioned problems in the alkaline transesterification reactions. More specifically, lipases catalyze the hydrolysis of esters, particularly long-chain TGs, generating free fatty acids, mono- and di-glycerides, and glycerol. The catalysis capabilities of lipases could be exploited in different biotechnological applications, including esterification, alcoholysis, aminolysis, acidolysis, and interesterification reactions in non-­ aqueous media (Aguieiras et al. 2015; Mohammadipanah et al. 2015). Moreover, lipases could be applied in transesterification reaction for biodiesel production in

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either immobilized or soluble free form, with the first attempt tried in 1990 (Tabatabaei et al. 2019a; Mittelbach 2015). This ecofriendly method allows direct biodiesel production via transesterification of low cost (i.e., quality) feedstock such as waste cooking oils with high FFA and water content under mild conditions (i.e., low energy consumption)(Tabatabaei et al. 2019a). In better word, there would be no requirements for feedstock pretreatment and catalyst removal in enzymatic catalysis process (Tabatabaei et al. 2019a). Despite their relatively high prices, the mentioned advantages of enzymatic catalysts have favored their industrial application in transesterification reaction by some companies such as Sunho Biodiesel Corporation (Taiwan), Piedmont Biofuel (USA), Lvming Co. Ltd. (China), and Hainabaichuan Co. Ltd. (Tabatabaei et al. 2019a). Commonly, lipases are extracted from biological sources, for example, yeasts and filamentous fungi. The most widely used lipases in biodiesel industry are those extracted from yeasts (e.g., Candida antarctica lipase B and Candida rugosa) and filamentous fungi (e.g., Rhizomucor miehei, Rhizopus oryzae, Thermomyces lanuginosus, Aspergillus niger, and Penicillium expansum) (Aarthy et al. 2014; Canet et  al. 2014; Contesini et  al. 2010; Tabatabaei et  al. 2019a; Li and Zong 2010; Rodrigues and Fernandez-Lafuente 2010; Sharma et  al. 2001; Tan et  al. 2010). These enzymes are commercially available in different forms, such as free (i.e., liquid formulations) and immobilized, whole cells (i.e., mixture of microorganism cells and enzymes), and solid enzymatic preparation using solid-state fermentation (Aguieiras et al. 2015; Tabatabaei et al. 2019a). Different types of plant oils (e.g., sunflower, soybean, rapeseed, olive, cotton, and palm), non-edible plant oils (e.g., jatropha, macauba, andiroba, babassu, castor bean, karanja, mahua, and crambe), and oil waste (such as dewaxed/degummed rice bran oil, waste cooking palm oil, soybean oil deodorizer distillate, degummed soybean oil, and recycled restaurant greases) have been used as the substrates for biodiesel production (yield, 70–95%) via lipase-catalyzed transesterification reaction (Aguieiras et  al. 2015; Carvalho et  al. 2013; Tabatabaei et  al. 2019a; Hama and Kondo 2013; Hernández-Martín and Otero 2008; Hsu et al. 2004; Jang et al. 2012; Kojima et al. 2004; Köse et al. 2002; Lai et al. 2005; Talukder et al. 2009; Valero 2012; Wang et al. 2006; Zheng et al. 2009). For instance, Nelson et al. (1996) investigated alcoholysis with various lipases in hexane and reported a methyl ester yield of 77.8% in methanolysis with lipase from Mucor miehei. In another study methyl esters were synthesized using Candida antarctica lipase as catalyst in a solvent-free system (Shimada et al. 1999). Overall, the lipase-catalyzed transesterification for biodiesel production is in the early stages, and its scale up faces many challenges, including high enzyme cost, the longer reaction time, and lipase inhibition by ethanol/methanol (Hama et al. 2018). However, it is worth it to solve these constraints as enzymatic transesterification has some superb advantages such as feasibilities of using low-grade oils as substrates, no soap formation, lower energy consumption, and easier downstream process requirement, compared to chemical approach.

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3.6  Conclusions The application of oleaginous fungi and yeasts as the source for lipid and/or lipase for biodiesel production is fascinating. Typically, SCO production by these fast-­ growing microorganisms delivers high lipid production rate and efficiency. Moreover, biodiesel feedstock production through this system provides oils resembling to those with 16–18 carbons from vegetable origin while having lower biodiesel tradeoffs and higher feedstock sustainability, i.e., lower land-use change and no food vs. fuel debate. On the downside, the production cost of SCOs is currently higher than that of traditional oil plants and animal fats. The main reasons could be necessity of aseptic conditions, high cost of the cultivation substrates, and high capital cost due to expensive fermentation equipment. Among them, substrate cost alone could contribute to as high as 40–80% of overall biodiesel production cost. Therefore, the economic feasibilities of SCO-based biodiesel production could be enhanced through assimilation of hydrophilic and hydrophobic organic wastes. However, commonly the chemical compositions of organic waste are not suitable for biomass and oil production by all oleaginous microorganisms. This highlights the need for isolation and identification of novel oleaginous yeasts and fungi, genetic modification of existing oleaginous yeasts and fungi, and/or fermentation optimization studies to efficiently convert organics waste to lipid carbon sources for each oleaginous fungus or yeast. Moreover, techno-economic outcomes as well as environmental feasibility of SCO-based biodiesel production could significantly be improved by further developing novel biodiesel production technologies. Some solutions may include direct transesterification of oleaginous microorganisms, in situ oil extraction and mass transfer intensified processes (e.g., microwave and ultrasonication), vortex microfluidic platform, high shear turbo thin film device, etc. Alternatively, oleaginous fungi and yeasts could be exploited as a good biological source for lipase production. The application of this biocatalyst in biodiesel production via transesterification have several advantages ranging from cheaper oil feedstock requirements to more simple downstream purification steps. Consequently, some companies such as Sunho Biodiesel Corporation, Piedmont Biofuel, Lvming Co. Ltd., and Hainabaichuan Co. Ltd., have developed their own enzymatic catalyzed biodiesel production despite the relatively high price of enzymes (Tabatabaei et al. 2019a). The production of cheaper and extremophile lipases via genetic engineering technologies or by screening new strains and development of new lipase-­based catalysts may further adsorb more commercial attentions on lipases for biodiesel production.

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Chapter 4

Fungal Biocontrol Agents as a New Source for Bioethanol Production Hamed Kazemi Shariat Panahi, Mona Dehhaghi, Gholamreza Salehi Jouzani, Rasoul Zare, Mortaza Aghbashlo, and Meisam Tabatabaei

4.1  Introduction Growing air pollution is known as a major threat to the environment and public health. It has been reported that air pollution will kill 4.5 million people each year by 2040 (Kazemi Shariat Panahi et al. 2019c). Toxic gaseous emissions from various industrial factories and the transportation sector are the main sources of air pollution (Hosseinzadeh-Bandbafha et al. 2018; Rajaeifar et al. 2019). On the other hand, limited supply of fossil fuels together with the world’s population growth has led to energy insecurity as well as global warming (due to greenhouse gas emissions) (Soltanian et al. 2019). A promising solution is the replacement of fossil fuels

We dedicate this chapter to the memories of our good friend and great mycologist Prof. Walter Gams who spent his life to present a meaningful and modern taxonomic concept of most fungal genera discussed in this chapter. He was also a source of inspiration for many young mycologists around the world. H. Kazemi Shariat Panahi · M. Dehhaghi Department of Microbial Biotechnology, School of Biology and Centre of Excellence in Phylogeny of Living Organisms, College of Science, University of Tehran, Tehran, Iran Faculty of Medicine and Health Sciences, Macquarie University, Sydney, NSW, Australia Biofuel Research Team (BRTeam), Karaj, Iran G. Salehi Jouzani Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran R. Zare (*) Iranian Research Institute of Plant Protection, Agricultural Research, Education and Extension Organization (AREEO), Tehran, Iran © Springer Nature Switzerland AG 2020 G. Salehi Jouzani et al. (eds.), Fungi in Fuel Biotechnology, Fungal Biology, https://doi.org/10.1007/978-3-030-44488-4_4

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with biofuels such as bioethanol, biogas, biobutanol, bio-crude oil, and biodiesel (Dehhaghi et al. 2019; Kazemi Shariat Panahi et al. 2019b, c, d; Rahimzadeh et al. 2018; Shirzad et  al. 2019; Tabatabaei et  al. 2019a). However, the conversion of agricultural crops (such as oil or cereal crops) into biofuels could put nations into hunger, raising food vs. fuel debate (Aghbashlo et al. 2017, 2018b, 2019). To avoid this, residual wastes must be identified, collected, and processed to serve as suitable and sustainable substrates for biofuel production (Aghbashlo et al. 2018a). Two such feedstocks are lignocellulose and chitin; however, the conversion of these second-generation feedstocks for biofuel production requires some extra procedures, compared to the first-generation feedstocks. For example, conversion of lignocellulose into bioethanol requires two extra steps, i.e., delignification and saccharification prior to ethanolic fermentation. These extra steps add extra expenses, translating into higher bioethanol production cost. To address this drawback, microorganisms such as filamentous fungi could be appreciated for on-site degradation of lignocellulose (mainly delignified) into reducing sugars or even subsequent bioethanol production through various production approaches including separate hydrolysis and fermentation, simultaneous saccharification and (co)fermentation, or consolidated bioprocessing. Amid this, the economic and environmental feasibility of bioethanol production from lignocelluloses could be increased further, through supporting sustainable agriculture by means of simultaneous production of fungal biocontrol agents. In fact, controlling plant pests is among the significant challenges in agriculture and food safety. Various chemical and physical strategies have been employed to control plant pests and diseases. Chemical pesticides are extensively used to overcome this issue. However, the adverse effects of excessive use of chemical pesticides on food chain have forced pest management researchers to develop alternatives with lower risks (Heydari and Pessarakli 2010). Recently, the World Health

M. Aghbashlo (*) Department of Mechanical Engineering of Agricultural Machinery, Faculty of Agricultural Engineering and Technology, College of Agriculture and Natural Resources, University of Tehran, Karaj, Iran e-mail: [email protected] M. Tabatabaei (*) Institute of Tropical Aquaculture and Fisheries (AKUATROP), Universiti Malaysia Terengganu, Terengganu, Malaysia Microbial Biotechnology Department, Agricultural Biotechnology Research Institute of Iran (ABRII), Agricultural Research, Education and Extension Organization (AREEO), Karaj, Iran e-mail: [email protected]

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Organization, and Food and Agriculture Organization of the United Nations have attempted to regulate the use of pesticides to protect the environment as well as human health (WHO 2016). Intense application of chemical pesticides, particularly fungicides in agriculture, has accumulated toxic compounds in plants and soil, which are harmful for the environment including human and animals. Moreover, resistance to these chemicals is rapidly occurring in pathogenic organisms. To overcome the mentioned challenges, an alternative is biological control to suppress plant pests, through which natural agents are applied to inhibit the plant pests or pathogens. The biocontrol agents generally have rapid proliferation rate in soil and exert positive effects on plants without causing toxicity to them or to humans. These bioagents (usually in combination with biofertilizers) could even improve plant growth and increase crop yield. For these reasons, biological control is known as an effective approach to achieve a more sustainable agriculture (Gawai 2018). Therefore, conversion of lignocellulose into bioethanol with the help of fungi that could also be applied as biocontrol agents at the end of fermentation could be a very innovative strategy.

4.2  Pest Control Strategies Basically, there are three biological control methods: natural conservation, inoculation/inoculative or classical method, and augmentation. In the natural control, the population of pests is reduced through their natural enemies. This strategy has been in function since the first evolution of terrestrial ecosystems over 500 million years ago. Natural conservation occurs in all the ecosystems without human interference and has a vital role in pest control in agriculture (Waage and Greathead 1988; Zare and Gams 2003). Humans can manipulate the natural conservation to achieve the highest performance efficiency of pests’ natural enemies. Inoculative biocontrol method involves the collection of the natural biocontrol agents, usually from their habitats, and their subsequent introduction into the environment where they are not found naturally in order to suppress the pests. This type of biocontrol method is mostly used to control pests, which have been introduced to a specific ecosystem where their natural enemies are absent. Inoculative biocontrol method is used approximately on 10% of the agricultural lands worldwide, and about 165 weed and pest species have permanently been brought under control (Bale et al. 2007; Cock et al. 2010). Augmentation biocontrol boosts the population of natural biological control agents to immediately suppress the native or non-native pests. This method has been known as an environmentally successful alternative to control the pests in various types of field crops such as cotton, maize, sugarcane, and soybean. In contrast to the natural and inoculation biocontrol methods which are performed using public funding, the augmentation is considered as a commercial project because it needs a large-scale production of natural biocontrol agents (Van Lenteren 2012).

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4.3  Types of Biopesticides 4.3.1  Insecticides Entomopathogens have been extensively used against insects as biocontrol agents in agriculture. The most important bio-insecticides are bacteria (mainly, Bacillus thuringiensis), viruses (e.g., Baculovirus spp.), entomogenous fungi (e.g., Metarhizium), and entomopathogenic nematodes (e.g., Steinernema spp.). Inundative release method is mostly applied for insect control, in which a large number of insect pathogens are applied to a high density of pest population for immediate suppression of pests. In fact, in this method, the pathogens of insect proliferate during several days and suppress the pest in a short time. However, in some cases, the effective insect control is achieved through secondary transmission (Cory and Franklin 2012). Bacillus spp. are the most important bacteria, which are applied for insect control. B. thuringiensis is a highly potent entomopathogen that is extensively used as bio-insecticide. In a favorable trophic environment, the bacterium proliferates rapidly and is in vegetative form. Insufficient carbon sources in the environment encourage it to enter the sporulation phase to produce Cry and Cyt proteins that are toxic for insects (Bravo et al. 2011). B. thuringiensis isolates are classified according to the serology tests as well as targets of their toxins. The most applicable bacterial serovars are san diego and tenebrionis against coleopteran larvae and aizawai and kurstaki against lepidopteran larvae (Siegwart et al. 2015). Several viruses have been introduced as insect biocontrol agents. However, baculoviruses have been characterized as the only practical insect pathogens (Suty 2010). Baculoviruses are a large group of DNA viruses, which can infect several insect species within Coleoptera, Diptera, Lepidoptera, and Trichoptera (Siegwart et al. 2015). Fungi are another promising source of insecticides, and so far over 700 species have been known to be entomopathogenic among which nine species Aschersonia aleyrodis, Beauveria bassiana, Beauveria brongniartii, Entomophaga maimaiga, Hirsutella thompsonii, Lecanicillium lecanii, Lagenidium giganteum, Metarhizium anisopliae, and Paecilomyces fumosoroseus comprise the most potent strains (Siegwart et al. 2015). To kill insects, fungal branching hypha must penetrate into the insect body through secreting degrading enzymes such as chitinases and cuticle-­ degrading protease (Castellanos-Moguel et al. 2007; Fang et al. 2005).

4.3.2  Herbicides Mostly, bio-herbicides refer to fungi and their bioactive metabolites; however, there are some reports of bacteria and viruses as bio-herbicides (Charudattan and Hiebert 2007; Daigle et al. 2002; Ferrell et al. 2008). Native or introduced biocontrol agents

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that can target weeds are usually used as the complementary agents to conventional biocontrol methods, particularly in areas where the pathogens have become resistant to chemical herbicides. Biocontrol of weeds are generally classified into three methods: classical, augmentative (inundative), and cultural methods. In the classical approach, a biocontrol agent is introduced into an environment where the biocontrol agent is not found naturally but can preserve its sustainability in the new environment. Controlling the Chondrilla juncea, known as rush skeleton weed, by using Puccinia chondrillina is an example of classical methods (Barton 2004). The inundative biocontrol includes the repeated applications of a bio-herbicide agent in order to decrease the weed population to a density in which the plant can be suppressed. The cultural biocontrol method includes several steps including crop rotation, land preparation, fallow management, sanitation, water management, and soil fertility preservation. Currently, there are strong reasons for using bio-herbicides, in particular microbial herbicides, instead of their chemical counterparts. The emergence of herbicide-resistant weeds, herbicide persistence in soil and crops, negative effects on non-target organisms, and non-selectivity are among the significant limitations of using the chemical herbicides (Guske et al. 2004; Sauerborn et al. 2007).

4.3.3  Fungicides There is a large variety of fungal diseases which occur in various parts of plants such as root, fruit, leaves, and stem. Most biocontrol methods of fungal diseases focus on soil-borne diseases. Accordingly, several biocontrol agents have been examined to decrease the fungal pathogens in soil (Heydari et al. 2007; Heydari and Pessarakli 2010). Several strains of Trichoderma harzianum  (T. harzianum) have been identified as potential biocontrol agents of plant diseases, particularly in the initial stages of the disease. It has been shown that increasing the population of T. harzianum to 5 × 105 cfu/g of soil significantly reduced the Pythium blight and root rot diseases (Heydari and Pessarakli 2010). Rhizoctonia and Sclerotinia are among the most important fungal pathogens and remaining their structures in soil can supply the initial inoculum. Therefore, controlling the initial inoculum in soil is the first step in suppression of fungal pathogens. Apart from Trichoderma, several bacterial species such as Bacillus subtilis, Bacillus cereus, Burkholderia cepacia, Pseudomonas fluorescens, Pseudomonas putida, and Pseudomonas aerofaciens have also been identified as effective biocontrol agents of soil-borne fungal diseases. By using these bacteria, various fungal pathogens such as Fusarium oxysporum, Fusarium solani, Gaummannomuces graminis, and Rhizoctonia solani have been successfully controlled (Kloepper et  al. 2004; Saravanakumar et  al. 2016; Shahraki et al. 2009).

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4.4  Fungal Biocontrols Fungi have received increasing interest as the biocontrol agents during the past several years, considering the number of commercial biocontrol products and potential candidates under development. Research on fungal biological control is rapidly growing with a focus on increasing plant productivity, protecting human and animal health, and improving food quality. Fungi possess significant characteristics such as high host specificity, ubiquitousness, rapid growth, destruction of host, and ease of culture and preservation in laboratory, making them suitable for application as promising biocontrol agents. The fungal strains approved as biocontrol agents by the Environment Protection Agency are listed in Table 4.1.

4.4.1  Fungal Species The most important fungal biocontrol genera are Aspergillus, Ampelomyces, Coniothyrium, Gliocladium, Fusarium, Paecilomyces, Pichia, and Trichoderma (Atehnkeng et  al. 2008; Gilardi et  al. 2008; Koumoutsi et  al. 2004) (Table  4.2). Among them, the genus Trichoderma consists of potent microorganisms capable of suppressing the pathogenic fungi as well as enhancing plant growth (Harman et al. 2004; Vinale et al. 2008). For instance, it has been reported that a conidial suspension of Trichoderma sp. rendered growth-promoting effect on sunflower and bean while could effectively inhibit Sclerotium rolfsii, i.e., the causal agent of root rot and seed rot diseases in these plants (Yaqub and Shahzad 2008). T. harzianum (ATCC 20476) was the first fungal strain introduced by the United States Environmental Protection Agency for controlling plant diseases (Gawai 2018).

4.4.2  Biocontrol Strategies Biological control is the consequence of various types of interactions between the pathogen and the biocontrol agent. Direct antagonisms, antibiosis, and competition are the main strategies involved in fungal biocontrol process. Direct antagonism includes the lysis and death of pest by biocontrol agent which is also known as hyperparasitism. In this relation, the fungi that are used as biocontrol agents are called mycoparasites (Jyoti and Singh 2016). Two types of mycoparasitism can be defined: (i) biotrophic mycoparasitism and (ii) necrotrophic mycoparasitism. In the former, the hyperparasite obtains nutrients from another organism without killing the host. In fact, this relation usually continues in a stable way. Apparently, this mycoparasitic interaction is not favorable for commercial biocontrol aims because the mass production of hyperparasite depends on its host cell as substrate. In the second form of mycoparasitism, the hyperparasite gains the

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Table 4.1  Commercial fungal biocontrol products Fungal strain Ampelomyces quisqualis M-10 Aspergillus flavus AF36 Gliocladium catenulatum JI446 Gliocladium virens GL-21 Trichoderma harzianum T-22

Commercial product AQ10 Alfa guard Prima stop soil guard

WRC-GL-21 and WRC-AP-1 RootShield

Plant disease Powdery mildew Aflatoxin

Pathogen Erysiphe cichoracearum Aspergillus flavus

Pythium, Fusarium, Phytophthora, Rhizoctonia Damping-off disease and root rot Root diseases

Soil-borne pathogens

Various fungi, parasitic nematodes Pythium, Fusarium, Rhizoctonia, Thielaviopsis, Cylindrocladium Pythium, Fusarium

Circle One Global, USA Kemira Agro Oy, Finland

W.R. Grace & Co. USA BioWorks, Geneva, N.Y., USA

T. harzianum T-22 T. harzianum T39

T-22G, T-22B

T. harzianum

Harzian 20, Harzian 10

T. harzianum

F-stop

T. harzianum

Supravit

Trichoderma spp.

Solsain, Hors-solsain, Plantsain

Root diseases

Trichoderma spp. Trichoderma spp. T. virens

ANTI-­ FUNGUS TY

–

Rhizoctonia, Pythium, Sclerotinia, and other sclerotic fungi –

–

–

Grondontsmettingen De Ceuster, Belgium Mycontrol, Israel

GlioGard and SoilGard Promot PlusWP Promot PlusDD

Damping-off diseases –

Pythium ultimum, Rhizoctonia solani –

Grace-Sierra Co., Maryland, USA Tan Quy, Vietnam

Trichoderma spp. T. koningii T. harzianum

Trichodex

Wilt, ear, and kernel rot Downy mildew, Botrytis cinerea white mold, powdery mildew – Pythium, Sclerotinia, Armillaria mellea Various pathogenic Damping-off fungi and seed rot diseases – –

Company EcoGen, USA

TGT Inc., N.Y., USA BioWorks, USA

Natural Plant Protection, Noguères, France Eastman Kodak Co., United States TGT Inc., N.Y., USA Bonegaard and Reitzel, Denmark Prestabiol, Montpellier, France

(continued)

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Table 4.1 (continued) Fungal strain Trichoderma spp. Trichoderma spp. Trichoderma spp.

Commercial product TRiB1

Plant disease –

TRICÔ-­ ĐHCT Vi – ĐK

Yellow leaf rot

T. virens

NLU-Tri

–

T. viride

Biobus 1.00WP

Trichoderma spp.

Bio – Humaxin Sen Vàng 6SC, Fulhumaxin 5.15SC BioSpark Trichoderma

Diseases in roots, crab spot, wilt Yellow leaves, root tumor

T. parceramosum, T. pseudokoningii, and T. harzianum

Root and wilt diseases

–

Pathogen –

Company National Institute of Plant, Vietnam Fusarium Can Tho University, Vietnam Pesticide Corp., Fusarium, Rhizoctonia solani, Vietnam Sclerotium rolfsii, Phytophthora palmivora, Pythium Various pathogenic Ho Chi Minh fungi University of Agriculture and Forestry, Vietnam Nam Bac, Vietnam Fusarium, Rhizoctonia solani, Phytophthora Fungi and An Hung Tuong, nematodes Vietnam

Various pathogenic BioSpark fungi Corporation, Philippines

nutrients from host spores or hyphae after killing the cells (Köhl et al. 2019). In the mycoparasitic interaction, the sequential processes that occur include identification of location, direct contact, recognition, penetration to the host, growth, lysis of pathogen, and exit. Various chemical interactions that occur during these processes mainly include production of lectins for initial recognition and contact, and release of several hydrolytic enzymes during penetration step. Fungal cell wall is the first barrier between mycoparasite and pathogen; therefore, production of hydrolytic enzymes, which can degrade the cell wall of pathogenic fungi, is a crucial step in mycoparasitic interactions. The main enzymes involved in lysis of fungal cell wall are chitinase, protease, N-acetyl glucosaminidase, and glucanase. Induction pattern of hydrolytic enzymes production varies among fungal strains, but generally the mycoparasite initially produces low amounts of chitinase. Following the release of oligomers due to cell wall degradation, chitinase production is highly induced, leading to complete lysis (Benítez et  al. 2004; Jyoti and Singh 2016). Among the

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Table 4.2  A summary of the biocontrol research works using main fungal species Fungi Non-toxigenic Aspergillus oryzae and Aspergillus sojae Atoxigenic Aspergillus flavus Atoxigenic A. flavus Atoxigenic A. flavus Atoxigenic A. flavus Atoxigenic A. flavus Fusarium oxysporum Fo-B2 F. oxysporum F. equiseti GF191 Fusarium equiseti GF183 F. oxysporum F221-B

Pichia membranifaciens Pichia guilliermondii Pichia anomala Pichia caribbica Pichia caribbica

Pichia anomala WRL-076 Pichia anomala Pichia membranifaciens

Pathogen Plant disease Toxin-producing fungi Toxin contamination

Host plant Cotton, corn, and peanuts

Aflatoxin contamination Aflatoxigenic strains Aflatoxin contamination Aflatoxigenic A. flavus Aflatoxin contamination Aflatoxigenic strains Aflatoxin contamination A. flavus Aflatoxin contamination F. oxysporum f.sp. Wilt of tomato lycopersici CU1 Fusarium oxysporum Fusarium wilt f.sp. cubense of banana Crown and root F. oxysporum f.sp. radicis-lycopersici rot Fusarium oxysporum Fusarium wilt f.sp. spinaciae of spinach Leaf lesions Curvularia lunata Wilt and root F. semitectum rot disease F. oxysporum f.sp. lactucae Rhizoctonia solani Rhizoctonia sp. Botrytis cinerea Gray mold disease Penicillium italicum Citrus blue mold Botrytis cinerea Postharvest Penicillium expansum diseases Rhizopus stolonifer Postharvest diseases Penicillium expansum Postharvest blue mold decay A. flavus Aflatoxin contamination Botrytis cinerea Postharvest diseases Monilinia fructicola Brown rot

Maize

A. flavus

References Dorner et al. (2000)

Atehnkeng et al. (2008) Corn Degola et al. (2011) Cashew nuts Al-Othman et al. (2013) Peanuts Alaniz et al. (2016) Maize Camiletti et al. (2018) Tomato Shishido et al. (2005) Banana Nel et al. (2006) Tomato Spinach Several crops

Grapevine

Horinouchi et al. (2007) Horinouchi et al. (2010) Thongkamngam and Jaenaksorn (2017)

Apple

Santos and Marquina (2004) Lahlali et al. (2011) Haïssam (2011)

Peach

Xu et al. (2013)

Apple

Li et al. (2014)

–

Hua et al. (2015)

Apple

Sare et al. (2018)

Plum fruit

Zhang et al. (2019)

Citrus

(continued)

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Table 4.2 (continued) Fungi Paecilomyces variotii P. variotii

P. variotii

Paecilomyces lilacinus PL251 P. lilacinus UP1 P. lilacinus

P. lilacinus P. lilacinus Pochonia chlamydosporia P. chlamydosporia

Pathogen Meloidogyne javanica

Plant disease Root disease

Pythium spinosum

Root disease

Charcoal rot disease Stalk rot Root-knot disease Root-knot disease Meloidogyne incognita Root-knot disease Aculus schlechtendali Leaves rusting, bronzing, and premature drying Meloidogyne incognita Root-knot disease Root-knot nematodes Root-knot disease Meloidogyne javanica Root-knot disease Biscogniauxia mediterranea Fusarium moniliforme Phytophthora cinnamomi Meloidogyne incognita

P. chlamydosporia

Globodera Heterodera Meloidogyne Nacobbus Meloidogyne javanica

P. chlamydosporia

Root-knot nematodes

Gliocladium virens Meloidogyne javanica G. virens

Pathogenic fungi

T. harzianum

Meloidogyne javanica

T. harzianum T. hamatum T. asperellum

Moniliophthora roreri

Host plant Various trees Soybean

Maize and corn

Tomato Tomato Apple

Vigna radiata Tomato Tomato

Root-knot disease

Various crops

Root-knot disease Root-knot disease Potato cyst Root-knot disease Seed-borne fungi diseases

Pistachio Potato

Eggplant

Lentil and black gram (Vigna mungo) seeds Root-knot galls Tomato Frosty pod rot disease

Cacao

References Al-Qasim et al. (2009) Al-Sheikh and Abdelzaher (2010) Rodrigo et al. (2017)

Kiewnick and Sikora (2006) Oclarit and Cumagun (2009) Demirci and Denizhan (2010)

Khan et al. (2019) Ahmed and Monjil (2019) Dallemole-­ Giaretta et al. (2012) Manzanilla-­ López et al. (2013) Ebadi et al. (2018) Dos Santos et al. (2019) Ashraf and Khan (2007) Agarwal et al. (2011)

Sahebani and Hadavi (2008) Bailey et al. (2008) (continued)

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Table 4.2 (continued) Fungi T. virens T. harzianum T. cerinum T. viride

Pathogen Rosellinia necatrix

Plant disease White root rot

Host plant Avocado

Soybean Fusarium oxysporum Root disease Pythium arrhenomanes T. brevicompactum Meloidogyne incognita Root-knot galls Tomato T. harzianum

F. oxysporum

T. asperellum

Pythium myriotylum

T. gamsii

Fusarium culmorum Fusarium graminearum Fusarium oxysporum f.sp. lycopersici

T. harzianum

Wilt of Cucumber cucumbers Root rot disease Cocoyam Fusarium head blight

Rice

Wilt of tomato

Tomato

T. asperellum

Phytophthora capsici

Root rot disease Pepper

T. asperellum

Colletotrichum gloeosporioides

Anthracnose

Mango

T. harzianum T. ghanense T. hamatum T. asperellum

Fusarium oxysporum f.sp. melonis

Wilt of melon

Melon

Phytophthora ramorum

Sudden oak death

Rhizoctonia solani

Bean plants

Fusarium oxysporum f.sp. lycopersici Rhizoctonia solani

Wilt of tomato

Oak and other species of trees Roots and lower stem diseases Tomato

Sheath blight

Rice

T. asperellum, T. harzianum, Trichoderma spp. T. asperellum T. asperellum T. longibrachiatum, T. harzianum T. gamsii T. harzianum T. viride

Colletotrichum gloeosporioides Panax notoginseng Meloidogyne javanica

Anthracnose

Papaya fruits Root rot disease Rice Root-knot galls Tomato

T. asperellum

Fusarium oxysporum f.sp. cucumerinum

Fusarium wilt disease

Cucumber

References Rosa and Herrera (2009) John et al. (2010)

Affokpon et al. (2011) Yang et al. (2011) Mbarga et al. (2012) Matarese et al. (2012) Sundaramoorthy and Balabaskar (2013) Segarra et al. (2013) de los Santos-­ Villalobos et al. (2013) Martínez-Medina et al. (2014) Widmer (2014)

Asad et al. (2014)

El_Komy et al. (2015) De França et al. (2015) Valenzuela et al. (2015) Chen et al. (2016) Al-Hazmi and TariqJaveed (2016) Saravanakumar et al. (2016) (continued)

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Table 4.2 (continued) Fungi T. harzianum

Pathogen Sclerotinia sclerotiorum Macrophomina phaseolina Ralstonia solanacearum Fusarium graminearum Fusarium oxysporum f.sp. lycopersici Sclerotium cepivorum

Plant disease Sclerotinia stem rot Soybean charcoal rot Tobacco bacterial wilt Fusarium stalk rot Tomato wilting

Tomato

White rot

Onion

Penicillium purpurogenum

White Yam Tuber

Yam

T. harzianum

Globodera pallida

Potato cyst

T. asperellum

Ramularia areola

Ramularia leaf spot Gray mold rot

T. harzianum T. harzianum T. harzianum T. asperellum T. album, T. harzianum, T. koningii, T. viride, and T. virens T. harzianum

T. harzianum

Botrytis cinerea, Mucor circinelloides, Aspergillus fumigatus, Aspergillus flavus, Rhizoctonia solani, Candida albicans T. atroviride Rhizoctonia solani Systemic disease T. harzianum Puccinia graminis Stem rust T. viride f.sp. tritici disease T. harzianum Fusarium Fusarium stalk graminearum rot Foot rot disease F. oxysporum, T. asperellum T. brevicompactum Rhizoctonia solani, Phytophthora capsici T. harzianum T. harzianum F. oxysporum Wilt disease T. asperellum T. asperellum

Trichoderma spp.

Fusarium verticillioides Corynespora cassiicola Curvularia aeria Pythium aphanidermatum

Fusarium ear rot Leaf spot

Pythium damping-off and root rot diseases

Host plant Soybean Soybean Tobacco Maize

References Zhang et al. (2016b) Khaledi and Taheri (2016) Yuan et al. (2016) Saravanakumar et al. (2017) Patel and Saraf (2017) Elshahawy et al. (2017)

Gwa and Abdulkadir (2017) Potato Contina et al. (2017) Cotton da Silva et al. (2017) Deng et al. Apples, oranges, and (2018) cucumbers

Cucumber Wheat Maize Black pepper and ginger Tomato Maize Lettuce

Tomato

Nawrocka et al. (2018) El-Sharkawy et al. (2018) Saravanakumar et al. (2018) Rai et al. (2019)

Bader et al. (2019) Veenstra et al. (2019) Baiyee et al. (2019) Elshahawy and El-Mohamedy (2019)

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parasitic fungi, Trichoderma and Clonostachys are the most studied fungi with high ability to produce a wide range of hydrolytic enzymes (Köhl et al. 2019). The second strategy, i.e., antibiosis, refers to a process in which fungi produce and release bioactive compounds to their surroundings that can kill pathogens. Most fungal species are able to produce secondary metabolites with antibiotic activity, particularly associated with their specific morphological differentiation steps (Jyoti and Singh 2016). Importantly, some fungal bioactive metabolites can affect the metabolism of plants, while others target the pathogens specifically. Antagonistic fungi belonging to the genera Trichoderma and Fusarium have been identified to produce volatile and non-volatile compounds. These compounds inhibit the colonization and the growth of pathogenic organisms and play important role in biological controls of several pathogens (Benítez et  al. 2004; Mathivanan et  al. 2008). Alamethicins, harzianic acid, peptaibols, massoilactone, viridian, glisoprenins, and heptelidic acid are among the important toxic metabolites of Trichoderma (Benítez et al. 2004). In general, Fusarium is a well-known plant pathogenic genus, several species of which cause root diseases. On the other hand, several Fusarium spp. such as F. decemcellulare, F. solani, F. longipes, and nonpathogenic F. oxysporum have been introduced as significant biological control microorganisms. Fusapyrone and deoxyfusapyrone isolated from Fusarium semitectum have been reported as strong antifungal compounds against various pathogenic fungi such as Alternaria alternata, Ascochyta rabiei, Aspergillus flavus, Botrytis cinerea, and Penicillium verrucosum (Mathivanan et al. 2008). Growth of plant pathogens highly depends on the available nutrients in the environment. Obligate biotrophic pathogens meet their nutritional requirements through infected host cells with no need for exogenous nutrient existence in their environment (Köhl et al. 2019). Most plant pathogens gain their essential nutrients by degrading the dead cells of plants, consuming their organic matter. Necrotrophic plant pathogens kill the plant tissue and then attack the cells to access the available nutrients. After induction of plant necrosis by pathogenic organisms, non-­ pathogenic saprophytic microorganisms also colonize the necrotic tissues and use the available nutrients; therefore, triggering a competition over nutrients. Some rare micronutrients such as Fe3+ have low solubility in soil; therefore, competition of microorganisms to access iron is critical (Jyoti and Singh 2016). Microorganisms obtain iron through production of low-molecular siderophores, which have high affinity for ferric iron. Therefore, microorganisms with a high ability to release siderophores and, hence, high affinity to iron can be considered as potential biocontrol candidates (Köhl et al. 2019; Whipps 2001). This strategy has been observed in some fungal species, for example, Trichoderma asperellum produces siderophores with a high affinity to iron and can control Fusarium wilt through scavenging iron (i.e., competition for iron) (Segarra et al. 2010). The yeast Metschnikowia pulcherrima can transform iron to a red pigment named pulcherrimin, suppressing the growth of Alternaria alternata, Botrytis cinerea, and Penicillium expansum by making iron unavailable in the environment (Köhl et  al. 2019; Saravanakumar et al. 2008).

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4.5  Bioethanol Production from Lignocellulosic Biomass Lignocellulose is made up of a carbohydrate portion (i.e., cellulose and hemicellulose) that is tightly bound (i.e., covalent and hydrogen bonds) and protected by an aromatic polymer (i.e., lignin) (Kazemi Shariat Panahi et al. 2019a). Three steps must be fulfilled for a successful conversion of lignocellulose into bioethanol, including (i) liberation of the carbohydrate portion from its aromatic protection, (ii) degradation of the liberated carbohydrates into free sugars, and (iii) fermentation of the produced sugars (i.e., pentoses and hexoses) into ethanol (Kazemi Shariat Panahi et al. 2019a). Lignocellulose-based commercial bioethanol plants are yet to find their real position in the market due to cost constraints. The high bioethanol production cost could be attributed to the recalcitrant structure of lignocellulosic biomass, requiring various chemical, physical, or physicochemical pretreatments in addition to high amounts of hydrolyzing enzymes to obtain reasonable sugar yields (Kazemi Shariat Panahi et al. 2019a). Moreover, a detoxification step is often required following the delignification process that adds extra expenses to lignocellulosic bioethanol production, compared to the first-generation bioethanol production plants. To decrease bioethanol cost from lignocellulosic biomass, on-site production of cellulases could be considered (Kazemi Shariat Panahi et al. 2019a). More specifically, the conventional process for bioethanol production from lignocellulosic biomass includes pretreatment of biomass, saccharification of delignified biomass to produce monomers, and fermentation of simple sugars to bioethanol using usual microorganisms such as Saccharomyces cerevisiae (S. cerevisiae), Escherichia coli, or Zymomonas mobilis. Apparently, elimination or combination of some of the mentioned steps could improve the economics of the process. For example, a consolidated bioprocessing could be adapted for conducting all the mentioned three steps in one reactor. It should be noted that considerable amounts of cellulases (30–50 mg/g of crystalline cellulose) and long retention times are still required for a complete saccharification of delignified biomass. Therefore, tons of enzymes would be consumed in a typical commercial lignocellulose-based bioethanol production plant. To meet such large enzyme quantity requirements, various microorganisms could be used for biological pretreatments or enzymes production (Hamedi et  al. 2015; Mohammadipanah et al. 2015; Tabatabaei et al. 2019b, c). Nevertheless, the most viable solution would probably be the exploitation of fungi because they have evolved robust secretion systems equipped with the Golgi complex and the endoplasmic reticulum within the cell cytosol (Xu et  al. 2009). This evolution allows them to secrete huge amounts of hydrolytic enzymes, compared to the most potent cellulolytic bacteria. For example, Trichoderma reesei could secrete >100  g cellulase/L of culture medium. A detailed explanation of various pretreatment techniques for preparation and delignification of different types of biomass and their subsequent saccharification methods has been provided in our previous report (Kazemi Shariat Panahi et al. 2019a).

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4.6  Bioethanol-Producing Biocontrol Fungi Overall, the application of some fungal species for enhanced production of biofuels (such as biogas and bioethanol) is well-known (Kazemi Shariat Panahi et al. 2019a; Tabatabaei et al. 2019b, c). In this section, some fungal biocontrol species that could be used during biological pretreatment and hydrolysis of lignocellulose (i.e., upstream) or bioethanol production step (i.e., mainstream) are presented and discussed.

4.6.1  Genus Fusarium Fusarium spp. are potent xylanase and cellulase producers, which could be used for saccharification of delignified feedstock (Bertonha et al. 2018). Some of the members of this genus are endophytic and phytopathogenic agents, and hence, are responsible for serious plant diseases such as Fusarium wilt of banana (see Chapt. 2). On the other hand, there are some members that are considered safe (non-­ pathogen) while rendering high hydrolyzing enzyme production capability. Interestingly, some Fusarium spp. (e.g., F. oxysporum) are even considered as potential ethanologens. The infiltration of the plant lignin barrier and hydrolyzing the carbohydrate portion into bioethanol allows conducting a consolidated bioprocessing in which a single reactor is constructed for simultaneous production of hydrolytic enzymes, hydrolysis of delignified biomass, and ethanolic fermentation (Kazemi Shariat Panahi et al. 2019a; Nugent et al. 2019). Following a sufficient aerobic growth phase, the proliferated F. oxysporum could ferment (under anaerobic or microaerobic conditions) sugars into bioethanol. Interestingly, wild-type F. oxysporum could tolerate high titers of ethanol (4.5–5%), high levels of sugars, and other inhibitors such as acetate (Ali et al. 2016; Hennessy et al. 2013). Indeed, this filamentous fungus could reduce acetate (i.e., one of the main inhibitory compounds produced by microorganisms during ethanolic fermentation) into bioethanol. However, for this purpose, F. oxysporum typically requires a limited oxygen supply (18–20%) (Ali et  al. 2016; Panagiotou et  al. 2005c). Moreover, this amount of oxygen allows the microorganism to keep growing. Ethanol concentrations as high as 80  mg/g of un-pretreated lignocellulose (i.e., wheat straw/bran) were achieved through a direct process using F. oxysporum strain 11C (Ali et al. 2012). Intriguingly, this F. oxysporum strain could deliver about four-­ time higher bioethanol yield (i.e., 3.26 g/g alkali-treated straw) with a corresponding theoretical yield of 80.2% if the lignocellulosic biomass was subjected to pretreatment (Ali et al. 2012). This could be attributed to the ability of F. oxysporum in secretion of three types of cellulase enzymes (i.e., endo- and exo-1,4-β-D-­ glucanases and β-glucosidase). More specifically, F. oxysporum could anaerobically (or under  microaerobic conditions) consume various types of biomass, while its enzymatic system could sufficiently tolerate the inhibitory compounds generated

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during lignocellulose pretreatment step as well as high ethanol concentrations (Paschos et al. 2015; Xiros et al. 2011). F. oxysporum could provide well-balanced and highly efficient enzymatic system (e.g., β-glucosidase, endoglucanase, xylanase, acetyl esterase) for hydrolyzing delignified biomass, delivering fermentation medium containing high enzyme titer but no free oligosaccharides (Panagiotou et  al. 2011). Interestingly, the secreted β-glucosidase by F. oxysporum strain F3 exhibits higher activity than endoglucanases, preventing the inhibition of cellulase by cellobiose (Panagiotou et al. 2005a). It should also be mentioned that endoglucanase from this strain has a low molecular mass, facilitating the penetration of this degrading enzyme through cellulose fibers. It was reported that this strain could aerobically thrive on cellulose at a specific growth rate of 0.023/h while delivering an ethanol yield of 0.35 g/g cellulose with a productivity rate of 0.044 g/L/h at the subsequent fermentation step (i.e., anaerobic conditions) (Panagiotou et al. 2005a). The ethanologenic characteristic of F. oxysporum strain 11C is linked to a large consortium of genes (91% efficiency) enzyme-saccharified hydrolysates of acid steam exploded sugarcane bagasse or plywood chips into bioethanol despite the presence of inhibitors such as acetic acid (4.8–7.65 g/L) and furaldehydes (e.g., 5-hydroxymethyl furfural, ~0.2–0.7 g/L; and furfural ~1.1–1.2 g/L) (Yuan et al. 2017). Interestingly, it was found that ethanolic fermentation at 42  °C could trigger genes encoding alcohol dehydrogenase 1-4, heat shock proteins (hsp90 and ssq1), and glyceraldehyde3-phosphate dehydrogenase (tdh2) in some P. kudriavzevii strains such as RZ8-1 for thermotolerance ability (Chamnipa et al. 2018). The mentioned abilities highlight the multi-stress-tolerance characteristic of some P. kudriavzevii strains, which could be efficiently exploited for bioethanol production from lignocellulosic biomass.

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Acid hydrolysis is a common pretreatment technique in lignocellulose-based bioethanol production plants. Therefore, the environmental burden and operating cost of the subsequent biological hydrolysis step could be minimized, while the efficiency of the process could be improved by using resistant microorganisms toward the extreme processing conditions (e.g., low pH and toxic compounds). One of the robust microorganisms for this purpose could be Pichia anomala (P. anomala  also known as Wickerhamomyces anomalus or Hansenula anomala), a yeast with innate stress resistance mechanisms. This yeast could grow about two times faster than S. cerevisiae (0.42/h vs. 0.22/h) when they are cultured in minimal media with a pH value of 3 (Fletcher et al. 2015). In contrast, S. cerevisiae was a more efficient ethanologen, producing more than 15 mmol/g dried cell weight/h bioethanol (i.e., about two times higher than that of P. anomala). Moreover, S. cerevisiae displayed slightly higher glucose uptake rate than P. anomala (i.e., 11.2 vs. 10.8 mol/g dried cell weight/h) on minimal media (Fletcher et al. 2015). Zha et al. (2013) reported that P. anomala (strain CBS 132101) could thrive in various biomass hydrolysates containing inhibitory compounds generated during lignocellulose pretreatment step. Additionally, this strain could successfully assimilate nitrate (as a nitrogen source) and xylose (a pentose sugar) for bioethanol production in the presence (20% air) or  the absence of oxygen. More specifically, the strain CBS 132101 did not exhibit a Pasteur effect, and relatively equal amounts of bioethanol were produced. However, the addition of oxygen by supplying air could result in faster growth rate (Zha et al. 2013). Kostas et al. (2016) measured the stress tolerance of P. anomala and investigated its metabolic activity against various synthetic minimal media and seaweed hydrolysates using the Biolog-Omnilog phenotypic microarray system. It was found that P. anomala had high metabolic outputs when individually assimilating fructose, rhamnose, mannitol, and xylose or seaweed hydrolysates. Accordingly, P. anomala could perform well on Chondrus crispus hydrolysate, producing 3.26  g/L bioethanol; however, a relatively long lag phase was observed. P. anomala could also be used for the biological pretreatment of lignocellulose. In this context, Anjaly Sukumaran Nair (Anjaly Sukumaran Nair 2011) inoculated wheat straw by P. anomala J121 and stored at 4 °C or 15 °C and 0.93 aw or 75.2% solid content for 30 days. A fungal colony count corresponding to 4.75 × 107 CFU/g was enumerated at the lower temperature (i.e., 4 °C). Compared to the uninoculated control, the population of the enterobacteria remained low during storage at higher temperature (i.e., 15 °C), supporting a biocontrol activity of P. anomala. Moreover, the fungal population of the sample remained monoculture (P. anomala), showing biocontrol activity against other fungi (yeasts and molds) (Anjaly Sukumaran Nair 2011). Following the storage pretreatment, an acid pretreatment was performed prior to simultaneous saccharification and fermentation for bioethanol production. The impact of the storage pretreatment on bioethanol production was evaluated by comparing the bioethanol yield with that obtained through a direct acid pretreatment, proving a 7.52% higher bioethanol yield. Moreover, the uninoculated wheat straw was also stored at the same conditions (i.e., 4 °C or 15 °C, 0.93 aw or 75.2% solid content, 30 d) and was then acid-hydrolyzed and fermented. Compared to the

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control, 3.9% higher and 6.17% lower bioethanol yield were obtained from the uninoculated wheat straw stored at 15 °C and 4 °C, respectively (Anjaly Sukumaran Nair 2011). Typically, P. anomala has a weak ability for lignin degradation, and therefore, a delignification process is often required prior to the assimilation of lignocellulose. A comparison of the raw and acid-treated potato peel waste with respect to ethanol production by P. anomala (X19) was conducted (Atitallah et al. 2019). It was found that delignification by acid treatment (30 °C, pH 4.8) prior to bioethanol production could improve its concentration by up to 16.65 times (i.e., 0.55 vs. 9.16 g/L).

4.6.3  Genus Paecilomyces Paecilomyces variotii (P. variotti)  and, to a lesser extent, Paecilomyces lilacinus  (P. lilacinus) are two important ethanologenic species in the nematophagous fungal genus of Paecilomyces. The attention toward P. variotii for bioethanol production is due to its highly glucose-tolerant β-glucosidases. This group of enzymes is a very important component of cellulase system for successful saccharification of cellulose through breaking β-glycosidic bonds. Therefore, the yield of saccharification (especially, in separate saccharification and fermentation approaches) will be significantly improved if β-glucosidases show high glucose tolerance. In 2010, such resistant enzyme was extracted from P. variotii MG3 (Job et  al. 2010). Soon after, it was found that P. variotii could survive and thrive many inhibitory compounds such as gallic acid, acetic acid, ethyl benzene, toluene, p- and m-xylenes, benzene, p-­coumaric acid, as well as phenol and some phenolic compounds (García-Peña et al. 2008; Sachan et al. 2006; Wang et al. 2010). Therefore, the huge capacity of specific detoxification of this fungus could be exploited for the bioremediation of toxic industrial effluents and by-products. In this regard, Pereira et  al. (2012) exploited the capability of P. variotii for the detoxification of eucalyptus spent sulfite liquor followed by its conversion into bioethanol by Pichia stipitis (yield, 0.24 g/g sugars; concentration 2.4 g/L). However, it was in 2014 that the ability of P. variotii (strain ATHUM 8891) for direct bioethanol production from lignocellulose through consolidated bioprocessing was evaluated for the first time (Zerva et al. 2014). The bioethanol production could be enhanced by supplementing nitrogen (e.g., nitrate-nitrogen), and the strain could efficiently ferment xylose and glucose into bioethanol (theoretical yield, 80%; concentration, 10.5 g/L) (Zerva et al. 2014). No diauxic behavior was observed during mixed sugars (i.e., xylose and glucose) fermentation; however, higher bioethanol concentration was recorded from xylose than glucose (3.24 vs. 6.46 g/L, respectively) during individual sugar fermentations. The ability for parallel fermentation of xylose and glucose by this fungus could increase the efficiency of bioethanol production (Zerva et al. 2014). P. lilacinus could optimally grow at 30 °C in the presence of ≥10% NaCl, converting glucose into bioethanol, CO2, acetate, and lactate with an approximate molar

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ratio of 5.4:3.72:1:0.15. Generally, microorganisms could be categorized into halotolerants (tolerating ≥10% salt), moderate halophiles (optimally growing at 2.9–14.6% salt), borderline extreme halophiles (optimally growing at 8.8–28.6% salt), and extreme halophiles (optimally growing at 14.6–30.4% salt) (Hamedi et al. 2015; Mohammadipanah et al. 2015). Accordingly, P. lilacinus could be classified as a moderate halophile. It is worth mentioning that this nematophagous fungus could not assimilate polysaccharides (such as cellulose) except laminarin and, to a lesser extent, xylan (Mountfort and Rhodes 1991). However, its ability to convert xylose and arabinose into bioethanol could be exploited for bioethanol production from mixed disaccharides, pentose, and hexose sugars (such as maltose and cellobiose, xylose, and glucose, respectively). Moreover, saline water or digested high salt content biomass could be assimilated by P. lilacinus for bioethanol production.

4.6.4  Genus Trichoderma Trichoderma is one of the most promising fungal genera for bioethanol production from lignocellulose due to its high cellulase production capacity. Trichoderma spp. (T. viride, T. longibrachiatum, and T. reesei) have long been identified as the prosperous source of industrial cellulases for bioethanol production. The marine Trichoderma spp. have also been found as a highly promising source of cellulases. Accordingly, Saravanakumar and Kathiresan (Saravanakumar and Kathiresan 2014) pretreated the sawdust (29 °C, pH 6.19, 8.16 IU/mL of cellulase) with cellulases obtained from Trichoderma estonicum for fermentable sugar production (glucose production yield, 78.56%), followed by their conversion into bioethanol by S. cerevisiae. Accordingly, it was observed that increasing temperature from 30 to 50 °C improved reducing sugar and glucose production by 95% and 70% during a 36-h hydrolysis process, respectively. This could be translated into a higher subsequent bioethanol production (Bu et al. 2019). Currently, commercial cellulase production from mutant strains of T. reesei are performed in several companies around the world, and the majority of R&D projects on bioethanol production from lignocellulosic biomass are dedicated to using cellulases of Trichoderma origin. Research on T. reesei was mainly initiated in the 1950s for secretion and extraction of hydrolytic enzymes with cellulose degradation capabilities (Viikari et al. 2012). In 1973 and 1979, two main issues including fossil fuel cost and dependence on Middle East politics created serious concerns in the Western countries. These concerns resulted in 20  years of extensive funding for scientific research, leading to key findings about cellulase production. Although several pilot experiments were designed throughout the world, an economical aspect of industrial production was not satisfactory. On the other hand, recent decreases in the cost of fossil fuels removed the motivations for further research and development in this topic. However, the enzyme production process received attention in some industries such as pulp, paper, detergent, and textile industries in which substrates did not need to be degraded completely (Viikari et al. 2012).

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Cellulases produced by T. reesei consist of three types of enzymes: (i) endoglucanases that cleave glycosidic linkage; (ii) cellobiohydrolases that hydrolyze the glycosidic linkages from the end of cellulose, producing cellobiose molecules; and (iii) β-glucosidases that convert cellobiose to glucose. Normally, the activity of β-glucosidases produced by T. reesei is low, leading to the accumulation of cellobiose and the subsequent suppression of endo- and exo-cellulases production (Zhang et  al. 2016a). In contrast, enhanced β-glucosidase activity could be achieved by using engineered T. reesei (such as strain Rut C30) containing an artificial transcription factor, increasing the release of glucose (Zhang et al. 2016a). It is known that some compounds could induce and/or change the composition of T. reesei cellulase cocktail. In this regard, Li et al. (2018) compared the cellulase profile of T. reesei (Rut C30) in the presence of a mixture of glucose and disaccharide as inducer. Although increasing the β-glucosidase production was achieved in response to the mixture of glucose and disaccharide, the activity of enzyme was still low. To address this issue, the gene aabgl1 encoding β-glucosidase in Aspergillus aculeatus was expressed in T. reesei (Rut C30), resulting in an increased β-glucosidase activity (i.e., 71-fold higher than that of the wild strain) (Li et al. 2018). The crude enzyme thereafter was utilized in the conversion of alkali-pretreated corn stover into bioethanol (54.2 g/L) through a 12 h simultaneous saccharification and fermentation with S. cerevisiae as ethanologen (Li et al. 2018). Kataria and Ghosh (Kataria and Ghosh 2011) studied the hydrolysis activity of an enzyme mixture from T. reesei on the degradation of Kans grass for its subsequent conversion into bioethanol using S. cerevisiae (yield, 0.46  g/g; theoretical yield, 91.98%). The greatest cellulase enzyme production (1.46 U/mL) was achieved at 28 °C in the presence of cellulose as the carbon source. There are two strategies for developing consolidated bioprocessing organisms, (i) conversion of a potent cellulase producer such as T. reesei into an organism with the ability to produce bioethanol and (ii) conversion of a powerful ethanologen organism like S. cerevisiae into a cellulose hydrolytic enzyme producer. In the former, the pathway required for bioethanol production must be introduced into the cellulolytic organism or improved if it naturally exists. Moreover, tolerance to ethanol must be checked in the wild strain. In the latter, on the other hand, the genes encoding cellulases must be introduced into the ethanologen organism to transfer the ability for degradation of lignocellulosic biomass into monomeric sugars. Trichoderma naturally possesses biochemical pathways required for bioethanol production; however, the bioethanol production must be improved with respect to yield and concentration. By introducing the related heterologous genes, bioethanol production may be enhanced as shown by Xu et al. (2015). According to their findings, increasing the activity of pyruvate decarboxylase and alcohol dehydrogenase, i.e., two key enzymes in bioethanol production pathway through transforming the genes ScPDC1 and ScADH1 from S. cerevisiae, improved the concentration of the produced bioethanol by the engineered strains of T. reesei, compared to the wild type. It is worth mentioning that T. reesei, as an obligate aerobic organism, could not be used as an efficient consolidated bioprocessing organism. Therefore, cellulose hydrolysis must be initiated under aerobic condition, then switching into anaerobic condition when the

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organism has adequate enzymes to initiate glycolysis or fermentation process. To address this issue, an efficient genetic engineering approach is needed to change or delete the oxygen-dependent transcriptional control of glycolysis pathway (Rautio et al. 2006). Similar to T. reesei, many Trichoderma spp. (such as T. harzianum) have superb cellulase production abilities, and hence, the potential for a consolidated bioprocessing of lignocellulose into bioethanol through genetic engineering techniques. However, the development of engineered T. reesei may be more successful than other engineered Trichoderma spp. due to the fact that (i) the required systems for gene manipulation and expression have already been developed and (ii) it has already been exploited at commercial scales for cellulase production, contributing to its more readily acceptance and adaptation by the industry (Xu et al. 2015).

4.6.5  Aspergillus oryzae Aspergillus oryzae (A. oryzae) is another biocontrol food-grade fungus capable of producing a broad spectrum of hydrolyzing enzymes. As an example, Sandhu et al. (2012) reported the production of carboxymethyl cellulase (i.e., endoglucanase), β-glucosidase, and xylanase, exopolygalacturonase by A. oryzae following glucose depletion. They extracted crude cellulase from the fungal culture medium which could be used for saccharification of C. reticulata. Accordingly, a 3-h saccharification process (50 °C, cellulase activity of 3 FPU/g dry substrate) led to the formation of sugars, mainly including glucose, 24.87  g/L; fructose, 21.98  g/L; sucrose, 10.86 g/L; and galacturonic acid, 6.56 g/L. In another study (Kotaka et al. 2008), two genes encoding endoglucanase were cloned from A. oryzae into S. cerevisiae. The recombinant enzyme (i.e., β-glucosidase) was successfully expressed on the cell surface of the engineered yeast (S. cerevisiae GRI-117-UK/pUDB7), allowing a direct bioethanol production from barley β-glucan (biomass, 20  g/L; retention time, 24 h; ethanol theoretical yield, 69.6%; and ethanol concentration, 7.94 g/L) and other substrates containing cellobiose. Genetic engineering techniques could also be used for performance improvements of A. oryzae cellulase. In this context, the modified cellulase cocktail from the recombinant A. oryzae could deliver a saccharification yield (glucose concentration, 752 mg/L) of up to 2.5 times higher than that of Cellic CTec2 from a bleached-kraft pulp (containing 73% w/w glucan) (Yamada et al. 2015). More specifically, the modified cellulase cocktail contained four types of cellulases including (i) endoglucanases, 99 U/mL; (ii) cellobiohydrolase I, 3.8 U/mL; (iii) cellobiohydrolase II, 1.9 U/mL; and (iv) β-glucosidases, 43 U/ mL. The application of this modified enzyme cocktail in a simultaneous saccharification and fermentation of kraft pulp slurry (3% solid content) with yeast as the ethanologen delivered a bioethanol yield of 9.7 g/L in 24 h (Yamada et al. 2015). A. oryzae could efficiently assimilate sugars (e.g., xylose) into bioethanol which is not metabolized by wild S. cerevisiae. Similar to other filamentous fungi (e.g., Fusarium spp.), A. oryzae biomass could be easily separated from the fermentation medium, particularly using distillers dried grains with solubles. The condensate could even be recycled back as inoculants into bioethanol production plants. The

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co-cultivation of A. oryzae with other molds or yeasts could improve sugars and bioethanol yields obtained from lignocellulose processing. On this basis, a 10-d consolidated bioprocessing was reported in which A. oryzae was paired with S. cerevisiae for a successful hydrolytic enzyme secretion (saccharification) and bioethanol production (37 g/L), respectively (Wilkinson et al. 2017). Recently, A. oryzae has been co-cultured with T. reesei for saccharifying steam-exploded sugarcane bagasse (Maehara et al. 2018). Compared to the pure cultivations, up to 50% higher amount of glucose could be prepared via the co-cultivation technique. Through a solid-state fermentation (32 °C, 70% moisture, 48 h), bioethanol productivity and theoretical yield of up to 4.77 g/L/h and 83.5% could be obtained by the mentioned fungal pairing in the presence of 0.5% w/w soybean protein (as lignin blockers, saving enzymes from unproductive adsorption) (Maehara et al. 2018).

4.6.6  Pochonia chlamydosporia After cellulose, the second most abundant biopolymer is chitin, mainly coming from aquaculture as wastes (Hamedi et al. 2015). This biopolymer could make up 60–80% of the weights of crab and shrimp bodies. Despite the great potential of chitin as biomass, only a few percent of it is currently recycled in the form of cheap crab and shrimp foods (Hamedi et  al. 2015). Chitinases and chitin deacetylases could convert chitin into chitosan through direct hydrolysis and deacetylation, respectively. For energy and environment protection purposes, entomopathogenic and/or nematophagous fungi could be exploited for the degradation of chitin/chitosan into reducing sugars, efficiently turning this abundant bio-residue into a promising bioethanol feedstock. On this basis, Aranda-Martinez et al. (2017) investigated the capabilities of P. chlamydosporia (a biocontrol fungus with nematophagous activity) (Moosavi et al. 2010; Zare and Gams 2003), for the degradation of chitin into reducing sugar and bioethanol production. The fungus could use acid-dissolved chitosan (pH 5.6, autoclaved at 120 °C for 20 min) as the sole nutrient source mainly under anaerobic conditions, showing a strong chitosanolytic activity with fungal colony radius to chitosan degradation radius of about 0.5 in 20 days. The production of the reducing sugars from 2 to 3 g/L pretreated-chitosan reached its maximum at around day nine. Interestingly, P. chlamydosporia could directly convert pretreated chitosan into bioethanol while showing a favorable ethanol tolerance (at least 9 g/L ethanol concentration) (Aranda-Martinez et al. 2017).

4.7  Concluding Remarks Some fungal biocontrol species could serve as promising platforms for producing various hydrolytic enzymes in large quantities, rendering natural tolerance to harsh conditions and lignin-derived toxic substances. These enzymes could be subsequently consumed for the saccharification of delignified biomass through a separate

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saccharification and fermentation process, while the fungal biomass could be used in various types of pesticides for pest control strategies. Fungal pretreatment could also be integrated with fermentation to allow simultaneous saccharification and fermentation. Moreover, the bioethanol production ability of some fungi such as F. oxysporum could be improved (through recombinant technology) and exploited for mixed fermentation of hexose and pentose either alone (as the only ethanologen) or in associations with other ethanologens (such as S. cerevisiae). The latter two strategies remove the repression signaled from the accumulation of end products (e.g., glucose, xylose, and cellobiose) on enzymatic reaction through their continuous assimilation into bioethanol, and hence, improving both sugars and consequently bioethanol production yields. It is worth mentioning that consolidated bioprocessing of lignocellulose into bioethanol is gaining more attention. If it appropriately matures, the technology could significantly reduce the capital and operating costs by using a single reactor for enzyme production, saccharification of delignified biomass, and bioethanol fermentation. Some filamentous fungal biocontrol species are promising microorganisms for successful conversion of lignocellulose into bioethanol under consolidated bioprocessing approach due to their efficient enzymatic systems that allows direct utilization of plant cell-wall biomass into bioethanol. In addition to their ability for large quantities of hydrolytic enzyme production, fungi are generally more appropriate for optimum enzymatic hydrolysis (40–50 °C) because they could thrive at higher temperatures (37  °C), compared to hydrolytic bacteria. In addition to the higher hydrolytic enzyme (particularly cellulases) production, filamentous fungi are also more appreciated than other microorganisms for bioethanol production due to the ease of separation and inoculum recycling. However, the bioethanol production rate and or yield as well as fungal growth rate must be sufficiently improved through genetic engineering techniques for an economically feasible consolidated bioprocessing. Moreover, for the successful simultaneous saccharification and (co)fermentation, or consolidated bioprocessing, some other physiological characteristics of fungal biocontrol species such as low ethanol tolerance may require modifications and improvements. Overall, careful selections of right fungal biocontrol species/strain for lignocellulose-based bioethanol production is crucial for obtaining the best possible outcomes. For example, T. reesei is one of the most potent cellulase-producing filamentous fungal biocontrol species; however, it cannot currently be utilized for consolidated bioprocessing of lignocellulose into bioethanol as the genes encoding glycolysis enzymes are only expressed in the presence of oxygen. Consequently, this prevents an efficient production of bioethanol from soluble sugars by this microorganism unless genetic engineering techniques are employed. Finally, the environmental and economic feasibilities of bioethanol production from lignocellulose could be improved by using resistant microorganisms toward the extreme processing conditions (e.g., low pH, high temperature, and high toxic compound concentration). On this basis, for example, P. kudriavzevii with an optimum growth temperature of up to 45 °C could minimize production costs by reducing contamination risk and cooling requirement while improving bioethanol

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productivity rate and product recovery. Similarly, P. anomala could grow at pH 3, decreasing contamination risk and minimizing the need for pH adjustment. Toxic-­ resistant fungi (e.g., P. variotii and P. kudriavzevii) could bioremediate inhibitory compounds that may not be assimilated by usual ethanologens, hence detoxifying substrates, allowing efficient bioethanol production.

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Chapter 5

Endophytic Fungi for Biodiesel Production Cristiano E. Rodrigues Reis, Heitor B. S. Bento, Ana K. F. Carvalho, Yan Yang, Heizir F. de Castro, and Bo Hu

5.1  Background The utilization of biomass as feedstock for generation of liquid fuels has been a reality for decades, having had its breakthrough growing during the world petroleum crises in the 1970s (Ratledge 1993). Throughout the last decades, there have been numerous advances in conversion of biomass and plant-based materials, like lipids to biofuels, in order to lessen the energy dependence on nonrenewable feedstock. More recently, however, the utilization of microbial-based biomass and lipids has been evaluated as a valuable alternative to vegetable crops within an energy context (Ratledge 2013). Primarily within the context of liquid bio-based fuels, ethanol and biodiesel stand as the biofuels with the largest production scales, which are correlated with a strong dependence on the production of sugarcane, maize, soybean, rapeseed, and palm, for example. Though numerous advances to regional economies have been made by the enhancement of agricultural practices by intensive agricultural production, a strong debate on the impacts of crop cultures for bioenergy has been thoroughly discussed in the political, social, and scientific spheres of society (Goldemberg et al. 2008; Hill et al. 2006). Among the many factors involved in the discussion on the feasibility of a crop-bioenergy industry, the debate on whether or not such bioenergy would provide energy independence is often raised as a pivotal point (Hill et al. 2006), as there are often numerous political and economic powers that dictate how the production of agricultural crops should

C. E. Rodrigues Reis · H. B. S. Bento · A. K. F. Carvalho · H. F. de Castro Chemical Engineering Department, Engineering School of Lorena, University of São Paulo, Lorena, São Paulo, Brazil Y. Yang · B. Hu (*) Department of Bioproducts and Biosystems Engineering, University of Minnesota, Saint Paul, MN, USA e-mail: [email protected] © Springer Nature Switzerland AG 2020 G. Salehi Jouzani et al. (eds.), Fungi in Fuel Biotechnology, Fungal Biology, https://doi.org/10.1007/978-3-030-44488-4_5

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be made, and therefore, these are related to a relatively uncertain scenario within a medium- to long-term future. The utilization of agricultural residues as bioenergy feedstock has also been thoroughly analyzed by the scientific community (Sánchez 2009), which still provides numerous groundbreaking discoveries. However, key discoveries that would enable greater feasibility on the utilization of lignocellulosic material as feedstock for bio-­ based liquid fuels still remain unclear, particularly on the understanding of biomass deconstruction on an industrial scale (Banerjee et al. 2010). As a complementing alternative to the utilization of renewable energy, the conversion of biomass and products derived from microorganisms stands as a favorable alternative, particularly in the case of oil- and fat-based biofuels (Ratledge 1993). Some species of microorganisms are able to convert carbon to lipids through their growth cycle, with special attention to some algal and fungal strains that are able to accumulate triacylglycerol-­ rich bodies within their cells (Sitepu et al. 2014). Though the accumulation of lipids in microbial cells is often triggered by biochemical pathways that are related to energy storage for eukaryotes, the understanding of the engineering and biological aspects governing the production of microbial oils has been widely explored in order to achieve high rates of lipid production to provide favorable numbers on their viability as feedstock for biofuel production (Martínez et al. 2015). Oleaginous microorganisms are often defined as those strains that are able to accumulate lipids above 20% of their biomass weight on a dry basis (Sitepu et al. 2014). A diverse number of microalgae species, as well as filamentous fungi and yeasts, have been thoroughly analyzed over the last decades with respect to their potential of accumulating lipid bodies, which are reported to be as high as 70 wt.% of their biomass in some species of fungi under high-stress cell cultures (Beales 2004). Particularly for fungi, the composition of lipids commonly obtained is of similar chemical composition to those of plant and animal oils and fats, but they do not compete or pose any direct effects on the utilization of food resources (Beales 2004). Many species of fungi can also utilize inexpensive carbon sources, such as lignocellulosic residues and other by-products, in order to produce and accumulate lipid bodies. Lipid-rich cells are often known as single cell oil (SCO) and have a typical short process cycle, not being subjected to weather and other factors involved in traditional crop production (Li et al. 2008; Liang and Jiang 2015). Fungal SCO has been widely studied since the 1970s, but only within the last decades, the conversion of SCO to liquid biofuels, like biodiesel, has been explored in greater depth (Ratledge 2013). For instance, over 1500 yeast species from more than 100 genera have been evaluated, and only about 30 of those are able to accumulate over 25% of their dry biomass weight as lipids (Botham and Ratledge 1978, 1979). Particularly, basidiomycetes and ascomycetes prevail among SCO-producing yeasts, with a focus on the Yarrowia, Candida, Rhodosporidium, Rhodotorula, Cryptococcus, and Lipomyces genera (Ratledge 2013). The exploitation of SCO production using filamentous fungi has a similar history to the yeast SCO production, albeit the focus of exploitation of the SCO production by filamentous fungi was primarily based on the high concentrations of polyunsaturated fatty acids (PUFA) in their lipid bodies (Gouda et  al. 2008; Martínez et  al. 2015; Xu et  al.

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2012). Mortierella alpina, one of the common strains used in fungal SCO production, is able to produce oils containing n-1, n-3, n-4, n-6, n-7, and n-9 PUFA, while the zygomycete Mucor circinelloides represents an outstanding model for the production of PUFA-rich SCO with nutraceutical value (Li et  al. 2008; Meng et  al. 2009; Vicente et al. 2010). Among some of the studied species for SCO production, the class of endophytic fungi has been recently considered as an interesting group to be analyzed for their related bioenergy value. Endophytic fungi or endophytes are fungi cells that commonly inhabit plant tissues and coexist in synergy with the plant cells (Strobel and Daisy 2003). It is estimated that endophytic fungi can be easily incorporated into the different stages of growth during a plant life cycle, being able to grow, for instance, within roots, stems, leaves, flowers, fruits, and seeds of plants (Strobel et  al. 1996). Though endophytes are commonly studied for their ecological role, especially on their interaction with the host plants, the understanding of the biotechnological prospect of their utilization remains an area with limited understanding. Endophytes provide a diversity of secondary metabolites, which are commonly absent in wild strains, and have been used as biological control agents to plant diseases, to provide bioactive metabolites for pharmaceuticals and other metabolites of chemical value (Stadler and Schulz 2009). Energy-related applications using endophytes are related to their production of SCO-like structures, as some endophytic fungi are able to produce metabolites of similar structure to their plant host, with a particular interest in oil-rich plants that are promising hosts for oleaginous fungal strains (Bhagobaty 2015). Within this context, this chapter summarizes the state-of-­ art of endophytic fungi within a bioenergy approach, with a particular focus on their role in the biodiesel production chain.

5.2  E  cology of Endophytic Fungi and Production of Enzymes and Other Metabolites The term “endophyte” encompasses all organisms living within plant tissues (Petrini 1986). From the late 1980s onwards, the knowledge on the biology of endophytic fungi had a major breakthrough in the scientific community, with critical reviews describing the role of evolution (Carroll 1988; Clay 1988), taxonomy (Petrini 1986), and the overall biology of these organisms (Carroll 1986; Clay 1990; Petrini 1991). The studies on the physiology and biology of grass endophytes, in sum with the evaluation of endophytes present in conifers, have led the understanding of term “endophytic fungi” as a class of mutualism. Petrini (1991) describes endophytes as being all those organisms that colonize symptomlessly the living internal tissues of their host during a more or less long period of their life. Considering the biodiversity of a given plant system, more than one endophytic species is likely to be present at a given time, which can naturally provide more than one isolated-prone strain at a particular plant tissue. However, while plant tissues host complex fungal

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communities with a functional organization, no biological model available has evaluated the interaction among endophytic species inhabiting the same tissue. The term “assemblage” (Petrini 1991) demonstrates that, while mutualistic relations occur in plant tissues, it is hypothesized that such interactions occur between endophyte and plant, not between different species of endophytes. From a taxonomic point of view, the endophytes present in roots and aerial plant organs belong to a vast majority to the Ascomycota group, while only a limited number of Basidiomycota or other taxa have been reported (Schulz and Boyle 2005). Endophytic fungi are virtually present in all organs of a given plant host, while some are even seed-borne (Carroll 1986). The formation of host-specific endophytic strains, for instance, is interpreted as a form of ecological adaptation, which may lead to different physiological traits within two or more morphologically indistinguishable strains of the same species. In addition, the specificity regarding organ colonization by endophytes is linked to adaptation to the particular microniche and the physiological conditions present in such organ (Beales 2004). While a large number of species can be practically isolated from a given host, only a small amount of those species are usually present as endophytic fungi. In other words, while the total number of taxa isolated from a given plant host may be large, only a small fraction of those can be considered dominant, which usually correlate with the fact of those having adapted to the endophytic life (Bhagobaty 2015). An example of the relationship between an endophyte and its host and the morphological aspects of an isolate is presented in Fig. 5.1. The successful colonization of a given plant host by a microbial species is dependent on a number of factors, such as temperature, nutrient availability, and the crucial factor related to the physical phenomenon of penetration onto the plant layers by the microorganism (Braga et al. 2016). The physical factor of adherence is linked to the availability of mechanical fractures of the protective tissues or by the enzymatic degradation of the cuticular and epidermal layers from the plant host. It has been demonstrated that most endophytes are able to utilize the nutrient sources present in their microenvironment, either present on the surfaces or on the cell wall of their host (Strobel et al. 2011). Evolutionary conditions led to the observation that most endophytes are able to not only have their cell growth supported by pectin and xylan but also to produce laccase and nonspecific peroxidases and commonly present lipolytic activity (Dey et al. 2011). Endophytes are also commonly reported as producers of extracellular cellulases and hemicellulases, while the presence of amylases is seldom reported on endophytic fungi (Strobel et al. 2004). Table 5.1 summarizes some of the species of endophytes and their hosts in regard to cellulase production. The production of enzymes by endophytes that are either dormant or nonexistent in wild strains is also reported in the literature, as is the case of production of some nonspecific C4 and C8 esterases in some Melanconium and Aspergillus isolates, which are relevant for the penetration of the epiderm layers of the plant host (Petrini 1991). However, isolates of the same species derived from the same host present consistency with respect to their enzymatic activities, which confirms the fact of the host-specific strain formation taking place within the same fungal species (Petrini

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Fig. 5.1  Morphological characteristics of endophytic fungi. (a) Microscopic aspect of hyphae and spores of endophyte isolated from Trichilia elegans. (b) Macroscopic visualization of endophytic fungus isolated from Trichilia elegans. (c) Histologic visualization of leaf showing endophyte hyphae. (Reproduced with permission from Corrêa et al. 2014)

1991). The presence of a given enzymatic activity may not be linked, though, to a particular endophytic fungal species, as, for instance, enzymes linked to the degradation of pectin and poly galacturonic acid are widespread among endophytes, as such enzymes are responsible for the partial degradation of the middle layer of the host cell wall (Saikkonen et al. 1998). In addition to the production of cellulases, another group of enzymes that are of special importance within the context of biofuel production is that of lipases. Lipases, which are a category of hydrolytic enzymes that act on the reversible reaction of hydrolysis of the ester bond present in tri-, di-, and monoacylglycerols, are of particular interest to the production of biodiesel, since these enzymes are able to catalyze either transesterification reaction of triacylglycerols to alkyl esters or the esterification reactions from free fatty acids with an alcohol. Part of the literature on lipase screening from endophytes is summarized in Table  5.2. The research conducted by Torres et al. (2003), for example, rendered the report of a lipase produced by endophytic Rhizopus oryzae, which was able to catalyze the esterification of free fatty acids in an iso-octane-containing reaction medium. Torres et al. (2003) reported the lipase with an optimum pH range between 4 and 7 with thermophilic behavior.

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Table 5.1  Cellulase and xylanase production by different endophytes and their respective host plants Host plant Costus igneus, Lawsonia inermis Coleus aromaticus Opuntia ficus-indica Mill. (Cactaceae) Opuntia ficus-indica Mill. (Cactaceae) Opuntia ficus-indica Mill. (Cactaceae) Opuntia ficus-indica Mill. (Cactaceae) Opuntia ficus-indica Mill. (Cactaceae) Osbeckia stellata

Penicillium aurantiogriseum Mortierella hyalina

Camellia caduca

Penicillium sp.

Acanthus ilicifolius

Alternaria chlamydospora Aspergillus sp. Acremonium zeae

Acrostichum aureum Zea mays

Endophytic fungus Cladosporium cladosporioides Curvularia vermiformis Acremonium terrícola Aspergillus japonicus Cladosporium cladosporioides Fusarium lateritium

Enzyme Cellulase

References Amirita et al. (2012)

Cellulase Cellulase, xylanase Cellulase, xylanase Cellulase, Xylanase Cellulase, xylanase Cellulase, xylanase Cellulase, xylanase Cellulase, xylanase Cellulase

Amirita et al. (2012) Bezerra et al. (2012)

Bhagobaty and Joshi (2012) Bhagobaty and Joshi (2012) Maria et al. (2005)

Cellulase Xylanase

Maria et al. (2005) Bischoff et al. (2009)

Bezerra et al. (2012) Bezerra et al. (2012) Bezerra et al. (2012) Bezerra et al. (2012)

A thorough analysis of the ecology of endophytic fungi was carried out using soybean husks and grains from croplands in the state of Minnesota (USA). The methodology described by Yang et al. (2014) followed harvesting and sterilization of soybean samples, milling, and incubation on potato dextrose agar plates at 27 °C. Internal transcribed spacer (ITS) regions identified the fungal hyphae isolated from the plates, from which genomic DNA was extracted. Amplified products were sequenced by Sanger sequencing and compared with sequences in the National Center for Biotechnology Information (NCBI) nucleotide collection. The study described that 33 fungal isolates were obtained from soybean samples taken during the summer, from ten different genera. The most abundant genera found in the samples were Fusarium, Alternaria, Penicillium, and Nigrospora, encompassing 18 different species, later confirming that similar species have been detected as endophytes in soybean samples. The authors also conducted soybean sample harvesting during the fall, which albeit promoted a similar distribution of genera, the frequency of culturable isolates decreased by approximately half when compared to the summer sample, that is, from 33 to 17, demonstrating indications for the spatial and temporal changes in the fungal community of endophytes. A. alternata strains were found abundantly in both samples, corroborating with previous reports on the fast growth and robustness of this fungal species. The plant-associated endophytes described by Schulz and Boyle (2005) have been widely isolated from a number of environments and curiously have been primarily recognized as pathogens to their host plants by

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Table 5.2  Examples of lipase-producing endophytic fungi and their respective plant host Species Rhizopus oryzae

Host Mediterranean plants

Cercospora kikuchii

Tithonia diversifolia

Aspergillus niger, Chalaropsis thielavioides, Colletotrichum gloeosporioides, Lasiodiplodia theobromae and Phoma glomerata Curvularia brachyspora Curvularia vermiformis Drechslera hawaiiensis Colletotrichum falcatum Phyllosticta sp.

Seeds (castor, coconut, neem, peanut, pongamia, rubber, and sesame) of Tamil Nadu (India) Adhatoda vasica Coleus aromaticus Adhatoda vasica Lawsonia inermis Adhatoda vasica, Lawsonia inermis Potentilla fulgens Osbeckia stellata Camellia caduca Schima khasiana Acrostichum aureum Acanthus ilicifolius Acrostichum aureum Acrostichum aureum Acanthus ilicifolius Acrostichum aureum Acanthus ilicifolius Bertholletia excelsa Theobroma cacao Amazon wood

Talaromyces flavus Mortierella hyalina Penicillium sp. Penicillium sp. Acremonium sp. Alternaria chlamydospora Alternaria sp. Aspergillus sp. Aspergillus sp. Fusarium sp. Pestalotiopsis sp. UEA_001 UEA_007 UEA_115

Lipase Mycelium-bound lipase and extracellular enzymes Extracellular enzymes Extracellular lipase

References Torres et al. (2003)

Costa-Silva et al. (2017) Venkatesagowda et al. (2012)

Extracellular lipase

Amirita et al. (2012)

Extracellular lipase

Bhagobaty and Joshi (2012)

Extracellular lipase

Maria et al. (2005)

Mycelium-bound Zonotto et al. lipase (2009)

other fungal community studies. However, as described by Schulz and Boyle (2005), endophytes usually maintain a finely balanced relationship with a healthy host, demonstrating no signs of pathogenicity as long as fungal virulence and host defense mechanisms are absent. Yang et al. (2014) reported the endophyte community present in soybean having Fusarium, Alternaria, Penicillium, Nigrospora, Cercospora, and Epicoccum. While, for instance, some Fusarium species are commonly found as plant pathogens, that is, producing a range of phytotoxic compounds, for example, fusaric acid, fumonisins, moniliformin, and some trichothecenes, that are known to cause adverse effects on the plants, such as necrosis and inhibition of seed germination and plant growth, other Fusarium species have been reported to actually support cell growth and promote control to plant diseases. Similar behavior is found among within some

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species of the genus Alternaria, which produces metabolites that control plant diseases, while other species are known to induce leaf blight and leaf spot diseases. Penicillium, Nigrospora, Cercospora, and Epicoccum also have been reported to produce phytochemicals that act as biocontrol agents for plant pathogens, while some other species from these genera are known to be producers of common mycotoxins. Penicillium, for instance, is known to be producers of citrinin, ochratoxin, and mycophenolic and penicillic acids, all of which are related to post-harvesting diseases in plants. It is important to highlight that the major research methods on fungal communities can be categorized as culture-dependent and culture-independent methods (Gherbawy and Voigt 2010). The first uses a number of prepared growth media for testing the development of fungal hyphae, being a rapid and low-cost method for recovering a large number of plant-associated fungal species, including endophytes. Culture-dependent methods, however, are linked to the successful growth of a few species that are able to have their growth supported in laboratory-prepared media. On the other hand, culture-independent methods (metagenomics techniques) allow full profiling of the complex microbial community and the evolutionary dynamics of the micro-environment (Gherbawy and Voigt 2010). Culture-independent methods may detect unculturable, low-abundance, and slow-growing microorganisms, which provide a less-biased picture of the biota when compared to culture-­dependent methods (Cocolin et al. 2007). Despite some of the technical advantages of the latter, culture-dependent methods are usually preferred by the scientific community when dealing with endophytes for SCO production, as slow-growing or unculturable endophytes may pose difficulties in scale-up operations (Yang et al. 2014).

5.3  B  iochemistry of SCO Production and Fatty Acid Profile of Endophytic Fungal SCO The accumulation of lipids in oleaginous fungi is linked to the limitation of a nutrient in the medium, commonly nitrogen, with a direct surplus of carbon (Botham and Ratledge 1979). While nitrogen presence is a key factor in regulating cell growth, as the production of proteins, amino acids, and nucleic acids is directly dependent on the presence of nitrogen compounds, its limitation decreases the kinetics of cell division, and the carbon flux is directed toward the synthesis of lipids, leading to the accumulation of triacylglycerols within intracellular lipid bodies (Li et al. 2008). The pathways leading to high amounts of lipids are dependent on the regulation of the biosynthetic pathways via the supply of the precursors acetyl-CoA, malonyl-­ CoA, and glycerol-3-phosphate, and NADPH as a cofactor. The de novo synthesis of fatty acids, which is the first step of intracellular lipid accumulation, occurs in the cytosol by the fatty acid synthetase complex, which activates the acyl carrier protein through the phosphopantetheine transferase enzyme by loading the coenzyme pantothenate. The fatty acid synthetase first loads acetyl-CoA on its beta-ketoacyl-acyl

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carrier protein synthase, and then it exerts activities on beta-ketoacyl-acyl carrier protein reductase, beta-hydroxy acyl-acyl carrier protein dehydratase, and enoyl-­ acyl carrier protein reductase enzymes. The set of reactions is repeated until palmitoyl-­acyl carrier protein is produced. The constant supply of acetyl-CoA, which first acts as an initial biosynthetic unit, and malonyl-CoA as the elongation unit, promotes elongation of two carbons at each reaction step, having the acetyl-­ CoA produced by citrate cleavage in the cytosol. Fatty acid synthesis in oleaginous yeasts and filamentous fungi generates final products through the fatty acids synthetase complex myristic or palmitic acids, depending on the species. Modifications of myristic or palmitic acids occur in the endoplasmic reticulum and are catalyzed by elongases, which promote elongation reactions, or endoplasmic reticulum desaturases, which are hydrophobic membrane-­ bound proteins. One of the most common desaturation modifications is carried out by Δ9-desaturases, which inserts the first double bond onto stearic or palmitic fatty acids, as well as Δ12-desaturases, which catalyze the formation of the second unsaturation on oleic acid in order to generate linoleic acid. The de novo synthesis generates fatty acid-CoA, which is esterified with sterol or glycerol to produce steryl esters and triacylglycerols, respectively. Oleaginous fungi carry steryl esters and triacylglycerols as energy storage in their lipid bodies. Triacylglycerols are mostly formed by sequential acylation of glycerol-3-phosphate through the action of diverse acyl transferases. Glycerol-3-phosphate is formed by the action of glycerol kinase on glycerol and is synthesized from dihydroxyacetone phosphate by metabolic reactions catalyzed by a glycerol-3-phosphate dehydrogenase. The formation of 1-acyl glycerol-3-phosphate is carried either by the addition of the first acyl group on the structure or through the reduction of acyl-­ dihydroxyacetone phosphate, catalyzed by an NADPH-dependent reductase. A subsequent acyl group is added through the catalytic action of a second acyltransferase, producing 1,2-diacyl glycerol-3-phosphate, from which the phosphate group is removed through phosphatide phosphatase isoenzymes, producing a diacylglycerol structure. Diacylglycerol is used as the direct precursor to triacylglycerol on the last step of the de novo synthesis, which is carried by using diverse acyl donors, such as acyl-CoA. The acyl-CoA reaction with the diacylglycerol structure is catalyzed by endoplasmic reticulum-bound enzymes with acyl transferase activity, which directly load the third acyl-CoA onto the structure (Tang et al. 2015). Neutral lipid structures, such as triacylglycerols, are stored in specialized hydrophobic compartments known as lipid bodies, which are assembled at specialized subdomains of the endoplasmic reticulum, and maintained between the two layers of the phospholipid membrane bilayer of the cell (Ratledge 2013). As oil-rich cells often saturate the space between the leaflets of the membrane bilayer of the endoplasmic reticulum, excess neutral lipid synthesis generates structures that bud off from the endoplasmic reticulum as a mature lipid body once it reaches its critical size (Ratledge 2013). In most oleaginous yeasts and filamentous fungi, triacylglycerols are the main constituent of lipid bodies, reaching average concentrations of 90% in regard to the total neutral lipid weight, whereas a smaller fraction is composed by steryl esters. Within the core of a lipid body, in which neutral lipids are

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surrounded by a phospholipid monolayer, a number of neighboring proteins play a key role in lipid metabolic utilization. For instance, whenever storage lipids are required to be used as energy sources, lipases and steryl ester hydrolases act in order to hydrolyze the triacylglycerol and steryl ester structures, in order to provide energy sources or building blocks for cellular membrane formation (Botham and Ratledge 1978).

5.4  E  ndophyte-Plant Interactions and Screening of Oil-Rich Plants Endophytes are usually in a latent state within their host plant during for extended periods of time, until micro-environmental conditions allow the growth of the fungal cells to either propagate or become pathogenic to the host. Interestingly, the colonization of plant hosts has been demonstrated to induce plant adaptation to stress factors, as demonstrated by Redman et al. (2002) and Giordano et al. (2009). Particularly, the host tolerance to biotic stress has been correlated with the presence of fungal-derived metabolites, which have shown antimicrobial activity (Saikkonen et al. 1998; Strobel and Daisy 2003). It is noteworthy to acknowledge that the relevance of endophytes to the scientific community had its cornerstone in the early 1990s with the detection of paclitaxel (Taxol®) produced by an endophyte isolated from Taxus brevifolia (Stierle et al. 1993). The Taxomyces andreanae isolate was able to produce Taxol®, an anticancer drug, which was produced until then at lower quantities and at a lower rate by the host plant. The production of Taxol® by T. andreanae served as an initial point for discussion on horizontal gene transferring between host and endophyte or vice versa (Li et al. 1998). In the years to follow, numerous herbal compounds produced by endophytes were claimed in the literature, with a clear particular interest in pharmacologically important metabolites (Amna et  al. 2006; Eyberger et al. 2006; Kour et al. 2008; Puri et al. 2005, 2006). Gunatilaka (2006) described the large potential of endophytic fungi as natural reservoirs of unique chemical structures that have been adapted through natural selection and evolution involved in the host plant protection and communication. Through gene sequencing studies on the genetic clusters involved in fungal metabolism, the total number of expected natural products exceeds by far the number of metabolites claimed in the literature, as many of these genes are not expressed under laboratory culture conditions. The lack of host stimuli in laboratory prepared media may serve as an explanation to the production of some natural metabolites occurring only in their natural micro-niche (Li et al. 1998). From a bioenergetic point of view, the interaction between oil-rich plants and microorganisms could provide adaptational advantages toward the discovery of oleaginous fungi. Genetic studies on horizontal gene transferring between endophytes and plant hosts have been focused so far on the understanding of secondary metabolites, many of those with particular interest of the pharmaceutical industry. The

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production of lipids by endophytes seems to follow a similar metabolic pathway than in wild strains, with little discussion on the literature regarding the genetic answers on higher lipid productivities by endophytic fungi. In addition to the lipid structures, some hydrocarbons have been detected by the endophytic fungi. Gliocladium roseum obtained from Eucryphia cordifolia, which may have some potential utility in the syntheses of diesel-like structures. Nonetheless, from the perspective of a microbial lipid biorefinery, one should take into consideration the high production of lipid storage within fungal cells with suitable fatty acid profiles and the ability to grow on low-cost substrates. The secretion of lignocellulosic-­degrading enzymes by endophytes can also be considered as an adaptative trait as these are not important for the colonization inside the plant tissues but also serve as an indication of culturability on various lignocellulosic residues. In fact, the production of SCO from the endophyte Microsphaeropsis sp., isolated from the oleaginous plant Sabina chinensis, using solid-state fermentation using wheat straw, has been one of the first demonstrations of the concept regarding SCO production using endophytes on lignocellulosic residues. One of the earliest demonstrations of the feasibility of producing SCO using endophytic fungi with a posterior interest in converting the lipids to biodiesel was carried out by Dey et  al. (2011). Two different endophytic fungi isolated from Ocimum sanctum (which is a medicinal shrub with high concentrations of volatile oil in its leaves and seeds) and Brassica juncea (an oleaginous crop) were tested as SCO producers using solid-state fermentation and submerged growth assays on rice straw and wheat bran. The fungal strains evaluated in the study were Colletotrichum sp. and Alternaria sp., which have been previously described as oleaginous microorganisms. The results demonstrated by Dey et  al. (2011) show that a typical medium for SCO production with a high carbon to nitrogen ratio (C:N) promotes higher lipid accumulation by both fungal species. In fact, the highest lipid yields were attained at high initial concentrations of glucose (100 g L−1) and low concentrations of peptone (1.8 g L−1). The authors also described that the concentration of phosphorus also play a role in determining the lipid yield in a high-glucose-­ containing medium, demonstrating that the optimum carbon to phosphorus (C:P) differs for each strain, as C:P of 86:1 to Colletotrichum sp. and C:P of 675:1 for Alternaria sp. were determined as the nutrient conditions that promoted the highest yields of lipid accumulation. The presence of oleaginous fungi from isolates obtained from soybean samples by Yang et al. (2014) was relatively abundant. The authors discuss that 13 of the 33 harvested isolates had lipid contents greater than 20% when grown in potato dextrose agar medium, 5 of which have demonstrated lipid contents greater than 40% in regard to the dry biomass weight. Particularly some Fusarium strains, F. sporotrichioides, F. acuminatum, and F. equiseti, demonstrated high lipid accumulation, reaching levels around 46.4%. Soil samples obtained near the soybean plants were also analyzed and had been described to contain oil-rich fungal species, such as F. equiseti, and two Mucorales, M. hiemalis and M. circinelloides. It is interesting to highlight the dual role of Fusarium strains in recent studies, which have simultaneously been reported as a species to be controlled, due to their known production

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of mycotoxins, while other studies have reported some Fusarium species as promising candidates for fungal SCO production. It is important to highlight that the isolation and harvesting of endophytes from oleaginous plants pose an alternative to the search of SCO-producing microorganisms from the traditional spots of oil-polluted and oil-rich environments. While oleaginous fungi represent a minor proportion of the total fungal population, the discoveries to happen as endophytes may represent significant advances to the field of SCO and its integration into the bioenergy industry.

5.5  P  erspectives of Endophytic SCOs as Feedstock for Microbial Biodiesel Current sustainable bioenergy concerns include not only the search for fossil fuels alternatives but also the fact that the novel renewable options must be industrially competitive, economically feasible, highly productive, and not affect other important industrial chains, as feed and food industries. In light of these, the second- and third-generation biofuels strive not to compete or endanger the essential food chain, by focusing on the use of nonedible raw materials as biofuel feedstock (Ahmad et al. 2017; Alaswad et al. 2015). Biodiesel is chemically composed of monoalkylated fatty acid esters usually obtained from renewable sources such as vegetable oils and animal fats. In order to meet the previously mentioned concerns, microbial oils have been highlighted as one of the most promising alternatives to the conventional biodiesel feedstock in order to meet the increasing biofuel demands around the world. SCO obtained from filamentous fungi and yeasts have several advantages over other lipid sources, that is, not requiring arable lands for cultivation and not competing with food production, besides their rapid growth, light independency, and easy scalability (Subhash and Mohan 2015). Among the microorganisms that can accumulate high levels of lipids, endophytic fungi may become a reliable source due to their enormous biodiversity and adaptability to unique habitats. Some studies showed that endophytic fungi are able to produce the same metabolites as their hosts (Peng and Chen 2007; Stierle et  al. 1993; Strobel et al. 1996); therefore, isolated endophytic fungi in oleaginous plants are expected to have the ability to accumulate oil. It has been reported that certain endophytic fungi have the ability to accumulate lipids remarkably up to 35% of dry biomass (Peng and Chen 2007). Fatty acid profile of microbial oils is very similar to those from plants, therefore offering the possibility of the same applications, for example, as lipid source for biodiesel production (Carvalho et al. 2015). Table 5.3 illustrates the SCO production by different endophytes and their respective host plants. Natural biological characteristics of endophytic fungi growth on their natural habitat have launched their potential to intrincically produce hydrolytic enzymes as cellullases, usually allowing the assimilation of lignocellulosic substrates

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Table 5.3  Oleaginous endophytic fungi and their respective host plant Oil content Endophytic fungus (wt. %) Microsphaeropsis 33.6 sp. Sclerocystis sp. 29.5

Oil yield (mg g−1 initial dry substrate) 42

Nigrospora sp.

21.3

23

Phomopsis sp.

34.8

26

Phomopsis sp.

31.6

27

Phomopsis sp.

27.4

23

35.0

34

29.4

32

Glycine max

Cephalosporium sp. Microsphaeropsis sp. Fusarium equiseti

47.6

NR

O. sanctum

Colletotrichum sp.

49.1

NR

B. juncea

Alternaria sp.

58.1

NR

Host plant Sabina chinensis Taxus chinensis Keteleeria evelyniana Keteleeria evelyniana Pinus massoniana Lamb Keteleeria davidiana Cupressus torulosa Sabina chinensis

35

References Peng and Chen (2007) Peng and Chen (2007) Peng and Chen (2007) Peng and Chen (2007) Peng and Chen (2007) Peng and Chen (2007) Peng and Chen (2007) Peng and Chen (2007) Yang et al. (2014) Dey et al. (2011) Dey et al. (2011)

(Bhagobaty 2015). Since substrate costs are related to up to 75% of the overall cost of microbial biodiesel production, the use of highly available agro-industrial lignocellulosic wastes is an outstanding alternative to reduce the microbial biodiesel production costs and insert the process in the biorefinery context, involving the transformation of a waste or residue into a valuable product (Carvalho et al. 2019). Dey et al. (2011) reported two endophytic fungal isolates, Colletotrichum sp. and Alternaria sp., capable of accumulating lipid contents up to 58% of their dry biomass. The produced lipids were adequate for biodiesel synthesis. Both endophytes also grew successfully on combined rice straw and wheat bran with good enzyme activity (1.21–2.51 FPU g−1ds) and lipid accumulation (60.32–84.30 mg g−1ds). Peng and Chen (2007) screened 141 isolates of endophytic fungi from stems of 7 oleaginous plant species analyzing their lipid accumulation capacity. About 49% of the total isolates presented lipid bodies in their hyphae, and 26 isolates (belonging to five genera: Microsphaeropsis, Phomopsis, Cephalosporium, Sclerocystis, and Nigrospora) showed higher lipid contents. Their lipid content ranged from 21.3% to 35% of dry cell weight when cultured in potato dextrose broth. All analyzed species were able to produce cellulase and microbial oil (19–42 mg g−1 initial dry substrate) when cultured on solid-state medium composed of steam-exploded wheat straw (20 wt.%), wheat bran (5 wt.%) and water (75%).

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Therefore, the interest in connecting the concept of a microbial biodiesel industry with the potential of endophytic fungi as producers of lipids, lipase, and cellulases presents a great deal of synergy. For example, while some strains and culture conditions do not promote high lipid contents, some Fusarium strains are reported in the literature to accumulate up to 47.56% of oil content, which offers great potentials for biodiesel production by directly utilizing lignocellulosic materials as feedstock (Yang and Hu 2018). Santos-Fo et al. (2011) showed the potential of using endophytic fungi isolated from a number of different species of tropical plants in the biodiesel production through direct transesterification by acid catalysis and methanol. While the literature concerning different methods of biodiesel production from endophyte-based lipids is relatively scarce when compared to wild strains and mutant fungi adapted to specific media, it is noteworthy to acknowledge that the scientific community may, in the near future, for instance, use the vast knowledge of lipase and cellulase engineering to connect with the potential of lipid accumulation by endophytes in order to provide low-cost and reliable pathways for advancing the science of biodiesel.

5.6  Conclusions Endophytic fungi through their interactions with plants produce different high-­ value bioproducts, such as pharmaceuticals, vitamins, industrial enzymes, SCOs, and other metabolites. Among these compounds, SCOs, lipase, and cellulases could be used for biofuel production. More specifically, lipase could be used for transesterification of oils into biodiesel. SCOs have been taken into account as an alternative to food- and feed-grade oils for biodiesel production to address the food vs. fuel conflict. Oil-rich plants have been reported as microenvironments rich in SCO-­ producing endophytic fungal strains. Recent studies have shown the high potentials of different endophytic fungi such as Alternaria, Microsphaeropsis, Fusarium, Colletotrichum, Phomopsis, Cephalosporium, Sclerocystis, and Nigrospora in economic production of SCOs (20–50% of their dry weight). These SCOs produced by endophytic fungi have similar fatty acid profiles to those of plant oils and are qualified for biodiesel production. Overall, it could be concluded that the promising capabilities of endophytic fungi in producing lipids and lipase could offer great potentials for economic production of third-generation biodiesel.

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Chapter 6

Wood-Rotting Fungi for Biofuel Production Ichiro Kamei

6.1  Overview of White-Rot Fungi and Brown-Rot Fungi Wood-rotting basidiomycetes have an important role in the carbon cycle as decomposers in forest ecosystems. Wood is decomposed by a variety of biological agents, including fungi, bacteria, and insects. Fungi colonize wood and degrade cell wall components to form brown or white rot (Fig. 6.1). Lignocellulosic materials including wood are renewable sources of feedstock for biofuel production. Cellulose, hemicellulose, and lignin are closely associated in cell wall, and covalent cross-­ linkages have been suggested to occur between lignin and polysaccharides. Cellulose is a linear and highly ordered polymer (containing crystalline and amorphous region) of cellobiose, the dimer of glucose, that presents over 50% of wood dry weight. To use the cellulose for chemical and biological conversion, the first step is the removal of lignin from lignocellulosic material. Lignin is the high-molecular weight aromatic polymer derived from oxidative coupling of monolignols; the three main components are p-coumaryl, coniferyl, and sinapyl alcohols. Delocalization of radicals in lignin monomer to couple at different sites gives complexed carbon-­ carbon and carbon-oxygen bonds in polymerized lignin such as β-O-4, β-5, β-β, 5–5, 5-O-4, and β-1 linkage. Lignin has the plant-specific composition, and its linkage motifs vary according to the plant species and growth stage. These complex chemical structures and the variety of lignin give the recalcitrance to the wood cell wall. Lignin-degrading basidiomycetes, collectively referred to as white-rot fungi, are observed frequently on forest litter and fallen trees. White-rot fungi are the only microbes which are capable of efficient depolymerization, degradation, and mineralization of all components of plant cell walls as shown in Fig. 6.1. White-rot fungi have a unique ability to decompose the recalcitrant wood lignin via the secretion of I. Kamei (*) Faculty of Agriculture, University of Miyazaki, Miyazaki, Japan e-mail: [email protected] © Springer Nature Switzerland AG 2020 G. Salehi Jouzani et al. (eds.), Fungi in Fuel Biotechnology, Fungal Biology, https://doi.org/10.1007/978-3-030-44488-4_6

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Fig. 6.1  The photograph of white-rotting wood and brown-rotting wood

extracellular lignin-degrading enzymes such as manganese peroxidase (MnP), lignin peroxidase (LiP), and versatile peroxidase (VP). The biochemical reaction mechanisms and genetics of lignin-degrading enzymes have well been explained by several review papers (Martínez et al. 2005; Kersten and Cullen 2007; Hammel and Cullen 2008; Lundell et al. 2010). MnP is considered to be one of the key enzymes involved in lignin degradation caused by white-rot fungi (Kuwahara et  al. 1984; Glenn and Gold 1985). MnP oxidizes Mn2+ to Mn3+ in an H2O2-dependent reaction, and Mn3+ organic acid chelates oxidize several lignin model compounds and synthetic lignin via the formation of a phenoxy radical (Wariishi et al. 1989a, b; Tuor et al. 1992; Wariishi et al. 1991). Additionally, MnP participates in lignin biodegradation via thiol and lipid-derived free radicals that are able to oxidize a variety of nonphenolic aromatic compounds (Wariishi et al. 1989a, b; Bao et al. 1994). LiP was isolated from white-rot fungus Phanerochaete chrysosporium (Tien and Kirk 1983; Glenn et al. 1983) and is characterized by its ability to oxidize high redox-­ potential aromatic compounds such as veratryl (3,4-dimethoxybenzyl) alcohol, methoxybenzenes, and non-phenolic lignin model dimers (Kuwahara et al. 1984; Kersten et al. 1985; Umezawa et al. 1986). The importance of LiP in the depolymerization of lignin was previously shown (Hammel and Moen 1991). VP was isolated from white-rot fungus Pleurotus eryngii and is able to oxidize both LiP and MnP substrates (Martínez et al. 1996). VP oxidizes Mn2+ to Mn3+, degrades the nonphenolic lignin model dimer, and oxidizes veratryl alcohol as LiP dose in an H2O2-­ dependent reaction (Caramelo et al. 1999; Heinfling et al. 1998). These extracellular oxidizing enzymes are apparently keys for degradation of lignin by fungi; however, these enzymes cannot penetrate into the compact structure of wood tissues due to their large molecular size. Therefore, involvement of small chemical oxidizers including activated oxygen species or enzyme mediators is considered. In contrast, brown-rot fungi have different degradation mechanisms for wood cell wall compared with white-rot fungi. The degraded wood develops characteristic cubical cracks as shown in Fig. 6.1, because of extensive depolymerization of cellulose. Lignin is also modified slightly during cell wall degradation by brown-rot

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fungi but remains as brownish residues in degraded cell wall regions. Because removal of lignin is not essential, it is difficult to explain this degradation phenomenon by general cellulolytic enzymatic (cellulase) reaction. The wood degradation mechanism of brown-rot fungi could be explained by two kinds of non-enzymatic reaction in the early stage of wood rotting. Brown-rot fungi secret oxalic and other organic acids to reduce the pH of wood and depolymerize hemicellulose and cellulose through acid hydrolysis, contributing to an increase in the porosity of the wood cell wall (Espejo and Agosin 1991; Green III and Highley 1997). Because brown-rot fungi do not contain ligninolytic enzymes, such as LiP, MnP, and VP, it has been proposed that they use an extracellular reaction of H2O2 with Fe2+, known as a Fenton reaction, to modify the structure of wood cell wall. Fenton reaction may lead to the formation of hydroxyl radicals, the strongest oxidant known in biological systems so far (Koenigs 1974; Hammel et al. 2002). It is recognized that hydroxyl radicals randomly attack the wood cell wall constituents causing non-enzymatic disruption of the lignocellulose matrix with internal cleavage of cellulose chains and lignin modification (Arantes et  al. 2012). It is considered that these non-­ enzymatic reactions occurring at early stages of wood degradation improve the accessibility of saccharification enzymes to polysaccharides contained in rotting wood. Compared with brown-rot fungi, white-rot fungi have been studied more extensively owing to their lignin-degrading (removal) activity for their biotechnological applications such as biopulping (pretreatment of woody material to make pulp), bioremediation (degradation and detoxification of environmental contaminants), and biorefineries.

6.2  B  iological Pretreatment for Lignocellulose Saccharification Lignocellulosic biomass usually includes forest biomass and wastes, agricultural residues, and energy crops, which are mostly constructed by cellulose, hemicelluloses, and lignin. These natural resources have been focused on as potential materials for the second-generation bioethanol production because of their beneficial characteristics, including high annual yield, abundance, and high concentrations of holocellulosic contents (Cai et al. 2017; Zabed et al. 2016). The second-generation bioethanol production (from cellulose and hemicellulose) is recognized as a key topic in energy science and research because the results could contribute not only to energy production but also to the reduction of environmental pollution. The primary research goal of second-generation bioethanol production is to increase the polysaccharide conversion to monosaccharides and fermentation yields (Zabed et al. 2017). Sulfuric acid hydrolysis is one of the easy methods to produce monosaccharide from polysaccharides cellulose and hemicellulose; however, the inhibitory compounds for fermentation such as furan derivatives (furfural and 5-hydroxymethyl-­2furaldehyde (5-HMF)), phenolic compounds (syringaldehyde, vanillin, and other

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phenols), and acetic acid are present in the hydrolysates. Therefore, more studies have been concentrated on enzymatic saccharification and not on acid hydrolysis. Bioethanol production from lignocellulosic materials is often complex, with three main steps, including pretreatment, saccharification, and fermentation. The first step in bioethanol production is pretreatment. Successful methods were identified as the technologies able to alter or remove compositional impediments and resistant structures of biomass in order to improve enzymatic saccharification by using mechanical, chemical, or biological methods. Several pretreatment methods have been studied for various sources of biomass (Sun and Cheng 2002; Conde-­ Mejía et al. 2012; Suhardi et al. 2013), including steam explosion, solvent extraction, and thermal pretreatment using acids or bases (Mosier et al. 2005). Taking sugarcane bagasse, one of the most abundant agricultural residues in the world generated at staggering 540 million tons annually, is an example of lignocellulose pretreatment. Pretreatment methods investigated for bagasse include acid pretreatment with various acids (Rodríguez-Chong et al. 2004; Gámez et al. 2006; Cheng et al. 2008; Hernández-Salas et al. 2009), steam explosion (Hernández-Salas et  al. 2009), alkaline treatment (Hernández-Salas et  al. 2009), alkaline dewaxing (Peng et al. 2009), biological treatment (Li et al. 2002a, b), wet oxidation (Martín et al. 2007), organic solvent pretreatment (Pasquini et al. 2005; Pereira et al. 2007), and liquid hot water pretreatment (Laser et  al. 2002). Although various physical (comminution, hydrothermolysis), chemical (acid, alkali, solvents, ozone), and biological pretreatment methods have been investigated over the years (Kumar et al. 2009), there is little agreement on the most suitable process. Because of its mild conditions, compared with chemical and physical pretreatments, biological pretreatment has been regarded as a potentially effective approach. Biological pretreatments of lignocelluloses using white-rot fungi have been increasingly examined. The most important point of biological pretreatment is how to degrade (remove) lignin selectively and maintain cellulose in the residual biomass. P. chrysosporium, which is one of the model white-rot fungi to study wood degradation mechanism and related enzymes as described above, shows non-selective lignin degradation (non-selective lignin-degrading fungus degrades lignin with cellulose). After pretreatment of biomass by P. chrysosporium to degrade lignin, the saccharification rate of residual biomass was decreased in several reports (Shi et al. 2009; Keller et al. 2003). In these reports, significant degradation of lignin was observed; however, sequential enzymatic saccharification rate (from cellulose to glucose) tended to decrease. It is considered that the accessible cellulose region, that is, amorphous region, is utilized through pretreatment by this fungus along with lignin. Therefore, the remaining cellulosic material treated by P. chrysosporium may contain the recalcitrant regions of lignocellulosic material, that is, the crystalline regions of cellulose. On the other hand, there are some white-rot fungi which have been reported as selective lignin-degrading fungi, showing strong lignin degradation while maintaining cellulose. Table 6.1 shows the effect of pretreatment for several types of biomass by the well-studied white-rot fungi, Ceriporiopsis subvermispora, Pleurotus ostreatus, Trametes versicolor, Irpex lacteus, and Echinodontium taxodii. C. subvermispora is

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Table 6.1  Effect of pretreatment of white-rot fungi on main components in biomass Fungus Biomass LL Ceriporiopsis subvermispora Corn stover 39.2 Rice straw 18 Wheat straw