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Cellulases in the Biofuel Industry
 0323994962, 9780323994965

Table of contents :
Cover
Cellulases in the Biofuel Industry
Preface
List of tables
List of figures
Copyright
Contents
Acknowledgments
1 Background
1.1 Introduction
1.2 Bioenergy and biofuels
1.3 Biomass
1.4 Cellulase enzyme
References
Relevant Websites
2 Worldwide scenario of biofuel production
2.1 Introduction
2.2 Status of biofuel production
2.2.1 United States
2.2.2 Brazil
2.2.3 European Union
2.2.4 Argentina
2.2.5 China
2.2.6 Australia
2.2.7 Canada
2.2.8 Thailand
2.2.9 Japan
2.2.10 India
References
Relevant websites
3 Generations of biofuels
3.1 Introduction
3.2 First generation biofuels
3.3 Second generation biofuels
3.4 Third generation biofuels
3.5 Fourth generation biofuels
References
Relevant websites
4 Challenges to biofuel production
4.1 Introduction
4.2 Challenges in lignocellulosic biomass conversion to biofuels/biochemicals
4.2.1 Feedstock production and logistics
4.2.2 Lignocellulosic biomass pretreatment
4.2.3 Enzymatic hydrolysis
4.2.4 Microbial fermentation and biomass
4.2.5 Biofuel cost
4.2.6 Water recycling
4.2.7 Generation of coproducts
4.2.8 Energy and environmental issues
References
Relevant websites
Further reading
5 Current production status of cellulases and challenges
5.1 Introduction
5.2 Production of cellulases
5.2.1 Submerged fermentations
5.2.2 Solid-state fermentation
5.2.3 Sequential solid-state fermentation and submerged fermentation
5.2.4 Use of mixed cultures
References
Further reading
6 Cellulase market scenario
6.1 Introduction
6.2 Market scenario
References
Relevant websites
7 Roles of cellulases in cellulose hydrolysis
7.1 Introduction
7.2 Cellulase enzyme systems for cellulose hydrolysis
7.2.1 Endoglucanase
7.2.2 Exoglucanase
7.2.3 β-Glucosidase
7.2.4 Other cellulases and accessory proteins
7.3 Synergy among cellulose degrading system
References
Further reading
8 Cellulases for biofuels production
8.1 Introduction
8.2 Bioethanol production
8.2.1 Pretreatment
8.2.2 Hydrolysis
8.2.3 Fermentation
8.2.4 Distillation
8.3 Factors affecting bioethanol production
8.4 Important developments in the production of cellulosic ethanol
8.4.1 Case study
8.4.1.1 Project ABBK in Hugoton, Kansas, USA
8.4.1.2 Clariant sunliquid plant Podari, Romania
8.4.1.3 Praj’s second generation (2G) cellulosic ethanol plant
8.5 Other biofuels made by assistance from enzymes
References
Relevant websites
Further reading
9 Advanced developments in production processes of cellulases
9.1 Introduction
9.2 Advanced development in production processes of cellulases
References
Further reading
10 Cellulases and auxiliary enzymes
10.1 Introduction
10.2 Cellulases
10.2.1 Cellobiohydrolase
10.2.2 endo-1,4-β-Glucanase
10.2.3 β-Glucosidase
10.2.4 Cellulase auxiliary enzymes
References
Further reading
11 Approaches to enhance cellulase production to improve biomass hydrolysis
11.1 Introduction
11.2 Thermostable cellulases
11.3 Isolation and screening of efficient thermostable cellulase producing fungi
11.4 Enhancement of thermal and pH stability of cellulases in the presence of nanomaterials
11.5 Recombinant DNA technology for increasing cellulase activity and efficacy
11.6 Use of suitable carbon source and necessity of pretreatment of lignocellulosic biomass
11.7 Optimization of medium and process parameters using statistical methods for improved production of cellulases
11.8 Improvements in production of cellulases via microbial fermentation processes
11.9 Microbial co-production of other important enzymes for the overall economy of the process
References
12 Future prospects
12.1 Feedstock production
12.2 Feedstock logistics
12.3 Biofuels production
12.4 Biofuels distribution
12.5 Biofuels end use
References
Relevant websites
Further reading
Index

Citation preview

Cellulases in the Biofuel Industry

Cellulases in the Biofuel Industry

Pratima Bajpai Consultant-Pulp and Paper, Kanpur, Uttar Pradesh, India

Preface There is a major international effort to develop renewable alternatives to fossil fuels. One approach is to produce a liquid fuel by enzymatically hydrolyzing carbohydrate polymers in biomass to sugars and fermenting them to ethanol. Cellulose is the main polymer in biomass and cellulases can hydrolyze it to cellobiose, which can be converted to glucose by β-glucosidases. Extensive research is being carried out to obtain cellulases with higher activity on pretreated biomass substrates by screening and sequencing new organisms, engineering cellulases with improved properties, and identifying proteins that can stimulate cellulases. Despite extensive research on cellulases, there are major gaps in our understanding of how they hydrolyze crystalline cellulose, act synergistically, and for which role they act in carbohydrate binding modules. This book presents cost-effective and current scenarios for cellulase production in the biofuel industry, including the most recent advancements for obtaining cellulases with higher activity on pretreated biomass substrates by screening and sequencing new organisms, engineering cellulases with improved properties, and identifying proteins that can stimulate cellulases. The mechanism and efficiency of the cellulase enzyme system on cellulose are discussed with the specific classification of each cellulase enzyme, as well as explanations of the limitations of cellulases in terms of their production processes, efficiency, and practical applications to biofuels. Various approaches to improve the production and efficiency of the cellulase enzyme system have been evaluated along with the current limitations that are hampering cost-effective production of cellulase and guidance on how these limitations might be resolved. Pratima Bajpai

xv

List of tables Table 1.1 Table 1.2 Table 1.3 Table 1.4 Table 1.5 Table 1.6 Table 1.7 Table 2.1 Table 2.2 Table 2.3 Table 2.4 Table 2.5 Table 2.6 Table 2.7 Table 2.8 Table 2.9 Table 2.10 Table 2.11 Table 3.1 Table 4.1 Table 4.2 Table 5.1 Table 5.2 Table 6.1 Table 7.1 Table 8.1 Table 8.2 Table 9.1 Table 10.1 Table 11.1 Table 11.2

Most common biomass feedstocks. Benefits of biofuels. Greenhouse gas emissions from various biofuels compared to gasoline. Classification of cellulase enzymes. Fungi having cellulolytic abilities. Bacteria having cellulolytic abilities. Actinomycetes having cellulolytic abilities. Biofuel yields by feedstock. Annual world fuel ethanol production (Mil. Gal.). Historical biorefinery count and production capacity. Annual US fuel ethanol production. US fuel ethanol plant production capacity as of January 1, 2021. US biodiesel plant production capacity as of January 1, 2021. Global commercial scale cellulosic ethanol plants. The status of the US commercial lignocellulosic ethanol facilities. Operating cellulosic ethanol plants in the US. Commercial-scale bioethanol plants in Australia. Biofuel key figures. Yield of biofuels from different feedstocks of second-generation biofuels. Processes for the pretreatment of lignocellulosic biomass. Cost area distribution of a general lignocellulosic greenfield facility. Advantages and disadvantages of solid-state and submerged fermentation. Cellulase production under submerged and solid-state fermentation. Commercial cellulases, their sources and suppliers. Modular architectures of cellulases from different bacteria. Advantages and drawbacks of potential organisms in lignocellulosicbased bioethanol fermentation. Comparison between Simultaneous saccharification and fermentation and Separate hydrolysis and fermentation. Benefits of thermostable cellulases. Cellulase or cellulase encoding genes present in microorganisms. Thermostable cellulases from various thermophilic microorganisms and their characteristics. Cellulases from various sources immobilized on different nanosupports and their improved properties/applications.

3 4 5 12 16 17 18 26 27 28 29 29 30 31 33 34 43 49 60 72 75 99 100 113 122 151 154 181 206 217 223

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List of figures Figure 1.1 Figure 1.2 Figure 1.3 Figure 1.4

Figure 1.5 Figure 1.6

Figure 1.7

Figure 1.8 Figure 2.1 Figure 3.1 Figure 3.2 Figure 3.3 Figure 3.4 Figure 3.5 Figure 3.6 Figure 3.7 Figure 3.8

Structure of lignocellulosic biomass and its biopolymers; cellulose, hemicellulose, and lignin. Lignocellulosic biomass along with their products. The cellulose polymer chain structure. Schematic representation for the macromolecular structure of lignin (Major monolignol units are colored as sinapyl alcohol-red, guaiacyl alcohol-blue, p-coumaryl alcohol-green). Structure of hemicellulose. Schematic representation of the synergistic action of cellobiohydrolases (CBHI, CBHII), endoglucanases (EG), and β-glucosidases (βG). Overview of the two strategies (free or cell-bound cellulase systems) for degrading cellulose. In free extracellular systems, endoglucanases and exoglucanases act synergistically, with the endoglucanase cutting amorphous cellulose providing chain ends for exoglucanases to release cellobiose. Then, β-glucosidaes complete the process of cellulose hydrolysis by releasing glucose. Also, cellodextrins released by endoglucanases can be further hydrolyzed by cellodextrinases. The carbohydrate binding domain directs the enzymes to their specific substrates. In the cellulosome system, all cellulases are anchored to a common scaffold but are generally thought to follow the same synergic mode of action. The scaffolding is bound to the cell membrane through the surface layer homology domain, while a network of dockerin and cohesin domains amplifies the number of cellulases bound to the same scaffolding unit. Lastly, a carbohydrate binding domain is responsible for the targeting of the whole complex to the substrate. Various strategies adopted for improving bioprocess of cellulase production. Renewable ethanol production by end use (https://www.clariant. com). Biofuels generations. Different generations of biofuels with their characteristics. Different feedstocks used in the first and second generation biorefinery for producing biofuels, biochemicals, food, and feed. First generation biofuels. Types of raw feedstock for second-generation biofuel. Biochemical conversions of second-generation feedstocks to biofuels. Biofuel generation from microalgae. Fourth generation biofuel production.

6 7 9 10

10 14

15

18 27 54 55 55 56 57 59 62 63

ix

x

List of figures

Figure 3.9 Figure 4.1 Figure 4.2 Figure 5.1

Figure 5.2 Figure 7.1 Figure 7.2

Figure 7.3

Figure 7.4

Figure 8.1 Figure 8.2 Figure 8.3 Figure 8.4 Figure 8.5 Figure 8.6 Figure 8.7

Figure 8.8 Figure 8.9 Figure 8.10 Figure 8.11 Figure 8.12 Figure 8.13

A schematic diagram of bioethanol production based on different generations. Chart of various steps involved in a biomass supply chain. Schematic representation of biological conversion of lignocellulosic components into various chemicals and biofuels. Diagram of different schemes for sequential SSF and SmF. The color of the arrows indicates the system to which they belong (green for A, red for B, and blue for C). Different production strategies for commercial production of cellulases (A, Off-site; B, On-site; C, Integrated; D, Consolidated). Enzymes involved in cellulose degradation. Schematic representation of the hydrolysis of cellulose by noncomplexed (A) and complexed (B) cellulase systems. a, cellulose; b, glucose; c, cellobiose; d, oligosaccharides; e, endoglucanase with carbohydrate-binding module (CBM); f, exoglucanase (acting on reducing ends) with CBM; g, exoglucanase (acting on nonreducing ends) with CBM; h, β-glucosidase; i, cellobiose/cellodextrin phosphorylase; j, S-layer homology module; k, CBM; l, type-I dockerin cohesion pair; m, type-II dockerin cohesin pair. The figure is not drawn to scale. Crystal structures of family 6 endoglucanase and exoglucanase. (A) The structure of endoglucanase Cel6A of Thermobifida fusca (PDB code: 1TML), which exhibits a deep cleft at the active site. (B) The structure of exoglucanase Cel6A of Humicola insolens (PDB code: 1BVW), in which the active site of it bears an extended loop that forms a tunnel. The simulation of the synergy between endoglucanase and exoglucanases in terms of substrate characteristics (degree of polymerization, A; and accessibility, B) and experimental conditions (enzyme loading, C; and reaction time, D). Schematic diagram of the synergic action of cellulases on cellulosic biomass hydrolysis. Role of cellulases during the complete biofuel production process. Schematic diagram for conversion of lignocellulosic biomass into ethanol. Processing route for bioethanol production. Schematic of pretreatment effect on lignocellulosic biomass. Separation process of bioethanol by extractive distillation. Clariant completes construction of first commercial Sunliquid cellulosic ethanol plant in Podari, Romania (https://www.clariant. com). Clariant’s flagship commercial Sunliquid cellulosic ethanol plant in Romania. (https://www.clariant.com). Sunliquid plant Podari (https://www.clariant.com). The Sunliquid process for the production of cellulosic ethanol from agricultural residues (https://www.clariant.com). Sunliquid—a fully integrated process design (https://www.clariant. com). Reproduced with permission. Sunliquid technology platform for the production of sustainable biobased products. Sun liquid technology: five licenses sold globally (https://www. clariant.com). Reproduced with permission.

64 70 79 101

102 120 123

125

131

140 141 141 145 146 157 161

161 162 163 163 165 165

List of figures

Figure 8.14 Figure 8.15 Figure 8.16 Figure 9.1 Figure 10.1

Figure 11.1 Figure 11.2 Figure 11.3 Figure 12.1

Reduction of Carbon dioxide emissions (https://www.clariant.com). Sunliquid: fully integrated process saves ~95% GHG emissions (https://www.clariant.com). Clariant Bioethanol Pilot Plant, Straubing, Germany (https://www. clariant.com). Cellulosome structure and assembly. Schematic diagram of the actions of cellulases and synergistic proteins in hydrolysis of cellulose. The C1-Cx lignocellulose degradation model for cellulose degradation. First, C1 factors, such as expansin, and lytic polysaccharide monooxygenases, which efficiently catalyze oxidative cleavage of glycosidic bonds in the recalcitrant polysaccharides of crystalline cellulose using molecular oxygen and the external electron donor, such as cellobiose dehydrogenase, ascorbic acid or gallic acid, creating nicking, swollen and disintegrated cellulosic structure, forming new initiation sites for conventional cellulases, namely, Cx factors. Second, the amorphous cellulose was hydrolyzed into monosaccharide by cellulase system. The endoglucanase acts on the amorphous (internal) region of the fibrils by cleavage of the β-glucosidic linkage, then the cellobiohydrolase releases cellobiose from the end of the polysaccharide chain, finally, β-glucosidase completes the degradation process by hydrolyzing cellobiose and other cellodextrins to glucose units. BGL, β-glucosidase; CBH I, cellobiohydrolase 1; CBH II, cellobiohydrolase 2; CDH, cellobiose dehydrogenase; C1 factor, cellulose hing domains (CBMs), plant expansins, bacterial expansins, and lytic polysaccharide monoxygenases; Cx factor, endoglucanase, cellobiohydrolase, and β-glucosidase; EG, endoglucanase; LPMO, lytic polysaccharide monooxygenase. Advantages of enzymatic hydrolysis carried out at elevated temperature. Strategies to increase cellulase efficiency and characteristics. Genetic manipulation strategies in fungi for improved cellulase production. Biomass-to-biofuels supply chain.

xi

166 166 167 188 207

215 226 227 244

Elsevier Radarweg 29, PO Box 211, 1000 AE Amsterdam, Netherlands The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States Copyright © 2023 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-323-99496-5 For Information on all Elsevier publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Susan Dennis Editorial Project Manager: Maria Elaine Desamero Production Project Manager: Bharatwaj Varatharajan Cover Designer: Victoria Pearson Typeset by MPS Limited, Chennai, India

Contents List of figures List of tables Preface Acknowledgments

1.

2.

3.

Background

ix xiii xv xvii 1

1.1 Introduction 1.2 Bioenergy and biofuels 1.3 Biomass 1.4 Cellulase enzyme References

1 2 5 11 19

Worldwide scenario of biofuel production

25

2.1 Introduction 2.2 Status of biofuel production 2.2.1 United States 2.2.2 Brazil 2.2.3 European Union 2.2.4 Argentina 2.2.5 China 2.2.6 Australia 2.2.7 Canada 2.2.8 Thailand 2.2.9 Japan 2.2.10 India References

25 28 28 33 37 39 39 41 43 45 46 47 50

Generations of biofuels

53

3.1 Introduction 3.2 First generation biofuels 3.3 Second generation biofuels 3.4 Third generation biofuels 3.5 Fourth generation biofuels References

53 56 58 61 62 64

v

vi

4.

5.

6.

7.

Contents

Challenges to biofuel production

67

4.1 Introduction 4.2 Challenges in lignocellulosic biomass conversion to biofuels/biochemicals 4.2.1 Feedstock production and logistics 4.2.2 Lignocellulosic biomass pretreatment 4.2.3 Enzymatic hydrolysis 4.2.4 Microbial fermentation and biomass 4.2.5 Biofuel cost 4.2.6 Water recycling 4.2.7 Generation of coproducts 4.2.8 Energy and environmental issues References Further reading

67 68 68 71 74 77 78 79 80 80 82 89

Current production status of cellulases and challenges

91 91 93 93 96

5.1 Introduction 5.2 Production of cellulases 5.2.1 Submerged fermentations 5.2.2 Solid-state fermentation 5.2.3 Sequential solid-state fermentation and submerged fermentation 5.2.4 Use of mixed cultures References Further reading

99 102 103 108

Cellulase market scenario

109

6.1 Introduction 6.2 Market scenario References

109 110 115

Roles of cellulases in cellulose hydrolysis

119

7.1 Introduction 7.2 Cellulase enzyme systems for cellulose hydrolysis 7.2.1 Endoglucanase 7.2.2 Exoglucanase 7.2.3 β-Glucosidase 7.2.4 Other cellulases and accessory proteins 7.3 Synergy among cellulose degrading system References Further reading

119 121 123 124 126 127 128 132 138

Contents

8.

9.

vii

Cellulases for biofuels production

139

8.1 Introduction 8.2 Bioethanol production 8.2.1 Pretreatment 8.2.2 Hydrolysis 8.2.3 Fermentation 8.2.4 Distillation 8.3 Factors affecting bioethanol production 8.4 Important developments in the production of cellulosic ethanol 8.4.1 Case study 8.5 Other biofuels made by assistance from enzymes References Further reading

139 140 144 147 149 155 156

Advanced developments in production processes of cellulases 9.1 Introduction 9.2 Advanced development in production processes of cellulases References Further reading

10. Cellulases and auxiliary enzymes 10.1 Introduction 10.2 Cellulases 10.2.1 Cellobiohydrolase 10.2.2 endo-1,4-β-Glucanase 10.2.3 β-Glucosidase 10.2.4 Cellulase auxiliary enzymes References Further reading

11. Approaches to enhance cellulase production to improve biomass hydrolysis 11.1 Introduction 11.2 Thermostable cellulases 11.3 Isolation and screening of efficient thermostable cellulase producing fungi 11.4 Enhancement of thermal and pH stability of cellulases in the presence of nanomaterials 11.5 Recombinant DNA technology for increasing cellulase activity and efficacy

157 159 167 168 176

179 179 180 190 194 197 197 197 198 199 200 202 208 212

213 213 214 219 220 222

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Contents

11.6 Use of suitable carbon source and necessity of pretreatment of lignocellulosic biomass 11.7 Optimization of medium and process parameters using statistical methods for improved production of cellulases 11.8 Improvements in production of cellulases via microbial fermentation processes 11.9 Microbial co-production of other important enzymes for the overall economy of the process References

12. Future prospects 12.1 Feedstock production 12.2 Feedstock logistics 12.3 Biofuels production 12.4 Biofuels distribution 12.5 Biofuels end use References Further reading Index

228 229 230 231 231 241 244 244 245 245 245 246 246 247

Acknowledgments I am grateful for the help received from many people and companies/ organizations for providing information. I am also thankful to various publishers for allowing me to use their material. My deepest appreciation is extended to Elsevier, Springer, RSC, ASM Publications, John Wiley & Sons, Hindawi, MDPI, IntechOpen, Enerdata, SpringerOpen, and other open-access journals and publications. My special thanks to the US Energy Information Administration (EIA), Enerdata, and Geoff Cooper, President & CEO, Renewable Fuels Association for granting permission to use their material. I would also like to offer my sincere thanks to Caroline Schmid, Global Marketing Manager, Clariant Produkte (Deutschland) GmbH for providing information on the Sunliquid process.

xvii

Chapter 1

Background 1.1

Introduction

The limited nature and speedy exhaustion of fossil fuels because of increasing global energy requirements is harmfully impacting the environment (Olguin-Maciel et al., 2020; Ali et al., 2019; Taghizadeh-Alisaraei et al., 2019; Nargotra et al., 2019; Patel et al., 2019; Popp et al., 2014). Replacing fossil fuels with biofuels has the potential to reduce some unwanted aspects of fossil fuel production and use, including conventional and greenhouse gas (GHG) pollutant emissions, exhaustible reduction of resources, and dependency upon unstable foreign suppliers. Therefore in theory, the production and use of biofuels could be sustained for an indefinite period (http://www. epa.gov). Biofuel production has substantially increased in the last twenty years with the aim of environmental protection and ensuring energy independence. Due to the increasing prices of fossil fuels, the production of biofuels has reached remarkable volumes over the last two decades. Worldwide biofuel production has increased ninefold between 2000 and 2020, in-spite of rigorous reduction in 2020, and should bounce back by 13% in 2021 over an expected recovery in oil demand and strengthened or maintained biofuel support policies. Bioethanol accounts for two third of the worldwide biofuel production and biodiesel for 32%. First-generation biofuels from sugarcane and corn based bioethanol and vegetable oil based biodiesel cover most of the worldwide production. Bioethanol production and consumption is dominated by North America—particularly the United States—and Latin America— particularly Brazil whereas biodiesel production and consumption is dominated by Asia and Europe. Worldwide biofuel consumption is expected to decline substantially by 2029, because of the likely reduction in fuel demand, bigger competition between transport technologies and the decarbonization development of public policies. First-generation biofuels will continue to take over global production by 2030, with advanced biofuels limited to 10% (expected share of 31% in the European Union thanks to the Renewable Energy Directive II directive) (https://www.enerdata.net/publications/executive-briefing/biofuels-market-dynamics.html). First-generation biofuels obtained from food raw materials such as sugarcane, cereal grains, root crops, and vegetable oils are becoming more and more competitive with food production. Over the last few years, there has Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00002-9 © 2023 Elsevier Inc. All rights reserved.

1

2

Cellulases in the Biofuel Industry

been a period of the active growth in the production of liquid biofuels. In 2018, the worldwide production of bioethanol and biodiesel together reached 167.9 billion liters. About 16.1% of maize grain, 1.7% of wheat grain, 3.3% of grain of other feed grains and 13.5% of vegetable oil were consumed (Kurowska et al., 2020). Biomass is a versatile renewable energy source. It can be directly converted into biofuels. Biomass offers several many benefits over fossil fuels. Biomass is carbon neutral and is always and extensively available as a renewable source of energy. It reduces the overdependence on fossil fuels and is less costly than fossil fuels. Biomass production generates income for the manufacturers, also generates lesser garbage in landfills and increases energy independence (Ali et al., 2019; Taghizadeh-Alisaraei et al., 2019; Brinkman et al., 2019; Muthuvelu et al., 2019). “Biomass is particularly important in providing raw materials for the production of renewable energy sources. The term biomass refers to all organic matter in the biosphere, of both plant and animal origin, and to materials obtained by its natural or artificial conversion (Muresan and Attia, 2017; Mehedintu et al., 2018; Contescu et al., 2018). By submitting biomass to biochemical, thermochemical and biological conversion processes, liquid and gas fuels are obtained (bioethanol, biodiesel, biogas). Up to now, biofuels have been produced mainly through alcohol fermentation of starch products (ethanol), municipal waste, sewage sludge and others (biogas), dry distillation of wood (methanol) and transesterification of higher fatty acids (biodiesel). Such fuels are counted as the first-generation ones and it is predicted that they will dominate for many years to come because they can be burnt in existing unmodified engines and their production is easy and economically viable. Currently, there are attempts to implement other renewable raw products in biofuel production, like cellulose, which are much more difficult to process, and to design more complex biotechnological methods” (Kurowska et al., 2020; Nigam and Singh, 2011; Hill et al., 2006). Some of the most common (and/or most promising) biomass feedstocks are listed in Table 1.1.

1.2

Bioenergy and biofuels

Bioenergy is energy obtained from biofuels. Biofuels are produced directly or indirectly from organic material—biomass—including plant materials and animal waste. The term biofuel refers to any liquid, gas, or solid fuel mostly produced from a renewable biomass resources. The Food and Agriculture Organization (FAO) reports that bioenergy covers about 10% of total world energy supply. Traditional unprocessed biomass such as fuel wood, charcoal and animal dung accounts for most of this and represents the major energy source for

Background Chapter | 1

3

TABLE 1.1 Most common biomass feedstocks. Grains and starch crops Sugarcane, corn, wheat, sugar beets, industrial sweet potatoes, etc. Agricultural residues Corn stover, wheat straw, rice straw, orchard prunings, etc. Forestry materials Logging residues, forest thinnings, etc. Energy crops Switchgrass, miscanthus, hybrid poplar, willow, algae, etc. Food waste Waste produce, food processing waste, etc. Urban and suburban wastes Municipal solid wastes (MSW), lawn wastes, wastewater treatment sludge, urban wood wastes, disaster debris, trap grease, yellow grease, waste cooking oil, etc. Animal byproducts Tallow, fish oil, manure, etc. Source: Based on https://www.eesi.org/topics/bioenergy-biofuels-biomass/description.

most of the people in developing countries who are using it mostly for heating and cooking. Highly developed and efficient conversion technologies are allowing the extraction of biofuels from wood, crops and waste material. Biofuels can be liquid, gaseous or solid, although the term is mostly used in a strict sense to refer only to liquid biofuels for transport. Biofuels may be obtained from agricultural and forestry residue, fast growing tree plantations and annual crops, fishery products or municipal wastes, and also from food industry and food service by-products and wastes (http://www.greenfacts.org). A differentiation is made between primary and secondary biofuels. Primary biofuels are utilized in an unprocessed form, mainly for cooking, heating or electricity production. These include fuel wood, wood chips and pellets and organic materials. Secondary biofuels result from processing of biomass. These include liquid biofuels such as biodiesel and ethanol and can be used in vehicles and industrial processes. Bioenergy is mostly used in homes (80%). In industry, it is used to a lesser extent (18%), whereas liquid biofuels used for transport are still playing a limited role (2%).

4

Cellulases in the Biofuel Industry

Although the production of liquid biofuels for transport has grown quickly over the last few years globally, it presently represents only 1% of total transport fuel consumption and only 0.2% 0.3% of total energy consumption. Examples of biofuel are bioethanol, biomethanol, biosynthetic gas (biosyngas), biodiesel, biogas, biochar, bio-oil, biohydrogen, and Fischer Tropsch produced liquids (Ali et al., 2019; Carrillo-Nieves et al., 2019). Biofuels serve as a bridge between the agricultural and energy markets as agricultural commodities are the important raw materials in biofuel production (Debnath and Giner, 2019). Main advantages and paybacks obtainable from the use of biofuels as a form of renewable fuel are presented in Table 1.2. Liquid biofuels are of especial interest because of the huge infrastructure already in place to use them, particularly for transportation. The liquid biofuel—ethanol is in greatest production. It is produced by fermentation of starch or sugar. United States and Brazil are among the leading producers of ethanol. In the United States, bioethanol is mostly produced from corn and it is normally blended with gasoline for producing “gasohol.” This contains 10% ethanol. In Brazil, ethanol is produced mostly from sugarcane. It is usually used as a 100% ethanol fuel or in gasoline blends containing 85%

TABLE 1.2 Benefits of biofuels. Biofuels are renewable and are carbon- and CO2/GHG-neutral during the progression of the life cycle Less GHG emissions are generated from the utilization of biofuels compared to FB fuels Biofuels are biodegradable, sustainable, and environmentally benign Biofuels are largely produced from locally available and accessible resources, applying safe production methods Production and utilization of biofuels enhance home-grown agricultural development and investment Biofuels provide improvements in the health and living conditions of people Biofuels create jobs and improvements in local livelihoods and reduce energy importation Economically, biofuel helps to stabilize energy prices, conserve foreign exchange, and generate employment at the macroeconomic level Household usage of biofuel does not trigger life-threatening health conditions, as opposed to FB fuels Source: Based on Awogbemi et al. (2021); Janampelli and Darbha (2019); Wu et al. (2020); Appavu et al. (2021); Navas et al. (2020); Darby and Callahan (2020); Smith (2019); Yaghoubi et al. (2019); Szabo´ (2019); Chintala (2019); Oyewole et al. (2019); Topcu and Tugcu (2020); Schuenemann and Kerr (2019); Mattioda et al. (2020); Siddiqui et al. (2019).

Background Chapter | 1

5

TABLE 1.3 Greenhouse gas emissions from various biofuels compared to gasoline. Fuel

Gasoline

Bioethanol 1G

Bioethanol 1G

Bioethanol 2G

Feedstock

Oil

Crop/Corn

Cane

Agricultural residues

Emissions

100%

48%

22%

14

Source: Based on https://sunliquid-project-fp7.eu/wp-content/uploads/2014/09/ factsheet_sunliquid_en.pdf.

ethanol. Unlike the “first-generation” ethanol biofuel produced from food crops, “second-generation” cellulosic ethanol is obtained from low-value biomass which includes wood chips, crop residues, and municipal waste (Hassan et al., 2018). It has a higher cellulose content. Cellulosic ethanol is mostly produced from sugarcane bagasse, or from a variety of grasses that can be grown on low-quality land. Bagasse is a waste product from sugar processing. Cellulosic ethanol is mainly used as a gasoline additive; the conversion rate is lower as compared to first-generation biofuels. Cellulosic ethanol from agricultural feedstocks and energy crops is usually considered to be environmentally sustainable because it provides higher reduction of GHG emissions and zero or low indirect emissions from land use change in comparison to traditional ethanol production from food and feed crops. Table 1.3 presents GHG emissions from various biofuels compared with gasoline.

1.3

Biomass

Biomass generally includes: plant-based woody biomass which are mostly lignocelluloses, plant-based non-woody biomass mostly starch, sugar and oils and animal/human based biomass which includes animal fats and proteins, slurry/slaughter wastes, house hold wastes, etc. Amongst the plantbased woody biomass, lignocellulosic biomass is considered as a potential resource for renewable energy. It is mostly used for land filling or simply burned off. Lignocellulosic biomass constitutes 60% of the plant cell wall. About 100 billion tons of plant dry material is produced in the world by photosynthetic activity yearly. The leftover of lignocellulosic biomass is mainly treated as waste therefore, extensive research has been conducted for the efficient exploitation of the lignocellulosic biomass for producing enzymes, biofuels, feeds, antioxidants etc. The major components of the lignocellulosic materials are cellulose, hemicellulose, and lignin. Fig. 1.1 shows the structure of lignocellulosic biomass. Fig. 1.2 shows the products obtained from lignocellulosic biomass.

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Cellulases in the Biofuel Industry

FIGURE 1.1 Structure of lignocellulosic biomass and its biopolymers; cellulose, hemicellulose, ´ ´ J.U., Hernandez-De ´ and lignin (Hern´andez-Beltr´an et al., 2019). Hernandez-Beltr an, Lira, I.O, ´ ´ F, Balagurusamy, N., 2019. Insight into Cruz-Santos, M.M, Saucedo-Luevanos, A, Hernandez-Ter an, pretreatment methods of lignocellulosic biomass to increase biogas yield: current state, challenges, and opportunities. Appl. Sci. Basel 9(18), 3721. https://doi.org/10.3390/app9183721. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

The composition of cellulose, hemicellulose, and lignin is found to vary from one plant species to another (Yusuf and Inambao, 2019; Bajpai, 2016, 2021; Shahzadi et al., 2014; Walker, 2010; Sher et al., 2021; Potters et al., 2010). For instance, hardwoods possess larger amounts of cellulose, while in the wheat straw and leaves, the higher amount of hemicelluloses are present. The ratios between different constituents within a single plant vary with the stage of growth, age, and other conditions. Lignocellulosic raw materials need destructive pretreatment to yield a substrate which is readily hydrolyzed by commercial cellulolytic enzymes, or by microorganisms producing enzyme, for releasing sugars for fermentation. In lignocelluloses, cellulose fiber strands are formed by cellulose linking to each other through hydrogen bonds. The cellulose structure inside the polymer is not homogenous. Crystalline regions are where cellulose

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FIGURE 1.2 Lignocellulosic biomass along with their products (Haq et al., 2021). Haq, I.U., Qaisar, K., Nawaz, A., Akram, F., Mukhtar, H., Zohu, X., et al., 2021. Advances in valorization of lignocellulosic biomass towards energy generation. Catalysts, 11, 309. https://doi.org/ 10.3390/catal11030309. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

nanofibrils are organized in order and compact, whereas amorphous regions are disordered and are easily hydrolyzed (Tran et al., 2019; Perez et al., 2002). Cellulose fibers are surrounded by hemicellulose and lignin. This structure naturally protects the polysaccharides from hydrolysis by enzymes and chemicals, thus raising a difficulty in both chemical and bioconversion of lignocellulose to other products, that is “ethanol” (Bajpai, 2021; Tran et al., 2019). Cellulose, hemicellulose, and lignin are tightly packed with each other, protecting them against attack of microorganisms and therefore make their degradation not easy. Cellulose and hemicelluloses can be converted into fermentable sugars which can be further converted into bioethanol and other value-added products (Singh et al., 2017). There are numerous ways for deconstruction of biomass such as thermochemical, chemical, or biological but the biological route by using enzyme is the most well-liked, as it is environment-friendly and sustainable. “Cellulose is considered the most copious and bountiful renewable sources for producing valuable products for energy sources, this property of cellulose makes it more important to use it for the production of value-added fuels by thermochemical or biochemical processes. This has been used for the production of biofuel especially for methane and bioethanol. Beside this, the cellulose has also wide range of application and use in different

8

Cellulases in the Biofuel Industry

industries such as food and beverage, animal feeds, detergent, agriculture, textile, pulp and paper industry. It has attracted the interest of industrial scientists, so these can be utilized for the production of several enzymes at both lab and industrial scale. It has also attracted the interest of scientists as they are taking advantage of using this low cost energy source (cellulose) for the production of biological products, that would add into the economy and energy security of the country. Cellulose usually occurs as fibers, densely packed with hydrogen bond and insoluble in water so it is very resistant to hydrolysis without the chemicals and mechanical degradation. Hence cellulose or other polysaccharide compounds can be converted into simple sugar or glucose units by the activity of cellulase enzymes. It is also a very stable part of plant cell wall matrix and biomass, that’s why plants take great benefit of it as it plays an important role to maintain the cell wall stability and integrity. The structure of cellulose determines the hydrolysis pathway and compactness in cellulosic structure makes it prone to the degradation. This compact and crystalline structure of cellulose is due to inter and intramolecular hydrogen bonding. This super crystalline structure of cellulose makes it resistant to hydrolysis even at very extreme reaction conditions, therefore, it must be pre-treated to convert the cellulosic biomass or plant cellulosic parts into simple sugars” (Sher et al., 2021). “Cellulose is a linear polysaccharide. In this polymer, D-glucose subunits are attached together by formation of β-1,4glycosidic linkages between individual glucose molecules. The molecular formula of cellulose is (C6H12O6)n. The n indicates the degree of polymerization (DP). It symbolizes the number of glucose subunits connected with each other. This number is varying from hundreds to thousands. Two glucose repeating units together are called cellobiose. In other words, this polymer is made by β-(1 - 4)-D-glucopyranose units in 4C1 conformation. It consists of long chains of anhydro-Dglucopyranose units (AGU) with each cellulose molecule having three hydroxyl groups per AGU with the exception of the terminal ends. Cellulose has both crystalline and amorphous regions in its structure in various proportions. Those regions are intertwined to form the structure of cellulose. There are four major crystalline forms, for instance, Iα, Iβ, II, and III. This crystalline structure is a result of intramolecular and intermolecular hydrogen bonding between glucose monomers in cellulose. These hydrogen bonds construct a huge network that directly contributes to the compact crystal structure of cellulose polymer. On the other hand, this strong intramolecular and intermolecular hydrogen bond formation leads to poor solubility of cellulose” (Tran et al., 2019; Ciolacu et al., 2011). Fig. 1.3 Shows the cellulose polymer chain structure. Lignin is the major structural component of lignocelluloses. It contains three different types of phenolic monomers: p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol (Fig. 1.4). It provides strength to the lignocellulosic biomass and hampers the action of hydrolytic enzymes by acting as a barricade. It is one of the most recalcitrant constituent of the lignocellulosic biomass because of its structural complexity (Arevalo-Gallegos et al., 2017; Gupta and Verma, 2015; Yao et al., 2015).

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FIGURE 1.3 The cellulose polymer chain structure (Suttie et al., 2017). Reproduced with permission from Suttie, E., Hill, C., Sandin, G., Kutnar, A., Ganne-Che´deville, C., Lowres, F., et al., 2017. Wood as bio-based building material. Performance of Bio-based Building Materials. ˇ & Leitgeb, M. (2021). ´ K., Knez, Z., Woodhead Publishing, pp. 21- 96, Elsevier and Vasic, Bioethanol production by enzymatic hydrolysis from different lignocellulosic sources. Molecules, 26 (3), 753. Distributed under the terms of the Creative Commons Attribution 4.0 International License.

Hemicellulose is the significant component of plant cell wall. It is the second most abundant polymer. It comprises of short linear and branched polymers (Fig. 1.5). Hemicellulose has an amorphous structure contrary to cellulose. It consists of several heteropolymers such as xylan, galactomannan,

10

Cellulases in the Biofuel Industry

FIGURE 1.4 Schematic representation for the macromolecular structure of lignin (Major monolignol units are colored as sinapyl alcohol-red, guaiacyl alcohol-blue, p-coumaryl alcoholgreen) (Karunarathna and Smith, 2020). Karunarathna, M.H., Smith, R.C., 2020. Valorization of lignin as a sustainable component of structural materials and composites: advances from 2011 to 2019. Sustainability, 12, 734. MDPI. https://DOI:10.3390/su12020734. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

–OOC H3CO

O

HO

H3C

OH O O

OH

O OH O

O HO O

O

O O

H3CO

O

O

HO

O

O O

OH

HO

FIGURE 1.5 Structure of hemicellulose (Machmudah et al., 2017). Reproduced with permission from Machmudah, S., Wahyudiono, Kanda, H., Goto, M., 2017. Hydrolysis of biopolymers in near-critical and subcritical water. In: Herminia, D., Maria Jesus, G.M. (Eds.), Water Extraction of Bioactive Compounds. Elsevier, Amsterdem, Netherlands, pp. 69 107.

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glucuronoxylan, arabinoxylan, glucomannan, and xyloglucan. Hardwood (e.g., dicot angiosperms) hemicelluloses mainly contain xylans, while glucomannans are main constituents in softwood (e.g., gymnosperms). As compared to cellulose, hemicellulose gets rapidly hydrolyzed because of its amorphous and branched nature. The production of bioenergy and bio-based materials from less expensive renewable lignocellulosic materials would bring advantages to the local economy, environment, and national energy security (Zhang, 2008).

1.4

Cellulase enzyme

Cellulase enzymes have been commercially available for more than 40 years and both basic and applied studies on cellulolytic enzymes have shown their biotechnological potential in several industries (Singh, 1999; Singh et al., 2007). Cellulase enzymes play an important role in the enzymatic hydrolysis of cellulosic polymers. Cellulase enzymes break down the cellulose of plant cell walls into simple sugars which can be fermented by microorganisms to fuels, mainly ethanol, and also chemicals, plastics, fibers, detergents, pharmaceuticals, and many other products. Lignin is a main obstruction on the way of efficient hydrolysis of biomass (Saini et al., 2016). It can be removed to some extent by alkaline pretreatment of biomass making cellulose and hemicellulose accessible to enzymes. Cellulose which constitute a main portion of the biomass, contains glucose linked with β, 1 4 linkage. Cellulase enzymes are able to hydrolyze cellulose into glucose, but hardly ever cellulose is present in pure form in nature, and is generally associated with hemicelluloses and lignin. Even hemicelluloses present a physical barrier for cellulases to access cellulose along with lignin (Volynets and Dahman, 2011). Therefore, cellulases along with xylanases are found to be more efficient for degradation of biomass. In actual fact, xylanases hydrolyze the hemicellulosic portion which makes the cellulose more accessible to cellulase. Therefore, xylanases when combined with cellulases show synergistic effect and release more fermentable sugars from the biomass (Hu et al., 2011). Table 1.4 shows classification of cellulase enzymes. “In the production of biofuels, cellulases are fundamental enzymes responsible for hydrolyzing cellulosic biomass into fermentable sugars. As cellulose is a crystalline unbranched polymer, several cellulases are needed to degrade it efficiently. Cellulases hydrolyze the β-1,4-D-glucan bonds, releasing cello-oligosaccharides, cellobiose, or glucose. The complete degradation of cellulose is carried out by an enzymatic complex, which includes endo-β-1,4-glucanases (EG EC 3.2.1.4), cellobiohydrolases (CBH EC

TABLE 1.4 Classification of cellulase enzymes (Roohi et al., 2019). Name of enzyme

EC number

Substrate

Site of cleavage

Product

Endo-1,4-β-d-glucan glucanohydrolase

EC 3.2.1.4

Cut at random at internal amorphous sites of cellulose, lichenin, and cereal β-d-glucans

Endo-1,4-β-dglucosidic linkages

Oligosaccharides of various lengths

Exoglucanase or 1,4-β-d-glucan cellobiohydrolases (cellobiohydrolases)

EC 3.2.1.91

Nonreducing ends of cellulose and cellotetraose

Exo-1,4-β-dglucosidic linkages

Cellobiose

Exoglucanases or 1,4β-d-oligoglucancellobiohydrolases

EC 3.2.1.74

Cellooligosaccharide, p-nitrophenyl-β-dcellobioside

Exo-1,4-β-dglucosidic linkages

Cellobiose

β-Glucosidases or β-d-glucoside glucohydrolases

EC 3.2.1.21

Terminal nonreducing ends of cellulose

Exo-1,4-β-dglucosidic linkages

β-d-Glucose

Cellobiose: orthophosphate α-d-glucosyl transferase

EC 2.4.1.49

Cellobiose

Phosphorolytic cleavage

β-d-Glucose

1,4-β-d-oligoglucan: orthophosphate α-dglucosyl transferase

EC 2.4.1.20

Cellodextrins ranging from cellotriose to cellohexoses

Phosphorolytic cleavage

β-d-Glucose

Cellobiose 2-epimerase

EC 5.1.3.11

Cellobiose

Exo-1,4-β-dglucosidic linkages 4-O-β-dGlucosylmannose

4-O-β-dGlucosylmannose

Source: Reproduced with permission from Roohi, B.R.K, Parveen, S., Khan, F., Zaheer, M.R., Kuddus, M., 2019. Chapter 8 Advancements in bioprocess technology for cellulase production. In: Srivastava, N., Srivastava, N., Mishra, P.K., Ramteke, P.W., Singh, R.L. (Eds.), New and Future Developments in Microbial Biotechnology and Bioengineering: From Cellulose to Cellulase: Strategies to Improve Biofuel Production. Elsevier.

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3.2.1.91 and EC 3.2.1.176), and β-glucosidases (BGL EC 3.2.1.21). Endoglucanases hydrolyze the glucose chain internally, releasing cellooligosaccharides and exposing additional free reducing and nonreducing ends. CBHs hydrolyze the exposed ends and release cellobiose, a strong CBH inhibitor, that will later be converted into glucose monomers. On the other hand, CBHs partially amorphize the crystalline regions of the cellulose matrix, which leads to easier substrate access for EGs. Finally, β-glucosidases cleave cellobiose generating glucose monomers. Hence, the degradation of cellulose is influenced by the synergistic action of the described enzymes” (Contreras et al., 2020; Kumar et al., 2008; Wang et al., 2012; Eriksson et al., 2002; Perez et al., 2002). Figs. 1.6 and 1.7 show schematic representation of the synergistic action of cellobiohydrolases (CBHI, CBHII), endoglucanases (EG), and β-glucosidases (βG) (Contreras et al., 2020). Cellulases are inducible enzymes, which are produced by broad variety of microorganisms including fungi, bacteria and actinomycetes (Tables 1.5 to 1.7). These microorganisms can be aerobic, anaerobic, mesophilic or thermophilic. Most widely studied cellulase producers are genera of Clostridium, Cellulomonas, Thermomonospora, Trichoderma, and Aspergillus (Henrissat et al., 1998; Kubicek, 1993; Sang-Mok and Koo, 2001; Sun and Cheng, 2002; Sukumaran et al., 2005; Kuhad et al., 2010, 2011). Among the microorganisms, fungi are the important producers of cellulases and account for about 80% of the cellulose hydrolysis on the Earth (Selig et al., 2015). Mostly, ascomycota, basidiomycota and deuteromycota members of the fungi contain efficient cellulolytic activities. Cellulase enzymes obtained from aerobic fungal microorganism are preferred commonly for industrial applications as they are extracellular and secreted in bulk during growth (Payne et al., 2015). Trichoderma reesi is the most extensively studied fungus. It has the ability to convert native cellulose to glucose (Gusakov, 2011). Additionally, Aspergillus, Humicola, Penicillum and Sclerotium fungi are considered potential candidates for industrial production of cellulolytic enzymes (Sreedharan et al., 2016; Gusakov and Sinitsyn, 2012). Actinomycetes genera for example Cellulomonas, Streptomyces and Thermomonospora also produce cellulolytic enzymes (Singh et al., 2016). “Structurally fungal cellulases are simpler as compared to bacterial cellulase systems, cellulosomes. Fungal cellulases typically have two separate domains: a catalytic domain (CD) and a cellulose binding module (CBM), which is joined by a short polylinker region to the catalytic domain at the N-terminal. The CBM is comprised of approximately 35 amino acids, and the linker region is rich in serine and threonine. The main difference between cellulosomes and free cellulase enzyme is in the component of

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Cellulases in the Biofuel Industry

Amorphous region

Cellobiohydrolase (GH 5, 6, 7, 9, 48)

HO O

OH HO O

O HO

O

OH

O

OH

OH

HO O

HO

O

OH O HO

O

O O OH HO

OH

O

OH O HO

O

O OH

HO OH

OH HO O

O

O HO

OH OH

OH

Cellobiohydrolase (GH 5, 6, 7, 9, 48)

Cellobiose HO

HO O OH

O

O

OH O HO

O

OH OH

HO

HO

HO O OH

OH O HO

Endoglucanase (GH 5-9, 12, 44, 45, 48, 51, 61, 74)

Cellooligosaccharides

O

OH

O HO

HO OH

O

OH

O

HO OH O

O

OH

O

OH O

OH HO O

OH

O

OH

HO

O

HO OH

O HO

O

O

O HO

HO

HO HO O

OH

OH HO O OH

O

O

HO

OH

HO

HO O OH

HO

OH

Crystalline region

O

OH O

HO

OH OH

O

HO

Cellodextrinase (GH 1,3)

HO

HO

HO

O

OH OH

HO

β-Glucosidase (GH 1,3)

Glucose HO O HO

OH OH

HO Cellobiose

Glucose

CBHI

EG

Glucose

βG

βG

CBHI

Cellobiose

FIGURE 1.6 Schematic representation of the synergistic action of cellobiohydrolases (CBHI, CBHII), endoglucanases (EG), and β-glucosidases (βG) (Gomes et al., 2016). Reproduced with permission from Gomes, E., de Souza, A.R., Orjuela, G.L., Da Silva, R., de Oliveira, T.B., Rodrigues, A., 2016. Applications and benefits of thermophilic microorganisms and their enzymes for industrial biotechnology. In: Schmoll, M., Dattenbo¨ck, C. (Eds.), Gene Expression Systems in Fungi: Advancements and Applications. Fungal Biology. Springer, Cham. Available from: https://doi.org/10.1007/978-3-319-27951-0_21 and Contreras, F., Pramanik, S., Rozhkova, A.M., Zorov, I.N., Korotkova, O., Sinitsyn, A.P., et al., 2020. Engineering robust cellulases for tailored lignocellulosic degradation cocktails. Int. J. Mol. Sci. 21(5), 1589 https://doi.org/ 10.3390/ijms21051589. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

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FIGURE 1.7 Overview of the two strategies (free or cell-bound cellulase systems) for degrading cellulose. In free extracellular systems, endoglucanases and exoglucanases act synergistically, with the endoglucanase cutting amorphous cellulose providing chain ends for exoglucanases to release cellobiose. Then, β-glucosidases complete the process of cellulose hydrolysis by releasing glucose. Also, cellodextrins released by endoglucanases can be further hydrolyzed by cellodextrinases. The carbohydrate binding domain directs the enzymes to their specific substrates. In the cellulosome system, all cellulases are anchored to a common scaffold but are generally thought to follow the same synergic mode of action. The scaffolding is bound to the cell membrane through the surface layer homology domain, while a network of dockerin and cohesin domains amplifies the number of cellulases bound to the same scaffolding unit. Lastly, a carbohydrate binding domain is responsible for the targeting of the whole complex to the substrate (Escuder-Rodr´ıguez et al., 2018). Escuder-Rodr´ıguez, J.J., DeCastro, ´ M.E., Rodr´ıguez-Belmonte, E., Becerra, M., Gonzalez-Siso, ´ M.E., Cerdan, M.I., 2018. Cellulases Thermophiles Found. Metagenomics; 6(3):66. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

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Cellulases in the Biofuel Industry

TABLE 1.5 Fungi having cellulolytic abilities. Soft rot fungi Aspergillus niger A. nidulans A. oryzae A. terreus Fusarium solani F. oxysporum Humicola insolens H. grisea Melanocarpus albomyces Penicillium brasilianum P. occitanis P. decumbans Trichoderma reesei T. longibrachiatum T. harzianum Chaetomium cellulyticum C. thermophillum Neurospora crassa P. fumigosum Thermoascus aurantiacus Mucor circinelloides P. janthinellum Paecilomyces inflatus P. echinulatum Trichoderma atroviride Brown rot fungi Coniophora puteana Lanzites trabeum Poria placenta Tyromyces palustris Fomitopsis sp. White rot fungi Phanerochaete chrysosporium Sporotrichum thermophile Trametes versicolor Agaricus arvensis Pleurotus ostreatus Phlebia gigantean Source: Based on Kuhad, R.C., Gupta, R., Khasa, Y.P., 2010. Bioethanol Production from Lignocellulosic Biomass: An Overview. In: Lal, B. (Ed.), Wealth from Waste. Teri Press, New Delhi, India.

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TABLE 1.6 Bacteria having cellulolytic abilities. Aerobic bacteria Acinetobacter junii A. amitratus Acidothermus cellulolyticus Anoxybacillus sp Bacillus subtilis B. pumilus B. amyloliquefaciens B. licheniformis B. circulan B. flexus Bacteriodes sp Cellulomonas biazotea Cellvibrio gilvus Eubacterium cellulosolvens Geobacillus sp Microbispora bispora Paenibacillus curdlanolyticus Pseudomonas cellulosa Salinivibrio sp Rhodothermus marinus Anaerobic bacteria Acetivibrio cellulolyticus Butyrivibrio fibrisolvens Clostridium thermocellum C. cellulolyticum C. acetobutylium C. papyrosolvens Fibrobacter succinogenes Ruminococcus albus Source: Based on Kuhad, R.C., Gupta, R., Khasa, Y.P., 2010. Bioethanol Production from Lignocellulosic Biomass: An Overview. In: Lal, B. (Ed.), Wealth from Waste. Teri Press, New Delhi, India.

cellulosomes-cohesin containing scaffolding and dockerin containing enzyme. The free cellulase contains cellulose binding domains (CBMs), which are replaced by a dockerin in cellulosomal complex, and a single scaffolding-born CBM directs the entire cellulosomes complex to cellulosic biomass” (Kuhad et al., 2011; Bayer et al., 1994, 1998, 2004; Percival Zhang et al., 2006). Fig. 1.8 shows various strategies adopted for improving bioprocess of cellulase production (Singhania et al., 2021).

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Cellulases in the Biofuel Industry

TABLE 1.7 Actinomycetes having cellulolytic abilities. Cellulomonas fimi C. bioazotea C. uda Streptomyces drozdowiczii S. lividans Thermomonospora fusca T. curvata Source: Based on Kuhad, R.C., Gupta, R., Khasa, Y.P., 2010. Bioethanol Production from Lignocellulosic Biomass: An Overview. In: Lal, B. (Ed.), Wealth from Waste. Teri Press, New Delhi, India.

FIGURE 1.8 Various strategies adopted for improving bioprocess of cellulase production. Reproduced with permission from Singhania, R.R., Ruiz, He´ctor A., Awasthi, M.K., Dong, C.-D., Chen, C.-W., Patel, A.K., 2021. Challenges in cellulase bioprocess for biofuel applications, Renew. Sustain. Energy Rev., Elsevier, 151, 111622.

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References Ali, M., Saleem, M., Khan, Z., Watson, I.A., 2019. The use of crop residues for biofuel production. In: Verma, D., Fortunati, E., Jain, S., Zhang, X. (Eds.), Biomass, Biopolymer-Based Materials, and Bioenergy. Woodhead Publishing, Oxford, UK, pp. 369 395. Appavu, P., Ramanan, M.V., Jayaraman, J., Venu, H., 2021. NOx emission reduction techniques in biodiesel-fuelled CI engine: a review. Aust. J. Mech. Eng. 18, 210 220. Arevalo-Gallegos, A., Ahmad, Z., Asgher, M., Parra-Saldivar, R., Iqbal, H.M.N., 2017. Lignocellulose: a sustainable material to produce value-added products with a zero waste approach- a review. Int. J. Biol. Macromol. 99, 30 318. Awogbemi, O., Kallon, D.V.V., Onuh, E.I., Aigbodion, V.S., 2021. An overview of the classification, production and utilization of biofuels for internal combustion engine applications. Energies 14, 5687. Bajpai, P., 2016. Pretreatment of lignocellulosic biomass for biofuel production. Springer Briefs in Molecular Science. Springer Nature, Basingstoke. Bajpai, P., 2021. Lignocellulosic Biomass in Biotechnology. Elsevier, Amsterdam. Bayer, E.A., Belaich, J.P., Shoham, Y., Lamed, R., 2004. The cellulosomes: multienzyme machines for degradation of plant cell wall polysaccharides. Annu. Rev. Microbiol. 58, 521 554. Bayer, E.A., Chanzy, H., Lamed, R., Shoham, Y., 1998. Cellulose, cellulases and cellulosomes. Curr. Opin. Struct. Biol. 8 (5), 548 557. Bayer, E.A., Morag, F., Lamed, R., 1994. The cellulosome—a treasure-trove for biotechnology. Trends Biotechnol. 12 (9), 379 386. Brinkman, M.L., Wicke, B., Faaij, A.P., Van Der Hilst, F., 2019. Projecting socio-economic impacts of bioenergy: current status and limitations of ex-ante quantification methods. Renew. Sustain. Energy Rev. 115, 109352. Carrillo-Nieves, D., Alan´ıs, M.J.R., Quiroz, R.D.L.C., Ruiz, H.A., Iqbal, H.M.N., Parra, R., 2019. Current status and future trends of bioethanol production from agro-industrial wastes in Mexico. Renew. Sustain. Energy Rev. 102, 63 74. Chintala, V., 2019. Coal vs biofuels: a social and economic assessment. Second. Third Gener. Feedstocks 513 529. Elsevier: London, UK. Ciolacu, D., Ciolacu, F., Popa, V.I., 2011. Amorphous cellulose—structure and characterization. Cellulose Chem. Technol.0 45 (1 2), 13 21. Contescu, C.I., Adhikari, S.P., Gallego, N.C., Evans, N.D., Biss, B.E., 2018. Activated carbons derived from high-temperature pyrolysis of lignocellulosic biomass. C. J. Carbon Res. 4, 51. Contreras, F., Pramanik, S., Rozhkova, A.M., Zorov, I.N., Korotkova, O., Sinitsyn, A.P., et al., 2020. Engineering robust cellulases for tailored lignocellulosic degradation cocktails. Int. J. Mol. Sci. 21 (5), 1589. Darby, H.M., Callahan, C.W., 2020. On-farm oil-based biodiesel production. Bioenergy 2020, 157 184. Elsevier: London, UK. Debnath, D., Giner, C., 2019. Interaction between biofuels and agricultural markets. In: Debnath, D., Babu, S. (Eds.), Biofuels, Bioenergy and Food Security. Elsevier, Amsterdam, The Netherlands, pp. 61 76. Eriksson, T., Karlsson, J., Tjerneld, F., 2002. A model explaining declining rate in hydrolysis of lignocelluloses substrates with cellobiohydrolase I (Cel7A) and endoglucanase I (Cel7B) of Trichoderma reesei. Appl. Biochem. Biotechnol. 2002 (101), 41 60. Escuder-Rodr´ıguez, J.J., DeCastro, M.E., Cerd´an, M.E., Rodr´ıguez-Belmonte, E., Becerra, M., Gonz´alez-Siso, M.I., 2018. Cellulases Thermophiles Found. Metagenomics 6 (3), 66.

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Topcu, M., Tugcu, C.T., 2020. The impact of renewable energy consumption on income inequality: evidence from developed countries. Renew. Energy 151, 1134 1140. Tran, T.T.A., Le, T.K.P., Mai, T.P., Nguyen, D.Q., 2019. Bioethanol production from lignocellulosic biomass. In: Yun, Y. (Ed.), Alcohol Fuels—Current Technologies and Future Prospect. Intech Open, London. ˇ Leitgeb, M., 2021. Bioethanol production by enzymatic hydrolysis from difVasi´c, K., Knez, Z., ferent lignocellulosic sources. Molecules 26 (3), 753. Volynets, B., Dahman, Y., 2011. Assessment of pretreatments and enzymatic hydrolysis of wheat straw as a sugar source for bioprocess industry. Int. J. Eng. Environ. 2 (3), 427 446. Walker, G.M., 2010. Bioethanol: Science and Technology of Fuel Alcohol. Ventus Publishing ApS, UK, ISBN 978-87-7681-681-0. Wang, M., Li, Z., Fang, X., Wang, L., Qu, Y., 2012. Cellulolytic enzyme production and enzymatic hydrolysis for second-generation bioethanol production. Biotechnol. China III: Biofuels Bioenergy 1 24. Springer: Berlin/Heidelberg, Germany. Wu, B., Tian, H., Hao, Y., Liu, S., Sun, Y., Bai, X., et al., 2020. Non-negligible stack emissions of non-criteria air pollutants from coal-fired power plants in China: condensable particulate matter and sulfur trioxide. Environ. Sci. Technol. 54, 6540 6550. Yaghoubi, J., Yazdanpanah, M., Komendantova, N., 2019. Iranian agriculture advisors’ perception and intention toward biofuel: green way toward energy security, rural development and climate change mitigation. Renew. Energy 130, 452 459. Yao, S., Yang, Y., Song, H., Wang, Y.A., Wan, H., 2015. Quantitative industrial analysis of lignocellulosic composition in typical agro-residues and extraction of inner hemicelluloses with ionic liquid. J. Sci. Ind. Res. 74, 58 63. Yusuf, A.A., Inambao, F.L., 2019. Bioethanol production techniques from lignocellulosic biomass as alternative fuel: a review. Int. J. Mech. Eng. Technol. 10, 34 71. Zhang, Y.H.P., 2008. Reviving the carbohydrate economy via multi-product lignocellulose biorefineries. J. Ind. Microbiol. Biotechnol. 35 (5), 367 375.

Relevant Websites https://www.epa.gov. https://www.enerdata.net/publications/executive-briefing/biofuels-market-dynamics.html. https://www.eesi.org/topics/bioenergy-biofuels-biomass/description. https://www.greenfacts.org. https://sunliquid-project-fp7.eu/wp-content/uploads/2014/09/factsheet_sunliquid_en.pdf.

Chapter 2

Worldwide scenario of biofuel production 2.1

Introduction

Bioenergy is currently drawing interest as a sustainable energy source that not only copes up with increasing prices of energy, but also provides income to deprived farmers and countryside communities around the globe. Increasing cost of fuel, rising energy demand, apprehensions over global warming and increased frankness to sustainable energy resources, domestic energy security and the drive for development into new markets for crops in the face of world trade outlooks are all factors motivating attention in increasing the usage of bioenergy. Among all forms of renewable energy used by human beings, bioenergy ranks number one. During the last twenty years, significant investment has been made to produce biofuels. In general, the global development and consumption of bioenergy and biofuels are gradually increasing, mainly in the cellulosic bioethanol and hydro treated vegetable oils sectors. Guo (2020) reported that by 2050, biofuels can provide up to 27% of world transportation fuel. United States and Brazil are the leading biofuel producers. These two countries account for 87% of worldwide production of biofuels driven by governmental support. Despite the intense interest in this area, there are not many players in this field. Biofuels are generally produced from organic matter obtained from living organisms or their products (Biomass). They can be used as a substitute to fossil fuels. Increase of prices of fuel and energy demand as well as global warming issues are the main reasons which drive massive interest in investigating natural and renewable sources for meeting the requirement of fuels and energy. Over the last few years, biofuels have been considered as an alternative to oils on global basis. The main elements behind their market development are their reduced carbon emissions as compared to conventional fuels and their positive effect on countryside development, along with the existing high prices of oil. Researchers are exploring a wide range of feedstock, mostly inedible crops and wastes for generating cost-efficient, high yield and environment friendly bioenergy having lowest amount of emissions. Table 2.1 shows biofuel yields by feedstock. Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00004-2 © 2023 Elsevier Inc. All rights reserved.

25

26

Cellulases in the Biofuel Industry

TABLE 2.1 Biofuel yields by feedstock. Ethanol (gallons per acre) Maize (U.S)

401

Maize (China)

213

Sugar cane (Brazil)

585

Sugar cane (India)

483

Cassava (Nigeria)

158

Cellulosic switch grass

1150

Biodiesel (gallons per acre) Soyabean (U.S)

59

Oil palm (Malaysia)

506

Rapeseed

110

Castor

140

Jatropha

170

Microalgae

5020

Source: Based on https://css.umich.edu/sites/default/files/Biofuels_CSS08-09_e2021_0.pdf; Chisti, 2007; United Nations Food and Agriculture Organization (2008); Oak Ridge National Laboratory (2005); Fulton (2006).

In 2020, the production of biofuels was 1677 thousand barrels of oil equivalent per day, as compared to the 187 thousand barrels of oil equivalent per day produced in 2000. Growth is mostly driven by policies which promote the production and use of biofuels because of the awareness that it might provide energy security and reduced emissions of greenhouse gases in relevant segments. Biofuels are advantageous because of its lower environmental impact as compared to fossil fuels and also its consumption of waste materials that would usually be thrown away (https://www.statista.com/statistics/274163/global-biofuel-production-in-oil-equivalent/). “According to the International Energy Agency (IEA) production of biofuels is expected to increase by 15% during the next five years, reaching 165 billion liters. It is predicted that by 2023, biofuels will account for about 90% of the renewables used in transport. Fuel ethanol accounts for about two third of biofuel production growth, whereas biodiesel and hydrotreated vegetable oil account for the rest. The countries in Asia account for most of the growth in biofuel output during the next five years. China, India and member states of the Association of Southeast Asian Nations represent half of the worldwide expansion in biofuel production. Latin America is responsible for an extra 45% of that growth, especially Brazil” (Bajpai, 2021; http://ethanolproducer.com/articles/15673/iea-predicts-growth-in-globalethanol-production-through-2023). The global production of

Worldwide scenario of biofuel production Chapter | 2

27

biofuel increased ninefold between 2000 and 2020. In 2021, the production will rebound in 2021 but will slightly remain below 2019 (https://www.enerdata.net/ publications/executive-briefing/biofuels-market-dynamics.html). Table 2.2 shows Annual World Fuel Ethanol Production (Mil. Gal.). Fig. 2.1 shows Renewable ethanol production by end use

TABLE 2.2 Annual world fuel ethanol production (Mil. Gal.). Region

2015

2016

2017

2018

2019

2020

% of world production

United States

14,807

15,413

15,936

16,091

15,778

13,926

53%

Brazil

7200

6750

6650

7990

8590

7930

30%

European Union

1360

1360

1420

1450

1370

1250

5%

China

770

670

800

770

1000

880

3%

India

190

280

200

430

510

515

2%

Canada

450

460

460

460

520

428

2%

Thailand

310

340

390

390

430

400

2%

Argentina

220

240

290

290

280

230

1%

Rest of World

393

487

454

529

522

500

2%

Total

25,700

26,000

26,600

28,400

29,000

26,059

Source: Reproduced with permission from https://ethanolrfa.org/markets-and-statistics/annual-ethanol-production.

FIGURE 2.1 Renewable ethanol production Reproduced with permission.

by end use

(https://www.clariant.com).

28

Cellulases in the Biofuel Industry

2.2

Status of biofuel production

Commercial production of bioenergy has seen steady growth in some countries. It dates back to the use of corn for production of ethanol (Msangi et al., 2007). Both biodiesel and bioethanol are produced all through the world. However, bioethanol is produced on a bigger scale as compared to biodiesel. Bioethanol is mostly produced and consumed in different parts of United States whereas biodiesel is produced in European Union. Production of ethanol on a large scale is being explored in African countries like Zimbabwe, South Africa, Malawi, Ghana and Kenya,. In general, it has been estimated that enough bioethanol is being produced in the world; can replace about 2% of total gasoline consumption (Banerjee et al., 2019). Production of ethanol from lignocelluloses is being developed. This is a new generation technology. This technology uses enzymes for producing ethanol and is being developed in North America, especially in Canada.

2.2.1

United States

The United States is the foremost biofuel producer in the world and has the growing and largest fuel ethanol market in the world. Table 2.3 shows historical biorefinery count and Table 2.4 shows annual US fuel ethanol production. Tables 2.5 and 2.6 show US Fuel Ethanol Plant Production Capacity as of January 1, 2021 and US Biodiesel Plant Production Capacity as of January 2021. In 2018, The United States contributed 45.5% of the world’s biofuel production. It is the leading producer of biogasoline in the world; has a 55.4% share which is equal to about 1047 thousand barrels/day (https://www.nsenergybusiness.com/features/top-biofuel-production-countries/). TABLE 2.3 Historical biorefinery count and production capacity. Year

Installed ethanol biorefineries

Total installed production capacity (mgy)

Average capacity per biorefinery (mgy)

2000

56

2007

36

2005

95

4294

45

2010

204

14,073

69

2015

214

15,594

73

2020

208

17,436

84

Source: Reproduced with permission from https://ethanolrfa.org/file/1007/ RFA_Outlook_2021_fin_low.pdf.

Worldwide scenario of biofuel production Chapter | 2

29

TABLE 2.4 Annual US fuel ethanol production. Year

Million (Gallons)

2011

13,929

2012

13,218

2013

13,293

2014

14,313

2015

14,807

2016

15,413

2017

15,936

2018

16,091

2019

15,776

2020

13,926

Source: Reproduced with permission from https://ethanolrfa.org/markets-and-statistics/annualethanol-production.

TABLE 2.5 US fuel ethanol plant production capacity as of January 1, 2021. Nameplate capacity PAD district

Number of plants

(MMgal/year)

(Mb/d)

PADD 1

4

347

23

PADD 2

178

16,271

1061

PADD 3

4

405

26

PADD 4

4

200

13

PADD 5

7

323

21

US Total

197

17,546

1145

Source: Reproduced with permission. Form EIA-819, Monthly Report of Biofuels, Fuels from NonBiogenic Wastes, Fuel Oxygenates, Isooctane, and Isooctene Fuel Ethanol Production Capacity is intended to measure estimated gallons of fuel alcohol that a plant is capable of producing over a period of one year (365 consecutive days) starting on the first day of each report month. https:// www.eia.gov/petroleum/ethanolcapacity.

The United States is also the leading producer of biodiesel in the world; has a share of 19.4%, which is equal to almost 136.18 thousand barrels/day as of end 2018.

30

Cellulases in the Biofuel Industry

TABLE 2.6 US biodiesel plant production capacity as of January 1, 2021. Production capacity PAD district

Number of plants

(MMgal/year)

(Mb/d)

PADD 1

13

152

10

PADD 2

37

1483

97

PADD 3

15

580

38

PADD 4

0

0

0

PADD 5

10

194

13

US Total

75

2409

157

Source: Reproduced with permission. Form EIA-819, Monthly Report of Biofuels, Fuels from NonBiogenic Wastes, Fuel Oxygenates, Isooctane, and Isooctene Biodiesel Production Capacity is intended to measure estimated gallons of biodiesel that a plant is capable of producing over a period of one year (365 consecutive days) starting on the first day of each report month. https:// www.eia.gov/biofuels/biodiesel/capacity/.

The United States mostly uses corn as the main raw material for producing bioethanol and soybeans for producing biodiesel. United States Department of Agriculture (USDA) has published a Grain Crushings and CoProducts Production report. According to this report, more than 5.55 billion bushels of corn were used for producing fuel ethanol in 2018. According to Energy Information Administration (EIA), the United States produced about 16.061 billion gallons of ethanol in 2018. Iowa being the leading state produced about 4.328 billion gallons per year. As of 1 January 2019, The EIA has reported 200 fuel ethanol production plants, with a nameplate capacity of 16,868 MMgal/year. Table 2.7 shows global commercial scale cellulosic ethanol plants and Table 2.8 shows the status of the commercial lignocellulosic ethanol facilities in United States (Zhang, 2019). “Cellulosic ethanol industry is still in its infancy. In the United States, the first commercial scale plants to produce cellulosic biofuels started operating in 2013. In the following 5 years, cellulosic ethanol production grew from 0 to 10 million gallons and most likely topping 15 million in 2018. However, that is far from the Renewable Fuel Standard’s original target of 7 billion gallons of cellulosic biofuel by 2018 and 16 billion by 2022. Of all five commercial cellulosic ethanol plants that were built/to be built in the US from 2010 to 2016, only POET’s Emmetsburg, Iowa facility is still in operation in 2019. In 2017, the total cellulosic ethanol produced was less than half the nameplate capacity (25 million gallons year—1) of this single plant (Table 2.9) (http://www.forbes. com). Currently the United States has a target of 136,260 million liters per year (ML/year) of renewable fuels production by 2022. This target is only

TABLE 2.7 Global commercial scale cellulosic ethanol plants (Padella et al., 2019). Company

Project

Country output

Capacity (ktons)

Status

Start-up year

Abengoa bioenergy biomass of Kansas, LLC

Commercial (acquired by Synata Bio Inc. [21])

US

75

Idle

2014

Aemetis

Aemetis Commercial

US

35

Planned

2019

Beta Renewables (acquired by Versalis [22])

Alpha

US

60

On hold

2018

Beta renewables (acquired by Versalis)

Energochemica

EU (Slovakia)

55

On hold

2017

Beta renewables (acquired by Versalis)

Fujiang bioproject

China

90

On hold

2018

Beta renewables 1 (acquired by Versalis)

IBP-Italian Bio Fuel

EU (Italy)

40

Idle

2013

Borregaard industries AS

ChemCell Ethanol

Norway

16

Operational

1938

Clariant

Clariant Romania

EU (Romania)

50

Under construction

2020

COFCO Zhaodong Co

COFCO Commercial

China

50

Planned

2018

DuPont

Commercial facility Iowa (acquired by VERBIO [23])

US

83

Idle

2016

Enviral

Clariant Slovakia

EU (Slovakia)

50

Planned

2021

Fiberight LLC

Commercial Plant

US

18

Under construction

2019 (Continued )

TABLE 2.7 (Continued) Company

Project

Country output

Capacity (ktons)

Status

Start-up year

GranBio

Bioflex 1

Brazil

65

Operational

2014

Henan Tianguan Group

Henan 2

China

30

Idle

2011

Ineos Bio

Ineos Bio Indian River County Facility (acquired by Alliance BioProducts in 2016 [24])

US

24

Idle

NA

Longlive Bio-technology Co. Ltd.

Longlive

China

60

Idle

2012

Maabjerg energy concept consortium

Flagship integrated biorefinery

EU (Denmark)

50

On hold

2018

POET-DSM Advanced Biofuels

Project Liberty

US

75

Operational

2014

Ra´ızen Energia

Brazil

Brazil

36

Operational

2015

St1 Biofuels Oy in cooperation with North European Bio Tech Oy

Cellunolix

EU (Finland)

40

Planned

2020

Source: Padella, M., O’Connell, A., Prussi, M., 2019. What is still limiting the deployment of cellulosic ethanol? Analysis of the current status of the sector. Appl. Sci. 9, 4523. (https://doi.org/10.3390/app9214523). This Table is distributed under the terms of the Creative Commons Attribution 4.0 International License (http:// creativecommons.org/licenses/by/4.0/).

Worldwide scenario of biofuel production Chapter | 2

33

TABLE 2.8 The status of the US commercial lignocellulosic ethanol facilities. Company

Location

DuPont Nevada

Iowa

Mascoma

Kinross, MI

POET LLC

Raw material

Capacity (mg/year)

Status

30

Sold to Verbio in Nov. 2018

Wood waste

20

Construction halted in 2013

Emmetsburg, IA

Corn stover

20 25

Operational in Sep. 2014

Abengoa Bioenergy

Hugoton, KS

Wheat straw

25 30

2013 2016 Bankrupt

BlueFire Ethanol

Fulton, MS

Multiple sources

20

Construction halted 2011

Source: Based on Zhang, C., 2019. Lignocellulosic ethanol: technology and economics. In: Yun, Y. (ed.), Alcohol Fuels Current Technologies and Future Prospect. IntechOpen Limited, London. https://doi.org/10.5772/ intechopen.86701.

achievable with a majority of this renewable fuel coming from lignocellulosic material, such as corn stover, wood, switch grass, wheat straw and purpose grown energy crops. Demonstration scale cellulosic ethanol plants are under construction as part of the government’s goal to make cellulosic ethanol cost competitive. The plants cover a wide variety of feedstocks, conversion technologies and plant configurations to help identify viable technologies and processes for full-scale commercialization. All demonstration plants, which are sized at 10% of a commercial-scale biorefinery, are expected to be operational soon” (Bajpai, 2021).

2.2.2

Brazil

“Brazil has been for decades the world’s largest producer and consumer of fuel ethanol, but was overtaken by United States in 2006. Brazil is the world’s top exporter. Brazil was the first country to produce bioethanol on a commercial scale, via their government’s Proalcool program that was started in 1975 for using fuel alcohol from sugar cane as a gasoline substitute in response to increasing prices of oil (Soccol et al., 2010). Brazil is now the world’s second largest producer in the world and is the largest exporter of fuel ethanol. Presently, Brazil has more than 80% of its vehicles running with bioethanol. Brazil’s total ethanol production for 2019 in Brazil was estimated at 34.45 billion liters, an increase of 4% in comparison to the revised figure for

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Cellulases in the Biofuel Industry

TABLE 2.9 Operating cellulosic ethanol plants in the US. Name of the company

Raw material

Scale

Year of operation

Capacity (mil gal)

American Process, Alpena, MI

Wood chips

Commercial scale

2012

0.95

American Process, Thomaston, GA

Wood chips

Commercial scale

2013

N/A

Calgren Renewable Fuels, Pixley, CA

Cow manure

Commercial scale

2015

N/A

DuPont Nevada, IO

Corn stover

Commercial scale

2015

30

Gulf Coast Energy Livingston, AL

Wood waste

Pilot scale

2009

20

Indian River Bioenergy Center Vero Beach, FL

Municipal solid waste

Commercial scale

2013

8

LanzaTech Soperton, GA

Wood waste

Pilot scale

2014

0.09

Pacific Ethanol Stockton, CA

Corn kernel fiber

Commercial scale

2015

0.75

Project LIBERTY (POET) Emmetsburg, IA

Corn stover, corn cobs, leaves, husk, stalk

Commercial scale

2014

20

Quad-County Galva, IA

Corn kernel fiber, corn

Commercial scale

2014

2

Renmatix Rome, NY

Wood chips, tall grasses, corn stover, bagasse

Demonstration

2008

N/A

Summit Natural Energy Cornelius, OR

Food processing and agricultural waste

Pilot scale

2009

N/A

Tyton Biofuels Raeford, NC

Tobacco waste

Pilot scale

2010

15

ZeaChem Boardman, OR

Wood

Demonstration

2013

0.25

Source: Based on Nguyen et al. (2017); Zhang (2019); Voorhis (2016); Voegele (2014, 2018); McGlashen (2013); POET-DSM (2018).

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2018. Total ethanol production from corn in 2019 was estimated at 1.4 billion liters, an increase of 609 million liters in comparison to 2018. Total cellulosic ethanol production was estimated at 45 million liters, and represents an unimportant share of total ethanol production in Brazil. No noteworthy changes have been made to the current status of advanced biofuels research and development and production. Total 2019 domestic demand for ethanol (fuel and other uses) was estimated at 33.93 billion liters, up 2.19 billion liters compared to revised figure for 2018. Ethanol is mostly used for fuel in Brazil. Brazil’s total ethanol exports were estimated at 1.8 billion liters, an increase of 11% in comparison to total exports in 2018 (1.62 billion liters). Brazil’s total ethanol imports for 2019 were 1.2 billion liters, a reduction of 495 million liters relative to the earlier year (1.695 billion liters). Ethanol imports are only for fuel use and originate almost totally from the United States” (Bajpai, 2021). Brazil is the second highest producer of biogasoline in the world having a 31.5% share which amounts to about 595.35 thousand barrels/day. Brazil is also the second highest producers of biodiesel, having a share of 14.1% equivalent to about 99,000 barrels/day (https://www.nsenergybusiness.com/ features/top-biofuel-production-countries/). The Latin American countries mainly use sugar cane for the production of fuel ethanol and soybeans for producing biodiesel. Bagasse, that remains after crushing the sugar cane, is extensively used as fuel in sugar mill co-generation plants for meeting the onsite energy demand. In some cases, bagasse is used to provide spare electricity for export purposes. “For 2019, total ethanol production was estimated at 34.445 billion liters. Total ethanol production for fuel use is estimated at 31.387 billion liters, up three % from the preceding calendar year. Total ethanol production from corn in 2019 was estimated at 1.4 billion liters, an increase of 609 million liters in comparison to 2018. Ethanol from corn represents four % of total ethanol production. There are presently ten plants which produce ethanol from corn in Brazil in the states of Mato Grosso and Goias and four plants under construction which should start operations in 1 2 years. The majority of the units are full-plant type, example dedicated corn-only. Some are flexplants, producing ethanol from sugarcane and corn. There are two new projects recently launched and at least five projects either in the licensing or financing stage. According to the Corn Ethanol National Union (UNEM) Brazil is producing 2.6 billion liters of corn ethanol in 2020 and is expected to produce 8 billion by 2028. Total cellulosic ethanol production for 2019 was at 45 million liters, an increase of 20 million liters in comparison to 2018, assuming that the existing plants are able to overcome the ongoing plant-level operational/mechanical challenges. This amount still represents an unimportant share of total ethanol production in Brazil” (Bajpai, 2021). “The total number of sugar-ethanol mills in 2019 is 370 units, one additional unit in comparison to the revised figure for 2018 (369 units). Hydrated

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Cellulases in the Biofuel Industry

ethanol production capacity for 2019 is reported unchanged from the revised figure for 2018 at 43.105 billion liters. This figure reflects the authorized hydrated ethanol production capacity of 233,000 m3 per day, as reported by ANP, and assumes an average of 185 crushing days. Ethanol installed industrial capacity depends on annual decisions made by individual plants to produce sugar and/or ethanol. Post contacts report that the industry responds to the ratio of 40:60 to switch between sugar and ethanol production or vice versa from harvest to harvest. Once producing units adjust their plants to produce a set ratio of sugar/ethanol in a given year, there is much less flexibility to change it during the crushing season (https://www.fas.usda.gov/data/ brazil-biofuels-annual-5). The number of plants producing bioethanol from sugarcane in Brazil is expected to increase in the next few years (Amorim et al., 2009; Basso and Rosa, 2010). In Brazil, ethanol blends are compulsary (E20 to E25) and anhydrous ethanol is also available from thousands of filling stations. Besides, there are 6 million flex-fuel vehicles in Brazil and 3 million are able to run on E100. In Brazil, Bioethanol now accounts for about 50% of the transport fuel market, where gasoline may now be regarded as the alternative fuel” (Bajpai, 2021). In Brazil, the first commercial-scale cellulosic ethanol plant was constructed at Sa˜o Miguel dos Campos, Alagoas. The production was started in September 2014. The existing production capacity is B22 m gallons annually. The plant is using technology developed by Beta Renewables / Biochemtex PROESA. The hydrolytic enzymes are supplied by Novozymes. For fermentation DSM yeasts are used. The cost of the plant was $190 m. On the co-generation system, $75 m was spent. In May 2013, the Brazilian Development Bank provided a $149 m loan to Bioflex Agroindustrial which is a subsidiary of GranBio, for building of the plant for converting bagasse and residual sugar cane straw into second generation ethanol. BNDES said “The use of straw and bagasse will allow industrial productivity of ethanol to reach around 10,000 L per hectare, corresponding to an increase of up to 45% compared to current levels.” GranBio plans to construct a new ethanol plant each year up to 2020 with a planned investment of US$1.7 billion. In December 2014, at the Costa Pinto sugarcane mill, production started at the $100 million, 40 MMly Ra´ızen Energia S/A commercial second generation ethanol plant. This plant has a capacity of 10MMgy. The technology developed by Iogen Energy which is a joint venture of Ra´ızen and Iogen Corporation, is used. Bagasse is used as feedstock for producing ethanol Enzymes are supplied by Novozymes. By 2024, Raizen is planning to produce up to 1 billion liters of bioethanol from cane straw and bagasse (https://www. etipbioenergy.eu/value-chains/conversion-technologies/advanced-technologies/ sugar-to-alcohols/commercial-cellulosic-ethanol-plants-in-brazil). Ra´ızen SA has planned to build its second cellulosic ethanol plant in the country. The production capacity of this plant is 82 million liters per year which

Worldwide scenario of biofuel production Chapter | 2

37

is two times the capacity of its first plant. This is the only company in the world which is operating two cellulosic ethanol plants on a commercial scale. It aims to supply to increasing worldwide demand for cellulosic biofuels. More than 90% of the output of the new plant has already been sold to global energy producer under long-term contracts. The new plant produces sugar, first generation ethanol, and power cogeneration from sugarcane biomass and also includes the company’s first biogas plant that was inaugurated in October 2020. This plant will be part of the Bonfim bioenergy park located in Guariba in the state of Sa˜o Paulo and will start by 2023, increasing Ra´ızen’s total installed capacity to 120 million liters per annum of cellulosic ethanol. (https://www.reuters.com/business/energy/brazils-razen-build-second-cellulosic-ethanol-plant-filing-2021-06-25/).

2.2.3

European Union

“The European Commission is supporting biofuels for reducing greenhouse gas emissions; boosting the decarbonization of transport fuels; diversifying fuel supply sources; offering new income opportunities in rural areas and developing long-term replacements for fossil fuel. In European Union, ethanol is mainly produced from sugar beets and wheat. Sugar beets prove to be a good feedstock for European bioethanol production. Because sugar beets have a much larger yield per hectare than wheat, the European Union currently produces 2 million more tons of sugar beet than wheat on 20 million less hectares of land. Additionally, sugar beets produce more ethanol per hectare: a hectare of sugar beets can produce 30 hectoliters more ethanol, on average, than wheat. Also, sugar beet ethanol is shown to have a more energy-efficient production process than wheat ethanol. Currently, the most important bioethanol producers are France, Spain, Germany, Sweden, Poland and Italy. In comparison with the United States and Brazil, European Union ethanol for fuel production is still very modest. These 2 countries are now in competition for the title of world biggest producer. For both these 2 countries the main driver is reducing dependency from foreign fossil fuel sources. In Europe, except for Sweden, and unlike the United States or Brazil, ethanol is not incorporated directly but transformed into ETBE (obtained by reacting isobutene, a liquefied petroleum gas, with ethanol) before being blended with gasoline. One reason for this regional particularity is the obligation to properly account for motor fuel properties such as volatility, since pure ethanol makes ethanol/gasoline blends more volatile. Another advantage of this practice is that it avoids separation of the alcohol and gasoline phases in the presence of traces of water. EU has set a 10% target for renewable energy use in transport for 2020. That target increases to 14% in 2030, with advanced biofuels counting double to the target. Taking double-counting into account, biofuels accounted for 7.1% of energy use in transportation last year and are expected to increase to 7.3%

38

Cellulases in the Biofuel Industry

this year. The increase is expected to be supported by increased imports (https:// www.greencarcongress.com/2019/08/20190812-fas.html). Fuel ethanol production reached about 5.443 billion liters in the European Union in 2018, which is higher from 5.38 billion liters in 2017. European Union fuel ethanol production increased to 5.505 billion liters in 2019. Fuel ethanol imports reached about 505 million liters in 2018, up from 238 million liters in 2017. Fuel ethanol imports are increased slightly in 2019, reaching 570 million liters. The European Union exported about 91 million liters of fuel ethanol last year, up from 41 million liters in 2017. Exports this year are expected to reach 95 million liters. Ethanol consumption has reached 7.22 billion liters in 2019, up from 7.082 billion liters in 2018 and 6.89 billion liters in 2017. The average ethanol blend rate in the European Union was 5.7% in 2018, up from 5.4% in 2017. The blend rate is expected to increase to 5.9% this year. European Union is presently having 58 first generation ethanol plants. This number has remained stable since 2017. The capacity of those facilities was 8.66 billion liters in 2018, up from 8.6 billion liters in 2014. Capacity remained at 8.66 billion liters through 2019. Capacity use is expected to be higher slightly, from 70% in 2017 and 2018 to 71% in 2019. The European Union also presently has two cellulosic ethanol refineries, having a joint capacity of 60 million liters” (Bajpai, 2021). “Sugar beet is expected to be the primarily feedstock for European Union ethanol production this year, reaching 8.145 million metric tons, followed by wheat with 5.665 million tons, corn with 5 million tons, triticale with 720,000 tons, barley with 430,000 tons and rye with 418,000 tons. Germany is expected to be the top European Union consumer of ethanol this year, with 1.505 billion liters, followed by the U.K. with 925 million liters and France with 880 million liters. Spain, Poland, the Netherlands, Italy and Sweden are also among the top eight ethanol consuming countries in the European Union” (http://ethanolproducer. com/articles/16422/reporteu-ethanol-consumption-to-increase-in-2019). France will become the main ethanol producer in the European Union. It is producing “1 billion liters, followed by Germany at 785 million liters, the United Kingdom with 695 million liters, Hungary with 645 million liters, Belgium with 645 million liters, the Netherlands with 565 million liters, Spain with 522 million liters, Poland with 265 million liters and Austria with 235 million liters” (ethanolproducer.com). Germany ranks third among the top biofuel producers in the world. In 2018, the production was 75.8 thousand barrels/day. It accounted for 2.9% of the global biofuel production capacity. German Association of Biodiesel Producers have reported that in 2018, 3.2 million tonnes of biodiesel was produced. The major raw materials used for production of this fuel are rapeseed and used cooking oil. The country is exporting major volumes of its biodiesel production. According to Union zur Fo¨rderung von Oel- und Proteinpflanzen (UFOP), the Netherlands is the top importer of fuel produced in Germany. Other importers of biodiesel are Austria, Belgium, Poland, and the United States.

Worldwide scenario of biofuel production Chapter | 2

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¨ lmu¨hle Hamburg The main producers of biodiesel in Germany is ADM O which is a subsidiary of the American group Archer Daniels Midland Company. ADM Rothensee, a subsidiary of ADM Hamburg Aktiengesellschaft, is involved in the origination of rapeseed from farmers, farmers’ associations, cooperatives, and local merchants.

2.2.4

Argentina

In 2018, Argentina produced 70.6 thousand barrels/day of biofuels. This accounted for 2.7% of the total production capacity worldwide. The country is mainly producing bioethanol and biodiesel. According to a USDA report, the country has nineteen bioethanol plants which produce 1.4 billion liters, with thirteen plants using sugarcane as raw material and the remaining plants use corn. “The 2006 Biofuels Law 26,093, enacted by Argentina, mandated a 5% mix of bioethanol in gasoline and biodiesel in diesel starting from 2010. The blend mandated a 12% mix for ethanol and a 10% mix for biodiesel since 2016. The country also passed a law in 2008 to promote the production of bioethanol from sugarcane” (https://www.nsenergybusiness.com/features/topbiofuel-production-countries/). Presently, Argentina is operating a total of 33 biodiesel plants. The largest plant can produce up to 700 million liters annually. ACA Bio Cooperativa Limitada, is investing $53 m for expansion of its existing ethanol production facility.

2.2.5

China

China is the world’s fourth-largest fuel ethanol producer and consumer after the United States, Brazil, and the European Union. The country accounted for 2.6% of total biofuel production last year. China has ambitious growth targets for second generation bioethanol from waste biomass (Zhenhong, 2006; Yan et al., 2010). In 2018, China produced 68 thousand barrels/day. Over the last year, China increased the production capacity by 258 million liters to a total of 5,258 million liters. Bioethanol production in China is mostly driven by it’s efforts to improve the air quality. In 2018, China imposed strict air pollution reduction measures across Tianjin, Beijing, and Hebei province. In 2020, China is using 10% ethanol-gasoline blend. It is also planning to shift renewable fuel production to commercial scale production of cellulosic ethanol by 2025. For avoiding reliance upon imports, China is required to increase fuel ethanol production from 12,670 to 19,005 million liters for meeting domestic demand. “In China bioethanol plants use corn, wheat and cassava. Sweet sorghum and sugar cane have future potential. Regarding second generation feedstocks, in

40

Cellulases in the Biofuel Industry

2008, COFCO built the first non-grain, 2 million tons cassava fuel ethanol factory in Guangxi. In 2012, Henan Tianguanbuilt built its first 10 k-ton cellulosic fuel ethanol pilot program. In 2013, Shandong Longlive built its 50 k-ton corn cob fuel ethanol production line. In 2014, Zonergy built its 30 k-ton sweet sorghum fuel ethanol factory” (https://noaw2020.eu/wp-content/uploads/2018/12/C1-1COFCO-Wuguoqing.pdf). “Ethanol gasoline is now being promoted all through China. Fuel ethanol production will be increased by 5 to 6 times on the base of present scale. A cellulosic fuel ethanol plant is being constructed at the capacity of 50,000 ton/y. By 2025—attempts will be made to realize large scale production of cellulosic fuel ethanol. The overall development of advanced biological liquid fuel technologies, equipment and the industry will reach an international advanced level, and an improved market oriented operating mechanism will be established. So far, Chinese government has made mandatory the use of E10 in nine provinces (at central and northern China) that account for about one-sixth of that country’s vehicles. Officials say that this mandate aims at reducing the oil demand—that is currently 40% imported—and also aims at improving air quality in big cities. However, Chinese authorities also frequently highlight the targets of helping stabilize grain prices and raising farmer’s income. In China, more than 80% of ethanol is produce from grains (corn, cassava, rice etc.), about 10% from sugar cane, 6% from paper pulp waste residue and the remainder is produced synthetically. So far, no significant amounts of biodiesel have been produced in China. There are no industrial scale biodiesel plants in the country. Fuel ethanol is exempted from consumption tax (5%) and value-added tax (17%). In the recent years, the fuel ethanol market in China has remained narrow minded. Imports were not allowed till 2015, and China hardly ever produced surplus volumes to export. As additional duties were implemented earlier this year, in China, fuel ethanol market backed away further from the global market. China is producing a wide range of ethanol products on a commercial-scale, including fuel ethanol, potable alcohol and industrial chemicals. Contrasting other major ethanol producers, China’s major end use market for ethanol is to produce industrial chemicals and not fuel ethanol. In the past, China’s ethanol output has followed national policy priorities. Since 2016, China’s corn processors, including fuel ethanol and industrial chemical producers, are enjoying the advantage of corn processing subsidies based on throughput volumes. In addition, China is expanding blending of ethanol with gasoline on a nationwide basis by doubling the number of administrative regions implementing E10, and investments supported by government for expanding the production capacity. Despite the strong central government support through policies and financial backing, ethanol sector of China faces near-term structural challenges and long term feedstock supply challenges for producing enough fuel ethanol for meeting ambitious E10 goals. Most experts consider China’s E10 consumption target to be virtually unachievable at the existing pace of market development” (Bajpai, 2021; https://apps.fas.usda.gov/newgainapi/api/report/ downloadreportbyfilename?filename 5 Biofuels%20Annual_Beijing_China% 20-%20Peoples%20Republic%20of_8-9-2019.pdf).

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“In 2019, fuel ethanol consumption was estimated at a record 4311 million liters (3.4 million tons), up 1397 million liters (1.1 million tons) from 2018 due to newly implemented provincial and municipal government directives for expanding fuel ethanol blending into gasoline supplies. All fuel ethanol in China’s transportation fuel supply must meet national standard GB18350 for denatured fuel ethanol. However, China does not administer a national standard for ethanol inclusion rates in retail gasoline formulations. Industry sources report that although many provinces and cities have adopted E10 in full, actual compliance and the standardization of retail gasoline fuel formulations vary greatly within a given transportation fuel market. MTBE competes with ethanol as a gasoline oxygenate for improving engine performance in China. Recent regulations under the Blue Sky Protection Plan (2018 2020) limit the use of MTBE, which further supports expanded ethanol demand. China is not producing ethanol-derived bio- ETBE (ethyl tert-butyl ether) in large scale. Fuel blending formulations incorporating ETBE need additional processing, which have not been used in China” (Bajpai, 2021).

2.2.6

Australia

The Australian biofuels market is currently in the development stage, with the government and private entities engaging in promotion of the technology across the country. Currently E5, E10, E15 and E85 are available throughout Australia as forms of unleaded petrol (Puri et al., 2012). Since 2000, the Australian government has been supporting fuel ethanol and has provided range of tax exemptions and production subsidies. Only three ethanol plants having a combined 440 million liters of annual capacity were operating last year, a number that remained steady for almost ten years and is expected to be maintained through 2020. The plants are expecting to operate at 53.4% capacity this year, which is down from 56.1% in 2019. Australian ethanol production is expected to reach 235 million liters this year, down from 247 million liters in 2019. Consumption is expected to be at 220 million liters this year, down from 239 million liters last year (http://www.biodieselmagazine.com/). The biodiesel production capacity is 100 million liters per year (0.17% of the global output). “The annual production of these two biofuels have an equivalent amount of energy as 375 million liters of diesel. Currently, domestic biodiesel production is approximately one per cent of total domestic diesel consumption, indicating biofuel’s extremely small market share. In addition, there is currently no aviation biofuel production in Australia” (https://arena. gov.au/assets/2019/11/biofuels-and-transport-an-australian-opportunity.pdf). Wheat, molasses, sorghum and barley are the major ethanol feedstock used in Australia. The average national blend rate for ethanol is expected to be at 1.4% this year, flat with 2018 and 2019. At present Australia is having three biodiesel plants having a combined capacity of 107 MMly of nameplate capacity, flat with 2019. The biodiesel

42

Cellulases in the Biofuel Industry

capacity was highest in 2013 when Australia was having seven biodiesel plants having a combined capacity of 312 MMly. This year, the biodiesel plants are expected to operate at 39.3% of capacity which is higher from 26.2% in 2019. Production of biodiesel is expected at 42 million liters in 2020. In 2019, it was 28 million liters. Consumption of biodiesel is expected at 26 million liters this year, flat with 2019. The average blend rate is expected to be at 0.2% this year which is flat with 2019. In 2014, the blend rate was highest at 3.8%. “The Australian car fleet can use substantial volumes of biofuel. In 2015, 95% of the 2.5 million cars in Queensland alone were compatible with ethanol. This figure has reached 98% in 2020. Most automakers are now manufacturing cars fit for higher ethanol fuel blends. Australian plants can presently only produce sufficient biofuel to satisfy 0.6% of the nation’s demand for transport fuel, which is very much behind the global average of 2.7%. This capacity has reduced in the last 10 to 15 years because of plant closures. In the meantime, existing plants are working below the full capacity. In Brazil car manufacturers have also shown the ability of industry for developing engines capable of running on high biofuel blends. Brazil now has more than 25 million “flex” vehicles, which can run on 0% to 100% ethanol. There is an increasing domestic awareness of Australia’s lagging biofuel production relative to the global average, and of the advantages and opportunities higher production would bring to the broader community (Bajpai, 2021). This awareness is resulting in state government initiatives like the Advance Queensland Biofutures 10-Year Roadmap and Action Plan, and the Bioenergy Roadmap for South Australia. A 40-fold expansion over the next 30 years is a significant challenge for the industry. Achieving it will require sustainable growth in feedstock resources, the application of new technologies and a consistent development of new production facilities. The investment required by production facilities alone is estimated at between $25 billion and $30 billion” (Bajpai, 2021; http://www.cefc.com.au). “The Australian biofuel industry is although small, it has immense experience using low value waste streams for remaining viable in periods of policy uncertainty and low consumer demand. But it is hindered by higher costs, juvenile technology, its small scale. Also a suitable policy and regulatory framework is lacking. Table 2.10 shows the biofuel plants in Australia, including production capacity. Especially, several biodiesel plants built in the early 2000s are not operating anymore. The shutting down of these plants is the result of many factors. These include reduced cost imports monopolizing the Cleaner Fuels Grant Scheme, increasing the price of raw material and inconsistent product quality. Australia has larger areas of cultivable and semi-cultivable land; established agricultural, forestry and engineering industries; important raw materials and superb solar resources. All of these give Australia an exclusive opportunity for meeting its projected biofuel requirements. In Australia, bioethanol is produced from grain and sugar cane molasses. Australia only produces small amounts of biodiesel to date. However, several industrial scale plants are in

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TABLE 2.10 Commercial-scale bioethanol plants in Australia. Company

Location

Feedstock

Production

Wilmar Sucrogen

Sarina, Qld

Sugarcane

60 ML/year bioethanol

United Petroleum

Dalby, Qld

Red sorghum

80 ML/year bioethanol

Manildra Group

Bomaderry, NSW

Waste starch

300 ML/year bioethanol

Source: CEFC and ARENA (2019). Biofuels and Transport: An Australian opportunity https://arena. gov.au/assets/2019/11/biofuels-and-transport-an-australian-opportunity.pdf.

the construction phase. The fuel ethanol industry is still waiting for the big breakthrough, but political support for the industry is growing. Australia is a net importer of crude oil, and biofuels are considered to be increasingly important in view of surging world oil prices. Advocates of ethanol production also cite benefits for the ailing domestic sugar cane industry” (Bajpai, 2021).

2.2.7

Canada

Canada is presently producing comparatively lower volumes of fuel ethanol. Several initiatives are ongoing for boosting the production considerably. “Production is expected to increase significantly in the next few years if current and announced biofuels programs are implemented. To meet Kyoto Protocol commitments, the country aims to replace 35% of its gasoline use with E10 blends, requiring production of 350 million gallons of ethanol. Seven new plants with total capacity of 200 million gallons are planned under the Ethanol Expansion Program. Ontario, Saskatchewan, and Manitoba are already promoting ethanol through production subsidies, tax breaks, and blending requirements. In Canada, ethanol is produced almost entirely from cereals. Gasoline mixed with ethanol is now available at over several gas bars across Canada from Quebec to the Pacific, including the Yukon Territory. In several regions, ethanol blends are available for bulk delivery for farm and fleet use. The federal government and many provinces are offering tax incentives. USDA Foreign Agricultural Service’s Global Agricultural Information Network provides an overview of biofuel industry of Canada and the country’s changing biofuels policy framework (https://www.fas.usda.gov/regions/canada). Canada is at present in the process of developing a Clean Fuel Standard, which will enact a carbon intensity strategy when accounting for the amount of blended renewable fuel. Once finalized, the Clean Fuel Standard will replace the volumetric approach presently in place under federal renewable fuels regulations of Canada Environment and Climate Change Canada released the regulatory design paper for the Clean Fuel Standard in December 2018. There are separate requirements for liquid, gaseous and solid fuels. The ECCC

44

Cellulases in the Biofuel Industry

released a benefit-cost analysis framework for the Clean Fuel Standard in February 2019. Information revealed on the Clean Fuel Standard regulations to date shows the program will reduce the carbon intensity of liquid fuels to be reduced by 10 grams of carbon dioxide equivalent per megajoule below the reference carbon intensity (year 2016) by 2030. Under the proposed CFS, separate carbon intensity requirements would be established for subsets of fuels in the following sectors: transportation, building requirements, and industry,” said the authors in the report. “The proposed CFS will not differentiate between crude oil types that are produced domestically or are imported. The federal government will maintain national blending mandates in the shortterm, establishing an “expiration date” for the volumetric requirements through consultations with stakeholders. Proposed Clean Fuel Standard regulations for the liquid fuels are published in 2019. Regulations for gaseous and solid fuel streams will be established at a later date. At present, federal renewable fuels regulations in Canada need fuel producers and importers to have an average ethanol content of at least 5% based on the volume of gasoline produced or imported. The federal regulations also need 2% renewable content based on the volume of diesel fuel and heating distillate oil fuel producers and importers produce or import. Several provincial blend mandates are also in place. Manitoba has an 8.5% ethanol blend mandate for gasoline, whereas Saskatchewan has a 7.5% mandate and British Columbia, Alberta, and Ontario each have a 5% mandate. For renewable blends in diesel, British Columbia and Ontario have a 4% mandate, whereas Manitoba, Alberta and Saskatchewan each have a 2% mandate. Canada produced 1.83 billion liters of fuel ethanol this year, up from 1.75 billion liters in 2018. Imports are expected to reach 1.37 billion liters this year, down from 1.39 billion liters in 2018. Fuel ethanol consumption is expected to reach 3.2 billion liters this year, up from 3.14 billion liters in 2018. The United States supplies essentially all of Canada’s fuel ethanol imports. Canada presently has 13 ethanol plants in operation, a number that has been stable since 2015. Nameplate capacity, however, is up. Capacity is expected to reach 2.15 billion liters this year, up from 1.97 billion liters in 2018 and 1.872 billion liters in 2017. Capacity use is expected to be at 85% this year, down from 89% in 2018 and 92% in 2017. The overall ethanol blend rate in Canada is expected to reach 6.6% this year, up from 6.4% in 2018 and 6.2% in 2017” (Bajpai, 2021). “Greenfield Global, the largest producer of ethanol in Canada, has taken steps for expanding production at its Varennes biorefinery following the introduction of draft regulations on the minimum renewable fuel volumes in the province of Quebec. Welcoming the Government of Quebec’s draft regulations on the minimum volume of renewable fuel to be blended in gasoline and diesel, the producer added that it could result in a major expansion of its production in the province. The new proposal would set blending thresholds of 10% renewable fuel in gasoline, and 2% renewable fuel in diesel by 2021. This would increase to 15% in gasoline and 4% in diesel by 2025” (gain.fas.usda.gov).

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“Greenfield applauds the Quebec Government for its ambitious targets and firm commitment to transition the province’s energy,” said Jean Roberge, executive vice president and managing director of renewable energy at Greenfield. “Biofuels are indeed a sector for the future. The government is demonstrating true leadership on climate change and taking steps necessary to effectively and efficiently increase the use and production of renewable fuels, like ethanol, in the province. Greenfield’s Varennes biorefinery, which is the first ethanol plant in Quebec, has been in operation since 2007. The facility produces the lowest carbon intensity ethanol in Canada, and also corn oil and distillers’ grain. The company is presently considering adapting emerging advanced biofuels technologies using non conventional feedstocks and processes, including cellulosic ethanol, renewable diesel and renewable natural gas, to further reduce the carbon intensity of its biofuels” (Bajpai, 2021). The first phase of Greenfield’s expansion study was completed last year, the second phase is now due to start with engineering, feedstock and environmental studies (https://greenfield. com/news/2019/greenfield-global-takes-next-steps-to-expand-production-at-itsvarennes-biorefinery-following-draft-regulation-on-the-minimum-renewablefuel-volumes-in-quebec/).

2.2.8

Thailand

The Thai government is interested in increasing consumption of ethanol and biodiesel produced locally from cassava. Thailand is producing cassava on a large scale. Attempts are being made to replace the octane-enhancing additive MTBE in gasoline with ethanol. Initially molasses was used for production of ethanol on a large scale but cassava was formally chosen as the major raw material. The price of sugar is very high therefore ethanol is not produced from sugarcane. In Thailand all premium gasoline are being replaced by E10. According to report from the United States Department of Agriculture’s Foreign Agricultural service “Thailand is relying upon domestic ethanol production, the country could get advantage from importing biofuels. Whereas the country allows imports of ethanol for industrial application, the country is not importing ethanol for use as a transportation fuel. Traders of ethanol in Thailand are needed to receive a permit from Ministry of Energy; but, to date, the MOE has never approved any imports of fuel ethanol due to sufficient supplies of locally produced ethanol.” Raw materials for production of ethanol in Thailand are sugar cane, molasses and cassava. Because of dearth of these raw materials, Thailand “will be forced to temporarily lower biofuel use targets or price surges when weather-related feedstock shortages occur, and the country’s lack of ethanol imports will stop it from meeting higher targets of ethanol use. It is also expected to see higher greenhouse gas emissions from land use change. Permitting some role for imports unlocks the full positive potential contribution biofuels can make.” The country is set to reduce it’s consumption goals

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for 2037. A new 20-year Alternative Energy Development Plan has been approved and “the government is in the process of reviewing ethanol and biodiesel consumption targets.” Those targets are expected to be lesser than the targets set out in the 2015 plan, because of the concerns over raw material supply. The new targets are expected to reduce the consumption target in 2037 to 2.4 billion liters which is 41% below the target set out in 2015. In 2019, the ethanol production rates increased, but not as fast as they once had. In 2017, the highest ethanol consumption growth rate at 12% was observed. In 2019, ethanol consumption has increased by 6% over 2018 levels. The reduced rate of ethanol production and use is because of the delay in the cessation of Octane 91 E10 sales. Despite of the delay, the ethanol-blend levels in the country have reached 13.5% in this year due to strong E20 sales (http://www.ethanolproducer.com/articles/16741/report-thailandcould-benefit-from-ethanol-imports).

2.2.9

Japan

Japan is one the major purchaser of motor gasoline in the world and is very much dependent upon imported oil. This country is using fuel ethanol or ETBE on a large-scale and has targeted to improve its energy security and reduce GHG emissions for accomplishing its Kyoto obligations. Blends of ethanol are now being used in some regions. The government has defined a mandate for E3 valid for the entire country and is planning for expansion of this mandate to E10 in next few years. But there is some opposition because of the smaller number of large scale ethanol suppliers and also because of the interests of oil companies preferring gasoline blends with ETBE rather than with fuel ethanol (Piacente 2006). In 2018, consumption of gasoline was 51 billion liters. “The pure ethanol equivalent of ETBE consumed, plus a small amount of ethanol consumed in direct blending, brings total fuel ethanol consumption to 890 million liters, and yields an effective national average blend rate of 1.8%” (gain.fas.usda.gov). “Japan is having one refinery which is producing about 0.2 million liters of bioethanol from rice. No change is expected to this level of production now. The refinery is situated in Niigata Prefecture and is operated by JA Zen-noh, the federation of agricultural cooperatives. It uses high yield rice grown mainly for biofuel production. The ethanol is used as part of an E3 blend, and the E3 gasoline is sold at six affiliated gas stations in the region of Niigata Prefecture. In 2010, Japan Biofuels Supply LLP started producing ETBE domestically. Each year, the company is producing 170 million liters of ETBE, using 72 million liters of ethanol. The company is fully dependent upon imported ethanol. Oil industry of Japan is now allowed to use United States ethanol for producing ETBE for meeting Japan’s biofuels mandate” (https://www.fas.usda.gov/data/japan-biofuels-annual-4).

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2.2.10 India India is one of the largest sugar producer in the world. India and the United States exchanged a memorandum for cooperation on development of biofuels in February 2009. The memorandum covered the production, consumption, distribution and marketing of biofuels in India. Most of the ethanol produced in India is used for industrial consumption. Lately, because of economic and strategic reasons, Indian government has seriously considered production of fuel ethanol and a mandate for E10 blends is presently effective in thirteen states. It is expected that in the near future, the mandate for E10 shall be effective in the entire country. The Indian Institute of Petroleum had performed studies using a 10% ethanol blend in gasoline and 15% ethanol in biodiesel. “In the coming years to come, the enormous rise in demand for biofuels is surely going to replace the liquid fuels which are already in use with ethanol which presently supplies over 95% of biofuels for transportation (Fulton et al., 2004). Currently, the most efficient ethanol production is based on dedicated energy crops like maize and sugarcane. At the same time, these dedicated ethanol crops may have greater effect on future food supply and demand (Msangi et al., 2007). First-generation Biofuels meet only up to 30% requirement of our country while the balance of almost 146 metric tonnes is met by import from other leading producers (Patni et al., 2011). India accounts for approximately 4% in production of bioethanol worldwide which is basically sugarcane derived. India started a mega biodiesel program based on Jatropha, amongst all other alternatives, which introduces a blend that contains 5% of biodiesel with fixed prices (Dufey, 2005). Biomass became the largest source of renewable energy in 2008 generating around 50 exajoules (EJ) (1200 million tonnes of oil equivalent) of bioenergy globally, which accounted for a 10% share of the total primary energy demand in the same period (Oyakhire and Mohammed, 2012). India registered an 85% increase in overall ethanol production in 2009. India holds only 0.3% share of biofuel production in 2010 globally. This includes 380 million liters of fuel ethanol and 45 million liters of biodiesel. It is worth noticing that India is the second largest sugarcane producer globally but holds only about 1% share of ethanol production globally. This can be attributed to the fact that 70% 80% of the cane produced in the country is utilized for production of sugar and the remaining 20% 30% for alternate sweeteners like Khandsari and Jaggery (Patni et al., 2011). Estimates indicate that India’s biofuel requirement of 0.5 billion gallons in 2012 will increase to 6.8 billion gallons by 2022 (Patni et al., 2011). Most of the biomass used presently is obtained from three main sources: forests, wastes and agriculture. This includes virgin wood from the conventional tree cutting, wood residues from wood processing industries and saw mills, agricultural energy crops, agricultural residues and wastes (Oyakhire and Mohammed, 2012). About 80% of ethanol (worth $173 million) in 2016 was imported from

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United States and was mostly classified as undenatured fuel at port of origin. Incidentally, 2016 import volume was the largest since 2009 (278 million liters) and almost double the volume of ethanol imported in 2015. National Biodiesel Mission (NBM) identified Jatropha (Jatropha curcas) as the most appropriate inedible oilseed for biodiesel production in order to achieve a proposed biodiesel blend of 20% with conventional diesel by 2017. That target was unachieved because of economic and agronomical constraints. By 2022, the Government of India intends to reduce its import of crude oil by 10% by several means such as increasing domestic output, promoting energy conservation and efficiency and also encouraging the use of other alternate fuels. Growth in the biofuel market will partly reduce import dependency of crude oils and encourage optimal use of other renewable energy resources, particularly when strong economic growth prospects drive higher demand for gasoline and petroleum products. Projected world primary energy demand is expected to rise to 600 1000 EJ in 2050 and various other scenarios indicate that the future bioenergy requirement of the country could reach up to 250 EJ/year, representing almost a quarter of the future global energy mix” (Banerjee et al., 2019). India has been promoting second generation bioethanol for achieving its E20 target. It can help meeting the objective of converting waste into energy and also reduce agricultural waste incineration. The second generation plant will play an important role in making ethanol available for blending. By 2025, ethanol production in India will increase three times to approximately 10 billion liters per year, according to Tarun Kapoor, secretary, Union Ministry of Petroleum and Natural Gas. The refineries plan to construct second generation bioethanol plants, according to Kushal Banerjee, chief general manager, Hindustan Petroleum Corporation Ltd (HPCL). HPCL has planned to set up four second generation ethanol plants which will convert agricultural waste into biofuel. This will reduce toxic air pollution in northern India. Furthermore, HPCL will build four plants to produce ethanol from surplus maize, rice and damaged grain. The first second generation ethanol biorefinery is being set up at Bathinda, Punjab. HPCL has made investment in Bangalore-based ASN Fuels for the establishment of a second generation Ethanol Pilot Plant at Tirupati in collaboration with IIT-Tirupati. DM Naveen Giri, CEO, ASN Fuels Pvt Ltd reported that they are working on production of ethanol from agricultural waste on a pilot scale which produces 10 L per day. They are now planning to scale up to 2000 L per day mid-scale plant. Intermittent thermal equalizers have been provided in the pilot plant for maintaining the desired pipeline temperature and to scrub the reacted carbon dioxide which can be bottled and sold (https://www.downtoearth.org.in/blog/energy/second-generation-bioethanol-it-is-time-to-launch-itheadlong-78507). Biofuel key figures are summarized in Table 2.11.

TABLE 2.11 Biofuel key figures. United States Leader in global biofuel production (2019) No. 1 in ethanol production (46% of global production) and No. 2 in biodiesel production (19%). 87% of biofuel production is bioethanol (mostly based on corn). Strong decline in bioethanol production in 2020 due to transport restrictions, limited decline for bioethanol (anti dumping measures enacted against Argentina and Indonesia). Expected recovery to its pre-crisis level by 2022, with slowing domestic petrol demand offset by higher exports to Canada, Brazil or India. Brazil No. 2 in ethanol production (28%, mainly based on sugar cane) and #4 in biodiesel production (12%, mainly based on soya). Expected production growth by 2029, thanks to the RenovaBio plan (since June 2020) that forces fuel suppliers to buy decarbonization certificates (CBIOs) from biofuel producers. European Union No. 1 in biodiesel production (over 30% in 2019), and No. 4 in ethanol production (5%). Still dependent on raw material imports (60% dependence) and on biofuel imports (from Argentina, Indonesia, Malaysia and China for biodiesel and from Ukraine, Brazil and the United States for ethanol). Expected decline in biodiesel consumption by 2029 (decrease in diesel vehicle sales, improved fuel efficiency and the competition from electric vehicles). RED II directive promotes advanced biofuels, which should impact the biofuel production and consumption structure in the EU. Asia China is the world’s 3rd largest producer of ethanol (8%) and 8th of biodiesel and HVH (2%). Lower demand in 2020 offset by the extension of ethanol blending obligations to new provinces (decided in 2017 to sell out corn stocks) The focus on electric mobility could contribute to slowing down the rise in ethanol production by 2029. Indonesia is the 3rd largest producer of biodiesel and HVH in 2019, but only the 21st ethanol producer. Biodiesel production rise in 2020 due to strengthened biodiesel blending obligation (from 20% to 30%) despite the 12% fall in diesel consumption Expected surge in biodiesel consumption by 2029 and fall in exports (EU ban on palm oil-based biodiesel): Indonesia may even become a net biodiesel importer in the coming decade to meet rising demand. Africa 5th largest ethanol producer despite a marginal production (and no biodiesel production) and net ethanol exporter. Some countries such as South Africa have introduced support measures but the share of biofuels in road transport in Africa should only average 1.5% 2.5% by 2040. Source: Reproduced with kind permission https://www.enerdata.net/publications/executivebriefing/biofuels-market-dynamics.html.

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References Amorim, H.V., Basso, L.C., Lopes, M.L., 2009. Sugar cane juice and molasses, beet molasses andsweet sorghum: composition and usage, The Alcohol Textbook, fifth ed. Nottingham University Press, pp. 39 46. Bajpai, P., 2021. Global production of bioethanol. Developments in Bioethanol. Green Energy and Technology. Springer, Singapore. Available from: https://doi.org/10.1007/978-981-158779-5_10. Banerjee, S., Kaushik, S., Tomar, R.S., 2019. Global scenario of biofuel production: past, present and future. In: Rastegari, A., Yadav, A., Gupta, A. (Eds.), Prospects of Renewable Bioprocessing in Future Energy Systems. Biofuel and Biorefinery Technologies, Vol 10. Springer, Cham. Available from: https://doi.org/10.1007/978-3-030-14463-0_18. Basso, L.C., Rosa, C.A., 2010. Sugar cane for potable and fuel ethanol. Proceedings of the Worldwide Distilled Spirits Conference 2008. Edinburgh. Nottingham University Press. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294 306. Dufey, A., 2005. International trade in biofuels: some preliminary data, presentation for the WWF Workshop “reaching a common position on biofuels”. Fulton, L., Howes, T., Hardy, J., 2004. Biofuels for transport, an international perspective, IEA study undertaken by the office of energy efficiency. Technol. R. D. 210. Paris. Fulton, L., 2006. Biodiesel: technology perspectives. In: Geneva UNCTAD Conference. Guo, M., 2020. The global scenario of biofuel production and development. In: Mitra, M., Nagchaudhuri, A. (Eds.), Practices and Perspectives in Sustainable Bioenergy. Green Energy and Technology. Springer, New Delhi. Available from: https://doi.org/10.1007/978-81-3223965-9_3. McGlashen, A., 2013. As key partner departs, future dims for Michigan cellulosic biofuel plant. https://energynews.us/2013/08/06/midwest/as-key-partnerdeparts-future-dims-for-michigancellulosic-biofuel-plant/ Msangi, S., Sulser, T., Rosegrant, M., Rowena, V., Claudia, R., 2007. Global scenarios for biofuels: impacts and implications for food security and water use. Biofuels Glob. Food Bal. 1 20. Nguyen, Q., Bowyer, J., Howe, J., Bratkovich, S., Groot, H., Pepke, E., et al., 2017. Global production of second generation biofuels: trends and influences. https://dovetailinc.org/upload/ tmp/1579558792.pdf Oak Ridge National Laboratory, 2005. Biofuels Switchgrass: Greener Energy Pastures. Oyakhire, O., Mohammed, M., 2012. Biofuels. Technical research report. Inst. Gas. Eng. Managers. Padella, M., O’Connell, A., Prussi, M., 2019. What is still limiting the deployment of cellulosic ethanol? Analysis of the current status of the sector. Appl. Sci. 9, 4523. Patni, N., Pillai, S.G., Dwivedi, A.H., 2011. Analysis of current scenario of biofuels in India specifically bio-diesel and bio-ethanol. International Conference on Current Trends in Technology. Nuicone, pp. 1 4. Piacente, E.A., 2006. Perspectives for Brazil in the Bio-Ethanol Market (MSc dissertation). State University of Campinas—Unicamp, Campinas. POET-DSM: Project Liberty, 2018. https://www.energy.gov/eere/bioenergy/poet-dsm-project-liberty, https://www.forbes.com/sites/rrapier/2018/02/11/cellulosicethanol-falling-far-short-ofthehype/#484c7df4505f Puri, M., Abraham, R.E., Barrow, C.J., 2012. Biofuel production: prospects, challenges and feedstock in Australia. Renew. Sustain. Energy Rev. 16 (8), 6022 6031. Elsevier.

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Soccol, C.R., Vandenberghe, L.P., Medeiros, A.B., Karp, S.G., Buckeridge, M., Ramos, L.P., et al., 2010. Bioethanol from lignocelluloses: status and perspectives in Brazil. Bioresour. Technol. Jul. 101 (13), 4820 4825. United Nations Food and Agriculture Organization, 2008. State Food Agriculture. Voegele, E., 2014. Bluefire announces EPC contract for mississippi project. http://ethanolproducer.com/articles/11521/bluefire-announcesepc-contract-for-mississippi-project Voegele, E., 2018. Verbio to buy DuPont cellulosic ethanol plant, convert it to RNG. http://biomassmagazine.com/articles/15743/verbio-to-buy-dupontcellulosic-ethanolplantconvert-it-to-rng Voorhis, D., 2016. Hugoton cellulosic ethanol plant sold out of bankruptcy. https://www.kansas. com/news/business/article119902263.html Yan, X., Inderwildi, O.R., King, D.A., 2010. Biofuels and synthetic fuels in the US and China: a review of well-to-well energy use and greenhouse gas emissions with the impact of land-use change. Energy Env. Sci. 3, 190 197. Zhang, C., 2019. Lignocellulosic ethanol: technology and economics. In: Yun, Y. (Ed.), Alcohol Fuels Current Technologies and Future Prospect. IntechOpen Limited, London. Available from: https://doi.org/10.5772/ intechopen.86701. Zhenhong, Y., 2006. Bio-fuels industry in China: utilization of ethanol and biodiesel in today and future. World Biofuels Symposium Beijing, China.

Relevant websites http://ethanolproducer.com/articles/15673/iea-predicts-growth-in-globalethanol-productionthrough-2023 http://ethanolproducer.com/articles/16422/reporteu-ethanol-consumption-to-increase-in-2019 http://www.ethanolproducer.com/articles/16741/report-thailandcould-benefit-from-ethanolimports https://apps.fas.usda.gov/newgainapi/api/report/downloadreportbyfilename?filename 5 Biofuels% 20Annual_Beijing_China%20-%20Peoples%20Republic%20of_8-9-2019.pdf https://arena.gov.au/assets/2019/11/biofuels-and-transport-an-australian-opportunity.pdf https://css.umich.edu/sites/default/files/Biofuels_CSS08-09_e2021_0.pdf https://ethanolrfa.org/markets-and-statistics/annual-ethanol-production https://noaw2020.eu/wp-content/uploads/2018/12/C1-1-COFCO-Wuguoqing.pdf https://www.downtoearth.org.in/blog/energy/second-generation-bioethanol-it-is-time-to-launchitheadlong-78507 https://www.eia.gov/biofuels/biodiesel/capacity/ https://www.eia.gov/petroleum/ethanolcapacity/ https://www.enerdata.net/publications/executive-briefing/biofuels-market-dynamics.html https://greenfield.com/news/2019/greenfield-global-takes-next-steps-to-expand-production-at-itsvarennes-biorefinery-following-draft-regulation-on-the-minimum-renewable-fuel-volumesin-quebec/ https://www.etipbioenergy.eu/value-chains/conversion-technologies/advanced-technologies/ sugar-to-alcohols/commercial-cellulosic-ethanol-plants-in-brazil https://www.fas.usda.gov/data/brazil-biofuels-annual-5 https://www.fas.usda.gov/data/japan-biofuels-annual-4 https://www.fas.usda.gov/regions/canada https://www.greencarcongress.com/2019/08/20190812-fas.html https://www.reuters.com/business/energy/brazils-razen-build-second-cellulosic-ethanol-plant-filing-2021-06-25/

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https://www.statista.com/statistics/274163/global-biofuel-production-in-oil-equivalent/; http://www. cefc.com.au https://www.nsenergybusiness.com/features/top-biofuel-production-countries/ http://www.forbes.com ethanolproducer.com gain.fas.usda.gov

Chapter 3

Generations of biofuels 3.1

Introduction

Biofuels are energy sources produced from biomass. Biofuels have been around for a long time, but coal and petroleum have been used mainly as energy sources because these are abundantly available, low priced and have a higher energy value. Fossil fuels such as petroleum and coal also come from biomass but they took millions of years to produce. Biofuels are making a re-emergence because of the increase in oil prices, diminishing fossil fuel reserves, the need to have a renewable, dependable source of energy and as a way to alleviate the effects of climate change (Olguin-Maciel et al., 2020). Biofuels are a renewable resource because they can grow again or never run out. In contrast, fossil fuels are nonrenewable as they require millions of years to form (https://agsci.oregonstate.edu/sites/agsci.oregonstate. edu/files/bioenergy/generations-of-biofuels-v1.3.pdf). There are four types of biofuels: first, second, third and fourth generation biofuels (Fig. 3.1; Khan et al., 2021). The characteristics of different generations of biofuels are presented in Fig. 3.2 (Sikarwar et al., 2017). Different types of feedstocks used in the first and second generation biorefinery for producing biofuels, biochemicals, food, and feed are shown in Fig. 3.3. First generation biofuels are also known as conventional biofuels. The feedstock for first generation biofuels are food crops. The sugars, starches and oils in sugarcane, corn, soy and palm are readily accessible. So, conversion of these feedstocks into biofuels simply involves either fermentation of the sugars or chopping up the fatty oils through transesterfication. Use of edible biomass increases the production cost and causes wasteful use of resources and energy spent in growing the crops. Use of edible biomass competes with food crops, needs large amounts of fertilizer and water, and big areas of crop- land (Rulli et al., 2016; Hayes et al., 2015). The second generation of biofuels are produced from inedible lignocellulosic biomass (Bhatia et al., 2017). Types of raw feedstock for secondgeneration biofuel are shown in Fig. 3.4 (Ganguly et al., 2021). Whereas this generation beats the shortcomings of the first generation, additional steps are needed for producing sufficient biofuels at a competitive price (Hayes et al., 2015). In the past few years, many studies have been conducted to attain this objective by using thermal, chemical, biological or enzymatic processes. Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00009-1 © 2023 Elsevier Inc. All rights reserved.

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FIGURE 3.1 Biofuels generations (Khan et al., 2021; Vasi´c et al., 2021). Khan, N., Sudhakar, K., Mamat, R., 2021. Role of biofuels in energy transition, green economy and carbon neutrality. ˇ Leitgeb, M., 2021. Bioethanol production by enzy´ K., Knez, Z., Sustainability 13, 12374; Vasic, matic hydrolysis from different lignocellulosic sources. Molecules 26 (3), 753. Distributed under the terms of the Creative Commons Attribution 4.0 International License.

Problems have been faced with all these conversion methods. But, chemical processes are found to be the most flexible (Faba et al., 2012). Combinations of several processes have also been examined for instance the production of

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FIGURE 3.2 Different generations of biofuels with their characteristics. Sikarwar, V.S., Zhao, M., Fennell, P.S., Shah, N., Anthony, E.J., 2017. Progress in biofuel production from gasification. Prog. Energy Combust. Sci. 61, 189 248. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

FIGURE 3.3 Different feedstocks used in the first and second generation biorefinery for producing biofuels, biochemicals, food, and feed (Balan, 2014). Balan, V., 2014. Current challenges in commercially producing biofuels from lignocellulosic biomass. 2014, Article ID 463074, 31p. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

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FIGURE 3.4 First generation biofuels. Khan, N., Sudhakar, K., Mamat, R., 2021. Role of biofuels in energy transition, green economy and carbon neutrality. Sustainability 13, 12374. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

sugars from biomass using a chemical method followed by biological or an enzymatic process (Faba et al., 2012). Third generation biofuels appear to be the best possibility for alternative fuel as they do not compete with food. However, there are still some challenges to overcome in making them commercially viable. Algae such as seaweed are photosynthetic plants; capture higher quantities of carbon dioxide and produce oxygen and also oil (Hayes et al., 2015). But, this type of biomass has some drawbacks. The cost is high and also the fact that biofuel produced from algae is less than that produced from other feedstocks. The major cause for this is that the oil produced by algae is very much unsaturated; it is more volatile particularly at higher temperatures, and so is degraded (Shah et al., 2018). Fourth generation technology combines genetically optimized feedstocks, designed to capture huge amounts of carbon, with genetically modified microorganisms, which are able to produce fuels efficiently. Key to the process is the capture and sequestration of carbon dioxide. Carbon sequestration is the process of capturing and storing atmospheric carbon dioxide. It is one method of reducing the amount of carbon dioxide in the atmosphere with an objective of reducing global climate change. This process makes fourth generation biofuels a carbon negative source of fuel. But, the weakness is carbon capture and sequestration technology, which continues to elude the coal industry.

3.2

First generation biofuels

First generation biofuels are also known as conventional biofuels. These are made from materials rich in sugar (sugarcane, sugarbeet, sweet sorghum), starch (corn and cassava) or oils (soybeans, rapeseed, coconut, sunflowers, and palms). These biofuels are produced through widely-understood technologies and processes, such

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FIGURE 3.5 Types of raw feedstock for second-generation biofuel. Reproduced with permission from Ganguly, P., Sarkhel, R., Das, P., 2021. The second- and third-generation biofuel technologies: comparative perspectives (Chapter 2). In: Dutta, S., Hussain, C.M. (Eds.), Sustainable Fuel Technologies Handbook, Academic Press, pp. 29 50.

as fermentation, distillation and transesterification (Fig. 3.5; Khan et al., 2021). These processes have been used for many years in several uses, such as producing alcohol. Fermentation of sugars and starches produces mainly ethanol. Butanol and propanol are produced in lesser quantities. Ethanol has one-third of the energy density of gasoline, but is presently used in several countries, as a gasoline additive. United States and Brazil have led the industrial production of ethanol fuel for many years. These countries together accounted for 85% of the world’s production in 2017. An advantage of ethanol is that it burns cleaner than gasoline and thus produces smaller amount of greenhouse gases. Another first generation biofuel, called biodiesel, is produced from vegetable oils, yellow grease, used cooking oils, or animal fats. The fuel is produced by transesterification. In this process fat or oil reacts with an alcohol to produce esters and glycerol. A catalyst is used for improving the rate of reaction and yield. As the reaction is reversible, excess alcohol is used for shifting the equilibrium towards the product side. Distillation is used to separate the main product from any of the by-products of the reactions. In place of petroleum diesel, biodiesel can be used in several diesel engines or in a mixture of the two. “Common first-generation biofuels include ethanol, other bioalcohols, biodiesel, bioethers, biogas, syngas etc. Ethanol represents the most common biofuel produced to date. Nowadays, 78% of biofuel’s total production contributed by bioethanol produced around 28 billion gallons per year from central corn in the

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United States and sugarcane in Brazil (Jambo et al., 2016; Boboescu et al., 2019). However, this generation’s biofuels increased production by raising questions due to its production-generating competition food production vs. fuels, arable lands, and biodiversity loss, in addition to being responsible for ecological degradation (Abdullah et al., 2019; Chowdhury and Loganathan, 2019). Studies have shown that biofuels obtained in this manner frequently do not contribute to greenhouse gas reduction, and they require a large amount of energy for their production” (Olguin-Maciel et al., 2020; Stolarski et al., 2015). Sugarcane (Brazil), molasses and corn (USA) are the main fuel crops used, as well as starchy materials such as sugarbeet, wheat, barley and rye. Crops containing starch have to be converted to sugars first. Oil seed rape is a very effective crop for producing biodiesel. Nevertheless, first generation biofuels have numerous associated problems. There is much debate over their actual advantage in reducing greenhouse gases and carbon dioxide emissions because of the fact that some biofuels can produce negative net energy gains, release more carbon in their production than their feedstock’s capture in their growth (http://energyfromwasteandwood.weebly.com/generations-of-biofuels.html). But, the most controversal issue with first generation biofuels is “fuel versus food.” As most of biofuels are produced directly from food crops, the increase in requirement for biofuels has lead to an increase in the volumes of crops being diverted away from the global food market. This has been held responsible for the worldwide increase in food prices over the past few years.

3.3

Second generation biofuels

Second Generation biofuels address several issues related to first generation biofuels. They are produced from non-food crops such as wood, organic waste, food crop waste and specific biomass crops such as perennial grass and fast-growing trees so eliminate the major issues with first generation biofuels (UN Report, 2007). Second Generation biofuels are also aimed at being more cost-efficient in relation to existing fossil fuels (http://energyfromwasteandwood.weebly.com/generations-of-biofuels.html). Life cycle assessments of second-generation biofuels have also shown that they will increase “net energy gains” overcoming another of the major limitations of first generation biofuels. Fig. 3.6 shows biochemical conversions of second-generation feedstocks to biofuels (Patinvoh and Taherzadeh, 2019). “Second generation biofuels do not compete between fuels and food crops as they come from distinct biomass. Second generation biofuels also produce higher energy yields per acre than first generation fuels. They allow for use of poor quality land where food crops may not be able to grow. The technology is quite immature, so it still has potential of cost reductions and increased production efficiency as scientific developments occur. But, some

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FIGURE 3.6 Biochemical conversions of second-generation feedstocks to biofuels. Reproduced with permission from Patinvoh, R.J., Taherzadeh, M.J., 2019. Chapter 9 Fermentation processes for second-generation biofuels. In: Basile, A., Dalena, F. (Eds.), Second and Third Generation of Feedstocks, Elsevier, pp. 241 272.

biomasses for second-generation biofuels still compete with land use since some of the biomass grows in the same climate as food crops. This leaves farmers and policy makers with the hard decision of which crop to grow. Cellulosic sources which grow alongside food crops could be used for biomass, such as corn stover (leaves, stalk, and stem of corn). However, this would take away too many nutrients from the soil and would need to be replenished through fertilizer. Furthermore, the process to produce second generation fuels is more complex than first generation biofuels as it needs pretreating the biomass to release the trapped sugars. This needs more energy and materials” (Datta et al., 2019). Technical barriers remain for second generation of biofuels (Sheldon, 2018). “The use of waste plant biomass has attracted researchers for a wide variety of uses such as feedstock to produce heat and electricity by direct burning (Naik et al., 2010; Nygaard et al., 2016) or as a raw material for wastewater treatments (Alalwan et al., 2018). But, using it as a low cost source of biofuel is very attractive (Westensee et al., 2018). A wide variety of discarded materials can be used as biofuel feedstock such as agriculture waste, poplar trees, willow and eucalyptus, miscanthus, switchgrass, reed canary grass, and wood and they mostly consist of plant cell walls whose primary components is polysaccharides (75%) (Westensee et al., 2018; LeBauer et al., 2018). These polysaccharides have a high sugar content which is preferred for biofuel production. But, agricultural by-products can provide only a limited proportion of the increased demand for biofuels” (Alalwan et al., 2019; Naik et al., 2010).

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“Second-generation biofuels address many of the issues related to firstgeneration biofuels. The prospects for fostering regional growth and improving the economic situation in developing regions is envisioned with secondgeneration biofuels. Around the world, several strategies for the production of second-generation biofuels are being considered. Still, the focus is mainly on two distinct paths, either the thermo or bio route generated by biomass of cellulose and lignin, tree surplus, and seasonal forage crop. Thermochemical manufacturing has the noteworthy advantage of higher flexibility of feedstock than biological production. The thermo route is focused on the heat processing of biomass under reduced oxidizing agent concentrations. Under the temperature range of 300 C 1000 C, the bottom range will mainly produce solid biofuel called biochar. At high temperatures, pyrolytic oil and syngas are the most concentrated substances in the middle range. The bio approach includes the pretreatment of lignocellulosic material, enzymatic hydrolysis, and the fermentation of sugars by specific strains of microorganism. It is more challenging to convert lignocellulose to reducing sugars than it is to convert starch. Biological, physical (thermal), or chemical catalysts are used for pretreating biomass in the biochemical pathway; therefore, improved advances in the development of second generation biofuels are hampered because of the chemical and structural properties of the extracellular matrix” (Khan et al., 2021). Table 3.1 shows yield of biofuels from different feedstocks of secondgeneration biofuels. TABLE 3.1 Yield of biofuels from different feedstocks of second-generation biofuels. Biofuels

Feedstock

Yield

Bioethanol

Sugarcane bagasse

165 g/kg

Fermentation, Acid hydrolysis (Sulfuric acid), Kluyveromyces sp. IIPE453, Fermentation at 50 C

Biodiesel

Palm oil

97 w/w

Transesterification, Sulfuric acid—5% v/w, 95 C/540 min

Biobutanol

Rice straw

5.52 g/L

Fermentation, C. sporogenes BE01, (37 C and 6.7 pH)

Biomethane

Corn Stover

135 dm3/kg VS

Anaerobic digestion, Cellulase (Spezyme CP) T 5 37 C 6 1 C, Time 5 30 days

Syngas

Corn Stover

H2: 26.9, CO: 24.7 CO2: 23.7 CH4: 15.3

Gasification, Fluidized bed gasifier GA: steam; T: 600 C 710 C

Based on Nolasco-Hipolito et al. (2001), Behera et al. (2015), Kovacic et al. (2017), Gottumukkala et al. (2013), Carpenter et al. (2010).

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Third generation biofuels

The Third Generation of biofuels take benefit of specially engineered energy crops such as algae as its energy source (Chisti, 2007). Algae are cultured to act as an inexpensivet, high-energy and totally renewable raw material. It is envisaged that algae has the potential to produce more energy per acre than conventional crops. Algae can also be grown using land and water not suitable for food production (https://agsci.oregonstate.edu/sites/agsci.oregonstate.edu/files/bioenergy/generations-of-biofuels-v1.3.pdf). The Third Generation of biofuels are more energy dense than first and second generation biofuels per area of harvest. Algae are beneficial in that it is able to grow in areas not suitable for first and second generation crops, which would alleviate pressure on water and arable land used. It can be grown using sewage, wastewater, saltwater, for instance oceans or salt lakes. Due to this, there would not be a requirement to use water that would otherwise be utilized for human consumption. But more research still needs to be done to improve the extraction process for making it financially competitive to petrodiesel and other petroleum-based fuels. Algae produces more energy per acre than conventional crops. Algae can also be grown using land and water not suitable for food production, so reducing the pressure on already exhausted water sources. Another advantage of algae based biofuels is that wide range of fuels like diesel, petrol and jet fuel can be produced. According to the IEA definition, third-generation biofuels are bio-based fuels produced from aquatic feedstock (usually algae) (Saladini et al., 2016). Algae are a promising alternative feedstock because of their high lipid and carbohydrate contents and better carbon dioxide absorption. Algae can be grown on wastewater and seawater. Unproductive dry lands and marginal farm lands do not compete with food crops on arable land or in freshwater environments (Saladini et al., 2016; Jambo et al., 2016). Algae is interesting for biofuels because it contains low level of lignin; has a higher growth rates and can produce biodiesel, butanol, methane and ethanol. Some green algal species can photolyze mediated biohydrogen production (Saladini et al., 2016; Jambo et al., 2016; Chowdhury and Loganathan, 2019; Datta et al., 2019; Khan and Fu, 2020). But, this type of biomass has drawbacks such as its higher initial investment for its production. The biofuel produced from algae is less stable than that produced from other sources. Furthermore, the high water quantity is also an issue when lipids have to be extracted from the algal biomass, which requires dewatering via either centrifugation or filtration before extraction of the lipids (Lee and Lavoie, 2013). Fig. 3.7 shows biofuel generation from microalgae (Khan et al., 2021).

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FIGURE 3.7 Biofuel generation from microalgae. Khan, N., Sudhakar, K., Mamat, R., 2021. Role of biofuels in energy transition, green economy and carbon neutrality. Sustainability 13, 12374. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

3.5

Fourth generation biofuels

Fourth generation biofuels are still in the developmental stage. These biofuels are produced using bioengineered microorganisms as microalgae, yeast, fungi, and cyanobacteria. Genetically modified crops consume more carbon dioxide from the environment than they release. These microorganisms are used to produce different fuels, including ethanol, butanol, hydrogen, methane, vegetable oil, biodiesel, isoprene, gasoline, and jet fuel (Alalwan et al., 2019). The research on fourth-generation biofuel started in 2006, and significant results have not been published yet (Datta et al., 2019). Fourth generation biofuels not only produce sustainable energy but also capture and store carbon dioxide. Biomass materials, which absorb carbon dioxide while growing, are converted into fuel using the same processes as second generation biofuels. This process differs from second and third generation biofuel production as at all stages of production the carbon dioxide is captured using processes such as oxy-fuel combustion (Schmetz et al., 2007). The carbon dioxide is geosequestered by storing it in old oil and gas fields or saline aquifers. This carbon capture makes fourth generation biofuel production carbon negative rather than simply carbon neutral, as it takes away more carbon than it produces. This system not only captures and stores carbon dioxide from the atmosphere but it also reduces carbon dioxide emissions by replacing the fossil fuels.

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FIGURE 3.8 Fourth generation biofuel production. Khan, N., Sudhakar, K., Mamat, R., 2021. Role of biofuels in energy transition, green economy and carbon neutrality. Sustainability 13, 12374. This Figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

“In fourth-generation biofuels, genetically modified microorganisms such as microalgae, yeast, fungi and cyanobacteria are utilized as sources. The ability of microorganisms to convert carbon dioxide to fuel through photosynthesis is utilized (Vassilev and Vassileva, 2016). The multiple advantages of microalgae such as their high growth rate and oil content and low structural complexity enhance their numerous commercial applications (Azizi et al., 2018). In addition to genetic modification, some fourth-generation technologies involve pyrolysis (in a temperature range between 400 C and 600 C) (Azizi et al., 2018), gasification, upgrading, and solar-to-fuel, pathways (Cuellar-Bermudez et al., 2015). The general purpose of these modifications is to improve the HC yield and create an artificial carbon sink to eliminate or minimize carbon emission. These technologies are still in early developmental stages” (Alalwan et al., 2019; Sikarwar et al., 2017). Fig. 3.8 shows fourth generation biofuel production (Khan et al., 2021). Fig. 3.9 represents a schematic diagram of the bioethanol produced by different generations of biofuels (Alalwan et al., 2019).

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FIGURE 3.9 A schematic diagram of bioethanol production based on different generations (Alalwan et al., 2019). Reproduced with permission.

References Abdullah, B., Muhammad, S.A.F.S., Shokravi, Z., Ismail, S., Kassim, K.A., Mahmood, A.N., et al., 2019. Fourth generation biofuel: a review on risks and mitigation strategies. Renew. Sustain. Energy Rev. 107, 37 50. Alalwan, H.A., Abbas, M.N., Abudi, Z.N., Alminshid, A.H., 2018. Adsorption of thallium ion (Tl 1 3) from aqueous solutions by rice husk in a fixed-bed column: experiment and prediction of breakthrough curves. Environ. Technol. Innov. 12, 1 13. November. Alalwan, H.A., Alminshid, A.H., Aljaafari, H., 2019. Promising evolution of biofuel generations. Subject review. Renew. Energy Focus. 28, 127 139. Azizi, K., Moraveji, M.K., Najafabadi, H.A., 2018. A review on bio-fuel production from microalgal biomass by using pyrolysis method. Renew. Sust. Energy Rev. 82 (P3), 3046 3059. Elsevier. Balan, V., 2014. Current Challenges in Commercially Producing Biofuels from Lignocellulosic Biomass 2014. Available from: https://doi.org/10.1155/2014/463074. Article ID 463074, 31p. Behera, S., Singh, R., Arora, R., Sharma, N.K., Shukla, M., Kumar, S., 2015. Scope of algae as third generation biofuels. Front. Bioeng. Biotechnol. 90. Bhatia, S.K., Kim, S.H., Yoon, J.J., Yang, Y.H., 2017. Current status and strategies for second generation biofuel production using microbial systems. Energy Convers. Manage. 148, 1142 1156. Boboescu, I.Z., Chemarin, F., Beigbeder, J.B., De Vasconcelos, B.R., Munirathinam, R., Ghislain, T., et al., 2019. Making next-generation biofuels and biocommodities a feasible reality. Curr. Opin. Green Sustain. Chem 20, 25 32.

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Carpenter, D.L., Bain, R.L., Davis, R.E., Dutta, A., Feik, C.J., Gaston, K.R., 2010. Pilot-scale gasification of corn stover, switch grass, wheat straw, and wood: 1. Parametric study and comparison with literature. Ind. Eng. Chem. Res. 49, 1859 1871. Chisti, Y., 2007. Biodiesel from microalgae. Biotechnol. Adv. 25, 294 306. Chowdhury, H., Loganathan, B., 2019. Third-generation biofuels from microalgae: a review. Curr. Opin. Green. Sustain. Chem. 20, 39 44. Cuellar-Bermudez, S.P., Garcia-Perez, J.S., Rittmann, B.E., Parra-Saldivar, R., 2015. Photosynthetic bioenergy utilizing CO2: an approach on flue gases utilization for third generation biofuels. J. Clean. Prod. 98, 53 65. Datta, A., Hossain, A., Roy, S., 2019. An overview on biofuels and their advantages and disadvantages. Asian J. Chem. 31, 1851 1858. Faba, L., Dıaz, E., Ordonez, S., 2012. Aqueous-phase furfural-acetone aldol condensation over basic mixed oxides. Appl. Catal. B 113, 201 211. Ganguly, P., Sarkhel, R., Das, P., 2021. The second- and third-generation biofuel technologies: comparative perspectives (Chapter 2). In: Dutta, S., Hussain, C.M. (Eds.), Sustainable Fuel Technologies Handbook. Academic Press, pp. 29 50. Gottumukkala, L.D., Parameswaran, B., Valappil, S.K., Mathiyazhakan, K., Pandey, A., Sukumaran, R.K., 2013. Biobutanol production from rice straw by a non acetone producing Clostridium sporogenes. Bioresour. Technol. 145, 182 BE187. Hayes, C.J., Burgess Jr, D.R., Manion, J.A., 2015. Combustion pathways of biofuel model compounds: a review of recent research and current challenges pertaining to first-, second-, and third-generation biofuels. In: Williams, I.H., Williams, N.H. (Eds.), Advances in Physical Organic Chemistry, Vol. 49. Elsevier, pp. 103 187. Jambo, S.A., Abdulla, R., Azhar, S.H.M., Marbawi, H., Gansau, J.A., Ravindra, P., 2016. A review on third generation bioethanol feedstock. Renew. Sustain. Energy Rev. 65, 756 769. Khan, S., Fu, P., 2020. Biotechnological perspectives on algae: a viable option for next generation biofuels. Curr. Opin. Biotechnol. 62, 146 152. Khan, N., Sudhakar, K., Mamat, R., 2021. Role of biofuels in energy transition, green economy and carbon neutrality. Sustainability 13, 12374. Available from: https://doi.org/10.3390/ su132212374. Kovacic, D., Kralik, D., Rupcic, S., Jovicic, D., Spajic, R., Tisma, M., 2017. Soybean straw, corn stover and sunflower stalk as possible substrates for biogas production in croatia: a review. Chem. Biochem. Eng. Q. 31 (3), 187 198. LeBauer, D., Kooper, R., Mulrooney, P., Rohde, S., Wang, D., Long, S.P., et al., 2018. BETYdb: a yield, trait, and ecosystem service database applied to second-generation bioenergy feedstock production. GCB Bioenergy 10, 61 71. Lee, R.A., Lavoie, J.M., 2013. From first to third-generation biofuels: challenges of producing a commodity from a biomass of increasing complexity. Anim. Front. 3, 6 11. Naik, S.N., Goud, V.V., Rout, P.K., Dalai, A.K., 2010. Production of first and second generation biofuels: a comprehensive review. Renew. Sustain. Energy Rev. 14 (2), 578 597. Nolasco-Hipolito, C., Kobayashi, G., Sonomoto, K., Ishizaki, A., 2001. Biodiesel production from crude palm oil and evaluation of butanol extraction and fuel properties. Process. Biochem. 37, 65 71. Nygaard, I., Dembele´, F., Daou, I., Mariko, A., Kamissoko, F., Coulibaly, N., et al., 2016. Lignocellulosic residues for production of electricity, biogas or second generation biofuel: a case study of technical and sustainable potential of rice straw in Mali Renew. Sustain. Energy Rev. 61, 202 212.

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Olguin-Maciel, E., Singh, A., Chable-Villacis, R., Tapia-Tussell, R., Ruiz, H.A., 2020. Consolidated bioprocessing, an innovative strategy towards sustainability for biofuels production from crop residues: an overview. Agronomy 10 (11), 1834. Available from: https:// doi.org/10.3390/agronomy10111834. Patinvoh, R.J., Taherzadeh, M.J., 2019. Chapter 9 - Fermentation processes for secondgeneration biofuels. In: Basile, A., Dalena, F. (Eds.), Second and Third Generation of Feedstocks. Elsevier, pp. 241 272. Rulli, M.C., Bellomi, D., Cazzoli, A., De Carolis, G., D’Odorico, P., 2016. The water-land-food nexus of first-generation biofuels. Sci. Rep. 6, 22521. Available from: https://doi.org/ 10.1038/srep22521. Saladini, F., Patrizi, N., Pulselli, F.M., Marchettini, N., Bastianoni, S., 2016. Guidelines for emergy evaluation of first, second and third generation biofuels. Renew. Sustain. Energy Rev. 66, 221 227. Schmetz, E., Ackiewicz, M., Tomlinson, G., White, C., Gray, D., 2007. Increasing security and reducing carbon emissions of the U.S. transportation sector: a transformational role for coal with biomass. National Energy Technology Laboratory, DOE/NETL- 2007/1298 August 24. Shah, S.H., Raja, I.A., Rizwan, M., Rashid, N., Mahmood, Q., Shah, F.A., et al., 2018. Potential of microalgal biodiesel production and its sustainability perspectives in Pakistan. Renew. Sustain. Energy Rev. 2018 (81), 76 92. Sheldon, R.A., 2018. Enzymatic conversion of first-and second-generation sugars. Biomass Green Chem. 169 189. Springer. Sikarwar, V.S., Zhao, M., Fennell, P.S., Shah, N., Anthony, E.J., 2017. Progress in biofuel production from gasification. Prog. Energy Combust. Sci. 61, 189 248. Stolarski, M.J., Krzyzaniak, M., Łuczynski, M., Załuski, D., Szczukowski, S., Tworkowski, J., et al., 2015. Lignocellulosic biomass from short rotation woody crops as a feedstock for second-generation bioethanol production. Ind. Crop. Prod. 75, 66 75. UN Report, 2007. Sustainable bioenergy: a framework for decision makers; April 2007 review of EU biofuels directive; public consultation exercise; April July 2006. ˇ Leitgeb, M., 2021. Bioethanol production by enzymatic hydrolysis from difVasi´c, K., Knez, Z., ferent lignocellulosic sources. Molecules 26 (3), 753. Vassilev, S.V., Vassileva, C.G., 2016. Composition, properties and challenges of algae biomass for biofuel application: an overview. Fuel 181, 1 33. Westensee, D.K., Rumbold, K., Harding, K.G., Sheridan, C.M., van Dyk, L.D., Simate, G.S., et al., 2018. The availability of second generation feedstocks for the treatment of acid mine drainage and to improve South Africa’s bio-based economy. Sci. Total Environ. 637 638, 132 136.

Relevant websites https://agsci.oregonstate.edu/sites/agsci.oregonstate.edu/files/bioenergy/generations-of-biofuelsv1.3.pdf http://energyfromwasteandwood.weebly.com/generations-of-biofuels.html

Chapter 4

Challenges to biofuel production 4.1

Introduction

The modern world is facing many challenges like security of energy supply, increase in the oil price, exhaustion of the resources and climatic changes. All these have an adverse effect on the environment. These challenges have induced significant advances in research and development of energy and fuels obtained from biomass. Therefore, on this point, biofuels appear to lessen these problems in a sustained manner. In the transport sector, biofuels have been considered as the most feasible options for reducing the emission of carbon dioxide. Moreover, biofuels can be effortlessly produced from the local resources. The production of biofuels has reached unprecedented volumes over the last 20 years due to the increase in price and the impact on the environment. Several countries all over the world have started to utilize biofuels on a national scale. Biofuels produced from biobased materials are a good option to petroleum based fuels. They offer many advantages to the environment and the society. Biofuels can be produced by several countries, need only minimum changes to retail distribution and end—use technologies. Biofuels are a partial response to global climatic change since they show the potential to stimulate rural development. In comparison to first generation biofuels, production of second generation biofuels is more difficult because of the intricacy of the biomass and issues related to production, harvesting, and transport of less dense biomass to centralized biorefineries. Besides this logistic challenge, other challenges as regards to processing steps in conversion of biomass to liquid transportation fuel still exist. At present, more than 80% of fuel requirement in the whole world is fulfilled by the petroleum and related fuel. According to International Energy Report 2014, the worldwide energy demand is expected to grow by 37% by 2040 (https://www.aa.com.tr/en/economy/global-energy-demand-to-grow-by-37by-2040-international-energy-agency/102390). Therefore, outstanding to limited and diminishing reserves of traditional petroleum fuels, extensive R & D is

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underway and best efforts are being made so that energy demand could easily be met with and some options could be found from sustainable resources. Carriquiry et al. (2011) have reported that second generation biofuels on an energy equivalent basis are more expensive by two to three times as compared to petroleum fuels. For reducing the production cost, many challenges in conversion of lignocellulosic biomass to biofuels and chemicals using biochemical platforms must be addressed (Hoekman, 2009; Menon and Rao, 2012; Luo et al., 2010). “These challenges are in the areas of feedstock production and logistics, development of energy efficient technologies (pretreatment, enzyme hydrolysis and microbial fermentation), coproducts development, establishment of biofuel and biochemical standards, biofuel distribution, societal acceptance, and minimization of environmental impact. All of these challenging areas need expertise in agronomy, biomass logistics and conversion, process engineering, chemistry, conversion technology, microbial fermentation, genetic engineering, economics, and environmental science” (Balan, 2014). Details about the challenges associated with bringing biofuels and biochemicals on the market are presented below.

4.2 Challenges in lignocellulosic biomass conversion to biofuels/biochemicals 4.2.1

Feedstock production and logistics

The cost of feedstocks considerably affects the cost of biofuel production. About 1 billion tons of biomass will be needed annually for displacing 30% of the United States current petroleum consumption. United States Department of Energy’s Office of the Biomass Program published two reports for evaluating the availability of 1 billion tons of feedstock in the United States (https://www1. eere.energy.gov/bioenergy/pdfs/final_billionton_vision_report2.pdf; https://www. energy.gov/eere/bioenergy/downloads/us-billion-ton-update-biomass-supply-bioenergy-and-bioproducts-industry). Three main feedstocks—forest and wood waste resources, agricultural residues and energy crops—were considered. Assumption of the yield were made based on the climatic conditions, crop management, and soil condition. Three different assumptions in yield increase by 2%, 3%, and 4% by year were considered in these studies. In 2012, 341 million tons of feedstock was available. About 30% came from forest residues and 70% from agricultural residues. The agricultural residues showed an acceptable increase whereas the forest biomass showed a marginal increase in yield, because of the projected advancements in agronomy practices. The dedicated energy crops will be one of the main feedstocks for producing bioenergy by 2022. A challenge will be to encourage farmers who grow grains for living to switch over to bioenergy feedstocks. Farmers can be given assurance of buy back guarantees to beat their reluctance and removing the threat of failure for selling their cultivated products.

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In 2030, 1047, 1164, and 1304 dry tons of biomass would be available, based on the three different yield increase assumptions—2%, 3%, and 4% respectively. Other potential high yield producers biomass feedstocks available for conversion are Erianthus, agave bagasse, Napier grass, date palm, and oil palm empty fruit bunch (Somerville et al., 2010). Additionally to these feedstocks, several industrial wastes can be utilized as potential feedstocks for production of biofuels and biochemicals. But, these materials are seasonal and these need to be available in huge quantities for use as raw material for biorefineries. Collection and harvesting of biomass is an essential step involving gathering and removal of the biomass from field that depends upon the state of biomass, that is woody, crop residue or grass. The end use of biomass and the moisture level also affect the way biomass is collected. Harvesting of biomass is an energy consuming process. This needs large machinery and large amount of fuel for transportation (McKendry, 2002). Machinery of different types is used to harvest biomass of different types. The cost of harvesting is affected by the type of machinery used (Thorsell et al., 2004). Woody biomass is harvested as felled-timber. It is then chipped or cut in to different lengths. Energy crops are generally harvested in one- pass using three steps—cutting, raking and baling (Sokhansanj and Hess, 2009). Agricultural residues are harvested by collecting and baling the biomass residue after harvesting the grains (Hoskinson et al., 2007). Most of the moisture conditioning of the agricultural residues is done in the field before baling (Hess et al., 2009). One of the major problem in biomass harvesting is soil contamination of biomass. Other challenges are the moisture level of biomass and the quantity of biomass that can be harvested from the field (Huggins et al., 2011). About one-third of biofuel production cost is associated with biomass cost. The cost of biomass is directly proportional to the yield (Duffy and Nanhou, 2002). This is affected by soil fertility, locality and genetics. Several plant breeding and genetic engineering methods have been used for increasing the yield of many potential energy feedstocks. Additionally, biomass yield can be improved by manipulation of the pathways in abiotic as well as biotic stress (Sticklen, 2006; Bhatnagar-Mathur et al., 2008). A combination of classical plant breeding, transgenesis and modern agricultural practices are being used for increasing the yield of biorefinery feedstocks. It is very important to get the safety ensured of these genetically modified crops for getting the public acceptance. The biomass supply chain includes many important processing steps, which are collection, storage, preprocessing involving densification by compaction, pelleting, and briquetting, transportation from field to biorefinery site and post processing. Fig. 4.1 shows various steps involved in a biomass supply chain (Singhvi and Gokhale, 2019). These supply chain steps will have a direct impact upon the cost of feedstock delivery. Difference in texture, seasonal availability, low bulk density and distribution over a large area

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FIGURE 4.1 Chart of various steps involved in a biomass supply chain. Reproduced with permission from Singhvi, M.S., Gokhale, D.V., 2019. Lignocellulosic biomass: hurdles and challenges in its valorization. Appl. Microbiol. Biotechnol. 103, 9305 9320.

are the main challenges being faced to transport the lignocellulosic biomass to biorefineries (Caputo et al., 2005). Biomass having low density occupies more volume and needs more transportation carrier space and therefore affects the transportation cost. Moisture level, distance from the field to biorefinery, available infrastructure and on-site technology, and the mode of transportation by rail or road also affects the transportation cost (Kumar et al., 2006; Badger, 2003). Biofuel cost depends very much upon the biorefinery size, which requires optimization based on the location and raw material (Kumar et al., 2003, 2005; Kumar and Sokhansanj, 2007). Development of successful biomass supply chains requires sustainable volumes of bulky biomass to be transported to the conversion facility at an economically feasible cost. This needs proper project planning and operations. A comprehensive report on guidelines for developing a sustainable biomass supply chain has been published (https://www.bioenergyconsult. com/tag/biomass-supply-chain). Woody biomass can be transported as whole tree residue, wood chip, bundle, pelleted or baled (square and round) forms. Agricultural biomass can be transported as loose, chopped, baled (square or round), or pelleted forms.

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The biomass density drastically changes depending upon the level of processing that the feedstock goes through (An et al., 2011; Zhu et al., 2011). When comparison is made with loose biomass, the density of pellets can increase to almost 8 times for woody biomass and 10 times for agricultural biomass. Biomass can be transported by trucks. If it is assumed that about 2000 tons of biomass/day for producing biofuels and biochemicals is required by a biorefinery (Kumar et al., 2003), transport of woody biomass or agricultural biomass as pellets would take 88 trips/day vs loose biomass, which would require 298 trips/day for agricultural biomass and 392 trips/day for woody biomass. The biomass transportation cost is expressed as a fixed cost and variable cost using round trip distance. There is an increase in transportation cost as the distance from the biorefinery site to biomass storage site increases (Sultana et al., 2010). Preservation of harvested biomass from rain or moisture has been carried out by storing in a building or through covering of polythene wraps before shipping to biorefinery.

4.2.2

Lignocellulosic biomass pretreatment

Lignocelluloses mainly consist of cellulose, hemicellulose, and lignin. The relative amount of these three polymers vary with different types of biomass. These polymers are associated with each other in a hetero-matrix to varying degrees and varying relative composition depending upon the type, species, and source of the biomass (Mood et al., 2013; Pauly and Keegstra, 2010). Biomasses from plants are obviously recalcitrant, having evolved to defend themselves against attacking microbes. The rate of decomposition of the plant biomasses is slow as these are recalcitrant, and may take several months to years for complete degradation of dead plants. Conversely, in a biorefinery, the conversion of biomass to biofuels is done in days. For increasing the accessibility of cellulose and hemicellulose, the lignin-hemicellulose complex cross-links must be broken. Several pretreatment processes have been developed (Cadoche and Lo´pez, 1989; Damaso et al., 2004; Gregg and Saddler, 1996; Itoh et al., 2003; Keller et al., 2003; Kim et al., 2003; Kuhad et al., 1997; Kuo and Lee, 2009a,b; Li et al., 2012a,b,c; Wang et al., 2011a,b; Zhang et al., 2009, 2012a,b, 2013; Zhao and Liu, 2012). Pretreatment plays an important role in lignocellulosic biomass conversion process. It alters the structure of lignocellulosic biomass and makes it accessible for enzymatic saccharification. Important pretreatment methods are listed in Table 4.1. Not all pretreatments are equally commercially viable. The ease with which a pretreatment process can be commercialized depends on the following factors (Ganguly et al., 2018; Balan, 2014): G

A pretreatment process which involves opening of the cell wall to bring lignin on the surface shows the potential to efficiently densify after

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TABLE 4.1 Processes for the pretreatment of lignocellulosic biomass. Acid hydrolysis Advantages Hydrolyzes hemicellulose to xylose and other sugars; alters lignin structure Increase in porosity/increased enzymatic hydrolysis Limitations and disadvantages High cost; equipment corrosion; formation of toxic substances; Generation of furfural/ hydroxymethyl furfural/need for recycling/costly Alkaline hydrolysis Advantages Removes hemicelluloses and lignin; Increases accessible surface area Limitations and disadvantages Long residence times required; irrecoverable salts formed and incorporated into biomass; Formation of salts of calcium and magnesium Organosolv Advantages Hydrolyzes lignin and hemicelluloses; Pure lignin obtained and used as value added product Limitations and disadvantages Solvents need to be drained from the reactor, evaporated, condensed, and recycled; high cost; solvents inhibit enzymatic hydrolysis AFEX Advantages Increases accessible surface area, removes lignin and hemicellulose to an extent; does not produce inhibitors for downstream processes; decrystallization of cellulose Limitations and disadvantages Not efficient for biomass with high lignin content; Costly Ammonia treatment Advantages Removal of lignin/ decrystallizing cellulose Limitations and disadvantages Removal of ammonia/costly Mechanical comminution Advantages Reduces cellulose crystallinity Limitations and disadvantages Power consumption usually higher than inherent biomass energy Steam explosion Advantages Causes hemicellulose degradation and lignin transformation; cost-effective Limitations and disadvantages Destruction of a portion of the xylan fraction CO2 explosion Advantages (Continued )

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TABLE 4.1 (Continued) Increases accessible surface area; cost-effective; does not cause formation of inhibitory compounds Limitations and disadvantages Does not modify lignin or hemicelluloses Pyrolysis Advantages Produces gas and liquid products Limitations and disadvantages High temperature; ash production Ozonolysis Advantages Reduces lignin content; does not produce toxic residues Limitations and disadvantages Lignin is damaged; cellulose/hemicelluloses unaltered; Large amount of ozone required; expensive Biological Advantages Degrades lignin and hemicelluloses; low energy requirements Limitations and disadvantages Rate of hydrolysis is very low; A part of fermentable sugars are utilized as carbon source Wet oxidation Advantages Treatment of wastes Limitations and disadvantages Costly Microwave Treatment Advantages Cheap/generates less pollution Limitations and disadvantages Degradation of cellulose/ hemicellulose Source: Based on Kumar et al. (2009), Chaturvedi and Verma (2013).

G

G

pretreatment without the addition of any binding agents and so is perfect for decentralized biomass processing. The durability of biomass for long term storage is also increased. AFEX, wet oxidation, and extrusion at high temperature fall under this category. Commercial viability increases if the densified, pretreated biomass has twofold application (animal feed, soil amendments, fertilizer and biomass composites) in addition to using them as biorefinery feedstock. Pretreatment process that needs large quantity of water for removing toxins from the pretreated biomass makes the process more costly and so less lucrative.

74 G

G

G

G

G

G

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Processes that can be scaled up for meeting the biorefinery requirement of handling more than 2000 tons/day or more are more feasible. The capital cost of pretreatment reactor increases based on the pretreatment conditions and the catalyst used. Processes that use lesser energy and inexpensive chemicals, reducing processing cost are more cost-effective. Processes which preserve lignin during pretreatment (as opposed to pretreatments such as alkaline hydrogen peroxide and ozonolysis which have the tendency to degrade lignin) and therefore the energy density of lignin are far more cost-effective. Pretreatments which require moderate temperatures are preferred from a cost perspective. Processes which use supercritical fluids (water and carbon dioxide) work at a very high pressure and need extra cost. Processes using less hazardous chemicals are preferred. For instance, chemicals such as hydrofluoric acids must undergo extra safety steps for avoiding accidents during biomass processing, which increases the processing cost. Recovery of catalyst during pretreatment is very important for the environment. Though this slightly increases the processing cost, there will be in general savings in energy, as lesser amount of chemicals is used during the process.

After pretreatment, the properties of the pretreated biomass (specific surface area, cellulose crystallinity index, degree of polymerization, lignin and acetyl content in biomass) may be significantly changed. Effective pretreatment method increases the enzymatic hydrolysis rate and significantly reduces the amount of enzymes required for converting the biomass into sugars, which can be used by microbes. As lignin is highly recalcitrant and is accountable for unproductive binding of enzymes, the sugar conversion efficiency is affected by the quantity of lignin present in pretreated biomass. Removal of lignin during the pretreatment process will allow recovery and reuse of enzymes. This will result in substantial cost savings. The pretreatment step is one of the most cost contributing and rate and yield limiting steps (Table 4.2). Pretreatment requires an investment which represents about 30% of the total equipment cost. One of the major challenges is the intrinsic recalcitrance of the lignocellulosic material which results in lower biomass to sugar yields and high pretreatment cost (Ganguly et al., 2018).

4.2.3

Enzymatic hydrolysis

Enzymes are secreted by microorganisms for degrading biomass to produce monomeric sugars for their own survival. Microorganisms use the biomass

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TABLE 4.2 Cost area distribution of a general lignocellulosic greenfield facility (Valdivia et al., 2016). Boiler, steam turbine generator

29%

Pretreatment

24%

Distillation

12%

Evaporation

8%

Utilities

6%

Biomass handling

5%

Saccharification

5%

Fermentation

4%

S/l separation

2%

Propagation

15%

Others

4%

Pretreatment and Boiler are the most cost-intensive areas. Source: Based on Valdivia, M., Galan, J.L., Laffarga, J., Ramos, J.L., 2016. Biofuels (2020): Biorefineries based on lignocellulosic materials. Microb. Biotechnol. 9 (5), 585 594. doi:10.1111/ 1751-7915.12387.

degrading enzymes in a natural environment in two ways. The first one is cellulosomal enzyme system (Bayer et al., 2013). Cellulosome complexes are intricate, multienzyme machines, produced by many cellulolytic microorganisms for the degradation of lignocelluloses. They consist of a complex of scaffoldin, which is the structural subunit, and a range of enzymatic subunits. The intersubunit interactions in these multienzyme complexes are mediated by cohesin and dockerin modules. The second one is the free enzyme system. The enzymes are secreted as individual components to act upon the biomass substrates (Hatakka and Hammel, 2010). The free enzyme system is easily produced and is commonly used for biomass conversion in biorefineries. Free biomass degrading enzymes are produced by commercial companies using fungi or bacteria. The microorganisms are fed with agricultural residues. This mixture contains several biomass degrading enzymes which include cellulase, hemicellulase, and pectinase enzymes (Zhang et al., 2012a). Cellulases constitute about 70% 85% of the mixture and hemicellulases and pectinases constitute the remaining 15% 30% depending on the organisms and the substrate. Another type of enzyme that may also play an important role in biomass deconstruction is ligninolytic enzymes which degrade lignin. Different types of enzymes are required to break different types of bonds during hydrolysis of lignocelluloses because of the complex network of cellulose and hemicellulose (Somerville et al., 2004). Such

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enzymes are called molecular scissors and produce specific monomeric sugars from complex carbohydrates. Numerous papers have been published on these enzymes and their synergistic action during hydrolysis (Gao et al., 2010, 2011; Banerjee et al., 2010; van Dyk and Pletschke, 2012). Cellulosehydrolyzing enzymes are classified into endoglucanase, cellobiohydrolase, and β-glucosidase. Endoglucanase produce random cuts at an internal position within cellulose fiber and release cellooligosaccharides. Exoglucanases/ cellobiohydrolase act on chain ends and release cellobiose which are then acted upon by β-glucosidase to release glucose (Xie et al., 2007). For that reason, cellulase can convert natural lignocellulose containing crystalline cellulose to glucose, but the rate of reaction is low. Trichoderma reseei which is a filamentous fungus produces high concentrations of enzymes up to 100 g/L and is generally used by Enzyme companies to produce biomass-degrading enzymes (Balan et al., 2013). Other thermophilic bacterial and fungal enzymes have recently been introduced to the market, which perform at higher temperatures for preventing contamination with Lactobacillus (Acharya and Chaudhary, 2012; Liszka et al., 2012). The amount of enzyme needed to hydrolyze lignocelluloses is onefold higher as compared to starch. This is due to the reason that lignocelluloses are naturally recalcitrant with a complex ultrastructure. Lignin in plant cell wall and degradation products generated during pretreatment deactivate enzymes and therefore increased quantity of enzymes are needed (Ximenes et al., 2010). Enzyme companies are making considerable progress in producing new generations of enzymes with higher specific activities and reduced cost using diverse process engineering and biotechnological methods (Alvira et al., 2013). But, technoeconomic analyses show that more research is required. Research is underway, for reducing the cost of enzymes (Hong et al., 2013). Several strategies have been explored for producing highly active enzymes. These include the following (Rizk et al., 2012; Horn et al., 2012; Dashtban et al., 2009; Song et al., 2012). G

G

G

Identification of novel multifunctional enzymes which are able to hydrolyze different types of polysaccharide linkages and have higher activity Production of novel enzymes by the use of different genetic engineering techniques Direct evolution of enzymes

Another strategy is expressing enzymes in plants that could be extracted and used after pretreatment of the extracted biomass for the production of fermentable sugars (Egelkrout et al., 2012). Enzymes can also be produced directly in biorefineries rather than producing them in a centralized location. On-site production of enzymes at biorefineries would avoid the requirement for concentration, storage, and shipping and would cut the production costs by the use of pretreated substrates already available at the biorefinery (Culbertson et al., 2013).

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The time required for complete hydrolysis of biomass to monomeric sugars depends upon the following factors (Chundawat et al., 2011; Zeng et al., 2014; Jeoh et al., 2007; Hall et al., 2010). G G G G G

Lignin content in biomass Pretreatment effectiveness Cellulose crystallinity Substrate concentration Enzyme activity

Removal of hemicelluloses during pretreatment drastically reduces the recalcitrance of biomass. This allows for a faster rate of sugar conversion (Hall et al., 2010; Yu et al., 2011; Ju et al., 2013). To avoid the long hydrolysis time, a new strategy was developed. The biomass is initially hydrolyzed for 24 hours and the sugars are removed after 24 hours and fermented separately. The remaining solids require further hydrolysis are added to fresh pretreated substrate in the same tank along with fresh enzymes for further hydrolysis. This strategy can help in reducing the biomass to sugar processing time (Jin et al., 2012a). As biomass degrading enzymes are costly, they should be recycled for reducing the processing cost. The methods that have been used for recycling the enzymes include immobilized enzymes on nanoparticles or polymeric matrices, ion exchange adsorption, and ultrafiltration. (Ansari and Husain, 2012; Mackenzie and Francis, 2013; Wu et al., 2010; Qi et al., 2011). About 44% cost of biofuel production in second generation ethanol production is from enzymes. On-site or near-site enzyme generation encourages access to the significant reduction in enzymes’ value up to 30% 70% carrying its interpreted purification and logistics (Singh et al., 2019; Khajeeram and Unrean, 2017). The cost of enzymes at 15% and 35% solid pretreated loading during enzymatic hydrolysis varies from 34.63% to 36.38%, respectively (Singh et al., 2019; Solarte-Toro et al., 2019).

4.2.4

Microbial fermentation and biomass

“Microbial fermentations convert sugars produced from lignocellulosic biomass into biofuels or biochemicals using bacteria, fungus or yeast.” This process can be performed separately from enzymatic hydrolysis, in combination with enzymatic hydrolysis (simultaneous saccharification and fermentation), or combine enzyme production and enzymatic hydrolysis (consolidated bioprocessing). Fermentation of glucose and xylose can be carried out separately or in combination (cofermentation, where both glucose and xylose are simultaneously converted). These strains do not naturally consume xylose but are made capable through genetic modification. Cofermentation is believed to be superior than separate fermentation in terms of cost savings. Many of challenging efforts have been made to

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genetically modify Saccharomyces cerevisiae and Zymomonas mobilis to ferment of xylose. Two xylose-fermenting pathways have been widely studied and engineered into S. cerevisiae namely, (1) xylose reductase (XR), xylitol dehydrogenase (XDH) pathway that exists in fungi and (2) xylose isomerase (XI) pathway that exists in bacteria. These two pathways convert xylose to xylulose. Some microbes, such as E. coli and Pichia stipitis, can natively ferment xylose. However, they typically have limitations, which reduce their effectiveness compared to S. cerevisiae or Z. mobilis (Wohlbach et al., 2011; Ha et al., 2010; Kuyper et al., 2004; Sonderegger et al., 2004; Asghari et al., 1996; Jin et al., 2012b). Another strategy adopted by Microbiogen (http://www.microbiogen.com/) involves fermenting glucose to ethanol under anaerobic conditions and converting xylose to yeast biomass under aerobic conditions using a native S. cerevisiae strain MBG3248, which was screened and adapted for propagation on xylose. This approach produced 173 L of ethanol and 134 kg of yeast protein from one ton of corn stover using dilute acid pretreatment technology. Lignocellulosic biomass can also be directly fermented to biofuels by CBP microbes, such as Clostridium thermocellum and Clostridium phytofermentans (Balan, 2014; Kollaras et al., 2011; Ng et al., 1981). In a biorefinery, microbial biomass is one of the potential coproducts (Sun and Cheng, 2002). In the batch fermentation process, part of the microbial biomass can be reused in the next batch (Lau et al., 2012). The left over biomass can be used as an animal feed if native organism is used. As most of the microorganisms used for fermentation in the second generation biorefineries are genetically modified organisms which can utilize glucose as well as xylose, there may be some potential directives for the utilization of microbial biomass as an animal feed. In the semi-continuous fast bioconversion with integrated recycle technology process, similar idea may apply (Jin et al., 2012a) where the microbial biomass is recycled every 24 hours for the next batch. Surplus cells are used for other uses. Moreover, separated cells have many degradation products of lignin and reduce the value of feed. Proteins from these microbes are processed and converted to amino acids by the use of acid hydrolysis. This strategy can be used for producing the chemicals which can be used for producing the biomaterials. Biological conversion of LCB components in to biofuels and different chemicals is shown in Fig. 4.2 (Singhvi and Gokhale, 2019).

4.2.5

Biofuel cost

“Several cost models have been developed in the past to understand the required capital investments. With various assumptions in place, when plant capacity (0 15,000 ton/day) was plotted against total production cost ($m23), it was found that a biorefinery should handle more than 2000 5000 ton/day in order to be economical (850 875 $m23). Plants that operated at more than 5000 ton/day

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FIGURE 4.2 Schematic representation of biological conversion of lignocellulosic components into various chemicals and biofuels. Reproduced with permission from Singhvi, M.S., Gokhale, D.V., 2019. Lignocellulosic biomass: hurdles and challenges in its valorization. Appl. Microbiol. Biotechnol. 103, 9305 9320.

showed a marginal increase in the production cost. According to some research findings, there is a steady decrease in the minimum ethanol selling price from 3.75 to 2.25 ($/gallon) as the plant size increased from 500 to 10,000 dry ton per day. The cost analysis is based on several factors like biomass transportation distance, type of biomass used and processing technology, efficiencies of different processing steps, type of biofuel, and coproducts production” (Balan 2014; Aden et al., 2002; Kaylen et al., 2000; Wright and Brown, 2007; Leboreiro and Hilaly, 2011, 2013; Argo et al., 2013).

4.2.6

Water recycling

Consumption of water is high in the production of biofuels using the sugar platform. For meeting the water requirements, key decisions must be made regarding the location of the biorefineries. In several cases, water is pumped from the ground, which adds substantial pressure on the local water resources (Bernardi et al., 2013; Yuliani et al., 2013). Recycling of the water helps to lessen this pressure, but requires additional investment. When recycling the water, it will be important to remove salts produced during neutralization, remove the organic material, and recycle the catalysts (Bernardi et al., 2013). Clarification is used

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for removing suspended solids and reducing chemical oxygen demand/biochemical oxygen demand in water. Many methods have been developed for treatment of water. These methods include coagulation, coagulation-flocculation, electrocoagulation, polymer resin filtration, and biological treatment. Among these methods, coagulation is a best approach for removing organics (Verenich and Kallas, 2001). Minimal water should be used in each processing step for reducing the cost of water recycling. In few cases, the selection of pretreatment method in a second generation biorefinery will be determined based on the availability of the water in the area. Colocating the first and second generation biorefineries will reduce waste water and the pressure on the water table (Balan et al., 2013; Dias et al., 2013).

4.2.7

Generation of coproducts

Generation of coproducts is very crucial for producing competitively priced biofuels. The first generation ethanol produced from corn consists of 67% dry mill based and 33% wet mill based operations. Generation of coproducts depend on the type of mill (Bothast and Schlicher, 2005). More ethanol is produced in the dry mill process as compared to the wet mill process. Dry mill process produces 2.8 gallon/bushels of corn whereas the wet mill process produces 2.5 gallon/bushels of corn. However, more coproducts are produced in the wet mill process. This results in more income. The corn dry mill industry produces dry distiller’s grains and soluble (DDGS) and carbon dioxide as the main coproducts which reduce the biofuel cost by 35% (Hettinga et al., 2009). The corn wet milling process involves, higher energy consumption and capital investment. This process produces multiple coproducts which include corn oil, corn gluten meal, corn gluten feed and carbon dioxide. For the second generation technology to be able to compete with the first generation technology, several coproducts should be produced which can be sold at a higher price. This will reduce the overall processing costs of biofuels (Stephen et al., 2012). From second generation biorefinery, some of the coproducts that could be produced are lignin, protein, microbial biomass (Tuck et al., 2012).

4.2.8

Energy and environmental issues

The success of a biorefinery will be assessed by the net energy produced using the different processing steps. Effectiveness will need to be strictly observed for maximizing the net gain in the energy produced in the biorefinery. In many technoeconomic evaluations of second generation biorefineries, lignin has been reported to be a good energy source for different processing steps. The unhydrolyzed solids rich in lignin are obtained as wet slurry and are dried before burned or gasified to produce energy. Hydrothermal

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pyrolysis technology uses wet biomass slurry for producing biooil and uses the heat recovered for several processing steps. If extraction of lignin is done during pretreatment and used for the production of value added products, the biorefineries will produce energy by burning part of the biomass which come at biorefineries (von Blottnitz and Curran, 2007; Gonc¸alves et al., 2013; Zhu and Zhuang, 2012; Kelloway and Daoutidis, 2013; Ruiz et al., 2013; Cherubini, 2010). Natural gas will be the main energy source for running biorefineries, and as the technology gets matured, the amount of natural gas usage will be gradually reduced. Colocation of biorefineries near thermal plants or use of energy from sustainable sources is another choice for getting heat and power. Addition of an anaerobic digestion facility near biorefinery will be advantageous that could treat the waste water and simultaneously the biogas produced can be utilized for various biorefinery operations (Yılmaz and Selim, 2013; Maclellan et al., 2013; Nizami and Ismail, 2013). Biofuels are helping in reducing greenhouse gas emissions and the related climatic change impact from transport so these are being promoted as a lowcarbon alternative to fossil fuels (Kendall and Yuan, 2013). In the future, when a new biorefinery is set up, many technologies will be explored based on their impact on the environment (Borrion et al., 2012). Few examples are air pollution caused by particulate emission during harvesting and grinding of biomass, noise pollution from explosive pretreatment processes, processes for producing pretreatment chemicals which generate greenhouse gas emissions, and discharge of pretreatment chemicals to the environment after processing. Life cycle analysis (LCA) is used for assessing the impact of these processing steps on the environment (Uihlein and Schebek, 2009; MartinezHern´andez et al., 2013). The challenges are in conducting the precise LCA analysis depending upon the data collected from the processes used in the biorefinery. Most of the companies are using LCA for assessing the environmental impact so that decisions can be taken for adjusting the process or areas that needs to be focused for reducing the emissions. The cost of establishing the biorefinery depends upon the emissions estimated by LCA. Big companies having capital to invest in a new biorefinery are making efforts to align different novel process technologies having many challenges to overcome (Ramaswamy et al., 2013). The selection of pretreatment method and lignocellulosic biomass can be decided based upon the availability of adequate amount of catalyst and feedstock in that area. The associated technologies are presently being scaled up to set up pilot plants for demonstrating the feasibility. Biomass logistics and technoeconomic evaluations are conducted at the same time for assessing the technology readiness level. Then environmental impact of different methods are assessed. Once suitable feedstock, pretreatment, and enzymes are used for producing low priced sugars, the selection of biofuels and biochemicals depends upon the market demand and more notably the biofuel

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policy defined by the local and federal government. The cost of biofuel processing must be kept lower using energy saving technologies and using lesser water in order to compete with the cost of petroleum fuels. Production of several coproducts in a biorefinery would help to reduce the cost of biofuel production (Balan, 2014). “NREL has made path breaking discoveries and inventions in LCB biorefineries, the technology is still in nascent stage. Considerable challenges remain to be addressed in the economical development of LCB biorefinery. The challenges are faced in every unit operation such as sustainable biomass supply chains, biomass pretreatment, fractionation, saccharification, and conversion of sugars and lignin to fuels and chemicals. One of the critical hurdles is the cost of feedstock which is dependent upon the supply chains and therefore, careful analysis and research on the current supply chains would contribute to a great extent to improve the efficiency of currently available supply chain pathways. In addition, transportation of LCB, pests, diseases, and land use have made the second generation biofuels expensive. The pretreatment step is one of the most expensive and rate and yield limiting steps. Pretreatment is a physical, chemical, physicochemical, or biological process which opens up the recalcitrance structure amenable to enzymatic attack. However, none of the pretreatment strategies was found to be eco-friendly and environmental friendly and hence, pretreatment of the biomass seems to be still unresolved problem. The future of LCB conversion to biofuels is expected to lie in the development of cost-effective pretreatment technologies for efficient hydrolysis of LCB even with using less enzyme doses. In addition, substantial reduction in the chemicals needed for pretreatment is necessary to have an economically/environmentally competitive process. Though several efforts have been devoted to develop enzyme cocktails, suitable and low-cost enzyme cocktails still remain as challenges in biomass processing. Different enzyme sources need to be tapped and evaluated for designing customized enzymatic cocktails. Metagenomic tools as well as bioprospection of the microbes could be applied to discover new sources of enzymes. From industrial point of view, the knowledge of enzymatic synergism in the enzyme cocktails is of great importance which may lead to rapid conversion of biomass, reducing the enzyme loads and thereby reducing the cost of LCB hydrolysis. The recent publications on GVL-pretreatment and CelA-type enzymes acting on raw biomass may help meet these challenges to make the biomass conversion technologies eco-friendly with affordable cost” (Singhvi and Gokhale, 2019).

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Relevant websites https:// www.bioenergyconsult.com/tag/biomass-supply-chain. https://www.aa.com.tr/en/economy/global-energy-demand-to-grow-by-37-by-2040-internationalenergy-agency/102390. https://www.energy.gov/eere/bioenergy/downloads/us-billion-ton-update-biomass-supply-bioenergy-and-bioproducts-industry. U.S. Billion-Ton Update: Biomass Supply for a Bioenergy and Bioproducts Industry. https://www1.eere.energy.gov/bioenergy/pdfs/final_billionton_vision_report2.pdf. Biomass as Feedstock for a Bioenergy and Bioproducts Industry: The Technical Feasibility of a BillionTon Annual Supply. http://www.microbiogen.com

Further reading ´ .D., Peralta-Ruiz, Y., Pardo, Y., Kafarov, V., 2013. Energy integration of Gonz´alez-Delgado, A bioethanol production process topology from microalgae biomass: evaluation of SSCF, SSF, acid hydrolysis and product purification alternatives. Chem. Eng. Trans. 35, 1069 1074. Vanholme, R., Morreel, K., Darrah, C., Oyarce, P., Grabber, J.H., Ralph, J., et al., 2012. Metabolic engineering of novel lignin in biomass crops. N. Phytologist 196 (4), 978 1000.

Chapter 5

Current production status of cellulases and challenges 5.1

Introduction

Global production of biofuel in 2020 reached 1677 thousand barrels of oil equivalent per day, in clear contrast to the 187 thousand barrels of oil equivalent per day that was produced in 2000 (http://www.statista.com). Growth has mostly been motivated by policies which promote the use and production of biofuels as they could help in reducing greenhouse gas emissions and the related climate change impact from transport. Biofuels can be advantageous because of its limited impact on the environment as compared to fossil fuels and also its utilization of waste materials which would generally be discarded. The biofuel market has been affected by blending mandates, fuel quality standards, sustainability criteria, and import tariffs. By 2024, the worldwide biofuel market is expected to reach a market size of 153.8 billion US dollars. The two most common types of biofuels in use today are ethanol and biodiesel. The most common ethanol feedstocks are sugarcane and coarse grain but this could vary regionally. Biodiesel is produced from oils or fats using transesterification and is the most common biofuel in Europe. The most commonly used feedstocks for production of biodiesel is vegetable oil, whereas nonagricultural feedstocks such as waste are becoming more pertinent in regions like Europe and the United States. Cellulases play an important role in the enzymatic hydrolysis of cellulosic polymers. These enzymes release monomeric sugars which can be fermented to ethanol. Cellulases are induced enzymes produced by a large variety of microbes including fungi and bacteria during their growth on cellulosic materials. These microbes can be aerobic, anaerobic, mesophilic, or thermophilic. The most widely studied cellulose producers are genera of Trichoderma, Aspergillus, Cellulomonas, Clostridium, and Thermomonospora (Kuhad et al., 2011). Cellulases are the most comprehensively studied enzymatic complex. These enzymes hydrolyze β -1,4 linkages in the cellulose structure to release glucose, cellobiose and cello-oligosaccharides. “The catalytic modules of cellulases have been classified into numerous families based on their amino acid sequences and crystal structures (Henrissat, 1991). Cellulases contain Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00008-X © 2023 Elsevier Inc. All rights reserved.

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noncatalytic carbohydrate-binding modules (CBMs) and/or other functionally known or unknown modules, which may be located at the N- or C-terminus of a catalytic module. In nature, complete cellulose hydrolysis is mediated by a combination of three main types of cellulases: 1. endoglucanases (EC 3.2.1.4) 2. exoglucanases, including cellobiohydrolases (CBHs) (EC 3.2.1.91) 3. β -glucosidase (BG) (EC 3.2.1.21) To hydrolyze and metabolize insoluble cellulose, the microorganisms must secrete the cellulases (possibly except BG) that are either free or cellsurface-bound. Cellulases are increasingly being used for a large variety of industrial purposes” (Liu and Kokare, 2017). Cellulases are projected to dominate worldwide market share of pulp and paper enzymes and may account more than 37% share by 2026. The growing concerns about exhaustion of crude oil and the emission of greenhouse gases have triggered off the production of bioethanol from lignocellulosic materials, specially through enzymatic hydrolysis of lignocelluloses—sugar platform (Bayer et al., 2007; Himmel et al., 1999; Zaldivar et al., 2001). But, the cost of cellulase enzyme for hydrolysis of pretreated lignocellulosic biomass needs to be reduced, and their catalytic efficiency should be further enhanced for making the process economically viable (Sheehan and Himmel, 1999). Engineering of cellulolytic enzymes with improved catalytic efficiency and increased thermostability is vital for commercializing lignocellulosic biorefinery. Activity of individual cellulases can be improved by using either directed evolution or rational design. However, improvements in performance of cellulase have been incremental, and no significant activity improvement has been observed so far. Further improvement on cellulase performance requires the improved understanding of mechanisms of cellulose hydrolysis and also the relationship of cellulase function, molecular structure as well as substrate characteristics. Many fungi—Aspergillus and Trichoderma spp., etc.—are the efficient producers of cellulases. These fungi are further classified as wood-rotting fungi (Yoon et al., 2014). Next, wood-rotting fungi can be differentiated into three categories, namely white-rot fungi, brown-rot fungi, and soft rot fungi. Among these, soft rot fungi are known as potential cellulase producers (e.g., Trichoderma reesei and Aspergillus niger). Besides, number of the white-rot fungi (e.g., Phanerochaete chrysosporium) and brown-rot fungi (e.g., Gloeophyllum trabeum) are also known for proficient production of cellulases (Rasmussen et al., 2010; Shrestha et al., 2008; Yoon et al., 2014). Although, the above-stated fungi show potential for effective production of cellulases, the hydrolysis of cellulosic biomasses using these cellulase enzymes is incomplete because of the lack of a particular cellulase component. For this reason, further research targeting to improve the efficiency of cellulases and their production is needed.

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5.2

93

Production of cellulases

Generally for commercial production of cellulases, T. reesei and A. niger fungi have been used. Both these fungi are mesophilic and grow well at 30 C to 35 C. Enzymes produced by thermophilic microorganisms perform well up to 90 C and are more favorable for industrial processes under severe conditions because of the faster reaction rate. The chances of contamination are decreased at higher temperatures (Biswas et al., 2014). Cellulases are produced by either solid-state fermentation (SSF) or submerged fermentation (SmF). SmF has been widely used for the industrial production of enzymes up to now, but SSF is rapidly gaining current scientific attention for the production of primary and secondary metabolites due to many environmental and apparent economic advantages it offers (Pandey, 1992, 1994, 2003). It presents potential opportunities for developing “green processes” using agroindustrial residues (Pandey and Soccol, 1998, 2000; Pandey et al., 2000, 2007). On an industrial scale, SSF is considered for the production of lowvalue enzymes like cellulases, xylanases, amylases, etc. The optimization of a fermentation process includes selection of media composition, type of cultivation, and process conditions regardless of the type of bioprocess. Significant endeavor and time needs to be expended for achieving these tasks. Once the organism has been chosen, the production process is developed.

5.2.1

Submerged fermentations

Most of the reports on microbial production of cellulase enzymes use SmF process and the broadly studied organism used for this purpose is T. reesei, which has been studied mostly in liquid media (Singhania et al., 2010). Bioreactors for SmF are well developed. Almost all the commercial enzyme producers are using the established technology of SmF because of better monitoring and also handling is easier. The industrial enzymes are produced in fermenters of 50 to 500 m3 capacity. In SmF, the medium is liquid which remains in contact with the microorganism. A supply of oxygen is necessary in SmF. There are four types of SmF processes: G G G G

batch culture fed-batch culture perfusion batch culture continuous culture

In the batch culture, the cells are grown in a fixed volume of nutrient culture medium under specific environmental conditions (e.g., nutrient type, temperature, pressure, and aeration). In continuous culture, during the exponential phase of microbial growth, fresh medium is added into the batch reactor. The medium containing the product is

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withdrawn from the reactor. Continuous culture gives near-balanced growth, with little fluctuation in nutrients, metabolites, cell numbers, or biomass. Extracellular enzymes are mostly recovered after cell removal (by separators or microfiltration, vacuum drum filtration) by ultrafiltration. If required, purification is conducted by using gel filtration or ion exchange. The final product is either a concentrated liquid with necessary preservatives like salts or polyols or alternately granulated to a nondusty dry product. It should be noted that enzymes are proteins, and so can cause and have caused in the past allergic reactions. So, protecting measures are essential in their production and application. A fed-batch process lies in between both of the above-stated processes (dos Reis et al., 2013). In this case, the concentrated components of the nutrients are slowly added to the batch culture. In perfusion batch culture, the addition of the culture and withdrawal of an equal volume of used cell-free medium is carried out. The perfusion method circulates medium through a growing culture. Allows simultaneous removal of waste, supply of nutrients, and harvesting of product. Perfusion processes also can offer considerably higher productivities in grams/l of bioreactor working volume per day. This enables the use of smaller bioreactor reducing capital investment. SmF is good in several cases because it can be easily handled and can be monitored with ease (dos Reis et al., 2013). In this type of fermentation, the nutrients and oxygen are easily dissolved in liquid and get dispersed evenly all over in the vessel, and due to this, heat and mass transfer takes place effectively (Biswas et al., 2014). Submerged batch process is extensively used for cellulase production; but higher yields of cellulase productions have been claimed using fed-batch techniques (Belghith et al., 2001; Ximenes et al., 2007). Fed-batch technique is also effective in reducing the negative effects of catabolic repression and viscosity (Esterbauer et al., 1991). Almost all companies are using submerged fed-batch fermentation process for producing low-cost cellulases since this technique can produce more than 100 g of crude cellulase/l of broth. It is estimated that cost of cellulase production based on dry protein weight can be as low as $10 to 40 per kilogram of dry protein weight. Most enzyme companies (e.g., Novozymes, Genencor, and Iogen) are producing commercial cellulases using Trichoderma and Aspergillus or their derivative strains whereas Dyadic International Inc. is using Chrysosporium lucknowense. Over the last few years, Genencor and Novozymes reported a 20to 30-fold reduction in cellulase production costs to 20 to 30 cents per gallon of cellulosic ethanol (Himmel et al., 2007). This is achieved mostly by 1. reducing the sugar costs from lactose to glucose plus a small amount of sophorose, 2. enzyme mixture for higher specific activity, and 3. better thermostability.

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But this achievement appears to be overemphasized. It is estimated that current cellulase costs may range from 1.00 to 1.50 US dollars per gallon of cellulosic ethanol. Production of cellulase is growth associated and is affected by several factors and their interactions may affect the productivity of cellulase (Tholudur et al., 1999). Out of the known inducers of cellulase genes, lactose has been reported to be the only economical additive in industrial fermentation media (Aro et al., 2005). In T. reesei a basal medium after Mandels and Reese (1957) has been mostly used with or without modifications. Carbon sources used in most of commercial cellulase fermentations are cellulosic biomass which includes straw, rice or wheat bran, bagasse, spent hulls of cereals and pulses, paper industry waste, and several other lignocellulosic residues (Adsul et al., 2004; Belghith et al., 2001; Heck et al., 2002; Reczey et al., 1996; Romero et al., 1999; Shen and Xia, 2004; Szijarto et al., 2004; Wen et al., 2005a, 2005b). Although most of the processes are batch processes, attempts have been made to produce cellulase in fed batch (Belghith et al., 2001; Ghose and Sahai, 1979) or continuous (Bailey and Tahtiharju, 2003; Ju and Afolabi, 1999; Schafner and Toledo 1992) mode, which supposably helps to supersede the repression which is caused by accumulation of reducing sugars. Increased fermentation time with a lower productivity is the main technical limitation in fermentative production of cellulases. Stirred-tank reactors (STRs) are most commonly used for the production of cellulases. In these types of reactors the main drawback is controlling the shear stress, which is often found to be harmful for the growth of filamentous fungi like T. reesei by damaging the filaments of the fungi. This damage of filament and disturbed growth result in lesser production of enzymes. Other bioreactors, that is airlift reactors, are better versions of the SmF technique. These bioreactors offer advantages over conventional systems, like simplicity of construction, reduced risk of contamination, and efficient gas liquid dispersion with reduced power consumption. Airlift reactors are usually operated under atmospheric pressure (Biswas et al., 2014). The rotating fibrous-bed reactor (RFBB) is a type of reactor that is different from the STR. Lan et al. (2013) investigated cellulase production with free cells of T. viride in a STR and immobilized cells in a rotating fibrousbed bioreactor (RFBB). When T. viride was immobilized on solid support made up of polypropylene for enzyme production, many drawbacks associated with STR use were overcome. Drawbacks like inhibition in growth of fungi by disintegration of mycelia, fouling of the pH and temperature probes, and clogging of sampling vent could be removed by the use of RFBB. A 35.5% higher FPase activity was obtained under the operating conditions employed using immobilized mycelia in RFBB as compared to STR (Biswas et al., 2014; Lan et al., 2013). T. viride mycelia form biofilm in RFBB

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during the process under low-shear conditions. This facilitates the production of enzyme by enhancing it. “Out of the three processes, the fed-batch fermentation method has an extra advantage of utilizing the highly concentrated substrate by adding substrate and other medium components sequentially at intermittent times. This reduces the effects of catabolite repression and viscosity, which consequently will not hinder the production process. Although, fewer studies have taken place that utilized this process, as usual the substrate used for enzyme production is highly complex in nature (more often lignocellulosic biomass) (dos Reis et al., 2013). A higher FPase activity, cellulase productivity, and yield were reported using Acremonium cellulolyticus mutant CF-2612 employing batch culture with 5% Solka Floc in a 2-L jar fermenter at 30 C, which was 18.0 U/mL, 150.0 FPU/L/h, and 360.0 FPU/g carbohydrate, respectively, and when fed-batch culture was used with Solka Floc, these values reached 34.6 U/mL, 240.3 FPU/L/h, and 346.0 FPU/g carbohydrate, respectively (Fang et al., 2009). Hydrophobin II is a lowfoaming agent, which has been used for cellulase production under SmF by T. reesei strain Rut-C30 on lactose medium (Bailey and Tahtiharju, 2003). dos Reis et al. (2013) reported maximum activities of FPase (8.3 U/mL), endoglucanases (37.3 U/mL), and xylanases (177 U/mL) using Penicillium echinulatum mutant strain S1 M29 under fed-batch cultivation with 40 g/L of cellulose. Commercially, SmF is the mostly utilized method for producing enzymes because it provides advantages such as a well-developed bioreactor equipped with techniques for monitoring and control; however, the SmF method has certain drawbacks, for example, the product yield achieved could be moderate, high cost, and generate large amounts of water as waste” (Abd-Elhalem et al., 2015; Hemansi et al., 2019).

5.2.2

Solid-state fermentation

This process occurs in the absence or near-absence of free water. The production cost of many enzymes when produced using SSF process is lower as compared to that produced in SmF. The SSF technique has emerged as an attractive technology, particularly for the production of several enzymes. The SSF process is used for the production of high value added products. Enzymes are the main products which are produced using SSF by using agro-industrial wastes as substrates (Behera and Ray, 2016). It is the technique in which microorganisms grow by attaching to solid substrates. This copies the natural environment of the microorganisms, in turn giving a productive outcome. Some of the advantages of SSF are higher fermentation productivity and product stability, higher end-concentration of products,

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insignificant catabolic repression, cultivation of microorganisms specialized for the water insoluble substrates or mixed cultivation of several fungi, and reduced demand on sterility because of the low water activity used in SSF (Behera and Ray, 2016; Yoon et al., 2014). For the production of cellulases, SSF is rapidly gaining interest as a costeffective technology and also for the bioconversion of lignocellulosic material using cellulolytic microorganisms (Pandey, 1992; Pandey et al., 2000, 2001, 2007). Tengerdy (1996) reported a 10-fold reduction in the production cost using SSF as compared to SmF. Pandey et al. (1999) also reported the production of cellulases using SSF. Despite the fact, several reports on production of cellulases using SSF are available, the commercial processes are still using the SmF technology (Sukumaran et al., 2006). “At the industrial scale, fermentations have been performed in different types of reactors as per the need. Tray bioreactors and drum-type reactors in batch culture or in continuous modes are some of the processes that are commonly followed at large scale (Behera and Ray, 2016). As reported by Vitcosque et al. (2012) an instrumented lab-scale bioreactor produced 0.55 IU/g FPase, 35.1 IU/g endoglucanase, and 47.7 IU/g xylanase using A. niger as inoculum with a fully automated online monitoring and controlling system by utilizing soybean meal substrate. Brijwani et al. (2010) reported use of a tray bioreactor for producing cellulases under a mixed culture fermentation technique by utilizing T. reesei and A. oryzae grown on soybean meal and wheat bran (4:1) having a 1 cm bed height and optimum operating conditions. An appropriate C:N ratio in the substrate was maintained, which aided improved cellulase titers and brought about the production of all three enzymes in a balanced ratio, which is required for complete conversion of biomass in biofuel production. Higher levels of BGL were reported by Dhillon et al. (2011) while fermenting apple pomace by employing A. niger and T. reesei in a tray bioreactor. Although the tray bioreactor is generally exploited commercially for SSF processes in fermentation industries there are a number of shortcomings such as proper heat transfer due to low thermal conductivity and bed height (Arora et al., 2018). A. niger FTCC 5003 in a laboratory using a PBR was carried out, which used palm kernel cake as substrate in the production of cellulase for 7 days and reported cellulase yield as high as 244.53 U/gds, using 100 g of palm kernel cake (Abdeshahian et al., 2011). Cellulase production was evaluated in terms of FPase activity by Trichoderma harzianum using empty fruit bunches, and in the case of Penicillium verruculosum COKE4E alkali-treated empty palm fruit bunch fiber was used as the sole carbon source in an RDB (Alam et al., 2009; Kim and Kim, 2012).

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The engineering principles used in an RDB reactor, specifically its air circulation and continuous mixing, make it demandable for biofuel production in pilot or lab scale using cellulosic materials. An enhanced productivity of ethanol (11.7 g/L/h) was seen when cassava starch was hydrolyzed using co-immobilized glucoamylase in an SSF process with simultaneous fermentation using Streptomyces cerevisiae in pectin gel in an FBR (Trovati et al., 2009). FBR was used to develop a process for the production of high bioethanol titer through fed-batch as well as simultaneous saccharification and fermentation of wild, nonedible cassava Manihot glaziovii. This allowed fermentation of up to 390 g/L of starch-derived glucose achieving a high bioethanol concentration of up to 190 g/L (24% v/v) with yields of around 94% of the theoretical value” (Hemansi et al., 2019; Moshi et al., 2014). For SSF operations, laboratory-scale column-type bioreactors have also been used (Behera and Ray, 2016). A. niger was used for the production of endoglucanases and grown using SSF process using a laboratory-scale bioreactor which was equipped with an online automated monitoring and control system. Maximum production of endoglucanase of 56.1 U/g dry substrate was achieved under optimum conditions. The optimized parameters included substrate moisture content of 72%, flow rate of 20 mL/min, and inlet air humidity of 70%, which resulted in improved endoglucanase activity (Farinas et al., 2011). SSF requires low-cost substrate for instance lignocellulosic waste and provides a platform for better interaction of microbes and the substrate which results in higher production of enzyme. SSF can be used as a substitute for SmF based on its economic analysis in almost every feature, including raw material, equipment and final cost of the product (Ashok et al., 2017). Advantages and disadvantages of SmF and SSF are presented in Table 5.1. “In general, the advantages linked to SSF are the result of its distinctive low water content. An environment in which water is not predominant, resembles natural growing environments and creates the perfect conditions for the proliferation of microorganisms, particularly filamentous fungi. Under such conditions, microbes can conquer practically any substrate and degrade it, even when nutrients are in low quantities. Furthermore, a low water level reduces the risk of contamination and evidently, represents less water expenditure. Additionally, an important feature usually overlooked, is that less water implies a smaller bioreactor volume which results in lower capital equipment cost (Shinkawa and Mitsuzawa, 2020). Finally, in many cases, microbes can directly utilize the substrates, with minimal or no pretreatment” (Lo´pez-Go´mez et al., 2020). Cellulase production by various fungi under SSF and SmF technology is presented in Table 5.2.

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TABLE 5.1 Advantages and disadvantages of solid-state and submerged fermentation. Solid-state fermentation Advantages Higher enzyme production due to improved interaction between microorganism and substrate used. Requires low operating cost Waste generated during fermentation utilized in other processes Space required (bioreactor volume) is small Disadvantages Problem faced during mixing Complication of process monitoring and control Incapable of being employed in large-scale production Product recovery and purification is less easy Submerged fermentation Advantages Easy process monitoring and control scaled up to industrial production capacity Better parametric control, i.e., controlling temperature, pH, oxygen, and nutrient availability in the reactor Product recovery and purification is easy Disadvantages Generation of diluted products High operating cost Space required (bioreactor volume) is large Not pragmatic Reproduced with permission from Castilho et al. (2009); Costa et al. (2018); de Castro and Sato (2015); Gutierrez-Uribe (2020); Leite et al. (2020); Lo´pez-Go´mez et al. (2020); Shinkawa and Mitsuzawa (2020); Verduzco-Oliva and Gutierrez-Uribe (2020); Viniegra-Gonz´alez and Favela-Torres (2006).

5.2.3 Sequential solid-state fermentation and submerged fermentation Cunha et al. (2012) studied sequential SSF and SmF of bagasse for producing cellulase enzymes, using a fungal strain of A. niger. This approach has twofold benefits of both the techniques, as cultivation was in part under solid state for an initial growth period and in part submerged for synthesis and secretion of cellulase enzymes. The results are encouraging as the cellulase enzyme produced by the developed method was about three times more and the same was also translated to a larger scale, that is, bubble column reactor of 5 L capacity. This sequential solid and submerged fermentation approach appears a promising alternative for current production of cellulase enzyme. Fig. 5.1 shows the potential schemes for sequential SSF-SmF (Lo´pezGo´mez and Venus, 2021). “System A shows the basic process in which a single organic residue is used for both, the generation of enzymes in SSF and as the substrate for hydrolysis and subsequent SmF (for example in

TABLE 5.2 Cellulase production under submerged and solid-state fermentation. Method

Strain

Substrate

Temp ( C)

Enzyme activity (U/g)a

Time (Days)

BGL

CMCase

FPase

References

SmF

Acremonium cellulolyticus CF-2612

Wet disk-milled rice straw

30

7

12.5

197.3

5.8

Hideno et al. (2011)

SSF

Aspergillus niger (KK2)

Rice straw

28

4 6

100

129

19.5

Kang et al. (2004)

SSF

Aspergillus niger NS2

Wheat bran

30

4

33

310

17

Bansal et al. (2012)

SmP

Aspergillus terreus

Oil palm fruit (empty) fiber

30

12

10.4

14.1

0.97

Shahriarinour et al. (2011)

SSF

Fusarium chlamydosporum HML 0278

Sugarcane bagasse plus wheat bran

30

4

135.2

281.8

95.2

Qin et al. (2010)

SmF’

Penicillium brasilianum 113 T 20888

Solka-Floc cellulose

30

9.6

1.09

98

0.68

Krough et al. (2004)

SSF

Penicillium chrysogenum QML-2

Corn stover powder and wheat bran

30

5

321.6

370.15

101.8

Zhang and Sang (2012)

SmP

Trichoderma harzianum IOC-4038

Partially delignified cellulignin

30

4

0.7

0.5

0.09

de Castro et al. (2010)

SSF

Trichoderma reesei Rut-C30

Wheat bran

30

5

38.1

103

5

Pre´vot et al. (2013)

SmP

Trichoderma viride NCIM 1051

Sugarcane bagasse treated with NaC1O2

30

10

0.3

21.8

0.9

Adsul et al. (2004)

aEnzyme activity expressed in U/mL. BGL, β-glucosidases; CMCase, carboxymethyl cellulase; FPase, filter paper activity; SmF, submerged fermentation; SSF, solid-state fermentation. Source: Reproduced with permission Hemansi, Chakraborty, S., Yadav, G., Saini, J.K., & Kuhad, R.C., 2019. Comparative study of cellulase production using submerged and solid-state fermentation (chapter 7). In: Srivastava, N., Srivastava M., Mishra, P.K., Ramteke, P.W., Lakhan, S.R. (Eds.), New and Future Developments in Microbial Biotechnology and Bioengineering, Elsevier, pp. 99 113.

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System C Substrate B

SSF B

Enzyme B

Enzyme C

SSF C

Substrate C

System B Substrate A

SSF A

Hydrolysis

Enzyme A

SmF

Product

System A

FIGURE 5.1 Diagram of different schemes for sequential SSF and SmF. The color of the arrows indicates the system to which they belong (green for A, red for B, and blue for C). Lo´pez-Go´mez and Venus (2021) (https://doi.org/10.3390/fermentation7020076). This figure is distributed under the terms of the Creative Commons Attribution 4.0 International License.

the process described in (Botella et al., 2009). System B shows a process which combines two substrates, one is used in the generation of the enzyme complex which later on hydrolyzes the second substrate (as in the case described in (Sun et al., 2014). System C tries to illustrate a more complex process, in which three substrates are involved. Such process could for example use substrate A to produce enzymes able to liberate carbohydrates, for example, cellulases, substrate B could be used to produce enzymes able to liberate nitrogen, for example, proteases. Later, those enzymes could be used to hydrolyze substrate 3, producing a feedstock for SmF. Naturally, this is only one of the multiple possible configurations. As discussed earlier, the breaking down of complex polymers in organic residues requires the action of several enzymes to be more effective. Conveniently, a vast amount of the literature is already related to the production of a wide variety of enzymes from numerous organic residues. Further studies should focus on the combination of SSF processes to produce a target cocktail of enzymes able to hydrolyze effectively a specific substrate. Additionally, since the multiple enzymes could have various optimum conditions, studies on the development of optimized hydrolysis processes will be required to guarantee that the cocktail as a whole works at its best. Such appropriate selection and combination of substrates, to attain enhanced enzymatic products, together with optimized hydrolysis processes, will further benefit the sequential process” (Lo´pezGo´mez and Venus, 2021; Steudler et al., 2019). Fig. 5.2 shows different production strategies for commercial production of cellulases.

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FIGURE 5.2 Different production strategies for commercial production of cellulases (A, Off-site; B, On-site; C, Integrated; D, Consolidated) (Singhania et al., 2021). Reproduced with permission Singhania, R.R., Ruiz, A., Awasthi, Dong, M.K., Cheng-Di, C., Chiu-Wen, P., Kumar, A., 2021. Challenges in cellulase bioprocess for biofuel applications, Renew. Sustain. Energy Rev. 151, 111622.

5.2.4

Use of mixed cultures

Co-culture technique can be an alternative for increasing the production under SSF and SmF. For co-culturing two microbes, synchronization is required. “Both organisms should have similar growing temperatures and nutritional requirements (carbon source). In the case of fungi, co-culturing is quite easy as compared to that of bacteria. Fungi normally coexist under SSF because it resembles their natural habitat where fungi grow symbiotically on solid substrates (Ho¨lker et al., 2004), whereas developing bacterial cocultures is problematic because each strain has different optimum conditions for growth and carbon source. Co-culturing fungi for cellulase has several advantages such as adaptability to the surroundings, higher yield, and substrate utilization compared to that of monocultures (Dashtban et al., 2009).

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Moreover, cellulases produced from co-culture are balanced in respect of producing a complete cellulase system in an appropriate amount. T. reesei and A. niger had been grown in a co-culture under SSF as well as SmF and the enzyme produced exploited at the commercial scale (Kolasa et al., 2014). In one of the reported co-culture approach, the lack of BGL in a T. reeseiderived cellulase system was overcome by co-culturing T. reesei with better BGL producing fungi, such as A. niger or A. phoenicis (Wen et al., 2005). Brijwani et al. (2010) co-cultured T. reesei (ATCC 26921) and A. oryzae (ATCC 12892), and optimized and scaled up the production of cellulase at tray bioreactor level. The cellulases produced under this co-culture were further utilized for hydrolyzing acid- and alkali-treated wheat straw. The results displayed enhanced release of monomeric sugar up to 48 hours during hydrolysis of biomass” (Hemansi et al., 2019).

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Biswas, R., Persad, A., Bisaria, V.S., 2014. Production of cellulolytic enzymes. Bioprocess. Renew. Resour. Commod. Bioprod. 1, 105 132. Botella, C., Diaz, A.B., Wang, R., Koutinas, A., Webb, C., 2009. Particulate bioprocessing: a novel process strategy for biorefineries. Process. Biochem. 44, 546 555. Brijwani, K., Oberoi, H.S., Vadlani, P.V., 2010. Production of a cellulolytic enzyme system in mixed-culture solid-state fermentation of soybean hulls supplemented with wheat bran. Process. Biochem. 45, 120 128. Castilho, L.R., Mitchell, D.A., Freire, D.M.G., 2009. Production of polyhydroxyalkanoates (PHAs) from waste materials and by-products by submerged and solid-state fermentation. Bioresour. Technol. 100, 5996 6009. Costa, J.A.V., Treichel, H., Kumar, V., Pandey, A., 2018. Advances in solid-state fermentation. Curr. Dev. Biotechnol. Bioeng. Elsevier: Amsterdam, The Netherlands, 2018. Cunha, F.M., Esperanc¸a, M.N., Zangirolami, T.C., Badino, A.C., Farinas, C.S., 2012. Sequential solid-state and submerged cultivation of Aspergillus niger on sugarcane bagasse for the production of cellulase. Bioresour. Technol. 112, 270 274. Dashtban, M., Schraft, H., Qin, W., 2009. Fungal bioconversion of lignocellulosic residues; opportunities & perspectives. Int. J. Biol. Sci. 5, 578 595. de Castro, A.M., Pedro, K.C.N.R., da Cruz, J.C., Ferreira, M.C., Leite, S.G.F., Pereira, N., 2010. Trichoderma harzianum IOC-4038: a promising strain for the production of a cellulolytic complex with significant β-glucosidase activity from sugarcane bagasse cellulignin. Appl. Biochem. Biotechnol. 162, 2111 2122. de Castro, R.J.S., Sato, H.H., 2015. Enzyme production by solid state fermentation: general aspects and an analysis of the physicochemical characteristics of substrates for agroindustrial wastes valorization. Waste Biomass Valoriz. 6, 1085 1. Dhillon, G.S., Oberoi, H.S., Kaur, S., Bansal, S., Brar, S.K., 2011. Value-addition of agricultural wastes for augmented cellulase and xylanase production through solid-state tray fermentation employing mixed-culture of fungi. Ind. Crop. Prod. 34, 1160 1167. dos Reis, L., Fontana, R.C., da Silva Delabona, P., da Silva Lima, D.J., Camassola, M., da Cruz Pradella, J.G., et al., 2013. Increased production of cellulases and xylanases by Penicillium echinulatum S1M29 in batch and fed-batch culture. Bioresour. Technol. 146, 597 603. Esterbauer, H., Steiner, W., Labudova, I., Hermann, A., Hayn, M., 1991. Production of Trichoderma cellulase in laboratory and pilot scale. Bioresour. Technol. 36, 51 65. Fang, X., Yano, S., Inoue, H., Sawayama, S., 2009. Strain improvement of Acremonium cellulolyticus for cellulase production by mutation. J. Biosci. Bioeng. 107, 256 261. Farinas, C.S., Vitcosque, G.L., Fonseca, R.F., Neto, V.B., Couri, S., 2011. Modeling the effects of solid state fermentation operating conditions on endoglucanase production using an instrumented bioreactor. Ind. Crop. Prod. 34, 1186 1192. Ghose, T.K., Sahai, V., 1979. Production of cellulases by Trichoderma reesei QM 9414 in fedbatch and continuous flow culture with cell recycle. Biotechnol. Bioeng. 21, 283 296. Heck, J.X., Hertz, P.F., Ayub, M.A.Z., 2002. Cellulase and xylanase production by isolated amazon bacillus strains using soybean industrial residue based solid-state cultivation. Braz. J. Microbiol. 33, 213 218. Hemansi, Chakraborty, S., Yadav, G., Saini, J.K., Kuhad, R.C., 2019. Comparative study of cellulase production using submerged and solid-state fermentation (chapter 7). In: Neha, Srivastava, Manish, Srivastava, Mishra, P.K., Ramteke, P.W., Ram Lakhan Singh (Eds.), New and Future Developments in Microbial Biotechnology and Bioengineering. Elsevier, pp. 99 113. Henrissat, B., 1991. A classification of glycosyl hydrolases based on amino acid sequence similarities. Biochem. J. 280 (Pt 2), 309 316.

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Further reading Pandey, A., Francis, F., Sabu, A., Soccol, C.R., 2004. General aspects of solid-state fermentation. In: Pandey, A. (Ed.), Concise Encyclopedia of Bioresource Technology. Haworth Press, New York, USA. Zhang, X.Z., Zhang, Y.H.P., 2013. Cellulases: characteristics, sources, production, and applications. In: Yang, Shang-Tian, El-Enshasy, Hesham A., Thongchul, Nuttha (Eds.), Bioprocessing Technologies in Biorefinery for Sustainable Production of Fuels, Chemicals, and Polymers, First Edition John Wiley & Sons, Inc.

Chapter 6

Cellulase market scenario 6.1

Introduction

Bioethanol is the most important biofuel worldwide in the transport sector, with 70.5% of the total shares and 142.6 billion liters produced in 2019. The major bioethanol producers are United States and Brazil. These countries account for 84% of the total global production. The European Union ranks third; produced 5443 million liters (5% of global production) in 2019 (RFA, 2020; Duque et al., 2021). Use of lignocellulosic biomass for bioethanol production in transport sector would make the cellulase enzyme the most demanded industrial enzyme, but present applications of cellulase enzyme in industries such as food and beverages, animal feed, fermentation, pulp and paper, agriculture, textile and detergent application itself generate millions of dollars’ worth of economy. Global cellulase market is expected to increase at a significant rate as it is extensively used in several industries. It also plays a very important role in hydrolysis of cellulose. It is also considered as an abundantly available renewable biological resource and low-cost energy source. “The global Cellulase market is valued at 1677.7 million USD in 2020 is expected to reach 2450.7 million USD by the end of 2026, growing at a CAGR of 5.5% during 2021 26.” (https://www. snntv.com/story/44585038/Cellulase). Present global players producing commercial grade cellulases include Novozymes, DowDuPont, DSM, Amano Enzyme, AB Enzymes, BIO-CAT, Primalco Ltd, Zhngrng Technlgy Crpratin Ltd., Shandng Lngda Bi-Prducts, Sunsn Industry Grup, Sinbis, Cdexis. North America accounted for the highest market share in the global cellulase market in 2017 (https://www.coherentmarketinsights.com/market-insight/ cellulase-market-2146). Asia Pacific region is expected to see a high demand for cellulases. Highest growth is expected to be seen particularly in India and China. Considerable growth will also be witnessed in Middle East and Africa, Eastern Europe and Latin America. (https://www.futuremarketinsights.com/ reports/cellulase-market). “The greatest potential of cellulolytic enzymes lies in ethanol production from biomass by enzymatic hydrolysis of cellulose but low titer cellulase production resulting into high cost of the enzyme is the major setback. A number of research groups are working on improving the cellulase production technologies as well as to improve their properties so as to increase the Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00001-7 © 2023 Elsevier Inc. All rights reserved.

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efficiency of cellulose hydrolysis of available cellulases by applying various biotechnological tools, and also to screen more potent cellulases, other than the available ones. The technologies developed from lignocellulosic biomass via cellulase hydrolysis promises environmental and economical sustainability in the long run along with non dependence on nonrenewable energy source” (Patel et al., 2011). As mentioned above, microbial cellulases are being used in a several industries where cellulase enzyme of varying degrees of purity is preferred. Though cellulase enzymes were initially examined several decades back for biomass bioconversion, this later became unappealing and the other industrial applications of the enzyme in food, animal feed, pulp and paper industry, textiles, detergents were primarily pursued. But, with the scarcity of fossil fuels and the increased requirement to find alternative sources for renewable energy and fuels, there is a renewed interest in the bioconversion of lignocellulosic biomass using cellulase and other lignocellulolytic enzymes. The development of cellulases, or more correctly, cellulolytic enzyme blends is an important factor for biorefinery. Enzymes can constitute up to 30% of the cost for producing biofuels from cellulosic sugars. For achieving commercial viability, the cost of enzyme for biomass depolymerization should not exceed US$ 0.10/l of ethanol product (Champreda et al., 2019; Patel and Shah, 2021).

6.2

Market scenario

Several cellulase enzymes are available in the market under different names or trademark for a variety of applications which could be also explored for hydrolysis of biomass (Srivastava et al., 2015). Cellulases are gaining huge attention as an industrially important enzyme with a wide range of applications. This is gradually increasing its demand in industries (Srivastava et al., 2018). Significantly different cost is reported in the literature for bioethanol production. Some researchers have reported that the cost of cellulase enzyme varied from $0.1 to $0.4/gal of ethanol, supporting that the existing technology was already profitable (Sassner et al., 2008; Aden et al., 2002; Wingren et al., 2005). Cellulase enzyme cost were reported $0.10/gal (Aden and Foust, 2009), $0.30/gal (Lynd et al., 2008), $0.32/gal (Dutta et al., 2010) and $0.35/gal (Klein et al., 2010). Conversely, other researchers reported that the cost of cellulase enzyme was still as high as up $0.69/gal ethanol (Kazi et al., 2010) or even $1.47/gal ethanol (Klein et al., 2012a) by quoting the current market prices of cellulases in textile, detergent, and animal feed industries (Phitsuwan et al., 2013). Such a high cost of cellulases certainly gave a negative evaluation on the viability of commercial cellulosic ethanol production using the available technology. On the other hand, on-site or near-site production of cellulases has been proposed as a promising way for significantly reducing the enzyme cost below $0.3/gal ethanol, due to its

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simplified purification and logistics, and also the potential inexpensive use of carbon source from lignocellulosic material (Merino and Cherry, 2007). Hong et al. (2013) made a comparison of the two enzyme production methods and observed that the on-site method reduced up to 30% of the cellulase enzyme cost in comparison to the off-site method. Takimura et al. (2013) observed that up to 70% reduction of cellulase enzyme cost was achieved using the enzyme produced by on-site method in comparison to that using the purchased enzyme. Due to the differences in the cost of enzymes, particularly for biofuel applications, there are many difficulties for techno-economic assessment and commercial production processes of biofuels. For the economical production of cellulases, several factors like low enzyme loading, high substrate loading, and a short hydrolysis period are important. These factors can significantly reduce the production cost of enzymes on a commercial platform (KleinMarcuschamer et al., 2012b). Several industries are producing cellulase enzymes on a commercial scale. Leading enzyme producers are Novozymes, Dupont (Genencore), Dyadiac, DSM, etc. These companies have made important contribution to reduce the cost of cellulase enzyme by several times. Genencor and Novozymes achieved 30-fold cost reduction and an increased enzyme activity (Saini et al., 2015). Genencor has introduced Accelerase1500, which is a cellulase mixture particularly for lignocellulosic industries. It is more efficient and cost- efficient than its predecessor-Accelerase1000 for second generation bioethanol industries. It makes certain almost complete conversion of cellobiose to glucose as it contains higher amount of β glucosidase activity than other commercially available cellulases today (Singhania et al., 2010; Penttila, 1998). For improving the conversion of both xylan and glucan, Genencor has also introduced AccelleraseXY, which is an accessory xylanase enzyme complex and AccelleraseXC which is an accessory xylanase/ cellulase enzyme mixture. Another AccelleraseBG, which is an accessory β glucosidase enzyme has been also introduced for improving the glucan conversion (Dupont Accellerase, 2019; Patel and Shah, 2021). There are several potential cellulase enzymes which can hydrolyze biomass along with β-glucosidases. Novozymes has various ranges of cellulases depending on their application. Novozymes also have cellulases for hydrolysis of biomass. Besides, Amano Enzyme Inc., Japan and MAP’s India are the other enzyme companies which have actively participated in the production of cellulases. Although, several enzyme producing companies around the world have participated in production and marketing of cellulases, but only few of them have developed cellulases for biomass conversion. “Novozymes are leading for cellulases for biomass hydrolysis. It has been a common practice these days that enzyme giant collaborates or are partnered with bioethanol producing companies, for example, DupontGenencore, POET-DSM, etc. The National Renewable Energy Laboratory of

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the United States has set their goals for reducing the cost of cellulases used in bioethanol production for which projects were initiated in 2000 with Genencor Corporation and Novozymes as contract partners. Genencor announced in 2004 that it has achieved an estimated cellulase cost in the range of $0.10 $0.20 per gallon of ethanol in the cost model of NREL (Genencor, 2004). Likewise, collaborative project between Novozymes and the NREL was able to lower the cellulases price for biomass to ethanol technology approximately $0.10 0.18/gal which is declined nearly 30-fold from estimated price in 2001. Novozymes predicted that their cellulases would enable to produce cost-effective second generation bioethanol by 2010. Moreover, Novozymes further announced to set up a production facility for cellulase worth $80 100 million in Nebraska” (Novozymes Press Release, June 23, 2008). The demand for cellulases is consistently on the rise due to its diverse applications. There are several other companies also involved in cellulase production for textile detergent, paper industries, and other industries. Genencor and Novozyme have played a significant role in bringing down the cost of cellulase several folds by their active research and are continuing to bring down the cost by adopting novel technologies. Novozyme has diverse range of cellulases available based on the application as CellusoftAP and CellusoftCR for bioblasting in textile mills, Carezyme and Celluclean for laundry in detergent, Denimax 601L for stonewash industry at low temperature as well as many others specific for particular application (Novozymes Press Release, June 23, 2008). Novozyme also announced the availability of cellulase preparation specifically for biomass hydrolysis last year, although no information is available on the source of production as well as availability of enzyme in the market. In Japan, Amano Enzyme Inc. actively appeared in enzyme production, later it was placed as a leading company among global enzyme producers. Most of the total global supply of industrial cellulase is manufactured in Europe, the United States, and Japan. Today, majority of the international cellulase manufacturing enterprises is intricated in their production as well in marketing for miscellaneous applications. However, only few of them are rising with quality cellulases production especially for biomass conversion and among established companies Genencor and Novozyme are probably the leaders” (Patel et al., 2011). Table 6.1 show the commercial cellulases produced by companies and their sources. Mostly they are genetically modified strains (Patel et al., 2011). For commercialization of cellulosic ethanol, a proper cellulase enzyme with low cost and high hydrolysis performance is very important because cellulase enzyme contributes to the major portion of the cost in lignocellulosic biorefining process (Gang et al., 2016; Klein et al., 2012a). In the last few decades, enormous efforts were made by the enzyme companies worldwide to develop many high-performance cellulase enzymes. These enzymes have been introduced into the market for practical use in the

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TABLE 6.1 Commercial cellulases, their sources and suppliers. Enzyme

Supplier

Source

Cellubrix (Celluclast)

Novozymes, Denmark

T. longibrachiatum and A. niger

Celtec2

Novozymes, Denmark

T. longibrachiatum and A. niger

Celtec3

Novozymes, Denmark

T. longibrachiatum and A. niger

Novozymes 188

Novozymes, Denmark

A. niger

Cellulase 2000L

Rhodia-Danisco (Vinay, France)

T. longibrachiatum and T. reesei

Rohament CL

Rohm-AB Enzymes (Rajamaki, Finland)

T. longibrachiatum and T. reesei

Viscostar 150L

Dyadic (Jupiter, United States)

T. longibrachiatum and T. reesei

Multifect CL

Genencor Intl. (S. San Francisco, CA)

T. reesei

Bio-feed beta L

Novozymes, Denmark

T. longibrachiatum/ T. reesei

Energex L

Novozymes, Denmark

T. longibrachiatum/ T. reesei

Ultraflo L

Novozymes, Denmark

T. longibrachiatum/ T. reesei

Viscozyme L

Novozymes, Denmark

T. longibrachiatum/ T. reesei

Cellulyve

50L Lyven (Colombelles, France)

T. longibrachiatum/ T. reesei

GC 440

Genencor-Danisco (Rochester, United States)

T. longibrachiatum/ T. reesei

GC 880

Genencor

T. longibrachiatum/ T. reesei

Spezyme CP

Genencor

T. longibrachiatum/ T. reesei

GC 220

Genencor

T. longibrachiatum/ T. reesei

Accelerase1500

Genencor

T. reesei

Cellulase AP30K

Amano Enzyme

A. niger

Cellulase TRL

Solvay Enzymes (Elkhart, IN)

T. reesei/ T. longibrachiatum (Continued )

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TABLE 6.1 (Continued) Enzyme

Supplier

Source

Econase CE

Alko-EDC (New York, NY)

T. reesei/ T. longibrachiatum

Cellulase TAP106

Amano Enzyme (Troy, VA)

T. viride

Biocellulase TRI

Quest Intl. (Sarasota, FL)

T. reesei/ T. longibrachiatum

Biocellulase A

Quest Intl.

A. niger

Ultra-Low Microbial (ULM)

Iogen (Ottawa, Canada)

T. reesei/ T. longibrachiatum

Source: Reproduced with permission Patel, A.K., Pandey, A., Singhania, R.R., 2011. Production of celluloytic enzymes for the hydrolysis of lignocellulosic biomass In book: Biofuels: Alternative Feedstocks and Conversion Processes (Ed.) Singhania RR, Elsevier.

production of cellulosic ethanol (Zhang and Bao, 2017). The latest products include the CTec series and Accellerase series from Novozymes and Genencor (now part of DuPont) respectively (Chen et al., 2016; Marcos et al., 2013). Chinese industrial enzyme companies have developed several home-based cellulase enzymes which are available for use in the production of cellulosic ethanol (Liu and Wang, 2014; Zhang et al., 2015, 2016). Codexis Inc., a manufacturer of enzymes for chemical, pharmaceutical and biofuel production launched CodeXyme 4 and CodeXyme 4X cellulase enzymes. These enzymes are used for the manufacturing of cellulosic sugars for producing biofuels and biobased chemicals. CodeXyme 4 and CodeXyme 4X are used for dilute acid pretreatments and hydrothermal pretreatments respectively (https://www.coherentmarketinsights.com/market-insight/cellulase-market-2146). Novozymes has developed Cellic CTec3 enzyme which is a mixture of cellulase and hemicellulase enzyme. Cellic CTec3 allows the most cost-effective conversion of pretreated lignocellulosic materials in to fermentable sugars as compared to other cellulase or enzyme complex commercially available for production of cellulosic ethanol (Saini et al., 2015). Cellic CTec3 HS works on different substrates, including acid hydrolyzed, auto-hydrolyzed, and alkaline pretreated substrates. Normally, only Cellic CTec3 HS will be required. But, if the pretreated feedstock of interest contains a substantial amount of hemicellulose, it is desirable to combine Cellic CTec3 HS with other lignocellulolytic enzymes for further boosting overall conversion yields and potentially reduce the harshness of pretreatment. The enzyme type and dosage depends on product, substrate type, pretreatment technology, and processing conditions.

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“Historically, cellulosic enzymes were significantly affected by feedback inhibition resulting from glucose production, which led to ineffective conversion. Later generations of enzymes, including Cellic CTec3 HS, were developed to eliminate this effect on the enzyme complex. With the introduction of Cellic CTec3 HS, there are more options available for consideration when selecting optimal process layout and targeting cost-efficient cellulose conversion. These include simultaneous saccharification and fermentation (SSF), separate hydrolysis and fermentation (SHF), and a modified version of these referred to as hybrid hydrolysis and fermentation (HHF)” (www.novozymes.com).

References Aden, A., Foust, T., 2009. Technoeconomic analysis of the dilute sulfuric acid and enzymatic hydrolysis process for the conversion of corn stover to ethanol. Cellulose 16, 535 545. Aden, A., Ruth, M., Ibsen, K., Jechura, J., Neeves, K., Sheehan, J., et al., 2002. Lignocellulosic biomass to ethanol process design and economics utilizing co-current dilute acid prehydrolysis and enzymatic hydrolysis for corn stover. Natl. Renew. Energy Lab. Gold. Champreda, V., Mhuantong, W., Lekakarn, H., Bunterngsook, B., Kanokratana, P., Zhao, X.Q., et al., 2019. Designing cellulolytic enzyme systems for biorefinery: from nature to application. J. Biosci. Bioeng. Dec. 128 (6), 637 654. Chen, X.W., Kuhn, E., Jennings, E.W., Nelson, R., Tao, L., Zhang, M., 2016. DMR (deacetylation and mechanical refning). processing of corn stover achieves high monomeric sugar concentrations (230 g/L). during enzymatic hydrolysis and high ethanol concentrations (.10% v/v). during fermentation without hydrolysate purifcation or concentration. Energy Env. Sci. 9, 1237 1245. Dupont Accellerases, 2019. Cellulase Enzyme Complex for Lignocellulosic Biomass Hydrolysis. Available at: http://www.accellerase.dupont.com. ´ lvarez, C., Dome´nech, P., Manzanares, P., Moreno, A.D., 2021. Advanced bioethaDuque, A., A nol production: from novel raw materials to integrated biorefineries. Processes 9 (2), 206. Dutta, A., Dowe, N., Ibsen, K.N., Schell, D.J., Aden, A., 2010. An economic comparison of different fermentation configurations to convert corn stover to ethanol using Z. mobilis and Saccharomyces. Biotechnol. Prog. 26, 64 72. Gang, L., Jian, Z., Jie, B., 2016. Cost evaluation of cellulase enzyme for industrial scale cellulosic ethanol production based on rigorous aspen plus modeling. Bioprocess. Biosyst. Eng. 39, 133 140. Genencor, 2004. Press release, 21 October 2004, genencor celebrates major progress in the conversion of biomass to ethanol. http://genencor.com/cms/connect/genencor/media_relations/ news/archive/2004/gen_211004_en.htm. Hong, Y., Nizami, A.S., Pourbafrani, M., Saville, B.A., MacLean, H.L., 2013. Impact of cellulase production on environmental and financial metrics for lignocellulosic ethanol. Biofuels Bioprod. Bioref. 7, 303 313. Kazi, F.K., Fortman, J., Anex, R., Kothandaraman, G., Hsu, D., Aden, A., et al., 2010. Technoeconomic Analysis of Biochemical Scenarios for Production of Cellulosic Ethanol. National Renewable Energy Laboratory, Golden, CO. Klein-Marcuschamer, D., Oleskowicz-Popiel, P., Simmons, B.A., Blanch, H.W., 2012b. The challenge of enzyme cost in the production of lignocellulosic biofuels. Biotechnol. Bioeng. 109, 1083 1087.

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Klein, M.D., Oleskowicz, P.P., Simmons, B.A., Blanch, H.W., 2010. Technoeconomic analysis of biofuels: a wiki-based platform for lignocellulosic biorefineries. Biomass Bioenerg. 34, 1914 1921. Klein, M.D., Oleskowicz, P.P., Simmons, B.A., Blanch, H.W., 2012a. The challenge of enzyme cost in the production of lignocellulosic biofuels. Biotechnol. Bioengg 109, 1083 1087. Liu, S., Wang, Q., 2014. Response surface optimization of enzymatic hydrolysis process of wet oxidation pretreated wood pulp waste. Cellul. Chem. Technol. 50, 803 809. Lynd, L.R., Laser, M.S., Bransby, D., Dale, B.E., Davison, B., Hamilton, R., et al., 2008. How biotech can transform biofuels. Nat. Biotech. 26, 169 172. Marcos, M., Gonz´alez-Benito, G., Coca, M., Bolado, S., Lucas, S., 2013. Optimization of the enzymatic hydrolysis conditions of steam-exploded wheat straw for maximum glucose and xylose recovery. J. Chem. Technol. Biotechnol. 88, 237 246. Merino, S.T., Cherry, J., 2007. Progress and challenges in enzyme development for biomass utilization. Adv. Biochem. Eng. Biotechnol. 108, 95 120. Novozymes Press Release, June 23, 2008. http://www.novozymes.com/en/MainStructure/ PressAndPublications/PressRelease/2008/New 1 Facility 1 in 1 Nebraska.htm. Patel, A.K., Pandey, A., Singhania, R.R., 2011. Production of celluloytic enzymes for the hydrolysis of lignocellulosic biomass. In: Singhania, R.R. (Ed.), Biofuels: Alternative Feedstocks and Conversion Processes. Elsevier. Patel, A., Shah, A.R., 2021. Integrated lignocellulosic biorefinery: gateway for production of second generation ethanol and value added products. J. Bioresour. Bioprod. 6 (2), 108 128. Penttila, M., 1998. Heterologous protein production in Trichoderma. In: Harman, G.E., Kubicek, C.P. (Eds.), Trichoderma and Gliocladium, 2. Taylor and Francis, London, pp. 367 382. Phitsuwan, P., Laohakunjit, N., Kerdchoechuen, O., Kyu, K.L., Ratanakanokchai, K., 2013. Present and potential applications of cellulases in agriculture, biotechnology, and bioenergy. Folia Microbiol. 58, 163 176. c. RFA, 2020. RFA’s Ethanol Industry Outlook; Renewable Fuels Association: Ellisvile, MO, USA, 2020; Available online: https://ethanolrfa.org/wp-content/uploads/2020/02/2020Outlook-Final-for-Website.pdf. Saini, J.K., Saini, R., Tewari, L., 2015. Lignocellulosic agriculture wastes as biomass feedstocks for second-generation bioethanol production: concepts and recent developments. 3 Biotech. 5, 337 353. Sassner, P., Galbe, M., Zacchi, G., 2008. Techno-economic evaluation of bioethanol production from three different lignocellulosic materials. Biomass Bioenerg. 32, 422 430. Singhania, R.R., Sukumarana, R.K., Patel, A.K., Larrocheb, C., Pandey, A., 2010. Advancement and comparative profiles in the production technologies using solid-state and submerged fermentation for microbial cellulases. Enzyme Microb. Tech. 46, 541 549. Srivastava, N., Srivastava, M., Mishra, P.K., Gupta, V.K., Molina, G., Rodr´ıguez-Couto, S., et al., 2018. Applications of fungal cellulases in biofuel production: advances and limitations. Renew. Sustain. Energy Rev. 82, 2379 2386. Srivastava, N., Srivastava, M., Mishra, P.K., Singh, P., Ramteke, P.W., 2015. Application of cellulases in biofuels industries: an overview. J. Biofuels Bioenergy 1 (1), 55 63. Takimura, O., Yanagida, T., Fujimoto, S., Minowa, T., 2013. Estimation of bioethanol production cost from rice straw by on-site enzyme production. J. JPA Pet. Inst. 56, 150 155. Wingren, A., Galbe, M., Roslander, C., Rudolf, A., Zacchi, G., 2005. Effect of reduction in yeast and enzyme concentrations in a simultaneous-saccharification-and-fermentation-based bioethanol process: technical and economic evaluation. Appl. Biochem. Biotech. 122, 485 499.

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Zhang, Q., Bao, J., 2017. Industrial cellulase performance in the simultaneous saccharification and co-fermentation (SSCF). of corn stover for high-titer ethanol production. Bioresour. Bioprocess. 4, 17. Zhang, H., Han, X., Wei, C., Bao, J., 2016. Oxidative production of xylonic acid using xylose in distillation stillage of cellulosic ethanol fermentation broth by gluconobacter oxydans. Bioresour. Technol. 224, 573 580. Zhang, J., Shao, S., Bao, J., 2015. Long term storage of dilute acid pretreated corn stover feedstock and ethanol fermentability evaluation. Bioresour. Technol. 201, 355 359.

Relevant websites https://www.snntv.com/story/44585038/Cellulase. https://www.futuremarketinsights.com/reports/cellulase-market. https://www.coherentmarketinsights.com/market-insight/cellulase-market-2146. www.novozymes.com.

Chapter 7

Roles of cellulases in cellulose hydrolysis 7.1

Introduction

Cellulase enzymes (cellulolytic enzymes) are hydrolytic enzymes which catalyze the hydrolysis of β-1,4 linkages in cellulose chains. These enzymes are produced by fungi, bacteria, protozoans, plants, and animals and perform the degradation of cellulases. Cellulose exists in a number of crystalline and amorphous topologies (Schwarz, 2001; Wilson, 2008, 2011). Native cellulose is insoluble and heterogenous which makes it recalcitrant for enzymatic hydrolysis. Microorganisms meet this challenge with the help of a multienzyme system. “Given the structure of cellulose, it practically makes it impossible for the enzyme to clasp cellulose into its substrate site and hence for a single enzyme to hydrolyze cellulose. This, together with its association with other polymers, makes cellulose containing material withstand harsh conditions making it hardy and resistant to degradation, hence its role as a structural and protective barrier. Cellulose, is therefore only hydrolyzed by a variety of simultaneously acting enzymes interacting with each other to bring complete hydrolysis. Consequently, true cellulolytic organisms produce a multiple-enzyme system. These multiple-enzyme systems act in synergy to bring effective hydrolysis of cellulose” (Lakhundi et al., 2015; Leschine, 1995; Singh and Hayashi, 1995; Xie et al., 2007). Cellulase enzymes possess noncatalytic carbohydrate-binding modules (CBMs) and/or other functionally known or unknown modules, which may be located at the N- or C-terminus of a catalytic module. Complete hydrolysis of cellulose is mediated by a combination of three different types of enzyme activities (Lynd et al., 2002; Haan et al., 2007; Teeri, 1997): 1. endoglucanases (EC 3.2.1.4) (1,4-β-D-glucan- 4-glucanohydrolase or carboxymethyl cellulase) 2. exoglucanases including cellobiohydrolases (CBHs) (EC 3.2.1.91) and cellodextrinase (EC 3.2.1.74) 3. β-glucosidase (BG) (EC 3.2.1.21) For complete and efficient hydrolysis of cellulose, the cooperative action of these three cellulolytic enzyme activities are required (Fig. 7.1). These Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00011-X © 2023 Elsevier Inc. All rights reserved.

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FIGURE 7.1 Enzymes involved in cellulose degradation (Kumla et al., 2020). Kumla, J., Suwannarach, N., Sujarit, K., Penkhrue, W., Kakumyan, P., Jatuwong, K., et al., 2020. Cultivation of mushrooms and their lignocellulolytic enzyme production through the utilization of agro-industrial waste. Molecules.; 25(12):2811. Distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/).

enzymes disturb the structure at the solid liquid interface and make the individual fibers available for hydrolysis. Based on the CAZy database classification, all three cellulase enzymes consist of members in separate GH families (Lombard et al., 2014). Cellulases are classified according to the depolymerization stage of the targeting substrate (Juturu and Wu, 2014; Sharma et al., 2016; Hasunuma et al., 2013; Szijarto et al., 2008; Teter et al., 2014; Bhardwaj et al., 2021). “Endoglucanases randomly hydrolyze internal sites of a cellulose chain and subsequently produce oligosaccharides of various lengths and thus generate new chain ends. Exoglucanases act progressively on the reducing and/or nonreducing ends producing either glucose, cellobiose and/or cellooligosaccharides. These soluble cellodextrins and cellobiose are then hydrolyzed by β-glucosidases to glucose. Endoglucanases have an open active site, as they are able to bind to the interior of the long cellulose fibers. This is in contrast to exocellulase, which have

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their active site in a tunnel and hence is consistent with their processive nature resulting in sequential release of cellobiose from the end of cellulose chain. The three types of enzymes act in a coordinated manner to hydrolyze cellulose. The amorphous regions within the cellulose fibers are first attacked by endoglucanase, creating sites for exoglucanases to proceed into the crystalline regions of the fiber. They also tend to act on microcrystalline cellulose, to apparently peel the cellulose chains off its microcrystalline structure. Lastly, β-glucosidases split cellobiose to glucose preventing the build-up of cellobiose which inhibits cellobiohydrolases. Cellulolytic activity of cellulases, not only differ in the way they act on cellulose but also in the way they bind to the crystalline surface of their insoluble substrate. In fact all enzymes that act on insoluble substrates contain two binding sites: the active site which is usually contained in the catalytic domain of the enzyme and the substrate binding site which is the part of a separately folded and functionally distinct carbohydrate/cellulose binding domain. The two domains are separated by linker peptide which acts as a flexible arm connecting the two parts together. Hence the structure of most cellulases includes a cellulose binding domain (CBD) and a catalytic domain (CD)” (Lakhundi et al., 2015; Wilson, 2011; Leschine, 1995; Teeri, 1997; Beguin, 1990; Bayer et al., 1998a; Lynd et al., 2002; Henrissat et al., 1998; Shoseyov et al., 2006; Schwarz, 2001).

7.2

Cellulase enzyme systems for cellulose hydrolysis

Microorganisms producing cellulases have evolved two strategies to use their cellulases: complexed cellulases and discrete noncomplexed cellulases (Lynd et al., 2002). Generally, many aerobic cellulase producing microorganisms degrade the cellulose by producing a set of individual cellulase enzymes each of which contains a CBM joined by a flexible linker peptide to the catalytic module. CBM is located in either the C-terminus or N-terminus of the catalytic module, whereas its location does not appear to be related to its function (Wilson, 2008). Fig. 7.2 shows hydrolysis of cellulose by noncomplexed (A) and complexed (B) cellulase systems. The modular architectures of cellulases from different bacteria are presented in Table 7.1. Conversely, most anaerobic microorganisms produce large (more than 1 million molecular mass) multienzyme complexes called cellulosomes. These are generally bound to the cell surface of the microorganisms (Bayer et al., 2004). In cellulosomes, only a few of the enzymes have a CBM, but the majority of them are found to be attached to the scaffoldin protein containing a CBM. A few anaerobic bacteria are able to produce free cellulases and cellulosomes (Doi and Kosugi, 2004; Gilad et al., 2003; Berger et al., 2007). Several researchers have reviewed the function and architecture of cellulosomes (Doi et al., 1998, 2003; Doi and Tamaru, 2001; Doi and Kosugi, 2004; Bayer et al., 1998a,b, 2004; Doi, 2008). The Carbohydrate-Active Enzymes database (CAZy) provides a list of the glycoside hydrolase families which is continuously updated (Cantarel et al., 2009).

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TABLE 7.1 Modular architectures of cellulases from different bacteria. Organism

Modular architecture

GenBank code

Anaerocellum thermophilu

GH9-(CBM3)3-GH48

ACM60955

A. thermophilum

GH9-(CBM3)3-GH5

ACM60953

Bacillus subtilis

GH5-CBM3

CAA82317

Clostridium phytofermentans

GH48-Ig-CBM3

ABX43721

C. phytofermentans

GH9-CBM3-(Ig)2-CBM3

ABX43720

Clostridium thermocellum

GH48-(Doc)2

AAA23226

C. thermocellum

GH26-GH5-CBM11-(Doc) 2

AAA23225

Clostridium cellulolyticum

GH48-Doc

ACL75108

Cellulomonas fimi

GH48-Fn3-CBM2

AAB00822

Thermobifida fusca

CBM2-Fn3-GH48

AAD39947

GH, glycoside hydrolase; CBM, carbohydrate-binding module; Ig, immunoglobulin-like domain; Doc, dockerin; Fn, fi bronectin-like domain. Reproduced with permission from Zhang, X.Z., Zhang, Y.H.P., 2013. Cellulases: characteristics, sources, production, and applications. In: Yang, S.-T., El-Enshasy, H.A., Thongchul, N. (Eds.), Bioprocessing Technologies in Biorefinery for Sustainable Production of Fuels, Chemicals, and Polymers. Available from: https://doi.org/10.1002/9781118642047.ch8.

Glycoside hydrolases, including cellulases, have been classified into 115 families based on crystal structures amino acid sequence similarities. “A large number of cellulase genes have now been cloned and characterized. They are found in 13 families (1, 3, 5, 6, 7, 8, 9, 12, 26, 44, 45, 51, and 48), and cellulase like activities have been also proposed for families 61 and 74 (Schulein, 2000). Based on the data in CAZy database, the three dimensional structures of more than 50 cellulases except the family 3 cellulases are available now. All of cellulases cleave β-1,4-glucosidic bonds, but they display a variety of topologies ranging from all β -sheet proteins to β/α-barrels to all α-helical proteins. Glycoside hydrolases cleave glucosidic bonds by using acid base catalysis. The hydrolysis is performed by two catalytic residues of the enzyme: a general acid (proton donor) and a nucleophile/base. Depending on the spatial position of these catalytic residues, hydrolysis occurs via retention or inversion of the anomeric configuration. For retaining cellulases, the anomeric C bearing the target glucosidic bond retains the same (substituent) configuration after a double-displacement hydrolysis with two key glycosylation/deglycosylation steps. By contrast, for inverting cellulases, the anomeric C invert its (substituent) configuration after a single nucleophilic displacement hydrolysis. The detailed catalytic mechanisms have been well characterized and reviewed” (Zhang and Zhang, 2013; Zechel and Withers, 2000; Vocadlo and Davies, 2008; Davies and Henrissat, 1995).

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FIGURE 7.2 Schematic representation of the hydrolysis of cellulose by noncomplexed (A) and complexed (B) cellulase systems. a, cellulose; b, glucose; c, cellobiose; d, oligosaccharides; e, endoglucanase with carbohydrate-binding module (CBM); f, exoglucanase (acting on reducing ends) with CBM; g, exoglucanase (acting on nonreducing ends) with CBM; h, β-glucosidase; i, cellobiose/cellodextrin phosphorylase; j, S-layer homology module; k, CBM; l, type-I dockerin cohesion pair; m, type-II dockerin cohesin pair. The figure is not drawn to scale. Reproduced with permission Zhang, X.Z., Zhang, Y.H.P., 2013. Cellulases: characteristics, sources, production, and applications. In Bioprocessing Technologies in Biorefinery for Sustainable Production of Fuels, Chemicals, and Polymers (eds S.-T. Yang, H.A. El-Enshasy and N. Thongchul). https://doi.org/ 10.1002/9781118642047.ch8.

7.2.1

Endoglucanase

Endoglucanase, or CMCase, cleave β-1,4-bonds of cellulose chains at random, producing new ends. Diverse endoglucanases are produced by archaea, bacteria, fungi, plants, and animals having different catalytic modules belonging to families 5 to 9, 12, 44, 45, 48, 51, and 74. Bacterial endoglucanases have multiple catalytic modules, CBMs, and other modules with functions not known whereas fungal endoglucanases have a catalytic module with or without a CBM (Table 7.1). The catalytic modules of most endoglucanases possess a cleft/grove-shaped active site, which allow the endoglucanases to bind and cut the cellulose chain to produce soluble cellodextrins or insoluble cellulose fragment and glucose. But, some endoglucanases are able to act progressively based upon their ability to hydrolyze crystalline cellulose and produce the main products as cellobiose or longer cellodextrins (Mejia-Castillo et al., 2008; Yoon et al., 2008; Li and Wilson, 2008; Zverlov et al., 2005; Cohen et al., 2005; Parsiegla et al., 2008). Endoglucanases are related to the GH family-5 and are categorized as endo-acting cellulases. These can slice β-1,4- glycosidic bonds from the

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internal structure. Furthermore, they are more active on the extra soluble amorphous region of the cellulose and reduce the polymerization rate by increasing the concentration of chain ends (Tom´as-Pejo´ et al., 2009). The structure of endoglucanases can be elucidated based upon the crystalline structure of Thermoascus aurantiacus (Juturu and Wu, 2014).

7.2.2

Exoglucanase

β-1,4-Exoglucanases or cellobiohydrolases (CBHs) are the most-studied exoglucanases. These enzymes are of considerable importance to cellulolytic system and facilitate the production of mostly cellobiose which is converted into glucose by β-glucosidases. Exoglucanases act upon the reducing or nonreducing ends of cellulose polysaccharide chains. Exoglucanases are able to work on microcrystalline cellulose efficiently, most likely peel the cellulose chains from the microcrystalline structure (Teeri, 1997). Different CBHs are produced by many fungi and bacteria, with catalytic modules belonging to families 5, 6, 7, 9, 48, and 74 glycoside hydrolases. “Aerobic fungal CBHs are in families 6 and 7; aerobic bacterial CBHs are in families 6 and 48; anaerobic fungal CBHs are in family 48; and anaerobic bacterial CBHs are in family 9 and 48. In other words, family 7 CBHs only originate from fungi, and family 48 CBHs mostly originate from bacteria. The most significant topological feature of CBHs’ catalytic module is the tunnel structure which is formed by two surface loops. The tunnel may cover the entirety (e.g., family 7 CBH) or part of the active site (e.g., family 48 CBH). Fig. 7.3 shows the crystal structures of family 6 endoglucanase and exoglucanase. Although these two enzymes share the similar folding, the active site of endoglucanase is the deep cleft structure while it is a tunnel for exoglucanase. The tunnel shape active site of exoglucanase facilitate the enzyme to hydrolyze cellulose in a distinctive processive manner (Koivula et al., 2002; Vocadlo and Davies, 2008). The glycoside hydrolase family 48 exoglucanases play important roles in crystalline cellulose hydrolysis mediated by bacterial cellulase systems. Their role is believed to be somewhat similar to that of the Trichoderma CBHI (Ce17A) (Teeri, 1997; Zhang et al., 2006). Family 48 exoglucanases is dominant catalytic components of cellulosomes, such as Clostridium cellulolyticum CelF (ReverbelLeroy et al., 1997), Clostridium cellulovorans ExgS (Liu and Doi, 1998), Clostridium jusui CelD (Kakiuchi et al., 1998), and Clostridium thermocellum CelS (Kruus et al., 1995; Wang et al., 1994), or as key noncomplexed cellulase components, such as Cellulomonas fimi CbhB (Shen et al., 1995), Clostridium stercorarium Avicelase II (Bronnenmeier et al., 1991), Thermobifida fusca Cel48 (Irwin et al., 2000), Paenibacillus barcinonensis BP-23 Cel48C (S´anchez et al., 2003), and Ruminococcus albus 8 Cel48A (Devillard et al., 2004). For example, C. thermocellum CelS accounts for B30% of the weight of the cellulosome isolated from the Avicel-grown culture but its level decreases to B10% of the weight of that isolated from cellobiose-grown culture, implying that CelS plays a

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FIGURE 7.3 Crystal structures of family 6 endoglucanase and exoglucanase. (A) The structure of endoglucanase Cel6A of Thermobifida fusca (PDB code: 1TML), which exhibits a deep cleft at the active site. (B) The structure of exoglucanase Cel6A of Humicola insolens (PDB code: 1BVW), in which the active site of it bears an extended loop that forms a tunnel. Reproduced with permission Zhang, X.Z., Zhang, Y.H.P., 2013. Cellulases: characteristics, sources, production, and applications. In Bioprocessing Technologies in Biorefinery for Sustainable Production of Fuels, Chemicals, and Polymers (eds S.-T. Yang, H.A. El-Enshasy and N. Thongchul). https:// doi.org/10.1002/9781118642047.ch8.

key role for crystalline cellulose hydrolysis” (Zhang and Zhang, 2013; Zhang and Lynd, 2004; Bayer et al., 1985). Exoglucanases (cellulose 1,4-β-cellobiosidases) are of two types—cellobiohydrolase I/CBHI (EC 3.2.1.176) and cellobiohydrolase II/CBHII (EC 3.2.1.91) (Cantarel et al., 2009). Additional functions such as endo-initiating action at the reducing ends of the cellulose fibers on CBHs have also been reported (Kurasin and Valjamae, 2011; Sukumaran et al., 2021). Exoglucanases are classified as the exo-acting cellulases due to their ability to cleave β-1,4-glycosidic bonds from the region of the chain ends and having a tunnel-shaped structure at the active site. Due to this tunnel shaped structure, the re-adhering of the separated molecules to the cellulosic crystalline structure is avoided (Divne et al., 1998). It is possible to understand the in depth structure of this enzyme based on the crystalline structure of Phanerochaete chrysosporium cellobiohydrolase (Cel7A). Endoglucanases play a significant role in transforming the solid structure of cellulose into sugars while exoglucanases are responsible for its solubilization. Furthermore, post-translational modifications show that endoglucanases possess 335 amino acid long peptide chain molds into a functional enzyme having an eightfold (β/α)8 barrel architecture. Alternatively, exoglucanases possess 431 amino acid long peptide chain molds into a functional enzyme which has a 3-dimensional β-jellyroll structure. Although, these perceptions are important for understanding the basic structure of these enzymes, more studies are needed on their molecular structure analysis for further modification and improving their effectiveness.

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“Two classes of exocellulases exist: one class attacks the nonreducing end of a cellulose molecule, cleaving off cellobiose residues, and all known members of this class are in family GH-6. Members of the other class attack the reducing end of a cellulose chain, cleaving off cellobiose residues, and all aerobic fungal members of this class are in family GH-7, whereas the bacterial members are in family GH-48. The anaerobic fungal reducing end attacking exocellulases are also in family GH-48, rather than in family GH7, which is surprising because family 7 exocellulases are more active than those in family GH-48. There are a few published reports claiming that enzymes from other families are exocellulases but these reports are not well documented and they are probably endocellulases. The most studied example is CBHA from C. thermocellum, which has a crystal structure, but unlike other exocellulases, its active site is not in a tunnel. Studies have shown that CBHA is an endocellulase like most other family 9 cellulases. There are several assays, including determination of the percentage of insoluble reducing sugars that are produced, lack of reduction in the viscosity of CMC, and the determination of the products of cellohexose hydrolysis, which noticeably distinguish between exocellulase and endocellulase activity. All of them should be used to give a clear result before an enzyme is called an exocellulase” (Wilson, 2009).

7.2.3

β-Glucosidase

β-Glucosidases (BG) hydrolyze soluble cellodextrins and cellobiose to glucose. This is an important step for the effective utilization of cellulose. BGs do not have a CBM. Activity of BG on insoluble cellulose is insignificant. Different BGs are present in all kinds of organisms, including bacteria, archaea, and eukaryotes, with different catalytic modules belonging to families 1, 3 and 9. BG degrade cellobiose, which is a known inhibitor of CBH and endoglucanases. Based on either experimental data or structural homology analysis, the stereochemistry of family 9 BG is of the inverting type and families 1 and 3 BGs are of the retaining type. Aerobic fungi mostly produce extracellular BG, and anaerobic bacteria keep their BG in cytoplasm. BG has a pocket-shaped active site. This allows them to connect with the nonreducing glucose unit and cut glucose off from cellobiose or cellodextrin. The structure of BG belonging to the family GH-1 and clanGH-A can be explained based on the crystalline structure of BG-A from Bacillus polymyxa (Juturu and Wu, 2014). The action of BG on oligosaccharides is considered as the rate-limiting step of the hydrolysis reaction as it catalyzes the rupture of β- glucosidic linkages for releasing glucose (Beguin and Aubert, 1994). BG is used along with other enzymes for producing reducing sugars during biofuel production from biomass (Lin and Tanaka, 2006). BGs are the biocatalysts for the final hydrolysis reaction in breaking the cellulose- the cleavage of terminal β-D-glucosyl residues to free D-glucose

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units (Leah et al., 1995). Cellobiose- the substrate of BG, can potentially inhibit exoglucanases; so its hydrolysis is important to free CBHs from product inhibition (Teugjas and Valjamae, 2013). BG also regulates the cellulolytic machinery through feedback inhibition by its product glucose. So, buildup of glucose can result in inhibition of BG, which inhibits CBHs and EGs. Therefore, BGs are regarded as the rate-limiting enzymes in the deconstruction of cellulosic biomass. “Many aerobic fungi secrete a β-glucosidase as part of their crude cellulase, whereas most cellulolytic aerobic bacteria do not, and their β-glucosidases are usually cytoplasmic. Some organisms, mainly anaerobic bacteria, contain a cytoplasmic cellobiose phosphorylase, also called dextrin phosphorylase, which converts cellobiose and soluble dextrins to glucose and glucose-1-phosphate, conserving the energy in the cellobiose linkage” (Wilson, 2009).

7.2.4

Other cellulases and accessory proteins

In the disintegration of cellulose fiber, several accessory proteins are involved (Sukumaran et al., 2021). Swollenins are one important class of such accessory proteins. These are analogous to plant expansins as they act upon the crystalline lattices of cellulose strands and disrupt the hydrogen bonds in them (Saloheimo et al., 2002). Basically, swollenins work with other non-hydrolytic swollenin like proteins and act as zippers which open up the crosslinking in cellulose microfibrils and make looser (Arantes and Saddler, 2010). It has been reported that “this protein has hydrolytic potential and a distinctive mode of action that has parallels to the mechanism of both EGs and CBHs (Andberg et al., 2015). Aside from swollenins, other proteins with the ability to generate amorphous regions and disrupt the cellulose chains include expansins, bacterial and fungal expansin-like proteins, loosenins etc. (Georgelis et al., 2015). Expansins function by disrupting the inter and intra H-bonds that bind the cellulose fibrils and other cell wall polysaccharides, thereby creating holes in the cell wall (Arantes and Saddler, 2010; Gourlay et al., 2013). A relatively newly discovered class of oxidative enzymes that act on cellulose are the Lytic Polysaccharide Monooxygenases (LPMOs) (Vaaje- Kolstad et al., 2010). LPMOs are distributed in four auxiliary activity (AA) families in the CAZy (carbohydrate-active enzyme) database1. 2. 3. 4.

cellulose- and hemicellulose-active fungal LPMOs (AA9), chitin- and cellulose-active bacterial LPMOs (AA10), chitin-active fungal LPMOs (AA11), fungal starch-degrading LPMOs (AA13) (Forsberg et al., 2016).

Based on their mode of action, bacterial and fungal LPMOs can be classified into three groups: Type 1, enzymes which oxidize the C1 carbon of the glycosidic bond; Type 2, those that oxidize the C4 carbon of the glycosidic bond;

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Type 3, the enzymes which can oxidize either C1 or C4 (Frandsen and Lo Leggio, 2016). Since EGs preferentially bind at the amorphous regions of cellulose, the ability of LPMOs to attack crystalline cellulose allows them to complement the glycoside hydrolases by functioning as endoenzymes that cleave crystalline cellulose surfaces. These enzymes affect cellulose cleavage through an oxidative mechanism involving molecular oxygen, a reductant and an active site Cu21 ion (Payne et al., 2015). Later studies have reported that the cellobiose dehydrogenaseLPMO system improves cellulose hydrolysis when used in combination with cellulases” (Sukumaran et al., 2021; Laurent et al., 2019).

7.3

Synergy among cellulose degrading system

Synergistic cooperation between cellulases is a precondition for effective degradation of cellulose (Woodward, 1991; Kostylev and Wilson, 2012). Reese et al. (1950) put forward the first explanation of synergism more than 50 years ago. Cellulase systems show higher collective activity than the sum of the activities of individual enzymes, a phenomenon known as synergism. In both anaerobic and aerobic organisms, certain type of cellulases can act synergistically on crystalline cellulose with the specific activity of some mixtures being ten times more than that of any single cellulase in the mixture. Synergism is generally observed only in the digestion of substrates containing crystalline cellulose, perhaps as there are only a few regions in this substrate which are accessible to each cellulase and cause access to the substrate to limit the rate of hydrolysis. It appears that synergism takes place only when two cellulases attack different regions of the cellulose surface and each one generates new sites of attack for other enzymes in the mixture. “Four forms of synergism have been reported: G

G G

G

Synergism between endoglucanase and exoglucanase, called endo-exo synergy Synergism between exoglucanases, called exo-exo synergy Synergism between exoglucanase and β-glucosidase to remove cellobiose that inhibits exoglucanase Synergism between catalytic and carbohydrate-binding domains

Fujita et al. (2004) reported the synergistic hydrolysis of amorphous cellulose by a yeast strain co-displaying endoglucanase II (EGII) and cellobiohydrolase II (CBHII) from Trichoderma reesei and Aspergillus aculeatus β-glucosidase 1 (BGL1). They observed higher hydrolytic activity by the strain co-displaying EGII and CBHII (1.3 mM reducing sugar were released in 60 hours) on amorphous cellulose than the strain displaying only EGII (0.5 mM reducing sugars were released in 60 hours) with the main hydrolysis product been cellobiose. The codisplay of BGL1 along with EGII and CBHII resulted in direct production of ethanol from amorphous cellulose. Ethanol

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was not produced from amorphous cellulose in the presence of only EGII and CBHII” (Lakhundi et al., 2015). Synergism between two endoglucanases CelY and CelZ from Erwinia chrysanthemiw was reported by Zhou and Ingram (2000). Efficacy of CelY and CelZ alone and in combination of both the enzymes using carboxymethyl cellulose (CMC) and amorphous cellulose as substrates was studied. Maximal synergy (1.8-fold) was seen for enzyme combinations containing typically CelZ; the ratio of enzyme activities produced was comparable to those produced by E. chrysanthemi. CelY and CelZ were fairly dissimilar in substrate preference. CelY was not able to hydrolyze soluble cellooligosaccharides but could hydrolyze CMC to fragments averaging 10.7 glucosyl units. On the contrary, CelZ was able to readily hydrolyze cellotetraose, cellopentaose, and amorphous cellulose to produce cellobiose and cellotriose as main products. CelZ hydrolyzed CMC to products averaging 3.6 glucosyl units. Combination of CelZ and CelY hydrolyzed CMC to fragments averaging 2.3 glucosyl units. Synergy did not require the simultaneous presence of both enzymes. Treatment of the substrate with CelY, enhanced the rate and degree of hydrolysis by CelZ. Full synergy was preserved by the sequential hydrolysis of CMC, if CelY was used as the first enzyme. A general mechanism has been put forward to elucidate the synergy between these two enzymes based mainly on differences in preference of substrate. Most cellulase enzymes contain CD and CBD which function separately but perform synergistically in the disruption and hydrolysis of the cellulose. The CBD makes the substrate more accessible to hydrolytic domain. It brings the catalytic module in close proximity for increasing the hydrolysis (Lynd et al., 2002; Reese et al., 1950; Teeri et al., 1998). It also plays a role in discarding the cellulose fragments from the cellulose surface by splitting of the cross linkages. It is also important to mention that cellulose degraders always appear to produce multiple enzymes of each class. Nogawa et al. (2001) reported that cellulase system of Trichoderma reesei consists of two exoglucanases (CBHI and CBHII), five endoglucanases (EGI, EGII, EGIII, EGIV and EGV) and two β-glucosidases (BGLI and BGLII). In Humicola insolens, at least seven cellulases including two cellobiohydrolases (CBHI and CBHII) and five endoglucanases (EGI, EGII, EGIII, EGV and EGVI) are present (Schulein, 1997). In Thermobifida fusca, three endoglucanases (E1, E2 and E5), two exoglucanases (E3 and E6) and a cellulase with both endo-exo activity (E4) are present (Lynd et al., 2002; Irwin et al., 1998). The production of multiple cellulases of same class may be because of the heterogenous nature of their substrate. Since the cellulose structure varies from being crystalline to amorphous with all degrees of order in between, therefore some of these enzymes are more efficient towards one form of cellulose whereas others are more efficient towards other forms. Furthermore, it also shows that though each individual β-1,4-glucosidic bond is chemically the same, the complex nature of the substrate and the environment in which

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they are present shows that they are not in identical context. So, one may anticipate to see synergy between cellulases of same class and also of different classes. The cocktail of these enzymes thereby acts in a synergistic manner in hydrolyzing cellulose. Besides cellulases, these organisms may also possess enzymes to degrade other polysaccharides which are associated with cellulose such as hemicelluloses, most probably as their breakdown is essential to get access to cellulose fibers (Xie et al., 2007). Synergistic effect with a mixture of cel45A (an endoglucanse), cel6A (an exoglucanase) from Humicola insolens and β-glucosidase from Penicillium brasilianum on amorphous cellulose was observed by Anderson et al. (2008). But on crystalline cellulose, these two enzymes inhibited each other because of the competition for binding sites on cellulose. Study with different enzymes by Valjamae et al. (1999) and Zhang and Lynd (2004) showed synergistic effect on crystalline cellulose but not on amorphous cellulose. The highest degrees of synergy have generally been observed with highly crystalline cellulose substrates while a lower degree of synergy has been shown by cellulose having a higher content of amorphous reason. Synergism between cellulase components from the same species, acting on different substrates, has been well studied. Although, synergism between cellobiohydrolase and endoglucanase components occurs but it is not necessary that cellobiohydrolase act synergistically with all endo-glucanase components equally. For example, during solubilization of cotton fiber, cellobiohydrolase from Trichoderma koningii acted synergistically with only two out of four isolated endo- glucanase components from this species (Wood and McCrae, 1978). Generally, the degree of synergy depends on the nature of the substrate, nature of enzymes, and the assay conditions (Kumar et al., 2018; Lynd et al., 2002; Van Dyk and Pletschke, 2012; Woodward, 1991; Zhang et al., 2010). Kogo et al. (2017) studied the enzymatic hydrolysis of NH4OH-treated rice straw by enzymes from T. reesei and Humicola insolens and found significant improvement in hydrolysis due to synergistic effect among combined enzyme preparations. Enzymes from T. reesei, H. insolens, and mixture (75%:25%, v/v) of T. reesei and H. insolens showed the hydrolysis yield 70.3%, 23.5%, and 79.8%, respectively. Synergy among exoglucanases and endoglucanases is very important for complete hydrolysis of cellulose. Several studies have been performed to study the optimal ratio between exoglucanases and endoglucanases for a highest synergy degree, and so on. However, it is not easy to draw any clear conclusion because of large variations in enzyme constituents, enzyme concentration, substrate concentration, product assays, reaction time, and so on. For seeking clearness, the functionally based kinetic model for enzymatic hydrolysis of pure cellulose by individual exoglucanase/endoglucanase has been developed by Zhang and Lynd (2006). “The model represents the actions of cellobiohydrolases I, cellobiohydrolase II, and endoglucanase I; and incorporates two measurable and physically interpretable substrate parameters: the degree of polymerization (DP) and the fraction of b-glucosidic bonds accessible to cellulase, Fa (Zhang

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and Lynd, 2004). Initial enzyme-limited reaction rates simulated by the model are consistent with several important behaviors reported in the literature, including the effects of substrate characteristics on exoglucanase and endoglucanase activities; the degree of endo/exoglucanase synergy; the endoglucanase partition coefficient on hydrolysis rates; and enzyme loading on relative reaction rates for different substrates. This is the first cellulase kinetic model involving a single set of kinetic parameters that is successfully applied to a variety of cellulosic substrates, and the first that describes more than one behavior associated with enzymatic hydrolysis. The model has potential utility for data accommodation and design of industrial processes, structuring, testing, and extending understanding of cellulase enzyme systems when experimental date are available, and providing guidance for functional design of cellulase systems at a molecular scale.” Fig. 7.4 shows the simulation of the synergy between endoglucanase and exoglucanases in terms of substrate characteristics and experimental conditions (Zhang and Lynd, 2006).

FIGURE 7.4 The simulation of the synergy between endoglucanase and exoglucanases in terms of substrate characteristics (degree of polymerization, A; and accessibility, B) and experimental conditions (enzyme loading, C; and reaction time, D). Reproduced with permission Zhang, Y.H.P., Himmel, M., Mielenz, J.R., 2006. Outlook for cellulase improvement: screening and selection strategies. Biotechnol. Adv. 24: 452 481.

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To describe synergism quantitatively, the degree of synergistic effect (DSE) is mostly used. DSE is defined as the ratio of the activity of synergistic mixture to the total of the activities of individual components. “Besides the nature of synergistic components, DSE has also been shown to depend on the molar ratio of the components; surface density of bound cellulases; and properties of cellulose like DP, crystallinity, and allomorph composition. Because cellulose properties can change with hydrolysis, the dependence of DSE on hydrolysis time has also been reported. Usually, a small molar fraction (a few percent) of EG results in the highest DSE, but the optimal fraction of EG depends on the nature of the EG as well as the substrate used. Data on the effect of the surface density of bound enzymes on DSE are controversial. Some studies have shown that DSE is highest in the case of low surface coverage, whereas others support the importance of proximity (i.e., high surface density) of synergistic components. Among cellulose substrates, the highest DSE is observed on substrates with high DP and intermediate crystallinity like filter paper or bacterial cellulose, whereas low or moderate DSE is observed on highly crystalline substrates. All of these findings indicate that endo-exo synergism is far more complex than appears from the conventional, starting point generation mechanism” (Jalak et al., 2012; Valjamae et al., 1999, 2001; Medve et al., 1994, 1998; Boisset et al., 2001; Henrissat et al., 1985; Woodward et al., 1988; Zhang and Lynd, 2004; Hoshino et al., 1997; Chundawat et al., 2011; Fenske et al., 1999; Riedel and Bronnenmeier, 1998; Samejima et al., 1998).

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Reese, E.T., Siu, R.G., Levinson, H.S., 1950. The biological degradation of soluble cellulose derivatives and its relationship to the mechanism of cellulose hydrolysis. J. Bacteriol. 59, 485 497. Reverbel-Leroy, C., Pages, S., Belaich, A., Belaich, J.-P., Tardif, C., 1997. The processive endoglucanase CelF, a major component of the Clostridium cellulolyticum cellulosome: purifi cation and characterization of the recombinant form. J. Bacteriol. 179, 46 52. Riedel, K., Bronnenmeier, K., 1998. Intramolecular synergism in an engineered exo-endo-1,4_-glucanase fusion protein. Mol. Microbiol. 28, 767 775. Saloheimo, M., Paloheimo, M., Hakola, S., Pere, J., Swanson, B., Nyyss€onen, E., et al., 2002. Swollenin, a Trichoderma reesei protein with sequence similarity to the plant expansins, exhibits disruption activity on cellulosic materials. Eur. J. Biochem. 269, 4202 4211. Samejima, M., Sugiyama, J., Igarashi, K., Eriksson, K.-E.L., 1998. Enzymatic hydrolysis of bacterial cellulose. Carbohydr. Res. 305, 281 288. S´anchez, M.M., Pastor, F.I.J., Diaz, P., 2003. Exo-mode of action of cellobiohydrolase Cel48C from Paenibacillus sp. BP-23. Eur. J. Biochem. 270, 2913 2919. Schulein, M., 2000. Protein engineering of cellulases. Biochim. Biophys. Acta 1543, 239 252. Schulein, M., 1997. Enzymatic properties of cellulases from Humicola insolens. Biotechnol. 57, 71 81. Schwarz, W.H., 2001. The cellulosome and cellulose degradation by anaerobic bacteria. Appl. Microbiol. Biotechnol. 2001 (56), 634 649. Sharma, A., Tewari, R., Rana, S.S., Soni, R., Soni, S.K., 2016. Cellulases: classification, methods of determination and industrial applications. Appl. Biochem. Biotechnol. 179 (8), 1346 1380. Shen, H., Gilkes, N.R., Kilburn, D.G., Miller, R.C.J., Warren, R.A., 1995. Cellobiohydrolase B, a second exo-cellobiohydrolase from the cellulolytic bacterium Cellulomonas fi mi. Biochem. J. 311, 67 74. Shoseyov, O., Shani, Z., Levy, I., 2006. Carbohydrate binding modules: biochemical properties and novel applications. Microb. Mol. Boil. Rev. 2006 (70), 283 295. Singh, A., Hayashi, K., 1995. Microbial cellulases: protien architechture, molecular properties and biosynthesis. Adv. Appl. Microbiol. 40, 1 44. Sukumaran, R.K., Christopher, M., Kooloth-Valappil, P., Sreeja-Raju, A., Mathew, R.M., Sankar, M., et al., 2021. Addressing challenges in production of cellulases for biomass hydrolysis: targeted interventions into the genetics of cellulase producing fungi. Bioresour. Technol. Jun. 329, 124746. Szijarto, N., Siika-Aho, M., Tenkanen, M., Alapuranen, M., Vehmaanpera, J., Reczey, K., et al., 2008. Hydrolysis of amorphous and crystalline cellulose by heterologously produced cellulases of Melanocarpus albomyces. J. Biotechnol. 136 (3 4), 140 147. Teeri, T.T., Koivula, A., Linder, M., Wohlfahrt, G., Divne, C., Jones, T.A., 1998. Trichoderma reesei cellobiohydrolases: why so efficient on crystalline cellulose? Biochem. Soc. Trans. 26, 173 178. Teeri, T.T., 1997. Crystalline cellulose degradation: new insight into the function of cellobiohydrolases. Trends Biotechnol. 1997 (15), 160 167. Teter, S.A., Sutton, K.B., Emme, B., 2014. Enzymatic processes and enzyme development in biorefining. Adv. Biorefineries 199 233. Woodhead Publishing, Sawston. Teugjas, H., Valjamae, P., 2013. Selecting β-glucosidases to support cellulases in cellulose saccharification. Biotechnol. Biofuels 6 (1), 105. Tom´as-Pejo´, E., Garc´ıa-Aparicio, M., Negro, M.J., Oliva, J.M., Ballesteros, M., 2009. Effect of different cellulase dosage on cell viability and ethanol production by Kluyveromeces marxianus in SSF process. Bioresour. Technol. 100, 890 895.

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Vaaje-Kolstad, G., Westereng, B., Horn, S.J., Liu, Z., Zhai, H., Sørlie, M., et al., 2010. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science 330 (6001), 219 222. Valjamae, P., Pettersson, G., Johansson, G., 2001. Mechanism of substrate inhibition in cellulose synergistic degradation. Eur. J. Biochem. 268, 4520 4526. Valjamae, P., Sild, V., Nutt, A., Petersson, G., Johansson, G., 1999. Acid hydrolysis of bacterial cellulose reveals different modes of synergistic action between cellobiohydrolase I and endoglucanase I. Eur. Biochem. 266, 327 334. Van Dyk, J.S., Pletschke, B.I., 2012. A review of lignocellulose bioconversion using enzymatic hydrolysis and synergistic cooperation between enzymes—factors affecting enzymes, conversion and synergy. Biotechnol. Adv. 30, 1458 1480. Vocadlo, D.J., Davies, G.J., 2008. Mechanistic insights into glycosidase chemistry. Curr. Opin. Chem. Biol. 12, 539 555. Wang, W.K., Kruus, K., Wu, J.H., 1994. Cloning and expression of the Clostridium thermocellum celS gene in Escherichia coli. Appl. Microbiol. Biotechnol. 42, 346 352. Wilson, D.B., 2008. Three microbial strategies for plant cell wall degradation. Ann. NY. Acad. Sci. 1125, 289 297. Wilson, D.B., 2009. Cellulases. Encyclopedia of Microbiology. , pp. 252 258. Elsevier. Wilson, D.B., 2011. Microbial diversity of cellulose hydrolysis. Curr. Opin. Microbiol. 14, 1 5. Woodward, J., 1991. Synergism in cellulase systems. Bioresour. Technol. 36 (1), 67 75. Woodward, J., Hayes, M.K., Lee, N.E., 1988. Hydrolysis of cellulose by saturating and nonsaturating concentrations of cellulase. Implic. synergism. Nat. Biotechnol. 6, 301 304. Wood, T.M., McCrae, S.I., 1978. The cellulase of Trichoderma koningii. Purification and properties of some endoglucanase components with special reference to their action on cellulose when acting alone and in synergism with the cellobiohydrolase. Biochem. J. 171 (1), 61 72. Xie, G., Bruce, D.C., Challacombe, J.F., Chertkov, O., Detter, J.C., Gilna, P., et al., 2007. Genome sequence of the cellulolytic gliding bacterium Cytophaga hutchinsonii. Appl. Env. Microbiol. Jun. 73 (11), 3536 3546. Yoon, J.J., Cha, C.J., Kim, Y.S., Kim, W., 2008. Degradation of cellulose by the major endoglucanase produced from the brown-rot fungus Fomitopsis pinicola. Biotechnol. Lett. 30, 1373 1378. Zechel, D.L., Withers, S.G., 2000. Glycosidase mechanisms: anatomy of a fi nely tuned catalyst. Acc. Chem. Res. 33, 11 18. Zhang, Y.H.P., Himmel, M., Mielenz, J.R., 2006. Outlook for cellulase improvement: screening and selection strategies. Biotechnol. Adv. 24, 452 481. Zhang, Y.H., Lynd, L.R., 2006. A functionally based model for hydrolysis of cellulose by fungal cellulase. Biotechnol. Bioeng. 94 (5), 888 898. Zhang, Y.H., Lynd, L.R., 2004. Toward an aggregated understanding of enzymatic hydrolysis of cellulose: non-complexed cellulase systems. Biotechnol. Bioeng. 2004 (88), 797 824. Zhang, X.Z., Sathitsuksanoh, N., Zhang, Y.H., 2010. Glycoside hydrolase family 9 processive endoglucanase from Clostridium phytofermentans: heterologous expression, characterization, and synergy with family 48 cellobiohydrolase. Bioresour. Technol. 101 (14), 5534 5538. Zhang, X.Z., Zhang, Y.H.P., 2013. Cellulases: characteristics, sources, production, and applications. In: Yang, S.-T., El-Enshasy, H.A., Thongchul, N. (Eds.), Bioprocessing Technologies in Biorefinery for Sustainable Production of Fuels, Chemicals, and Polymers. Available from: https://doi.org/10.1002/9781118642047.ch8.

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Zhou, S., Ingram, L.O., 2000. Synergistic hydrolysis of carboxymethyl cellulose and acidswollen cellulose by two endoglucanases (CelZ and CleY) from Erwinia chrysanthemi. Bacteriol 182, 5676 5682. Zverlov, V.V., Schantz, N., Schwarz, W.H., 2005. A major new component in the cellulosome of Clostridium thermocellum is a processive endo-beta-1,4-glucanase producing cellotetraose. FEMS Microbiol. Lett. 249, 353 358.

Further reading Sinnott, M.L., Zou, J.-Y., Kleywegt, G.J., Szardenings, M., Sta˚hlberg, J., Jones, T.A., 2002. The active site of cellobiohydrolase Cel6A from Trichoderma reesei: the roles of aspartic acids D221 and. J. Am. Chem. Soc. 124, 10015 10024.

Chapter 8

Cellulases for biofuels production 8.1

Introduction

Depletion of fossil fuel resources and the escalating requirement of alternative sources for renewable energy have developed an enormous interest in production of cellulase enzymes (Pandey et al., 2012). Cellulolytic enzymes are finding extensive applications in the biofuel industries. Bioconversion of lignocellulosic biomass using cellulase and other enzymes are one of the important areas for the commercialization of biofuels. The enzymatic conversion of renewable lignocellulosic biomass into biofuels is an environment friendly and sustainable option to fossil-derived fuels (Srivastava et al., 2015; He et al., 2014). But, economic feasibility is one of the major constraints which is limiting its practical realization up to now. For this reason, research endeavors on bioconversion of lignocellulosic wastes into biofuels are directed towards developing the cost-effective processes which may be able to compete with the existing processes (Yuan et al., 2011; Anthony et al., 2003). Production of biofuels from lignocellulosic biomass holds great promise due to the following reasons: G G G

Plentifulness Extensive availability Low cost of cellulosic materials

“Cellulase enzymes play a very important role in the enzymatic hydrolysis of cellulosic polymers to release monomeric fermentable sugars to produce biofuels (Falkoski et al., 2013). These enzymes hydrolyze β-1,4-D-glucan linkages in the cellulose structure and produce glucose, cellobiose and cellooligosaccharides. This is the most widely studied enzymatic complex including endoglucanases (EG; EC 3.2.1.4), cellobiohydrolases (CBH; EC 3.2.1.91) and β-glucosidases (BGL; EC 3.2.1.21) (Rawat et al., 2014). Additionally, endoglucanases release nicks in the polymeric structure of cellulose, showing reducing and non-reducing ends while cellobiohydrolases produce cellooligosaccharides as well as cellobiose units by acting on reducing and non-reducing ends. Meanwhile, β-glucosidases cleave cellobiose to release monomeric sugar molecules during the hydrolysis reactions (Srivastava et al., 2018). Therefore,

Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00007-8 © 2023 Elsevier Inc. All rights reserved.

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FIGURE 8.1 Schematic diagram of the synergic action of cellulases on cellulosic biomass hydrolysis. Singh, A., Patel, A.K., Adsul, M., Singhania, R.R., 2017. Genetic modification: a tool for enhancing cellulase secretion. Biofuel Res. J. 14, 600610. Distributed under the terms and conditions of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0/).

a complete cellulase system is required for the synergic action to further convert cellulose into monomeric sugars for the effective production of biofuels. Cellulase is an enzyme of industrial significance and subsidizes around 20% overall market of enzyme around the world (Srivastava et al., 2014). Further, it is expected that the demand for this enzyme will be highly motivated by the commercial biofuel production industries in the near future (Behera and Ray, 2016). Therefore, production and efficiency of cellulase enzyme have become one of the main attentive points to be focused on an industrial scale” (Srivastava et al., 2018). Schematic diagram of the synergistic action of cellulase enzymes on cellulosic biomass hydrolysis is shown in Fig. 8.1 (Singh et al., 2017). In Fig. 8.2 the role of cellulase enzymes during the complete biofuel production process is shown and in Fig. 8.3, the schematic diagram of biofuels production from lignocellulosic biomass is presented.

8.2

Bioethanol production

Over the past decade, a lot of interest has been shown in the production of fuel ethanol due to increased environmental concern, high prices of crude oil

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Cellulase System

Endoglucanase

Cellobiohydrolase

Beta-glucosidase

Cleave cellubiose to release glucose molecules Cleaving reducing and non-reducing end of cellulose

Release celluoligosacchrides and cellubiose

Synergic action

Bioethanol Biohydrogen Hydrolysis of cellulosic biomass

Sugar released

Fermentation

FIGURE 8.2 Role of cellulases during the complete biofuel production process. Reproduced with permission Srivastava, N., Srivastava, M., Mishra, P.K., Gupta, V.K., Molina, G., Rodr´ıguez-Couto, S, et al., 2018. Applications of fungal cellulases in biofuel production: advances and limitations. Renew. Sust. Energy Rev. 82, 2379 2386.

Lignocellulosic biomass (e.g., corn stover)

OH OH O HO

HO O

Enzyme

O

O

OH OH

Fermentaon H

O

OH

Cellulose

OH

OH HO

2C2H5OH + 2CO2

OH

n

Glucose

Ethanol

FIGURE 8.3 Schematic diagram for conversion of lignocellulosic biomass into ethanol (Mensah et al., 2021). Reproduced with permission Mensah, M.B., Jumpah, H., Boadi, N.O., Awudza, J.A.M., 2021. Assessment of quantities and composition of corn stover in Ghana and their conversion into bioethanol, Sci. Afr. 12, e00731.

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and also, by the restriction in certain regions of the gasoline additive methyl tert-butyl ether (MTBE), which can be interchanged directly with ethanol (Kirk et al., 2002). “In the transport sector, bioethanol is the most important biofuel worldwide, with 70.5% of the total shares and 142.6 billion liters produced in 2019 (RFA, 2020). The United States and Brazil are the major bioethanol producers, accounting for 84% of the total production worldwide. The European Union is third in place reaching 5443 million liters (5% of global production) in 2019. Currently, fuel ethanol is mainly produced from food-related crops such as sugarcane, corn, wheat, and barley. This production has been known as first-generation (1G) bioethanol and the technology is mature at the industrial level. In 2019, 94% of the total bioethanol production in the United States was based on corn starch” (Duque et al., 2021). Bioethanol is considered a potential substitute for the conventional gasoline and can be utilized directly in vehicles or mixed with the gasoline, thus reducing emissions of greenhouse gases and use of gasoline (https://afdc. energy.gov/). “For direct application (E100), the timing (and electronic control system if in use) of the gasoline engine is adjusted, and larger gasoline tank is used. However, the use of bioethanol (E100) is usually characterized with difficulty in starting the engine at a low temperature or during the cold weather due to higher heat of vaporization required. The blending of bioethanol with gasoline might not require modifying the engine, rather it will help to enhance ignition or engine performance. The most commonly used blends are E85 and E10. Advantages of bioethanol include high-octane rating resulting to increased engine efficiency and performance, low boiling point, broad flammability, higher compression ratio and heat of vaporization, comparable energy content, reduced burning time and lean burn engine (Carrillo-Nieves et al., 2019). The disadvantages include high production cost resulting from high cost of feedstock, enzymes, detoxification and ethanol recovery, respectively. Bioethanol possesses a low volumetric energy density, meaning that more volume of bioethanol/km (up 50%) will be consumed compared to the conventional gasoline (https://afdc.energy.gov/). The use of bioethanol in engines might require frequent replacing the engine parts as the bioethanol has the capacity to degrade some elastomers and cause corrosion of metals (Dahman et al., 2019). However, in attempt to reduce the cost of production, lignocellulosic biomass is being considered as feedstocks because of availability and low cost of acquisition” (Edeh, 2020). Production of first generation bioethanol created a strong debate around the harmful impact that the widespread utilization of cereal grain in United States and Europe may have on food security on a worldwide scale. Consequently, many directives have been put into action in different areas of the world for establishing sustainability criteria aiming at protecting the use of land for food and feed and ensuring that biofuels involve measurable greenhouse gas savings as compared to fossil fuels. Furthermore, in case of European Union, “the new Directive on Renewable Energy (RED II)

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establishes that conventional food-based biofuels, including bioethanol, will be limited at a maximum of 7% of final consumption of energy in the road and rail transport sector in 2030. This scenario has paved the way for the introduction of the so-called advanced biofuels that are produced from nonfood biomass materials such as lignocellulosic feedstocks. Lignocellulosic bioethanol can be produced from the cellulosic fraction of a great variety of lignocellulosic feedstocks through the biochemical conversion pathway, which is based in hydrolysis and fermentation processes. This cellulosic ethanol is also known as second generation bioethanol or advanced ethanol” (Duque et al., 2021; Sindhu et al., 2019). Improved enzymes are being developed which can facilitate the use of inexpensive and partially utilized substrates like lignocelluloses, to make bioethanol more competitive with fossil-derived fuels. The cost of enzymes required to convert lignocelluloses into an appropriate fermentation feedstock is a main issue, and the recent work is focusing on developing enzymes with higher activity and stability and also their efficient production. In USA, Department of Energy has initiated program for supporting these developments, encouraged by the general emphasis on reduction of pollution and the necessity to work towards the fulfillment of the Kyoto protocol. There are two different methods for bioethanol production from biomass (Achinas and Euverink, 2016). 2 2

Biochemical conversion Thermochemical conversion

Both these methods conclude into fragments of lignin, hemicellulose and cellulose via degradation of lignocelluloses. Polysaccharides get hydrolyzed into sugars and then converted into ethanol (Gamage et al., 2010; Demirbas, 2007; Chandel et al., 2007). But, these conversion techniques are not same. At the present time, the biochemical method is the most commonly used (Achinas and Euverink, 2016; Fehrenbacher, 2009). The thermochemical method involves gasification of feedstock at 800 C in the presence of a catalyst (Mu et al., 2010). This method entails high level of heat and produces synthesis gas (syngas) which is a fuel gas mixture. It consist mainly hydrogen, carbon monoxide, and very often some carbon dioxide. At a temperature of 300 C, syngas can be chemically converted into a mixture of alcohols using MoS2 as the catalyst. Ethanol is separated from the mixture using distillation (US Department of Energy, Office of Energy Efficiency & Renewable Energy, Colorado, 2007). On the other hand, syngas can also be further processed into ethanol using Zymomonas mobilis, Saccharomyces cerevisiae or Clostridium ljungdahlii (Liu et al., 2014a,b; Younesi et al., 2005). Quite the opposite to the thermochemical pathway towards syngas, the biochemical method involves mild physical and/or thermochemical pretreatment, and biological pretreatment using hydrolytic enzymes to degrade

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hemicelluloses and cellulose. The physical and/or thermochemical pretreatment are mostly used to overwhelm contumacious substances and enhance the accessibility/ availability of cellulose to hemicellulase and cellulase enzymes in the biological pretreatment for producing the monomeric sugars (Yang and Wyman, 2008; US Department of Energy, Office of Energy Efficiency & Renewable Energy, Colorado, 2011). The upstream process involves hydrolysis of cellulose and breakdown of hemicelluloses into monomeric sugars. The sugars are then converted into ethanol using the fermentation process and pure ethanol is produced using the distillation process (Chandel et al., 2007; Zhu et al., 2009). Simultaneously, the recalcitrant by-product, lignin can be combusted and converted into heat and power (Gamage et al., 2010). Generally, conversion of biomass to ethanol includes the following steps (Spatari et al., 2010; S´anchez and Cardona, 2008) 1. 2. 3. 4.

Pretreatment Enzymatic hydrolysis Fermentation Distillation The processing route of bioethanol is shown in Fig. 8.4.

8.2.1

Pretreatment

Lignocellulosic biomass composed of cellulose, hemicelluloses and lignin is one of the most promising substrates for sustainable production of biofuel. The problem is a matter of how to break the bonds of this molecule for converting it into fermentable sugars. Actually, this is undeniably the type of raw material which is the most difficult to process. In the lignocellulosic biomass, lignin binds together protein, pectin, and the two types of polysaccharides, cellulose and hemicellulose. Lignin resists attack by the microorganisms and imparts strength to the plant. “The pretreatment of lignocellulosic biomass through various methods helps to release cellulose usually embedded in a matrix of polymers consisting of lignin and hemicellulose by disrupting the original structure (Fig. 8.5). With this, cellulose is separated from the polymer matrix and is more accessible for enzymatic hydrolysis, thereby resulting in increased sugar yields greater than 90% (theoretical yield) using feedstocks such as grasses, corn and wood (Ceballos, 2018). This means that cellulose is more susceptible to enzymatic hydrolysis when its crystalline structure is disrupted. Without the disruption, enzymes bind on the surface of the lignin and not the cellulose chains impeding enzymatic hydrolysis” (Edeh, 2020). Pretreatment increases the surface accessibility of polysaccharides to hydrolytic enzymes. It breaks down the lignin structure and disrupts the crystalline structure of cellulose,

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FIGURE 8.4 Processing route for bioethanol production (Raven et al., 2019). Reproduced with permission from Raven, S., Srivastava, C., Kaushik, H., Hesuh, V., Tiwari, A. (2019). Fungal Cellulases: New Avenues in Biofuel Production. In: Srivastava, M., Srivastava, N., Ramteke, P., Mishra, P. (eds) Approaches to Enhance Industrial Production of Fungal Cellulases. Fungal ˇ Leitgeb, M. (2021). Bioethanol Production by ´ K., Knez, Z., Biology. Springer, Cham and Vasic, Enzymatic Hydrolysis from Different Lignocellulosic Sources. Molecules, 26(3), 753. Distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/).

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Pretreatment

Hemicellulose FIGURE 8.5 Schematic of pretreatment effect on lignocellulosic biomass (Mosier et al., 2005a,b). Based on Mosier, N., Hendrickson, R., Brewer, M., Ho, N., Sedlak, M., Dreshel, R., 2005a. Industrial scale-up of pH-controlled liquid hot water pretreatment of corn fiber for fuel ethanol production. Appl. Biochem. Biotechnol. 125, 77 97. Mosier, N., Wyman, C.E., Dale, B. E., Elander, R., Lee, Y.Y., Holtzapple, M.T., 2005b. Features of promising technologies for pretreatment of lignocellulosic biomass. Bioresour. Technol. 96, 673 686.

so that the acids or enzymes are able to access and hydrolyze the cellulose without any difficulty (Bajpai, 2016; Achinas and Euverink, 2016). “Pretreatment can be the most expensive process in biomass-to-fuels conversion but it has great potential for improvements in efficiency and lowering of costs through further research and development. Pretreatment is an important tool for biomass-to-biofuels conversion processes. Pretreatment can be carried out using different methods such as mechanical combination, steam explosion, ammonia fiber explosion, acid or alkaline pretreatment and biological treatment, organosolv pretreatment etc. Most notably, a collaborative team called consortium for applied fundamentals and innovation (CAFI) funded by the Department of Energy and Department of Agriculture has formed and focused on several leading pretreatment technologies, including dilute (sulfuric) acid pretreatment, flow-through pretreatment, ammonia fiber expansion (AFEX), ammonia recycle percolation (ARP), and lime pretreatment for the past several years” (Bajpai, 2016; Ang et al., 2012; Brink, 1994; Cadoche and Lo´pez, 1989; Damaso et al., 2004; Gregg and Saddler, 1996; Itoh et al., 2003; Keller et al., 2003; Kim et al., 2003, 2006; Kuo and Lee, 2009a,b; Takacs et al., 2000; Wyman et al., 2009; Zhang et al., 2009, 2012, 2013a,b; Zhao and Liu, 2012; Moxley et al., 2008). Hydrolysis and downstream processing can be optimized by using an efficient pretreatment method. The basic treatment methods include physical and thermochemical processes which disrupt the recalcitrant materials and enable the cellulose to undergo hydrolysis with higher efficacy and reduced

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energy requirement (Zheng et al., 2009). The selection of pretreatment process for each feedstock should be done according to its characteristics. Treatment of agricultural biomass differs from woody biomass because of its physical properties and chemical composition (Zhu and Pan, 2010). Dissimilar to agricultural biomass, woody biomass needs higher amount of energy to reach size reduction for further enzymatic saccharification. “Toxic compounds have also to be considered for evaluating the pretreatment cost. Different substances may act as inhibitors of microorganisms that are used in the ethanol fermentation. These inhibitors include phenolic compounds, furans (furfurals and 5-HMF), aliphatic acids and inorganics compounds (iron, chromium or nickel). Several alternative measures can be taken to avoid problems caused by inhibitors. The detoxification process is an important step which can affect the pretreatment performance. General feedstock versatility and toxic inhibitors produced have to be considered on the pretreatment efficiency in order to reach optimal conditions” (Achinas and Euverink, 2016; Wyman et al., 2005; Jo¨nsson et al., 2015; Zhu and Pan, 2010; Zhu et al., 2010; Laser et al., 2002; Dey et al., 2020).

8.2.2

Hydrolysis

Enzymatic treatment is done after pretreatment to hydrolyze cellulose and hemicellulose (Pirzadah et al., 2014). Enzymatic hydrolysis is an effective approach in comparison to acidic or alkaline hydrolysis; it is environment friendly, economical and energy efficient (Taherzadeh and Karimi, 2007; Ferreira et al., 2009). Moreover, inhibitory or toxic by-products are not formed. Enzymatic hydrolysis is conducted using cellulases and hemicellulases which are exceptionally substrate specific (Banerjee et al., 2010). These enzymes degrade the lignocellulosic material into simple sugars. Optimum temperature and pH for cellulase activity are in the range of 40 C 50 C and pH 4 5 respectively (Neves et al., 2007). For xylanase, the optimum temperature is 50 C and the pH is 4 5 (Park et al., 2002). Cellulases and hemicellulases are produced by several microorganisms including fungi and bacteria. Trichoderma species is the predominant source of fungi producing industrial grade cellulolytic enzymes (endoglucanase, exoglucanase, and β-glucosidase). These enzymes act synergistically to hydrolyze the biomass (Eggeman and Elander, 2005). Several studies have shown that lignin is a source of sustainable energy and value added products. Furthermore, it was found that certain metals like Ca and Mg significantly increased enzymatic hydrolysis via lignin-metal complexation (Liu et al., 2010; Eggeman and Elander, 2005). Belkacemi and Hamoudi (2003) conducted enzymatic hydrolysis of filtrate obtained from pretreatment of corn stalks with steam using commercial enzyme containing cellulase and β-D-xylosidase from Aspergillus niger. The hydrolysis was performed using free enzymes in aqueous substrate solution

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at a pH of 5 and temperature of 30 C. About 90% saccharification was achieved using 0.12 units of cellulase per mg of dissolved solids present in the filtrate after 10 hours. Chen et al. (2008) conducted similar study. They used a mixture of T. reesei ZU-02 cellulase and A. niger ZU-07 cellobiase. The hydrolysis yield improved to 81.2% with cellobiase activity enhanced to 10 CBU/g substrate. Enzymatic hydrolysis improved by the addition of surfactant Tween-80. In fed-batch hydrolysis, the hydrolysis yield was 83.3% and the reducing sugars concentration reached 89.5 g/L. In the hydrolysate, 56.7 g/L glucose, 23.6 g/L xylose, and 5.7 g/L arabinose was found to be present, which is suitable for using in subsequent fermentation. Furthermore, there are certain adjuvants which improve the hydrolysis process. When polyethylene glycol was added, the enzymatic conversion of lignocellulosic biomass increased from 42% to 78% at 16 hours at an optimum temperature of 50 C. Hydrolysis yield increased by 7.5% when Tween 80 was added at a dose level of 5 g/L (Bo¨rjesson et al., 2007). T. reesei degraded 68.21% of rice straw pretreated with alkali while 73.96% conversion was achieved from enzymatic hydrolysis of rice straw pretreated by alkali assisted with photocatalysis technology (Xu et al.,1998). Wheat straw treated with alkaline peroxide gave 96.75% yield after enzymatic hydrolysis while atmospheric autocatalytic organosolv pretreated wet wheat straw showed more than 75% yield (Saha and Cotta, 2006). Eriksson et al. (2002) reported that hydrolysis of biomass can also be improved by the use of certain additives (surfactants and bovine serum albumin) in order to block lignin interaction with cellulase enzymes. Sewalt et al. (1997) reported that the harmful effect of lignin on cellulase enzymes can be overcome by ammonization and distinct nitrogen compounds. Cao et al. (1996) developed simultaneous approach of enzymatic treatment performed with the engineered cofermentation microbial process known as simultaneous saccharification and fermentation (SSF) for efficient hydrolysis. By the use of genetically engineered, xylose fermenting yeast Saccharomyces 1400 (pLNH33), an ethanol concentration of 47 g/L was achieved in 36 hours with 84% yield in the batch fermentation of a glucose-xylose mixture from corn cob. Furthermore, an ethanol concentration of 45 g/L was achieved in 48 hours with 86% yield using SSF process. This process gained importance during the late 1970s for its potential to reduce toxic by-product and improved bioethanol production (Bisaria and Ghose, 1981). Separate hydrolysis and fermentation (SHF) is another approach for biomass hydrolysis. This process has certain limitations. It involves the inhibition of the hydrolytic enzymes (cellulases) by saccharide products such as glucose and cellobiose which results in a slower process and a reduced yield of fermentable sugars. Unlike SHF, the SSF process keeps the sugar level very low to cause any visible inhibition of cellulases (Kumar et al., 2009; Spindler et al., 1991).

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The cellulose fraction releases glucose and the hemicellulose fraction releases pentose sugars such as xylose which is the second most abundant sugar in lignocellulosic biomass after glucose. Glucose is fermented into ethanol with ease, but for xylose, special microorganisms are required for fermentation. The second generation holds great benefits involving the fermentation of biomass in form of agricultural waste materials. However, there are some challenges to overcome like effective pretreatment and fermentation technologies along with environment friendly technology (Olsson and Hahn-Ha¨gerdal, 1996). The type of pretreatment affects the performance of the hydrolysis process (G´ırio et al., 2010; Chandel et al., 2007). There are two different types of hydrolysis processes which involve acidic reaction with sulfuric acid or enzymatic reaction. The acidic reaction can be hydrolysis with dilute acid or with concentrated acid. A high temperature of 200 C 240 C is used to disrupt cellulose crystals in dilute acid hydrolysis (Xiang et al., 2003). In case of concentrated acid hydrolysis high amount of free sugars (80%) and reduced concentrations of inhibitors are produced. So this process is more effective in comparison to dilute acid hydrolysis. But, the drawback of this process is that it needs large amount of acid (Torget et al., 1990; Hamelinck et al., 2005). When acid hydrolysis is used, the incident of chemical dehydration occurs on monosaccharides which results in the appearance of other compounds such as aldehydes (Sun and Cheng, 2002). This particular issue has driven the researchers to focus on enzymatic hydrolysis. Effective pretreatment is fundamental to an efficient enzymatic hydrolysis (Hendriks and Zeeman, 2009).

8.2.3

Fermentation

Biomass processing (pretreatment and hydrolysis) is an important step in optimizing the fermentation process (Chandel et al., 2007; Gamage et al., 2010). Pretreatment produces the saccharified biomass which can be fermented by using many potent microorganisms (Pirzadah et al., 2014). But, the main impediment in lignocellulosic biofuel technology involves the lack of potent microorganisms possessing the ability to ferment both hexose and pentose sugars (Gamage et al., 2010; Talebnia et al., 2010). An ideal microorganism should fulfill the following criteria: 1. 2. 3. 4. 5.

Wider substrate utilization Higher ethanol yield Able to bear high titre of ethanol and temperature Capability to tolerate toxic by-products or inhibitors present in hydrolysate Possess cellulolytic activity

Hexose sugars are readily fermented into ethanol but fermentation of pentose sugars is not common among microorganisms (Toivolla et al., 1984). Only a few microorganisms are able to convert pentose sugars into ethanol. The wellknown microorganism S. cereviseae is able to ferment only hexose sugars into

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ethanol. In the recent years, novel yeasts such as Pichia stipitis, Candida shehatae, and Pachysolan tannophilus have been reported which possess the ability to ferment both pentose and hexose sugars into ethanol. Thermophilic microorganisms are the ideal candidates for ethanol production on an industrial scale. These microorganisms possess the ability to withstand higher temperature and toxic by-products produces during the fermentation process. Table 8.1 shows advantages and drawbacks of potential organisms in lignocellulosic-based bioethanol fermentation. As mentioned before, the major techniques used in the fermentation of biomass hydrolysates are SSF and SHF. SSF is a superior, efficient and costeffective technique for the production of ethanol in comparison to SHF as it avoids the formation of inhibitory by-products and does not involve the use of separate reactors. But, any variation in optimum conditions of enzymes for hydrolysis and fermentation reduces its efficacy (Neves et al., 2007). Ethanol yield coefficient is found to be higher under SSF than SHF due to more conversion of pentose sugars (xylose) into xylitol (Buaban et al., 2010). Table 8.2 shows comparison between Simultaneous saccharification and fermentation and Separate hydrolysis and fermentation. Aside from SSF or SHF, there are other methods used for fermentation of biomass hydrolysates. These methods include consolidated bioprocessing (CBP) and simultaneous saccharification and cofermentation (SSCF) (Cardona and S´anchez, 2007). CBP is conducted inside a single reactor. Several steps—production of cellulase enzyme, biomass hydrolysis, and ethanol fermentation—are performed simultaneously (Bjerre et al., 1996). This process is also termed as direct microbial conversion (DMC) and involves either mono- or co-culture of microorganisms to degrade cellulose into ethanol. Furthermore, CBP method is inexpensive, easily available, and does not involve any costly inputs (Hamelinck et al., 2005). Microorganisms used for the conversion of biomass into ethanol include Fusarium oxysporum, Neurospora crassa, Paecilomyces species and Clostridium thermocellum, One of the important drawback related to this method is that its effectiveness is very low due to poor ethanol yield coefficient and longer fermentation time (3 12 days) (Szczodrak and Fiedurek, 1996). Sree et al. (1999) used S. cereviseae strain-VS3 (thermotolerant microorganisms) under SSF for production of ethanol using potato and sweet sorghum as a substrate. Now methods are focusing on developing recombination yeast which can utilize all forms of sugars thus improving ethanol production and reducing the operating costs. Two approaches are suggested in this concern: G

G

The first one involves modification of the genetic makeup of particular microorganisms and their metabolic pathways (ethanologens additional pentose metabolic pathways). Second one involves genetically engineering microorganisms in such a way so that they can ferment both pentose and hexose sugars (Dien et al., 2003).

TABLE 8.1 Advantages and drawbacks of potential organisms in lignocellulosic-based bioethanol fermentation (Garver and Liu, 2014). Species

Advantages

S. cerevisiae (Facultative anaerobic yeast)

G

G G

G

Candida shehatae (Microaerophilic yeast)

G

Drawbacks

Naturally adapted to ethanol fermentation High alcohol yield (90%) High tolerance to ethanol (up to 10% v/v) and chemical inhibitors Amenability to genetic modifications

G

Ferment xylose

G

G

G G

G

Zymomonas mobilis (Ethanologenic gramnegative bacteria)

Ethanol yield surpasses S. cerevisiae (97% of the theoretical) G

G

G

G

High ethanol tolerance (up to 14% v/v) High ethanol productivity (fivefold more than S. cerevisiae volumetric productivity) Amenability to genetic modification Does not require additional oxygen

G

G G

References

Not able to ferment xylose and arabinose sugars Not able to survive high temperature of enzyme hydrolysis

Gamage et al. (2010), Hahn-Hagerdal et al. (2007), Jorgensen (2009), McMillan (1994), Rogers et al. (2007), Talebnia et al. (2010)

Low tolerance to ethanol Low yield of ethanol Require microaerophilic conditions Does not ferment xylose at low pH

Banerjee et al. (2010), Ligthelm et al. (1988), McMillan (1994), Zaldivar et al. (2001)

Not able to ferment xylose sugars Low tolerance to inhibitors Neutral pH range

Balat and Balat (2008), Herrero (1983), Liu et al. (2010), McMillan (1994)

(Continued )

TABLE 8.1 (Continued) Species

Advantages

Pichia stipites (Facultative anaerobic yeast)

G

G G

G

Best performance xylose fermentation Ethanol yield (82%) Able to ferment most of cellulosic-material sugars including glucose, galactose, and cellobiose Possess cellulase enzymes favorable to SSF process

Drawbacks G

G

G

G

G

Pachysolen tannophilus (Aerobic fungus)

G

Ferment xylose

G G

G

Escherichia coli (Mesophilic gram-negative Bacteria)

G

G

Ability to use both pentose and hexose sugars Amenability for genetic modifications

G

G G

G G

G

References

Intolerant to a high concentration of ethanol above 40 g/L Does not ferment xylose at low pH Sensitive to chemical inhibitors Requires microaerophilic conditions to reach peak performance Reassimilates formed ethanol

Jeffries et al. (2007), McMillan (1994), Nigam (2001), Shupe and Liu (2009), Zaldivar et al. (2001)

Low yield of ethanol Require microaerophilic conditions Does not ferment xylose at low pH

Zaldivar et al. (2001), Zayed and Meyer (1996)

Repression catabolism interfere to cofermentation Limited ethanol tolerance Narrow pH and temperature growth range Production of organic acids Genetic stability not proved yet Low tolerance to inhibitors and ethanol

Gamage et al. (2010), Liu et al. (2010), Weber and Boles (2010), Zayed and Meyer (1996)

Kluveromyces marxianus (Thermophilic yeast)

G

G G G G

G

G G

G

G

Thermophilic bacteria Thermoanaerobacterium saccharolyticum Thermoanaerobacter ethanolicus Clostridium thermocellum (Extreme anaerobic bacteria)

G

G

G G G

Able to grow at a high temperature above 52 C Suitable for SSF/CBP process Reduces cooling cost Reduces contamination Ferments a broad spectrum of sugars Amenability to genetic modifications

G

Resistance to an extremely high temperature of 700C Suitable for SSCombF/CBP processing Ferment a variety of sugars Display cellulolytic activity Amenability to genetic modification

G

G G

Excess of sugars affect its alcohol yield Low ethanol tolerance Fermentation of xylose is poor and leads mainly to the formation of xylitol

Banat et al. (1992), Kumar et al. (2009), Weber and Boles (2010)

Low tolerance to ethanol

Georgieva et al. (2008), Kumar et al. (2009), Lynd et al. (2002), Shaw et al. (2008), Zeikus et al. (1981)

Source: Reproduced with permission Garver, M.P., Liu, S., 2014. Development of thermochemical and biochemical technologies for biorefineries, In Bioenergy Research: Advances and Applications, Editor(s): VK. Gupta, MG Tuohy, CP Kubicek, J Saddler, F Xu, Elsevier, pp. 457 488.

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TABLE 8.2 Comparison between Simultaneous saccharification and fermentation and Separate hydrolysis and fermentation. Simultaneous saccharification and fermentation (SSF) Features and benefits Low costs Higher ethanol yields due to removal of end product inhibition of saccharification step Limitations Difference in optimum temperature conditions of enzyme for hydrolysis and fermentation Balat and Balat (2008) Separate hydrolysis and fermentation (SHF) Features and benefits Reduces the number of reactors required Each step can be processed at its optimal operating conditions Separate steps minimize interaction between the steps Limitations End product inhibition minimizes the yield of ethanol. Chance of contamination due to long period process S´anchez and Cardona (2008) Source: Based on Pirzadah et al. (2014).

Several genetically modified microorganisms such as P. stipitis NRRLY-7124 (Nigam, 2001), P. stipitis BCC15191 (Buaban et al., 2010), recombinant such as C. shehatae NCL-3501 (Abbi et al., 1996), E. coli KO11 (Takahashi et al., 2000), S. cerevisiae ATCC 26603 (Moniruzzaman, 1995) have been developed. Though several methods have been developed for improving ethanol production from lignocellulosic material but there are still some obstacles that should be addressed. These include the following: 1. Development of more effective pretreatment method for lignocellulosic materials 2. Maintaining a stable performance of genetically engineered microorganisms in industrial-scale fermentation operation (Ho et al., 1999) 3. Integration of optimal components into economical ethanol production system (Dien et al., 2003). Fermentation process involves either batch, fed batch, or continuous process depending upon several parameters such as, type of lignocellulosic hydrolysate, enzyme kinetics and economic inputs. Fermentation of hexoses by Saccharomyces cerevisiae for production of ethanol in an immobilized cell reactor has been successfully conducted (Godia et al., 1987). Talebnia et al. (2010) recommended Clostridium sp. and Thermoanaerobacter sp. (anaerobic hemophilic bacteria) for fermentation at higher temperatures. Other genetically engineered thermotolerant microorganisms included are Z. mobilis, Candida lusitanieae and K. marxianus (Bjerre et al., 1996).

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S. cerevisiae generally converts sugars into ethanol under anaerobic conditions at a temperature of 30 C. In this pathway other by-products are also produced in the form of carbon dioxide and N-based compounds. High ethanol yield is obtained from S. cerevisiae (90% of the theoretical yield) from sugars (Kumar et al., 2009; Claassen et al., 1999). S. cerevisiae, extensively used in starch and sucrose based first generation bioethanol production, is the most suitable candidate for cellulosic ethanol production because of its ability to efficiently ferment hexose under anaerobic conditions, higher tolerance to ethanol as well as inhibitory compounds present in lignocellulosic hydrolysates. However, the weakness of S. cerevisiae is that in can ferment only hexose sugars. Therefore, the interest for microorganisms acting on pentoses is increasing (Mart´ın et al., 2002). Extensive research has been conducted to develop microorganisms which can ferment pentoses and hexoses synchronously available from the hemicellulosic fraction and tolerate inhibitory conditions. In the recent years, research is focusing on efficient methods such as SSF in order to establish a consolidated bioprocessing so that hydrolysis and fermentation take place in a single reactor. This results in cost reduction. The formation of higher amount of inhibitory compounds is also avoided. Whilst, there are several microorganisms which can convert sugars to ethanol as well as the use of one microorganism appears promising for efficient fermentation, their limitation from the view point of ethanol yield, higher tolerance to chemical inhibitors and temperature is still evident in several demonstrated projects (Ladisch et al., 2010). Addition of surfactants to cellulase enzymes along with the augmentation of other enzymes such as xylanases or Lytic polysaccharide monooxygenases has been found to be an effective method for increasing the digestibility of structural carbohydrates. Use of genetically engineered S. cerevisiae and also other notable microorganisms, results in improved fermentation. These microorganisms can ferment other monomers such as xylose and withstand high temperatures which is more appropriate for SSF processes.

8.2.4

Distillation

The fermentation of monomeric sugars is generally followed by recovery of ethanol from the fermentation broth. The end-product is a mixture of ethanol water and needs further separation using a distillation process (Edeh, 2020). Fractional distillation is a very common for separating ethanol from water based on their different volatilities. The distillation column is heated and from the top of the column the ethanol is collected as it’s boiling point is lower (78.3 C) than the boiling point of water (100 C). But, the concentration of the ethanol distillate is about 92% and further dehydration is needed for obtaining 99% ethanol (Cardona and S´anchez, 2007). “Usually, the water content of the broth is reduced to approximately 0.5% by volume

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enabling the formation of anhydrous ethanol with a minimum of 99.5% by volume. This operation is constrained by the azeotropic nature of ethanolwater solution and can be carried out based on the principle of distillation (i.e., leveraging the difference in boiling point of the components of the solution). The problem with the azeotropic solution is overcome by using a separating agent which alters the relative volatility of the key component. The techniques used in the recovery of pure ethanol from the fermentation broth include adsorption distillation, azeotropic distillation, diffusion distillation, extractive distillation, vacuum distillation, membrane distillation and chemical dehydration. The conventional techniques include azeotropic distillation, liquid-liquid extraction and extractive distillation (Nitsche and Gbadamosi, 2017). Extractive distillation is the most predominantly used for large scale operations. There are some other techniques that are gaining traction for future use especially due to less energy requirement. These are pervaporation and salt distillation” (Edeh, 2020; Nagy et al., 2015). Fig. 8.6 shows the separation process of bioethanol by extractive distillation (Damayanti et al., 2021).

8.3

Factors affecting bioethanol production

Following factors affect the production of ethanol (Zabed et al., 2014): G G G G G G

Sugar concentration Temperature Fermentation time pH Inoculum size Agitation rate

“High temperature could denature the enzymes and reduce their activity. The ideal temperature for the fermentation of biomass is 20 C 35 C (Liu and Shen, 2008). Near 30 C is an optimum temperature for the free cells of S. cerevisiae although cells which are immobilized have marginally greater optimum temperature. The optimum yield of bioethanol production could be achieved using a concentration of 150 g/L (Zabed et al., 2014). The pH of the broth also affects the production of bioethanol because, it impacts on the bacterial contamination, yeast growth, fermentation rate and by-product formation. The optimum range of pH for the fermentation of the biomass using Saccharomyces cerevisiae is 4.0 5.0. When the pH is less than 4.0, a longer incubation period is required and at a pH above 5.0, ethanol concentration is significantly reduced. To optimize the yield of bioethanol, another factor to be considered is the agitation rate. The higher the agitation rate, the higher the quantity of ethanol produced. For fermentation using yeast cells, the commonly used agitation rate is 150 200 rpm. Excess agitation rate may limit the metabolic activities of the cells” (Edeh, 2020; Zabed et al., 2014).

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FIGURE 8.6 Separation process of bioethanol by extractive distillation (Damayanti et al., 2021) Damayanti, D., Supriyadi, D., Amelia, D., Saputri, D.R., Devi, Y.L.L., Auriyani, W.A., Wu, H.S. Conversion of Lignocellulose for Bioethanol Production, Applied in Bio-Polyethylene Terephthalate. Polymers 2021, 13, 2886. https://doi.org/10.3390/polym13172886. Distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/).

8.4 Important developments in the production of cellulosic ethanol First generation ethanol is produced from sugars and starchy biomass such as grain and corn. In Brazil and India, it is mainly produced from sugarcane and in the USA, it is produced from starchy biomass. The sugars and starchy biomass are generally edible in nature and therefore there is a food-vs-fuel conflict. Sugars can be directly converted into ethanol but starches are first converted to fermentable sugars with the help of enzymes from malt or molds. This process is well-known but higher prices of the raw material and the ethics about using food products for fuel are the main issues. The raw material used for production of second generation ethanol is lignocellulosic biomass such as wood, straw, and agricultural residues, which are generally available as wastes. These types of biomass are low-cost but the process technology is more complex than converting sugar and starch (Fan et al., 1987; Badger, 2002).

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Considerable investment has been made to develop the commercial processes using the thermochemical and biochemical conversion technologies for ethanol. An excellent review has been published by Johnson et al. (2010) on the status of commercial production of ethanol from lignocellulosic biomass. Burning of ethanol produced from cellulose produces 87% less emissions than burning petrol, whereas burning of ethanol from cereals produces about 28% less emissions (Nicola et al., 2011). Ethanol produced from cellulose contains 16 times the energy required to produce it, petrol only 5 times and ethanol from maize only 1.3 times. Ethanol from cellulose holds great promise because cellulose is abundantly available and the cost is relatively lower. Considerable investment into research, pilot and demonstration plants is going on for developing commercial processes using the biochemical and thermochemical conversion technologies for ethanol. “US has a target of 136,260 million liters per year (ML/yr) of renewable fuels production by 2022. This target is only achievable with a majority of this renewable fuel coming from lignocellulosic material, such as wood, corn stover, switch grass, wheat straw and purpose grown energy crops. Demonstration-scale cellulosic ethanol plants are under construction as part of the government’s objective to make cellulosic ethanol cost competitive. The plants cover a wide variety of feedstocks, conversion technologies and plant configurations to help identify viable technologies and processes for full-scale commercialization. All demonstration plants, which are sized at 10% of a commercial-scale biorefinery, are expected to be operational soon. Commercial-scale plants are in the planning stages. Demonstration and commercial plants include—Abengoa Alico, Alltech, American Energy Enterprises (AEE), Bluefire Ethanol, Coskata, Flambeau River Papers, Park Falls, Wisconsin, Fulcrum-Bioenergy, Sierra Biofuels Plant, ICM, Mascoma, The Wisconsin Rapids, Pacific Ethanol, Red Shield Environmental (RSE), The BioGasol process, Poet, Pure Energy & Raven BioFuels, Range Fuels, Verenium, Virent. Several efforts are underway in North America to commercially produce ethanol from wood and other cellulosic materials as a primary product. NREL and its partners say that the research conducted in this area is an important step toward realizing the potential of biorefineries (www.ethanol.org/ documents/6-05_Cellulosic_Ethanol.pdf). Biorefineries, analogous to today’s oil refineries, will use plant and waste materials to produce an array of fuels and chemicals—not just ethanol. Biorefineries will extend the value-added chain beyond the production of renewable fuel only. Progress towards a commercially viable biorefinery depends on the development of real-world processes for biomass conversion. With these new technologies for the production of cellulosic ethanol, its promise becomes closer to reality with each passing day. Cellulosic ethanol is on track to be cost competitive with corn-based ethanol, a development that could drive the fuel’s production, according to an industry survey conducted by Bloomberg New Energy Finance. The survey

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focused on 11 major producers in the cellulosic ethanol industry, all of which use a technique known as enzymatic hydrolysis to break down and convert the complex sugars in non-food crop matter, and a fermentation stage to convert the material into ethanol, BNEF said. Project capital expenditures, feedstock and enzymes used in the production process are still the largest costs of running a cellulosic ethanol plant, the respondents said in the survey. But technology has pushed operating costs lower” (Bajpai, 2018).

8.4.1

Case study

8.4.1.1 Project ABBK in Hugoton, Kansas, USA The Abengoa Bioenergy Biomass of Kansas (ABBK) is a company of Abengoa Bioenergy in Hugoton, Kansas. It is the third cellulosic ethanol facility and was co-funded by the US DOE to start commercial production since July 2013 (Dina et al., 2013; https://www.renewable-technology.com/ news/newsabengoa-opens-cellulosic-ethanol-plant-in-kansas-us-4410806/). This innovative process consists of steam explosion combined with biomass fractionation, fermentation and distillation for ethanol recovery. This process uses agricultural crop residues which would otherwise remain idle feedstock. Different types of agricultural residues—corn stover, wheat straw, and seasonal grasses are converted into sugars, which are fermented, distilled, and dehydrated into fuel ethanol which is shipped to distribution centers in the region. In this plant, 1100 t of dry biomass/day for bioethanol production, that is 350,000 t/a, for the annual ethanol production capacity of nearly 80,000 t are used. The biomass includes more than 80% corn stover, wheat straw and milo stubble and collected within radius of 80 km in the amount of 15% of the available potential. There are three pricing options: cash on a dry ton basis; cash (little less) plus payment tied to the Chicago Board of Trade price of ethanol; smaller amount of cash plus the nutrient replacement program provided by ash from the Hugoton plant. One of the pricing options that the farmers obtain is $15 per dry ton of biomass, whereby ABBK provides harvesting and the amount for the farmers is only revenue for the biomass (Martinov et al., 2015). ABBK’s Hugoton facility, consists of three major areas: 1. Enzymatic hydrolysis section, where corn-stover is converted to cellulosic ethanol; 2. Cogeneration section, creates steam and electricity needed to operate the integrated biorefinery facility; and 3. Wastewater treatment section, used to treat process and utility wastewater streams for water reuse and to minimize discharge. The conversion process employed by ABBK at the Hugoton site is based on a dilute acid, enzymatic hydrolysis technology designed to convert lignocellulosic biomass into cellulosic ethanol. The facility was designed to

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process approximately 930 dry tons per day (DTPD) of feedstock resulting in an annual production of 25 MMGPY of cellulosic ethanol. The carbohydrate fraction, cellulose and hemicellulose components, are converted into monomeric sugars and then converted via fermentation into cellulosic ethanol by employing the conversion technology developed at ABNT. The lignin fraction will be used as a boiler fuel in the cogeneration section. The major processing areas in the enzymatic hydrolysis section include feedstock handling; pretreatment to make the feedstock material amenable for the conversion operations; saccharification to produce monomeric sugars from the pretreated feedstocks; fermentation is used to produce cellulosic ethanol from the monomeric sugars; distillation is used to separate and concentrate ethanol to very high purity levels; and stillage area is used to recover lignin and solids from wet waste streams. The biomass cogeneration plant was designed to produce the process steam and electric power required by the facility utilizing the lignin recovered from the enzymatic hydrolysis section and with supplemented ground biomass from the feedstock handling area. The cogeneration section is capable of producing up to 21 MW of electric power which is sufficient to power itself and provide excess clean renewable power to the local Stevens County community (Leon, 2017). The wastewater treatment section has been designed to handle about 140,000 lb. per day of total chemical-oxygen-demand (COD) process effluent stream. Due to the presence of nutrients in the effluent, ABBK’s agronomist studies supported mixing of the effluent water with utility water and ultimately used for land application. This land application of the water was permitted by the State of Kansas. ABBK has contracted professional biomass harvesting and removal firms for doing all the logistics. Agronomic professionals and engineers are also hired for adjusting harvesting and transportation procedures according to the requirement. The harvesters cut, rake, bale and transport the material from the fields. Contract-farmer has no expenditure associated with the removal of biomass. Alternately, if farmer has the manpower, equipment and time to harvest and store from the contracted fields, ABBK pays the farmer for this. The up to date facility also has an electricity cogeneration component which allows it to operate as a self-sufficient renewable energy producer. By using residual biomass solids from the ethanol conversion process, the plant produces 21 megawatts of electricity which is enough to power itself and provide excess clean renewable power to the local Stevens County community. US Department of Energy provided $132.4 million loan and $97 million grant for the construction of this LCB plant. As compared to gasoline, the LCB produced at ABBK enables GHG emission reduction of more than 60%. ABBK created approximately 300 construction jobs, and is expected to support 65 permanent jobs. It will cut 132,000 metric tons of carbon emissions annually—equivalent to taking 28,000 cars off the road.

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8.4.1.2 Clariant sunliquid plant Podari, Romania Innovative specialty chemical company Clariant based in Switzerland has successfully developed an efficient and economic process for the production of cellulosic ethanol—the Sunliquid process. Clariant has completed the construction of first commercial Sunliquid plant for the production of cellulosic ethanol from agricultural residues in Podari, Romania (Figs. 8.7 8.9). The EUR 140 million facility (about $162 million USD) will process approximately 250,000 tons of straw to produce approx. 50,000 tons of cellulosic ethanol per annum. The plant in Podari, Dolj county, is built on a 10-hectare area on which construction was initiated in 2019 with up to 800 workers onsite. Contracts with more than 300 local farmers have been signed to ensure the supply of the necessary feedstock. The sun liquid process converts agricultural residues into cellulosic ethanol, a second-generation biofuel, also

FIGURE 8.7 Clariant completes construction of first commercial Sunliquid cellulosic ethanol plant in Podari, Romania (https://www.clariant.com). Reproduced with permission.

FIGURE 8.8 Clariant’s flagship commercial Sunliquid cellulosic ethanol plant in Romania. (https://www.clariant.com). Reproduced with permission.

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FIGURE 8.9 Sunliquid plant Podari (https://www.clariant.com). Reproduced with permission.

referred to as advanced biofuel. This process is designed for industrial plants with a production capacity of 50,000 to 150,000 ton of cellulosic ethanol per year. Five license agreements have already been signed in Slovakia, Poland, Bulgaria and two in China. Sunliquid process is divided into four technological steps (Figs. 8.10 and 8.11) (Hortsch and Corvo, 2020): The straw is first shredded and thermally pretreated to open the stable lignocellulose structure and make it easier for the enzymes to access the sugar chains. Conventional processes use chemicals during the pretreatment. With the sun liquid technology, however, the pretreatment process has been optimized so that no chemicals are needed. This process enables optimal enzymatic hydrolysis immediately afterward. Purification processes are no longer required, costs for chemicals are saved, and the process is made safer and more environmentally friendly. The straw fibers are divided into two batches: special microorganisms are applied to a tiny fraction and use the straw as a nutrient source. The organisms rapidly produce large quantities of feedstockand process-specific enzymes—these act as a kind of biological scissors, which break up the long cellulose chains. The enzymes are then added back to the central part of the raw material. They liquefy the straw and split its cellulose and hemicellulose components into the following sugar types: glucose, xylose, and arabinose. The insoluble lignin, the woody part of the straw, is extracted and burned to generate energy that is returned to the process.

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FIGURE 8.10 The Sunliquid process for the production of cellulosic ethanol from agricultural residues (https://www.clariant.com). Reproduced with permission.

FIGURE 8.11 Sunliquid—a fully integrated process design (https://www.clariant.com). Reproduced with permission.

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Together with other energy-rich by-products, the lignin is the most important source of energy for the overall energy-efficient process. Specially developed fermentation organisms are added to the resulting sugar solution, which simultaneously converts all types of sugar to ethanol and ensures an optimal yield during fermentation and the utilization of all associated by-products. In the final step, the water is removed with the help of an optimized, highly energyefficient distillation and purification method. Pure ethanol can now be further processed into biofuel or other chemicals. The only other by-product left is the vinasse. Until now, enzymes have been the most expensive component in production costs. This is precisely where the real innovative character of the sun liquid technology becomes apparent. In the sun liquid process, costs are reduced to a minimum through process-integrated enzyme production directly onsite in the production plant. The enzymes are produced at the place and time where they are needed. There are no costs for transport, storage, and processing, and there is no dependence on enzyme suppliers. Because they are individually adapted to the raw material used and the prevailing conditions, the process is also much more efficient than with standard enzymes. It can be flexibly applied to the respective regionally available raw materials, from wheat straw to corn straw to sugarcane bagasse and energy crops, such as switchgrass. With these optimized enzymes, the process achieves high sugar yields. Besides, the process can convert not only glucose—an easily degradable C6 sugar—but also xylose and arabinose, which are more challenging to use C5 sugars (Hortsch and Corvo, 2020).

Enzyme production facilities form an integral part of the Sunliquid plant. No additional substrate costs are incurred as the enzymes are directly produced on a small portion of the pretreated raw material. Moreover, there are no extra costs for formulation and logistics and producers are no longer dependent on external enzyme suppliers. Until now, enzyme costs constitute the major part of the process costs. Using integrated enzyme production, these are significantly reduced and as a result, the costs involved in the Sunliquid process are linked mostly to the raw materials. “A Sunliquid plant with an annual capacity of 50,000 tons requires approximately 227,000 tons of straw. Based on 10% land use, this gives rise to an area of some 4700 km2 or a collection radius for straw of a good 35 km. This is enough to operate a fleet of 62,500 vehicles (approximately the number of gas-powered vehicles in Germany) with an average mileage, yet with almost zero CO2 emission in its total balance—and without any additional investment in new infrastructures” (https://sunliquid-project-fp7. eu/wp-content/uploads/2014/09/factsheet_sunliquid_en.pdf). The bioethanol produced by the sun liquid process helps decarbonize the transport sector by providing up to 95% CO2 savings compared to fossil fuel, and by as much as 120% if carbon sequestration is considered and used as part of the production process. In other words, a sun liquid plant with an

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output of 50,000 tons per year like the Podari plant can help avoid B120,000 tons of CO2 emissions per year, equivalent to the annual CO2 emissions of B35,000 cars. The sun liquid process is a prime example of a circular economy solution. Instead of being discarded or burned on the fields, agricultural residues are utilized without any associated environmental risks. The bioethanol produced replaces fossil resources and has the potential to be used as building block for future production of biobased chemicals. Fig. 8.12 shows Sunliquid technology platform for the production of sustainable biobased products. The technology has been sold to 5 countries (Fig. 8.13). Fig. 8.14 shows reduction of carbon dioxide emissions with Sunliquid technology. Fig. 8.15 shows that Sunliquid fully integrated process saves B95% GHG emissions. Fig. 8.16 shows Clariant Bioethanol Pilot Plant, Straubing, Germany.

FIGURE 8.12 Sunliquid technology platform for the production of sustainable biobased products. Reproduced with permission Hortsch, R., Corvo, P., 2020. The biorefinery concept: producing cellulosic ethanol from agricultural residues. Chem. Ing. Tech. 92 (11), 1803 1809. 14

Media Conference Call - sunliquid Plant Construction Completed 15.10.2021

sunliquid® technology: Innovation leadership with 5 licenses sold globally 1st EU License Slovakia

2nd China License China

1G 2G

Sino-Dan



Enviral: Slovakia’s biggest EtOH producer



Sino-Dan Jianye: major Chinese agriculture company



Plant capacity: 50,000 tpa cellulosic ethanol



Plant capacity: 25,000 tpa cellulosic ethanol



Raw material: wheat straw



Raw material: corn stover

2nd EU License Poland

Oil 2G



ORLEN Południe: Poland's largest petroleum company



Plant capacity: 25,000 tpa cellulosic ethanol



Raw material: wheat straw

3rd EU License Bulgaria

ETA BIO

2G

1st China License China

Guozhen



Eta Bio Ltd: leading Bulgarian agricultural company



Plant capacity: 50,000 tpa cellulosic ethanol



Licensee: Anhui Guozhen Group & Chemtex Chemical Engineering



Raw material: wheat straw



Plant capacity: 50,000 tpa cellulosic ethanol



Raw material: wheat straw, corn stover

FIGURE 8.13 Sun liquid technology: five licenses sold globally (https://www.clariant.com). Reproduced with permission.

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Circular thinking drives innovation in biofuels 15.10.2021

sunliquid® technology helps to significantly reduce the CO2 emissions in the transport sector EXAMPLE CALCULATION FOR ONE 50KTA CELLULOSIC ETHANOL PLANT INPUT DATA

95%

CALCULATED SAVINGS*

Plant size:

50,000 tons Ethanol/year

Average emission sunliquid® bioethanol:

4.7 g CO2/MJ

Fossil fuel comparator (default REDII):

94 g CO2/MJ

CO2 Reduction

~120,000 tons CO2/year Saving ~35,000 cars emission per year

>95% CO2 savings of sunliquid® biofuel compared to gasoline

* Assumptions: The average fuel economy for new 2017 model year cars, light trucks and SUVs in the United States was 24.9 mpgUS (9.4 L/100 km), 10.000 km/year driving; 8.7 tonsCO2/capita/year in EU27 (Eurostat) Reference: "Highlights of the Automotive Trends Report". US EPA. March 2019. Retrieved 23 June 2019.

FIGURE 8.14 Reduction of Carbon dioxide emissions (https://www.clariant.com). Reproduced with permission.

1

3

2 Straw

Enzymes & Microorganisms

ENZYME PRODUCTION ORGANISM

4

5

Fermentation organisms

FERMENTATION

PURIFICATION

BIOETHANOL

SACCHARIFICATION

PRE-TREATMENT

LIGNIN

C5 & C6 SUGARS

CO2

VINASSE

WATER

FIGURE 8.15 Sunliquid: fully integrated process saves B95% GHG emissions (https://www. clariant.com). Reproduced with permission.

8.4.1.3 Praj’s second generation (2G) cellulosic ethanol plant Praj’s state of the art second generation ethanol pilot plant facility is operational since 2009. This facility has tested more than 450 MT of biomass such as corn cob, cane bagasse, corn Stover, Empty fruit bunches (EFB), Rice straw, etc. Rigorous testing and 800,000 man-hours of technology

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Demonstration Plant Straubing Capacity: 1,000 t/a EtOH; ~4,500 t/a feedstock (with scaled-down commercial design reproducing all process steps) FIGURE 8.16 Clariant Bioethanol Pilot Plant, Straubing, Germany (https://www.clariant.com). Reproduced with permission.

development efforts enabled us to scale the “Enfinity” to 1 million liters per annum capacity. PRAJ’s End to-end 2G “Smart Bio refinery” solution based on “Enfinity” technology will process multiple feed stocks and shall produce multiple products like bio ethanol, bio butanol, bio chemicals, power, Bio CNG, CO2 etc.

8.5

Other biofuels made by assistance from enzymes

“Biofuels include products made via sustainable processing; substantiated by reducing the need for energy from fossil fuel, obtaining better production efficiencies and reducing environmental impact. Biodiesel is an example of such a product having combustion properties like petro-diesel. Biogas is a renewable energy source resulting from biomass—mainly waste products

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from industrial or agricultural production. A biorefinery is a facility that integrates biomass conversion processes and equipment to produce fuels, power, and value-added chemicals from biomass. Enzymatic catalysis is needed as the way to a sustainable, selective and mild production technique. Biodiesel is methyl or ethyl esters of fatty acids made from renewable biological resources: vegetable oils or animal fat. The esters are typically made by catalytic reactions of free fatty acids (FFA) or triglycerides with alcohols, preferably methanol or ethanol. The overall reaction is a sequence of consecutive and reversible reactions, in which diglycerides and monoglycerides are formed as intermediate compounds. The complete stoichiometric reaction requires 1 mol of triglycerides and 3 mol of alcohol. The reaction is reversible and therefore excess alcohol is used to shift the equilibrium to the products’ side. Methanol and ethanol are frequently used in the process. Transesterification as an industrial process is generally carried out by heating an excess of the alcohol under different reaction conditions in the presence of an acid or a base, or by heterogeneous catalysts such as metal oxides or carbonates, or by a lipase enzyme. The biodiesel yield in the transesterification process is affected by process parameters like moisture, content of free fatty acids (FFAs), reaction time, reaction temperature, catalyst type and molar ratio of alcohol to oil” (Bajpai, 2018).

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(Eds.), Recent Advances in Bioenergy Research, III. Sardar Swaran Singh National Institute of Renewable Energy, Kapurthala, India, pp. 121 129. Srivastava, N., Rawat, R., Oberoi, H.S., Ramteke, P.W., 2015. A review on fuel ethanol production from lignocellulosic biomass. Int. J. Green Energy 12, 949 960. Srivastava, N., Srivastava, M., Mishra, P.K., Gupta, V.K., Molina, G., Rodr´ıguez-Couto, S., et al., 2018. Applications of fungal cellulases in biofuel production: advances and limitations. Renew. Sust. Energy Rev. 82, 2379 2386. Sun, Y., Cheng, J., 2002. Hydrolysis of lignocellulosic materials for ethanol production: A review. Bioresour Technol 83, 1 11. Szczodrak, J., Fiedurek, J., 1996. Technology for conversion of lignocellulosic biomass to ethanol. Biomass Bioenergy 10 (5), 367 375. Taherzadeh, M.J., Karimi, K., 2007. Acid based hydrolysis processes for ethanol from lignocellulosic materials: a review. BioResources 2 (3), 472 499. Takacs, E., Wojnarovits, L., Foldavary, C., Hargagittai, P., Borsa, J., Sajo, I., 2000. Effect of combined gamma-irradiation and alkali treatment on cotton cellulose. Radiat. Phys. Chem. 57, 339. Takahashi, C.M., Lima, K.G.C., Takahashi, D.F., Alterthum, F., 2000. Fermentation of sugarcane bagasse hemicellulosic hydrolysate and sugar mixtures to ethanol by recombinant Escherichia coli KO11. World J. Microbiol. Biotechnol. 16, 829 834. Talebnia, F., Karakashev, D., Angelidaki, I., 2010. Production of bioethanol from wheat straw: an overview on pretreatment, hydrolysis and fermentation. Bioresour. Technol. 101 (13), 4744 4753. Toivolla, A., Yarrow, D., Van-den-bosch, E., Van-dijken, J.P., Sheffers, W.A., 1984. Alcoholic fermentation of D-xylose by yeasts. Appl. Microbiol. Biotechnol. 47, 1221 1223. Torget, R.W., Werdene, P.J., Himmel, M.E., Grohmann, K., 1990. Dilute acid pretreatment of short rotation woody and herbaceous crops. Appl. Biochem. Biotechnol. 24 25, 115 126. U.S. Department of Energy, Office of Energy Efficiency & Renewable Energy, Colorado, 2007. National Renewable Energy Laboratory (NREL) Thermochemical ethanol via indirect gasification and mixed alcohol synthesis of lignocellulosic biomass. Technical report NREL/TP510 41168. U.S. Department of Energy, Office of Energy Efficiency & Renewable Energy, Colorado, 2011. National Renewable Energy Laboratory (NREL) Process design and economics for biochemical conversion of lignocellulosic biomass to ethanol: Dilute-acid pretreatment and enzymatic hydrolysis of corn stover. Technical report NREL/TP-5100 47764. ˇ Leitgeb, M., 2021. Bioethanol production by enzymatic hydrolysis from difVasi´c, K., Knez, Z., ferent lignocellulosic sources. Molecules 26 (3), 753. Weber, C., Boles, E., 2010. Sugar-hungry yeast to boost biofuel production. Sci. News Sci. Dly. 92, 881 882. Wyman, C.E., Dale, B.E., Elander, R.T., Holtzapple, M., Ladisch, M.R., Lee, Y.Y., 2005. Coordinated development of leading biomass pretreatment technologies. Bioresour. Technol. 96, 1959 1966. Wyman, C.E., Dale, B.E., Elander, R.T., 2009. Comparative sugar recovery and fermentation data following pretreatment of poplar wood by leading technologies. Biotech. Prog. 25, 333 339. Xiang, Q., Lee, Y.Y., Pettersson, P.O., Torget, R.W., 2003. Heterogeneous aspects of acid hydrolysis of α-cellulose. In: Davison, B.H., Lee, J.W., Finkelstein, M., McMillan, J.D. (Eds.), Biotechnology for Fuels and Chemicals. Applied Biochemistry and Biotechnology. Humana Press, Totowa, NJ, pp. 505 514.

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Xu, J., Takakuwa, N., Nogawa, M., Okada, H., Morikawa, Y., 1998. A third xylanase from Trichoderma reesei PC-3 7. Appl. Microbiol. Biotechnol. 49, 18 724. Yang, B., Wyman, C.E., 2008. Preteatment: the key to unlocking low-cost cellulosic ethanol. Biofuels Bioprod. Biorefin. 2, 26 40. Younesi, H., Najafpour, G., Mohamed, A.R., 2005. Ethanol and acetate production from synthesis gas via fermentation processes using anaerobic bacterium. Clostridium ljungdahlii Biochem. Eng. J. 27, 110 119. Yuan, P., Meng, K., Huang, H., Shi, P., Luo, H., Yang, P., et al., 2011. A novel acidic and lowtemperature- active endo-polygalacturonase from Penicillium sp. CGMCC 1669 with potential for application in apple juice clarification. Food Chem. 129, 1369 1375. Zabed, H., Faruq, G., Sahu, J.N., Azirun, M.S., Hashim, R., Boyce, A.N., 2014. Bioethanol production from fermentable sugar juice. Sci. World J. 2014, 1 11. Zaldivar, J., Nielsen, J., Olsson, L., 2001. Fuel ethanol production from lignocellulose: a challenge for metabolic engineering and process integration. Appl. Microbiol. Biotechnol. 56, 17 34. Zayed, G., Meyer, O., 1996. The single-batch bioconversion of wheat straw to ethanol employing the fungus Trichoderma viride and the yeast Pachysolen tannophytus. Appl. Microbiol. Biotechnol. 45, 551 555. Zeikus, J., Ben-Bassat, A., Ng, T., Lamed, R., 1981. Thermophilic ethanol fermentation. Basic Life Sci 18, 441 461. Zhang, S., Marechal, F., Gassner, M., Perin-Levasseur, Z., Qi, W., Ren, Z., et al., 2009. Process modelling and integration of fuel ethanol production from lignocellulosic biomass based on double acid hydrolysis. Energy Fuels . Zhang, Z., O’Hara, I.M., Doherty, W.O.S., 2012. Pretreatment of sugarcane bagasse by acid-catalysed process in aqueous ionic liquid solutions. Bioresour. Technol. 120, 149 156. Zhang, T., Kumar, R., Wyman, C.E., 2013a. Sugar yields from dilute oxalic acid pretreatment of maple wood compared to those with other dilute acids and hot water. Carbohydr. Polym. 92, 334 344. Zhang, D.S., Yang, Q., Zhu, J.Y., Pan, X.J., 2013b. Sulfite (SPORL) pretreatment of switchgrass for enzymatic saccharification. Bioresour. Technol. 129, 127 134. Zhao, X., Liu, D., 2012. Fractionating pretreatment of sugarcane bagasse by aqueous formic acid with direct recycle of spent liquor to increase cellulose digestibility-the Formiline process. Bioresour. Technol. 117, 25 32. Zheng, Y., Pan, Z., Zhang, R., 2009. Overview of biomass pretreatment for cellulosic ethanol production. Int. J. Agric. Biol. Eng. 2, 51 68. Zhu, J.Y., Pan, H.J., 2010. Woody biomass pretreatment for cellulosic ethanol production: technology and energy consumption evaluation. Bioresour. Technol. 101, 4992 5002. Zhu, J.Y., Wang, G.S., Pan, X.J., Gleisner, R., 2009. Specific surface to evaluate the efficiencies of milling and pretreatment of wood for enzymatic saccharification. Chem. Eng. Sci. 64, 474 485. Zhu, J.Y., Xuejun, P., Zalesny, R.S., 2010. Pretreatment of woody biomass for biofuel production: energy efficiency, technologies, and recalcitrance. Appl. Microbiol. Biotechnol. 87, 847 857.

Relevant websites http://www.ethanol.org/documents/6-05_Cellulosic_Ethanol.pdf. https://www.clariant.com.

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https://www.renewable-technology.com/news/newsabengoa-opens-cellulosic-ethanol-plant-inkansas-us-4410806/. https://sunliquid-project-fp7.eu/wp-content/uploads/2014/09/factsheet_sunliquid_en.pdf. https://www.abengoa.com/web/en/novedades/hugoton/imagenes/. https://www.afdc.energy.gov/uploads/publication/ethanol_basics.pdf; 2015.

Further reading Abdi, B., 2019. Indian oil to invest Rs 700 crore in setting up 2G ethanol plant at Panipat refinery. ETEnergyWorld. https://energy.economictimes.indiatimes.com/news/oil-and-gas/indianoil-to-invest-rs-700-crore-in-setting-up-2g-ethanol-plant-at-panipatrefinery/69872795. Adhanom, B.T., 2019. Sugarcane bagasse-based bioethanol producing biorefinery plant simulation and techno-economic analysis (master’s thesis). Addis Ababa University, http://etd.aau. edu.et/handle/123456789/19345. Bayrakcı, A.G., Kocar, G., 2014. Second-generation bioethanol production from water hyacinth and duckweed in Izmir: a case study. Renew. Sust. Energy Rev. 30, 306 316. Brown, A., Waldhelm, L., Landalv, I., Saddler, J., Edadian, M., McMillan, J.D., et al., 2020. Advanced biofuels—potential for cost reduction. Chaturvedi, V., Verma, P., 2013. An overview of key pretreatment processes employed for bioconversion of lignocellulosic biomass into biofuels and value added products. Biotech 3, 415 431. Coughlin, K., Fridley, D., 2008. Physical Energy Accounting in California: A Case Study of Cellulosic Ethanol Production. United States. https://doi.org/10.2172/953684.https://www. osti.gov/servlets/purl/953684. Du, R., Huang, R., Su, R., Zhang, M., Wang, M., Yang, J., et al., 2013. Enzymatic hydrolysis of lignocellulose: SEC-MALLS analysis and reaction mechanism. RSC Adv 3, 1871 1877. Ethanol Basics (Fact Sheet). Clean cities, energy efficiency & renewable energy (EERE), 2015. https://www.afdc.energy.gov/uploads/publication/ethanol_basics.pdf. Ganguli, S., Somani, A., Motkuri, R.K., Bloyd, C., 2018. India alternative fuel infrastructure: the potential for second-generation biofuel technology. U.S. Department of Energy’s Office of Scientific and Technical Information. https://www.osti.gov/servlets/purl/1530891/. Gnansounou, E., Dauriat, A., 2010. Techno-economic analysis of lignocellulosic ethanol: a review. Bioresour. Technol. 101 (13), 4980 4991. Available from: https://doi.org/10.1016/j. biortech.2010.02.009. Hassan, S.S., Williams, G.A., Jaiswal, A.K., 2018. Lignocellulosic biorefineries in Europe: current state and prospects. Trends Biotechnol . Available from: https://doi.org/10.1016/j. tibtech.2018.07.002. Haus, S., Bjo¨rnsson, L., Bo¨rjesson, P., 2020. Lignocellulosic ethanol in a greenhouse gas emission reduction obligation system—a case study of swedish sawdust based-ethanol production. Energies 13 (5), 1048. Liu, S., 2008. A kinetic model on autocatalytic reactions in woody biomass hydrolysis. J. Biobased Mater. Bioenergy 2 (5), 135 147. Liu, S., 2010. Woody biomass: niche position as a source of sustainable renewable chemicals and energy and kinetics of hot-water extraction/hydrolysis. J. Biotech. Adv. 28, 563 582. Mupondwa, E., Li, X., Tabil, L., 2017. Large-scale commercial production of cellulosic ethanol from agricultural residues: a case study of wheat straw in the Canadian Prairies. Biofuels. Bioprod. Bioref. 11, 955 970. Available from: https://doi.org/10.1002/bbb.1800.

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TATA project win order for bioethanol plant in India. https://biofuels-news.com/news/tata-projects-wins-order-for-bioethanol-plantin-india/. Zhou, Y., Searle, S., Anup, S., 2021. Techno-economic analysis of cellulosic ethanol in India using agricultural residues, International Council on Clean Transportation. https://theicct. org/sites/default/files/publications/cellulosic-ethanol-analysis-india-jun2021.pdf.

Chapter 9

Advanced developments in production processes of cellulases 9.1

Introduction

Cellulase enzymes are one of the extensively used industrial enzymes which are available on a commercial scale for more than 40 years. These are induced enzymes produced by several microorganisms including fungi and bacteria during their growth on cellulosic substrates. Aerobic as well as anaerobic microorganisms are found to produce these enzymes, but aerobic cellulolytic fungi, viz., Trichoderma viride and T. reesei, are widely used. Cellulase enzymes are presently the third largest industrial enzyme worldwide, by dollar volume. For several decades, cellulase enzymes have played an important role as biocatalysts. They are being used in many industries - textile, paper and pulp, laundry and detergent, agriculture, medicine, food and feed industry, brewing and agriculture (Patel et al., 2019; Jayasekara and Ratnayake, 2019). Cellulase enzymes are expected to become the biggest volume industrial enzyme, if ethanol, butanol, or some other fermentation product of sugars, produced from biomass by enzymes, become a main transportation fuel. According to Global Cellulase (CAS 9012-54-8) Market Research Report published in 2018, the application of cellulase enzymes in industries is significantly rising. Asia-Pacific is the largest user of cellulase enzymes. In 2016, the cellulase market demand was 29.71% in animal feed, 26.37% in food and beverages, and 13.77% in the textile industry. During the 2018 25 period, applications of cellulase enzymes is expected to reach 2300 million USD by 2025, growing at a compound annual growth rate of 5.5%. Novozymes and DuPont, major producers of cellulase enzymes are supplying these enzymes to the global market for various commercial applications (Jayasekara and Ratnayake, 2019). Almost all the industrial cellulase enzymes are produced from aerobic cellulolytic fungi (Schulein, 1998). “This is due to the ability of engineered strains of these organisms to produce exceptionally large amounts of crude cellulase (over 100 g/L), the comparatively higher specific activity of crude cellulase on crystalline cellulose, and the ability to genetically modify these strains to tailor Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00012-1 © 2023 Elsevier Inc. All rights reserved.

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the set of enzymes they produce, so as to give optimal activity for specific applications. The DOE has been supporting industrial research to reduce the cost of cellulase enzymes for production of bioethanol for a number of years” (Wilson, 2009b). Bioconversion of lignocellulosic materials using cellulase and other enzymes is the key area for the commercialization of biofuels (Wilson, 2009a; Magrey et al., 2018). Many researchers around the world are constantly working in this area. Higher cost and lower production have always been a main limitation which should be overcome by the use of novel and multipurpose strategies. In this background, use of low-priced raw materials as the substrates, use of genetically modified microorganisms, use of efficient crude thermostable/thermophilic enzymes are some of the major factors which can significantly improve the production of cellulases (Srivastava et al., 2015a). Presently, many countries have implemented policies regarding production of cellulosic ethanol and have set targets to shift the biomass resource from sugar or starchy substrates to lignocellulosic substrates (Ejaz et al., 2021).

9.2 Advanced development in production processes of cellulases There is a huge demand of enzymes which are more active and stable at higher temperature and substrate-specific (Vaishnav et al., 2018; Singh et al., 2017). Thermostable enzymes are able to catalyze reaction under harsh conditions found in industrial processing. So these enzymes offer robust catalyst. “Use and development of molecular biology techniques, permitting genetic analysis and gene transfer for recombinant production, led to dramatically increased activities in the field of thermostable enzymes during the 1990. This also stimulated isolation of a number of microbes from thermal environments in order to access enzymes that could significantly increase the window for enzymatic bioprocess operations. The greatest potential of cellulolytic enzymes lies in ethanol production from biomass by enzymatic hydrolysis of cellulose but low thermostability and low titer of cellulase production resulting into high cost of the enzyme is the major set-back. A number of research groups are working on cellulase to improve its thermostability so as to be able to perform hydrolysis at elevated temperatures which would eventually increase the efficiency of cellulose hydrolysis. The technologies developed from lignocellulosic biomass via cellulose hydrolysis promise environmental and economical sustainability in the long run along with non-dependence on nonrenewable energy source” (Patel et al., 2019). Use of thermostable enzymes using a variety of microorganisms for complete or partial hydrolysis of cellulose has been examined (Srivastava et al., 2015a). These researchers have reported the utilization of thermostable

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cellulases over cellulases for efficient hydrolysis. For bioprocessing industries, thermostable cellulases are regarded as best candidates as the hydrolysis of lignocellulosic materials with thermostable cellulases increases the rate of reaction, the rate of bioavailability of organic compounds, the diffusion coefficient and the solubility of the substrate. The viscosity, contamination and the cost is reduced. Therefore, thermostable cellulase enzymes considerably help in completing the hydrolysis reactions (Srivastava et al., 2018; Khelila and Cheba, 2014). Furthermore, thermostable cellulase enzymes possessing these properties can also be considered as the model system for studying temperature stability and enzymatic activity which would surely open a new opportunity for protein engineering (Haki and Rakshit, 2003). Thermostable cellulases from various thermophilic microorganism have been reported (Aro et al., 2005; Liu et al., 2013; Karnaouri et al., 2014; de Vries and Visser, 2001; Singhania et al., 2010; Bayer et al., 2007; Schulein, 1998; Yazdi et al., 1990). Table 9.1 shows benefits of thermostable cellulases. These benefits result in increasing the economic exploitation of cellulase enzymes; which is one of the main costs adding component to the bioethanol technology (Vaishnav et al., 2018; Singhania et al., 2010). Ang et al. (2013) studied production of thermostable cellulases from Aspergillus fumigatus SK1. These thermostable cellulases showed higher FPU activity with effective hydrolysis for reduced time period. Thermostable enzymes are also able to reduce the hydrolysis time (Srivastava et al., 2015b; Dutta et al., 2014). Among several advanced approaches studied for production of cellulase enzyme, solid-state fermentation (SSF) is a preferable production method (Pandey, 1994). The SSF is conducted in the absence of free water in large amount but in the presence of sufficient amount of moisture which gives support for the growth of fungi on lignocellulosic material (Pandey et al., 2000; Singhania et al., 2010). Due to this, the dewatering cost during the downstream processing can be substantially reduced. Though, on an

TABLE 9.1 Benefits of thermostable cellulases. Reduced hydrolysis times Decreased risk of contamination Facilitates recovery of volatile products such as ethanol Lower costs for cooling after thermal pretreatment Reduction in enzyme needed for hydrolysis Increased rate of reaction Source: Based on Viikari et al. (2007); Abdel-Banat et al. (2010); Taylor et al. (2009); Turner et al. (2007); Yeoman et al. (2010).

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industrial scale, cellulase production is conducted using the submerged fermentation process, but because of the lower yield and higher cost, this process is not cost-effective. Conversely, SSF can be up-scaled for larger volume production. Moreover, SSF is beneficial for high enzyme concentration and productivity and also lower requirement of the sterility equipment (Holker et al., 2004). In addition, the crude enzyme obtained from the SSF process can be directly used for the hydrolysis of the lignocellulosic material. Furthermore, in comparison to the submerged fermentation process, the production cost of SSF can be reduced by ten times (Raghavarao et al., 2003) because of reduced energy requirement (Holker et al., 2004) and suitable inexpensive lignocellulosic material (Singhania et al., 2010). SSF provides proper condition for filamentous fungi because these can be grown easily (Holker et al., 2004). Filamentous fungi for instance T. reesei, A. niger, A. fumigates can be grown for production of cellulases using the SSF conditions. Several researchers have reported effective production of cellulase enzymes using SSF (Chandra et al., 2010; Srivastava et al., 2014; Ang et al., 2013). SSF has been suggested as a cost-effective method for producing several biochemicals and industrially important enzymes by using lignocellulosic biomass as substrate (Uncu and Cekmecelioglu, 2011). Many reports are available for cost-effective production of cellulases using SSF with lignocellulosic biomass as substrate (Dhillon et al., 2012). It requires simple equipment with reduced energy requirements resulting in higher enzyme yield and considerably reduced processing cost. SSF and SmF using lignocellulosic biomass as substrates is an added benefit to the process based on economic concern and promoting the search for efficient substrates. So, various approaches are being used for reducing the production cost of enzyme, such as SSF using lignocellulosic biomass as carbon source and enzyme inducer (Bhardwaj et al., 2021). Tengerdy (1998) reported that SSF was found to be mostly suitable for production of lignocellulolytic enzymes for several agricultural biotechnological applications. To demonstrate this, production of cellulase in SmF and SSF was compared. Cellulase yields are normally about 10 g/L, and the average fermentation cost in a stirred tank reactor is B$200/m3 in SmF. Therefore, the cost of production in SmF is B$20/kg. In SSF, the average production is about 10 mg/g substrate and the average fermentation cost is just about $25/mt. Therefore, the unit cost of cellulase produced by SSF is only $0.2/kg (Tengerdy, 1996, 1998; Pandey et al., 2000). Apart from SSF, improvement of production of cellulases depends upon selection of lignocellulosic substrates used for SSF. Lignocellulosic materials constituted of renewable substrates are found in enormous amount and considered as the potential substrates for production of biofuels. These biomasses are obtained as waste products of agricultural practices, particularly from various agriculture based industries (Perez et al., 2002). These biomasses are the central point of modern industries as these are natural and

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renewable resources of energy. Furthermore, these lignocellulosic materials can be efficiently converted into different value-added products such as cellulases, sugars generation for biofuels production and also inexpensive energy sources for fermentation and production of enzyme (Iqbal et al., 2013; Asgher et al., 2013). Therefore, use of appropriate lignocellulosic material under SSF can improve the production of cellulases to a larger extent. Besides SSF and appropriate substrate, cellulase production can be further improved by the use of genetically modified organisms (GMOs). Though, several filamentous fungi are able to produce cellulase enzymes but because of the lower yield, production on a commercial scale will not be viable. Introduction of high cellulase producing gene into thermophilic microorganism can improve the production of cellulases, but not much information is available in the literature. So, a thorough understanding about the genetics of the organisms is required. “Improvements in specific activities of cellulose can be possible via cellulase engineering which depend on rational design. The concept of artificial designing of cellulase seems to be more promising for cellulases having desired features. In addition with genetic modification, co-culture concept is advantageous for cellulase production under the SSF. Improvement in cellulase production can be achieved via co-culture of different fungi in the single medium (Kalyani et al., 2013). Co-culture also has several advantages, for example, higher productivity, adaptability and substrate utilization compared to pure culture (Holker et al., 2004). The enzymes system obtained via co-culturing of different fungi may interact with each other and forms a complete cellulase system. The performance of co-culturing fungi for cellulase production has also been reported by (Hu et al., 2011). These authors found that the lignocellulosic components were depolymerized to a greater extent when the fungi were co-cultured on lignocellulosic substrate during the SSF and showed better efficiency. In one of the study by (Kalyani et al., 2013), β-glucosidase activity was recovered by co-culturing of Sistotrema brinkmannii and Agaricus arvensis. In addition to various approaches to improve the cellulase production on an industrial scale, number of different cofactors such as addition of metal ions can also enhance the cellulase production, significantly (Srivastava et al., 2014). Recently, concept of nanomaterials has aroused as a new era in the revolution of renewable energy production. Dutta et al. (2014) reported enhanced cellulase production in the presence of hydroxyapatite nanoparticles. In this study, an improved thermal stability of cellulose was achieved along with reducing sugars at the hydrolysis temperature of 80 C, when rice husk/rice straw was used as the substrates. In one of the study by (Srivastava et al., 2015b) an improved cellulase production, thermal stability as well as sugar productivity in the presence of Fe3O4/alginate nanocomposite has been reported. In this study, a higher yield of cellulases using A. fumigatus AA001 under the SSF was achieved in the presence of Fe3O4/alginate nanocomposite. Further, cellulase production and its thermal

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stability were also improved in the presence of Fe3O4 nanoparticle and Fe3O4/ alginate nanocomposite. The results reported by the authors clearly exposed that nanoparticles can play an important role to improve the cellulase production as well as the entire bioconversion process. In addition, an improved cellulase production and thermal stability has also been reported in the presence of nickel cobaltite (NiCo2O4) nanoparticle under the SSF using the thermotolerant A. fumigatus NS (Class: Eurotiomycetes) (Srivastava et al., 2014). Thus, nanoparticles may be considered as one of the key factor to enhance the cellulase production in the near future” (Magrey et al., 2018). Apart from the abovementioned studies, Ansari and Husain (2012), Shakeel and Qayyum (2012) and Verma et al. (2013) have reported an improvement of cellulase production, thermal stability and also its hydrolysis efficiency in the presence of nanoparticles. Ansari and Husain (2012) immobilized cellulase enzyme in the presence of magnetic nanoparticles. Enzymes immobilized on nanoparticles showed a broad ranging working pH and temperature and high thermal stability in comparison to the native enzymes. Nanoparticle based immobilization showed following important features: 1. Nano-enzyme particles can be easily synthesized in high solid content without the use of surfactants and toxic reagents 2. Well defined and homogenous core-shell nanoparticles with a thick enzyme shell can be obtained 3. Particle size can be easily tailored within utility limits. Furthermore, with the increasing attention given to cascade enzymatic reaction and in vitro synthetic biology, it appears that co-immobilization of multi-enzymes could be obtained on these nanoparticles Verma et al. (2013) immobilized β-glucosidase on a nanoscale carrier for potential application in biofuel production. β-Glucosidase from A. niger was immobilized to functionalized magnetic nanoparticles by covalent binding. Immobilized nanoparticles showed 93% immobilization binding. The optimum pH of free enzyme was 4.0 and immobilized enzyme was 6.0. However, the temperature optima was same at 60 C. Michaelis constant was 3.5 for free enzyme and 4.3 mM for immobilized β-glucosidase. Immobilized enzyme showed higher thermostability at 70  C. The immobilized nanoparticle-enzyme conjugate was found to retain more than 50% enzyme activity up to the 16th cycle. Maximum amount of glucose from hydrolysis of cellobiose by immobilized β-glucosidase was obtained in 16 hours. Although, nanoparticles show improved production of cellulase, thermal stability, and hydrolysis efficiency, the mechanism is not well known. So, emphasis should be made on production of cellulases using the nanomaterials. Genomic sequencing of cellulolytic organisms has been conducted during the last decade. Important new information about how microorganisms

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degrade cellulose has been obtained from genomic sequences. “The sequences of the aerobic microorganisms: Hypocrea jecorina (Tricoderma reesei), Phanerochaete chrysosporium, and Thermofida fusca, all contain multiple cellulase genes, most of which encode a carbohydrate binding module (CBM), and several processive cellulase genes are present in each organism (Martinez et al., 2004; Lykidis et al., 2007). The genome sequences of Clostridium thermocellum, Ruminococcus albus and Ruminococcus flavifaciens all contain scaffolding genes and multiple cellulases genes that encode docerin domains, consistent with the presence of cellulosomes in these anaerobic bacteria and several processive cellulase genes are found among the docerin encoding genes (Bayer et al., 2008). There are three cellulolytic microorganisms whose genomes do not contain known genes for processive cellulases or docerin domains or scaffoldings: Cytophiga hutchinsonii is an aerobic cellulolytic bacterium that is tightly bound to cellulose fibers during growth on cellulose (Xie et al., 2007), while Fibrobacter succinogenes is an anaerobic cellulolytic bacterium that also is tightly bound to cellulose fibers (Qi et al., 2007). From their genome sequences these organisms do not use either the free cellulase or the cellulosomal mechanism to degrade cellulose, so they must use a novel mechanism (Wilson, 2008). Finally Postia plancenta is an aerobic brown rot fungus that appears to produce hydrogen peroxide and Fe (II) ions that generate OH radicals that carry out cellulose depolymerization (Martinez et al., 2009). Further research is needed on each of these organisms to determine the detailed mechanisms that they use to completely metabolize cellulose. Metagenomics is also being used to try to identify new cellulases and a major study of DNA isolated from the microorganisms in termite guts was reported (Warnecke et al., 2007). About one hundred hydrolases related to cellulose degradation were identified including members of eight cellulose families; however, no members of families containing exocellulase genes were present. This might be due to the fact that termites chew up the biomass into very fine particles that may be easier to degrade then other forms of cellulose. It is interesting that screening of genes for cellulase activity, either from isolated organisms or from DNA libraries from various environmental samples has not identified any new cellulase families in the past few years. One novel hydrolase containing both a glucanase and a xylanase was found in a library isolated from soil” (Magrey et al., 2018; Nam et al., 2009). A number of proteins have been identified which modify cellulose and improve its hydrolysis by cellulases. One is a class of plant proteins called expansions (Carey and Cosgrove, 2007). Another is a fungal protein which shows some homology to expansin called swollenin (Yao et al., 2008). “Expansins are a class of plant proteins which interact with and modify cell walls and/or cell wall components by an unknown activity, thought to result in expansion, slippage, or lengthening of cell wall structures. These twodomain proteins consist of a domain homologous to the GH45 EG catalytic

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core and a second domain homologous to Group II grass pollen allergens; both domains have no known catalytic function and display no detectable hydrolytic activity on lignocellulosic or model substrates (expansins are thoroughly reviewed by Sampedro and Cosgrove, 2005). T. reesei has been shown to express a protein, named swollenin, which has sequence homology to plant expansins and displays a similar mysterious disruptive effect on cellulosic substrates. Isolated T. reesei swollenin has been shown, without cause to formation of detectable reducing ends, to weaken filter paper and to affect superstructural changes in cotton fibrils by light and atomic force microscopy (Saloheimo et al., 2002). Despite the apparent lack of direct lytic activity on cellulose, the addition of T. reesei swollenin significantly increases the breakdown of filter paper by cellulases (Wang et al., 2010; Ja¨ger et al., 2011). Swollenin from A. fumigatus is reported to have weak lytic activity on CMC but no apparent hydrolytic activity on microcrystalline cellulose, though treatment of microcrystalline cellulose with A. fumigatus swollenin reduced apparent microcrystalline cellulose particle size and potentiated breakdown of the cellulose by hydrolytic cellulases” (Sweeney and Xu, 2012; Chen et al., 2010). The purified swollenin showed slight hydrolytic activities on xylan and yeast cell wall glucan, whereas no noticeable activities on carboxymethyl cellulose, filter paper, cotton fiber, or cellulose powder CF11 were seen. These results show that although swollenins maintain unidentified glycohydrolytic activities, it is inactive against β-1,4-glycosidic bonds in cellulose (Yao et al., 2008). Bjerkandera adusta which is a basidiomycete produces a similar protein, loosinin, which increases cellulase activity on cotton and agave bagasse (Quiroz-Castan˜eda et al., 2011). Bacterial species, including Bacillus subtilis and Hahella chejuensis (Kerff et al., 2008; Lee et al., 2010) also produce expansin-like molecules. Like fungal swollenins, bacterial expansins change cellulose fiber structure and support the breakdown of cellulose by hydrolases without showing noticeable direct hydrolase activity (Lee et al., 2010; Kim et al., 2009). Structural and mutational studies of expansin EXLX1 from B. substilis have thrown some light on these proteins, showing that many clustered residues on GH45-like domain are needed for EXLX1’s cell wall modifying activity, and that the second domain is possibly a new type of CBM (Kerff et al., 2008; Georgelis et al., 2011). Further study will be needed for determining the mechanism by which these molecules increase conversion of lignocelluloses and allow industrial use of this class of protein. CIP Proteins CIP1 and CIP2 (cellulose induced protein-1 and -2, respectively) were first reported in a transcriptional analysis of T. reesei. Both contain a CBM and are co-regulated with known cellulase enzymes (Foreman et al., 2003). The function of cellulose induced protein-1 is unknown, although it is claimed that cellulose induced protein-1 from T. reesei has weaker activity on p-nitrophenyl β-D-cellobioside (Foreman et al., 2011) and some synergistic activity with both

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GH61 and swollenin (Scott et al., 2011) cellulose induced protein-2, found in both T. reesei and Schizophyllum commune, has been shown to be an esterase which cleaves the methyl ester of 4-O-methyl-D-glucuronic acid (Li et al., 2007). This enzyme, now classified as the first member of CE15 family (EC 3.1.1.-), possibly acts in the cleavage of hemicellulose-lignin crosslinks. More investigation of both the functions and the potential of these enzymes in commercial applications are required. “While many microorganisms secrete biomass degrading enzymes into their environment, other microbes, particularly anaerobic biomass degrading microbes, use cell-surface linked enzymes to breakdown lignocellulosic materials. Cellulosomes are arrays of multiple cellulase and hemicellulase proteins, assembled by specific interactions between dockerin domains on the enzyme and cohesins bound to structural scaffoldings on the microbial surface (reviewed extensively in Ding et al., 2008; Gilbert, 2007; Bayer et al., 2007; Fontes and Gilbert, 2010). This spatial clustering of multiple lignocellulose degrading enzymes results in an increased synergy between lytic activities (Moraı¨s et al., 2011). It has been shown that recombinant cellulosomes can be transplanted to other industrially useful organisms, such as S. cerevisiae (Tsai et al., 2010; Lilly et al., 2009) and B. subtilis (Anderson et al., 2011). The ability of cellulosomes to cluster activities may present unique capabilities, both in synergistic breakdown of a substrate and in targeted degradation of specific biomass components” (Sweeney and Xu, 2012). The assembly of different components of cellulosomes on the surface of the cell is shown in Fig. 9.1 (Arora et al., 2015). In Bacillus subtilis, an expansin like protein has been identified and its structure has been determined. Expansin protein was shown to stimulate corn stover hydrolysis by crude cellulase enzyme (Kerff et al., 2008; Kim et al., 2009). Many microorganisms secrete proteins which only contain CBMs and two T. fusca proteins (E7, E8) have been purified and shown to stimulate low concentration of cellulases (Moser et al., 2008). Finally there are the family 61 proteins stated before. Several efforts have been made to model the cellulase catalyzed hydrolysis of crystalline cellulose but still it is not known about this process to create a true mechanistic model. A detailed mechanistic model of amorphous cellulose hydrolysis by crude cellulase was presented by Peri et al. (2007). This model fits their experimental results very well. There are three major strategies which are being used to engineer cellulase enzymes with higher activity on crystalline cellulose: directed evolution, rational design, and increasing thermostability of cellulases by either of the previous methods, which may also lead to higher activity. “Engineering more thermostable enzymes is relatively straightforward and there are some general approaches that can be applied to any enzyme for which a large number of related sequences are known, as is true for most cellulases (Heinzelman et al., 2009). A paper describes evolving T. Reesei Cel12A

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CELLULOSE CARBOHYDRATE BINDING MODULE

CELLULOSOMAL ENZYME DOCKERIN

TYPE-I COHESIN INTERACTION SCAFFOLDIN

DOCKERIN

TYPE-II COHESIN INTERACTION SURFACE LAYER HOMOLOGY

BACTERIAL CELL FIGURE 9.1 Cellulosome structure and assembly. Arora, R., Behera, S., Sharma, N.K., Kumar, S., 2015. Bioprospecting thermostable cellulosomes for efficient biofuel production from lignocellulosic biomass. Bioresour. Bioprocess. 2, 38. https://doi.org/10.1186/s40643-015-00664. Distributed under the terms of the Creative Commons Attribution 4.0 International License.

for enhanced thermostability while another evolved a family 5 endoglucanase with higher activity on CMC but it had no activity on crystalline cellulose (Lin et al., 2009). At this time there are no published reports of engineered cellulases with major (greater than 1.5-fold) increases in activity on crystalline cellulose. Furthermore, to be useful on an industrial process the improved enzyme has to increase the activity of a synergistic mixture containing several cellulases and in several cases mutant enzymes with higher activity do not do this (Zhang et al., 2000). At this time, it is not clear why this is happening but it has been shown for several exocellulases. Another surprising result is that an improved processive endocellulase

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catalytic domain, produced by combining two site directed mutations, that showed higher activity in synergistic mixtures then the wild type catalytic domain, did not show higher activity on crystalline cellulose then wild type intact enzyme when the missing domains were added back to form the intact mutant enzyme. This result seems surprising but it shows that activity on crystalline cellulose may involve interactions between the catalytic domain and the carbohydrate binding module (CBM) that go beyond the CBM simply anchoring the catalytic domain to the cellulose (Esteghlalian et al., 2001). Directed evolution of cellulases with improved activity on crystalline cellulose requires that the mutant cellulases be screened on a crystalline substrate not on CMC as most mutations that increase CMCase activity decrease activity on crystalline cellulose. Furthermore, the native enzyme should be utilized, not the catalytic domain given the above result. Finally any improved enzymes need to be tested in the appropriate synergistic mixture on the actual substrate for the final process in order to be certain that they will be useful. A problem with directed evolution is that it can only be used to screen potential single or with a massive screen potential double mutations, since the mutant library size required to include most possible larger multiple mutations is too large. Rational design does not have this limitation, but it does require a detail understanding of structure functional relationships for cellulase crystalline cellulose activity that is still lacking. If we can gain a clear understanding of exactly how cellulases hydrolyze crystalline cellulose it should be possible to design enzymes with multiple changes that have higher activity on specific biomass substrates. Another approach to engineering more active cellulose degrading enzymes is to create optimized cellulosomes by synthesizing hybrid scaffolding molecules that contain cohesins with different binding specificity from different organisms. The exact composition and geometry of the enzymes in a cellulosome can be controlled by attaching the appropriate docerin domain to each enzyme in the cellulosome. In one experiment, the six T. fusca cellulases produced during growth on cellulose were modified by removing their family 2 CBM domain and replacing it with a docerin domain, thus converting a free cellulase of interesting findings from this approach but it did not produce a cellulosome with increased cellulase activity over the free cellulase system (Caspi et al., 2008). Another experiment involved adding CBM domains to two of the key cellulosomal enzymes. This increased the activity of each enzyme when it was bound to an artificial scaffolding containing one cohesin but various designer cellulosome containing the modified enzymes and other cellulases all had lower activity on crystalline cellulase than comparable designer cellulosomes containing the wild type enzymes (Mingardon et al., 2007). These results do not invalidate the possibility of improving the activity of cellulosomes by the designer approach but we need to understand more about how the enzymes on cellulosomes interact to degrade crystalline cellulose before we can create better cellulosomes” (Wilson, 2009b).

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Further reading Ajeje, S.B., Hu, Y., Song, G., Peter, S.B., Afful, R.G., Sun, F., et al., 2021. Thermostable cellulases/xylanases from thermophilic and hyperthermophilic microorganisms: current perspective. Front. Bioeng. Biotechnol. 9, 794304. Available from: https://doi.org/10.3389/fbioe. 2021.794304. Akram, F., Haq, I.U., 2020. Overexpression and characterization of TnCel12B, a hyperthermophilic GH12 Endo-1,4-β-glucanase cloned from thermotoga naphthophila RKU-10T. Anal. Biochem. 599, 113741. Assareh, R., Shahbani Zahiri, H., Akbari Noghabi, K., Aminzadeh, S., Bakhshi khaniki, G., 2012. Characterization of the newly isolated Geobacillus Sp. T1, the efficient cellulaseproducer on untreated barley and wheat straws. Bioresour. Technol. 120, 99 105. Boyce, A., Walsh, G., 2018. Expression and characterisation of a thermophilic Endo-1,4-β-glucanase from sulfolobus shibatae of potential industrial application. Mol. Biol. Rep. 45, 2201 2211.

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Fusco, F.A., Fiorentino, G., Pedone, E., Contursi, P., Bartolucci, S., Limauro, D., 2018. Biochemical characterization of a novel thermostable β-glucosidase from dictyoglomus turgidum. Int. J. Biol. Macromol. 113, 783 791. Global Cellulase (CAS 9012-54-8) Market by Type (EG, CBH, BG), By Application (Animal Feed, Textile Industry, Food & Beverages, Biofuels, Others) And By Region (North America, Latin America, Europe, Asia Pacific and Middle East & Africa), Forecast To 2028. Grassick, A., Murray, P.G., Thompson, R., Collins, C.M., Byrnes, L., Birrane, G., et al., 2004. Three-dimensional structure of a thermostable native cellobiohydrolase, CBH IB, and molecular characterization of the Cel7 gene from the filamentous fungus. Talaromyces emersonii. Eur. J. Biochem. 271, 4495 4506. Hong, J., Tamaki, H., Kumagai, H., 2007. Cloning and functional expression of thermostable β-glucosidase gene from thermoascus aurantiacus. Appl. Microbiol. Biotechnol. 73, 1331 1339. Kar, B., Verma, P., den Haan, R., Sharma, A.K., 2017. Characterization of a recombinant thermostable β-glucosidase from Putranjiva roxburghii expressed in Saccharomyces cerevisiae and its use for efficient biomass conversion. Process. Biochem. 63, 66 75. Murray, P., Aro, N., Collins, C., Grassick, A., Penttila, M., Saloheimo, M., et al., 2004. Expression in Trichoderma reesei and characterisation of a thermostable family 3 β-glucosidase from the moderately thermophilic fungus Talaromyces emersonii. Protein Expr. Purif. 38, 248 257. Olajuyigbe, F.M., Ogunyewo, O.A., 2016. Enhanced production and physicochemical properties of thermostable crude cellulase from sporothrix carnis grown on corn cob. Biocatal. Agric. Biotechnol. 7, 110 117. Potprommanee, L., Wang, X., Han, Y., Nyobe, D., Peng, Y., Huang, Q., et al., 2017. Characterization of a thermophilic cellulase from geobacillus Sp. HTA426, an efficient cellulase-producer on alkali pretreated of lignocellulosic biomass. PLoS One 12, e0175004. Shi, R., Li, Z., Ye, Q., Xu, J., Liu, Y., 2013. Heterologous expression and characterization of a novel thermo-halotolerant endoglucanase Cel5H from dictyoglomus thermophilum. Bioresour. Technol. 142, 338 344. Singh, R., Kumar, R., Bishnoi, K., Bishnoi, N.R., 2009. Optimization of synergistic parameters for thermostable cellulase activity of aspergillus heteromorphus using response surface methodology. Biochem. Eng. J. 48, 28 35. Wang, K., Luo, H., Bai, Y., Shi, P., Huang, H., Xue, X., et al., 2014. A thermophilic endo-1,4β-glucanase from Talaromyces emersonii CBS3940.64 with broad substrate specificity and great application potentials. Appl. Microbiol. Biotechnol. 98, 7051 7060. Yan, Q., Hua, C., Yang, S., Li, Y., Jiang, Z., 2012. High level expression of extracellular secretion of a β-glucosidase gene (PtBglu3) from Paecilomyces thermophila in Pichia pastoris. Protein Expr. Purif. 84, 64 72. Yang, G., Yang, D., Wang, X., Cao, W., 2021. A novel thermostable cellulase-producing Bacillus licheniformis A5 acts synergistically with Bacillus subtilis B2 to improve degradation of Chinese distillers’ grains. Bioresour. Technol. 325, 124729.

Chapter 10

Cellulases and auxiliary enzymes 10.1 Introduction Lignocellulosic biomass is the most economical and highly renewable natural resource in the world. It can be used as a renewable raw material for the production of biofuels and value-added chemicals, and biorefinery of lignocellulosic materials contributes significantly to the sustainable development of economy. However, production of fermentable sugars from lignocellulosic substrates is suffering from higher cost and reduced effectiveness of lignocellulolytic enzymes. Filamentous fungi, particularly Trichoderma reesei, are the most efficient producers of lignocellulolytic enzymes, and enhancement of production efficiency of enzymes and optimization of enzyme composition are of immense importance for biorefinery of lignocellulosic materials. Efficient production of an optimized combination of lignocellulolytic enzymes with enhanced hydrolytic efficiency will significantly contribute to production of biofuels from lignocellulosic material (Zhang et al., 2019). Some proteins have been identified, during hydrolysis of lignocellulosic biomass with enzymes, which can loosen the packaging of cellulose fibril network in a non-hydrolytic manner. This process is called amorphogenesis. These proteins act in a synergistic manner together cellulases, thus increasing the accessibility of cellulose to the enzymes. For this reason, these helper proteins are called amorphogenesis-inducing agents.

10.2 Cellulases Cellulose-hydrolyzing cellulases (Cels) comprise catalytic modules and carbohydrate-binding modules. Catalytic modules belong to 14 GH families whereas carbohydrate-binding modules belong to 15 carbohydrate-binding module families. Functionally, cellulases include 1,4-β-D-glucan-cellobiohydrolase (CBH) (EC 3.2.1.91), endo-1,4-β-D-glucanase (EG) EC (3.2.1.4), and β-glucosidase (BG) (EC 3.2.1.21) (Hasunuma et al., 2013). The different substrate specificity of these enzymes allow them to act in a synergistic manner on cellulose. The products of endoglucanases are cellodextrans and cellobiohydrolases, are cellobiose. These products inhibit the activity of Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00006-6 © 2023 Elsevier Inc. All rights reserved.

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enzyme. Therefore, effective hydrolysis of cellulose necessiates the presence of β-glucosidases which cuts the final glycosidic bonds and produces glucose (end product). Most of the cellulase enzymes are found to be active at temperature 20 C 60 C, and pH 4 8 and but some cellulase enzymes from extremophiles are able to extend their activity/stability far beyond this temperature and pH range (Xu, 2011). A few other important enzymes catalyzing the reversible phosphorolytic cleavage and epimerization are also grouped as part of the cellulase enzyme complex. “Cellobiose phosphorylase or cellobiase (orthophosphate α-D-glucosyl transferase, EC 2.4.1.20) catalyzes reversible phosphorolytic cleavage of cellobiose to glucose. Cellodextrin phosphorylase (1,4-β-D-oligoglucan orthophosphate α-D-glucosyl transferase, EC 2.4.1.49) catalyzes the conversion of cellodextrins (cellotriose to cellohexose) to glucose. Cellodextrin phosphorylase does not act on cellobiose. Cellobiose epimerase (EC 5.1.3.11) catalyzes the epimerization of disaccharides like cellobiose into 4-O-β-D-glucosylmannose” (Bhardwaj et al., 2021; Sharma et al., 2016). The potential of these enzymes as biocatalysts for conversion of biomass on an industrial scale has yet to be thoroughly investigated. “Cellulases hydrolyze cellulose which is a linear polysaccharide molecule containing repeated β (1-4) linked D-glucopyranosyl (Glc) units. Glucose and short cellodextrins are formed. The reaction is conducted mostly by cellulase enzymes. These include no less than two exo-β-glucanases or cellobiohydrolases (CBHs; EC 3.2.1.91) (CBH I and CBH II), four endo-β glucanases (EGs; EC 3.2.1.4) (EG I, EG II, EG III, EG V), and one β-glucosidase (βG; EC 3.2.1.21). Cellulose has a fairly simple composition (anhydro-Glc units only) and morphology (largely amorphous and monoclinic Iβ or triclinic Iα crystalline) but there exists a huge natural diversity of cellulase enzymes with catalytic modules belonging to 14 GH families for accommodating four main reactions forms and diverse types of synergies” (Bajpai, 2021; Duncan and Schilling, 2010; Zhang et al., 2010).

10.2.1 Cellobiohydrolase Cellobiohydrolase (CBH) is an exocellulase which degrades cellulose by hydrolyzing the 1,4-β-D-glycosidic bonds. CBH cleaves two to four units from the ends of cellulose. There are two types of CBH CBH I and CBH II. CBH I cleaves progressively from the reducing end of cellulose whereas CBH II cleaves progressively from the nonreducing end of cellulose. The exo-acting CBH I is called Cel7A. The endo-acting CBH I is called Cel7B. CBH Cel6A contains an active site within a tunnel that opens and closes in response to ligand binding. “CBHs constitute the bulk of several cellulolytic mixtures from fungi. The enzymes have active sites that are like tunnel and they chop cellobiose units from the end of the cellulose chains. There are two types of CBH enzymes. CBH I cleaves progressively from the reducing

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end, whereas CBH II cleaves progressively from the nonreducing end of cellulose. The exo-acting CBH I is termed Cel7A. The endo-acting CBH I is termed Cel7B. The active site in Cellobiohydrolase Cel6A opens and closes in response to ligand binding. In fungi, CBH I and CBH II activities are associated with GH7 and GH6 families, respectively. Many fungal CBHs are associated with type 1 carbohydrate-binding modules, and these may assist the catalytic core with processivity (Bajpai, 2021; Beckham et al., 2010; Tavagnacco et al., 2011; Teter et al., 2014). Degradation of crystalline cellulose is performed mostly by CBHs. Therefore, these enzymes are very important for the degradation of lignocelluloses. Archetypical CBHs are present in GH6 and 7, and also 48, families. GH7 CBH is present in most of the cellulolytic fungi. GH6 CBH is present in several cellulase-producing fungi. Amongst secreted enzymes of cellulaseproducing fungi, up to about 70% wt or so are CBHs (Herpoe¨l-Gimbert et al., 2008; Sipos et al., 2010; Chundawat et al., 2011). GH7 CBH is specific toward the reducing end of a cellulose chain. In contrast, GH6 CBH is specific to the nonreducing end of a cellulose chain. These specificities are of opposite types make GH7 and 6 CBHs extremely synergistic and cooperative in the degradation of their substrate” (Bajpai, 2021). “The CBH catalytic core features tunnel-like active sites, a topology that equips CBH with the ability to hydrolyze cellulose processively: it threads into the end of a cellulose chain through its active site, cleaves off a cellobiosyl unit, glides down the chain, and starts the next hydrolysis step. A CBM may assist the catalytic core with processivity. Such processive reactions, plus the insolubility of the cellulose substrate, makes CBH kinetics deviant from the Michaelis-Menten model, and show significant fractal and local jamming effect. Processive CBH movement can be obstructed by kinks or other impediments on the cellulose surface; and as such it has been suggested that k(off) values may be a major factor in CBH efficiency. GH7 CBH I has about ten anhydro-Glc-binding subsites in its active site, in which cellulose or cellodextrin is bound and activated by means of hydrogen bonding and π-stacking with major amino acid residues. Many CBHs have also CBMs in addition to the catalytic core which is an important key in action on CBH on crystalline cellulose” (Sweeney and Xu, 2012; Beckham et al., 2010; Vocadlo and Davies, 2008; Warden et al., 2011; Xu and Ding, 2007; Igarashi et al., 2011; Kurasin and Va¨ljama¨e, 2011; Liu et al., 2011; Praestgaard et al., 2011).

10.2.2 endo-1,4-β-Glucanase “Degradation of amorphous cellulose can be carried out by EGs (EC 3.2.1.4). Unlike CBH, EG hydrolyzes internal glycosidic bonds in cellulose in a random, on-off fashion. Such dynamics make EG well-suited to less orderly or partially shielded cellulose parts, generating new cellulose chain

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ends for CBH action. A few EGs can act processively on crystalline cellulose. There is a significant synergism between CBH and EG, and their co-presence and cooperation are determinant for highly efficient enzymatic systems of industrial biomass-conversion” (Sweeney and Xu, 2012; Wilson, 2008; Li et al., 2010). EGs are found to be present in several organisms. Different EGs have a catalytic core which belong to more than ten GH families, of which GH5, 7, 9, 12, 45, and 48 are representative. “Typical cellulolytic fungi secrete EGs at 20% wt level in their secretomes. Also known as EG-I, II, III, and V, respectively, GH7, 5, 12, and 45 EG are most common in natural fungal cellulase mixes. Most cellulolytic fungi and bacteria produce numerous EGs. Although they all act on the same cellulose substrate, they do so through differing mechanisms inverting for GH6, 9, 45, and 48 EGs; retaining for GH5, 7, 12 EGs. Such EG plurality may relate to different EGs’ side-activities on hemicellulose in degrading complex lignocellulose, or synergism between processive and conventional EGs” (Sweeney and Xu, 2012; Herpoe¨l-Gimbert et al., 2008; Chundawat et al., 2011; Vlasenko et al., 2010).

10.2.3 β-Glucosidase β-Glucosidase is an important component of cellulase enzyme complex that is necessary for complete hydrolysis of cellulose into glucose. β-glucosidase gets inhibited by its product glucose. It is a main bottleneck in the efficient conversion of biomass by cellulases. For avoiding this problem many strategies have been used along with its production strategies and general properties. It plays a very important role in the production of ethanol from biomass through enzymatic method. Therefore many improvements have taken place in the commercial production of cellulases for hydrolysis of biomass, which contains higher and improved β-glucosidases to efficiently convert biomass. Most of the fungi producing cellulases are short of one or the other components of cellulase needed to effectively breakdown the cellulosic fibers. It is needed for a deficient enzyme to be added so that all the components are present in optimal concentrations. Possibility exists that a deficient component be over-expressed in base enzyme producing organism for getting desired combination having all the components of cellulase enzyme. The choice of enzyme component to be expressed or over-expressed is generally based on the absence of that enzyme component in base preparations. Most of the industrial production of cellulase enzyme is conducted by recombinant fungal strains. Novozyme Inc. used T. reesei as parent strain and isolated effective cellulolytic and non-cellulolytic genes from other organism and transferred them into this strain for effective preparation of enzyme mixture in a single organism (McFarland et al., 2007). Likewise, Dyadic’s strain was also a recombinant which was found to produce few accessory enzymes.

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Dotsenko et al. (2015) reported that “co-fermentation of Penicillium verruculosum (basic strain) with another P. verruculosum with heterologouslyexpressed Aspergillus sp. β glucosidase produced up to 19% of β glucosidase as compared with 4% in the basic strain alone. This cofermented enzyme preparation led to 113% increased hydrolysis of avicel than the basic strain enzyme preparation.” In another study by Bulakhov et al. (2017), “the enzyme preparations produced by recombinant P. verruculosum strains, expressing the heterologous AnBGL (β glucosidase from A. niger) or TrLPMO LPMO from T. reesei under the control of the glucoamylase gene promoter in a starch-containing medium, proved to be more effective in hydrolysis of a lignocellulosic substrate than the control enzyme preparations without the heterologous enzymes. The enzyme composition containing both AnBGL and TrLPMO showed the highest performance in lignocellulose hydrolysis, providing a background for developing a fungal strain capable of expressing both heterologous enzymes simultaneously” (Singh et al., 2017). The β glucosidase genes from several yeast, mold, bacterial, plant, and animal systems have been cloned and expressed in E. coli as well as eukaryotic hosts for instance Pichia stipitis, Saccharomyces cerevisiae, and filamentous fungi. Filamentous fungi are found to be a good producer of β-glucosidases and a number of β-glucosidases from glycosyl hydrolase family 3 and 1 have been purified and characterized from different fungi. In spite of being good producers of the enzyme, reports on cloning of the genes encoding β-glucosidase from fungi are not many because of complexities. Even Saccharomyces and Pichia have also been used as host for expressing eukaryotic β-glucosidases genes. Many thermotolerant β-glucosidases from bacteria have been cloned and expressed in Escherichia coli. A β-glucosidase I coding sequence from P. decumbens was ligated with the CBH 1 promoter of T. reesei which is extensively used in enzyme industry for improving β-glucosidase activity in T. reesei in order to efficiently degrade cellulose (Ma et al., 2011). “The ligated sequences of β-glucosidase I coding sequence and CBH I promoter sequence were introduced into the genome of T. reesei RUT-C-30 through Agrobacterium-mediated transformation. Two transformed strain were selected based on their β-glucosidase activity and filter paper activity, which were respectively, 6 8 and 30 folds higher compared with the wild strain. Furthermore, heterologously expressed pBGL1 was purified and added to enzyme complex secreted by T. reesei RUT-C30 and was observed that during saccharification of pretreated cornstalk, the yield of glucose increased by up to 80%. This result showed that heterologous expression of a BGL in T. reesei could produce a balanced cellulase blend (Ma et al., 2011). Transformation was done in cellulolytic filamentous fungus T. reesei with the help of a plasmid carrying a dominant selectable marker; the

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acetamidase gene (amdS) or the argB gene of A. nidulans, complementing respective argB mutation of T. reesei. Transformation frequency reached up to 600 transformants per pg of transforming DNA. Co-transformed unselected DNA efficiency was high (approx. 80%). Transformed DNA was found integrated at different locations, often in multiple tandem copies in the T. reesei genome. In addition to this, the E. coli β-glucosidase was expressed in T. reesei in enzymatically-active form from the A. nidulans’s gpd promoter” (Singh et al., 2017; Penttila¨ et al., 1987).

10.2.4 Cellulase auxiliary enzymes Expansins are phytoproteins which are able to loosen the plant cell wall and disrupt the cellulose crystal structure (Marowa et al., 2016) while swollenins are expansin derivatives from fungi (e.g., T. reesei, A. fumigatus, etc) and bacteria (e.g., Bacillus subtilis). Swollenin also show crystal-disruption activity on cellulosic materials (Nakashima et al., 2014; Ward and Penttilla, 2002). There is evidence that these non-hydrolytic accessory proteins can improve the cellulase activity because these proteins are able to disrupt the hydrogen bonds for reducing cellulose crystallinity and increasing the cellulase accessibility to enzymes (Harris et al., 2014). Expansins and expansin-like proteins were firstly isolated from plants as cell wall-loosening factors (Cosgrove, 2000). Their mechanisms of action are not fully known. It has been suggested that the rupture of hydrogen bonds may be involved (Sampedro and Cosgrove, 2005). Expansins are modular proteins containing two discrete domains connected by a short linker. The structure resembles to that of cellulases. But, as compared to the catalytic domain of glycoside hydrolase 45, the N-terminal domain of expansin lacks catalytic activity (Payne et al., 2015; Georgelis et al., 2015); the C-terminal domain is similar to certain carbohydrate-binding modules. Both the domains are needed for the complete cell wallloosening activity of expansins (Sampedro and Cosgrove, 2005; Duan et al., 2018). Mcqueen-Mason and Cosgrove (1994) proposed that expansins are able to break the hydrogen bonding between plant cell wall polysaccharides without hydrolyzing. “In 2002, the first expansin-like protein swollenin from fungi was discovered in T. reesei (Saloheimo et al., 2002). Fungal swollenins show sequence similarity to expansins and are often referred to as expansin-like proteins. In fact, swollenins was able to disrupt cotton fibers and filter paper structures on a microscopic level without detectable reducing sugars (Saloheimo et al., 2002). Over the last two decades, more than 10 types of swollenins have been reported (Yao et al., 2008; Zhou et al., 2011; Kang et al., 2013; Santos et al., 2017). The Expansin engineering Database (ExED1), has recently been released to the

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public (Lohoff et al., 2020). Generally, expansins are no longer than 250 amino acids and have a two-domain structure. The primary domain of expansins resembles the glycoside hydrolase family 45 (GH45) and this homology preserves certain sequence features of the GH45 catalytic site (Bhardwaj et al., 2020). The second domain has a characteristic flat aromatic-rich surface and is homologous to group-2 grass pollen allergens. Some studies have proposed that this domain functions as a CBM (Cosgrove, 2000; Georgelis et al., 2012). Within the expansin, the two domains are interconnected by a short linker, and both domains are required for plant cell-wall loosening activity. However, the structural discrepancies between swollenins and expansins lead to functional differences. For example, swollenins have an additional CBM domain, making them homologous to fungal cellulases in the N-terminal (Eibinger et al., 2016). In other cellulases, CBMs direct the binding of the enzymes to cellulosic surfaces and enhance lignocellulose degradation (Hoffre´n et al., 1995; Velikodvorskaia et al., 2013; Maharjan et al., 2018). Additionally, the Oglycosylation of linkers in swollenins may also have an effect on the enzyme binding to surfaces” (Zhang et al., 2021; Amore et al., 2017). For the first time, the synergistic effect of co-displayed cellulase and expansin-like protein on a S. cerevisiae cell surface was reported by Nakatani et al. (2013). About 2.9 fold higher degradation activity on phosphoric acid-swollen cellulose in comparison with the activity of cellulaseexpressing strain only was observed. Fused B. subtilis expansin and Clostridium thermocellum endoglucanase for the degradation of highly crystalline cellulose were studied by Nakashima et al. (2014). They observed about 35% digestibility by the fused proteins. The use of these accessory enzymes in cellulase mixtures for industrial applications is liable for improving the amount of reduced sugar attainable from lignocellulosic materials (Obeng et al., 2017). Swollenin contains N-terminal CBM linked to a C-terminal expansin-like domain (Saloheimo et al., 2002). Swollenin from T. reesei showed a distinctive mode of action and resemblance with both endoglucanases and cellobiohydrolases (Andberg et al., 2015). But, Swollenin does not possess any hydrolytic activity in itself. Gourlay et al. (2013) reported that swollenin synergistically improved endoxylanase rather than endoglucanase or cellobiohydrolase activity during hydrolysis of pretreated corn stover with enzyme. This is because swollenin rendered xylan part of lignocellulose more available for degradation by xylanases, thus indirectly exposing cellulose to the cellulases. Zhou et al. (2011) were able to successfully express swollenin in A. niger, and reported that the simultaneous incubation of swollenin with cellulases results in a important synergistic increase in cellulose hydrolysis. This synergy further improved when cellulose was pretreated with swollenin.

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Zhang et al. (2021) reported that TlSWO, (swollenin protein) from Talaromyces leycettanus JCM12802, is an acidic and mesophilic swollenin. It shows activity towards lichenan, barley β-glucan, sodium carboxymethyl cellulose and laminarin. A larger increase in glucose yield was seen when cellulose substrates were incubated with TlSWO and cellobiohydrolases. Furthermore, TlSWO showed synergistic effect on cellobiohydrolase when using PCS and PASC as substrates. But, no important synergistic effect was seen between TlSWO and endoglucanases, showing that TlSWO has superior coordination with cellobiohydrolases. “Cooperative action of expansins on improving the performance of cellulases and hemicellulases on hydrolysis of cellulosic and hemicellulosic substrates has been demonstrated. However, their synergism on cellulose hydrolysis was shown to be significant mostly under the conditions with low cellulase dosage. Several studies have demonstrated that the capability of plant and microbial expansins on enhancing the efficiency of cellulose hydrolysis by cellulases under specific conditions. For example, Zea h protein purified from fresh postharvest corn stover showed synergistic interaction with cellulase and increased degradation of filter paper (Han and Chen, 2007). An expansin from Oryzae sativa was shown to increase the activity of cellulase up to 2.4 times compared to the enzyme alone, at low cellulase loading (Seki et al., 2015). An expansin-like protein named swollenin isolated from T. reesei was shown to disrupt and swell cotton fiber (Saloheimo et al., 2002). In addition, Kim and coworkers demonstrated that BsEXLX1, the first bacterial expansin-like protein described from Bacillus subtilis enhanced the enzymatic activity of cellulase on hydrolysis of filter paper by 240% (Kim et al., 2009). An active ternary cellulolytic system comprising T. reesei cellulase (Celluclast, Novozymes), a selected in-house crude enzyme preparation from A. aculeatus BCC199 containing a complex mixture of cellulases and hemicellulases, and a recombinant Bacillus subtilis expansin was optimized using a systematic experimental design approach (Suwannarangsee et al., 2012). This optimized ternary enzyme complex produced reducing sugar products up to twofolds greater than a commercial cellulase from T. reesei at the same filter paper unit (FPU) loading” (Zhang et al., 2019). In the recent years, discovery of lytic polysaccharide monooxygenases (LPMOs) has been regarded as the major revolution in cellulose degradation. “Research has been conducted on the role of non-hydrolytic auxiliary enzymes involved in cellulose hydrolysis and instead of endo-exo synergism, the LPMOs were used in solubilization of crystalline cellulose by oxidative cleavage of glycosidic bonds. LPMOs improve the saccharification by providing new sites for cellobiohydrolases and β-glucosidase actions. These enzymes introduced a new, oxidative mechanism to

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polysaccharide degradation. In cellulose degradation, it is presumed that LPMOs act on the surface of crystalline cellulose fibrils, thus rendering them more accessible to cellulases (Harris et al., 2010; Levasseur et al., 2013; Eibinger et al., 2014). Interestingly, these enzymes can derive the electrons required for this process from lignin via long-range electron transfer (Westereng et al., 2015). In the carbohydrate-active enzyme (CAZy) database, the LPMOs have been classified as members of families CBM33 and family GH61. But now again, these have been re-classified to auxiliary activities (AA) as opposed to their previous classification as GH61 and are found in the AA families 9 11 and 13” (Singh et al., 2017; Lombard et al., 2014). Mhuantong et al. (2015) conducted the metagenomic survey of sugarcane bagasse metagenome and reported the presence of GH61 family auxiliary enzymes (AA1, AA3, and AA9). DNA stable-isotope probing was used by Verastegui et al. (2014) for assessing the functionally active soil bacterial communities and their glycoside hydrolase genes from disparate Canadian soils. This functional annotation of metagenome also showed the presence of GH61 auxiliary enzymes. The occurrence of these auxiliary enzymes in metagenome would help to initiate new ideas on the mechanism of conversion of cellulosic biomass by enzymes. The knowledge gathered about this newly characterized protein will improve the overall hydrolytic potential of enzyme systems. M. thermophila which was earlier known as C. lucknowense C1 expressed amazingly higher LPMOs (Visser et al., 2011). It has high number of LPMOs genes than any other powerful producers of cellulases like T. reesei or A. niger (Table 10.1). A. nidulans also has high number of LPMOs genes but generally not all are expressed in their natural forms and therefore required to be expressed. Jagadeeswaran et al. (2016) cloned AN3046, an AA9 LPMO in A. nidulans which is active on cellulose, in protein expressing vector (pEXPYR) by ligation free cloning. Mass spectrometric analysis was conducted and it was found that the enzyme was active on hemicellulose xyloglucan. The AN3046 LPMO was found to synergistically degrade sorghum stover with other hydrolases. “M. thermophila is found to produce thermostable cellulase with higher G:C content in the protein coding genes suggesting the potential adaptability to higher temperatures. About 75% of M. thermophila codons have a higher GC content at the third nucleotide position (GC3) compared with the corresponding mesophilic counterpart (Berka et al., 2011). Enzymes from thermophillic strain when expressed in mesophillic host tend to retain their thermostable properties (Berka et al., 1998; Murray et al., 2004). Therefore,

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TABLE 10.1 Cellulase or cellulase encoding genes present in microorganisms. Microorganisms

Cellulase/genes

References

Trichoderma reesei

2 CBHs, 8 EGs,7 BGLs, 16 hemicellulases, 9 LPMOs genes

Aro et al. (2005), To¨rro¨nen et al. (1994)

Humicola insolens

2 CBHs,5 EGs lacking CBM

Schulein (1997)

Nerospora crassa

3 CBHs, 4 EGs, and 1 BGLs

Yazdi et al. (1990)

Penicillium decumbens

3 CBHs, 11 EGs, and 11 BGLs

Liu et al. (2013)

Myceliophthora thermophila

8 EGs,7 CBHs, 9 BGLs, 25 LPMOs genes

Karnaouri et al. (2014)

Aspergillus niger

2 CBHs, 2 EGs, 5BGLs, 7 LPMOs genes

de Vries and Visser (2001), Singhania et al. (2011)

Clostridium thermocellum

12 endo and exo cellulases

Bayer et al. (1998)

Source: Singh, A., Patel, A., Adsul, M., Mathur, A., Singhania, R., 2017. Genetic modification: a tool for enhancing cellulase secretion. Biofuel Res. J., 4(2), 600_610. https://doi.org/10.18331/ BRJ2017.4.2.5. Distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/).

it would be good to clone and express heterologous thermophillic cellulases into fast and uniform growing fungal strain which could be less viscous when grown in fermenter. M. thermophila has been genetically modified and used for industrial cellulase production by Dyadic because of its capacity of producing 100 g/L protein and also better efficacy of the enzyme” (Singh et al., 2017). Several LPMOs from different microbial origins, mostly AA9 from fungi with few from AA10 bacteria have been found to enhance cellulase activity in the hydrolysis of pure cellulose and pretreated lignocellulosic biomass (Zhang et al., 2019). AA9 LPMO is present typically in cellulolytic fungi and exists as a single domain or contain a CBM1. It attacks the inaccessible highly crystalline surface on cellulose before the action of the hydrolases. The initial attack of AA9 generates more accessible sites for cellulases, leading to improved enzymatic activity (Quinlan et al., 2011). The AA9 LPMO enzyme is now a main component in Cellic Ctec enzyme series which is an effective cellulase preparation from Novozymes. The actions of synergistic proteins in the hydrolysis of cellulose are presented in Fig. 10.1.

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FIGURE 10.1 Schematic diagram of the actions of cellulases and synergistic proteins in hydrolysis of cellulose. The C1-Cx lignocellulose degradation model for cellulose degradation. First, C1 factors, such as expansin, and lytic polysaccharide monooxygenases, which efficiently catalyze oxidative cleavage of glycosidic bonds in the recalcitrant polysaccharides of crystalline cellulose using molecular oxygen and the external electron donor, such as cellobiose dehydrogenase, ascorbic acid or gallic acid, creating nicking, swollen and disintegrated cellulosic structure, forming new initiation sites for conventional cellulases, namely, Cx factors. Second, the amorphous cellulose was hydrolyzed into monosaccharide by cellulase system. The endoglucanase acts on the amorphous (internal) region of the fibrils by cleavage of the β-glucosidic linkage, then the cellobiohydrolase releases cellobiose from the end of the polysaccharide chain, finally, β-glucosidase completes the degradation process by hydrolyzing cellobiose and other cellodextrins to glucose units. BGL, β-glucosidase; CBH I, cellobiohydrolase 1; CBH II, cellobiohydrolase 2; CDH, cellobiose dehydrogenase; C1 factor, cellulose hing domains (CBMs), plant expansins, bacterial expansins, and lytic polysaccharide monoxygenases; Cx factor, endoglucanase, cellobiohydrolase, and β-glucosidase; EG, endoglucanase; LPMO, lytic polysaccharide monooxygenase. Reproduced with permission Zhang, F., Bunterngsook, B., Li, J.X., Zhao, X.Q., Champreda, V., Liu, C.G., et al., 2019. Regulation and production of lignocellulolytic enzymes from Trichoderma reesei for biofuels production. In: Li, Y., Ge, X., editors. Advances in Bioenergy, vol. 4. Elsevier; p. 79 119.

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Liu, Y.S., Baker, J.O., Zeng, Y., Himmel, M.E., Haas, T., Ding, S.Y., 2011. Cellobiohydrolase hydrolyzes crystalline cellulose on hydrophobic faces. J. Biol. Chem. 286, 11195 11201. Liu, G., Zhang, L., Wei, X., Zou, G., Qin, Y., Ma, L., et al., 2013. Genomic and secretomic analyses reveal unique features of the lignocellulolytic enzyme system of Penicillium decumbens. PLoS One 8 (2), e55185. Li, Y., Irwin, D.C., Wilson, D.B., 2010. Increased crystalline cellulose activity via combinations of amino acid changes in the family 9 catalytic domain and family 3c cellulose binding module of Thermobifida fusca Cel9A. Appl. Environ. Microbiol. 76, 2582 2588. Lohoff, C., Buchholz, P.C.F., Roes-Hill, M.L., Pleiss, J., 2020. Expansin engineering database: a navigation and classification tool for expansins and homologues. Proteins 2020, 26001. Lombard, V., Ramulu, H.G., Drula, E., Coutinho, P.M., Henrissat, B., 2014. The carbohydrateactive enzymes database (CAZy) in 2013. Nucleic Acids Res. 42 (D1), D490 D495. Maharjan, A., Alkotaini, B., Kim, B.S., 2018. Fusion of carbohydrate binding modules to bifunctional cellulase to enhance binding affinity and cellulolytic activity. Biotechnol. Bioproc. E. 23, 79 85. Marowa, P., Ding, A., Kong, Y., 2016. Expansins: roles in plant growth and potential applications in crop improvement. Plant. Cell Rep. 35, 949 965. Available from: https://doi.org/ 10.1007/s00299-016-1948-4. Ma, L., Zhang, J., Zou, G., Wang, C., Zhou, Z., 2011. Improvement of cellulase activity in Trichoderma reesei by heterologous expression of a beta-glucosidase gene from Penicillium decumbens. Enzyme Microb. Technol. 49 (4), 366 371. McFarland, K.C., Ding, H., Teter, S., Vlasenko, E., Xu, F., Cherry, J., 2007. Development of improved cellulase mixtures in a single production organism. In: Eggleston, G., Vercellotti, J.R. (Eds.), Industrial Application of Enzymes on Carbohydrate-Based Material. ACS Symposium Series, 2. American Chemical Society, pp. 19 31. Mcqueen-Mason, S., Cosgrove, D.J., 1994. Disruption of hydrogen bonding between plant cell wall polymers by proteins that induce wall extension. PNAS 91, 6574 6578. Available from: https://doi.org/10.1073/pnas.91.14.6574. Mhuantong, W., Charoensawan, V., Kanokratana, P., Tangphatsornruang, S., Champreda, V., 2015. Comparative analysis of sugarcane bagasse metagenome reveals unique and conserved biomass-degrading enzymes among lignocellulolytic microbial communities. Biotechnol. Biofuels 8 (1), 16. Murray, P., Aro, N., Collins, C., Grassick, A., Penttila¨, M., Saloheimo, M., et al., 2004. Expression in Trichoderma reesei and characterisation of a thermostable family 3 β-glucosidase from the moderately thermophilic fungus Talaromyces emersonii. Protein Expr. Purif. 38 (2), 248 257. Nakashima, K., Endo, K., Shibasaki-kitakawa, N., Yonemoto, T., 2014. A fusion enzyme consisting of bacterial expansin and endoglucanase for the degradation of highly crystalline cellulose. RSC Adv. 4, 43815 43820. Nakatani, Y., Yamada, R., Ogino, C., Kondo, A., 2013. Synergetic effect of yeast cell-surface expression of cellulase and expansin-like protein on direct ethanol production from cellulose. Microb. Cell Fact. 12, 66. Obeng, E.M., Adam, S.N.N., Budiman, C., Ongkudon, C.M., Maas, R., Jose, J., 2017. Lignocellulases: a review of emerging and developing enzymes, systems, and practices. Bioresour. Bioprocess. 4, 16. Payne, C.M., Knott, B.C., Mayes, H.B., Hansson, H., Himmel, M.E., Sandgren, M., et al., 2015. Fungal cellulases. Chem. Rev. 115, 1308. Penttila¨, M., Nevalainen, H., Ra¨tto¨, M., Salminen, E., Knowles, J., 1987. A versatile transformation system for the cellulolytic filamentous fungus Trichoderma reesei. Gene. 61 (2), 155 164.

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Praestgaard, E., Elmerdahl, J., Murphy, L., Nymand, S., McFarland, K.C., Borch, K., et al., 2011. A kinetic model for the burst phase of processive cellulases. FEBS J. 278 (9), 1547 1560. Quinlan, R.J., Sweeney, M.D., Leggio, L.L., Otten, H., Poulsen, J.C.N., Johansen, K.S. et al., 2011. Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Natl. Acad. Sci. 108(37), 15079 15084. Saloheimo, M., Paloheimo, M., Hakola, S., Pere, J., Swanson, B., Nyysso¨nen, E., et al., 2002. Swollenin, a Trichoderma reesei protein with sequence similarity to the plant expansins, exhibits disruption activity on cellulosic materials. Eur. J. Biochem. 269 (17), 4202 4211. Sampedro, J., Cosgrove, D.J., 2005. The expansin superfamily. Genome Biol. 6, 242. Santos, C.A., Ferreira-Filho, J.A., O’Donovan, A., Gupta, V.K., Tuohy, M.G., Souza, A.P., 2017. Production of a recombinant swollenin from Trichoderma harzianum in Escherichia coli and its potential synergistic role in biomass degradation. Microb. Cell Fact. 16, 0697. Schulein, M., 1997. Enzymatic properties of cellulases from Humicola insolens. J. Biotechnol. 57 (1 3), 71 81. Seki, Y., Kikuchi, Y., Yoshimoto, R., Aburai, K., Kanai, Y., Ruike, T., et al., 2015. Promotion of crystalline cellulose degradation by expansins from Oryza sativa. Planta 241 (1), 83 93. Sharma, A., Tewari, R., Rana, S.S., Soni, R., Soni, S.K., 2016. Cellulases: classification, methods of determination and industrial applications. Appl. Biochem. Biotechnol. 179 (8), 1346 1380. Singhania, R.R., Sukumaran, R.K., Rajasree, K.P., Mathew, A., Gottumukkala, L., Pandey, A., 2011. Properties of a major β-glucosidase-BGL1 from Aspergillus niger NII-08121 expressed differentially in response to carbon sources. Process. Biochem. 46 (7), 1521 1524. Singh, A., Patel, A., Adsul, M., Mathur, A., Singhania, R., 2017. Genetic modification: a tool for enhancing cellulase secretion. Biofuel Res. J. 4 (2), 600 610. Available from: https:// doi.org/10.18331/BRJ2017.4.2.5. Sipos, B., Benko, Z., Dienes, D., Reczey, K., Viikari, L., Siika-aho, M., 2010. Characterisation of specific activities and hydrolytic properties of cell-wall-degrading enzymes produced by Trichoderma reesei Rut C30 on different carbon sources. Appl. Biochem. Biotechnol. 161, 347 364. Suwannarangsee, S., Bunterngsook, B., Arnthong, J., Paemanee, A., Thamchaipenet, A., Eurwilaichitr, L., et al., 2012. Optimisation of synergistic biomass-degrading enzyme systems for efficient rice straw hydrolysis using an experimental mixture design. Bioresour. Technol. 119, 252 261. Sweeney, M.D., Xu, F., 2012. Biomass converting enzymes as industrial biocatalysts for fuels and chemicals: recent developments. Catalysts 2, 244 263. Tavagnacco, L., Mason, P.E., Schnupf, U., Pitici, F., Zhong, L., Himmel, M.E., et al., 2011. Sugar-binding sites on the surface of the carbohydrate-binding module of CBH I from Trichoderma reesei. Carbohydr. Res. 346 (6), 839 846. Teter, S.A., Sutton, K.B., Emme, B., 2014. Enzymatic processes and enzyme development in biorefining. In: Waldron, K.W. (Ed.), Advances in Biorefineries. Elsevier, Amsterdam, pp. 199 233. To¨rro¨nen, A., Harkki, A., Rouvinen, J., 1994. Three-dimensional structure of endo-1,4-beta-xylanase II from Trichoderma reesei: two conformational states in the active site. EMBO J. 13 (11), 2493 2501. Velikodvorskaia, G.A., Chekanovskaia, L.A., Lunina, N.A., Sergienko, O.V., Lunin, V.G., Dvortsov, I.A., et al., 2013. The family 28 carbohydrate-binding module of the thermostable endo-1,4-beta-glucanase CelD Caldicellulosiruptor bescii maximizes the enzyme’s activity and binds irreversibly to amorphous cellulose. Mol. Biol. (Mosk.) 47 (4), 667 673.

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Verastegui, Y., Cheng, J., Engel, K., Kolczynski, D., Mortimer, S., Lavigne, J., et al., 2014. Multisubstrate isotope labeling and metagenomic analysis of active soil bacterial communities. MBio 5 (4), e01157-14. Visser, H., Joosten, V., Punt, P.J., Gusakov, A.V., Olson, P.T., Joosten, R., et al., 2011. Development of a mature fungal technology and production platform for industrial enzymes based on a Myceliophthora thermophila isolate, previously known as Chrysosporium lucknowense C1. Ind. Biotechnol. 7 (3), 214 223. Vlasenko, E., Schu¨lein, M., Cherry, J., Xu, F., 2010. Substrate specificity of family 5, 6, 7, 9, 12, and 45 endoglucanases. Bioresour. Technol. 101, 2405 2411. Vocadlo, D.J., Davies, G.J., 2008. Mechanistic insights into glycosidase chemistry. Curr. Opin. Chem. Biol. 12, 539 555. Warden, A.C., Little, B.A., Haritos, V.S., 2011. A cellular automaton model of crystalline cellulose hydrolysis by cellulases. Biotechnol. Biofuels 4, 39. Ward, M., Penttilla, M., 2002. Swollenin, a Trichoderma reesei protein with sequence similarity to the plant expansins, exhibits disruption activity on cellulosic materials. Eur. J. Biochem. 269 (17), 4202 4211. Westereng, B., Cannella, D., Agger, J.W., Jorgensen, H., Andersen, M.L., Eijsink, V.G.H., et al., 2015. ). Enzymatic cellulose oxidation is linked to lignin by long range electron transfer. Sci. Rep. 5, 18561. Wilson, D.B., 2008. Three microbial strategies for plant cell wall degradation. Ann. N. Y. Acad. Sci. 1125, 289 297. Xu, F., Ding, H., 2007. A new kinetic model for heterogeneous (or spatially confined) enzymatic catalysis: contributions from the fractal and jamming (overcrowding) effects. Appl. Catal. A: Gen. 317, 70 81. Xu, F., 2011. Enzymatic degradation of lignocellulosic biomass. In: Tao, J., Kazlauskas, R. (Eds.), Biocatalysis for Green Chemistry and Chemical Process Development. Available from: https://doi.org/10.1002/9781118028308.ch14. Yazdi, M.T., Woodward, J.R., Radford, A., 1990. Cellulase production by Neurospora crassa: the enzyme of the complex and their regulation. Enzyme Microb. Technol. 12 (2), 116 119. Zhang, F., Bunterngsook, B., Li, J.X., Zhao, X.Q., Champreda, V., Liu, C.G., et al., 2019. Regulation and production of lignocellulolytic enzymes from Trichoderma reesei for biofuels production. In: Li, Y., Ge, X. (Eds.), Advances in Bioenergy, vol. 4. Elsevier, pp. 79 119. Zhang, M., Su, R., Qi, W., He, Z., 2010. Enhanced enzymatic hydrolysis of lignocellulose by optimizing enzyme complexes. Appl. Biochem. Biotechnol. 160, 1407 1414. Zhang, H., Wang, Y., Brunecky, R., Yao, B., Xie, X., Zheng, F., et al., 2021. A swollenin from Talaromyces leycettanus JCM12802 enhances cellulase hydrolysis toward various substrates. Front. Microbiol. 12, 658096. Available from: https://doi.org/10.3389/fmicb.2021.658096. Zhou, Q., Lv, X., Zhang, X., Meng, X., Chen, G., Liu, W., 2011. Evaluation of swollenin from Trichoderma pseudokoningii as a potential synergistic factor in the enzymatic hydrolysis of cellulose with low cellulase loadings. World J. Microb. Biot. 27, 1905 1910.

Further reading Georgelis, N., Tabuchi, A., Nikolaidis, N., Cosgrove, D.J., 2011. Structure function analysis of the bacterial expansin EXLX1. J. Biol. Chem. 2011 (286), 16814 16823.

Chapter 11

Approaches to enhance cellulase production to improve biomass hydrolysis 11.1 Introduction The bioconversion of lignocelluloses into monosaccharides is crucial for ensuring the continuous production of value-added products and biofuels. Enzymatic method, which shows a higher yield, reduced energy requirement, and improved selectivity, could be the most effective and environment friendly method for the conversion of complex lignocellulosic biomass to fermentable sugars. This is expected to make cellulase and xylanase enzymes the most demanded enzymes in industry. The widespread nature of thermophilic microbes allows them to propagate on different types of substrates and produce significant amounts of cellulase and xylanase enzymes, which makes them an important source of thermostable enzymes. Lignocellulolytic enzymes deconstruct lignocelluloses by enzymic depolymerization of holocellulose into simple sugars. Enzymatic methods offer several benefits over physical and chemical pretreatments as they are environment friendly, and do not require detrimental chemicals like acids or bases. Among all the known processes, the enzymatic hydrolysis of lignocellulosic material using cellulases and xylanases is best due to its improved specificity, and no loss of substrate. Furthermore toxic substances are not produced (Mohsin et al., 2019). Several strategies have been used for increasing cost-efficient and economically viable production of cellulases using sustainable approaches. These include use of thermostable enzymes, investigating different types of substrates and pretreatment of lignocelluloses before utilizing them for producing enzymes. The optimization of medium composition and the role of different medium constituents can be looked at. Furthermore, different process parameters have to be investigated for increasing the production of cellulases (Verma and Kumar, 2020). Also, improvement in production of cellulases via microbial fermentation processes and microbial co-production of other important products is needed to enhance the economics of the overall process. For increasing effectiveness of cellulases via reusability concept Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00003-0 © 2023 Elsevier Inc. All rights reserved.

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and use of various molecular strategies need to be investigated to enhance the activity and effectiveness of cellulases.

11.2 Thermostable cellulases Thermostable enzymes are stable and active at high temperatures. These enzymes have a wide range of industrial and biotechnological applications and are more suitable for the harsh conditions due to their thermal stability. Cellulases are the major enzyme needed for the hydrolysis of lignocellulosic materials. But, the enzymatic process has been hindered by the cost of enzymes (Ariaeenejad et al., 2021; Mukasekuru et al., 2020; Pihlajaniemi et al., 2021). The cost of enzymes for producing ethanol from corn stover was found to be $0.68/gal if the sugars could be converted at maximum theoretical yields, and $1.47/gal if the yields were based on saccharification and fermentation yields reported in the literature (Klein-Marcuschamer et al., 2012). Despite of the significant advances made in this area, another study reports the average cost of ethanol production from lignocellulosic biomass in the range from $1.91 to $3.48 per gallon ethanol (Saini et al., 2020). Gregory (2007) emphasized that the cost of enzyme should be reduced to 3 to 4 cents per gallon of ethanol before commercialization of the cellulosic ethanol technology. Improved enzyme activity at elevated temperature in the trade-off between stability and activity has been proposed as a possible strategy for reducing the dose of enzyme and therefore the cost of enzymes. These constraints could be alleviated by the use of extremely thermostable enzymes from thermophilic and hyperthermophilic microorganisms (Viikari et al., 2007; Yeoman et al., 2010). Moreover, saccharification conducted at higher temperature is very beneficial as shown in Fig. 11.1 (Ajeje et al., 2021). Concerning processes with high specificities and configurations of processes, thermostable enzymes show higher stability, appreciably reducing the amount of enzymes required for hydrolysis and reducing the reaction time (Karnaouri et al., 2019; Yang et al., 2018). At higher temperatures, a reduction in viscosity increases the diffusion of the substrate for an efficient enzymatic reaction (Bhalla et al., 2013). Thermostable enzymes enhance the rate of reaction and reduce contamination risk when comparison is made with enzymes obtained from mesophilic microorganisms. Furthermore, these enzymes improve the solubility of the substrate and recoveries of volatile compounds (Arora et al., 2015; Yadav et al., 2018). Before enzymatic treatment, lignocellulosic material is subjected to acidic or alkaline pretreatment. This step is followed by neutralization which can be avoided by the use of thermo-acidophilic and thermo-alkalophilic enzymes (Bhalla et al., 2013). Quite the opposite to mesophilic enzymes, enzymes produced from thermophilic microbes are very active and very stable in the presence of alcoholic substances, organic solvents and detergents, thus negating the use of costly

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FIGURE 11.1 Advantages of enzymatic hydrolysis carried out at elevated temperature. Ajeje S.B., Hu Y., Song G., Peter S.B., Afful R.G., Sun F., et al. 2021. Thermostable cellulases/xylanases from thermophilic and hyperthermophilic microorganisms: current perspective. Front. Bioeng. Biotechnol. 9, 794304. https://doi.org/10.3389/fbioe.2021.794304. This figure is distributed under the terms of the Creative Commons Attribution License (CC BY).

industrial catalysts which are used in diverse industries (Arora et al., 2015; Bala and Singh, 2019; Prasad, 2019). Limited information is available on thermostable enzymes with higher specific activities. Also, not much information is available on improving the production and thermostability of enzymes from thermophilic microorganisms. Thermostable enzymes are used in a variety of applications. These thermostable enzymes, have received considerable attention recently, particularly in degradation of biomass (Thapa et al., 2020). Several thermophilic fungi are found to produce highly thermostable cellulases, including Talaromyces emersonii (Murray et al., 2001), Sporotrichum sp. (Ishihara et al., 1999), Thermoascus aurantiacus (Gomes et al., 2000), and Syncephalastrum racemosum (Wonganu et al., 2008). These fungi show maximum activity in the range of 70 C80 C. Purified cellulases produced by these eukaryotic microorganisms have been characterized (Patel et al., 2019). A gene TeEgl5A that encodes a highly thermostable β-endoglucanase was isolated from a thermophilic fungi

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Talaromyces emersonii CBS394.64 and it was overexpressed in Pichia pastoris. After purification, the recombinant β-endoglucanase shows optimal relative activity at a temperature of 90 C and pH of 4.5. It has very high stability at 70 C and over a wide pH range of 1.010.0, and it is very resistant to the majority of metal ions, proteases, and sodium dodecyl sulfate. TeEgl5A possesses a wide range of substrate specificity and shows increased activity against polymers that contain β-1,4-glycosidic bonds and β-1,3-glycosidic bonds (Wang et al., 2014). Thermoascus aurantiacus possess the ability to secrete a highly thermostable cellulase for biomass deconstruction (Mohsin et al., 2019). The best cellulase producer among fungi was thought to be Trichoderma sp. but it can be susceptibility to product inhibition (Akram et al., 2018). Both novel thermostable β-glucosidases from filamentous fungi were expressed in Trichoderma reesei. They both have an optimum temperature at 60 C and pH 5.0 both enzymes were highly thermostable. Enzymes from CEL3a and CEL3b were incubated at pH 5.0; and 60 C they retained 98% and 88% of their relative activity after 6 hours of incubation (Colabardini et al., 2016). The cellulase activity of Sporotrichum thermophile (Coutts and Smith, 1976) and Talaromyces emersonii (Folan and Coughlan, 1978) is almost the same as the cellulase activity of a mesophilic fungi T. reesei. Compared to the relative cellulase activity of T. viridae, some thermophilic fungi C. thermophile, S. thermophile, and T. aurantiacus were identified to produce cellulase that is twice or thrice greater in activity (Akram et al., 2018). There have been more significant temperature stable proteins in thermophilic bacteria and hyperthermophilic archaea than in thermophilic fungi (Ajeje et al., 2021; Patel et al., 2019). Thermostable cellulases from different thermophilic microorganisms and their characteristics are shown in Table 11.1. Thermostable enzymes are found both from mesophilic and thermophilic microorganisms (Srivastava et al., 2015). Endoglucanases (mol. wt. from 30 to 100 kDa) obtained from thermophilic fungi are thermostable. These enzymes have a carbohydrate content of 2.0% to 50% and show optimum activities in the temperature range of 55 C to 80 C. Exoglucanases (mol. wt. from 40 to 70 kDa) show optimum activities in the temperature range of 50 C to 75 C. These are glycoproteins and thermostable by nature. Furthermore, the molecular weights of thermophilic cellulase enzymes show a wide range (30250 kDa) and the carbohydrates are in the range from 2% to 50%. Besides, thermophilic cellulase enzymes show excellent thermostability and are stable at 60 C and have long half-lives at 70 C, 80 C, and 90 C (Li et al., 2011). In addition, the understanding of function and nature of thermostable enzymes in fungi is relatively weaker in comparison to bacteria. So, thorough studies and characterization of amino acid residues which affect the thermophilic nature and thermal stability of fungal cellulases are immediately needed in order to have a better understanding of their

TABLE 11.1 Thermostable cellulases from various thermophilic microorganisms and their characteristics. Microorganism

Enzyme

Optimum pH

Optimum temperature

Specific activity

Thermostability/half-life

References

Geobacillus sp. T1

Cellulase

6.5

70 C

ND

Stable for 1 h at 60 C

Assareh et al. (2012)





Thermotoga naphthophila RKU-10 T

endo-1,4β-glucanase

6

90 C

1664 U mg21

Half-life of 180 min at 95 C

Akram and Haq (2020)

Dictyoglomus turgidum

β-glucosidase

5.4

80 C

160 U mg21

After incubation at 70 C for 2 h, it retained 70% of its activity

Fusco et al. (2018)

Dictyoglomus thermophilum

endo-1,4β-glucanase

5.0

50 C85 C

7.47 6 0.06 U/mg

It retained 80% of its relative activity after incubation at 50 C for 135 days

Shi et al. (2013)

Sulfolobus shibatae

endo-1,4β-glucanase

35

95 C100 C

ND

Retained 98, 90, and 84% of its activity at 75 C, 80 C, and 85 C, respectively after 120 min

Boyce and Walsh (2018)

Bacillus licheniformis A5

Cellulase

6.0

50 C

ND

Retained 82% of its activity after 120 min at 80 C

Yang et al. (2021)

Aspergillus heteromorphic

Cellulase

4.5

60 C

ND

Retained 60.0% of its activity after 1 h at 90 C

Singh et al. (2009)

Sporothrix carnis

Cellulase

5.0

80 C

ND

Retained 75% of its activity after 300 min at 80 C

Olajuyigbe and Ogunyewo (2016) (Continued )

TABLE 11.1 (Continued) Microorganism

Enzyme

Optimum pH

Optimum temperature

Specific activity

Thermostability/half-life

References

Paecilomyces thermophila

β-glucosidase

6.0

65 C

ND

Putranjiva roxburghii (PRGH1)

β-glucosidase

5.0

65 C

ND

75 C for 60 min

Kar et al. (2017)

Thermoascus aurantiacus

β-glucosidase

5.0

70 C

23.3 U/mg

It maintained 70% of its relative activity after incubation at 60 C for 1 h

Hong et al. (2007)

Geobacillus sp. HTA426

Cellulase

7.0

60 C

ND

Stable at 50 C70 C for 300 min

Potprommanee et al. (2017)

Talaromyces emersonii

β-glucosidase

71.5 C

482.8 U/mg

Half-life of 62 min at 65 C

Murray et al. (2004)

Talaromyces emersonii

Cellobiohydrolase

5.0

68 C

ND

Half-life of 68 min at 80 C

Grassick et al. (2004)

Talaromyces emersonii CBS394.64

Endo-1,4β-glucanase

4.5

90 C

ND

Highly thermostable at 70 C

Wang et al. (2014)

Yan et al. (2012)

Source: Ajeje S.B., Hu Y., Song G., Peter S.B., Afful R.G., Sun F., et al. 2021. Thermostable cellulases/xylanases from thermophilic and hyperthermophilic microorganisms: current perspective. Front. Bioeng. Biotechnol. 9, 794304. https://doi.org/10.3389/fbioe.2021.794304. This table is distributed under the terms of the Creative Commons Attribution License (CC BY).

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involvement for improving the thermostability of fungal cellulases. Thermostable cellulase enzymes are considered as best candidates for bioprocessing industries as the hydrolysis of lignocellulosic biomass with thermostable cellulase enzymes increases the rate of reaction, the diffusion coefficient, the bioavailability of organic compounds and the solubility of the substrate, reduction in the viscosity and contamination and also the cost. As a result, thermostable cellulase enzymes considerably help in complete hydrolysis reactions (Khelila and Cheba, 2014). Furthermore, thermostable cellulase enzymes having these properties can also be considered as the model system for studying temperature stability and enzymatic activity which would surely open a new opportunity for protein engineering (Haki and Rakshit, 2003). Properties of thermostable cellulases from different fungi have been reported by several researchers (Ang et al., 2013; Badal, 2002; Bakalova et al., 2002; Gaind and Singh, 2015; Grewal et al., 2017; Kupski et al., 2014; Murashima et al., 2006; Narra et al., 2014; Olajuyigbe and Ogunyewo, 2016; Pereira et al., 2015; Saqib et al., 2010; Sikandar et al., 2017; Srivastava et al., 2014; Takashima et al., 1999; Vasconcellos et al., 2015; Zhou et al., 2017).

11.3 Isolation and screening of efficient thermostable cellulase producing fungi “Thermophilic microorganisms having growth temperature ranges from 50 C to 80 C are potential candidates to produce highly active and thermostable cellulase systems (Zambare et al., 2011; Liang et al., 2011). However, several mesophilic microorganisms have also been reported to produce a considerable amount of thermostable cellulases (Gao et al., 2008). Several bacteria and fungi have been known and reported as potential producers of thermostable cellulases. Nevertheless, the latter are the most suited microorganisms for cellulose degradation and the production of cellulolytic enzyme systems (Rani et al., 2004). Additionally, fungi produce larger amount of cellulases than bacteria because of the large embedded accumulation of mycelium which is easy to separate and hence it helps to reduce the cost of separation process from the fermentation medium. Filamentous fungi characterized under the category of soft rots and white rots are preferred for the enzymatic degradation of cellulose. These filamentous fungi species produce large volumes of efficient cellulases (Himmel et al., 2007; Zhang et al., 2006). Thus, isolation and characterization of soft and white-rot fungi produce a complete cellulase system that can efficiently improve the biomass conversion process and enzyme production. Besides large volumes and selective or complete cellulase production, these species of fungi also show a wide range of temperature and pH stability which make them superior for biomass degradation (Indira et al., 2016). Thermophilic or thermostable enzymes obtained from the thermophilic or thermotolerant fungi can make the overall biofuel production

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process more flexible. Thermophilic fungi are promising sources of cellulases due to the production of cellulase at higher temperatures, high activity and thermal stability at relatively higher temperatures. Thermophilic fungi may also offer higher rates of cellulase production than the mesophile one” (Coutts and Smith, 1976; Srivastava et al., 2018). Production of cellulases from thermophilic fungi has been reported by many researchers. Cellulase production rate of thermophilic fungi such as Thermoascus aurantiacus, Sporotrichum thermophile and Chaetomium thermophile, were found to be higher by two or three times in comparison to that of mesophilic fungi such as T. viride (Acharya and Chaudhary, 2012; Folan and Coughlan, 1978). Sixteen cellulase producing thermophilic fungi from four fungal orders (three ascomycete orders Sordariales, Eurotiales and Onygenales and one zygomycete order Mucorales) were reported by Busk and Lange (2013). Production of endoglucanases and β-glucosidases from the thermophilic fungus Myceliophthora thermophila JCP 14 were reported by Pereira et al. (2015). Production of cellulases from the thermophilic fungi Myriococcum albomyces and Humicola insolens was reported by Borkar (2016). Despite the fact that, thermophilic fungi producing thermostable cellulases have been extensively reported, more studies are needed for further exploring the thermophilic fungi and the mechanisms of cellulase production at molecular level. Furthermore, isolation and screening of different thermophilic fungi from different habitats and orders are required for obtaining thermally-stable cellulase production and cost-efficient biomass conversion processes for producing biofuels.

11.4 Enhancement of thermal and pH stability of cellulases in the presence of nanomaterials Nanomaterials are expected to play a significant role in improving the thermal and pH stability of the cellulase enzymes considerably because of many exceptional chemical and physical properties (Pandurangan and Kim, 2015). The important properties of nanomaterials are presented below (Ansari and Husain, 2012; Willner et al., 2006): G G G G

High surface-to-volume ratio High catalytic efficiency High surface reaction activity Strong adsorption ability

These properties can improve the thermal and pH stability of the cellulases. Furthermore, the huge surface area of nanomaterials may contribute as the matrix for the enzyme immobilization leading to superior stability. Also, the multipoint attachment of the enzyme molecules on the surface of the nanomaterials can reduce the protein unfolding. This would result in better stability of the enzyme involved on the surface of nanoparticles (Kamyar et al., 2011; Zhu et al., 2014).

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Mukhopadhyay et al. (2012) observed improved thermostability of cellulases in the presence of hydroxyapatite nanoparticles. The production and thermal stability of cellulases were improved in the presence of nickelcobaltite (NiCo2O4) nanoparticles, Fe3O4 nanoparticles and Fe3O4/alginate nanocomposites and zinc oxide (ZnO) nanoparticles (Srivastava et al., 2014; 2015, 2016). Verma et al. (2013) observed found improved thermostability when β-glucosidases were immobilized on magnetic nanoparticles. Zhang et al. (2015a, 2015b) reported that immobilization of cellulases on functionalized magnetic nanospheres (produced by co-condensation of tetraethylorthosilicate with three different amino-silanes: 3-(2-aminoethylamino propyl)-triethoxysilane (AEAPTES), 3-(2-aminoethylamino propyl)-trimethoxysilane (AEAPTMES) and 3-aminopropyltriethoxysilane (APTES)) increased stability of cellulase enzyme. Baskar et al. (2016) used cellulase bound magnetic nanoparticles as biocatalyst for the hydrolysis of Sesbania aculeate biomass. Under optimum conditions, the maximum ethanol yield of 5.31 g/L was obtained using Sesbania aculeate biomass hydrolysate. Reusability study showed that the nanobiocatalyst was efficient for producing ethanol. Phadtare et al. (2014), conducted synthesis of polyurethane microsphereAuNP “coreshell” structures and this nanocomposite was used for the immobilizing endoglucanases. The immobilized enzyme showed better thermal stability and maintained its activity as compared to the free enzyme. The high surface area of the host AuNPs made immobilized cellulase “quasi free.” The immobilized enzyme showed better heat stability and was easier to reuse. Carbodiimide activated Fe3O4 MNPs for covalent immobilization of cellulases were used by Jordan et al. (2011). The maximum enzyme binding was 90% at loadings of 1 to 2 mg. The enzyme-to-support saturation point took place at a weight ratio of 0.02. Immobilized enzyme showed very high heat and storage stability and the optimum temperature was 50 C. The ionic forces between enzyme and support shifted the optimum pH from 4.0 to 5.0. Liao et al. (2008) used a new soluble and easily separable nanocarrier, polyvinyl alcohol (PVA)/Fe2O3 NPs for immobilizing cellulase enzyme. The immobilized cellulase retained 50% activity after using for five times. In another study, Liao et al. (2010) used PVA/Fe2O3 NPs bound cellulases along with wet ball milling for degrading microcrystalline cellulose. It was able to produce 1.89 mg/mL glucose, at least 3-times than the sum of individual yield. The immobilized cellulase enzyme retained 40% of its original activity after using for four cycles. Use of immobilized cellulase enzyme along with wet ball milling shows high potential in improving the efficiency of cellulose hydrolysis. Ho et al. (2008), developed a one-step method for producing cellulaseimmobilized NPs that made up of well-defined poly(methyl methacrylate)

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cores and cellulose shells. The immobilized cellulase showed a broader pH range and improved thermal stability. This technique has opened new possibility for the immobilization of thermostable enzyme to form nanoenzyme particles. Zhou (2009) immobilized cellulases on chitosan (CS)-coated MNPs modified by α-ketoglutaric acid. The pH range of immobilized enzyme was found to be broader and it showed lesser loss of activity than the soluble enzyme. The optimum temperature for immobilized enzyme was 50 C and that of the free enzyme was 40 C. The immobilized enzyme showed higher thermal and storage stability. Table 11.2 shows cellulases from various sources immobilized on different nanosupports and their improved properties/applications. Use of nanomaterials for improving the quality of cellulases may play an important role to make the use of such enzymatic system more cost-effective on an industrial scale.

11.5 Recombinant DNA technology for increasing cellulase activity and efficacy To further reduce the cost of cellulose-based ethanol production, improvement of microbial strains can serve as a potential method for overproduction of cellulases. “A filamentous fungus T. reesei (Teleomorph Hypocrea jecorina) is the most widely studied organism for the complete set of cellulolytic enzymes production. Subsequently, its cellulolytic potential was documented in the late 1960s, and several studies have been performed for developing the mutants capable of producing efficient cellulases using conventional mutagenesis with physical and/or chemical mutagens (Chand et al., 2005; Zhang et al., 2006). Many high cellulase-producing mutants have been reported, which are currently being used on a commercial scale. Their genetic basis responsible for enhanced production of cellulases is not known properly (Fujii et al., 2010; Zhang et al., 2006). A widely used mutant strain RUT C30 was observed lacking a genomic fragment of 85 kb and missing B30 genes that are involved in various biological processes while using genome walking in combination with complex oligonucleotides for determining the loci deletion (Seidl et al., 2008). The cloning of mutated genes using physical and chemical mutagens is a tedious process, hence, a reason for slow progress in mutations characterization at the molecular level (Gehring et al., 2000). Hence, using practical insertional mutagenesis approaches for discovering gene function in terms of cellulase formation can improve the strain properties and help understand the mechanisms responsible for the high cellulase production. The insertional mutagenesis has advantages, such as the mutated genes are tagged with inserted elements that can be used to identify the flanking sequences and the disrupted genes” (Bhardwaj et al., 2021; Jeong et al., 2002).

TABLE 11.2 Cellulases from various sources immobilized on different nanosupports and their improved properties/ applications (Husain, 2017). Name of enzyme

Name of support

Method of immobilization

Properties

References

Cellulase

CS-MNPs

Covalent Binding

High heat & storage stability

Zhou (2009)

Cellulase

CDI-Fe3O4 MNPs

Covalent binding

Better catalytic efficiency, stability and reusability

Wang et al. (2006); Jordan et al. (2011); Jordan and Theegala (2014)

Cellulase

Supermagnetic MNPs & GA

Covalent binding

High affinity and activity in a broad range of pH & temperature; effectively hydrolyzed steam-exploded corn stalks

Xu et al. (2011)

Cellulase

PVA/Fe2O3

Adsorption

Good reusability

Liao et al. (2010); Ho et al. (2008).

Cellulase

CS- Fe3O4 & GA

Covalent binding

Very high loading, stability over a broad range of pH & temperature, & reusability

Zang et al. (2014)

Cellulase

Functionalized MWCNTs

Adsorption

High binding efficiency & loading, reusability

Mubarak et al. (2014)

Cellulase from T. reesei

Activated magnetic support

Covalent binding

km decreased, High temperature-optima, good reusability

Ahmad and Sardar (2014)

(Continued )

TABLE 11.2 (Continued) Name of enzyme

Name of support

Method of immobilization

Properties

References

Cellulase

Molecular imprinted supermagnetic Fe3O4@SiO2 NPs

Adsorption

Higher catalytic efficiency & temperature optima, better thermal stability

Li et al. (2014)

Cellulase

Functionalized magnetic nanosphere, APTES & GA

Entrapment

Very high loading, stability and reusability, effective use in biofuel production

Zhang et al. (2015a, 2015b); Huang et al. (2015)

Cellulase from A. niger

β-cyclodextrin-MNPs via silanization & reductive amidation

Covalent binding

Increased continuous hydrolysis of raw straw, high biding efficiency, stability & reusability

Huang et al. (2015)

Cellulase A. fumigates

MnO2 NPs

Adsorption

High stability in a broad range of pH & temperatures, high heat stability, reusability & cellulose hydrolysis

Cherian et al. (2015)

Cellulase

AgNPs

Adsorption

Quite efficient in cellulose hydrolysis

Salunke et al. (2015)

Cellulase

AgNPs & AuNPs

Adsorption

High heat stability & reusability

Mishra and Sardar (2015).

Cellulase

CLEA-amine functionalized Fe3O4@silica coreshell MNPs

Covalent binding

Improved heat & operational stability, & reusability

Khorshidi et al. (2016)

Cellulase A. niger

TiO2 NPs & 3-APTES

Adsorption & Covalent binding

Covalently bound enzyme found superior in stability & reusability than adsorbed enzyme

Abraham et al. (2014)

Cellulase

Fe3O4@SiO2 NPs

Adsorption

High immobilization yield, halflife & reusability

Harmoko et al. (2016)

Cellulase

Vinyl functionalized cubic MS

Adsorption

NPs support far superior in activity, stability & reusability

Tao et al. (2016)

T. reesei cellulase

CS-MNPs & GA

Covalent binding

Better thermal & storage stability & very high CMC hydrolyzing reusability

S´anchez-Ram´ırez et al. (2017)

Cellulase

MNPs

Adsorption

Effective bioethanol production from Sesbania aculeate biomass

Baskar et al. (2016)

Cellulase

Nano-PEGylated GO

Covalent binding

Efficiently hydrolyzed raw straw slurry

Xu et al. (2016)

Cellulase

AF-CoFe2O4-MNPs, EDS & NHS

Covalent binding

Superior thermal stability & good reusability

Bohara et al. (2016)

Cellulase

Attapulgite@CS (ATP@CS) NC & GA

Covalent binding

High pH & heat stability & reusability, effective hydrolysis of wheat straw

Yang et al. (2016)

Cellulase

Silica-coated MNPs

Adsorption

High colloidal stability, loading & reusability

Roth et al. (2016)

Cellulase

CS-Fe3O4

Covalent binding

Improved stability and reusability

Selvam et al., 2016

Source: Husain, Q., 2017. Nanomaterials immobilized cellulolytic enzymes and their industrial applications: a literature review. JSM Biochem. Mol. Biol. 4 (3), 1029. This table is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/).

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A flow diagram of strategies used to increase the cellulase efficiency and their characteristics are presented in Fig. 11.2 (Bhardwaj et al., 2021) and Fig. 11.3 (Mondal et al., 2022). Genetic engineering is considered as the modern technology for producing cellulases on an industrial scale. Cloning and expression of genomic information available in a thermophilic organism can be expressed in a fast growing thermotolerant/ thermophilic or mesophilic organism and thereby, providing promise for producing specific stable enzymatic systems which may be effective even in crude form (Haki and Rakshit, 2003). Some thermophilic fungi are found to produce small amount of cellulases at higher temperatures, and considering this factor, thermophilic and mesophilic fungi can be a potential option for expression of thermophilic cellulases. Many types of cellulases have been overexpressed in yeasts. Pichia pastoris, have been used for producing recombinant cellulases (Lindenmuth and McDonald, 2011; Sriyapai et al., 2011). Gao et al. (2012) found that separation and purification of thermostable enzymes were easier when they were overexpressed in a heterologous host by using affinity tags, such as His-Tags, which make their production costeffective from commercial point of view (Liu et al., 2013).

FIGURE 11.2 Strategies to increase cellulase efficiency and characteristics. Bhardwaj, N., Kumar, B., Agrawal, K., Verma P., 2021. Current perspective on production and applications of microbial cellulases: a review. Bioresour. Bioprocess. 8, 95. https://doi.org/10.1186/s40643-02100447-6. This figure is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/).

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FIGURE 11.3 Genetic manipulation strategies in fungi for improved cellulase production. Reproduced with permission from Mondal, S., Halder, S.K., Mondal, K.C. (2022). Tailoring in fungi for next generation cellulase production with special reference to CRISPR/CAS system. Syst Microbiol and Biomanuf 2, 113129 (2022). https://doi.org/10.1007/s43393-021-00045-9.

Carbon catabolite repression is a specific and important mechanism which controls metabolic processes in microorganisms (Yang et al., 2015). In most fungi, Cys2His2-type transcription factor CRE1 is the key regulator of carbon catabolite repression. The function of CRE1 in Myceliophthora thermophila ATCC42464 (thermophilic fungus) was studied by Yang et al. (2015) via RNA interference. The filter paper activity and β-1,4- endoglucanase activities were found to be 3.76 and 1.31-fold higher, respectively, in comparison to the parent strain in 6 days. Also, β-1,4-exoglucanase and cellobiase activities were found to be 2.64 and 5.59-fold higher, respectively, in comparison to the parent strain in 5 days. But, in comparison to the mesophilic fungi, studies related to the regulation of cellulase gene expression in thermophilic fungi are limited and need to be thoroughly investigated. The ethanol fermenting genes such as pyruvate decarboxylase (pdc) and alcohol dehydrogenase II (adh II) were cloned from Zymomonas mobilis and transformed into three different cellulolytic bacteria, namely Erwinia chrysanthemi, Proteus mirabilis JV and Enterobacter cloacae JV, and their ethanol production capability was studied. “Recombinant E. cloacae JV was found to produce 4.5% and 3.5% (v/v) ethanol, respectively, when CMC and 4% NaOH pretreated bagasse were used as substrates, whereas recombinant P. mirabilis and E. chrysanthemi with the same substrates could only produce 4%, 3.5%, 1%, and 1.5% of ethanol, respectively. The recombinant E. cloacae strain produced twofold higher percentage of ethanol than the wild type. The recombinant E. cloacae strain could be improved further by increasing its ethanol tolerance

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capability through media optimization and also by combining multigene cellulase expression for enhancing ethanol production from various types of lignocellulosic biomass so that it can be used for industrial level ethanol production” (Piriya et al., 2012).

11.6 Use of suitable carbon source and necessity of pretreatment of lignocellulosic biomass Carbon source used in production of enzyme could account for more than 50% of the total enzyme cost, if it were pure glucose (Ellila¨ et al., 2017). The rising demand for cellulases in several industries has prompted researchers to explore several resources which can reduce the cost of production and be sustainable. “Microbial strains can be used to produce cellulase enzymes utilizing LCB obtained from forestry and agricultural residues as costeffective raw materials that are abundantly available in nature (Kumar et al., 2008). Lignocellulosic plant residues can be an effective alternative to the costly carbon sources as they are cheap, renewable, abundant, and good nutrients sources for the microorganisms involved in cellulase production (Saini et al., 2017). Many lignocellulosic residues, such as wheat bran, sugarcane bagasse, rice straw, wheat straw, grape stalk, seeds, fruit pomace, corn cob, and soy bran, have been evaluated in the production of cellulases (Jampala et al., 2017; Masutti et al., 2015). Further, variation in atmospheric conditions results in different plant diversity and compositional structure, thus causing variations in the available waste. In Brazil, sugarcane bagasse is an effective alternative to the expensive carbon sources due to its large availability at the sugarcane mills (Vasconcellos et al., 2015). Additionally, the substrates used in the production process act as an enzyme inducer, and their source microorganism may produce enzymatic cocktails with diverse catalytic abilities for cellulose breakdown (Li et al., 2017; Pandey et al., 2016). Similarly, lignocellulosic substrates can be used as a suitable alternative to commercial inducers by producing a number of enzymes leading to better cellulose hydrolysis (Cunha et al., 2012). The use of waste biomass for cellulase production could reduce the production cost and partly address the environmental LCB disposal problems (Gomes et al., 2015). Although celluloses are available in considerable quantities in the LCB, their accessibility to microbes is poor due to their natural recalcitrance properties in biomass. Thus, before using LCB as the substrate for cellulase production, it can be subjected to pretreatment to improve cellulose accessibility to microorganisms (Saini et al., 2017). After optimization and a suitable fermentation process, pretreatment of biomass can enhance the production of enzymes” (Bhardwaj et al., 2021). The major objective of pretreatment is to overcome the recalcitrance of lignocellulosic biomass, for separating the cellulose from the matrix polymers, and to make it more accessible for enzymatic hydrolysis

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(Rodriguez-Zuniga et al., 2014). Pretreatment is the most expensive step in the conversion of lignocellulosic substrate to ethanol and accounts for almost $0.30/gallon of ethanol produced (Houghton et al., 2006). Pretreatment technologies are generally classified into physical, chemical, physicochemical, and biological. The use of these methods mostly depends upon the physicochemical characteristics of the raw materials. Chemical and thermochemical pretreatment methods, are generally used for many applications (Alvira et al., 2010). The main problem associated with the pretreatment methods is that the inhibitors, such as furfural and 5-hydroxymethylfurfural, etc. are generated (Scordia et al., 2013). Therefore, the type of pretreatment method greatly affects the growth of fungi as the inhibitors inhibit the growth of fungi. Many efforts have been made to minimize the inhibition (Almeida et al., 2007). Improved production of enzyme was reported by Vasconcellos et al. (2015). The phenolic compounds known inhibitors of cellulase production, were removed. Sonication treatment of fermentation broth has been proposed for improved production of cellulases, xylanases, and pectinases using different types of microorganisms (Delgado-Povedano and De Castro, 2015; Jalal and Leong, 2018). Improved production of glucose from Bacillus licheniformis α-amylase was reported by using ultrasound treatment at 100% amplitude for 1 minute to the sorghum grain slurry (Shewale and Pandit, 2009). The amylase enzyme obtained by using this method showed improved amyloglucosidase enzyme saccharification by 8%.

11.7 Optimization of medium and process parameters using statistical methods for improved production of cellulases Several studies have been conducted for increasing the production of cellulases using one factor at a time (OFAT) and different statistical approaches (Bhardwaj et al., 2017). Optimization of the media constituents and the physicochemical process involved in the production of cellulases improved enzyme yield. A threefold increase in cellulase production was achieved by Response Surface Methodology (RSM) model which is a well-known statistical tool (Shajahan et al., 2017). RSM was used for optimizing the cellulase production for evaluating the interactions of independent physicochemical parameters in Schizophyllum commune COC and A. aneurinilyticus strain BKT-9 (Kumar et al., 2018; Srinubabu et al., 2007). RSM is a multivariate strategy with several advantages like lesser number of investigational runs, better explanation of the statistical potentials, and individual and interactive properties of the involved parameters. “The central composite design is the most commonly used fractional factorial design used in the response surface model. In this design, the center points are augmented with a group of axial points called star points.

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With this design, quickly first-order and second-order terms can be estimated” (Bhattacharya, 2021). Srinubabu et al. (2007) optimized the environmental and nutritional factors for production of cellulases from bacteria obtained from Dal Lake, urban freshwater Himalayan Lake. A new cellulase-producing bacteria Bacillus licheniformis KY962963, an epiphytic bacteria of marine algae Chlorococcum sp. were reported by Shah et al. (2021). The nutritional and ecological factors for improved production of cellulases using Bacillus licheniformis KY962963 were optimized using the simultaneous application of “PlackettBurman” design and OFAT approach. It was found that moisture content of 75%, dipotassium hydrogen phosphate concentration of 2 g/L, incubation time and temperature of 3 days and 35 C, were found to be optimum for improved production of cellulases.

11.8 Improvements in production of cellulases via microbial fermentation processes The solid-state fermentation (SSF) and submerged fermentation (SmF) are the two major processes used for the production of cellulases and other enzymes such as xylanases, proteases, and pectinases using different microorganisms (Mrudula and Murugammal, 2011; Pant et al., 2015). Lignocellulosic biomass can be used as substrates which act as an inducer for enzyme production during fermentation. SmF possesses many benefits in process control as abundant water reduces the temperature, and nutrients and oxygen concentration gradients. Also the enzyme recovery is easier in SmF. Furthermore, SmF process can serve as a successful technology for developing the fermentation processes for producing enzyme on a commercial scale (Colla et al., 2016; Hansen et al., 2015). In SSF, the degradation of an organic substrate takes place aerobically in the absence of free water for producing the desired end-product. The optimum parameters differ widely based on the fermentation process. Also, different type of end products can be produced by the use of same substrate in varying operational conditions or by using different types of microbial strains (De Castro and Sato, 2015). SSF method is found to be economical for the production of different important industrial enzymes and many other biochemicals by the use of lignocellulosic biomass as substrate (Uncu and Cekmecelioglu, 2011). There are many reports for cost-effective production of cellulases from lignocellulosic biomass using SSF as substrate (Dhillon et al., 2012). SSF process requires simple equipment with reduced energy requirement resulting in higher enzyme yield and substantially lesser processing cost. So, various approaches have been used for reducing the production cost of enzyme, such as SSF using lignocellulosic biomass as substrate and enzyme inducer.

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11.9 Microbial co-production of other important enzymes for the overall economy of the process Lignocellulosic biomass is rich in lignin, cellulose, and hemicellulose constituents and many decomposer fungal strains are able to produce cellulases, hemicellulases and ligninolytic enzymes concurrently for their natural growth. Therefore, simultaneous production of lignocellulolytic enzymes has been proposed for reducing the overall cost of lignocellulolytic biorefinery. Simultaneous production of cellulases and xylanases from T. reesei NCIM 1186 and its improvement via using desirability-based multi-objective optimization method was suggested by Jampala et al. (2017). Ultrasound treatment of fermentation broth has a positive effect on the co-production of cellulolytic, hemicellulolytic, pectinolytic and fibrinolytic enzymes using different microorganisms (Avhad and Rathod, 2015; Delgado-Povedano and De Castro, 2015; Jalal and Leong, 2018). Avhad and Rathod (2015) reported that use of ultrasonication treatment for inducing the enzyme production from microorganisms, increased yield of fibrinolytic enzyme. The process involved 12 hours of growth of Bacillus sphaericus MTCC 3672 with optimized ultrasound irradiation at 25 kHz for 10 minutes with 40% duty cycle and 160 W power. Optimized ultrasonication treatment resulted in 1.48-fold increase in fibrinolytic enzyme yield as compared to nonsonicated fermentation. The effect of the ultrasonic treatment in improving the fermentative coproduction of cellulases, xylanases, and pectinases enzymes from Bacillus subtilis ABDR01 has been reported by Yadav et al. (2020). Treatment of fermentation broth at ultrasound power of 90 W using 25 kHz frequency with 70% duty cycle for 5 minutes gave the maximum production of cellulases (22.17 U/mL), xylanases (137.95 U/mL), and pectinases (87.82 U/mL) at the short bacterial growth phase of only 6 hours. Ultrasonic treatment of fermentation broth caused cell cluster disaggregation and improves the uptake of nutrients along with maintaining the cellular integrity of microorganisms. This process significantly increased the biomass concentration and end-product of the fermentation process.

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Chapter 12

Future prospects The interest in biofuels is growing rapidly because of the reasons of energy security, diversity, sustainability and reduction in greenhouse gas emissions. It is expected that biofuel will play a major role in meeting the world’s energy requirement in the future. Biofuels have been broadly accepted as alternative fuels for the transportation sector for enhancing the performance of transport vehicles (Bajpai, 2021). In the recent years, United States has passed regulations and implemented aggressive goals for encouraging increased use of biofuels. Individual States in United States particularly California are taking even stronger positions with respect to biofuels. Preliminary endeavors focused mostly on bioethanol, produced by fermentation of sugars from grains (particularly corn) (Hoekman, 2009). Now research is focusing on “Second Generation Biofuels” which are produced from different types of biomass resources using a broad range of conversion technologies. Over many years of development, some of the leading producers of biofuel in the world have covered a lot of ground in developing and improving the used technologies. The first-generation technologies developed in the beginning provided manufacturing of the three major types of biofuel: bioethanol, biodiesel, and biogas from starch-containing agricultural products as feedstocks. At the start of the 21st century, many new strategies were developed, which led to the expansion of biofuel production and appearance of second- and the third-generation biofuel (Guo et al., 2015; Lopes et al., 2016; Saracevic et al., 2019; Titova, 2019). In the recent years, important developments have been made for improving the production and efficacy of cellulases for low-priced biofuel production. Although cellulases have various applications in the industry, improving the efficacy, reduction in cost and energy requirements are always important parameters for producing biofuels from cellulosic biomass. Isolation, screening, and cultivation of thermophilic/thermotolerant fungi for obtaining thermotolerant/ thermostable cellulases on an industrial scale for producing biofuels are still a roll-back factor. Production of enzyme by submerged fermentation is also costly for production of biofuel from biomass. Aside from enzyme purification and separation of end products, incomplete understanding of protein engineering and its implementation are also adding an extra impediment in the overall Cellulases in the Biofuel Industry. DOI: https://doi.org/10.1016/B978-0-323-99496-5.00010-8 © 2023 Elsevier Inc. All rights reserved.

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process of bioconversion. So, there is requirement of extensive research on the production and optimization of cost-effective cellulase enzymes from potential fungi using solid state fermentation and biomass as an inexpensive process. Furthermore, study on more thermostable cellulases their stability and protein engineering could also support for obtaining novel and more inexpensive processes for production of biofuel in comparison to the existing processes (Srivastava et al., 2018). Top enzyme producers are extensively working to provide cost-efficient and superior cellulase and hemicellulase cocktails for making cellulosic biofuel a realism. An improved product called Cellic Ctec3 was launched in 2014 by Novozymes. This was an improved version from the earlier Cellic Ctec2 version. The efficiency of production of biomass from ethanol was 1.5 times higher than the earlier version. About 50 kg of the new cocktail is required for making one ton of ethanol from biomass (https://www.novozymes.com/en/news/news-archive/2012/02/advanced-biofuels-becoming-reality-with-novozymes-new-enzyme-technology). Extensive research has been conducted during the last decade to find reasonably priced, sustainable supply of biofuels. Although several issues remain to be solved, good progress has been made which is judged by the large projects initiated and the plants being built. Further research is focusing on developing better quality, inexpensive cocktails of cellulases/hemicellulases. “However, the success of the enzymatic approach to second-generation biofuels is not dependent solely on enzyme producers. It will also depend greatly on how those enzymes are best deployed and, thus, equally heavy reliance is being placed on all the auxiliary processes from feedstock supply and pretreatment to opening up markets in the automobile industry for ethanol addition” (Carrigan, 2016). The lignocellulosic biomass, being the most abundant and bio-renewable biomass on earth, has received significant attention because of its noncompetitiveness with food supplies. Thus, lignocellulosic biomass is considered as the most suitable substrate for producing biofuels. “The high-value products from lignocellulosic biomass increase profitability; the high volume fuels help meet the energy demands and reduce GHG emissions from traditional power plant facilities. Though NREL has made path-breaking discoveries and inventions in lignocellulosic biomass biorefineries, the technology is still in nascent stage. The considerable challenges remain to be addressed in the economical development of lignocellulosic biomass biorefinery. The challenges are faced in every unit operation (step) such as sustainable biomass supply chains, biomass pretreatment, fractionation, saccharification, and conversion of sugars and lignin to fuels and chemicals. One of the critical hurdles is the cost of feedstock, which is dependent on the supply chains and hence, careful analysis and research on the current supply chains would greatly contribute to improve the efficiency of currently available supply chain pathways. In addition, transportation of lignocellulosic biomass, pests,

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diseases, and land use have made the 2G biofuels expensive. The pretreatment step is one of the most cost contributing and rate- and yield-limiting steps. Pretreatment is a physical, chemical, physicochemical, or biological process which opens up the recalcitrance structure amenable to enzymatic attack. However, none of the pretreatment strategies was found to be eco-friendly and environmental friendly and hence, pretreatment of the biomass seems to be still unresolved problem. The future of lignocellulosic biomass conversion to biofuels is expected to lie in the development of cost-effective pretreatment technologies for efficient hydrolysis of lignocellulosic biomass even with using less enzyme doses. In addition, substantial reduction in the chemicals needed for pretreatment is necessary to have an economically/environmentally competitive process. Though many efforts have been devoted to developing enzyme cocktails, suitable and low-cost enzyme cocktails still remain as challenges in biomass processing. Different enzyme sources need to be tapped and evaluated for designing customized enzymatic cocktails. Metagenomic tools as well as bioprospection of the microbes could be applied to discover new sources of enzymes. From industrial point of view, the knowledge of enzymatic synergism in the enzyme cocktails is of great importance which may lead to rapid conversion of biomass, reducing the enzyme loads and thereby reducing the cost of lignocellulosic biomass hydrolysis. The recent publications on GVL-pretreatment and CelA-type enzymes acting on raw biomass may help meet these challenges to make the biomass conversion technologies eco-friendly with affordable cost. Though cellulosic ethanol has been recognized as a major biofuel, alternative fuels such as alkanes and alkenes are of particular interest since they are superior biofuels to ethanol due to highenergy content (30%). Efforts are being made to recommercialize the butanol production from CBE process using engineered Clostridium strains. Finally, the integration of all the processes including physical, chemical, and biological to produce bulk as well as high-value chemicals from all the three components of lignocellulosic biomass may be the appropriate strategy to reduce the cost of ethanol production” (Singhvi, and Gokhale, 2019). Production of bioethanol from lignocellulosic biomass is not cost-effective at the moment. There are some barriers which need to be addressed for efficiently converting lignocellulosic biomass to ethanol. These include biomass feedstock, pretreatment, saccharification of cellulose and hemicellulose matrix, and simultaneous fermentation of hexoses and pentoses. Regarding biomass feedstock major obstructions are supply, cost, handling, and harvesting. The main obstacles with respect to conversion technology involve processing of biomass, convenient and cost-efficient pretreatment process for detaching cellulose and hemicelluloses from their complex with lignin. Another problem is to develop a satisfactory hydrolysis process for depolymerization of cellulose and hemicelluloses for producing higher amounts of simple sugars. Enzymatic hydrolysis is considered as a potent phenomenon for saccharification of complex polymer. Presently, research and development is focusing on cellulase engineering for

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FIGURE 12.1 Biomass-to-biofuels supply chain (Ying et al., 2020; https://doi.org/10.3390/ en13071799). Ying, H.P., Phun Chien, C.B., Yee Van, F., 2020. Operational management implemented in biofuel upstream supply chain and downstream international trading: current issues in Southeast Asia. Energies 13 (7), 1799. Distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/).

reducing the cost of ethanol production. Finally, the challenges associated with fermentation process are co- fermentation of pentoses and hexoses besides the use of potent and efficiently engineered microbial cultures. Further work should focus on inhibiting the generation of inhibitors (furfural, acetic acid) during biomass processing which acts as major obstacles to reduce the hydrolysis efficiency and fermentation of biomass. For combating the challenges associated with lignocellulosic biofuel technology, novel strategies such as genetic engineering, cellulase engineering should be used so that technology for production of ethanol from lignocellulosic biomass may be effectively developed, optimized, and commercialized in the near future. Development of a robust and sustainable biofuels industry will require a large and complex supply chain (US DOE, 2007; Ying et al., 2020). Five major elements in this supply chain are shown in Fig. 12.1.

12.1 Feedstock production Large industries must develop to produce sufficient volumes (and sustainable supplies) of biomass feedstocks. Waste materials can contribute a significant portion initially, but with increasing demand, more deliberately grown feedstocks will be required. Bioengineering will be necessary to develop improved crops (lower water requirements, faster growth, disease resistant, improved harvestability, etc.).

12.2 Feedstock logistics New crops (switchgrass, forest thinnings, algae, etc.) require development of equipment and methodologies for cost-effective harvesting, storage, and

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preprocessing. Transportation costs are a current barrier. Year-around production of biofuels requires constant, reliable feedstock supply. Effective separation of waste feedstocks remains problematic.

12.3 Biofuels production Further processing improvements are necessary for cost-effective operation. Integrated biorefineries must be developed and deployed. Effective utilization of coproducts (including heat and electrical power) is necessary for economic operation.

12.4 Biofuels distribution Compatibility with current infrastructure components (storage, blending, transportation, and dispensing) is critical. Quality control procedures must be implemented to ensure that biofuels meet all applicable product specifications.

12.5 Biofuels end use Availability of biofuels-compatible vehicles must be expanded. To encourage consumer satisfaction, these vehicles must offer performance equivalent to conventional vehicles. A final, overriding consideration in this entire supply chain is economics. Biofuels to become commercially successful on a large scale will require favorable economics at each point along this chain (Hoekman, 2009). The biofuels industry is undergoing rapid growth and transformation. Driven largely by concerns over energy security and greenhouse gases, national and state policies are being developed and implemented to promote much greater utilization of biofuels. At the present time, biofuels are dominated by corn derived ethanol, but much greater emphasis is now being directed toward Second Generation Biofuels produced from lignocellulosic material and triglycerides. Both thermochemical and biochemical conversion technologies for producing biofuels are advancing rapidly. Further improvements are expected in the near future as a result of substantial investments by both public and private sectors (Hoekman, 2009). No single best strategy for biofuels is expected to materialize, as optimization depends upon the regional feedstocks, existing infrastructure, integration with other industrial plants (including petroleum refineries) and other considerations which vary from place-to-place. The natural resources are adequate to produce considerable quantity of second generation biofuels which would possibly be sufficient to displace 30% to 50% of current petroleum-derived fuels over the next 30 to 50 years. But, great care must be taken for ensuring that this transformation to biofuels will be affordable and sustainable with negligible undesirable environmental impact.

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Cellulases in the Biofuel Industry

References Bajpai, P., 2021. Developments in Bioethanol. Green Energy and Technology. Springer, Singapore. Available from: https://doi.org/10.1007/978-981-15-8779-5_11. Carrigan, J., 2016. Chapter 15 Applications of cellulase in biofuel industry. In: Gupta, V.K. (Ed.), New and Future Developments in Microbial Biotechnology and Bioengineering, Elsevier, pp. 177 184. Guo, M., Song, W., Buhain, J., 2015. Bioenergy and biofuels: history, status, and perspective. Renew. Sustain. Energy Rev. 42, 712 725. Hoekman, S.K., 2009. Biofuels in the U.S. challenges and opportunities. Renew. Energy 34 (1), 14 22. Lopes, M.L., Paulillo, S.C., Godoy, A., Cherubin, R.A., Lorenzi, M.S., Giometti, F.H., et al., 2016. Ethanol production in Brazil: a bridge between science and industry. Braz. J. Microbiol. 47 (Supplement 1), 64 76. Saracevic, E., Fru¨hauf, S., Miltner, A., Karnpakdee, K., Munk, B., Lebuhn, M., et al., 2019. Utilization of food and agricultural residues for a flexible biogas production: process stability and effects on needed biogas storage capacities. Energies 12, 2678. Srivastava, N., Srivastava, M., Mishra, P.K., Gupta, V.K., Molina, G., Rod´rıguez-Couto, S., et al., 2018. Applications of fungal cellulases in biofuel production: advances and limitations. Renew. Sust. Energy Rev. 82, 23792386. Singhvi, M.S., Gokhale, D.V., 2019. Lignocellulosic biomass: hurdles and challenges in its valorization. Appl. Microbiol. Biotechnol. 103, 9305 9320. Titova, E.S., 2019. Biofuel application as a factor of sustainable development ensuring: the case of Russia. Energies 12 (20), 3948. US DOE, 2007. Biomass multi-year program plan. US DOE Office of the Biomass Program, October 2007. Ying, H.P., Phun Chien, C.B., Yee Van, F., 2020. Operational management implemented in biofuel upstream supply chain and downstream international trading: current issues in Southeast Asia. Energies 13 (7), 1799. Available from: https://doi.org/10.3390/en13071799.

Relevant websites https://www.novozymes.com/en/news/news-archive/2012/02/advanced-biofuels-becoming-reality-withnovozymes-new-enzyme-technology. https://www.eesi.org/files/mypp_april_2011.pdf.

Further reading Gamage, J., Howard, L., Zisheng, Z., 2010. Bioethanol production from lignocellulosic biomass. J Biobased Mater Bioenergy 4, 3 11.

Index Note: Page numbers followed by “f” and “t” refer to figures and tables, respectively.

A Amorphogenesis-inducing agents, 197

B

β-glucosidase cellulase enzyme complex, 200 cellulolytic filamentous fungus T. reesei, 201 202 P. decumbens, 201 Biochar, 60 Biodiesel, 56 57 Bioenergy, 2 5 Bioethanol distillation, 155 156 fermentation, 149 155 hydrolysis, 147 149, 154t lignocellulosic-based bioethanol fermentation, 151t pretreatment, 144 147 effect on lignocellulosic biomass, 146f production, 109 biofuel process, 141f lignocellulosic biomass conversion into ethanol, 141f processing route for, 145f saccharification and fermentation, 154t Biofuels, 2 5 benefits of, 4t compared to gasoline, 5t cost, chemicals and biofuels, 79f distribution, 245 end use, 245 first-generation, 1 2 made by assistance from enzymes, 167 168 production, 245 Biomass, 5 11 cellulose polymer chain structure, 9f hemicellulose structure, 10f

lignocellulosic, 6f, 7f macromolecular structure of lignin, 10f Biomass feedstocks, 3t Biomass-to-o-biofuels supply chain, 244f

C Cellic Ctec3, 114 115, 242 Cellulase auxiliary enzymes encoding genes present in microorganisms, 206t lytic polysaccharide monooxygenases (LPMOs), 204 205 M. thermophila, 205 and synergistic proteins in hydrolysis of cellulose, 207f Cellulase enzyme, 11 18 actinomycetes having cellulolytic abilities, 18t bacteria having cellulolytic abilities, 17t bioprocess of cellulase production, 18f classification of, 12t for degrading cellulose, 15f fungi having cellulolytic abilities, 16t synergistic action of cellobiohydrolases, 14f Cellulase enzyme systems and accessory proteins, 127 128 for cellulose hydrolysis, 121 128 from different bacteria, 122t endoglucanase, 123 124 exoglucanase, 124 126 family 6 endoglucanase and exoglucanase, 125f hydrolysis of cellulose by noncomplexed, 123f Cellulase market scenario Cellic CTec3 HS, 114 115 global market, 109 international manufacturing, 112 market scenario, 110 115 sources and suppliers, 113t

247

248

Index

Cellulase production to improve biomass hydrolysis, enhancing DNA technology, 222 228 isolation and screening of efficient thermostable cellulase, 219 220 microbial co-production, 231 via microbial fermentation processes, 230 pretreatment of lignocellulosic biomass, 228 229 process parameters using statistical methods, 229 230 suitable carbon source, 228 229 thermal and pH stability of, 220 222 thermostable cellulases, 214 219 Cellulases and auxiliary enzymes cellobiohydrolase, 198 199 cellulase auxiliary enzymes, 202 207 encoding genes, 206t endo-1,4-β-glucanase, 199 200 synergistic proteins in hydrolysis, 207f Cellulases for biofuels production bioethanol production, 140 156 factors affecting bioethanol, 156 lignocellulosic biomass conversion into ethanol, 141f process, 141f processing route for bioethanol, 145f synergistic action of, 140, 140f Cellulases in cellulose hydrolysis roles degradation, enzymes in, 120f enzyme systems for, 121 128 synergy among degrading system, 128 132 Cellulases via microbial fermentation processes, 230 Cellulosomes, 121 Challenges to biofuel production biofuel cost, 78 79 energy and environmental issues, 80 82 enzymatic hydrolysis, 74 77 feedstock production and logistics, 68 71 generation of coproducts, 80 lignocellulosic biomass conversion to biofuels, 68 82 pretreatment, 71 74 microbial fermentation and biomass, 77 78 water recycling, 79 80 Conventional biofuels, 53, 56 57 Current production status of cellulases and challenges sequential solid-state fermentation, 99 101 solid-state fermentation, 96 98 submerged fermentation, 99 101

submerged fermentations, 93 96 use of mixed cultures, 102 103

D Dextrin phosphorylase, 127

E Energy and environmental issues, 80 82 Enzymatic hydrolysis cost area distribution, 75t Trichoderma reseei, 76 Exocellulase, 126

F Feedstock logistics, 244 245 Feedstock production, 244 biomass supply chain, 70f and logistics, 68 71 First generation biofuels, 56 58 Fourth generation biofuels, 62 63 bioethanol production based on, 64f production, 63f

G Generation of coproducts, 80 Generations of biofuels, 53, 54f characteristics, 55f first generation, 53, 56 58 fourth generation, 56, 62 63 second generation, 53 56, 58 60 third generation, 56, 61 Greenhouse gas emissions, 5t

I Isolation and screening of thermostable cellulase producing fungi, 219 220

L Lignocellulosic biomass conversion to biofuels/biocemicals, 68 82 Lignocellulosic biomass pretreatment, 71 74 necessity of, 228 229

M Microbial fermentation and biomass, 77 78 Molecular scissors, 74 76

Index

P pH stability of cellulases, enhancing, 220 222 Production of cellulosic ethanol case study, 159 167 Clariant Sunliquid plant Podari, Romania, 161 165 Praj’s second generation (2G) cellulosic ethanol plant, 166 167 Project ABBK in Hugoton, Kansas, USA, 159 160 developments in, 157 167 Production processes of cellulases, 180 189 Bacillus subtilis, 187 benefits of thermostable cellulases, 181t bioconversion of lignocellulosic materials, 180 Bjerkandera adusta, 186 cellulosome structure and assembly, 188f engineer enzymes, 187 189

R Recombinant DNA technology for increasing cellulase activity and efficacy, 222 228

S Screening of efficient thermostable cellulase producing fungi isolation and, 219 220 production of cellulases, 220 Second generation biofuels, 58 60 biochemical conversions of, 59f raw feedstock for, 57f yield of biofuels from different feedstocks of, 60t Separate hydrolysis and fermentation (SHF), 148 Sequential solid-state fermentation cellulase production under submerged, 100t commercial production of cellulases, 102f sequential SSF and SmF, 101f and submerged, 99 101 Simultaneous saccharification and fermentation (SSF), 148 Solid-state fermentation (SSF), 230 advantages and disadvantages of, 98, 99t cellulase production, 97 high value added products, 96 97 operations, 98

249

Status of biofuel production Argentina, 39 Australia, 41 43 annual production, 41 commercial-scale bioethanol plants in, 43t Brazil, 33 37 first commercial-scale cellulosic ethanol plant, 36 GranBio plans, 36 operating cellulosic ethanol plants in the US, 34t Canada, 43 45 Clean Fuel Standard, 43 44 Greenfield Global, 44 45 China, 39 41 bioethanol plants, 39 40 fuel ethanol consumption, 41 European Union, 37 39 German Association of Biodiesel Producers, 38 renewable energy use, 37 38 India, 47 49 biofuel key figures, 49t first second generation ethanol biorefinery, 48 largest sugar producer, 47 second generation bioethanol, 48 Japan, 46 Thailand, 45 46 production of ethanol in, 45 46 United States, 28 33 annual US fuel ethanol production, 29t biorefinery count and production capacity, 28t global commercial scale cellulosic ethanol plants, 31t US biodiesel plant production capacity, 30t US commercial lignocellulosic ethanol facilities, 33t US fuel ethanol plant production capacity, 29t Submerged fermentation (SmF), 230 batch process, 94 fed-batch process, 94 rotating fibrous-bed reactor (RFBB), 95 96 stirred-tank reactors (STRs), 95 Swollenin, 185 186 Synergism, 128

250

Index

Synergy among cellulose degrading system CelY and CelZ, 129 exoglucanases and endoglucanases, 130 131, 131f Humicola insolens, 129 synergism forms, 128 Thermobifida fusca, 129

thermo-acidophilic and thermo-alkalophilic enzymes, 214 216 Trichoderma reesei, 216 Third generation biofuels, 61 algae, 61 generation from microalgae, 62f IEA definition, 61

T

W

Thermal stability of cellulases, enhancing, 220 222 Thermostable cellulases, 181 concerning processes, 214 216 hydrolysis carried out at elevated temperature, 215f

Water recycling, 79 80 Worldwide scenario of biofuel production annual world fuel ethanol production, 27t production status, 28 49 renewable ethanol production by end use, 27f yields by feedstock, 26t