Liquid biofuel production 9781119459835, 1119459834, 9781119459859, 1119459850, 9781119459866, 1119459869

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Liquid Biofuel Production

Scrivener Publishing 100 Cummings Center, Suite 541J Beverly, MA Publishers at Scrivener Martin Scrivener ([email protected]) Phillip Carmical ([email protected])

Liquid Biofuel Production

Edited by

Lalit Kumar Singh and Gaurav Chaudhary

This edition first published 2019 by John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA and Scrivener Publishing LLC, 100 Cummings Center, Suite 541J, Beverly, MA 01915, USA © 2019 Scrivener Publishing LLC For more information about Scrivener publications please visit www.scrivenerpublishing.com. All rights reserved. No part of this publication may be reproduced, stored in a retrieval system, or transmitted, in any form or by any means, electronic, mechanical, photocopying, recording, or otherwise, except as permitted by law. Advice on how to obtain permission to reuse material from this title is available at http://www.wiley.com/go/permissions. Wiley Global Headquarters 111 River Street, Hoboken, NJ 07030, USA For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Limit of Liability/Disclaimer of Warranty While the publisher and authors have used their best efforts in preparing this work, they make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation any implied warranties of merchantability or fitness for a particular purpose. No warranty may be created or extended by sales representatives, written sales materials, or promotional statements for this work. The fact that an organization, website, or product is referred to in this work as a citation and/or potential source of further information does not mean that the publisher and authors endorse the information or services the organization, website, or product may provide or recommendations it may make. This work is sold with the understanding that the publisher is not engaged in rendering professional services. The advice and strategies contained herein may not be suitable for your situation. You should consult with a specialist where appropriate. Neither the publisher nor authors shall be liable for any loss of profit or any other commercial damages, including but not limited to special, incidental, consequential, or other damages. Further, readers should be aware that websites listed in this work may have changed or disappeared between when this work was written and when it is read. Library of Congress Cataloging-in-Publication Data ISBN 978-1-119-45987-3 Cover image: Levulinic acid compound | Ibreakstock | Dreamstime.com Waste Plant | Belish | Dreamstime.com Cover design by Kris Hackerott Set in size of 11pt and Minion Pro by Manila Typesetting Company, Makati, Philippines Printed in the USA 10 9 8 7 6 5 4 3 2 1

Contents 1

Process Engineering Biofuel Production Opubo Gbanaye Benebo 1.1 Biofuel Production Background 1.1.1 General Limitations 1.1.2 Limitation of Cashcrop Raw Material 1.1.3 Limitations of Algae Raw Materials Remediation 1.1.4 Limitations Remediation 1.2 Process Engineering Liquid Biofuel Production 1.2.1 Algae Cultivation Assessment 1.2.2 Algal Cultivation Inefficiencies Remediation 1.2.3 Technology Development 1.2.4 Lessons from the Algae Biofuel Industry Collapse 1.2.5 Process Development Norms 1.2.6 Research Team 1.2.7 Alga Cultivation General Issues 1.2.8 Biofuel Process Technology 1.3 Algal Cultivation Process Technology 1.3.1 Cellular Reaction Kinetics Analysis 1.3.2 Cultivation Bench-Scale Model Design 1.3.3 Cultivation Bioreactor 1.3.4 Concentrator Harvesting of Cells 1.3.5 Cell Rupture Technology 1.3.6 BioFeedstock Separation Process 1.3.7 Bench-Scale Cultivation Process Technology 1.3.8 Process Technology Financial Viability Design 1.3.9 Process Technology Sustainability Engineering 1.3.10 Process Technology Optimization Engineering 1.3.11 Base Cultivation Process Technology

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Contents 1.4 Algal Biomass Biorefinery Process Engineering 1.4.1 Resourcing Algal Biomass 1.4.2 Microbes Nutrients-Feed Production 1.4.3 Fermentation Process Technology 1.4.4 Biodiesel Process Technology 1.4.5 Biorefinery Process Technology 1.4.6 Engineering Cost Impact Analysis Acknowledgment About the Author References

2 A Renewable Source of Hydrocarbons and High Value Co-Products from Algal Biomass Abhishek Walia, Samriti Sharma and Saruchi 2.1 Introduction 2.2 Algal Biomass Production 2.2.1 Growth Conditions 2.2.1.1 Temperature 2.2.1.2 Light Intensity 2.2.1.3 pH 2.2.1.4 Aeration and Mixing 2.2.1.5 Salinity 2.2.2 Photoautotrophic Production 2.2.2.1 Open Pond Production Pathway 2.2.2.2 Closed Photobioreactor Systems 2.2.3 Harvesting and Dewatering of Algal Biomass 2.2.3.1 Flocculation 2.2.3.2 Chemical Flocculation 2.2.3.3 Electroflocculation 2.2.3.4 Biofloculation 2.2.3.5 Magnetic Separation of Algae 2.2.3.6 Dissolved Air Flotation 2.2.3.7 Filtration 2.2.3.8 Centrifugation 2.2.3.9 Attachment/Biofilm-Based Systems 2.3 Developments in Algal Cultivation for Fuel By Using Different Production System 2.3.1 Stirred Tank Photobioreactor 2.3.2 Vertical Tubular Photobioreactors 2.3.2.1 Bubble Column 2.3.2.2 Airlift Reactors

26 27 28 28 29 29 30 32 33 34 35 36 38 38 38 38 39 39 39 40 40 40 42 42 42 42 43 43 43 43 43 44 44 45 45 45 46

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2.4

2.5

2.6

2.7 2.8 2.9 2.10

2.3.3 Horizontal Tubular Photobioreactors 2.3.4 Flat Panel Photobioreactor Algal Biofuels – Feedstock of the Future 2.4.1 Biohydrogen 2.4.2 Biobutanol 2.4.3 Jet Fuel 2.4.4 Biogas 2.4.5 Bioethanol Biofuel Pathways 2.5.1 Thermo-Chemical Conversion 2.5.2 Biochemical Conversion 2.5.3 Alcoholic Fermentation 2.5.4 Biophotolysis High Value Co-Products from Algal Biomass 2.6.1 Algae in Human Nutrition 2.6.2 Algae in Animal and Aquaculture Feed 2.6.3 Algae as Fertilizer 2.6.4 Algae as Recombinant Protein 2.6.5 Algae as Polyunsaturated Fatty Acids (PUFAs) Microalgae in Wastewater Treatment Economics of Algae Cultivation Problems and Potential of Alga-Culture Conclusion References

3 Waste Biomass Utilization for Liquid Fuels: Challenges & Solution Sourish Bhattacharya, Surajbhan Sevda, Pooja Bachani, Vamsi Bharadwaj and Sandhya Mishra 3.1 Introduction 3.2 Waste Biomass and its Types 3.3 Major Waste Biomass Conversion Routes 3.4 Metabolic Engineering in Yeast for Accumulation of C5 Sugars along with C6 Sugars 3.5 Genetic Engineering for Improved Xylose Fermentation by Yeasts 3.6 Biofuel from Microalgae through Mixotrophic Approach Utilizing Lignocellulosic Hydrolysate 3.7 Conclusion References

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4 Biofuel Production from Lignocellulosic Feedstock via Thermochemical Routes Long T. Duong, Phuet Prasertcharoensuk and Anh N. Phan 4.1 Introduction 4.2 Fast Pyrolysis 4.2.1 Principles 4.2.2 Reactors 4.2.2.1 Bubbling Fluid Bed 4.2.2.2 Circulating Fluid Bed 4.2.2.3 Rotating Cone 4.2.2.4 Ablative Pyrolysis 4.2.2.5 Screw Reactor 4.2.2.6 Other Reaction Systems 4.2.3 Bio-Oil Composition and Properties 4.2.4 Factors Affecting on Biomass Pyrolysis 4.2.4.1 Feedstock 4.2.4.2 Biomass Pre-Treatment 4.2.4.3 Temperature and Carrier Gas Flow Rate 4.3 Bio-Oil Upgrading 4.3.1 Hydrodeoxygenation 4.3.2 Catalytic Cracking 4.3.3 Fast Hydropyrolysis 4.3.4 Cold Plasma 4.4 Gasification 4.4.1 Types of Gasifier 4.4.1.1 Fixed Bed Gasifier 4.4.1.2 Fluidized Bed Gasifier 4.4.1.3 Entrained Flow Gasifier 4.4.2 Influence of Operating Parameters on Gasification Process 4.4.2.1 Equivalence Ratio 4.4.2.2 Steam to Biomass Ratio 4.4.2.3 Gasifying Agents 4.4.2.4 Gasification Temperature 4.5 Fischer-Tropsch Synthesis 4.5.1 Fischer-Tropsch Reactors 4.5.1.1 Multi-Tubular Fixed Bed 4.5.1.2 Slurry Bubble Column 4.5.1.3 Fluidized Bed

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Contents ix 4.5.2 Catalysts 4.5.3 Influence of Operating Parameters on Fisher-Tropsch Synthesis 4.6 Summary References 5

Exploring the Potential of Carbohydrate Rich Algal Biomass as Feedstock for Bioethanol Production Jaskiran Kaur and Yogalakshmi K.N. 5.1 Introduction 5.2 Microalgae and Macroalgae as Bioethanol Feedstock 5.3 Process Involved for Production of Bioethanol from Algae 5.4 Algal Biomass Cultivation 5.4.1 Open Pond Systems 5.4.2 Closed Photobioreactors (PBR) 5.5 Pretreatment of Algal Biomass 5.5.1 Physical Pretreatment 5.5.2 Chemical Pretreatment 5.5.3 Biological Pretreatment 5.6 Fermentation of Algal Hydrolysate 5.7 Distillation 5.8 Manipulation of Algal Biomass 5.9 Pros and Cons of Bioethanol Production from Algae 5.10 Conclusions References

6 Development of Acid-Base-Enzyme Pretreatment and Hydrolysis of Palm Oil Mill Effluent for Bioethanol Production Nibedita Deb, Md. Zahangir Alam, Maan Fahmi Rashid Al-khatib and Amal Elgharbawy 6.1 Introduction 6.2 Biomass Energy 6.3 Palm Oil Mill Effluent (POME) 6.4 Pome Characterization 6.5 Pretreatment 6.5.1 Physical and Physicochemical Pretreatment 6.5.2 Chemical Pretreatment 6.5.3 Biological Pretreatment 6.6 Hydrolysis 6.6.1 Concentrated Acid Hydrolysis

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Contents 6.6.2 Dilute Acid Hydrolysis 6.6.3 Base Hydrolysis 6.6.4 Enzymatic Hydrolysis 6.6.5 Cellulase Enzymes Hydrolysis Fermentation Process Bioethanol 6.8.1 Lignocellulosic Bioethanol 6.8.2 Bioethanol Production by Fermentation of Sugars 6.8.3 Bioethanol Determined by GC/MS from POME Hydrolysate Conclusion Acknowledgment References

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7 Technological Barriers in Biobutanol Production Arpita Prasad, Shivani Thakur, Swati Sharma, Shivani Saxena and Vijay Kumar Garlapati 7.1 Introduction 7.2 Production Technologies of Biobutanol 7.3 Lignocellulosic Materials for Bio-Butanol Production 7.4 Natural Producers of Biobutanol 7.5 Main Obstacles in the Biobutanol Production 7.5.1 Approaches to Overcome the Obstacles 7.6 Engineered Pathways towards a Better Solventogenic Producer 7.6.1 Engineered Pathways in Bacteria 7.6.2 Engineered Pathways in Yeast 7.7 In-Situ Butanol Recovery Integrated with Batch and Fed-Batch Fermentation 7.8 Future Prospects 7.9 Conclusions References

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6.9 6.10

8 Biobutanol: Research Breakthrough for its Commercial Interest Sandip B. Bankar, Pranhita R. Nimbalkar, Manisha A. Khedkar and Prakash V. Chavan 8.1 Introduction 8.2 Butanol: Next-Generation Liquid Fuel 8.3 Routes of Butanol Production 8.3.1 Chemical Route 8.3.2 Biological Route

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Contents xi 8.4 Microbial ABE Production 8.4.1 Microbial Strains 8.4.2 Biosynthetic Pathways of Clostridia 8.5 Feedstocks Used in ABE Fermentation Process 8.6 Saccharification and Detoxification Processes 8.7 Strain Engineering and Developments in Butanol Production 8.8 Bioreactor Operations 8.9 Butanol Separation Techniques 8.9.1 Extraction 8.9.2 Gas Stripping 8.9.3 Pervaporation 8.9.4 Perstraction 8.9.5 Adsorption 8.9.6 Hybrid Separation Process 8.10 Techno-Economic Assessment 8.11 Current Status and Future Prospective References 9 Potential and Prospects of Biobutanol Production from Agricultural Residues Shuvashish Behera, Koushalya S, Sachin Kumar and Jafar Ali B M 9.1 Introduction 9.2 Agricultural Residues 9.2.1 Husk 9.2.2 Straw 9.2.2.1 Wheat Straw 9.2.2.2 Rice Straw 9.2.2.3 Barley Straw 9.2.3 Bagasse 9.3 ABE Fermentation 9.3.1 Butanolgenic Microorganisms 9.3.2 Fermentation 9.3.3 ABE Pathway 9.3.3.1 Acid Producing Phase 9.3.3.2 Solvent Producing Phase 9.4 Challenges 9.4.1 Strict Anaerobic Nature 9.4.2 Tolerance to Solvent 9.4.3 Sensitivity of Acids 9.4.4 Shifting of pH

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Contents 9.5 Future Prospects and Conclusions Acknowledgments References

10 State of Art Strategies for Biodiesel Production: Bioengineering Approaches Irem Deniz, Bahar Aslanbay and Esra Imamoglu 10.1 Introduction 10.2 Biodiesel and Microalgal Biorefineries 10.2.1 Microalgae 10.2.2 Microalgae and Biodiesel 10.2.3 Selection of Microalgal Strain for Biodiesel Production 10.2.4 Microalgae Cultivation 10.2.5 Harvesting and Lipid Extraction 10.2.6 Conversion of Microalgal Oil to Biodiesel 10.3 Metabolic Engineering Approaches for Biodiesel Production 10.4 Novel Photobioreactor Designs for Biodiesel Production 10.5 Advanced Photobioreactor Configurations and Kinetics 10.6 Conclusions References 11 Bio-Oil Production from Algal Feedstock Naveen Dwivedi and Shubha Dwivedi 11.1 Introduction 11.1.1 Microalgae 11.1.2 Classification of Microalgae 11.1.3 Algae Growth 11.2 Technologies Used for the Production of Bio-Oil from Algal Biomass 11.3 Properties of Bio-Oils 11.4 Uses of Bio-Oils 11.5 Up-Gradation of Bio-Oil to Biodiesel along with Recent Developments 11.5.1 Esterification/Alcoholysis 11.5.2 Solvent Addition 11.5.3 Emulsification 11.5.4 Hydrotreating/Hydro Deoxygenation 11.5.5 Hydro-Cracking 11.5.6 Zeolite Cracking

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Contents xiii 11.6 Conclusion References 12 Effect of Upgrading Techniques on Fuel Properties and Composition of Bio-Oil Krushna Prasad Shadangi and Kaustubha Mohanty 12.1 Introduction 12.2 Bio-Oil and its Properties 12.3 Upgrading of Bio-Oil 12.3.1 Catalytic Pyrolysis 12.3.2 In-Situ versus Ex-Situ Catalytic Pyrolysis Process 12.3.3 Hydrodeoxygenation 12.3.4 Hydrogenation 12.3.5 Steam Reforming 12.3.6 Emulsification 12.3.7 Esterification 12.4 Conclusion References Index

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1 Process Engineering Biofuel Production Opubo Gbanaye Benebo Aningba Technology Development Co NY, USA Current address: Aningba Technology Development Company, Inc., Lewes, DE, USA

Abstract The reasons for the slow growth of the liquid biofuel industry are explained and the approach of process engineering the biofuel production is suggested as apparently the most viable approach for overcoming the limitations of the heretofore failed approaches and invariably for developing the biofuel industry. The significance of the Bench-Scale Model as the foundational mechanism for evolving a translation of the Bench Top experiments results into Engineering Process design is emphasized, and the development of a Base Process Technology derivative of the Bench-Scale Model design is presented for guidance, not only for the biofuel industry but also for the entire biotechnology industry. Further, the various biofuel Market-Participation process technologies so developed are then integrated into an Algal Biomass Biochemical Pathway Biorefinery, as developed for deployment. Keywords: Process engineering, algae cultivation, airlift, process development, biofuel process technology, reaction kinetics

1.1 Biofuel Production Background Liquid biofuel of the form bioalcohol; bioethanol and biobutanol; and biodiesel mostly have been assessed as alternative sources of energy to fossil fuels being derived from biological matter and as such renewable without added net GHG emission into the atmosphere. While there has been a significant focus on the industrial production of the biofuel for large-scale use by industries, the performance has been less than expected or projected Email: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (1–34) © 2019 Scrivener Publishing LLC

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over the years. Many ventures that got started with the object of attaining critical mass and invariably acquiring enduring existence never quite got there, and instead flamed out and went belly-up. These productions started off mostly with cashcrops: sugarcane, etc., as first-generation biofuel production operations that were mostly driven by organic sugars mostly from carbohydrates and aimed to produce mostly ethanol or for sharper distinction, bioethanol. The biofuel production raw materials scope was then expanded with the use of lignin and biomass wastes such as leaves, grasses, harvest stucks wastes, and others still for the production of bioethanol but with a different technology of Cellulosic Biofuel production. Subsequently, Algae as raw material also become part of the scope of resources for the production of biofuel with both carbohydrates and lipids deemed as raw materials for the biofuel production. Then, of course, almost all of these biofuel production operations suffer one form of limitation or another. In most cases these limitations have led to either the idling of the process plant or outright bankruptcy filing. However, during the time-span when these first-generation biofuel production operations were slowly going bankrupt, advances were also being made that fixed some of the sources of the limitations of the different process technologies adopted for the production.

1.1.1 General Limitations Some of the limitations of the various biofuel production operations are common to all and some are specific to the particular production operation. Underrating the general or common limitations and constructing solutions to them advances the production operation collectively. In that respect then, a delineation of each of the common limitations should be constructive. The vast sizes of land surface space needed to support the production operations is one of the common limitations of the biofuel production operations. In every case of the production means, vast amounts of land space is required. The primary reason for that is the need to avail a large surface area for irradiance of light during daylight hours. The landspace often required for growing the raw materials is just too expansive because of the need for horizontal layouts of farms. The dedicated growing of switchgrass for raw material source to support cellulosic biofuel production without any doubt demands large expanses of switchgrass farms from which the grass can be readily harvested for production purposes, and the larger the need for production volume the larger the space needed to grow switchgrass. Growing the switchgrass, of

Process Engineering Biofuel Production 3 course, is essential for the purposes of production leveling as well as production stability. The other forms of raw materials such as biomass wastes are just not stable, being available intermittently. Of course, this situation also applies to the use of cashcrops for the source of glucose raw material for the production of bioalcohol. Similarly the raceways, ponds and other facilities for cultivating algae for the extraction of pilids and carbohydrates also require large expanses of land both directly and indirectly. Heterotrophic algae generally require large expanses of land indirectly because they require glucose and even carbohydrates as substrates and both forms of raw materials have already been noted as requiring large-size farm activities. Phototrophic algae, on the other hand, directly require large expanses of land, as by the very nature of their existence they require irradiance of light during certain periods of the day. Phototrophic algae execute a pattern of dynamics all day during the hours of light. Notably, phototrophic microalgae trace cyclic motions all day – floating towards the surface of the fluid in which it exists and then recycling away, only to repeat the process. Quite as significant though is that for a given orientation of the alga the cyclical motion causes the Cylinder of Exclusion to itself trace another geometric region that the alga cell then requires to support survival. Moreover, a projection of the space vertically up to the surface of the liquid also defines a specific surface area of the liquid surface that the phototrophic alga cell most appropriate to itself for the acquisition of sunlight to drive its carbon fixing reactions. Consequent on the need for vast large expanses for irradiance is also the exposure of surface for water and moisture evaporation. As a result the growing of raw materials for biofuel production is also attended with the need to provide large volumes of water supply to the raw materials growth process. With respect to most algae cultivation vessels, except for the closed vessels, the evaporation of the water is proportional to the size of the surface area, and because this surface area expands as the production volume demand increases, the evaporation also increases, resulting in high cost of operation. Admittedly, of course, closed cultivation vessels allow almost zero evaporation; the demand of water does not necessarily get abated by the vessel being closed. The reason is simple; the closed vessels either cause a mitigation of the irradiance and therefore demand more space to meet the production need or Brag-diffracts the irradiance and therefore require larger cultivation vessels to compensate for the loss of irradiance and the consequent need for more water for the extra vessel volume demand. Both large expanse of land and large volume of water require the use of a correspondingly large quantity of fertilizers or macro-nutrients and

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micro-nutrients, which results in one form or another to phosphorus depletion. The open-ended sourcing of nutrients for the growth and cultivation of the raw materials of biofuel production operations places demand on the sustainability of the operations. The larger the production volume that must be met as per the results of Break-Even analysis, the more the nutrients that must be fed into the production operation: more water volume means more nutrients in order to keep the level concentration up to the design specification; the more the land space needed for farming cashcrops or switchgrass the more nutrients must be used to cover the surface of the land in order to maintain the same level of concentration in the topsoil of the farm. In either case more nutrients are required to meet the need of the production volume. The growing and harvesting and collecting of the switchgrass plants and transporting to the grinder and ultimately feeding into the process entails several downtimes. Agreed that the growth period and the harvesting could be overlapped with periods of existing inventory depletion, yet the operation still requires strict management and then there is always the possibility of natural damage on the switchgrass farm. Similarly the cultivation of algae as is currently operated has several downtimes such as the intermittent harvesting, the transportation of the harvest to centrifuge location and then centrifuging, then cell rupture operation and then biorefinery of the products.

1.1.2 Limitation of Cashcrop Raw Material Cashcrop-based biofuel production operations have unique characteristics that are just as worth noting. Primary amongst the limitations is the competition for food that leads to sharp price increases often to a level beyond what was assessed in the Break-Even Analysis in such times for which such analysis was even performed. A consequence of such sharp price increases has been the erosion of the Financial Viability that may have been assessed for such production operations. The idling of plants only to restart after drops in prices in those cases in which price drop occurred, has the effect of lack of production stability. Financial viability generally requires steady performance of the production over the duration of the lifespan of the company as used in the Depreciation analysis. Irregular performance necessarily introduces disruption of operations and gradually loss of effectiveness and productivity. The competition with food operations over the raw material cashcrops also results in engendering some reservation for support of biofuel

Process Engineering Biofuel Production 5 production which necessarily needed govermental subsides at the initial stages to get off the ground and become a viable operation, but that still leaves the long-term solution to the competition. With population growth and corresponding increasing demand for biofuel, there is never the potential for generating enough cashcrops to meet the needs of both biofuel production and food for the populace. So with the erosion of support for biofuel as an alternative to fossil fuel comes the elimination of the financial subsidies for the biofuel industry and hence the erosion of the Financial Viability of the biofuel production. The Financial Viability of biofuel production operations of this category prevailing from this sharp rise in prices results invariably in accumulation of financial loss in operations leading to idling, shutdowns of operations and outright bankruptcies of the companies.

1.1.3 Limitations of Algae Raw Materials Remediation Biofuel production based on the use of algal lipids and carbohydrates also has its unique characteristics that are just as worth noting. The unique limitations, however, are applicable only to the phototrophic algae which have different needs from the heterotrophic algae, the needs of which have been noted under the General Limitations, Section 1.1. One primary limitation of the cultivation of phototrophic algae for biofuel production raw material has been the low concentration of carbon dioxide in the atmosphere from which the the gas is absorbed into the suspension for the utilization of the algae. A competing limitation is that the cultivation vessels tend to suffer extensive scatter [1] of the light irra- diance on the surface exposed to the open. The Light scatter dynamics essentially follows the pattern of Brag Diffraction consequent on the Crystalline Microbial Arrays formation resulting from cell-interactive dynamics from which the cells form array structures that are crystalline equivalent in form [1] and because these arrays are consistent with Bravais lattice, the cells effectively support this lattice form, Microbial Lattice Arrays, MLAs, in the context of the CMAs. As a consequence of the array formation, the light incidence on the surface are primarily Bragg diffracted out of the cultivation vessel with very little quantity of the light actually being transmitted into the reaction medium [1].

1.1.4 Limitations Remediation Remediation of the various limitations in the production of liquid biofuel undoubtedly will advance the development of the industry. So then there is

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the need to address the limitations that may be retarding the growth of the industry. Several approaches have been adopted that in combination define a growth thrust for biofuel production operations. Any remediation of the limitations elicited for each of the production operation must necessarily have at the core the capacity to embody most of the remediation. The most obvious solution to any of the limitations is the transformation of the biofuel production operation into continuous operation, because it is generally known that continuous operations tend to eliminate all downtimes that characterize Batch Operations. So the elimination of all batch steps is a significant solution. This solution results in Continuous Flow Production: in this regard the transformation designs operations of the form of chemical process but uses a bioreactor in place of the characteristic chemical reactor and other specialized process-specific equipment; and so entails the use of process equipment configuration bioreactor to support the continuous production. Effectively all raw material production mechanisms that have the dynamics of farming therefore are disqualified as an option of remediation. The remediation of the need for large landspace is just as critical towards enabling the advancing of the biofuel production. One significant approach would be to use bioreactors of stackable design, such that the bioreactors collectively occupy a small landspace while growing upwards. The need for the stackable bioreactor design is consequent on the large number of bioreactors needed to support continuous production operations. An operation requiring the effluent of a bioreactor per hour requires 24 bioreactors per day, and because each bioreactor takes about 8 days to be ready for effluent discharge, places the number of bioreactors needed for a continuous process at 192 minimum; and any production volume increase requirement will only increase the bioreactor count from 192 bioreactors. So then the use of a bioreactors of stackable design is critical to the remediation of landspace demand. The extensive use of nutrients, both macro-nutrients and micronutrients, as a limitation is mostly in the use of fossil fuels products as the base chemical in the production process. As such the object of using biofuel as an alternative fuel is not realizable unless the nutrients productions do not involve the use of chemicals based on fossil fuel derivatives. This dependency has generally been referred to as the NPK: Nitrogen (N), Phosphorus (P) and Potassium (K); dependency. As such, a more self-contained biofuel production operation would be more consistent with the goals of alternative fuel source to fossil fuels. In effect the remediation of that limitation can only be obtained provided the nutrients used in the production operations are obtained from the raw materials of the biofuel production or the

Process Engineering Biofuel Production 7 by-products of the production. Addressing direct response to the argument of NPK dependency, in effect a limitation to the object of the alternative fuel program, a necessary remediation is the recycling of the nutrients or at least the raw materials by which the nutrients were produced, and to then produce the nutrients from the same recycled raw materials. In that regard there are, in fact, several entry-points—meaning the first step of the Process Chemistry or Process Biochemistry or even Process Microbiology—by which to develop an Algal Phosphorus Recycling Process technology, and an extraction process research [2] proffers one entry-point into the phosphorus recycling technology development, and being continuous further supports the adoption of the Continuous Operations supports Nutrients Recycling Process Designs. Amongst the raw materials delineated so far as being used for the biofuel production, one needs to be chosen as the preferred for the objects of biofuel production. A criteria set as may be developed from the Remediation methods, has the need for complete recovery and recycle of the nutrients and is overriding because of the need to virtually eliminate any loss of phosphorus so as not to have to tap into the reserve needed to support life of the growing population. In this regard then, the use of any form of landbased farm is excluded, which would be cashcrops and switchgrass, and either one would require mixing the nutrients with the soil of the farms. The only raw material then left to use becomes algae. Though chosen first and foremost on the basis of the preservation of human life, algae also offer the added advantage of serving as a raw material for all the needs of being a continuous operations. First, algae as a source of Starch/Sugar stands to provide raw materials for the bioalcohol biofuel sector of the biofuel industry; second, as a source also of lipids algae also offers the raw materials for the biodiesel biofuel sector of the biofuel industry, effectively serving the entire liquid biofuel industry. Then, of course, the bioalcohol produced with algae carbohydrates is also a default raw material for the production of diesel, and in effect propagating its raw material provisioning through the entire liquid biofuel industry. On the other hand, the bioalcohol fermentation process requires nutrients to be fed to the bacteria and such nutrients could be produced with the waste biomass of algae after the extraction of the carbohydrate and the lipids which are essentially carbon dioxide and water in origin, hence preserving the nutrients consumed during the cultivation process for recycle. Indeed, this gives one continuous backward “Supply Chain” necessary to be developed through the Algae Cultivation Industry, effectively integrating the industry as to yield only the products that are based on carbon dioxide and water: The Bioalcohol Biofuel Industry needs nutrients for the bacteria which is best

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acquired from the Biomass Biofertilizer Industry; The Algal Lipids/Oils Biodiesel Industry needs ethanol as raw material which is best got from the Algal Bioalcohol Industry. Further, though not often undertaken in a traditional Process Technology deployment setting, Algal cultivation is very well-suited to the Process Technology format, and hence is a good raw material for continuous production operations, and proffers the Growth Thrust for the liquid biofuel industry. Incidentally, the most advances derivative of both related and unrelated efforts have been made in the production of algal biofuel production. The landspace use maximization requires stackable bioreactor; Biological Process Technology allows stacking of the reactors, albeit well-designed reactors. Also an incidental benefit of the Biological Process is that harvesting is no longer necessary as the reactor effluent must necesarily flow into the next process equipment whatever that may be and on to the next equipment until the final product flows into storage tanks, effectively eliminating the downtimes and handling costs.

1.2 Process Engineering Liquid Biofuel Production In the context of the remediations considered for limitations of the various Biofuel production operations, algae as raw material seems to offer the most promise, and accordingly is focused on here as the raw materials on which the advances are been cobbled together to elicit a growth thrust for the production of liquid biofuels. In this regard suggest designs of the form of chemical process but using bioreactor and other specialized process-specific equipment, and so the use of process equipment configuration bioreactor to support continuous production. By and large the process engineering must be based on the experimental and established approaches by which the cultivation of algae has so far been performed. So then the object of the process engineering is to replace the hodgepodge of equipment and machines with off-the-shelf process equipment with standardized designs to support reproducibility, repeatability, lower cost from economy of scale, and rapid replacement.

1.2.1 Algae Cultivation Assessment Effectively, every algae cultivation system or operation necessarily provides the needs for supporting the metabolism with respect to the classification of the alga species enabling the algae existence in as near its natural form

Process Engineering Biofuel Production 9 of existence as possible. Every algae cultivation operation effectively consists of chosen feed type and environment that support the life of the alga species. Several forms of vessels and environments have been contrived and applied at different times and in different settings for the cultivation of algae; the efficacy of the design of each of the vessels is best elicited with assessment of the performance of the vessel in context of use; and so some of the commonly used vessels have been evaluated for intrinsic limitations demanding remediation: Paddle-Wheel Circulation Cultivation Raceway A raceway is a rectangular channel that is constructed into a loop of the form of a raceway. At an appropriate point within the loop, a wheel is emplaced which is connected to a drive motor that drives the wheel paddles that should effectively churn the suspension when poured into it. Usually this channel is about 5ft wide, 0.5 f deep and as long as the designer use-specifies. Though used quite often and mostly as a form of algae farming facility, the operation and control of the vessels are limited by the difficulty in matching the production rate and the harvesting rate. Given any volume of the algal suspension as may be calculated, then the quantity of suspension withdrawn at each cycle of harvest is equal to the computed volume. The implementation of the harvesting process that effectively balances the harvest volume and the growth rate is not always trivial. It has been almost impossible to accomplish this balance dynamically. Airlift Algae Suspension Circulation Cultivation Tubes Banks This vessel is simply a bank of interconnected glass tubes that are organized such that liquid flows through the full length beginning from one open end point and start point. By design one open end of the tube is affixed to a tank at a bottom outlet, while the other open end is connected to the lateral side of a vertical tube column to the base of which is affixed an air bubbling outlet through which air is pumped into the column. The tubes are irradiated with light either by exposure to the sun being placed outdoors or by artificial light produced with LED and Light bulbs and other forms of light generators. The light beam is simply incident on the

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Liquid Biofuel Production glass tubes without consideration for the angle of incidence relative to the direction of the fluid flow. At this end of tube connection, the liquid is lifted with gas bubbles into the holding tank, and the liquid is aerated during the uplift into the tank. Inside the tank the fluid then flows out through the bottom outlet into the interconnected tube. The algae in the suspension being exposed to light during the travel through the tubes utilize the dissolved carbon dioxide and nutrients in the suspension and both grow bigger in size as well as replicate. A portion of the suspension is then harvested through the holding tank. However, there are several obvious performance shortcomings attendant to this vessel. The shortcomings occur from just about every aspect of the design though by default. Given the volume of the algal suspension then the quantity of suspension withdrawn at each cycle of harvest is equal to the computed volume. The implementation of the harvesting process that effectively balances the harvest volume and the growth rate is not always trivial. Bubble Column Cultivation Tanks This vessel is in essence a glass tank with air sparger affixed inside it and an impeller also affixed. The sparger is connected through the base of the tank to an air bubbling inlet through which air is pumped into the sparger. The tank walls are irradiated with light either by exposure to the sun from being placed outdoors or by artificial light produced with LED, light bulbs or other forms of light generators. The light beam is simply incident on the glass tubes without consideration for the angle of incidence relative to the Global Coordinate System. The significant advantage with this reactor over the earlier ones is that the alga cells circulate in a form similar to their natural motion, though admittedly, the circulation is not uniform as some of the cells enter their circulation upward motion before actually getting to the bottom of the reactor. Bubble Airlift Suspension Recirculating Tanks This vessel is in essence a glass tank with an inner shorter cylinder resulting in a nested concentric cylinders; and air sparger is affixed inside inner cylinder, and an impeller also affixed. The Sparger is connected through the base of the tank to an air bubbling inlet through which air is pumped

Process Engineering Biofuel Production 11 into the sparger. The outer-tank outer surface walls are irradiated with light either by exposure to the sun from being placed outdoors or by artificial light produced with LED, light bulbs or other forms of light generators. The light beam is simply incident on the glass tubes without consideration for the angle of incidence relative to the Global Coordinate System. The significant advantage with this reactor over the earlier ones is that the alga cells circulate in a form similar to their natural motion, and even more importantly the circulation is uniform as every alga cell enters its circulation upward motion only after actually getting to the bottom of the reactor. The above listing, though not exhaustive in the least, of the various types of vessels, refers to the most commonly used forms. Each one, however, has its limitations, as observed already, and it is the remediation of these limitations that significantly advanced the development of technologies for the production of algal biomass.

1.2.2 Algal Cultivation Inefficiencies Remediation Having noted the sources of inefficiencies in the cultivation of algae, one of the considerations of the process engineering of algae for biofuel production is the process design rationale that necessarily designs in circumvention of these sources of inefficiencies. A significant inefficiency is the Bragg diffracting of incident light on the surface of the cultivations vessels as observed. In effect, the Light scatter dynamics of Bragg diffracting of incidence light suggests the use of special cultivation bioreactor designs that minimize light scattering at least completely avoids the Bragg Scattering characteristics of these vessels. In effect, a form of Cultivation Bioreactor Engineering is required for which some specifications may have to be set to the end of meeting normal ordinary standard. Further to ensure that grazers not intended to be in the algal suspension do not get into the medium and ravage the cultivation the bioreactor must be closed, and impenetrable to extraneous microbes to the extent possible. A necessary approach then is the design of the bioreactor as a composite regular design and as a radiosterilizer such that whenever continual monitoring of the bioreactor for invasion by grazers reveals such contamination, the radiosterilization can be implemented dynamically.

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The recycling of the waste algal biomass is crucial to the sustenance of life through the conservation of phosphorus that every engineering effort must be made to support the aspect of the process engineering. Accordingly it is expected that the process of decomposition cum transformation of biotic wastes into natural biofertilizers must be carefully studied and replicated in the laboratory setting. The expectation is that this bioprocess chemistry will result in the design of a biotechnology subprocess as the stages of the conversion of the biotic waste into natural Biofertilizers is duplicated with a form of bioreactors in itself. The extraction of the nutrients from the waste biomass presents an entirely different consideration. Because there is no reference from which to determine a bioprocess chemistry, the engineering of the process for the production of nutrients will have to be creatively developed. Even then the likely approach would involve the development of a bioprocess chemistry and the development of the subprocess that supports that chemistry, although such would entail one biochemical reactor or more, as the biomass must necessarily be specially reacted within a sequence of enzymatic contexts en route to producing the nutrients. The restriction of the utilization of water during the cultivation of the algae is enabled with the use of process equipment configuration design of the bioreactors. In effect the bioreactor is designed according to the principles of chemical reactor engineering. A vessel that is closed allows for monitoring of all essential variables, and supports the suspended state of the microbes through the duration of the batch-reaction time. As per earlier observations for low-cost operations, the reactor must utilize sunlight and distilled water which is used within the process such that prevents contamination with grazer and extraneous microbes; the process must be closed and circulating to conserve the distilled water used for the production.

1.2.3 Technology Development Admitting algae technologies as process technology requires a fundamental rethink about the development of the algae biofuel process technology development approach such that there is consistency with the process technology development norm. The fundamental rethink is critical because heretofore the algae biofuel industry has not followed the standard technology development process; rather the industry has adopted the process commercialization path often adopted in the biological industry. That the approach did not align with the process industry approach is readily surmised from an analysis of recent failed algal bio-fuel ventures.

Process Engineering Biofuel Production 13 Further, the process technologies as attempted commercialization by not following the process development, the consequential operations also could not support the Process Industry norm of one 2-week maintenancedowntime a year. As a matter of standard operation of the process industry, the process technology operations must prevail over a period of a whole year with only one 2-week downtime. Clearly, cultivating algae for the biofuel industry must adopt a growth reactor of such equipment configuration design that meets the design specifications: by the requirement of permissible downtime of 2 weeks a year, the reactor fabrication material must not be wettable, as to prevent the natural stickiness of the algae cells to the reactor internal surface that necessitates frequent cleaning and frequent downtime.

1.2.4 Lessons from the Algae Biofuel Industry Collapse In almost all cases, the sale of assets of failed biofuel ventures show assetsbase of less than $100 million and no Process Pilot Plant, though as fluidbased production operation and hence a process venture meant the deployment of a biological process plant. Yet none of the ventures had a process pilot plant, which shows that the base process technology was never developed. Indeed, the Semi-Commercial Pilot Plant, which any of these ventures would have constructed to start off the Execution Phase [3] of the Startup Stage, would have cost about $100 million. The nonexistence of a Process Pilot Plant shows that the technology development steps were not followed, suggesting these venture-developers never adopted normal venture planning for developing process ventures, and the stages of technology development, short-circuited the technology development process and so never developed a venture-specific base technology and as such could never plan for break-even production volume to support the operations. The motivation for failing to follow the development path, however, is endemic in the biological industry up till now. Generally, for the biological industry, including the medical industry, experiments are performed in the laboratory within some general sense of expectation, and should a success obtain then the procedure is simply to scale up the laboratory set up and produce more. That process often amounted to commercialization in that industry. However, that approach often is simply empirical and there is usually not enough understanding of the science that is brought to bear on the engineering of the technology that is necessarily consequential on the experimental observation. That biological industry approach of commercializing production, however, is not consistent with the process technology industry. While in fact

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the approach is successful in an industry where there will always be a consumer to buy the product at any cost that compensates the inefficiencies of the operations development, that is not the case in the process industry, which being very competitive requires optimization of the engineering of the technology such that wastes and inefficiencies are eliminated before commercialization of the process.

1.2.5 Process Development Norms So without exception, an Algae Process technology development endeavour must necessarily begin at the Bench-Scale Model Stage and develop the essential base technology. Normally, a venture acquires the experimental data of innovations of biological scientists, and develops at the Bench-scale a base technology, for the design and construction of a Pilot Plant, that through optimization becomes the small scale of the would-be Commercial Production Plant. The norm for process technology development has been primarily of three stages: the Bench Top Scale development, the Bench-Scale Model Development, Pilot Plant Engineering stage. Only after the Pilot Plant Stage has been successfully developed does a commercialization begin. In general each of these stages can not be circumvented and should not be as each stage forces up a certain dynamic of the technology that is not obvious otherwise. Although these three stages are the primarily recognized ones in the process industry, these steps, however, actually mask the detailed steps that have been discovered over the years and have been catalogued as the standard [4] by the National Aeronautical and Space Agency (NASA). These development-wide applicable steps consist of ten levels [5] but have generally been repackaged into the three process industry accepted steps [3], and should in actuality be followed for consistency across the board. All development begins with the Bench-Top Research Experimental Design for the production operations. In the case of algae cultivation which has been simply enlarged without guidance by Engineering Methods of Scale-Up, ponds, raceways, bubble columns, etc., followed by harvesting using centrifuge and flocculation, cell rupture using venturi, etc., and a cellular components separation constitute the experimental design. So then for all intents and purposes this public domain information then defines the experimental design on the process engineering must take off and evolve into a commercial plant design.

Process Engineering Biofuel Production 15 The first stage of the process development is the Bench-Scale Model stage; at this stage of the process technology the goal is the translation of the laboratory experiments into a semblance of a continuous process technology. The cultivation of algae with intermittent harvesting by one group producing algae biomass followed by another group trying to produce something else has large downtimes and expensive handling costs in between operations and hence takes away from any possibility of financial viability. The design objective is to replace the glassware and vessels used at the laboratory experiments with bench-scale process equipment consistent with standard off-the-shelf process equipment. In effect, each task of the laboratory experiments is replaced with some form of process equipment, after the completion of the choices of the process equipment in substitute of the laboratory task. The choice of the process equipment, of course, is guided by the prevailing understanding of the science defining the specific task of the laboratory experiment. After the scientific characterization of the process of the Bench-Scale Model, together with the implementation of such equipment changes as may be appropriate, then the engineering activities move into the construction, operation and optimization of the Pilot Process Plant. The technology development now focuses on the Pilot Plant Engineering, which sort of builds up a larger scale of the operational Bench-Scale Model, and into this process Plant are integrated various devices and instruments which incrementally upgrades the technology to the level of commercial process plant. Operations conditions are changed and equipments redesigned or substituted otherwise as necessary until optimal conditions are determined, at which time the resulting Pilot Process Plant becomes a mini-Commercial Large Scale Process Plant. These three stages then are the evolution path through which the process engineering must be developed in order that technology viability could be properly evaluated and established, leading to commercial viability as well as financial viability [3].

1.2.6 Research Team The task of developing Biological Process Technology is often approached as a one-man project, but that is in fact not realistic. Often, some people have given the impression that an entrepreneur being given a helping hand with some funding or incubation space is all that is required to get him/her off the ground, but nothing can be more inaccurate. Many years experience has taught me differently.

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The fact is any process development task must be approached as a team effort, and the entrepreneur must know the specific class of the development team he/she fits in, and then form the team by attracting competent staff to join the venture. Indeed, every development team must consist of three classes of developers: the Technology Conceptualists, the Validation Experimentalists, and the Technology Optimizers. A team that is deficient in any one of these classes is bound to fail. The fact is that the conceptualists are strictly scientific thinkers without regard to the construction of the equipment that makes manifest the conceptualizations. Usually the Validation Experimentalists who understand the scientific underpinning of the concept design work with the conceptualists to build the equipment from the concept – often designing around the limitations of the direct implementation of the concepts; and after the validation the equipment is then handed over to the optimization developers to undertake the design optimization. Often these people are very different in their thinking orientation towards technologies: mostly the optimizers cannot conceptualize, and cannot validate concepts; the Validation Experimentalists may with some level of push perform optimization but not very often and not willingly; the Conceptualists often are just that, conceptualists, though occasionally there might be one who can also do experimental validation but not too often. Each reactor design is also very specific to the objects of the design. The conceptualists will evolve the design to the end object, the experimentalists will validate each one, the optimizer will optimize, the Biochemist develops the Pathways, the Kineticist develops the Reaction Rate Equations, the Optical Physicist develops the optics, and the EM-Physicist undertakes the quantum electrodynamics analysis of the interaction between the EM-waves and the electron of the Photosystems.

1.2.7 Alga Cultivation General Issues As is now widely known, the one raw material for Biodiesel Processes that guarantees profitability and production stability is algal oil, ordinarily extracted from a category of algae that must be deliberately grown under optimal operating conditions. So then any Biodiesel Process Technology must necessarily also resolve on the means of making the algae available for the purposes of extracting the oil. Interestingly, the algae from which the oil is extracted could become available to the Biodiesel Process by either being grown with a technology that is integrated as a subprocess of the entire Biodiesel Process Technology, or being independently grown by vendors using stand-alone process

Process Engineering Biofuel Production 17 technology separate from the Biodiesel Process. In any case, the key factors of consideration in any such operation must derive from the mind-set of operating with only unavoidable cost. In keeping with that mind-set, the considerations must then center on the cost of operation of a simple alga production operation as reverse engineered from posts on the public domain. The factors based on such abstract reverse engineering include among others, break-even point dependency on design rationale, cost of electricity for LED and other forms of artificial lights, the cost of fertilizers, and raw material provisioning. The cost reduction posture towards these cost factors, though entrepreneur-specific, can be achieved by some obvious means. With respect to optical energy source, sunlight instead of LED lights should be utilized. This posture certainly eliminates the initial and repeat purchases of LED lights, and the cost of electricity consumed by such. Besides the white light maintains the optical energy utilization capacity of the algae at 100%. Further, the use of the sunlight causes the algae to operate through the cycles: Periods of Light hours and periods of Dark hours. Further, given the current sustained price pressure on fuels, competitive Market Participation is predicated on comparable pricing of the Biodiesel. Comparative competitiveness analysis—while neglecting the scale that supports break-even as well as profitability—suggests a specific set of integrated technologies is essential for price competitiveness: analysis of profitability has shown that algal oil-based Biodiesel is price competitive only if produced with a core Biodiesel Process that is further augmented with three essential technologies: Biofertilizers Process Technology, Solar Power Technology and Biofuel Combustion Power Technology. So then, whether the technology by which the algae is produced is a subprocess technology integrated into the overall Biodiesel Process or is a stand-alone process technology of an algae biomass vendor, any process for Biodiesel production must necessarily embody this minimal set of technologies. While for the integrated subprocess conformance can be forced to ensure low production cost, for the vendor technology, the issue is a bit intrusive. Indeed, getting vendors to use features-conformant technology imposes a level of control that is significant in being burdensome. So then algae growth cooperatives guarantee the only viable option for vendor supply chain in permitting adoption of non-intrusive conformant technology.

1.2.8 Biofuel Process Technology With completion of the delineations and analysis of the factors that are of significance in the process engineering of the liquid biofuel based on algal

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biologics, the synthesis of the results of the analysis is then undertaken in engineering of the consequent biofuel process design. As it is, there are two forms of liquid biofuel: Bioalcohol biofuel and Biodiesel biofuel, of which the latter is of more complexity and embodies the former. Therefore it follows that the first tier liquid biofuel process design must be the bioalcohol biofuel, and correspondingly the Biofuel Process Technologies are the Bioalcohol Process Technology. Invariably the second-tier liquid biofuel process design is the Biodiesel Process Technology. At the basic design the bioalcohol process design utilizes the carbohydrates extracted from the cellular materials of the effluents of the cultivation bioreactors and converts it into alcohol, preferably biobutanol, although the most commonly produced is the bioethanol. The biodiesel process design at the basic integration has the algal oil extracted from the algae effluents of the cultivation bioreactors and combines then with the bioalcohol of the first-tier biofuel production process and produces biodiesel. These design considerations invariably suggest that the technology development of the process engineering must consist of at least three technology development tasks: Alga Cultivation Process Technology, Bioalcohol Biofuel Bioreactor Technology and Biodiesel Biofuel Reactor Technology; the approaches to which are presented. However, because significant advances and developments have been recorded in the public domain for the Biodiesel Biofuel Reactor Technology, the presentation skips the technology.

1.3 Algal Cultivation Process Technology The process technology for the algae cultivation then is simply the conversion of the known production operations: Cultivation: growing algae; Harvesting: collecting the algae from the growth-fluid; Cell Rupture: the disemboweling the cells; and BioFeedstock Extraction: separating out the lipids or carbohydrates, or both from the cellular biologics suspension. The initial task then is the partitioning of the production tasks into process units. Rationally, the cultivation step can be made a process unit, and as is the BioFeedstock Extraction. Further, while as a continuous operation there is no more need for Harvesting as such has become automatic being the effluent of the Cultivation Process unit. So a Harvestor – now just a concentrator. The Cell-Rupture operation is the only one left to be merged with either of the process units, and which careful consideration

Process Engineering Biofuel Production 19 suggests being merged into the extraction unit to collectively become Bioproduct Extraction Process Unit. Obviously, the basic algae lipids and starch production tasks are engineered into the Cultivation Process Unit and BioFeedstock Extraction Process Unit.

1.3.1 Cellular Reaction Kinetics Analysis A very simple example is this: carbon dioxide diffuses into the environment of the algae and by osmotic pressure diffuses through the cytoplasm membrane and into the cell and onwards into the chloroplast. In the chloroplast it is processed into a glucose-type metabolite; this then diffuses out of the chloroplast back into the cytoplasm where it participates in both Catabolic and Anabolic Reactions with the latter producing intermediaries that serve reactants in the Biosynthesis reactions that cause the massgrowth and cell-divisions. However, representing this observation mathematically in terms of the cellular reaction rate equations for use in the evaluation of bioreactors and in turn in a process technology has been hindered by several challenges, all of which must be addressed only through extensive research and development: • The reaction mechanism by which the optical energy actually gets transferred into the ADP and to form the ATP that drives the Calvins Cycle reactions; • The sequences of reactions that transform the molecular CO2 into glucose-type metabolite; • The possible Denaturation-reactions in an algae cell and which are driven by acidity of the environment, etc.; • The chemical potential in the chloroplast at which the diffusion of the metabolite out of the chloroplast into cytoplasm occurs; • The number of channels (as transporters or symporters) through which the metabolites diffuse out of the chloroplast; • The metabolites captured in the cytoplasm, with vesicles or just plain substances in the cytoplasm; • The transport-mechanics by which the enzymes travel from their biologics of production to the surface of the chloroplast to begin the reacting with the metabolite. There are just so many things that the engineer still needs to know, that we do not know as yet. Research and development therefore is essential

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and must continue to be supported. Yet some significant progress could still be made in determining the operating conditions that enable optimal operations: The use of Composited Reactions Mass Action Kinetics Model [6] rate expression relation can be derived somewhat for current application even if imprecise. Then, of course, because biofuels are either in the form of bioalcohol or biodiesel, three reactions orders have been sequenced to accomplish this goal and correspondingly, to that end, three reactors have been developed but are still under continual improvement design: Batch Photoradiation Stirred Bioreactors for algae cultivation; Immobilized Microbes Carriers Packed Bed Bioreactor for fermentation; Algal Oil Transesterification Reactor for biodiesel formation.

1.3.2 Cultivation Bench-Scale Model Design By the process units definitions then the Bench-Scale Model must now consist of Cultivation Bioreactor, Concentrator, Cell Rupturer and BioFeedStock Extraction. These constitute the initial set of process equipments to be assembled to form the process technology. Of course, one of the advantages of development of a process technology is the option to use off-the-shelf process equipment which standardizes the equipment as well as ensure Commercial Viability [3]. For one thing, replacement of equipment no longer becomes a challenge. Further, at evaluation of BreakEven Analysis, equipment viable sizes become readily available and get procured. So a critical growth thrust of the algal industry is enabled by the systematization of production operations into process technology for continuous flow operations; these process equipment must be selected from as standardized off-the-shelf equipment as possible. Given the context of the development of the technology with offthe-shelf process equipment, the elicitation of the advances that have been made with respect to liquid biofuel production then should integrate designs that are already in the public domain as a means for entrepreneurs to begin the venture as these collectively define referenceable advances made in the development of process engineering of biofuel production. So then each of the process equipment proffered as equivalent or substitute for the task of the biofuel technology evaluation by way of the laboratory experiments is reviewed in terms of available public domain design. These advances, however, have also suggested a certain set of specifications that needs to be defined as guides.

Process Engineering Biofuel Production 21

1.3.3 Cultivation Bioreactor Several designs of algae cultivation bioreactors are available and known and can be licensed or procured for use. However, such a bioreactor must necessarily have the process equipment configuration design such that all parameters can be readily evaluated from measurements of variable.

1.3.4 Concentrator Harvesting of Cells Harvesting the algae cells has to be done with Centrifuging, with the final end product being a mucilaginous mass of algae cells not very suitable for subsequent operations except for drying. For the purposes of process engineering, however, there is the need to have the algae effluent stream of the cultivation bioreactors be simply concentrated, which effectively suggests that the a Concentrator instead of a Centrifuge would be a better process equipment. However, Concentrators that are in the public domain in the process industry could be thought of as Filtration Equipment. Although admittedly a purely continuous process equipment offers the prospects of highest productivity, and is advised as the recommended path to adopt, many algae biomass producers would still use centrifuging to collect the biomass for product. In those cases, where the Centrifuge can be set to operate over a set period of time, as determined through a matrix quadratic relation [1], such an option enables the continued deployment of Centrifuges for the concentration as well. Besides, although, the implementation of the operating time span for the harvesting process may entail several steps, depending on the degr ee of concentration that is preferred, yet there just may be some harvesting situations such that small quantity of fluid is left with the cells instead of an end-product of gooey mass, and only the use of the Centrifuge would achieve that state. During such times it is helpful to calculate the time-span for which the centrifuge should be run. The method developed for making such estimates of time is based on the inter-particle or inter-cell spacing computation matrix function quadratic equation that allows the time-span to be determined [1].

1.3.5 Cell Rupture Technology One of the devices for the rupturing of algae cells is said to be the Venturi. Generally venturi [7] is a flow pressure reducer, with a pressure significantly becoming lower than before entry into the venturi. The use of the venturi as Cell Rupture Device of the algae biomass leverages the pressure differential

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between the inlet and outlet sections of the venturi technology resulting correspondingly in higher internal pressure at the oulet and causing bursting of the cells. Although somewhat functional, the venturi extraction approach, however, is not very effective, because while in fact cells are usually ruptured as expected, the cells are not always disemboweled and as a result some lipids are left inside some of the cells. So, depending on the venturi design, variable quantities of the lipid may be lost as part of the post-extraction biomass waste. Yet quite significantly with respect to the performance, the extraction process is not implosive as would be derivative of compressive pressure differential, as would be the case for algae cells deep inside the Earth which would also have been subjected to intense uniform heating. So an effective design guideline of Cell Rupture devices design can be justifiably based on the dynamics as the natural processes regarding the formation of the fossil fuel: increase the internal temperature and external pressure simultaneously as to implode the cells and then cause a diffusive accumulation of the oil that invariably leads to the formation of the oil-wells. Effectively, the one design specification, essentially, is accomplish the same end result as the natural process. Most significant of all is that the process is conceptually reenacting a natural process and means that both the developing of the high temperature and the high pressure must occur virtually simultaneously. This dynamic is the specification by which all devices or equipment used for the extraction of lipids from algae must be designed. The one design specification is enabled inducing Thermodynamic high temperature and pressure uniformly on the algae cells, such that the equipment must cause an implosion instead of explosion of the cell, as the former will keep the contents localized while the latter will scatter them. Effectively, the equipment must as first step do in minutes the task of accumulation of the oil that naturally took millions of years as with the fossil fuels. The process of concentrating the microbes, however, is accomplished with the Concentrator such that the algae cells are yet in a fluid enough state as to be able to effectively apply the high pressure and temperature. So then the specification-compliant Algal Oil Extraction Process must create an environment that allows the simultaneous application of both high temperature and pressure on the algae cells. Moreover, the environment must provide a sort container capacity such that the lipids can form a pool through diffusive accumulation.

1.3.6

BioFeedstock Separation Process

The Cell Rupture effluent rationally would flow into the Feedstock Separation Process, which by performance must separate both the starch and the lipids from the residual biomass, and then the separation

Process Engineering Biofuel Production 23 of the lipids from the starch for use respectively in biodiesel reactor and fermentation bioreactor. By this evaluation the Separation Process Technology must at a minimum consist of two types of process equipment. A concept in the public domain that could be useful is the Algae Cells flocculation, which entails neutralizing the electrostatic charges on the cell resulting in the sedimentation of the cells. The sedimentation process of course could be further accelerated with a Centrifuge. Of course, the feed for the Flocculation Process is the effluent of a Decantation Process that extracts the biomass feedstock of starch and lipids.

1.3.7

Bench-Scale Cultivation Process Technology

Although the basic tasks constituting the laboratory experiments have been effectively defined with substitute process equipment, such design may not as yet be viable for several reasons including three most important ones: Financial viability of the design may have to be evaluated and the design reengineered for the Financial viability, Sustainability engineering of the design may also be necessary and must be performed as well to ensure Sustainability, and finally Optimization Engineering of the basic integrated equipment may yet be necessary. The Financial viability ensures that the process is vested with production stability; Sustainability engineering ensures that the overall operation does not have to potential for negative impaction of life; Optimization ensures maximal productivity of operation. Of course, such optimization may consider extraneous but relevant factors prior to integrating a functional form of the process technology, even abstract in construction.

1.3.8

Process Technology Financial Viability Design

Several factors have been considered with respect to the Financial Viability design of the Base Process Technology, and the object of these factors have been to both reduce the cultivation cost as well as lower the investment cost. The optimal low-cost algae production operation for algae cultivation Process is determined to have two subprocesses: Biofertilizer Process, Solar Power Technology. The Biofertilizer Process is by design a mechanism for the recycling of the biofertilizers and nutrients fed the current algae effluent of the bioreactor during the cultivation period, so the materials could be fed to the algae currently in the cultivation bioreactors. Besides, the direct production of the biofertilizer for the cultivation is likely to result in the use of

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contaminant-free fertilizers, as fertilizers seem to have contaminants of trace amounts of heavy metals [8]. The Solar Power Technology is anticipated to have the energy demand met by the extraction of Solar Optical and Solar Thermal energies. Because the processing of the algae cultivation entails energy use which may be viewed as a sort of energy input into the cultivation system, the input of available energy into the system enhances the energy density analysis of the output products.

1.3.9

Process Technology Sustainability Engineering

The process Engineering of Sustainability of the Base Technology arises from and is driven most by concerns expressed regarding algae biomass production operations. The concerns as variously expressed address the sourcing of the carbon dioxide, water, and nutrients and fertilizers. The source of the carbon dioxide as well as the purity is significant even if the biomass produced from the algae are used in the production of biofuel, and that the biofuel would be combusted in an engine and vented into that atmosphere for human breathing poses a level of danger to humanity. Sustainability then calls for the use of clean carbon dioxide stream. Effectively, a Multi-Stage Air Separation Purification Technology for the cleaning process is suggested as enabling of Sustainability, and is of critical consideration. The sourcing of water as raw material is crucial only because of the conditions of rolling drought that seem prevalent nowadays and such that affect different regions unpre- dictably. Further, water as raw material is expected to be provisioned subject to certain specifications: Purity and free-of-grazers that eat algae and free-of-bacteria that will make the performance unpredictable; that is necessary in order to ensure performance reproducibility and repeatability. So even with the availability of water, the specifications on the quality of the water for growing algae must still be addressed. Obviously therefore, algae growth process would have to use distilled water, and also use radiolytic reactors irradiating the water at frequencies that destroy contaminants coupled with filtration process. The utilization of phosphorus nutrients for the cultivation of algae biomass for biofuel has also received some attention, albeit all objections, on the ground that human life support requires availability of phosphorus for human existence than in the cells of algae for the production of biofuel. However, human existence also demands the provisioning of energy from Sustainable sources, and biofuel offers one such source, quite possibly even the source with the most Sustainable Carbon Footprint. Such definitive analysis, of course, obtains only with the analysis of the

Process Engineering Biofuel Production 25 full stream of energy usage for the production the source. Of course, the objections have actually been raised with respect to the dependency of the algae on the NPK – Nitrogen, Phosphorus and Potassium, based on their chemical nomenclature. In addressing direct response to the argument of NPK dependency there are in fact several entry-points of Sustainability engineering—meaning the first step of the Process Chemistry or Process Biochemistry or even Process Microbiology—by which to develop an Algal Phosphorus Recycling Process technology. Yet a fairly good support for the endeavor development [2] in the public domain is one entry-point into the phosphorus recycling technology development, and can be leveraged effectively for such technology.

1.3.10

Process Technology Optimization Engineering

Further specification can only be derived of the dynamics endemic of the algae and so warranting a review of the natural existence of algae. Also algae usually exist in shallow quiescent waters but with variable visibility through the depth and to absorb light-energy, navigate the short distance up towards the surface and back away from it, and do so with cyclicality. Inferential of this endemicity, the microbes do not suffer dynamic contortions that likely distort the dynamic balance between cytoplasmic electrochemical—and mechanical—potentials that supports life. Biomimicry of the algae growth dynamics, however, directly incorporates all aspects of the endemicity in the design, making Biomimicry a critical specification. The reactor has depth along the direction of incidence of sunlight so that the algae may have navigable depth to keep out of the irradiation at will. Further, the medium is not stirred to prevent deformations. Designing for biomimicry has been deemed implementable with a special four-reactor system, though any set of sequential combinations of the four reactors should also invariably meet the specification, and therefore should also enable algae extraction under conditions of uniform density and size. So, by and large an optimal Algae Cultivation Process Technology is a five-bioreactor system process which must include Radiolytic Algae Culture PBR, Continuous Replication Growth PBR, Algae Size-Stabilization PBR, and a set of PBRs for Reaction-specific Amplification in each of which a group of specific Biochemical Reactions is targeted for acceleration; and the fifth bioreactor being the “Bioproduct Bioreactor,” so-called because by design, the algae in this reactor preferentially produce the raw material for the target bioproduct. In particular, the bioreactor has to operate under conditions of “no replication” or “Cell-Function Maintenance” such that virtually all carbon dioxide diffusing into the cell is converted into starch,

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sugar or lipid, without regard to the proportion of conversion as each is suited for biofuel production. Of course, the design of such bioreactor is very daunting because the engineering entails very careful design application of established sciences consistent with the principles of Applied Science. Besides, the reactor must maintain that condition within a single region of large volume but with continuous flow. Of course, there may be several of each type of PBR of each of the five types of reactors. Any Biological Process Technology that does not consist of these minimum sets of PBRs just might struggle along or outright fail.

1.3.11

Base Cultivation Process Technology

The Algae Cultivation Base Process Technology, as composed from the various processes that must be performed with the technology, consists of that process scope that spans all of the Bench-Scale Technology as described in Section 3.7 and further augmented with Biofertilizer Process Technology, Solar Power Technology, Multi-Stage Air Separation Purification Technology, Biomass Nutrients Recovery Technology, and Biomimicry decomposed five-set Cultivation Reactors. Of course, depending on the situation more subprocess units may be added, but these define the essential set of processes for the cultivation of algae biomass for Biorefinery operation.

1.4 Algal Biomass Biorefinery Process Engineering Utilizing algal biomass as feedstock for the operation of Biorefinery can be one of two biorefineries: Bioalcohol Biofuel Biorefinery, Biodiesel Biofuel Biorefinery; and of course the special composite of both biorefineries to produce yet another biorefinery. The Biodiesel Biofuel Biorefinery, however, embodies the Bioalcohol Biofuel Biorefinery as a consequence of the integrated use of the products of the processes at different stages. Strictly the Algal Biomass Bioalcohol Biofuel Process is simply an integrated combination of the Commercial Algae Cultivation Process Technology derivative of the base technology and a Fermentation Process Technology. By extension, Algal Biomass Biodiesel Biofuel Process Technology is also simply an integrated combination of the Bioalcohol Biofuel Process Technology and a Biodiesel Process Technology. Each process technology, of course, does not qualify as a Biorefinery. However, additional raw materials consumed in the production of bioalcohol and the need of recovery biomass wastes define the tasks that must be addressed in process engineering the associated Biorefinery that

Process Engineering Biofuel Production 27 embodies a more complex design of a process than just a mere biofuel process. There are issues that demand the production of additional biochemical products in addition to the production of the biofuel. As is accepted [9], the cell components are continually decomposition while also being built. Effectively there is a dynamic balance under which only cellular function maintenance is supported. So there is the need to supply nutrients to the bacteria cells for feed to support cell maintenance. The analysis of the nutrients required to support bacterial sustenance has been performed [10]; there are about fourteen substances; however, most of these substances are produced with means of chemical pathway. So it is necessary to produce these nutrients from the biomass waste of the algae cultivation process. In effect, while the main biomass feed that is required for the production of biofuel are the starch and lipids, the need for the production of nutrients for the bacteria expands the need to include the waste biomass as well. Similarly with the production of Biodiesel, the option of using ethanol as raw material in the production of the biodiesel sets the stage for further integrated design as per the definition of biofinery, resulting in the necessity integrating the bioalcohol biorefinery to the Biodiesel Biofuel Process technology. However, the production of biodiesel has glycerol as by-product which has also been determined as utilized by bacteria for the fermentation of ethanol as well. The feed raw materials for the Biorefineries, however, is the algal biomass growth operations inexpensively; the task of developing a biorefinery then starts with the stipulation of that source even if only in the abstract; and more so at the pilot plant level and is presented as required.

1.4.1

Resourcing Algal Biomass

The approach simply is this: retrofit previously used process equipment integrated into the form of the algae cultivation process technology. More specifically purchase a used Fermentation Reactor from any used equipment reseller. Many such reactors are suitable as Bubble-Column Algae Growth Reactor. The basic equipment design of the retrofit engineering is well documented [6] as are the Descriptive Reactor Analysis equations for analyzing reactor designs, making it easy to leap forward in the development of Algae Process Technology. Next proceed to retrofit it with an optical system for irradiating the reaction medium with light. Be sure to have a light port in the cap through which light beam would be transmitted into the optical system. Next conduct computational analysis of the process in validation of the design based on the equipment geometry and

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configuration, and determine the optimal operating conditions for the specific algae, and quality of light. Finally start running the process within the region of the optimal operating conditions as determined from the computational analysis. The operational optimal conditions should not be too far from that determined computationally; and once determined, the process is up and running.

1.4.2

Microbes Nutrients-Feed Production

There is a need to supply nutrients to the bacteria cells for feed to support cell maintenance. So it is necessary to produce these nutrients from the biomass waste of the algae cultivation process. In effect, a subprocess must be developed for the production of the nutrients. Although there is no public domain design of such process, a limited form of such a process could be inferred based on the raw materials available for the production. Using the entire biomass of the algae less the starch and the lipids, and the organic nature of the raw materials the subprocess would have to consist of several enzymatic bioreactors, some of which would break down the cellular components and others to synthesize the desired nutrients. Of course, integrated with these would be Separation process units for isolating various components prior to the consumption of each in a different reaction. The integration of the separations process and the reactors are likely to be mixed as both sequential and concurrent in assemblage. The final effluent of the subprocess, of course, is specified as the nutrient ot nutrients such as is the object of the subprocess development.

1.4.3

Fermentation Process Technology

Regarding the technology of the Fermentation Process, for production operations, which is a continuous operating process plant, one of the considerations has been the conservative use of phosphorus because of its essential nature in the support of all lives, and most particularly of human lives. There is therefore the need to conserve the phosphorus of the nutrients feed that is provided to the bacteria to support cellular functions. Although recycling the waste bacterial biomass offers one option for the conservation of the phosphorus, a more controlled approach is the use of a Packed Bed Bioreactor [9] because such eliminates the need for the cycling while still enabling the conservative use of the phosphorus. Both continuous operation and Batch operation of the Packed Bed Reactor is possible. The batch operations procedure is rather trivial. The

Process Engineering Biofuel Production 29 operation of the components follows this progression: Basically, during operation, the feed mixed with other supplementary additives is charged into the Fermentation Reactor to initiate and sustain fermentation. The Fermentation reaction is allowed to proceed up to a level of completion as determined by the reaction time allowed, either by the batch operation or by the residence time in the reactor. At the set time the fermentation reaction product is transferred to the distillation system for distillation. The product of the distillation process should yield grain alcohol.

1.4.4

Biodiesel Process Technology

A process engineering map of the bio-diesel process chemistry into the production process defines the operational process. Even then, operating the process begins very simply with having the algal lipids oil pumped into the Transesterification Reactor. Simultaneously, alcohol ethanol is pumped into another Flow Stirred Tank Reactor where it is reacted with alkaline solution to produce the corresponding ethoxide and then pumped into the Transesterification Reactor where it reacts with the oil at a high temperature to produce bio-diesel, glycerine and soap. At the end of the reaction time, these are pumped into a Separator Process, where upon cooling, the mixture is separated. The result of the transesterification reaction is two immiscible liquids: Bio-diesel-Soap Solution, and Glycerine; and possibly excess alcohol. Very often, the soap and the bio-diesel are homogeneously mixed, while the glycerine is not, and separates out after a while.

1.4.5

Biorefinery Process Technology

Although understandably the bioalcohol process could be operated as a stand alone, the biorefinery being presented here nonetheless is an integrated design, where such reveals the utilization of the glycerol of the biodiesel process in recycle for production of ethanol with the advantage of the extraction of the ethanol used in the production of the biodiesel. In effect, after a certain volume of bioalcohol has been supplied to the biodiesel process, the rest can become available for sale. In the context of biorefinery operation, the lipids and starch of the algae cultivation constitute the feedstock for the biorefinery, and are characterized as such. The Biorefinery [11] aims to the extent possible to develop a Biochemical Pathway distinctly different from Chemical pathway [12] and recycle used process equipment including reimagining the use of available equipment with reengineering of equipment whenever possible.

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By design, the biorefinery [13] quite succinctly, has the algal biomass consisting of the residual-biomass, lipids and starch extracted fed as raw materials by separate streams: the residual-biomass flows into the Nutrients Process, N-P; the starch stream flows into the Fermentation Process, SF-P; and the lipids stream flows into the Biodiesel Process, B-P. Internally Glycerine produced in B-P flows into the Fermentation Process, GF-P, being utilized in the fermentation of ethanol as a sort of recycle of the ethanol from the Distillation Process, FE-D, that got pre-mixed with with the hydroxide in AH-M and then fed into B-P. The nutrients from N-P are partially fed to the SF-P and GF-P and the balance sent to storage as product. the balance of the ethanol from FE-D is also sent to storage as bioalcohol biofuel and all the biodiesel from B-P sent to storage as biodiesel biofuel. The integrated operation, as biorefinery [13] produces nutrients in addition to biofuels, and most of all, in satisfaction of the qualification for biofinery, provides another avenue through the nutrients from which additional bio-products could be produced.

1.4.6

Engineering Cost Impact Analysis

The experimental validations of the designs of a Biological Process Technologies composing the Alga Biomass Biorefinery of Figure 1.1, of course, do not conclude the process engineering necessarily. There is the requirement that the process be also cost effective, and generally be able to support the production operation of the corporation to be built on its functionality. Accordingly, the first complete concept design is necessarily subjected to cost optimization through Break-Even Analysis. Admittedly, by the standard approach of production Break-Even Analysis, the proprietary process is first designed and then a list price is evaluated based on the company pricing policy, which often is determined relative to the cost of unit production. However, with Biological Process production, the luxury of pricing policy-based list pricing is not available to the company. Developing Biological Process Technology for competitive market entry is rather tricky, because of the nuances of the relatively slow rate of performance of the bioreactors that must be designed to drive the process. So while with Manufacturing—and Chemical—Technology processes the products obtain rather quickly, with Biological Technology processes this is not the case, so sizing the related process production plant takes a certain sort of “reverse engineering”. Rather the company must research the market and evaluate a market-supportable price, say, Market Price. Then using this

Process Engineering Biofuel Production 31

N-P

GF

SF SF-P

GF-P

FE-D

NaOH

B-P AH-M

Ethanol

Figure 1.1 EnhKnow

Biodiesel

Nutrients

Algal Biomass Biochemical Pathway Biorefinery.

Market Price the company then constructs a Revenue Curve, and develops a biological technology, the corresponding production plant and constructs (based on the process production plant) a Production Cost Curve against the Revenue Curve. Both curves are then extrapolated until a Break-Even Point is reached, and then extended beyond to determine a profitable production volume. Many algae ventures are not operating at their optimal operating conditions and as a consequence the prices of algal products are still deemed high. The main source of this is the relatively higher cost of production due to the very low efficiency of the bioreactors deployed. Now Rational Analyses of current cultivation vessels have, in fact, shown that about 70%-75% of the insolation on vessels is wasted. So then the productivity of the current vessels are between 25% and 30%, correspondingly, 75% to 70% revenue loss is suffered by these algae ventures, without the entrepreneurs even realizing the losses [1]. Recovery of the revenue loss, however, is still possible with using bioreactor for process technologies, the structured design method of which is also proffered, and the substantiation is as follows.

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By a formula [14] of Production Management, the production cost of any Algae Production Operations could be evaluated from

CP = Dp(RcCr + Pc) + OE

(4.1)

where CP is the Production Cost, Rc is bioreactors count, Cr is cost of reactor, Dp is linear annual depreciation factor, and Pc is the cost of rest of process. The significant factor here is the Dp which is about 0.04 for a linear depreciation of capital investment, CI, and for eliciting the import of that factor let the Operating Expense, OE be reduced by, say, 50% (0.5) and the amount reallocated into the CI in purchase of process technology bioreactor and the CP becomes:

CP = Dp(RcCr + Pc + 0.5OpEx) + 0.5OE

(4.2)

As can be seen immediately the contribution of the increase to the CP is just Dp(0.5 OE) which is about 0.02 OE, so the reallocation of the cost to the CI lowers the daily, monthly and yearly production cost by 48% of the pre-allocation OE. Rational analysis suggests that if in addition, the reallocation were to increase production, then revenue growth also gets to be realized; and even if there is no production increase but the bioreactors count, Rc, drops due to higher efficiency of the newer bioreactors (purchased with the 0.5 OE funds) resulting from optimal utilization of raw materials the overall contribution of CI may even drop leading to an even lower production cost such as may obtain with using bioreactor for process technologies the design approach of which is comprised. Of course, now it is known that all that happens. The Process Technology for Algal Biomass Bioalcohol Biofuel Process was specifically designed designed for Engineering Cost Analysis and then Venture Startup Funding assessment. Using prices of equipment in the public domain, the capital investment for the fully operational Process Pilot Plant stage or Venture Execution Phase is guess-estimated at between about $87 million and $130 million. This investment need, of course, is but an estimate worthy of use only as a guide.

Acknowledgment The author is grateful to his associates in working with the collaboration to put together this compilation of the developments that have been taking place since 2009 and are still ongoing engineering technology research and

Process Engineering Biofuel Production 33 development made for Aningba Research and Development Companies, at the moment an unincorporated informal organization through which technologies are being developed for subsequent deployment and licensing.

About the Author Opubo G Benebo has engaged in various roles in the founding of a diversified corporation with Company Business being the development, manufacture and marketing of advanced engineering technologies enabling the ignition of chemical and biochemical reactions in air pollutants incinerators, and in medical and diagnostic technologies, respectively; and so has engaged in entrepreneurial development of base- technologies for supporting chemical and metabolic reactions utilizing electron-particle beam for ignition, as well as the development of software for the computational design and control of such technologies. Opubo G Benebo also authors analysis reports through an issues-analysis company engaged in the analysis of socio-economic issues including societal lifestyles with intrinsic economic values. He is the author of several books on bioreactors including Algae Cultivation Bioreactor Analysis; Bioreaction Analysis; Kinetics and Parameters of Bioreactions in Production Bioreactors; Batch Microbiological Reactors Designs; Bioreactors Computational Designs Structures; Designing Batch Fermentation Reactors; Design Basics; Venture Offer Prospectus; Reforming Business Plan into Offer Prospectus and Offering Memorandum; Securing Venture Financing; An Entrepreneur’s Guide to Startup Strategy-Driven Financing; Founding Business Entities; An Entrepreneur’s Guide to Strategic Planning, Start-ups & Financing; Writing Business Plans; The Ultimate Guide; Demons & African-Warriors; Rational Reflections on Four African Demonic Folklores. He is a Chemical Engineer with B. Applied-Sc and MSc, and is also the founder of a very competitively positioned software company with a dynamic growth-thrust. Founding the software company was part of an overall entrepreneurial endeavour at the founding of the diversified corporation.

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Generally, Opubo G Benebo may be reached through his email address, [email protected], associated with the business operations through which he sustains his entrepreneurial endeavours of the founding and managing of corporations.

References 1. Benebo, O.G., Algae Cultivation Bioreactor Analysis, Okumaye Publishing Co, Inc, USA, 2017. 2. Torrice, M., How to get the good stuff out of chicken manure: Environmental engineers look for sustainable ways to deal with the waste in Maryland. Chem. Eng. News, 94, 21–22, 2016. 3. Benebo, O.G., Business Entities Venture-Startups: An Entrepreneur’s Guide to Planning & Managing Business Startups, Okumaye Publishing, USA, 2016. 4. Mankins, J.C., Technology Readiness Levels, A white paper, Advanced Concepts Office, NASA Office of Space Access and Technology, USA, 1995. 5. Graettinger, C.P., Garcia, S., Siviy, J., Schenk, R.J., Van Syckle, P.J., Using the technology readiness levels scale to support technology management in the DoD’s ATD/STO environments, Special report: A findings and recommendations report, CECOM RDEC STCD, Army CECOM 2002. 6. Benebo, O.G., Batch Microbiological Reactors Designs: Bioreactors Computational Design Structures, Okumaye Publishing Company, Inc., USA, 2015. 7. Greenkorn, R.A. and Kessler, D.P., Transfer Operations, McGraw-Hill, USA, 1972. 8. Mortvedt, J.J., Heavy metal contaminants in inorganic and organic fertilizers. Fert. Res., 43, 55–61, 1995. 9. Benebo, O.G., Designing Batch Fermentation Reactors: Design Basics, Okumaye Publishing Company, Inc., USA, 2013. 10. Neidhart, F.C., Bloch, P.L., Smith, D.F., Culture medium for enterobacteria. J. Bacteriol., 119, 3, 736–747, 1974. 11. Shuangning Xiu, B.Z. and Shahbazi, A., Biorefinery processes for biomass conversion to liquid fuel, in: Biofuel’s Engineering Process Technology, Dr. M.A.D.S. Bernardes (Ed.), InTech Open Science, 2011. DOI: 10.5772/16417. Available from: https://www.intechopen.com/books/biofuel-s-engineering-process-technology/ biorefinery-processes-for-biomass-conversion-to-liquid-fuel 12. Tao, D., Knoshaug, E.P., Davis, R., Laurens, L.M.L., Wychen, S.V., Pienkos, P.T., Nagle, N., Combined algal processing: A novel integrated biorefinery process to produce algal biofuels and bioproducts. Algal Res., 19, 316–323, 2016. 13. de Jong, E. and Jungmeier, G., Industrial Biorefineries and White Biotechnology, Elsevier, Netherlands, 2015. 14. Abramowitz, I., Production Management: Concepts and Analysis for Operation and Control, The Ronald Press Company, USA, 1967.

2 A Renewable Source of Hydrocarbons and High Value Co-Products from Algal Biomass Abhishek Walia1*, Samriti Sharma2 and Saruchi3 1

Department of Microbiology, College of Basic Sciences, CSKHPKV, Palampur, HP, India 2 Department of Biotechnology, Chandigarh Group of Colleges, Chandigarh, India 3 Department of Biotechnology, CT Group of Institutions, Shahpur Campus, Jalandhar, Punjab, India

Abstract In today’s world fossil fuels are decreasing day by day, and there is an urgent requirement to find a substitute fuel to satisfy the energy needs of the people. Around the globe, an earnest interest in alternative, sustainable fuels and feedstocks is developing. Algae have attracted considerable attention as a possible biomass for producing sustainable biofuel. Algae-based biofuels and their co-products offer abundant support in helping to meet the standards of renewable fuels. In contrast with first- and second-era feedstocks, green growth can give an option of feedstock to worldwide bio-refineries for high yield of biodiesel, ethanol and aeronautics fuels without compromising food supplies, rainforests or arable land. The global market for algal biomass is poised for explosive growth in the next ten years. Algae is attracting increased investment and interest from biofuels, petroleum, and agribusiness industries. In this chapter, we explicate the approaches for making algal biodiesel economically competitive with respect to petrodiesel, different algal production systems, and a pathway for producing liquid transportation fuels from algae based on the current state of the technologies. Algae-based biofuels and their high-value co-products offer abundant support in helping to meet the standards of green renewable fuels and could gradually replace an ample percentage of fossil fuels necessary to meet the current growing energy demand. Keywords: Algae, biofuels, biodiesel, photobioreactor, open ponds, pathways, co-products *Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (35–72) © 2019 Scrivener Publishing LLC

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2.1 Introduction Advanced and innovative technologies for the use of different sources of energy must be established to replace fossil fuels. Biofuels are important as an alternative option for fossil fuels. With an increase in the world population, the growing energy demand has put lots of pressure in the nonrenewable resources of the Earth. The total world consumption of essential energy including oil, coal, natural gas, nuclear energy and hydroelectricity was calculated at 11,295 million tonnes of oil proportional [1]. Out of these essential energy sources oil (35%) and coal (29%) shared maximum percentage followed by natural gas (24%) [2]. Reducing the global greenhouse gas concentration to a level below the current level of 500 ppm CO2 would need some drastic reductions in emissions to roll out of all fossil fuels in developed countries by 2050, if there is no change in the reduction of emissions in developing countries [3, 4]. The use of fossil fuels results in the increased accumulation of greenhouse gases (GHGs) in the environment [5, 6]. The transportation sector majorly contributes to global fossil fuel CO2 emissions to the atmosphere (21%), followed by power production. With average economic growth of 3.2% per year to 2030, transport sector contribution would be 23% to total anthropogenic GHG emissions in 2030 [3]. Therefore, there is an increase in the use of biofuels in the world. Thirty billion liters of biofuels are used annually in Europe, and North and South America. According to IEA estimates, bioethanol and biodiesel have the capability to reach 10% of world transport fuel by 2025 [7, 8]. Therefore, several research efforts have been made to develop some alternative sources of energy like first- and second-generation biofuels. Firstgeneration biofuels depend on sugar-rich plants like sugarcane, sugarbeet, maize, rapeseed oil, etc., which has placed remarkable pressure on global food market and food security. This leaves a question mark regarding their possibility to replace fossil fuels, due to competition with food and fiber production which uses the same agricultural land for their production. First-generation biofuels sources were used for nearly three decades but found inadequate to fulfill the rising world requirements. Instead, they posed a threat to global food security, which necessitated a gradual transition to second-generation biofuels. In second-generation biofuels, raw material used for producing biofuels is from lignocellulosic agricultural residues, forest residues, wood processing waste and non-food crop feed stock rather than from food crops. This resolves some of the problems caused by using first-generation biofuels.

A Renewable Source of Hydrocarbons

37

However, there is concern over availability of land and required conversion technologies, which are not so effective and have not touched the scales for commercial exploitation [9]. It is true that first- and second-generation biofuels have immense potential as an alternative to fossil fuels but they have a negative effect on the global ecosystem. Algal biomass, called a third-generation biofuel, has immense potential to replace the non-renewable energy sources. Scientists from all over the world indicate that algal biomass offers great potential which could be harnessed to produce future green transport fuel and also used in the sequestration of CO2 [10]. Algae refer to microorganisms, from microscopic cyanobacteria to macroscopic giant kelp. Algae use sunlight for their growth and convert it into energy in the same manner as do plants. However, algae impart numerous advantages that can be harnessed to produce algal biofuels for future needs. Productivity of algae is high and also has a faster growth rate as compared to other terrestrial energy crops. Algae can be easily cultivated on land not suitable for agricultural crops without giving any competition to food market and security. Algal species can grow on fresh, brackish sea and even in sewage water. Algae can produce up to 60% oil per dry weight of biomass under such conditions [11]. Extraction of algal biomass directly into biodiesel using trans-esterification is an environment-friendly process [12]. Among algal fuels’ attractive characteristics are that they can be grown with minimal impact on fresh water resources, can be produced using saline and wastewater, have a high flash point, and are biodegradable and relatively harmless to the environment if spilled. Algae, as compared to other second-generation biofuels, cost more per unit mass due to its high capital and operating costs, but the fuel yield is 10 to 100 times more per unit area. According to the United States Department of Energy, algae required 15,000 square miles (39,000 km2), which means that only 0.42% of the area of the United States is required to replace all the petroleum fuel. This is seven times less than the area used for corn harvesting in the United States in 2000. Therefore, algae have gained increased attention as a potential producer of biodiesel along with other lipid-based biofuels and several other high-value co-products [13, 14]. In this way, it is important to find out the means for reduction in the cost of algal biofuel processes, by using cheap and cost-effective raw materials and producing high-value co-products so that it can also compete with the cheap fossil fuels presently in the market. With increasing health consciousness among people, research has been diverted and focused on developing novel products with functional ingredients from algae. Algae deliver some high-value compounds along with biofuel, which imparts good health benefits as polyunsaturated fatty acids, polysaccharides,

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pigments, minerals, vitamins, enzymes, and bioactive peptides. Algal biomass can also be used as a source of biofertilizer to increase the growth of plants. This chapter briefly reviews the potential of algae as a source of hydrocarbon and production of high-value co-products.

2.2 Algal Biomass Production There are different pathways for the algal biomass production. We know that phototrophic algae under natural growth conditions absorb sunlight and nutrients from the aquatic habitats and assimilate carbon-dioxide from the air. There is benefit to using natural conditions, especially sunlight, as a free natural energy resource for commercial production; however, there is also limitation of this natural pathway, as there are seasonal variations in sunlight and it may be limited due to diurnal cycles. Because of limitations in the natural condition for algal growth, artificial means like fluorescent lamps are being employed [15, 16]. This artificial lighting is beneficial for continuous production, but it requires significantly higher energy output. The electricity supply for this artificial energy is employed from fossil fuels; thus the primary aim of producing price-competitive biofuel from the algal is negated and thereby also increases the carbon footprints system [17, 18].

2.2.1 Growth Conditions Algae are photosynthetic organisms, so they require sunlight and CO2 to increase biomass. The optimal ranges of these factors are:

2.2.1.1 Temperature Most commonly cultured species of micro-algae tolerate temperatures between 16 and 27°C. Temperatures lower than 16°C will slow down growth, whereas those higher than 35°C are lethal for a number of species [19, 20].

2.2.1.2 Light Intensity Light intensity plays an important role, but the requirements vary greatly with the culture depth and the density of the algal culture: at higher depths and cell concentrations the light intensity must be increased to penetrate through the culture (e.g., 1,000 lux is suitable for some small lab flasks, but 5,000–10,000 might be required for larger volumes) [19, 21].

A Renewable Source of Hydrocarbons

39

Light may be natural or supplied by fluorescent tubes. Too high light intensity (e.g., direct sunlight, small container close to artificial light) may result in photo-inhibition. Also, overheating due to both natural and artificial illumination should be avoided. Fluorescent tubes emitting either in the blue or the red light spectrum should be preferred as these are the most active portions of the light spectrum for photosynthesis. The duration of artificial illumination should be a minimum of 18 hours of light per day, although cultivated phytoplankton develops normally under constant illumination [22–24].

2.2.1.3 pH The pH range for most cultured algal species is between 7 and 9, with the optimum range being 8.2–8.7 [25].

2.2.1.4 Aeration and Mixing Mixing is necessary to prevent sedimentation of the algae, to ensure that all cells of the population are equally exposed to the light and nutrients, to avoid thermal stratification (e.g., in outdoor cultures) and to improve gas exchange between the culture medium and the air. The latter is of primary importance as the air contains the carbon source for photosynthesis in the form of carbon dioxide. For very dense cultures, the CO2 originating from the air (containing 0.03% CO2) bubbled through the culture limits the algal growth and pure carbon dioxide may be supplemented to the air supply (e.g., at a rate of 1% of the volume of air). CO2 addition furthermore buffers the water against pH changes as a result of the CO2 /HCO3 balance [26, 27]. Depending on the scale of the culture system, mixing is achieved by stirring daily by hand (test tubes, Erlenmeyer’s), aerating (bags, tanks), or using paddle wheels and jet pumps in open ponds. However, it should be noted that not all algal species can tolerate vigorous mixing and it is necessary to know or experiment to create the best algae growing conditions [28].

2.2.1.5 Salinity Marine phytoplanktons are extremely tolerant to changes in salinity. The best algae growing conditions for most species is at a salinity level that is slightly lower than that of their native habitat, which is obtained by diluting sea water with tap water. Salinities of 20–24 gL–1 have been found to be optimal [23].

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2.2.2 Photoautotrophic Production There are different pathways for the production algal biomass, but photoautotrophic production pathway is one of the most economically feasible for large-scale production of algal biomass. There are two systems deployed based on closed photobioreactor technologies and the open system. The viability of both systems depends upon the algal strain and the costs of water and land [29].

2.2.2.1 Open Pond Production Pathway This pathway for the algal cultivation has been used since the 1950s. This open pond system can further can further be classified into natural waters and artificial pond (raceway ponds) or containers. The raceway ponds are generally closed loop; these are oval shaped and have recirculation channel and are 0.2–0.5m deep. There is proper provision of mixing and circulation so as to stabilize algae growth and productivity. The raceway ponds are built with concrete. Nowadays compacted earth lined ponds utilizing white plastic have also been used [30, 31]. Algal broth and nutrients are added in the paddlewheel and then circulated with the help of closed loop to the harvest extraction point in continuous production culture. Sedimentation is prevented by the paddlewheel. Surface air fulfilled the requirement of CO2, but in the case of submerged culture aerator may be used to enhance the absorption of CO2. This is the cheaper way of large-scale algal biomass production and also has lower energy requirements. The maintenance and cleaning are easier in the open pond system. The main problem with the open pond system is to maintain the highly selective environment due to the threat of contamination and pollution from other algal species. It is possible to maintain the monoculture by maintaining an extreme environment, but only few algal trains are suitable for this, like Chlorella and Spirulina [32–34].

2.2.2.2 Closed Photobioreactor Systems This system overcomes the major problems which are associated with the open pond system. Pollution and contamination problem with the open pond system are not a problem with the closed photobioreactor system, so this system can be used for the production of high-value product in cosmetic and pharmaceutical sectors. Single species of microalgae can be used for a prolonged period without the risk of contamination. The harvesting cost is less and productivity is higher in a closed photobioreactor system.

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The setting up cost of a closed system is higher than that of the open pond system. To capture the sunlight, an array of straight glass/plastic tubes are arranged horizontally, vertically, inclined. Proper recirculation of the algal culture is done with the help of a mechanical pump. Agitation is required for the proper mixing and exchange of different gases and nutrients in the system. There are different types of photo bioreactor systems like flat-plate photobioreactor, tubular, column and closed photobioreactors [35]. Flat-plate photobioreactors are the earliest form of the closed system. The flat-plate photobioreactors have a larger exposed surface area for illumination and they also have high density of photoautotrophic cells. These are generally made up of transparent materials, so that they can capture a large amount of solar energy and the dense culture flows across the plate. These are well suitable for the large-scale biomass culture of the algal cell, because of high photosynthetic efficiency and squat accumulation of dissolved oxygen [36]. Tubular photobioreactor are more suitable for outdoor mass culture; the length of the tubes is the design limitation, as it is dependent on the O2 accumulation potential, pH variation and CO2 depletion. These are not indefinitely scale-up, so large-scale production plants are formed by integrating a different multiple reactor unit. The open pond system is less efficient as compared to the closed photobioreactor system. There are several factors which affect the productivity of the open pond system, like temperature fluctuation, evaporation loss, incompetent mixing, deficiency of CO2 and light limitation. Evaporation loss leads to cooling, which changes the composition of the growth medium, and this has an injurious effect on algae growth. It is very difficult to control the temperature fluctuation due to seasonal variations and diurnal cycles. Light cannot penetrate in the depth and there is also CO2 deficiency due to diffusion; both of these also reduced the productivity of the algal biomass [37]. Closed photobioreactor systems overcome the limitation of the open pond system. The biomass growth (required 40–50% carbon) algae depend on a sufficient supply of a carbon source and light to carry out photosynthesis, but in case of microalgae may assume many types of metabolisms like autotrophic, heterotrophic, mixotrophic, photoheterotrophic and are capable of a metabolic shift as a response to changes in the environmental conditions. There are different other sources, rather than the organic carbon or substrate, which are required for the growth of the algae, like vitamins, salts, nitrogen and phosphorous. They also required the optimal parameters oxygen, carbon-dioxide, pH, temperature, light intensity for the formation of the desired product [38, 39].

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2.2.3 Harvesting and Dewatering of Algal Biomass Micro-algal strains due to their microscopic nature (typically 3-20 microns) and their low concentrations (typically less than 2 g algae/L) are difficult to harvest. The standard algae culture has approximately 1g of algae in 5,000g of water. So the standard algae cultivation has approximately 99.98% water and only 0.02% algae. Due to high water content, harvesting and dewatering of algae become expensive and can account for 30% of total production costs. Methods for the harvesting of algae include concentration through centrifugation, foam fractionation, flocculation, membrane filtration and ultrasonic separation. All these harvesting methods contribute to the total cost of algae biomass. [40, 41].

2.2.3.1 Flocculation Flocculation is the agglomeration of algae cells. This can cause cells to fall out of suspension. In general, algae cells carry negative charges on the surface, which can cause the cells to form large clumps or flocs when neutralized. These flocs can be more readily separated from the growth medium. Many different means have been investigated to induce flocculation of algal cells.

2.2.3.2 Chemical Flocculation Inorganic molecules, such as aluminum sulfate, ferric sulfide or lime, can neutralize the cells’ charge or reduce it and thus the cells form clumps. One other method of chemical flocculation is the use of highly charged organic molecules known as polyelectrolytes. In addition to neutralizing the charge these polyelectrolytes can also physically link cells together, which helps form very stable flocs. They also do not have the toxicity that many of the inorganic molecules cause. This makes them a much more attractive chemical option for flocculation [42].

2.2.3.3 Electroflocculation It is the process of forming flocs by introducing an electric current into the culture. This method is very effective, resulting in very high separation efficiencies. Electrofloculation can also be done in very large cultures without consuming a high amount of electricity. This has made this method increasingly popular.

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2.2.3.4 Biofloculation This can be stimulated by limiting nitrogen or altering pH and dissolved O2 levels. Addition of another algae like Scenedesmus obliquus also leads to flocculation. Although this method does not need any additional cost for flocculants, it generally requires a longer time [43].

2.2.3.5 Magnetic Separation of Algae This method is also based on flocculation principle. Magnetic nanno particles (MNPs) are mixed in appropriate doses to the cell culture and magnetic field is applied externally. The cells aggregate towards magnet, leaving a clear liquid medium which is decanted to collect biomass. This method requires less time and is economic since these MNPs can be reused [21].

2.2.3.6 Dissolved Air Flotation DAF is often coupled with some form of flocculation treatment. The cells are usually flocculated first and then air is bubbled through the liquid causing the flocs to float to the surface for easier harvesting. The bubble geometry can play a large role in the efficiency of DAF.

2.2.3.7 Filtration Filtration is the action of flowing particles onto a screen, which separates substances according to particle and screen pore size. Filtration can be an effective method for harvesting larger strains of algae. The algae Spirulina sp. can be simply filtered from the culture medium. Although filtration can be inexpensive and an effective method for harvesting algae, it has the major issue of filter fouling and clogging, which limit its application in large-scale cultivation [19, 20].

2.2.3.8 Centrifugation Centrifugation is a widely used separation method used in many liquidsolid separations. This technology utilizes centrifugal forces to separate substances of different densities. Separation efficiency is dependent upon the size of desired algal species. Although very effective, centrifugation is considered unfeasible in large-scale algae culture facilities due to the high capital and operational costs [44, 45].

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2.2.3.9 Attachment/Biofilm-Based Systems These systems are designed to address the issue of high harvesting costs. In these culture systems, algae are encouraged to attach to a substrate. Once attached, cell proliferation occurs and a biofilm forms. This biofilm is ideal for harvesting because it is already held together and can be easily scraped from the substrate and separated from the culture medium [46].

2.3 Developments in Algal Cultivation for Fuel By Using Different Production System Algae are an important resource of biofuels and bioenergy production. The main work has been conducted on improvement of algal strain selection, maintenance of pure species, growth and cultivation of algae biomass [47]. Algae lack the distinct organs, contain only green chlorophyll, which is masked by photosynthetic pigments that give them a distinguished color. These photosynthetic pigments are made up of four different types of pigments: red, brown, blue and gold. Some algae are microscopic (phytoplankton) and able to float in surface waters due to high lipid content while other are macroscopic (seaweeds) which attached to rocks and other structures. The macroalgae have not been fully utilized as a biofuel resource. The algae are cultivated from a variety of systems from open air ponds to closed bioreactors under controlled environmental conditions. The temperature requirement for algal growth is dependent upon the species and strain cultured. A temperature lower than 16°C will normally reduces algal growth, whereas a temperature more than 35°C is lethal for a number of species. The optimal temperature for algal (phytoplankton) growth ranged from 20–30°C. Open pond systems are used for the majority of algae especially for high oil content algae. The open pond system has been categorized into two types; natural waters (lakes, lagoons and ponds) and artificial ponds or container. The major advantage of the open pond system is that it is easy to operate or construct and has minimum operating cost. However, the limiting factors in open pond systems are poor light utilization by microalgal cells, poor diffusion of CO2 to the atmosphere, evaporation losses and requirement of large land area which make it susceptible to environmental fluctuations. Closed bioreactors are closed, controlled systems with equipment that provide ideal conditions for high algae cultivation productivity. The commercial cultivation of algae mainly prefers closed photobioreactors because it facilitates better control of the pure culture by providing

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optimal growth conditions such as temperature, exposure of lights, mixing, culture density, pH levels, gas supply, amount of carbon dioxide and water. Various type of Photobioreactors are discussed below.

2.3.1 Stirred Tank Photobioreactor A stirred-type bioreactor is a conventional bioreactor where mixing is achieved by mechanical agitation. The core components of a stirred tank bioreactor are the impeller or agitator, which perform a wide range of functions, including aeration, heat and mass transfer, and mixing for homogenization [48]. Sometimes baffles are used to reduce vortexing through stirring. In a stirred type bioreactor 70–80% volume of the bioreactor is filled with liquid which allows adequate headspace for liquid droplets from the exhausted gases or to accommodate any foam that may develop. In order to prevent excess foaming, supplementary impellers are installed for foam breaking. CO2 is aerated at the bottom of bioreactors through air sparger. Stirred tank photobioreactor have high mass transfer rate, which leads to lower incidence of dark zones inside the reactor and higher biomass productivity. The major disadvantage associated with this method is low surface area to volume ratio, which decreases light harvesting efficiency [49]. Moreover, mechanical agitation produces excess of heat than the sparging of compressed gas; hence stirred tank photobioreactors are difficult and expensive to operate and maintain. Modifications in stirredtype photobioreactors can be made by developing new oxygenation which reduce shear, exploiting different protective agents, enhancing existing impellers and agitators.

2.3.2 Vertical Tubular Photobioreactors A vertical tubular photobioreactor is the most suitable bioreactor because it allows better penetration of light through transparent vertical tubes. These photobioreactors have a large surface area which allows cultures to be circulated either with an air pump or by an airlift system. Based on the mode of liquid flow, vertical tubular photobioreactors are categorized into two groups: bubble column and airlift reactors.

2.3.2.1 Bubble Column The alternative to a stirred reactor is a bubble column photobioreactor in which agitation and aeration is achieved by gas sparging. Bubble column photobioreactors are used extensively on commercial scale for production

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of baker’s yeast, vinegar, beer and for the treatment of wastewater. The bubble column has no internal structure, thus fluid flow is driven by bubbles. Sometimes perforated horizontal plates are used to break down or to redistribute the coalesced bubbles produced from sparger. The behavior of bubbles will decide the column hydrodynamics and mass transfer characteristics. Low gas flow rates will be responsible for homogeneous flow and bubbles are evenly distributed across the column cross section; there is no or little back mixing of the gas phase. While during fast or heterogeneous gas flow rate flow, bubbles and liquid tend to rise up from the center of column while a corresponding down flow of liquid occurs near the walls [48]. The external source of light is used in bubble column photobioreactors which affects the photosynthetic efficiency [50]. This bioreactor has several advantages including low capital cost, high surface area to volume ratio, heat and mass transfer, homogenous culture environment, lack of moving parts, and the efficient release of O2 and residual gas mixtures.

2.3.2.2 Airlift Reactors The airlift photobioreactor is a modification of the bubble column photobioreactor but it has two interconnecting zones, the riser (up flowing) and the down comer (down flowing) streams. Gas is aerated through the riser, resulting in gas holdup, driving liquid upward and forming the dark zone. An air bubble drags the liquid from the riser zone inside the concentric tube up to the outside zone. In this way the liquid is recycled downward externally, and finally back to the riser zone; this is the process of “airlift” and received the “light flash effect” to enhance photosynthetic efficiency of illuminated algal cells. This type of liquid circulation is known as internal loop, whereas in the case of external loop riser or dark zone is set outside the vertical tube while short horizontal sections are set at the top and bottom to separate the riser and down comer, forcing the liquid cycle and causing the “light flash effect”. The liquid is circulated in the reactor due to the density difference between the riser and down comer [48]. The mixing of gas is usually better in external loop than in internal loop reactors. The airlift reactor has several advantages, such as high mass transfer efficiency, low energy consumption, good for immobilization of algae on moving particles and well mixing with low shear stress.

2.3.3 Horizontal Tubular Photobioreactors A horizontal tubular photobioreactor is the most used reactor. It is different from a vertical tubular photobioreactor in many ways, particularly with

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the amount of gas dispersion, surface to volume ratio, the nature of the fluid movement, the gas-liquid mass transfer characteristics and the internal irradiance levels. Assembly of horizontal tubular photobioreactor contained multiple tubes arranged in different orientation such as horizontal, spiral, helicoidal, inclined and their variations, but all orientations basically work in the same way. Aside from the arrangement of tubes, tubular photobioreactors vary in the tube length, circulation system, flow velocity and geometric configuration of the light receiver. The diameter of tubes mainly ranged from 10mm to 60 mm, and lengths of up to several hundred meters. The main purpose to introduce multiple tubes is to achieve high surface to volume ratio (above 100/m), which is one of the main advantages of this design [51]. When the diameter of the tubes increases, the surface to volume ratio decreases and this factor has a strong impact on the culture growth. The focusing effect (distribute the light homogenously) prevents the mutual shading and increases radiation intensity. The major disadvantage associated with this method is that the accumulation of O2 to inhibitory levels (above air saturation) generally inhibits photosynthesis in microalgae [52]. Although horizontal tubular photobioreactors are generally considered as the best reactor for scalable and practicable culture system, according to Sanchez Miron et al. [52], it is not economically feasible due to the requirement of cooling system, high O2 and high light intensity results in photo inhibition.

2.3.4 Flat Panel Photobioreactor Flat panel photobioreactors use simple geometry which reduces light penetration depth through the culture suface [53]. A flat panel photobioreactor is used by various researchers for mass cultivation of different algae. A flat panel photobioreactor contain a frame which is covered by a transparent plate on both sides. Algal cell suspension is circulated through a pump. The main features of the flat panel photobioreactor are high surface to volume ratio, absence of mechanical devices for cell suspension and the vertical or tilted inclination from the horizontal of the channels. Moreover, the degassing or gas exchange mechanism of the culture is performed by bubbling air from the base of each channel. The plexi-glass alveolar plates of 16mm thickness are used in a flat panel photobioreactor, owing to high surface to volume ratio [54]. It has been reported that the high photosynthetic efficiencies can be achieved in the flat panel photobioreactor due to the large illumination surface area [38]. The inhibition effect due to accumulation of dissolved O2 concentration is lower in flat-plate photobioreactors as compared to horizontal tubular photobioreactors. But they give lower areal yields as compared to tubular photobioreactors because they have very short penetration depths and do not offer light dilution

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which leads to photo-inhibition of microalgal growth [55]. Flat panel photobioreactors are suitable for small-scale production but not for commercial scale due to the requirement of support materials or compartments, problems in controlling culture temperature and the difficulties associated with hydrodynamic stress resulting from aeration, a problem that has never been reported in tubular reactors [16]. Moreover, there are multiple issues such as high stress damage associated with aeration; sterilization issues; biofouling on surface; and incompatibility with off the shelf industrial fermentation equipment [56]. There are a number of barriers in the commercialization of biofuel production through algae. The most significant is the consumption of energy in the production processes, like energy required for separation of the algal products from the aqueous medium and use of fertilizers and water for algal growth. Most of the algal strains contribute to greenhouse gas emission, mainly due to the current energy inputs required for algal biofuel production. The development of a more enhanced system for low energy separation processes and use of waste water or recovery of nutrients are the main areas for future research. The improved energy can be recovered through combustion or anaerobic digestion from available (non-oil) biomass by ignoring biodiesel. In one of the new technologies, non-oil components of algal biomass will recycle to provide nutrients and energy input to growth and processing of the algae. The downstream processing of algal biomass such as conversion into fuel and high-value coproducts poses a different set of challenges. Through processing technologies such as gasification, pyrolysis, catalytic cracking, anaerobic digestion, and enzymatic or chemical transesterification, whole algal extracts or algal biomass can be converted into different fuels, including ethanol, kerosene, jet fuel, biogas and bio-hydrogen [11, 57–59]. Nowadays highly interesting areas include the development of higher efficiency harvesting and dewatering technologies, improved high-value product extraction and downstream processing, and development of novel method suitable for wet algae.

2.4 Algal Biofuels – Feedstock of the Future Microalgae have received considerable attention due to potential feedstock for producing sustainable transport fuels (biofuels). Based upon the species and cultivation conditions, microalgae produce useful quantities of polysaccharides (sugars) and triacylglycerides (fats). These raw materials (sugars and fats) are used in the production of transport fuels such as bioethanol and biodiesel. Microalgae also produce proteins which could be used as a source of animal feed, whereas some species of microalgae

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produce commercially valuable compounds such as pigments and pharmaceuticals. As a renewable source microalgae have several advantages like high rate productivity, CO2 production, low water requirement for its cultivation as compare to terrestrial plants, cultivated in different environmental conditions, high oil content (20–25% dry matter content), do not compete with food production in the biofuel production, can be used as a source of food or fertilizer, sometimes nutrients required for growth are obtained from wastewater [5, 11, 60–62]. Microalgae biomass can be further converted into biofuels through different routes, for example, fermentation to bio-ethanol and bio-hydrogen and conversion at high temperature to produce bio-crude oil or transesterification of lipids to biodiesel [63]. There are different algal species which produced wide varieties of products:

2.4.1 Biohydrogen The production of hydrogen through biological routes is a promising mechanism because they are environmentally friendly. Currently hydrogen gas is produced by steam reformation process of fossil fuels which increase pollution and continuous decrease in the availability of fossil fuels [64]. Large-scale electrolysis of water can also produce hydrogen gas but it costs more electricity than hydrogen gas as yield. Biological hydrogen production through bacteria, a group called purple non-sulphur bacteria can extract hydrogen from carbohydrates by using light whereas green sulphur bacteria from H2S or S 2O3 . This option is only possible if wastewater is present along with these components [65]. The algae can make hydrogen gas by utilizing sunlight and water, although only in the complete absence of oxygen. Hydrogen production through biophotolysis or photofermentation is a two-step process. Microalgae having high amount of sugar are used in the production of bioethanol. These sugars decomposed to a simple sugar called “glucose” which is fermented to produced ethanol and other components. The ethanol will undergoes distillation or filtration processes to remove water so that it can be used as an ingredient with petrol.

2.4.2 Biobutanol Biobutanol can be used as biofuel, derived through fermentation of the carbohydrates in either micro- or macroalgae. Biobutanol production provides an alternative green solution to replace fossil fuels and reduced environmental issue [66]. The fermentation process of Biobutanol production is essentially based on genus Clostridium because this bacteria has the ability to utilize simple or complex sugars like pentose, hexose, etc. This fermentation

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technology is known as acetone-butanol-ethanol process (ABE) due to the names of main products in 3:6:1. The butanol is about 3% of final concentration in products. During production of biobutanol some factors are considered such as cost of raw material and its pretreatment, small amount of product obtained and its toxicity, and cost of product recovery from fermentation broth. The extracted sugars are sometimes pretreated with lignocellulosic biomass and are used as substrate to make fermentation process economical. The Biobutanol fermentation process has certain limitations such as substrate inhibition, slow growth, butanol toxicity in the medium, and hence, lower cell density. Apart from these limitations, the yield of Biobutanol is also affected by other end products such as acetone and ethanol. To overcome these problems, researchers have developed genetic engineered microbial strains capable with improved Biobutanol yield.

2.4.3 Jet Fuel Airlines have suffered from a number of problems such as an increase in the price of petroleum-based jet fuel, which has tripled over the last 7 years, dependency on imported petroleum oil and deterioration of climate due to greenhouse gas emission. The production of jet fuel through biological sources, also known as bio-jet fuel is an alternative fuel which is less dependent on and greener than petroleum-based jet fuel. The bio-jet fuel has low volume per unit energy and could reduce flight-related GHG emissions by over 60% compared to petroleum-based jet fuels. The microalgae grow in the presence of water, light, carbon dioxide and nutrients, e.g., nitrogen, phosphorus, and potassium. Grown algae are harvested and used to extract oil using hot hexane solvent. The extracted oil is converted to biodiesel which is then used to produce microalgae bio-jet fuel [67]. The rapid growth of microalgae could produce up to 58,700 L oil per hectare (6,275 gal oil/acre) which is higher oil productivity than other crops. The airlines ran the first test of algae-based jet fuel in January 8, 2009, which showed that no modification to the engine was required because of its low flash point and low freezing point [3].

2.4.4 Biogas Biogas is produced through anaerobic digestion of organic material with macro and microalgae [68]. Anaerobic digestion contains about 60–70% biomethane, while the rest is mainly CO2, which can be fed back to the algae. Algae has the ability to remove organic contaminants, heavy metals, nutrients, pathogens from domestic wastewater or convert raw material into high-value chemicals (algae metabolites) or biogas [69]. A photosynthetic

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process of algae produces oxygen which reduces the requirement of external aeration required in aerobic treatment of hazardous pollutants. Accurate selection of algal strain and its proper cultivation produces O2 which is utilized by acclimatized bacteria to biodegrade hazardous pollutants such as phenolics, organic solvents and polycyclic aromatic hydrocarbons [69]. The microalgae can be protected from effluent toxicity or light utilization efficiency can be optimized through well-mixed photobioreactors. The high biomass concentration of algae in photobioreactors can be useful to protect microalgae from light inhibition whereas at low biomass concentration, light intensity should be optimized to avoid mutual shading and dark respiration. Photobioreactors can be designed as closed (tubular, flat plate) or open systems (stabilization ponds or high rate algal ponds). Closed systems are efficiently used; however, they are costly to construct or operate. The main advantage of biogas production from algae is that it can use wet biomass, reducing the need for drying and nutrients can be recovered from digested biomass in liquid and solid phase.

2.4.5 Bioethanol Bioethanol can be used biofuels, derived through fermentation of the sugars in either micro or macroalgae contains more energy per molecule than ethanol which can replace part of the fossil-derived petrol. It is less corrosive to internal combustion engines. In some algal species, starch content of more than 50% has been reported. The cellulose and hemicelluloses of algae can be hydrolysed to sugar which is then converted into ethanol, creating the possibility of utilizing larger part of algal dry matter to ethanol [70]. Algae have some beneficial characteristics over woody biomass because of the absence of lignin, uniform and consisted composition, and lack of specific functional parts such as roots and leaves. The cell walls of algae are mainly made up of polysaccharides which can be hydrolyzed into sugar. Another algaespecific technology is the genetic modification of algae to produce ethanol from sunlight and CO2 [71]. Ethanol production through or from algae have interesting prospects but currently it is only in the initial phase of research. More analyses of the system are required to develop a full-scale production system.

2.5 Biofuel Pathways There are different pathways to producing biofuels from biomass feedstock. There are different steps for biofuel production like cultivation, collection

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and processing. The microalgae were cultivated in saline and fresh water in the open and closed pond system, then algal species which have higher lipid contents were harvested and lipids were released by lysing the algae. The recovered lipids are then converted by chemical processes into hydrocarbon fuel. These hydrocarbon fuels are free from oxygen, so these fuels are suitable for aviation and transportation. The different processes are required for the conversion of algal biomass to energy and these are generally classified into two categories: thermochemical and biochemical conversion. The choice of the conversion depends upon the quantity and type of feedstock, desired energy form, economic and preferred end product (Figure 2.1) [72].

2.5.1 Thermo-Chemical Conversion This category covers the thermal decomposition of organic components into biomass and then fuel products. There are different processes involved in this like direct combustion, thermochemical liquefaction, gasification and pyrolysis. These processes yield high oil and high heating value products like bio-oil, bio syngas, etc. [72–75].

2.5.2 Biochemical Conversion There are different biological processes, which were used for the conversion of biomass into important biofuels like anaerobic digestion, alcoholic fermentation, and biophotolysis. Anaerobic digestion converted organic Carbohydrate

Fermentation

Bioethanol

Pyrolysis

Bio-oil

Gasification

Bio-syngas

Anaerobic Digestion

Biogas

Biophotolysis

Biohydrogen

Algal Biomass

Product Formed Polyunsaturated fatty acid Pigment Protein

Food supplement Pharmacy & Cosmetics

Figure 2.1 Flow chart showing different pathways and products formed by algae.

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wastes into biogas. The biogas primarily consists of methane, carbondioxide and traces of other gases like hydrogen, sulphide. This process involves breakdown of organic matter with the help of microorganisms and then produce a gas having energy content about 20–40% of the lower heating value of the feedstock. This process is better for high moisture content organic waste. There are three sequential stages of anaerobic digestion: hydrolysis, fermentation and methanogenesis [72, 76].

2.5.3 Alcoholic Fermentation This biomass containing sugar, starch or cellulose is converted into ethanol by alcoholic fermentation. The sugar containing biomass is mixed with water and kept warm in a large fermenter. Yeast converted these biomass into ethanol. A distillation process is required to remove the water and other impurities. C. vulgaris microalgae is a good source of ethanol due to high content of starch [72, 76].

2.5.4 Biophotolysis Hydrogen is a clean and efficient energy source and naturally occurs in the molecules. Microalgae have the necessary genetic, enzymatic and metabolic characteristic to produce hydrogen gas. Microalgae act as an electron donor in the carbon-dioxide fixation process or evolved in both dark and light reaction and produce hydrogen in anaerobic condition. Microalgae converts H2O molecules into H+ and O2, then H+ ions are subsequently converted into H2b by hydrogenase enzymes in anaerobic condition [77–81].

2.6 High Value Co-Products from Algal Biomass The world has the experience of using algae for a wide array of products. Marine microalgae have very high content of protein, lipid, vitamins, pigment, anti-oxidants, fatty acids and different types of nutrient supplements, so it was mainly used as a healthy food. There are varieties of algal products available in the market and many more algal products are constantly discovered. The recent interest in use of the algal biomass is in the field of biofuel. There is growing interest in exploiting the algal biomass to produce co-product having great value. Commercialization of algal and algalbased high-value co-products depends upon the market and economic factors. As demand for these products increases day by day, in recent years

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research into increasing the production of algal product has also increased so as to meet the market demand. The algal market is expected to achieve 27,552.11 tons by 2024, extending at a CAGR of 5.32% in the period from 2016 to 2024. The retail value of the micro-algae product was estimated by Pulz and Gross (2004) at US$ 5–6.5 billion, out of which the health food sector generated from algal products was US$ 1.25–2.5 billion, followed by the production of docosahexanoic acid (DHA) at US$ 1.5 billion and aquaculture, US$ 700 million [82–84].

2.6.1 Algae in Human Nutrition The algae species like Chlorella vulgaris, Spirulina maxima, Haematococcus pluvialis and Dunaliella salina are gaining interest as they are rich in carbohydrates, protein, vitamins like A, C, B1, B2, B6, niacin, iodine, potassium, iron, magnesium, calcium and minerals. Because of this they are broadly utilized as dietary supplements. Utilization of algae as a human food started 4,000 years ago in China and Japan, and today Japan is the prime consumer of algae-based product. The method of cultivating leafy algae was given by the Japanese. Porphyra the red algae (nori), Chlorella and Athrospira (Spirulina) have very high content of proteins and their cell wall consists of polysaccharide, which are digestible by human beings; their production required largescale farming practice [85]. Spirulina also produces yellow white protein and phycobiliprotein and these are used in many food industries. The cost of Spirulina is US$20/kg worldwide and the market requirement of Spirulina is about 10,000 metric tons per year. Chlorella is used as a human health supplement; its market value is about US$44/Kg and about 2,000 tons of Chlorella-based product was consumed every year. These are competing with the other plant- and animal-based proteins like soya, milk, eggs, etc. One of the most important health benefits of algal protein is that it also boosts the immune system, reduces fatigue, detoxes the body, improves digestion, increases energy level and appetite and improves the functions of liver, kidney and cardiovascular, so use of algal products solves the problem of malnutrition [86, 87].

2.6.2 Algae in Animal and Aquaculture Feed The growth of the aquaculture depends upon the protein source or accessibility of the fishmeal. Fishmeal is the main limiting factor for the development of aquaculture. The estimated cost of fishmeal is $1,700/ton

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in 2013. 30–60% of the cost of aquaculture operation depends upon the feed. The unavailability of fishmeal leads to the financial vulnerability of operating the aquaculture. The microalgae is one of the best solutions to this problem, cultivated microalgae have all the food supplements that are essential for the production of many farmed fishes, many aquaculture species and shellfish. There is no requirement of processing; some algae can directly be used for nurturing all stages of marine gastropods, larvae of penaeid shrimp, marine fish species and zooplankton. If compared to the $1,700 per ton that fishmeal costs, the algal farming cost is much less (US$400 to $600 per metric ton). Different combinations and concentrate of microalgae blends are available in the market, which are used in aquaculture, e.g., Shell fish diet 1800 is a mixture of five different marine microalgae [88, 89]. In most of the area where there is scarcity of green grasses, livestock farmers use straw and concentrate feed for their animals. This leads to infertility and many diseases in these animals due to lack of proper nutrients. The use of algae with straw and concentrate in the feed surely helps economic livestock production as algae is rich in many proteins and nutrients. Algae is rich in Omega 3 with EPA, so fish growing on algal-based feed have better health than fish grown with soy- or corn-based fishmeal. Chlorella was used as feed in sheep and pigs in different trials and on successful completion of these trials, Chlorella was cultivated commercially as a livestock feed for the production of feed additives. The worldwide animal feed market is about $20 billion, which is much higher than the fishmeal. Both aqua and animal culture producers depend upon the formulated feed being a balanced diet for the best growth, quality and health of the farmed animal [90–92].

2.6.3 Algae as Fertilizer The algal biomass after processing and extracting oil/carbohydrate can be used as bio-fertilizer for plants. There are large varieties of the microalgae, which act as nitrogen-fixator and as soil conditioner. Utilization of the bio-fertilizer in agriculture fields could expand the amount of the carbon, which is accumulated in the soil and this helps in decreasing the greenhouse gas emission by depleting the necessity of fossil-derived fuels by recovering the nitrogen: phosphorus: potassium from the stream of wastewater. Traditionally Cyanobacteria were used in the rice field as an organic source of nitrogen. Blue green algae is available in the market as the brand name ‘Algalization’ and is used as a biofertilizer. It is creating an eco-friendly agro-ecosystem, as it is capable of fixing nitrogen in an anaerobic condition, which brings 25–30 KgN/hectare/season, thus enhancing

56

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the crop yield by 10–15%. It was found that cyanobacteria inoculation in barley, oat, cotton, tomato, radish, sugarcane, chilli, maize and lettuce crops enhance the production. Seaweeds have high mineral content and they enhanced the retain capacity of the soil, so these are used worldwide as stimulant and bio-fertilizer. They are helpful in soil conditioning and this led to increase in rooting, draught and salt resistant, superior crop and fruit yield, enhanced the resistance from bacteria, viruses and fungi and increased the photosynthetic activity. So because of these qualities the demand of algal-based bio-fertilizers increasing day by day. The microalgae Spirulina are exploited as iopesticidesas, because they produce different types of allelochemicals like majusculamide-C. Chlorella vulgaris have phytoprotective effects on grape seedlings, which were infected by Xiphinema nematodes [82, 93, 94].

2.6.4 Algae as Recombinant Protein Astaxanthin, β-carotene and C-phycocyanin are the recombinant protein of algae. β-carotene has a large number of applications such as food coloring agent, additives in cosmetics, Pro-vitamin source, etc. The most suitable biological source of β-carotene is the D. salina and it can produce about 14% dry weight. Commercially more than 90% of β-carotene is chemically synthesized [88, 95]. The ataxanthin is used in the field of cosmetics, feed industries and in the nutraceuticals sector. It is a very good antioxidant; it protects human health from UV-light, improves the immune system, is anti-inflamatory, and a source of pro-vitamin A, hormone precursor and very good coloring agents. H. pluvialis microalgae is a good source of astaxanthin and is preferred over the synthetic astaxanthin because of large deposition of the natural pigment. Also, consumer demand is higher for the natural pigment. The natural market of astaxanthin is about $7,150 per/Kg. C-phytocyanin is found in cyanobacteria, cryptophytes and rhodophytes and is a major photosynthetic blue pigment. It is light harvesting proteins called phycobiliproteins [96]. It is used as a nutrient for both humans and animals. It is used as a natural dye for pharmaceuticals, cosmetics and food due to its antioxidative properties [82, 97].

2.6.5 Algae as Polyunsaturated Fatty Acids (PUFAs) Algae are a very good source of polyunsaturated fatty acids. It is essential for human development and physiology. It has been proven that PUFAs reduced the risk of cardiovascular disease. Fish and fish oil are the main

A Renewable Source of Hydrocarbons

57

source of the PUFAs, but they can’t be used as food additive due to fish odour, accumulation of toxins, lesser oxidative stability and the fact that they have a good deal of fatty acids. Vegetarians do not use them in their  diet;  in  all  such conditions algae are better options for PUFAs [83, 84, 98].

2.7 Microalgae in Wastewater Treatment Wastewater provides ideal environmental conditions for the growth of a wide range of microorganisms, especially viruses, bacteria and protozoa. The majority of the microorganisms are harmless and they can be used in biological sewage treatment; sometimes sewage also contains harmful microorganism such as pathogenic microorganisms from sick individuals and a symptomic carrier. The pathogenic microorganisms are mainly viruses, bacteria and protozoa. The viruses are responsible for hepatitis, Bacteria for cholera, typhoid, tuberculosis whereas protozoa are for dysentery, and their eggs (Parasitic worms) are also found in sewage [99]. The extent to which coliform organisms are removed from sewage represents efficiency of disinfecting sewage. The tertiary treatment for removal of pollutant can be accomplished by biological or through chemical methods. But a biological treatment appeared to be the best as compared to chemical treatment because of cost effectiveness and the fact that it does not produce secondary pollutants. The main aim of tertiary is to remove phosphate, ammonia and nitrate which are about four times more expensive than primary treatment. Algal cultures have been used for wastewater treatment for 75 years, which has allowed the mass production of different strains Chlorella and Dunaliella. Microalgae use inorganic nitrogen, phosphorus, heavy metals and some toxic organic compounds for their growth and do not produce secondary pollution. The other beneficial effects are the production of oxygen and disinfecting effect (due to increase in pH during photosynthesis). The microalgae are used for wastewater because of its photosynthetic capabilities of converting solar energy into useful biomass and incorporating nutrients such as nitrogen and phosphorus causing eutrophication [100]. Palmer [101] surveyed microalgal genera on wastewater stabilization ponds and found frequently occuring algaes were Chlorella, Ankistrodesmus, Scenedesmus, Euglena, Chlamydomonas, Oscillatoria, Micractinuim and Golenkinia. In Central Asia, six lagoon systems of algal were surveyed by Erganshev and Tajiev [102]. Long-term data of their analysis revealed that the Chlorophyta was dominant both in variety and quantity followed by

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Cyanophyta, Bascilhariophyta and Euglenophyta. Palmar [103] listed algae on the basis of their tolerance to organic pollutant which belongs to 60 genera and 80 species. The most tolerant genera were found to be Chlorella, Scenedesmus, Chlamydomonas, Oscillatoria, Euglena, Nitzschia, Naviculer and Stigeoclonium. More than 1,000 algal taxa had been reported which are pollutant tolerant and belongs to 240 genera, 725 species and 125 varieties and forms. The substantial requirements of microalgae for wastewater treatment systems is the land [104], So efforts are being made to use hyperconcentrated algal cultures. This proves to be highly efficient for the removal of Phosphate and Nitrogen within very short periods of time, e.g., less than 1h [105]. The efficient algal system can treat agro-industrial wastes [106, 107], Livestock wastes [108], human sewage [109], industrial wastes [110], piggery effluent [111], and the waste from food processing factories. The algal system can also treat toxic minerals such as bromine, arsenic, scandium, tin, mercury, cadmium and lead.

2.8 Economics of Algae Cultivation The in-depth analysis of production costs showed that the majority of costs are due to the price of inputs, production of algal biomass and harvesting, whereas the total production is sensitive to variations in annual areal productivity, plant production capacity, algal lipid content and the price of carbon [112]. Davis et al. [113] attempted to combine data of major US national laboratories such as data as resource assessment, technoeconomic and life cycle analysis models into a harmonized baseline model which represents near-term production of algal biodiesel. From this, they have found that the model is sensitive to estimate algal lipid content and composition production and downstream process (Table 2.1). The major challenge associated with economic feasibility is the supply cost of CO2 [114]. It can be overcome if the algal plant is situated close to a point source where CO2 is released as waste product (Carbon supply will be free). The problem associated with flue gas is the costs associated with transport and distribution of gas and it will also contaminate algal productivity [115]. The usages of water, energy, nutrients and recycling have an impact on cost and environment. The recycling process mainly involves the gasification of oil-extracted algal biomass into crude oil and electricity or through anaerobic digestion [113, 116–118]. Although there is extensive

Packed bed reactor

Ponds

Packed bed reactor

Ponds

Ponds

Tubular Packed bed reactor

2.

3.

4.

5.

6.

Cultivation method

1.

Sr. no.

4100 t year-1 (100 ha)

10 MG lipid year-1 (1950.58 ha)

41 t ha-1 year-1

25%

25 g m-2 day-1

10 MG lipid year-1 (1950.58 ha)

2100 t year-1 (100 ha)

25%

1.25 kg m-3 day-1

500 ha

500 ha

Scale

21 t ha-1 year-1

40%

40%

Lipid content

24 g m-2 day-1

40-90 g m day-1

-2

Biomass productivity

5.51

6.57

USD per kg algal biomass

Table 2.1 Estimation of cost of algal biodiesel production along with key assumptions.

2.25

4.78

4-7.50

25-33

USD per L algal lipid or biodiesel

[122]

[122]

[124]

[124]

[123]

[123]

(Continued)

References

A Renewable Source of Hydrocarbons 59

Flat-panel Packed bed reactor

Packed bed reactor

Packed bed reactor

Packed bed reactor

Ponds

Ponds

Ponds

Packed bed reactor

8.

9.

10.

11.

12.

13.

14.

Cultivation method

7.

Sr. no.

20–50% 20–50% 25% 50% 20–40%

0.65–1.95 kg m-2 day-1

20–30 g m-2 day-1

25 g m-2 day-1

30–60 g m-2 day-1 or

80–120 t ha-1 year-1

100000 t year-1

400 ha

10 MG lipid year-1

333.3 ha

30 m3

90 t ha-1 year-1

64 t ha year

200 t year-1

-1

90 t ha-1 year-1

Scale 6400 t year (100 ha)

Lipid content

-1

-1

Biomass productivity

89

16.2

7.91

USD per kg algal biomass

1.25–2.5

0.42–0.97

3.54

1.3–2.53

2.29–4.34

USD per L algal lipid or biodiesel

Table 2.1 Estimation of cost of algal biodiesel production along with key assumptions. (Continued)

[112]

[118]

[126]

[121]

[121]

[125]

[125]

[122]

References

60 Liquid Biofuel Production

A Renewable Source of Hydrocarbons

61

publication regarding anaerobic digestion of human and animal wastes, much less information is available regarding anaerobic digestion of algal biomass [119, 120]. Recently, harvesting is considered as a major area for large economic implications [114] but in-depth analysis conducted by Davis et al. [113] showed that 95% harvesting, 85% extraction and 78–85% conversion efficiencies resulted in 65% conversion of algal lipids to fuel. The economics can be substantially improved if the conversion efficiency is improved by harvesting method such as flocculation with centrifugation [118], settling of algal biomass followed by air flotation and centrifugation [113], belt filter press [121], and origin oil’s electro water separation technology [112] have been proposed as viable options. There are various technologies which have potential to revolutionize biomass production, but all are demonstrated at small scale. From example, Norsker et al. [122], estimate the production cost between 5.5–7.9  USD kg-1 biomass which can decrease 60% of price of biomass production such as 1.7 USD kg-1 dry weight for open ponds and 0.9 USD kg-1 dry weight in closed reactors through improvement in solar radiation by site selection, free sources of nutrients and CO2 and an increase in photosynthetic efficiency (Figure 2.2). The overall sentiment of the literature was summarized by Brownbridge et al. [112] with the conclusion that a plant which has algal biodiesel as its primary product can be commercially feasible with current technologies, prices and forecasts. The fundamental understanding of the limiting factors of algal biodiesel will help in accurately assessing the economic feasibility which are mainly areal productivity, nutrients supplies, CO2 and energy requirements in cultivation and down streaming processing.

2.9 Problems and Potential of Alga-Culture The algae established symbiotic relationships with bacteria in aquatic ecosystems. Algae support the aerobic bacterial oxidation of organic matter by the release of oxygen through photosynthesis whereas released CO2 and nutrients are used for algal growth (Table 2.2). The main nutrient sources utilized by algal biomass are ammonium, carbon dioxide and orthophosphate. Oswald [127], and Grobbelaar et al. [128], observed the total amount of released oxygen through algal biomass is 1.5g O2/l and 1.9 g O2/l, respectively. Arceivala [129], calculated that 4-6% solar radiations are useful for treating the pond at 40 N latitude, which produces algal biomass at rate

62

Liquid Biofuel Production Algae and site selection Light

Water

CO2

Nutrients

Cultivation of algae Algal effluent 2.7%

Culture recycle

Harvesting Algal slurry Biomass processing (Dewatering, thickening, filtering and drying)

Algal cake TSS15-25%

Oil extraction by cell disruption and oil extraction

Biodiesel production

Figure 2.2 Microalgae Biodiesel value chain stages.

of 80 kg O2/l ha day. Dry weight of algal cell is due to protein bounded N2 (45–60%) and phosphorus which is essential for the synthesis of phospholipids, nucleic acids and phosphate esters. Algae remove nitrogen and phosphorus from nutrient load within a few hours to a few days for its own growth and development [105]. In comparison to other treatment methods, oxidative ponds are considered effective for nutrients removable because it supports the growth of some species. Increasing pH and BOD causes the sedimentation of phosphorus, ammonia and hydrogen sulphur removal. In algal pond high pH can also lead to pathogen disinfection [130]. The heavy metals removal by algae shows changes in species such as lead by Chlamydomonas, Cadmium, copper and zinc by Chlorella vulgaris, Chrome by Oscillatoria and molybdenum by Scenedesmus chlorelloides [131–135].

A Renewable Source of Hydrocarbons

63

Table 2.2 Advantages and disadvantages of algae in biodiesel production. Sr. no.

Advantage

Disadvantages

1

Algae fuel contains no sulfur, non-toxic in nature

Relatively new technology

2

Highly bio-degradable

Produces unstable biodiesel with many polyunsaturates

3

High Yield per acre (7-31 times greater than palm oil).

Poor biodiesel performance

4

Grow anywhere

5

Rapid growth rates

6

Reduce carbon emissions based on where it’s grown

7

Higher Polyunsaturated content suitable for cold climate

8

Extracts of algal oil can be used as livestock feed and even processed into ethanol

2.10 Conclusion First- and second-generation biofuels resource limitations clearly indicate that they are inadequate to meet the current global demand of future transport fuels. Even so, the third-generation biofuels produced from algae still need more advanced research and development activities to decrease the cost of algal biomass production to a satisfactory level that could compete well with cheap fossil fuels available in the market. Bio-fixation and wastewater treatment technologies are economically feasible when compared to algal fuels. In algal biofuels, production systems like open pond cultivation method, being much cheaper to construct and operate, can be scaled up to several hectares and thus appear to be the method of choice for commercial microalgae production. Algae can produce and release different types of metabolites and bioactive compounds, which shows wide applications in industries like food, pharmaceutical, cosmetics, and agriculture. Production of algal biofuels will become competitive, if we consider the production of these higher value co-products such as bio-fertilizers,

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human feed, animal feed, pigments, vitamins, cosmetics, biopolymers, etc. To make the process more feasible in terms of cost, many companies are showing interest in entering into algae-based value-added products, which helps in minimizing the high cost of biodiesel production. Hence, it appears that the combination of algal fuels with carbon bio-fixation, wastewater treatment, and other high value co-products could possibly assist as a cost-effective and eco-friendly stage to achieve the aforesaid goals.

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3 Waste Biomass Utilization for Liquid Fuels: Challenges & Solution Sourish Bhattacharya1*, Surajbhan Sevda3, Pooja Bachani1,2, Vamsi Bharadwaj1,2 and Sandhya Mishra1 1

Division of Biotechnology and Phycology, CSIR - Central Salt and Marine Chemicals Research Institute, Bhavnagar, India 2 Academy of Scientific & Innovative Research (AcSIR), CSIR - Central Salt and Marine Chemicals Research Institute, Bhavnagar, India 3 Department of Bioscience and Biotechnology, Indian Institute of Technology, Guwahati, India

Abstract In the present scenario, liquid fuels are consumed globally at a rate of 95.24 barrels per day, which is increasing day by day due to continuous population growth. To meet this exigency, the focus is laid on utilizing waste biomass for generating sufficient liquid fuels, e.g., biobutanol, bioethanol and biodiesel, in a cost-effective and eco-friendly manner. Hydrolysate of bagasse, cane trash and rice husk containing reducing sugars may be utilized for bioethanol production as well as growing microalgae for generating microalgal biodiesel at commercial scale. Similarly, hydrolysate of sweet sorghum bagasse, cassava bagasse and sugarcane bagasse can be utilized for bioethanol production at commercial scale. One of the major problems with lignocellulosic hydrolysate is the presence of non-fermentable pentose sugars, which can be solved by developing a model organism that can ferment all sorts of sugars for generating bioethanol and biobutanol. However, development of oleaginous yeast through metabolic engineering which can ferment all sugars (fermentable and non-fermentable sugars) and can accumulate lipids for generating biodiesel may be a sustainable approach. For increasing microbial lipid accumulation, overexpression of TAG biosynthesis-enhancing enzymes, regulation of TAG bypass routes and blocking competing pathways are better routes. Similarly, a novel microalgal strain may

*Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (73–88) © 2019 Scrivener Publishing LLC

73

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be considered which may utilize all fermentable and non-fermentable sugars for generation of microalgal biodiesel as well as bio-ethanol and bio-butanol from the hydrolysate of the residual biomass. Keywords: Waste biomass, biofuel, bio-ethanol, metabolic engineering, yeast fermentation

3.1 Introduction Lignocellulosic biomass is one of the abundant organic mass in the hemisphere, which is about 50% of total biomass present in the biosphere [1]. However, the major portion of hemicellulose consists of pentose sugars and its conversion to alcohol is a major challenge through fermentation. In the past few years, some developments pertaining to conversion of pentose sugars to alcohol have been made, but still there are certain limitations to such technologies with respect to commercially viable processes [1]. One of the major problems encountered during direct fermentation of lignocellulosic hydrolysate to alcohol is fermentation of xylose and existence of microbial inhibitors in the hydrolysate [2]. For decades, researchers have used metabolic engineering in yeast S. cerevisiae for improving fermentation efficiency, which involves a modified genome shuffling for xylose/pentose sugar fermentation by S. cerevisiae, recursive DNA shuffling with recombination of P. stipitis genome with S. cerevisiae, and development of recombinant yeast strain ScF2 which can utilize xylose for ethanol production [3–6]. A major component of lignocellulosic biomass hydrolysate is xylose [7, 8]. Xylose fermentation is needed for effective conversion of lignocelluloses into fuels and chemicals [9–13]. Few yeasts are known to ferment xylose directly to ethanol; however, the rate and yield needs to be improved for its commercialization. Most of the wild-type strains of Saccharomyces cerevisiae are unable to convert D-xylose or pentose sugars [13]. The presence of glucose in microalgal hydrolysate represses xylose utilization, so, altering glucose regulation will lead to maximizing conversion of xylose to ethanol. For optimal production, a low amount of oxygen is required for xylose utilization. Respiration can reduce ethanol yields, so the role of oxygen must be better understood and respiration/aeration must be reduced in order to improve ethanol production. At present, researchers have also developed various engineering strategies to improve ability of yeast to ferment D-xylose to ethanol. Mixotrophic and heterotrophic growth of Chlorella sp. utilizing organic carbon from agricultural wastes is a sustainable process to produce biofuel feedstock with advantage of high biomass productivity, less production age and cost-effective carbon substrate [14–21]. Although, Chlorella sp. utilizes

Waste Biomass Utilization for Liquid Fuels 75 all sorts of organic carbon for its growth and lipid production, there are certain species of Chlorella which can utilize non-fermentable pentose sugars such as D-xylose e.g. Chlorella sorokianana can utilize D-xylose for its growth [22]. Since D-xylose is the second most abundant sugar present in lignocellulosic biomass, its effective utilization is desired for cost-effective microalgal biofuel production through mixotrophic approach.

3.2 Waste Biomass and its Types Biomass is derived directly or indirectly from plants or any other source that can be utilized as the energy or any materials in considerable amount. The resource base includes thousands of plant species, terrestrial and Table 3.1 The major crops and waste utilized for bioenergy generation. Crop

Waste

1

Coconut

Fronds, husk, shell

2

Coffee

Hull, husk, ground

3

Corn

Cob, stover, stalks, leaves

4

Cotton

Stalks

5

Nuts

Hulls

6

Peanuts

Shells

7

Rice

Hull/husk, straw, stalks

8

Sugarcane

Bagasse

9

Agricultural crops

Mixed agricultural crops, not limited to crop waste

10

Mixed type

Agricultural crops and waste including non-organic wastes

11

Microalgal spent biomass

After extraction of oil from microalgal biomass, carbohydrates present in the microalgal biomass can be utilized for bioethanol and biobutanol generation.

Source: UNEP (2009) Converting Waste Agricultural Biomass into a Resource. Compendium of Technologies. The International Environmental Technology Centre. Osaka/Shiga, 441.

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aquatic, various agricultural, forestry, industrial residue, forest waste, sewage and animal wastes. Table 3.1 shows various types of crops through large-scale energy plantation, can be one of the most promising energy biomass. For conversion to energy, various types of biomass may be utilized such as wood, napier grass, rapeseed, water hyacinth, giant kelp, Chlorella, saw dust wood chip, rice straw, rice husk, kitchen garbage, pulp sludge, animal dung, etc.

3.3 Major Waste Biomass Conversion Routes There are various types of waste biomass which may be categorized under cellulosic waste biomass, non-cellulosic biomass and agricultural waste. Table 3.2 shows various routes for waste biomass conversion to energy [23]. Table 3.2 Processes used for cellulosic waste biomass conversion to energy. Technology

Conversion process type

Biomass waste

Energy or fuel produced

Biodiesel production

Chemical

Rapeseed, soy beans waste, vegetable oil

Biodiesel

Direct combustion

Thermochemical

Agricultural waste, mixed waste

Heat stream Electricity

Ethanol production

Biochemical (aerobic)

Sugar or starch crops, wood waste, pulp sludge, rice and corn straw

Bioethanol

Gasification

Thermochemical

Agriculture waste, mixed waste

Low or medium producer gas

Methanol

Thermochemical

Agriculture waste, mixed waste

Methanol

Pyrolysis

Thermochemical

Agriculture waste, municipal solid waste

Synthetic fuel, oil (Biooil/ Biocrude), Charcoal

Waste Biomass Utilization for Liquid Fuels 77

3.4 Metabolic Engineering in Yeast for Accumulation of C5 Sugars along with C6 Sugars One of the major advantages of recent technologies over classical genetic techniques is that just one characteristic can be precisely modified, without affecting other properties. Molecular biology approaches have introduced a new dimension. The expression of heterologous genes has substantially increased such possibilities as well. In the past 20 years, impressive progress has been made in developing recombinant strains of Saccharomyces cerevisiae for its industrial applications. The strain development targets are categorized into two broad categories: (1) Improvement of fermentation performance and simplification of the process and (2) Improvement of product quality, e.g., organoleptic and hygienic characteristics.

3.5 Genetic Engineering for Improved Xylose Fermentation by Yeasts Lignocellulosic hydrolysates usually contain glucose, which is known to be a repressor of xylose utilization process, so, by altering glucose regulation maximum xylose conversion could be achieved. Along with glucose, oxygen is another tuning factor of xylose utilization as it requires low amount of oxygen. By considering the detrimental effect of respiration, proper understanding of oxygen involvement in the xylose fermentation process is required. Also, by reducing the respiration rate, ethanol production can be augmented. This is a major problem in the bioethanol industry as the second most abundant carbohydrate in the lignocellulosic biomass hydrolysate is xylose and at present researchers have developed various engineering strategies to improve the ability of yeast to ferment D-xylose to ethanol. Direct conversion of xylose to ethanol can be achieved by improving the wild-type strain of S. cerevisiae by modified genome shuffling method. Recombinant yeast strains were constructed by recursive DNA shuffling with the recombination of the entire genome of Pichia stipitis with that of S. cerevisiae. A potential recombinant yeast strain ScF2 capable of efficiently utilizing xylose was

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obtained by two rounds of genome shuffling which may consume higher xylose concentration for rapid ethanol fermentation as compared to other recombinant strain. Demeke et al. [24], designed an expression cassette (able to utilize D-xylose- and L-arabinose) which was inserted in two copies in the genome of ethanol Red strain (an industrial yeast strain used for bioethanol production). The gene expression cassettes containing 13 genes including Clostridium phytofermentans XylA, encoding D-xylose isomerase (XI), and enzymes of the pentose phosphate pathway were integrated into both the alleles of the PK2 locus (Figure 3.1). The PK2 gene encodes a replaceable glucose-repressed second isoform of pyruvate kinase, which is generally expressed for utilization of non-fermentable carbon sources. Recently another development was made by Wei et al. [25], in the area of engineering S. cerevisiae for effective utilization of xylose wherein to reduce xylitol formation, a new strategy was demonstrated that increased carbon flux towards target products by controlling the process of xylitol secretion. The genomic DNA of the D452-2 strain using primer pairs FPS1-f and FPS1-r, was cloned into the yeast integrative plasmid pRS403 under the control of theTDH3 promoter and CYC1 terminator using SpeI and SalI restriction enzyme sites, yielding the plasmid pRS403-FPS1. S. cerevisiae strain for producing xylitol was constructed by expressing only XR encoded by the XYL1gene from S. stipitis and was given the strain name D10. PS1gene in the D10 strain is deleted to evaluate the effect of xylitol excretion. Thereafter, the PS1 gene present in the D10 strain was deleted to evaluate the effect of xylitol excretion and used for the fermentation studies, which showed all consumed xylose into xylitol within 80 hrs. FPS1 deletion reduced xylitol production by 21% to 30% and increased ethanol yields by 3% to 10% under various fermentation conditions. Another important development was engineering of fast xylosefermenting Saccharomyces cerevisiae through rational design and adaptive evolution. In this case, a xylose-fermenting yeast was constructed by genome integration of xylose-utilizing genes and adaptive evolution, containing Piromyces XYLA to enable the host strain to convert xylose to xylulose; Endogenous genes (XKS1, RKI1, RPE1, TKL1, and TAL1) were over expressed to accelerate conversion of xylulose to ethanol; and Candida intermedia GXF1, which encodes a xylose transporter, was introduced at the GRE3 locus to improve xylose uptake (Figure 3.2). The fermentation studies of the best evolved strain CIBTS0735 consumed 80 g/l glucose and

Waste Biomass Utilization for Liquid Fuels 79 vector pHD22 - arabinose cassette

vector pHD8 - xylose cassette

(a)

(b)

(c)

(d)

10000 bps

(e)

20000 bps

30000 bps

Recombinant industrial strain

Mutagenesis followed by selection of xylose growing mutants Efficient sporulation and mass mating Selection of xylose growing and inhibitor tolerant variants Serial transfer for efficient xylose fermentation

Selection of best xylose fermenting and inhibitor tolerant strain

Figure 3.1 General strategy used for development of a strain with high xylose fermentation capacity and high inhibitor tolerance (Source: [24]). The gene expression cassettes containing 13 genes including Clostridium phytofermentans XylA, encoding D-xylose isomerase (XI), and enzymes of the pentose phosphate pathway were integrated into both the alleles of the PK2 locus. The PK2 gene encodes a replaceable glucoserepressed second isoform of pyruvate kinase, which is generally expressed for utilization of non-fermentable carbon sources.

80

Liquid Biofuel Production S. cerevisiae CCTCC M94055 arg1: :pCpA1/G-XI ty1: : pYIE2-Ty-XI δ: :loxP-zeo-loxPADH1p-XKS1-XKS1tTPI1p-TAL1-TAL1tPGK1p-RPE1-RPE1tFBA1p-TKL1-TKL1tPDC1p-RKI1-RKI1t CIBTS0525

adaptive evolution & single-colony isolation

CIBTS0555

gre3: :pYIE2-GXF1 CIBTS0573 adaptive evolution & single-colony isolation CIBTS0735

Figure 3.2 Protocol for strain development (Source: [26]).

40 g/l xylose media within 24 hours having biomass yield 0.63 g DCW/l and 53 g/l ethanol yield.

3.6 Biofuel from Microalgae through Mixotrophic Approach Utilizing Lignocellulosic Hydrolysate Microalgae are prokaryotic or eukaryotic photosynthetic microorganisms present in a wide range of environmental conditions, both in an aquatic and terrestrial ecosystem. Microalgae as a potential source of biofuel production has many advantages over traditional biofuel crops. Microalgae can be potentially grown on a marginal land with wastewater/seawater for their cultivation. They are not only used for biofuel production, but also for biological sequestration of metals greenhouse gases. Recently, various strains of Chlorella sp. have been studied for CO2 sequestration in order to reduce the effects of global warming with a focus on open cultivation [27, 28]. It is estimated that more than 50,000 species of microalgae exist and they are known to grow rapidly under a variety of environmental conditions and survive under a harsh environment. Thus, it is possible to adapt a species to suit local environments or specific growth conditions. Hence, they can be grown on lands not suitable for agriculture and they could tolerate high salt content.

Waste Biomass Utilization for Liquid Fuels 81 Microalgae can be grown through heterotrophic and mixotrophic cultivation utilizing glucose present in lignocellulosic hydrolysate. Heterotrophic cultivation of Neochloris oleoabundans showed the ability to grow on 10 g/L glucose. Various species of Chlorella grown heterotrophically utilizing glucose with an increased growth rate of 40–85% compared to autotrophic cultures [29, 30]. Chlorella strains upon xylose enhancement were able to grow on pentose sugars through phototrophic cultivation showing improved growth on xylose and no growth changes on arabinose. Rhamnose is having a potential to support Chlorella vulgaris growth at 1.64 g/L, with Chlorella culture at a concentration of 1.5 g/L xylose and being 20% smaller when compared to 1.8 g/L glucose [31, 32]. Microalgal growth can be enhanced by lignocellulosic hydrolysate when accompanied with combination of suitable strain and cultivation conditions, which will stimulate biomass productivity in mixotrophic and heterotrophic cultures. Improved growth of some strains and increase in lipid and biomass productivity in the presence of sugars or acetates is possible in some robust strains [33–35], but in some of the strains biomass productivity may increase, but lipid productivity and other value added compound may decrease [30, 36]. Moreover, cultivation conditions such as availability of light strongly influences an increase in microalgae culture growth [37]. In addition to sugar and acetates, other substances such as phenolics and furans are also constituents of lignocellulose hydrolysates [38–41]. Phenolic compounds can be stimulatory or inhibitory for microalgae, but depends more on its structure and concentration, as well as on microalgal strains used [42–44]. Furans originate from sugars during pretreatment of lignocellulosic biomass. Pre-adaptation is a critical step to increase resistance in microalgal strains when it encounters a concentration of furan above threshold level in the cultivation medium enriched with lignocellulose derived compounds [45]. Finally, the composition of lignocellulosic hydrolysates is dependent on the lignocellulose treatment method. Methods and process hydrolysis conditions should be selected in such a way to achieve optimal feedstock substrates for microalgal cultivation with better biomass yield. Microalgae can potentially provide fuels in various forms [46, 63]. The algal biomass can be utilized for combustion; further, the crude oil taken from the microalgae can be utilized for direct combustion for supplementing other transportation fuels [47], anaerobic digestion of the microalgal biomass can produce biogas [48, 49] and hydrogen, fermentation of the carbohydrates derived from microalgae can be used for ethanol production

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[50, 51], bioethanol can also be produced directly through microalgal photosynthesis [52, 53]. As compared to terrestrial crops, which take a season to grow and contain a maximum of about 5–20% of dry oil content, microalgae grow quickly and contain higher oil content [54]. The major bottleneck for the production of biofuel from microalgae at a commercial scale is the generation of biomass. This is necessary for the process to be economically feasible and to ensure an uninterrupted supply for the consumer markets [55]. The basic necessities for the cultivation of microalgae are light, CO2, minerals and water. However, the specific requirements are dependent on the species or strain of microalgae that is being cultivated. Microalgae could be mass cultivated in open systems or in closed photobioreactors [56, 57]. Mass cultivation of algae in open ponds has been practiced since the 1950s and is the oldest industrial system [56]. Large-scale open ponds can be constructed with plastic, concrete and bricks for mass cultivation [56, 57]. Basically, these cultivation systems require relatively low construction and operating cost and pilot systems can be made on lands that have degraded and are unfit for agriculture [57, 58]. However, such ponds consume energy, which is required for circulation/agitation. Along with carbon source, microalgae require nitrogen, which is required for their primary metabolism. Lipids are synthesized as a food reserve that may be required during periods of nutritional stress. Phosphorus, in the form of phosphates, is a major nutrient for growth of algae. In addition to these components, trace metals such as Mg, Ca, Mn, Zn, Cu, Fe and vitamins are needed for higher productivity [59–61] as well as an efficient photobioreactor for microalgal cultivation [62]. Further, Jatropha biodiesel waste residues i.e. hydrolysate of deoiled cake of Jatropha (JOCH) and crude glycerol coproduct stream (GL7 and GL8) may be added along with seawater diluted with tap water (1:2) for growing Chlorella variabilis in an efficient manner [63].

3.7 Conclusion Lignocellulosic biomass hydrolysate is a potential source of xylose which may be used for the production of bioethanol through fine tuning of yeast that is capable of utilizing xylose. A number of attempts have been made for improving the industrial strain S. cerevisiae for economic conversion of xylose utilization. Metabolic engineering has played an important role in increasing rate and yield of bioethanol production. However, non-fermentable sugars present in the lignocellulosic hydrolysate may be used for mixotrophic cultivation of microalgae for its biomass generation. Further, lipids may be

Waste Biomass Utilization for Liquid Fuels 83 extracted from the microalgal biomass which can be converted into biodiesel if the lipid profile is suitable. The spent microalgal biomass is an important source of carbohydrates and the spent microalgal hydrolysate may be used for bioethanol and bio-butanol production through microbial fermentation.

References 1. Chandel, A.K., Singh, O.V., da Silva, S.S., Detoxification of lignocellulosic hydrolysates for improved bioethanol production, INTECH Open Access Publisher, London, UK., 2011. 2. Hahn-HäGerdal, B., Lindén, T., Senac, T., Skoog, K., Ethanolic fermentation of pentoses in lignocellulose hydrolysates. Appl. Biochem. Biotechnol., 28, 1, 131–144, 1991. 3. Chiang, S.J., Strain improvement for fermentation and biocatalysis processes by genetic engineering technology. J. Ind. Microbiol. Biotechnol., 31, 3, 99–108, 2004. 4. Dequin, S., The potential of genetic engineering for improving brewing, wine-making and baking yeasts. Appl. Microbiol. Biotechnol., 56, 5–6, 577– 588, 2001. 5. Lin, Y. and Tanaka, S., Ethanol fermentation from biomass resources: Current state and prospects. Appl. Microbiol. Biotechnol., 69, 6, 627–642, 2006. 6. Jeffries, T.W. and Shi, N.Q., Genetic engineering for improved xylose fermentation by yeasts, in: Recent Progress in Bioconversion of Lignocellulosics, pp. 117–161, Springer, Verlag Berlin Heidelberg, 1999. 7. Sukumaran, R.K., Singhania, R.R., Mathew, G.M., Pandey, A., Cellulase production using biomass feed stock and its application in lignocellulose saccharification for bio-ethanol production. Renew. Energy, 34, 2, 421–424, 2009. 8. Maurelli, L., Ionata, E., La Cara, F., Morana, A., Chestnut shell as unexploited source of fermentable sugars: Effect of different pretreatment methods on enzymatic saccharification. Appl. Biochem. Biotechnol., 170, 5, 1104–1118, 2013. 9. Gupthar, A.S., Segregation of altered parental properties in fusions between Saccharomyces cerevisiae and the D-xylose fermenting yeasts Candida shehatae and Pichia stipitis. Can. J. Microbiol., 38, 12, 1233–1237, 1992. 10. Ness, J.E., Welch, M., Giver, L., Bueno, M., Cherry, J.R., Borchert, T.V., Minshull, J., DNA shuffling of subgenomic sequences of subtilisin. Nat. Biotechnol., 17, 9, 893–896, 1999. 11. Heluane, H., Defigueroa, L.I.C., Vázquez, F., Fusion of yeast protoplasts and isolated nuclei of Fusarium moniliforme. Acta Biotechnol., 18, 4, 353–359, 1998. 12. Kordowska-Wiater, M. O. N. I. K. A. and Targoński, Z. D. Z. I. S. Ł. A. W., Application of Saccharomyces cerevisiae and Pichia stipitis karyoductants to the production of ethanol from xylose. Acta Microbiol. Polonica, 50, 3–4, 291–299, 2000.

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13. Zhang, W. and Geng, A., Improved ethanol production by a xylose-fermenting recombinant yeast strain constructed through a modified genome shuffling method. Biotechnol. Biofuels, 5, 1, 1, 2012. 14. Amin, S., Review on biofuel oil and gas production processes from microalgae. Energy Convers. Manage., 50, 7, 1834–1840, 2009. 15. Chen, G.Q. and Chen, F., Growing phototrophic cells without light. Biotechnol. Lett, 28, 9, 607–616, 2006. 16. Liang, Y., Sarkany, N., Cui, Y., Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnol. Lett., 31, 7, 1043–1049, 2009. 17. Zheng, Y., Chi, Z., Lucker, B., Chen, S., Two-stage heterotrophic and phototrophic culture strategy for algal biomass and lipid production. Bioresour. Technol., 103, 1, 484–488, 2012. 18. Garcí, M.C., Sevilla, J.F., Fernández, F.A., Grima, E.M., Camacho, F.G., Mixotrophic growth of Phaeodactylum tricornutum on glycerol: Growth rate and fatty acid profile. J. Appl. Phycol., 12, 3–5, 239–248, 2000. 19. Bhatnagar, A., Chinnasamy, S., Singh, M., Das, K.C., Renewable biomass production by mixotrophic algae in the presence of various carbon sources and wastewaters. Appl. Energy, 88, 10, 3425–3431, 2011. 20. Heredia-Arroyo, T., Wei, W., Ruan, R., Hu, B., Mixotrophic cultivation of Chlorella vulgaris and its potential application for the oil accumulation from non-sugar materials. Biomass Bioenergy, 35, 5, 2245–2253, 2011. 21. Abad, S. and Turon, X., Valorization of biodiesel derived glycerol as a carbon source to obtain added-value metabolites: Focus on polyunsaturated fatty acids. Biotechnol. Adv., 30, 3, 733–741, 2012. 22. Zheng, Y., Yu, X., Li, T., Xiong, X., Chen, S., Induction of D-xylose uptake and expression of NAD(P)H-linked xylose reductase and NADP+-linked xylitol dehydrogenase in the oleaginous microalga Chlorella sorokiniana. Biotechnol. Biofuels, 7, 1, 1, 2014. 23. Dtie, U.N.E.P., Converting waste agricultural biomass into a resource, in: Compendium of Technologies, United Nations Environment Programme, Osaka, 2009. 24. Demeke, M.M., Dietz, H., Li, Y., Foulquié-Moreno, M.R., Mutturi, S., Deprez, S., Verplaetse, A., Development of a D-xylose fermenting and inhibitor tolerant industrial Saccharomyces cerevisiae strain with high performance in lignocellulose hydrolysates using metabolic and evolutionary engineering. Biotechnol. Biofuels, 6, 1, 1, 2013. 25. Wei, N., Xu, H., Kim, S.R., Jin, Y.S., Deletion of FPS1, encoding aquaglyceroporin Fps1p, improves xylose fermentation by engineered Saccharomyces cerevisiae. Appl. Environ. Microbiol., 79, 10, 3193–3201, 2013. 26. Diao, L., Liu, Y., Qia, F., Yang, J., Jiang, Y., Yang, S., Construction of fast xylose-fermenting yeast based on industrial ethanol-producing diploid Saccharomyces cerevisiae by rational design and adaptive evolution. BMC Biotech., 13, 110, 2013.

Waste Biomass Utilization for Liquid Fuels 85 27. Kleinheinz, G.T. and Keffer, J.E., Use of Chlorella vulgaris for carbon dioxide mitigation in a photobioreactor. J. Ind. Microbiol. Biotechnol., 29, 5, 275–280, 2002. 28. Yanagi, M., Watanabe, Y., Saiki, H., CO2 fixation by Chlorella sp. HA-1 and its utilization. Energy Convers. Manage., 36, 6, 713–716, 1995. 29. Sun, N., Wang, Y., Li, Y.T., Huang, J.C., Chen, F., Sugar-based growth, astaxanthin accumulation and carotenogenic transcription of heterotrophic Chlorella zofingiensis (Chlorophyta). Process Biochem., 43, 11, 1288–1292, 2008. 30. Kim, D.G. and Hur, S.B., Growth and fatty acid composition of three heterotrophic Chlorella species. Algae, 28, 1, 101–109, 2013. 31. Hawkins, R.L., Utilization of xylose for growth by the eukaryotic alga, Chlorella. Curr. Microbiol., 38, 6, 360–363, 1999. 32. Gim, G.H., Kim, J.K., Kim, H.S., Kathiravan, M.N., Yang, H., Jeong, S.H., Kim, S.W., Comparison of biomass production and total lipid content of freshwater green microalgae cultivated under various culture conditions. Bioprocess Biosyst. Eng., 37, 2, 99–106, 2014. 33. Li, T., Zheng, Y., Yu, L., Chen, S., Mixotrophic cultivation of a Chlorella sorokiniana strain for enhanced biomass and lipid production. Biomass Bioenergy, 66, 204–213, 2014. 34. Wang, H., Fu, R., Pei, G., A study on lipid production of the mixotrophic microalgae Phaeodactylum tricornutum on various carbon sources. Afr. J. Microbiol. Res., 6, 5, 1041–1047, 2012. 35. Orosa, M., Franqueira, D., Cid, A., Abalde, J., Carotenoid accumulation in Haematococcus pluvialis in mixotrophic growth. Biotechnol. Lett., 23, 5, 373– 378, 2001. 36. Cordero, B.F., Obraztsova, I., Couso, I., Leon, R., Vargas, M.A., Rodriguez, H., Enhancement of lutein production in Chlorella sorokiniana (Chorophyta) by improvement of culture conditions and random mutagenesis. Mar. Drugs, 9, 1607–1624, 2011. 37. El-sheekh, M.M., Bedaiwy, M.Y., Osman, M.E., Ismail, M.M., Mixotrophic and heterotrophic growth of some microalgae using extract of fungal-treated wheat bran. Int. J. of Recycl. Org. Waste Agric., 1, 1, 1–9, 2012. 38. Jönsson, L.J., Palmqvist, E., Nilvebrant, N.O., Hahn-Hägerdal, B., Detoxification of wood hydrolysates with laccase and peroxidase from the white-rot fungus Trametes versicolor. Appl. Microbiol. Biotechnol., 49, 6, 691–697, 1998. 39. Mussatto, S.I., Dragone, G., Roberto, I.C., Ferulic and p-coumaric acids extraction by alkaline hydrolysis of brewer’s spent grain. Ind. Crops Prod., 25, 2, 231–237, 2007. 40. Taherzadeh, M.J. and Karimi, K., Acid-based hydrolysis processes for ethanol from lignocellulosic materials: A review. BioResources, 2, 3, 472–499, 2007. 41. Taherzadeh, M.J., Niklasson, C., Lidén, G., Conversion of dilute-acid hydrolyzates of spruce and birch to ethanol by fed-batch fermentation. Bioresour. Technol., 69, 1, 59–66, 1999.

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42. Bajguz, A., Czerpak, R., Piotrowska, A., Polecka, M., Effect of isomers of hydroxybenzoic acid on the growth and metabolism of Chlorella vulgaris Beijerinck (Chlorophyceae). Acta Societatis Botanicorum Poloniae, 70, 4, 253–259, 2001. 43. Larson, L.J., Effect of phenolic acids on growth of Chlorella pyrenoidosa. Hydrobiologia, 183, 3, 217–222, 1989. 44. Kamaya, Y., Tsuboi, S., Takada, T., Suzuki, K., Growth stimulation and inhibition effects of 4-hydroxybenzoic acid and some related compounds on the freshwater green alga Pseudokirchneriella subcapitata. Arch. Environ. Contam. Toxicol., 51, 4, 537–541, 2006. 45. Liang, Y., Zhao, X., Chi., Z., Rover, M., Johnston, P., Brown, R., Jarboe, L., Wen, Z., Utilization of acetic acid-rich pyrolytic bio-oil by microalga Chlamydomonas reinhardtii: Reducing bio-oil toxicity and enhancing algal toxicity tolerance. Bioresour. Technol., 133, 500–506, 2013. 46. Kröger, M. and Müller-Langer, F., Review on possible algal-biofuel production processes. Biofuels, 3, 3, 333–349, 2012. 47. Lestari, S., Mäki-Arvela, P., Beltramini, J., Lu, G.Q., Murzin, D.Y., Transforming triglycerides and fatty acids into biofuels. ChemSusChem, 2, 12, 1109–1119, 2009. 48. Zamalloa, C., Vulsteke, E., Albrecht, J., Verstraete, W., The techno-economic potential of renewable energy through the anaerobic digestion of microalgae. Bioresour. Technol., 102, 2, 1149–1158, 2011. 49. Markou, G., Angelidaki, I., Georgakakis, D., Carbohydrate-enriched cyanobacterial biomass as feedstock for bio-methane production through anaerobic digestion. Fuel, 111, 872–879, 2013. 50. Matsumoto, M., Yokouchi, H., Suzuki, N., Ohata, H., Matsunaga, T., Saccharification of marine microalgae using marine bacteria for ethanol production. Appl. Biochem. Biotechnol., 105, 247–254, 2003. https://doi.org/10.1385/ ABAB:105:1-3:247. 51. Ho, S.H., Kondo, A., Hasunuma, T., Chang, J.S., Engineering strategies for improving the CO2 fixation and carbohydrate productivity of Scenedesmus obliquus CNW-N used for bioethanol fermentation. Bioresour. Technol., 143, 163–171, 2013. 52. Williams, D., Algenol biofuels announces plan to build and operate a pilotscale algae-based integrated biorefinery. J. Can. Pet. Technol., 48, 8, 6–8, 2009. 53. Lü, J., Sheahan, C., Fu, P., Metabolic engineering of algae for fourth generation biofuels production. Energy Environ. Sci., 4, 7, 2451–2466, 2011. 54. Chisti, Y., Biodiesel from microalgae. Biotechnol. Adv., 25, 3, 294–306, 2007. 55. Brennan, L. and Owende, P., Biofuels from microalgae-a review of technologies for production, processing, and extractions of biofuels and co-products. Renew. Sustain. Energy Rev., 14, 2, 557–577, 2010. 56. Oswald, W.J., Microalgae and wastewater treatment, in: Microalgal Biotechnology, A. Borowitzka and L.J. Borowitzka (Eds.), pp. 305–328, Cambridge University Press, Cambridge, 1992.

Waste Biomass Utilization for Liquid Fuels 87 57. Tredici, M.R., Mass production of microalgae: Photobioreactors, in: Handbook of Microalgal Culture: Biotechnology and Applied Phycology, vol. 1, pp. 178–214, 2004. 58. Chen, F. and Johns, M.R., Relationship between substrate inhibition and maintenance energy of Chlamydomonas reinhardtii in heterotrophic culture. J. Appl. Phycol., 8, 1, 15–19, 1996. 59. Liu, J., Chen, F., Huang, J., Microalgae as feedstocks for biodiesel production, INTECH Open Access Publisher, London, UK., 2011. 60. Mata, T.M., Martins, A.A., Caetano, N.S., Microalgae for biodiesel production and other applications: A review. Renew. Sustain. Energy Rev., 14, 1, 217–232, 2010. 61. Lardon, L., Helias, A., Sialve, B., Steyer, J.P., Bernard, O., Life-cycle assessment of biodiesel production from microalgae. Environ. Sci. Technol., 43, 17, 6475–6481, 2009. 62. Sevda, S., Bhattacharya, S., Abu Reesh, I.M., Bhuvanesh, S., Sreekrishnan, T.R., Challenges in the design and operation of an efficient photobioreactor for microalgae cultivation and hydrogen production, in: Biohydrogen Production: Sustainability of Current Technology and Future Perspective, pp. 147–162, 2017. 63. Ghosh, P.K., Mishra, S.C.P., Gandhi, M.R., Upadhyay, S.C., Mishra, S.K., Pancha, I., Shrivastav, A.V., Jain, D., Shethia, B., Maiti, S. and Zala, K.S., Integrated process for the production of oil bearing Chlorella variabilis for lipid extraction utilizing by-products of Jatropha methyl ester (JME) production. U.S. Patent 8, 741, 628, 2014.

4 Biofuel Production from Lignocellulosic Feedstock via Thermochemical Routes Long T. Duong, Phuet Prasertcharoensuk and Anh N. Phan* School of Engineering, Newcastle University, Newcastle Upon Tyne, UK

Abstract Increasing concerns over greenhouse gas emissions associated with fossil fuel usage that have negative impacts on climate change, insecurity in energy, depletion of natural resources have led to enormous attention towards renewable and sustainable resources. Lignocellulosic materials (waste wood, forest/agricultural residues, and municipal solid waste) have been considered as potential feedstocks to produce biofuels that can replace petroleum-based products in transport sector, which currently relies mainly on fossil fuel (up to 90%) and contribute up to 25-30% emissions. Biofuels derived from lignocellulosic materials can be produced via chemical, biological or thermochemical routes. In this chapter, biofuel production via thermochemical routes such as pyrolysis and gasification are explored. Pre-treatment technologies and operating parameters affecting the processes and product properties are discussed. Due to chemical and thermal instability, high acidity and water content of liquid derived from pyrolysis, pyrolysis liquid cannot be directly used as transportation fuel. Therefore, upgrading of pyrolysis liquid is also focused. Keywords: Pyrolysis, bio-oil upgrading, hydrodeoxygenation, catalytic cracking, hydropyrolysis, cold plasma, gasification, Fischer-Tropsch

4.1

Introduction

Global warming due to air pollutions, i.e. CO2 emission associated with the use of fossil fuels in industrial and transportation sectors has led the rise of sea level, consequently the disappearance of coastal areas [1].

*Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (89–166) © 2019 Scrivener Publishing LLC

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Thus, alternative green and sustainable fuels must be sought to replace petroleum-based sources. Waste/residue biomass including agricultural and forest, municipal solid/industrial waste is the promising feedstock to produce biofuel to reduce fuel production costs, eliminate competition between land set for food and that for fuel/energy and contribute to waste management strategy. Many countries have set their ambitious goal of using biomass/waste for partially replacing fossil fuels e.g. increasing the biofuel consumption to 36 billion gallons by 2022 from 9 billion gallons in 2008 in the USA [2] and the European Union setting 27% of its final energy from renewable sources by 2030 [3]. Lignocellulosic biomass containing cellulose (25-50 wt%), hemicellulose (15-40 wt%), lignin (10-40 wt%), extractives (0-15 wt%) and a small amount of inorganic mineral matter [4] can be used as feedstock to produce biofuels via biological/chemical, combination (biochemical) or thermochemical approaches. Biochemical processes convert cellulose and hemicellulose in the feedstock to sugars that can be then fermented either into alcohols (bioethanol or biobutanol) [5] or into hydrocarbons i.e. alkanes/alkenes [6] in the presence of enzymes or other microorganisms. Furfural and its derivatives i.e. hydroxymethylfufural (HMF) are also produced from sugars via chemical processes that are complex, involved a number of steps and required long reaction time [7–10]. Either biological/chemical or combination process cannot convert all major components in biomass. In contrast, advanced thermochemical processes can utilise all three major components in lignocellulosic materials via a short reaction time [5, 8]. However, they are not suitable for feedstock with moisture content more than 20% due to high energy required for drying process [11]. Advanced thermochemical processes such as pyrolysis and gasification are the common technologies that can be used to produce intermediate products, which can be further processed to generate fuels for transportation sector or internal combustion engines. When lignocellulosic biomass decomposes under elevated temperatures, i.e. 400-650 °C, in an inert environment (pyrolysis), it can produce a mixture of up to 300 compounds depending on conditions (Table  4.1). These compounds can be either rapidly quenched to obtain high liquid yield or further decomposed and/or partially oxidised (gasification process) to form hydrogen or synthetic gas that can be processed to produce liquid fuel via Fischer-Tropsch (FT) synthesis or methanol. In pyrolysis, there are three fractions: gas, liquid (bio-oil) and solid (biochar) formed with their proportion and properties strongly depending upon the nature of feedstock, particle size, heating rate, operating temperature and reactor configuration [14–16]. For instance, pyrolysis of biomass containing a high proportion of cellulose or hemicelluloses produces

Biofuel from Lignocellulosic Feedstock 91 Table 4.1 Typical products derived from fast pyrolysis of biomass constituents [12, 13]. Composition

Products

Hemicellulose

Acetic acid, furfural, furan.

Cellulose

Levoglucosan, 5-hydroxymethylfurfural, hydroxyacetaldehyde, acetol, formaldehyde.

Lignin

Small amount of phenol monomer (including phenol, cresol, guaiacol, syringol), large amount of oligomer with the range molecular weight from hundreds to thousands of Daltons.

Extractives

Fatty acid, rosin acid.

higher liquid yield than that containing lignin. Depending upon the heating rates, pyrolysis can be categorised into slow pyrolysis (heating rates < 100 °C/s) and fast pyrolysis (heating rates 100-1000 °C/s) of which the latter is applied for liquid production (up to 75-80 wt%). Liquid derived from pyrolysis cannot be used directly as transportation fuel due to its chemical and thermal instability, high water (15-35 wt%) and oxygen content (30-40 wt%), which need to be upgraded. Upgrading process remains challenges due to the complexity of the pyrolysis products [17], rapid catalyst deactivation and high temperature & pressure requirements [18]. Gasification is a partial oxidation process to convert lignocellulosic material into a mixture of mainly H2 and CO (known as synthetic gas or syngas) together with small amounts of CH4, CO2, N2, char, ash, tar, oils etc. at a temperature range of 900-1500 °C [11]. The proportion of components in the syngas product is strongly influenced by the types of a gasifier and its operating conditions i.e. gasifying agent, equivalence ratio of gasifying agent to feedstock and properties of feedstock [19–21]. Typically, the gas derived from biomass gasification contains approximately 28-40 vol% CO, 22-32 vol% H2, 10-16 vol% CO2, 3-11 vol% CH4, 0.15-0.24 g/ Nm3 tar (high molecular weight compounds that are condensed into sticky substances when the gas is cooled down below 50 oC) and other impurities [22–24]. With tar formation and low ratio of H2/CO (typically less than one for biomass gasification) [25], the syngas needs to be cleaned before using in gas turbines and internal combustion engines for heat and electricity generation [26, 27] and/or upgraded via steam reforming to obtain a high

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molar ratio of H2/CO, i.e. above 2 for chemical synthesis (i.e. methanol) and liquid hydrocarbons via FT synthesis [28, 29]. Pyrolysis plays an important role in biomass gasification because it affects the quality of the syngas due to high volatile matter (75-85 wt%) in biomass compared to that in coal (20-30 wt%). The volatiles and a small fraction of bio-char from the pyrolysis step react with gasifying agents (O2, H2O, CO2, air or their mixture) at a temperature range of 900-1500 °C [30, 31]. The uniform mixing between volatiles and gasifying agents and high temperature are important to minimise concentration of tar in the gas products. The remaining bio-char and gas from pyrolysis and oxidation stages are further converted to H2, CO, small amount of CH4 and CO2 via heterogeneous and homogeneous reactions [11, 19, 32]. This chapter is to comprehend the main aspects of advanced thermal processes including fast pyrolysis/bio-oil upgrading and gasification/FT for biofuel production. Apart from conventional upgrading methods of pyrolysis liquid, this chapter also focuses new advanced approaches such as oxygen removal in bio-oil by cold plasma and fast hydropyrolysis.

4.2 Fast Pyrolysis 4.2.1 Principles Fast pyrolysis is carried out at a temperature of around 500 °C with high heating rates (100-1000 °C/s). In order to obtain high heating rates/uniform heating, biomass needs to be finely ground, i.e. less than 3 mm [33, 34] and dried to a moisture content below 10%. As a result, biomass decomposes rapidly into vapours, aerosols, and a small amount of char. The design should allow not only to have a short residence time of volatiles (typically < 2 s) but also to rapidly remove the char in order to minimise secondary (cracking) reactions in order to maximise the liquid yield. After the volatiles are rapidly quenched, a dark brown liquid is formed with its heating value (17-19 MJ/kg) of about half that of petroleum-based fuel oil.

4.2.2 Reactors Although fast pyrolysis was firstly investigated in 1875, a significant progress in developing bio-oil production has only been dated back from the 1980s [35]. To date, a wide range of reactors such as bubbling and circulating fluidized beds, ablative, entrainment, rotating cone, auger and vacuum have been studied. The key features of these types of fast pyrolysis reactor are summarized in Table 4.2. Among these types, the fluidized beds have

Status

Commercial

Commercial

Commercial

Laboratory

Laboratory

Pilot

None

Property

Fluid bed

Circulating fluid bed

Rotating cone

Entrained flow

Ablative

Screw/Auger

Vacuun

60 wt%

60 wt%

75 wt%

60 wt%

70 wt%

75 wt%

75 wt%

Bio-oil yield on dry biomass

High

Medium

High

Medium

High

High

Medium

Complexity

Large

Medium

Large

Small

Small

Small

Small

Feed size

Low

Low

Low

High

Low

High

High

Inert gas requirement

Large

Small

Small

Medium

Small

Medium

Medium

Specific reactor size

Table 4.2 Overview of fast pyrolysis reactor characteristics for bio-oil production (adapted from [39]).

Difficult

Medium

Difficult

Easy

Medium

Easy

Easy

Scale-up

Medium

High

High

Low

High

Low

Low

Gas quality

Biofuel from Lignocellulosic Feedstock 93

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Liquid Biofuel Production

received the most attention for fast pyrolysis due to their excellent heat and mass transfer characteristics, simplicity of operation, and relative ease of scale-up [35]. The updated fast pyrolysis plants and projects for bio-oil production are also illustrated in Table 4.3.

4.2.2.1 Bubbling Fluid Bed A bubbling fluid bed (BFB) with approximately 1 mm diameter quartz sand bed material is the most common reactor type for liquid production from pyrolysis [36, 37]. Volatiles are quickly separated from the char in a cyclone and quenched in a condenser to collect liquid (Figure 4.1). An electrostatic precipitator is typically used to collect aerosols, which are incompletely depolymerised lignin fragments existing as a liquid with high molecular weight. The BFB pyrolyser provides high liquid yield of typically 70-75 wt% (based on dry wood feedstock). Small biomass particle sizes of less than 2-3 mm are needed in order to achieve high heat transfer between hot sand bed material and biomass [38]. Hot vapour residence time, about ~ 1 s, is controlled by the fluidising gas flow rate. As char acts as a cracking catalyst at high temperatures, rapid and effective char separation is crucial, which is achieved via cyclones. The main disadvantage of the BFB is the high flow rate of inert gas for fluidisation. To enhance energy efficiency, a fraction of non-condensable gas from pyrolysis is compressed and recycled for the process. Therefore, gas cleaning steps are required to avoid blockage of heat exchangers, and blowers [40].

4.2.2.2 Circulating Fluid Bed A circulating fluid bed (CFB) reactor has many of the features of the BFB described above. However, the residence time of the fluidizing bed material (i.e. sand) is similar to that of char and vapours. Due to high gas velocity, char particles are more attrited, leading to high char content in the collected bio-oil; therefore, the process requires more extensive char removal [41]. As shown in Figure 4.2, biomass, which is continuously conveyed into the reactor, contacts with hot inert sand particles and then the decomposition takes place. The char and sand are carried out of the reactor and sent to a combustor chamber where the char particles are combusted. The main advantage of the CFB compared to the BFB is the direct heat supply to the biomass by recirculation of sand from the combustion chamber. Similarly to the BFB, the problem in the CFB operation is high flow rates of inert gas.

Location

Circulating fluidized bed Circulating fluidized bed

Aracruz, Espirito Santo, Brazil

Vienna, Georgia

Port-Cartier, Quebec, Canada

Joensuu, Finland

Ensyn – Fibria

Ensyn – Renova Capital Partners

Ensyn – Arbec Forest Products – Groupe Rémabec

Fortum – VALMET

Fluidized bed

Circulating fluidized bed

USA

Circulating fluidized bed, catalytic

Technology

KiOR/Inaeris Technologies

Feeding Capacity of >1000 kg/h

Host organization

10000

8333

16667

18333

20 833

Capacity (kg fry feed/h)

Fuel

Fuel

Fuel

Fuel

Catalytic biooil for HDO

Application

Table 4.3 Fast pyrolysis bio-oil production processes updated to 2017 (> 100 kg/h) [49–52].

2013

2018

–

–

2014

Operation year

(Continued)

Operational

Commissioning

In design phase

In design phase

Dormant

Status

Biofuel from Lignocellulosic Feedstock 95

Location

Hengelo, The Netherlands

Renfrew, Ontario, Canada

Malaysia

Canada

Rhinelander, Wisconsin, USA

Rhinelander, Wisconsin, USA

Host organization

BTG BioLiquids – EMPYRO

Ensyn

BTG

ABRI Tech.

Red Arrow/ Ensyn – Kerry

Red Arrow/ Ensyn

Circulating fluidized bed

Circulating Fluidized bed

Auger

Rotating cone

Circulating fluidized bed

Rotating cone

Technology

Separation of chemicals and fuel

Separation of chemicals and fuel

1667

1250

Fuel

Fuel

Fuel

Fuel

Application

2000

2000

2500

5000

Capacity (kg fry feed/h)

2002

2014

2012

2005

2006

2015

Operation year

Table 4.3 Fast pyrolysis bio-oil production processes updated to 2017 (> 100 kg/h) [49–52]. (Continued)

(Continued)

Operational

Operational

Dormant

Dormant

Operational

Operational

Status

96 Liquid Biofuel Production

Location

Auger

Fluidized bed (mobile)

Germany

Canada

USA

Tempere, Finland

Karlsruhe Institute of Technology (KIT)

Agri-Therm/ University of Western Ontario

KiOR/Inaeris Technologies

Valmet

Fluidized bed

Circulating fluidized bed, catalytic

Circulating fluidized bed

Canada

Technology

Ensyn

Feeding Capacity = 100-1000 kg/h

Host organization

300

417

420

500

625

Capacity (kg fry feed/h)

Fuel

Catalytic biooil for HDO

Chemical feedstock

Feedstock for gasification

Fuel and chemicals

Application

2008

2008

2009

2009

1995

Operation year

Table 4.3 Fast pyrolysis bio-oil production processes updated to 2017 (> 100 kg/h) [49–52]. (Continued)

(Continued)

Operational

Dormant

Operational

Operational

Operational

Status

Biofuel from Lignocellulosic Feedstock 97

Location

UK

Germany

USA

The Netherlands

China

Germany

Host organization

Biomass Engineering Ltd.

Pytec

Virginia Tech

BTG

University of Science Technology of China, Hefei

Fraunhofer UMSICHT

Ablative

Fluidized bed

Rotating cone

Fluidized bed

Ablative

Fluidized bed

Technology

100

120

200

250

250

250

Capacity (kg fry feed/h)

Fuel and chemicals

Fuel

Fuel and chemicals

Fuel

Fuel

Fuel and chemicals

Application

2015

–

–

–

2009

2012

Operation year

Table 4.3 Fast pyrolysis bio-oil production processes updated to 2017 (> 100 kg/h) [49–52]. (Continued)

Operational

Operational

Operational

Dormant

Dormant

Dormant

Status

98 Liquid Biofuel Production

Biofuel from Lignocellulosic Feedstock 99 Gas For recycle, export or flare

Cyclone(s) Pyrolyser Electrostatic precipitator

Quench cooler

Biomass

Char For heat or export

Bio-oil

Gas recycle

Recycle gas heater and/or oxidiser

Figure 4.1 Bubbling fluid bed reactor with electrostatic precipitator [39].

Cyclone(s) Pyrolyser

Flue gas

Gas for recycle, export or flare Quench cooler

Biomass Sand & char

Electrostatic precipitator

Hot sand

Air Combustor Ash Recycle gas heater and/or oxidiser

Figure 4.2 Circulating fluid bed reactor [39].

Bio-oil Gas recycle

100

Liquid Biofuel Production

4.2.2.3 Rotating Cone A rotating cone reactor is a relatively recent development [40]. The key features of the rotating cone reactor are that the hot sand and biomass are mixed at the base of the rotating cone and the centrifugation drives the mixture up the inner wall of the rotating heated cone. There is no auxiliary fluidizing gas in this reactor compared to the BFB and CFB. The volatiles withdrawn from reactor by a small amount of carrier gas are collected and then processed conventionally (i.e. a combination of a condenser and a demister). Sand and char are dropped into a fluidized bed surrounding the cone and then they are transported to a combustor for burning char. The hot sand is recirculated to the pyrolyser. The process and the rotating cone reactor are described in Figure 4.3. The liquid yield of up to 70 wt% can be obtained [42]. The main advantage of a rotating cone reactor is the absence of carrier gas and there is no requirement of gas recycling. However, the process operation is more complex because an integrated control of three subsystems is required: a rotating cone pyrolyser, a riser for sand recycling, and a bubbling bed char combustor.

4.2.2.4 Ablative Pyrolysis In ablative pyrolysis as shown in Figure 4.4, the hot reactor wall (approximately 600 °C) is in contact with “melt” biomass under pressure created by centrifugal forces. As biomass/partially charred particles are moved away, the molten layer vaporises to products that are similar to those derived Char combustor Pyrolysis gases and vapours Sawdust feed Sand

Flue gas

Biomass Ash

Hot sand

Condenser

Gas

Vapours

Rotating cone Reactor

Sand & char

Air

Figure 4.3 Rotating cone pyrolysis reactor and integrated process [33].

Bio-oil storage

Biofuel from Lignocellulosic Feedstock 101 Biomass in Heat

Vapours out Heat

Char out Heat

Heat

Figure 4.4 Ablative fast pyrolysis reactor [39].

from fluidized beds. The rate of the decomposition is strongly influenced by the pressure of the wood onto the heated surface, the relative velocity of the wood, the heat exchange surface and the reactor surface temperature. The main advantage of this technology is that large particles (up to 20 mm [43]) can be used as reaction rates are not limited by heat transfer through the biomass particles. Furthermore, there is no requirement for carrier gas, therefore the equipment size is smaller than BFB or CFB and the reaction system is more compact. In addition, the absence of the fluidising gas substantially increases the partial pressure of the condensable vapours, leading to more efficient condensation and smaller equipment. However, there are two major limitations in ablative pyrolysis: (i) limited heat transfer from hot gases to the ablative surface and (ii) difficulties in contacting feedstock owning diverse morphologies (particle shape, structure, and density) with the ablative surface [44]. Therefore, scale-up of ablative pyrolysis is difficult.

4.2.2.5 Screw Reactor A screw pyrolyser uses a hot heat carrier loop with a mechanically fluidized bed without an auxiliary fluidizing gas (Figure 4.5). Heating can be obtained via different types of carrier such as hot sand, steel or ceramic balls, or external sources [45]. The hot vapour residence times can be in the range of 5-30 s depending on the design and size of reactor [33, 46]. The main advantage of the screw reactors is that it can easily process materials that are difficult to handle or highly heterogeneous. Moreover, the size of the bio-oil condensation system is smaller than the fluidized beds due to the lack of fluidizing gas. However, the liquid yield (about 60 wt%) is lower than that obtained in the fluidized beds and the liquid

102

Liquid Biofuel Production Biomass

Fast pyrolysis

Pyrolysis Gas Char

Condensate(s)

Sand Heat carrier Biosyncrude

Figure 4.5 Screw pyrolyser [47].

is often phase separated due to containing high water content caused by cracking processes under long residence time of volatiles and their contact time with char particles.

4.2.2.6 Other Reaction Systems 4.2.2.6.1 Entrained Flow In principle, entrained flow fast pyrolysis is a simple technology that biomass particles (1-5 mm) are fed to a down-flow heated reactor. Unlike many other fast pyrolysis reactors described above, no extra hot solid material is used to transport and heat the biomass particles. The development of this technology has not been successfully due to poor heat transfer between hot gas and biomass particles. To obtain efficient heat transfer, high gas flow rate is required, consequently the plant size is larger than BFB or CFB and it is difficult to collect liquid from the low vapour partial pressure. The liquid yield is around 40-55 wt% [40, 41], which is much lower than that obtained in the fluidized beds (70-75 wt%).

4.2.2.6.2 Vacuum Pyrolysis Vacuum pyrolysis developed by the University of Laval and Pyrovac, Canada, is a combination of slow and fast pyrolysis conditions [48]. The heat transfer rate to and through the solid biomass is much slower and the vapour residence time is much shorter than those in the aforementioned reactors. Large particles size solids are slowly heated to a temperature of 450 °C while the gas is removed from the hot zone quickly under a pressure of 20-100 kPa [33, 40]. The liquid yield is around 35-50 wt%. Operating

Biofuel from Lignocellulosic Feedstock 103 under vacuum pressure makes the process complex and costly because of the use of very large vessels and piping systems. The advantages of the process are that (i) it can process large particles compared to other fast pyrolysis reactors and (ii) there is less char in the liquid because no carrier gas is required leading to no further separation process.

4.2.3 Bio-Oil Composition and Properties Pyrolysis liquid, known as bio-oil, is a dark red-brown to almost black liquid depending upon its chemical composition and the presence of micro-carbon particles. Bio-oil is the mixture of hundreds of various oxygenated compounds of which most of them have low concentrations (below 1 wt%). These compounds account for about 45 wt% and are typically categorised into acids, alcohols, aldehydes, ketons, esters, sugars, furans, phenols, guaiacols, syringols and miscellaneous oxygenates (Misc Oxy) as shown in Figure 4.6. As presented in Table 4.4, bio-oil has high water content, typically 15-35 wt%. Water cannot be removed via physical methods such as centrifugation and distillation because it exists in a micro-emulsion and bio-oil is chemical and thermal instability [54, 55]. As soon as the bio-oil is heated up above 50 °C, it is solidified. The high water content decreases the heating value of biooil, thereby reducing the local combustion temperature and the combustion 40

Hemicellulose and cellulose

Low wt% High wt%

35 Acids: Formic Acetic Propanoic

30

Misc Oxy: Glycolaldehyde Acetol

wt (%)

25 Alcohols: Methanol Ethanol Ethylene Glycol

20 15

Esters: Methyl formate Butyrolactone Angelicalactone

10

Aldehydes: Formaldehyde Acetaldheyde Ethanedial

Phenols: Phenol DiOH-benzene Dimeth-phenol

Sugars: Anhydroglucose Cellobiose Furans: Fructose Furfurol Glucose HMF

Lignin

Guaiacols: Isoeugenol Eugenol Methyl guaiacol

Furfural Ketones: Acetone

5

s Ph en ol s Gu ai ac ol s Sy rin go ls

an Fu r

s Al co ho ls Ke to ne s Al de hy de s M isc Ox y Su ga rs

te r Es

Ac id

s

0

Figure 4.6 Typical chemical composition in bio-oil. Reprinted with permission from American Chemical Society, 2006, ref. [53].

104

Liquid Biofuel Production

Table 4.4 Typical properties of bio-oil in comparison with burner fuels.

Property

Unit

Bio-oil [50]

Residue-containing burner fuels (class E-H according to British Standard BS2869:2017) [60, 61]

Higher heating value, HHV

MJ/kg

14-19

42.7-44.0

Water

wt%

20-30

0.5-1.0

2-3

0 mg KOH/g

pH Kinematic viscosity

mm2/s

14-40 @40 °C

8.2-56.0 @100 °C

Density at 15°C

kg/dm3

1.11-1.30

0.93-0.99

Pour point

°C

-36 ÷ -9

–

Flash point

°C

40-110

66

Solids

wt%

40%) such as food waste and algae [75]. It operates at a temperature range of 200-260 °C and autogenerous pressure up to 5000  kPa. The purpose of HTC is to stabilise material and increase carbon and energy contents [75]. Three products are generated during HTC including gas, aqueous phase, and solid fuel [76, 77]. Around 55-90 wt% solid can be obtained from HTC, containing 80-95% the energy content of the original biomass [78]. However, commercial-scaled HTC is challenging due to pressurized operating conditions. Chemical pre-treatment such as washing/leaching is used to remove inorganic compounds such as alkali metals before pyrolysis to improve the yield and quality of bio-oil. Alkali metals in the feedstock act as catalysts for cracking processes that lead to increase water content and

Biofuel from Lignocellulosic Feedstock 107 Table 4.5 Properties and benefits associated with torrefaction of biomass [69, 73, 74]. Improved properties

Benefits

Size reduction and feeding characteristics (i.e. homogeneity, grindability)

- Lower energy required for size reduction - Lower operating cost of milling equipment - Increase homogeneity

High energy content

- High efficient thermo-chemical conversion processes (i.e. combustion) - Low logistics cost (storage/ transportation)

Low moisture content and hydrophobicity

- Low mass loss compared to biomass - Low risk of biological degradation - Better blending for various biomass

Reduced emissions during storage

- Low emission of CO, CO2 and CH4 off-gases

decrease the bio-oil yield. Water is used to wash alkali sulphates, carbonates, and chlorides [79–81] whereas hydrochloric acid leaches carbonates and sulphates of alkaline earth and other metals [82]. Organic compounds of Mg, Ca, K and Na can be leached by ammonia or water [83, 84]. However, washing/leaching could lead to the loss of hemicelluloses and cellulose through hydrolysis [33] and create large amount of waste water that need to be treated. Compared to physical and chemical pre-treatments, biological methods are much slower but less energy-consumed and greater environmental friendly [4]. Pyrolysis performance was improved with fungal pre-treated lignocellulosic feedstock in terms of selective decomposition of lignin during pyrolysis [4], reducing pyrolysis temperature and the emission of gas contaminant (i.e. SOx) [85], increasing yield of aromatic products in the presence ZSM-5 [86]. It was also reported [87] that enzymes hydrolysis of lignin can enhance the production of phenols and aromatic hydrocarbons. The choice of a pre-treatment method prior to fast pyrolysis requires a consideration of several factors such as costs and technologies available for grinding, drying, transporting, storing, handling, and upgrading. Table 4.6 summarizes properties and product yields when pre-treating biomass before pyrolysis.

T (°C)

500

450

425-550

Reactor type

FBR

Auger

FBR Water 24 h Water 20 min Water + agitate, 1 min

Palm fruit bunches

Palm fruit bunches

Steam explosion

Loblolly pine

Palm fruit bunches

0.5% NaOH

Loblolly pine

–

0.5% H2SO4

Loblolly pine

Palm fruit bunches

–

3.68

2.14

1.03

5.36

0.64

2.49

0.71

0.39

0.1% H2SO4

Hemlock

Loblolly pine

–

Hemlock

51

58

72.4

49.8

44

49

63

54

72.4

56.5

75.9

0.1% H2SO4

Poplar

Bio-oil yield (wt%) 65.8

Feed ash (wt%)

–

Pre-treatment

Poplar

Feedstock

Table 4.6 Properties of bio-oils from various raw and pre-treated biomass feedstocks (adapted from [69]).

16

13

11.0

15.1

29.3

55.6

29.5

20.8

 

 

 

 

(Continued)

Water in oil (wt%)

108 Liquid Biofuel Production

450

600

500

480 

Auger 

Pyroprobe

FBR

FBR

– Drying @180 °C Torrefaction @230 °C Torrefaction @270 °C

Clean pine

Clean pine

Clean pine

Clean pine

HTC @190 °C

Eucalyptus

–

Lignin residue –

Hot water + hydrolysis

Maple wood

Eucalyptus

–

2% H2SO4

Corn stalks

Maple wood

–

Pre-treatment

Corn stalks

Feedstock

Note: FBR – fluidized bed reactor.

T (°C)

Reactor type

0.59

0.41

0.49

0.41

0.95

4.5

Feed ash (wt%)

51

58

58

65

9.8

10

11.0

14.0

18

26

59 68

 

 

 

21

27

Water in oil (wt%)

44.8

66.6

66.7

46

35

Bio-oil yield (wt%)

Table 4.6 Properties of bio-oils from various raw and pre-treated biomass feedstocks (adapted from [69]). (Continued)

Biofuel from Lignocellulosic Feedstock 109

110

Liquid Biofuel Production

4.2.4.3 Temperature and Carrier Gas Flow Rate Pyrolysis temperature plays an important role in the process, affecting product yields and properties. Increasing pyrolysis temperature increases the rate of decomposition. A temperature range of 450-550 °C with a short residence time (typically < 2 s) is commonly used to minimise secondary cracking. Pyrolysis temperature affects the degree of the decomposition of biomass to volatiles (primary cracking), and volatiles & char to smaller molecule weight compounds (secondary cracking) [88–90]. The secondary cracking is dominant at temperatures above 550 °C, generating high gas fraction [16]. At temperature below 400 °C, the decomposition process of biomass is incompleted, leading to a low yield of bio-oil. However, there is no clear correlation between pyrolysis temperature and properties of biooil [66, 91]. The carrier gas flow rate, i.e. nitrogen flow rate, is one of the most important parameters in fast pyrolysis as it determines hot vapour residence time and residence of time of biomass in the reactor (for fluidized beds) and affects the rate of heat transfer from hot environment into biomass. In principle, a short hot vapour residence time favours for bio-oil production because it minimises further decomposition of desirable products. It was reported that a combination of low pyrolysis temperatures (400-550 °C) and short hot vapour residence time (typically < 2 s) is preferable for high liquid yield production (> 60 wt%) [16, 33]. However, there is a limit of hot vapour and biomass residence time to ensure good heat transfer at biomass particle surface. Residence time for the decomposition of biomass particles must be longer than the vapour residence time to achieve a complete decomposition for a high bio-oil yield [92]. The carrier gas flow rate also influences physical properties of bio-oil. A high flow rate provides bio-oil with low water content but high viscosity, density and heating value [66, 93] because it minimises the cracking processes. Solid content in bio-oil decreases with high carrier gas flow rate because the ability to separate solid particles from vapour stream by a reverse flow cyclone is more effective [66, 94]. However, increasing carrier gas flow rate could decrease partial pressure of low molecular weight compounds in vapour products, therefore reducing the bio-oil yield due to the difficulties in condensation of these components [12, 95]. Carrier gases containing CO, CO2, H2, and light hydrocarbons (reactive gases), affect to properties of bio-oil [96, 97]. It was reported [97] that bio-oils produced under the reactive gas atmospheres, are richer in aromatic hydrocarbons than those produced under N2; however, the yield of organics in bio-oil decreases up to 23%.

Biofuel from Lignocellulosic Feedstock 111

4.3 Bio-Oil Upgrading Although bio-oil derived from pyrolysis process could potentially be used for heat/power generation or fuels for transportation. However, its chemically and thermally unstable properties due to high oxygenated compounds, upgrading is required. A number of methods can be used to improve the bio-oil properties, e.g. physical (hot filter, adding solvents, emulsion and vacuum distillation) and chemical methods (hydrotreating, catalytic cracking, esterification, and gasification). Among these, two main routes have been extensively investigated such as catalytic hydrotreating or hydrodeoxygenation (HDO) and catalytic cracking, evidenced by a large number of references listed in current review papers [33, 98–100].

4.3.1 Hydrodeoxygenation Hydrodeoxygenation (HDO) is a process to remove oxygen via dehydration in the presence of hydrogen and catalyst at high pressures (up to 300 bar) and temperatures (250-450 °C) [101, 102]. The HDO process generates mainly naphtha-like products that can be either used directly as drop-in fuels or further refined in existing refining facilities to produce conventional fuels such as gasoline and/or diesel. Various reactions occur in the HDO process such as cracking, hydrocracking, decarbonylation, decarboxylation, hydrogenation, hydrodeoxygenation, polymerisation and condensation [103–105]. With a typical elemental composition of bio-oil obtained from fast pyrolysis (C: 55-65 wt%, O: 28-40 wt%, and H: 5-7 wt% [98, 106]), a general reaction of the HDO process can be simplified and displayed as in Equation 4.1 [99]:

C1H1.4O0.4 + 0.7H2

1" CH2" + 0.4H2O

(4.1)

where “CH2” represents an unspecified hydrocarbon product. The reaction is exothermic with an overall heat of reaction of 2.4 MJ/ kg [107]. For a complete deoxygenation (based on the stoichiometry of Equation 4.1), the maximum yield of hydrocarbons could be around 56-58 wt% [108]. However, it is difficult to achieve due to many other side reactions occurring simultaneously in the HDO process (i.e. polymerisation, condensation), therefore the final products with the lower O/C ratio (0.11) compared to the untreated bio-oil (0.56) [106] still contain a certain percentage of oxygen.

112

Liquid Biofuel Production

High pressure in the HDO process is required to (i) increase mass transfer due to low solubility of hydrogen and bio-oil, (ii) prevent evaporation due to the presence of water and (iii) avoid coke formation [99, 106, 109]. A large excess of hydrogen is required, i.e. 35-40 mol H2/kg of bio-oil (800 litre H2/kg of bio-oil) [110] or 600-1000 litre H2/kg of bio-oil [111] compared to a theoretical amount of hydrogen of around 25 mol/kg (500 litre H2/kg of bio-oil) for the complete deoxygenation. To achieve a high degree of deoxygenation (> 50%), long reaction time is needed [40]. In a continuous flow reactor, the oxygen content of the upgraded oil decreases around 11% when reducing liquid hour space velocity (LHSV) from 0.70 h-1 to 0.25 h-1 over a Pd/C catalyst at 140 bar and 340 °C [110]. To achieve oxygen content in the liquid below 20 wt%, LHSV must be in a range of 0.1-1.5 h-1 [112] for continuous processes and 3-4 h for batch system [18, 105, 113]. It was reported [114] that deep HDO of pyrolysis oil encounters major challenges with coking formation at high temperatures (> 300 °C). Therefore, a two-stage hydrotreating of bio-oil is implemented to overcome the aforementioned issues for bio-oil upgrading [100, 115]. - Stage 1 – Stabilization: This stage reduces the reactivity of functional groups such as aldehydes, ketones, and double C=C bonds and transforms sugars into more “stable” components [106, 116]. It is commonly performed at temperatures below 250 °C and 34-140 bar of hydrogen in the presence of a catalyst [112, 116]. The degree of deoxygenation (DOD) is about 50% mainly due to the thermal decomposition [100]. - Stage 2 – Deep HDO is carried out at a temperature range of 350-450 °C and high pressure 70-200 bar of hydrogen. HDO operated at high temperatures is feasible to achieve the degree of deoxygenation of typically 95% [106, 117]. Catalysts initially tested for HDO of bio-oil were the traditional hydrodesulfurization (HDS)-based catalysts such as CoMo and NiMo supported on alumina or alumina silicate. Based on traditional HDS processes, HDO has been tested with a wide range of operating conditions, various types of reactors (such as batch and fixed bed reactors) and feedstock. The level of deoxygenation on the HDS-based catalysts is 80-90% depending on the severity of the process. The life span of the catalyst is from several hours [113, 118] to several days [100]. However, catalytic activity decreases over time due to the transformation of catalysts from sulphide to oxide [99]. Hence, H2S or dimethyl sulphide is co-fed to the system to generate

Biofuel from Lignocellulosic Feedstock 113 sulphide catalysts and to stabilise catalysts [99, 119], which leads to an increase in the sulphur content in the product oil [120, 121]. Furthermore, alumina support has low durability in high water environment such as biooil and high acid content of Al2O3 leads to the formation of coke on the catalyst surface [122]. Transition metal catalysts (Ru, Pd, Rh and Pt), common metals (Ni, Fe, Pb, Zn, or mixtures), and oxides of metals (CuO, NiO) supported on ZrO2, SiO2 or activated carbon have been examined for HDO of bio-oil [105, 106, 123–127]. The performance of these catalysts are equivalent to or even better than that of the HDS catalysts [107]. Compared to conventional HDS catalysts, precious metal catalysts (such as Ru, Rh, Pd, Pt) exhibit higher catalytic activity, which can be used at lower temperatures in stage 1. The formation of coke is slow due to the low reaction temperatures and the support is not acidic, therefore prolonging the life span of the catalysts. Although precious metals are potential catalysts for the HDO process due to their catalyst activities and high resistance to fouling, their high price causes a low economic efficiency of HDO process. Therefore, these catalysts are mainly used for the stabilization stage and traditional HDS-based catalysts are used for the deep HDO stage [128, 129]. The effects of various catalysts and reaction conditions can be seen in Van Krevelen plot shown in Figure 4.7 in which there are two distinct areas of the

0.6 Bio-oil (dry)

0.5 HPTT-line

Molar O/C

0.4

Ru/Al2O3

+H2

0.3

Pt/C NiMo/Al2O3

CoMo/Al2O3 Ru/C

HPTT 300

Ru/TiO2

0.2 –H2O

Pd/C

Charcoal

Deep HDO NiMo/Al2O3

0.1

CoMo/Al2O3 Ru/Al2O3

0.0 0.0

Mild HDO

0.5

1.0 Molar H/C

Pt/C

Ru/TiO2 Ru/C Pd/C

1.5

2.0

Figure 4.7 Van Krevelen plot based on the elemental compositions (dry basis) of the mild and deep HDO over various catalysts. Reprinted with permission from American Chemical Society, 2009, ref. [105].

114

Liquid Biofuel Production

mild and deep HDO oils. A high degree of deoxygenation, corresponding to ratios of H/C = 1-1.5 and O/C = 0.03-0.11 can only be achieved with a suitable catalyst under severe conditions, i.e. 350 °C and 200 bar. After HDO, bio-oil properties are significantly improved after HDO in terms of oxygen content, viscosity, and pH. HDO upgraded bio-oil can be directly used for a number of applications including combustion in turbines or engines. However, it requires further upgrading in refining processes such as distillation and hydrocracking to produce transportation fuels [130]. One of the most challenges for the HDO process is a large amount of hydrogen required. Although hydrogen can be produced from biomass or coal via gasification, splitting water and steam reforming of natural gas, it is commercially produced via steam reforming of methane at high temperatures (> 700 °C), which accounts for about 95% of the hydrogen used today in the U.S [131]. Although water electrolysis is recognized as an easy and primary route for the production of pure hydrogen, it is not easy to scale up to industrial plans due to technical and economic problems (i.e. production cost of 1 kg of H2 via this route is 3-6 times higher than that from steam reforming of methane) [132, 133]. Recently, hydrogen production from biomass has gained attention due to the reduction of life cycle greenhouse gas emissions from 26% (via dark fermentation [134]) to nearly 100% (via gasification [131]) compared to steam reforming of methane. It is also reported that the estimated production cost of hydrogen from biomass via gasification is similar the same as that from steam reforming of methane in the industrial scale [132, 133]. Therefore, to increase the sustainability and affordability, hydrogen should be generated from sustainable and renewable sources such as waste biomass/residues.

4.3.2 Catalytic Cracking In catalytic cracking, the oxygen is removed in the forms of carbon dioxide, carbon monoxide, water and short-chain oxygenates via several reactions such as cracking, decarboxylation, decarbonylation, dehydration and water gas shift. The process is carried out in the presence of a catalyst, mainly zeolites, without hydrogen at atmospheric pressure and a temperature range of 300-600 °C depending on a type of reactor [135]. The general reaction can be described in Equation 4.2 [99]:

C1H1.4O0.4

0.9CH1.2 + 0.1CO2 + 0.2H2O

where “CH1.2” is an unspecified hydrocarbon product.

(4.2)

Biofuel from Lignocellulosic Feedstock 115 The catalytic cracking process can be operated in either liquid or vapour phase. It can be integrated or coupled in a pyrolysis process (Figure 4.8) or used as a stand-alone process for upgrading. The integrated catalytic pyrolysis and coupled vapour upgrading have been mostly studied in order to utilise the heat of hot vapour stream [136]. In these cases, catalysts are held in a fixed bed after a pyrolyser or integrated in a fluidized bed pyrolyser. Zeolites are commonly used due to their durability to high temperatures and mechanical forces [33]. However, the catalysts are rapidly deactivated after several hours by coking [135] and required frequent regeneration, normally in a fluid catalytic cracking (FCC) unit. A relatively high residence time, i.e. LHSV around 2 h−1 [53], is required to achieve an acceptable degree of deoxygenation in catalytic cracking process. The catalytic cracking upgraded bio-oil has a higher heating value (21-36 MJ/kg) than original bio-oil (16-19 MJ/kg) and a lower ratio of H/C ratio (0.551.20) compared to HDO upgraded bio-oil [99, 137, 138]. It contains a high proportion of aromatic compounds with the oxygen content decreased to 13-24% [135]. The selection of an appropriate catalyst plays an important role in the formation of high value compounds (phenolics, monoaromatic hydrocarbons) and in the elimination of undesirable compounds (acids, aldehydes, ketones). Zeolites, particularly ZSM-5, effectively convert biomass to above 30 wt% aromatics [140] and selectively deoxygenate pyrolysis vapours thereby decreasing the O/C ratio [141]. The use of zeolite cracking catalysts also results in the formation of small molecules such as methane, ethylene and propylene [142]. It was found that the deoxygenation effectiveness was Catalytically treated vapours

Catalytically treated vapours

Char out

Spent catalyst out

In situ reactor (continuous)

Biomass in (a) in-situ

Ex situ reactor (continuous) Primary pyrolysis vapours

Spent catalyst out

Catalyst in

Pyrolysis reactor (non-catalytic)

Catalyst in

Biomass

(b) ex-situ

Figure 4.8 Integration (a) and coupling (b) catalytic cracking into pyrolysis process [139].

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in the order of Ca-Y > Mg-Y > Na-Y = H-ZSM-5 > H-Y [143], indicating that metal impregnated zeolites have a higher potential to remove oxygen than conventional acidic zeolites. This was also confirmed by Mullen et al. [144], where Ca-Y was found to be more effective than β-zeolite with regards to deoxygenation. Although the quality of bio-oil improves in catalytic cracking, it has not meet the transport fuel quality standards. Therefore, the products need to be further upgraded via the HDO process before integrating with the production processes of gasoline/diesel. This indicates that catalytic pyrolysis should be used as the first step to reduce oxygen content in pyrolysis liquid and the amount of hydrogen required for upgrading bio-oil into hydrocarbons.

4.3.3 Fast Hydropyrolysis Fast hydropyrolysis is a process in which the decomposition of biomass occurs at high heating rates (~ 500 °C/s) in a H2 environment [145]. It can only be performed in a system that allows a short vapour residence times i.e. a few seconds, such as fluidized bed reactors, cyclone reactors, and micropyrolysis systems, i.e. Py–GC/MS (pyroprobe) [146]. H2 can be used alone or diluted in an inert gas such as N2. In hydropyrolysis, hydrogen radicals generated from H2 gas react with biomass-derived volatiles in the presence of a catalyst to remove oxygen in the form of water, CO, and CO2 to produce hydrocarbons. In addition, the hydrogen radicals could stabilise many reactive volatile intermediates, which in turn prevent the polymerization of these intermediates, thereby lowering the possibility of coke formation on the catalyst [147, 148]. One of the advantages of hydropyrolysis is that the operating pressures (around 30 bar) are much lower than that in HDO (up to 300 bar) [149, 150]. Another interesting characteristic of hydropyrolysis is that the process is exothermic, which helps sustain endothermal pyrolysis reactions taking place inside the reactor [149]. As shown in Figure 4.9, four types of fast hydropyrolysis process have been developed with their characteristics summarized in Table 4.7. In general, fast hydropyrolysis produces twice as high hydrocarbon yield as catalytic fast pyrolysis with less problems of catalyst deactivation due to the presence of H2. The primary products are aromatics alongside other products (alkanes and cycloalkanes) in the presence of a secondary upgrading unit (ex-situ upgrading). In some cases, the liquid contains less than 0.5 wt% oxygen [149], and liquid yield up to 70 wt% [145].

Biofuel from Lignocellulosic Feedstock 117 (a)

(c) Oxygenated volatiles

Oxygenated volatiles

Fast hydropyrolysis reactor Biomass Feed

Fast hydropyrolysis reactor

Inert particles (bed) Distributor plate

Biomass feed

Biomass feed

Distributor plate Hydrocarbons

(d) Hydrocarbons

Hydrocarbons

Catalytic fast hydropyrolysis reactor

Inert particles (bed)

H2 feed

H2 feed

(b)

Hydrotreating unit with catalyst

Hydrotreating unit with catalyst Catalytic particles (bed)

Catalytic fast hydropyrolysis reactor

Distributor plate

Biomass feed

Catalytic particles (bed) Distributor plate Hydrocarbons

H2 feed

H2 feed

Figure 4.9 Possible configurations of the fast hydropyrolysis process: (a) Non-catalytic fast hydropyrolysis; (b) Catalytic fast hydropyrolysis; (c) Non-catalytic fast hydropyrolysis with ex-situ hydrotreating; and (d) Catalytic fast hydropyrolysis with ex-situ hydrotreating [146].

4.3.4 Cold Plasma Plasma is defined as a partially ionized gas including ions, atoms, metastable and free radicals that creates a quasi-neutral media [157]. There are two types of plasma: thermal and non-thermal plasmas or cold plasma. The main disadvantage of thermal plasmas is their high energy consumption [158, 159] to create an extremely hot medium of ions, electrons and molecules (to temperatures of 5000-10000 °C). The reactor temperature is very high, therefore cooling of the electrodes is required to reduce their thermal erosion and it is not feasible to incorporate in catalytic processes. For cold plasmas, external energy (up to a few hundred of Watts) is mainly transferred to electrons to raise their temperature (rather than the bulk gas temperature) into the range of 10000-100000 °C (equivalent to 1-10 eV) whilst ions and neutral species (radicals, molecules) remain at near ambient temperature. The role of cold plasmas is not to provide energy to the system, but to generate radicals and excited species, resulting in a low energy consumption.

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Liquid Biofuel Production

Table 4.7 Summary of possible configurations of the fast hydropyrolysis process. Type of process

Characteristics

Observation

Non-catalytic fast hydropyrolysis [151–153]

- Requirements: H2 gas, no catalysts, operating conditions: 500 °C and 0-30 bar of H2 pressure.

- Reduce yields of all oxygenated compounds was observed. - A slight increase in phenolic yields. - Increase low molecular species and yields of permanent gases, reducing the yield of liquid products when operated at high pressures.

Catalytic fast hydropyrolysis [147, 149, 151, 154–156]

- Fast hydropyrolysis is performed in the presence of catalysts but without ex-situ upgrade. - Pyrolysis reactor is a circulating fluidized bed reactor operated at 0-21 bar of H2 pressure and 300-650 °C depending on the type of feedstock. - Various catalysts tested such proprietary catalysts of RTI International, HZSM-5 support with Ni, Co, Mo and Pt; Ni/Al2O3, CoMo/Al2O3, Ni/LY.

- Can achieve oxygen content of 4.2 wt% with a low liquid yield of 8.7 wt%. - Major products in the liquid phase are aromatic hydrocarbons (up to 94%) including benzene derivatives, substituted naphthalenes, and polycyclic aromatic hydrocarbons. - An addition of a supported metal increases the rate of aromatics production.

(Continued)

Cold plasma-catalysis (known as cold plasma-assisted catalysis) refers to the combination of plasma with a material that has heterogeneously catalytic properties. This hybridization can change the mechanism of a chemical process by altering reaction pathways [160]. There are many types of cold plasma such as dielectric barrier discharge (DBD), corona discharge and spark discharge, where DBD is more efficient in generation of radicals in the gas phase [157]

Biofuel from Lignocellulosic Feedstock 119 Table 4.7 Summary of possible configurations of the fast hydropyrolysis process. (Continued) Type of process

Characteristics

Observation

Non-catalytic fast hydropyrolysis with ex-situ hydrotreating [145, 152, 153, 156]

- A hydrotreating unit is placed after pyrolyzer. - Pyrolysis reactor is a cyclone operated at 27-54 bar of H2 pressure, 480-580 °C, and the product vapours upgraded in the catalytic bed is at 300-375 °C. - Catalysts: Al2O3, 2% Ru/Al2O3, 2% Pt/Al2O3, 5 wt% Pt - 2.5 wt% Mo/ MWCNT

- Main products: aromatic compounds, a small amount of alkanes and cycloalkanes - Pt, Pd and Ni is found to be effectively for hydrogenation. Oxygen is removed via dehydration - The secondary hydrotreating unit is required to produce hydrocarbons.

Catalytic fast hydropyrolysis with ex-situ hydrotreating [149, 150]

- Fast hydropyrolysis reactor includes a catalyst, and the produced vapours are upgraded in a secondary packed bed unit. - Pyrolysis reactor: fluidized bed reactors operated at 20-35 bar H2 pressure and 343-469 °C, and the hydrotreating temperature ranges from 343 to 399 °C.

- Major products: hydrocarbons in the gasoline and diesel range with the yield of about 28 wt%. - Typical fuel properties of the product are: 0.3 total acid number (TAN), RON 84 and less than 0.4 wt% oxygen. - Life span of catalysts: 750 h. - However, the effect of having catalysts in both reactors remains unclear.

and simple in design and operation. Cold plasmas have been used in reforming hydrocarbons to produce hydrogen [158, 159] and in ozone regeneration. Recently, several research groups have attempted to apply cold plasma with or without catalysts for oxygen removal in bio-oil or model compounds as summarized in Table 4.8. Taghvaeit et al. [162, 164] applied cold plasma using a DBD containing metal catalysts such as Ni, Co, Mo, Re and Pt with the support of Al2O3 to produce aromatics from model compounds of lignin derived bio-oil

Feedstock

Anisole – Lignin derived model compound

Anisole – Lignin derived model compound

No.

1

2

Without catalysts

Dielectric Barrier Discharge

He

He

Reactor type Carrier gas

Commercial Dielectric Barrier catalysts: Discharge Ni-Mo/Al2O3; Pt/Al2O3; Co-Mo/ Al2O3; Pt-Re/ Al2O3; and Al2O3.

Catalyst

Effects of carrier gas flow rate, liquid anisole feed flow rate, and reactor length on the reactor performance were investigated: - High flow rate reduced anisole conversion. It also decreased selectivity of phenol but increasing selectivity of benzene. - Increasing the feed flow rate led to a decrease in anisole conversion as well as selectivity of benzene and phenol. - Anisole conversion peaked at an outer electrode length.

Effects of applied voltage, activities of different types of catalysts and catalyst deactivation were investigated: - Major compounds in liquid product including benzene, phenol, 2-methylphenol, 4-methylphenol. - Activities: Ni-Mo/Al2O3 > Pt/Al2O3 > Co-Mo/Al2O3 > Pt-Re/Al2O3 > Al2O3 > plasma alone. - Higher voltage, higher conversion

Note

Table 4.8 Studies on upgrading bio-oil with integration of cold plasma.

[162]

[161]

Ref.

(Continued)

2014

2013

Year

120 Liquid Biofuel Production

Feedstock

Anisole – Lignin derived model compound

No.

3

Without catalysts

Catalyst Dielectric Barrier Discharge

Reactor type He, Ar or H2

Carrier gas Effects of different types of carrier gases, voltage and pulse frequency were investigated: - Major compounds in liquid product including benzene, phenol, 2-methylphenol, 4-methylphenol. Small amount of o-xylene, p-xylene, cyclohexane, methylcyclohexane, hexanal, decane, and methanol. - Anisole conversion: He > H2 > Ar - Phenol is abundant product with selectivity of 49.7% (Ar), 44.4% (H2), 56.4% (He). - Selectivity for benzene in H2 plasma is higher than those in Ar and He. - Small amount of cyclohexane, especially in H2 plasma.

Note

Table 4.8 Studies on upgrading bio-oil with integration of cold plasma. (Continued)

[163]

Ref.

(Continued)

2014

Year

Biofuel from Lignocellulosic Feedstock 121

Feedstock

4-Methylanisole – Lignin derived model compound

No.

4

Without catalysts

Catalyst Dielectric Barrier Discharge

Reactor type Mixture of Ar and H2

Carrier gas Effects of different composition of carrier gas mixture, voltage and pulse frequency were investigated: - The major products were 4-methylphenol, 2,4-dimethylphenol and p-xylene. - Conversion of 4-methylanisole reduced when increasing % of H2 in carrier gas. - Increasing the voltage, selectivity for 4-methylphenol and 2,4-dimethylphenol have a descending trend, while the trend for other products is ascending. - By increasing voltage and frequency, both conversion and discharge power increased.

Note

Table 4.8 Studies on upgrading bio-oil with integration of cold plasma. (Continued)

[157]

Ref.

(Continued)

2015

Year

122 Liquid Biofuel Production

Feedstock

4-Methylanisole – Lignin derived model compound

No.

5

Commercial catalysts: 2 type of Ni-Mo/ Al2O3; Pt-Cl/ Al2O3; Co-Mo/ Al2O3; Pt-Re/ Al2O3; Ni/Al2O3 and Al2O3

Catalyst Dielectric Barrier Discharge

Reactor type Ar

Carrier gas Effects of various catalysts and discharge power (DP) were investigated: - Conversion decreased in the following order: Pt-Cl/Al2O3 > Ni-Mo/Al2O3 (1) > Ni-Mo/Al2O3 (2) > Co-Mo/Al2O3 > Pt-Re/Al2O3 > Ni/Al2O3 > Al2O3 > plasma alone. - The plasma catalysis system has a lower discharge power, electrode temperature and breakdown voltage compared to plasma alone. - The main products were 4-methylphenol, 2,4-dimethylphenol and p-xylene.

Note

Table 4.8 Studies on upgrading bio-oil with integration of cold plasma. (Continued)

[164]

Ref.

(Continued)

2015

Year

Biofuel from Lignocellulosic Feedstock 123

Feedstock

4-Methylanisole – Lignin derived model compound

No.

6

Reactor type Corona discharge

Catalyst

Without catalysts

Ar

Carrier gas Effects of pulse repetition frequency, gap distance, pin number, carrier gas flower rate and plate of electrode diameter were investigated: - BTX and phenol with maximum selectivity of about 15% and 46%. - Demethylation, transalkylation, hydrogenolysis and demethoxylation and methane decomposition are the main chemical reactions. - Selectivity for BTX and phenol enhanced when increasing frequency or gap distance or pin number. - Selectivity of phenol and BTX peaked at Ar flow rate of 300 ml/min. - Higher PED was more favourable for phenol and BTX production

Note

Table 4.8 Studies on upgrading bio-oil with integration of cold plasma. (Continued)

[165]

2017

(Continued)

Ref.

Year

124 Liquid Biofuel Production

Feedstock

Rape straw

No.

7

Commercial HZSM-5

Catalyst

Reactor type Bio-oil vapour likes as carrier gas.

Carrier gas

Authors investigated the impact of catalyst temperature, catalyst height and discharge power on the yield and properties of bio-oil: - Cold plasma slightly decreased oil yield but increased quality of bio-oil. - NTP could completely eliminate fibrous coke; and decrease the attachment of graphite coke, as well as improve anticoking performance of the HZSM-5 catalyst.

Note

Table 4.8 Studies on upgrading bio-oil with integration of cold plasma. (Continued) Ref. [166]

Year 2017

Biofuel from Lignocellulosic Feedstock 125

126

Liquid Biofuel Production

Table 4.9 Effect of cold plasma on bio-oil catalytic upgrading [166]. Parameter

NTP + HZSM-5

HZSM-5

Oxygen content, wt%

19.79

26.25

High heating value, MJ/kg

33.14

29.50

pH value

4.98

4.47

Heavies, wt%

3.15

6.38

Chemical composition (relative content), % Aromatics

91.95

85.35

Hydrocarbons

22.48

14.17

Acids

0.00

1.14

such as anisole and 4-methylanisole. It was found that the conversion was up to 99% with the maximum degree of deoxygenation of 47%. Aromatic compounds such as benzene and xylene were found in the liquid product (Table 4.8). The effect of cold plasma on bio-oil upgrading is shown in Table 4.9. The quality of upgraded bio-oil was improved significantly, particularly oxygen content and heating value when applying cold plasma catalysis upgrading compared to a conventional HZSM-5 catalytic upgrading method. Furthermore, the amount of coke deposits on the catalyst decreased from 5.9% to 2.1% [166].

4.4 Gasification Gasification is a partial oxidation process to produce synthetic gas (syngas), which is a mixture of mainly H2 and CO with small amount of CH4, CO2, N2, tars at a temperature range of 900-1500 °C [11]. The syngas composition is strongly influenced by the types of a gasifier and operating conditions such as temperature, gasifying agents (O2, CO2, air or steam), particle size, properties of biomass feedstock and equivalence ratio [11, 167]. The syngas can be used to generate heat and electricity using gas turbines and internal combustion engines [26, 168] or feedstock for chemical processes, i.e. methanol and liquid hydrocarbons via FT synthesis. Depending on applications, the syngas needs to be cleaned and/or upgraded to meet requirements, i.e. particulates, tar, H2S and AAEM metals as shown in Table 4.10.

< 50

< 100 – –

< 30

Pt-Co/SiO2 > Co/SiO2 [263]. The selectivity of C5+ hydrocarbons and CO conversion increases when doping Rhenium (Re) on Co/Al2O3, Co/TiO2 and Co/SiO2 catalysts [264]. An addition of alkali metal ions (i.e. Na+ and K+) in Fe catalysts promotes the water gas shift reaction and activity of Fe catalysts whereas Cs+, Rb+ and Li+ have negative impacts [265].

Biofuel from Lignocellulosic Feedstock 145 Table 4.14 The advantages and disadvantages of different types of catalysts in Fischer-Tropsch synthesis [225, 232, 237, 239–245]. Catalyst

Advantage

Disadvantage

Fe

- Relatively low cost and availability. - Promote water-gas shift reaction. - Possible to use syngas with H2/CO ratio less than 2. - Operated at a wide range of temperature (210-350 °C). - Suitable to produces aromatic compounds and alcohols.

-

Ni

- High hydrogenation activity. - Suitable for methane formation.

- High cost. - During the reaction Ni carbonyls are formed (highly toxic).

Co

- High activity (up to 60-70% conversion per single pass). - High selectivity to long chain alkanes. - High deactivation resistance. - Long life time. - Suitable for biodiesel generation. - No water-gas shift reaction occurred.

- Excessive CH4 formation at high temperature. - Low selectivity to oxygenate products and alkene. - Difficulty to regeneration of deactivated catalysts. - Low hydrogenolysis and low shift activity at a high H2/CO ratio in syngas. - High investment cost. - Required syngas with high H2/ CO ratio (2.06-2.16).

Ru

- The highest active catalysts for FT synthesis. - High hydrogenation activity, when alkanes are preferred as the main product.

- High cost.

Low hydrogenation activity. Low selectivity to methane. Low deactivated resistance. Sensitive to the presence of sulphur in the syngas.

4.5.3 Influence of Operating Parameters on Fisher-Tropsch Synthesis The effects of operating conditions such as temperature, pressure, syngas composition etc. on the product selectivity are shown in Table 4.15. The

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Liquid Biofuel Production

conversions of H2 and CO and the selectivity of long chain hydrocarbons (C5+) increase with decreasing reaction temperature [266–268]. This could be because (i) light hydrocarbons (i.e. CH4) have higher activation energy than heavy hydrocarbons (C5+), resulting in an increased selectivity of light hydrocarbons (CH4) with increasing temperatures and (ii) high temperature promotes CO dissociation and provides more C atoms on the catalyst surface, leading to the formation of light hydrocarbons [269]. High H2/CO ratios in the syngas favour the chain termination therefore promoting the selectivity of light hydrocarbon (C1-C4) whereas low H2/CO ratios are preferential for the chain growth and the production of long chain hydrocarbons (C5+) [269, 270]. This was because the coverage of CO increases whereas the coverage of H2 decreases with decreasing H2/ CO ratios [271]. According to Tian et al. [269] and Mirzaei et al. [272], the coverage of CO increased significantly from 0.21-0.57 while that of H2 dramatically decreased from 0.59 to 0.15 when decreasing a H2/CO ratio from 5.0 to 1.0, leading to more heavy hydrocarbons (C5+) formation. The pressure can affect the reactivity of catalyst and product distribution in the FT process. The CO conversion increases with increasing total pressure in the system, resulting in an increase chain growth probability, favouring the selectivity of long chain hydrocarbon products (C5+) [273]. Tian et al. [269] reported that CH4 formation decreased while the C5+ increased with increasing total pressure from 1.5 MPa to 3.0 MPa. An increase in pressure enhances the collisions between reactants and the catalyst surface. The CO coverage decreased from 0.55 to 0.23, whilst the H2 coverage increased from 0.30 to 0.44 when total pressure decreased from 3.0 to 1.0 MPa. This is due to high desorption energy of CO. However, high Table 4.15 Influence of FT operating conditions on product selectivity [217, 276]. Operating parameter being increased Selectivity parameter

Temperature

Pressure

H2/CO ratio

Carbon number distribution (chain growth)

Lower α value

Higher α value

Lower α value

CH4 selectivity

Increased

Decreased

Increased

Alkene selectivity

–

–

Decreased

Oxygenate selectivity

–

Increased

Decreased

Aromatic selectivity

Increased

–

Decreased

Biofuel from Lignocellulosic Feedstock 147 total pressure could cause the lower CO conversion and reduce the rates of reactant adsorption process due to the mass transfer limitations of the liquid products on the catalyst surface [274, 275].

4.6 Summary Advanced thermochemical conversion including fast pyrolysis and gasification shows great potential for converting biomass waste/residues into liquid fuels for transportation sector to replace fossil-based fuels. Although high yield could obtain in a short reaction time (70-75 wt% based on dry biomass), the bio-oil derived from fast pyrolysis has very different properties to conventional gasoline/diesel such as high water content and oxygen content, acidity, and chemical and thermal instability. The fluidized bed reactors are commonly used for fast pyrolysis due to its heat and mass transfer properties superior to others. Over the last five years, four commercial fast pyrolysis plants are commissioned and operated in Canada, Finland, the United States, and the Netherlands for production of bio-oil as feedstock for boiler applications. Hydrodeoxygenation (HDO) and catalytic cracking are the two main routes that are used to remove oxygen from pyrolysis liquid. In HDO, twostage process is applied to overcome the issues such as coking formation or plugging at lines. In catalytic cracking, heat of pyrolysis zone or bio-oil vapour stream can be utilised by embedding catalyst into the pyrolysis reactor (in-situ) or coupling in a separate reactor (ex-situ). However, the quality of bio-oil derived from catalytic cracking upgraded is not meet fuel quality standards. Therefore, this method can be used as the first step to reduce oxygen content in pyrolysis which then further upgraded using HDO. Although some remarkable results have achieved in upgrading of biooil for transportation fuels, it has been only in laboratory scales. The main challenges are rapid catalyst deactivation, high temperatures and pressure (HDO process) and large amount of hydrogen required for upgrading processes. Integration and intensification processes such as fast hydropyrolysis and cold plasma technologies to improve quality of bio-oil have become promising approaches to reduce the operation costs and the complexity of upgrading system. Cold plasma technologies have been tested with model compounds and found that oxygen removal can be carried out at lower temperatures and pressures over a much short reaction time (few seconds to minutes compared to several hours). Combining cold plasma and catalysts enhances significantly the process, i.e. prolonged catalyst lifetime, and high degree of oxygen removal. However, energy efficiency of cold plasma

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Liquid Biofuel Production

assisted process needs to be assessed and therefore more research needs to be done to optimise the design of cold plasma. Gasification of biomass has been intensively studied to convert biomass into syngas or H2. The main challenge is the quality of syngas such as low H2/CO ratio (< 1) and impurities such as tars. Therefore, a fully understanding the effects of operating conditions and types of gasifier could improve gasification process to enhance the ratio of H2/CO (≥ 2) for liquid hydrocarbon production via a FT synthesis and to minimise tar formation. Furthermore, hydrogen from gasification can be used for upgrading of biooil in the HDO. This would enhance the sustainability of biofuel production from biomass as hydrogen is currently produced via steam reforming of methane. Although, the FT synthesis is a commercialised process to produce biofuel (liquid hydrocarbons) using Fe and Co catalysts, challenges still remain such as low selectivity of C5+, high pressure requirements and catalyst deactivation. Developing advanced catalysts, i.e. applied bimetallic catalysts (Co/Ni and/or Co/Fe) with noble metals promotes (Pd, Pt and Ru) have attracted attention in the FT synthesis.

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Biofuel from Lignocellulosic Feedstock 165 254. Li, J., Jacobs, G., Das, T., Zhang, Y., Davis, B., Fischer–Tropsch synthesis: Effect of water on the catalytic properties of a Co/SiO2 catalyst. Appl. Catal., A, 236, 67–76, 2002. 255. Tavasoli, A., Sadagiani, K., Khorashe, F., Seifkordi, A.A., Rohani, A.A., Nakhaeipour, A., Cobalt supported on carbon nanotubes - A promising novel Fischer-Tropsch synthesis catalyst. Fuel Process. Technol., 89, 491–498, 2008. 256. Reuel, R.C. and Bartholomew, C.H., Effects of support and dispersion on the CO hydrogenation activity/selectivity properties of cobalt. J. Catal., 85, 78–88, 1984. 257. Oh, J.-H., Bae, J.W., Park, S.-J., Khanna, P.K., Jun, K.-W., Slurry-phase Fischer–Tropsch synthesis using Co/γ-Al2O3, Co/SiO2 and Co/TiO2: Effect of support on catalyst aggregation. Catal. Lett., 130, 403–409, 2009. 258. Khodakov, A.Y., Chu, W., Fongarland, P., Advances in the development of novel cobalt Fischer–Tropsch catalysts for synthesis of long-chain hydrocarbons and clean fuels. Chem. Rev., 107, 1692–1744, 2007. 259. Enache, D.I., Roy-Auberger, M., Revel, R., Differences in the characteristics and catalytic properties of cobalt-based Fischer–Tropsch catalysts supported on zirconia and alumina. Appl. Catal., A, 268, 51–60, 2004. 260. Feyzi, M., Irandoust, M., Mirzaei, A.A., Effects of promoters and calcination conditions on the catalytic performance of iron–manganese catalysts for Fischer–Tropsch synthesis. Fuel Process. Technol., 92, 1136–1143, 2011. 261. Morales, F., de Groot, F.M.F., Gijzeman, O.L.J., Mens, A., Stephan, O., Weckhuysen, B.M., Mn promotion effects in Co/TiO2 Fischer–Tropsch catalysts as investigated by XPS and STEM-EELS. J. Catal., 230, 301–308, 2005. 262. Morales, F., de Smit, E., de Groot, F.M.F., Visser, T., Weckhuysen, B.M., Effects of manganese oxide promoter on the CO and H2 adsorption properties of titaniasupported cobalt Fischer–Tropsch catalysts. J. Catal., 246, 91–99, 2007. 263. Tsubaki, N., Sun, S., Fujimoto, K., Different Functions of the Noble Metals Added to Cobalt Catalysts for Fischer–Tropsch Synthesis. J. Catal., 199, 236– 246, 2001. 264. Storsæter, S., Borg, Ø., Blekkan, E.A., Holmen, A., Study of the effect of water on Fischer–Tropsch synthesis over supported cobalt catalysts. J. Catal., 231, 405–419, 2005. 265. Ngantsoue-Hoc, W., Zhang, Y., O’Brien, R.J., Luo, M., Davis, B.H., Fischer– Tropsch synthesis: Activity and selectivity for Group I alkali promoted ironbased catalysts. Appl. Catal., A, 236, 77–89, 2002. 266. Dry, M.E., Practical and theoretical aspects of the catalytic Fischer-Tropsch process. Appl. Catal., A, 138, 319–344, 1996. 267. Claeys, M. and van Steen, E., Chapter 8 - Basic studies, in: Studies in Surface Science and Catalysis, A. Steynberg and M. Dry (Eds.), pp. 601–680, Elsevier, The Netherlands, 2004. 268. Zhu, X., Lu, X., Liu, X., Hildebrandt, D., Glasser, D., Heat transfer study with and without Fischer-Tropsch reaction in a fixed bed reactor with TiO2, SiO2, and SiC supported cobalt catalysts. Chem. Eng. J., 247, 75–84, 2014.

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269. Tian, L., Huo, C.-F., Cao, D.-B., Yang, Y., Xu, J., Wu, B.-S., Xiang, H.-W., Xu, Y.-Y., Li, Y.-W., Effects of reaction conditions on iron-catalyzed Fischer– Tropsch synthesis: A kinetic Monte Carlo study. J. Mol. Struct. Theochem, 941, 30–35, 2010. 270. Kwack, S.-H., Park, M.-J., Bae, J.W., Ha, K.-S., Jun, K.-W., Development of a kinetic model of the Fischer–Tropsch synthesis reaction with a cobalt-based catalyst. React. Kinet. Mech. Catal., 104, 483–502, 2011. 271. Zhang, X., Qian, W., Zhang, H., Sun, Q., Ying, W., Effect of the operation parameters on the Fischer–Tropsch synthesis in fluidized bed reactors. Chin. J. Chem. Eng., 26, 245–251, 2018. 272. Mirzaei, A.A., Shirzadi, B., Atashi, H., Mansouri, M., Modeling and operating conditions optimization of Fischer–Tropsch synthesis in a fixed-bed reactor. J. Ind. Eng. Chem., 18, 1515–1521, 2012. 273. Sauciuc, A., Abosteif, Z., Weber, G., Potetz, A., Rauch, R., Hofbauer, H., Schaub, G., Dumitrescu, L., Influence of operating conditions on the performance of biomass-based Fischer–Tropsch synthesis. Biomass Convers. Biorefin., 2, 253–263, 2012. 274. Feyzi, M., Mirzaei, A.A., Bozorgzadeh, H.R., Effects of preparation and operation conditions on precipitated iron nickel catalysts for Fischer-Tropsch synthesis. J. Nat. Gas Chem., 19, 341–353, 2010. 275. Jalama, K., Effect of operating pressure on Fischer-Tropsch synthesis kinetics over titania-supported cobalt catalyst, in: The World Congress on Engineering and Computer Science, Springer Singapore, Singapore, 2017. 276. Klerk, A.D., Chapter 4 - Fischer–Tropsch synthesis, in: Fischer-Tropsch Refining, A.D. Klerk (Ed.), pp. 73–103, Wiley-VCH, Singapore, 2011.

5 Exploring the Potential of Carbohydrate Rich Algal Biomass as Feedstock for Bioethanol Production Jaskiran Kaur and Yogalakshmi K.N.* Centre for Environmental Sciences and Technology, School of Environment and Earth Sciences, Central University of Punjab, Bathinda, India

Abstract Transport fuels in the 21st century are scarce to fulfill the demands of the ever-increasing human population. Moreover, pollution problems, particularly global warming, that arise with the continuous usage shifted the research focus towards development of ecofriendly fuels. Bioethanol, a biofuel used in the form of a fuel blend emerged as a more promising transport fuel. During the past decades, bioethanol was produced from feedstocks like starch and sugar crops and lignocellulosic biomass. However food insecurity, water shortage and limited land availability problems associated with these feedstocks necessitated the development of sustainable feedstock. Carbohydrate-rich algae having inherent ability to grow rapidly in wastewater or saltwater thereby eliminating the need of fresh water would be considered as a suitable option. This chapter evaluates the different algal strains used in bioethanol production and various steps (pretreatment, fermentation and distillation) used for bioethanol production from algal biomass. In addition, strategies for engineering algal strains using genetic techniques for optimizing bioethanol production is also discussed in the chapter. Keywords: Global warming, transport fuels, biofuel, bioethanol, algae, pretreatment, fermentation, distillation

*Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (167–196) © 2019 Scrivener Publishing LLC

167

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5.1 Introduction The human population has developed at the speed of a hare in the 21st century while energy reserves has continued snailing along. Continuous supply of energy is the need of the hour in the transportation and industrial sectors. As per recent reports, the transport system alone consumed about 29% of energy resources in the US in 2016 [1]. Fossil fuels are considered as potential energy carriers. However, qualms over a rise in petroleum prices coupled with the greenhouse gases emissions linked with the burning of fossil fuels necessitated the development of renewable energy sources [2]. Biofuels, viz. bioethanol, are considered a vital energy source to combat the problem of greenhouse gas emissions and global warming. The biodegradability and less toxicity of bioethanol makes bioethanol an appropriate replacement of the fossil reserves. The demands of bioethanol particularly in the form of fuel blend has risen in recent years. The quantity of bioethanol blend with gasoline consumed for transportation in the United States increased from 1.4 billion gallons in 1995 to 14.4 billion gallons in 2016 [3]. Bioethanol can be produced from various feedstocks. On the basis of type of feedstocks, bioethanol can be grouped into three categories, namely first-generation, second-generation and third-generation bioethanol. First-generation, also termed conventional, bioethanol is primarily derived from food crops like starch, sugars, wheat, and sorghum. The problems of habitat loss, biodiversity reduction, soil erosion, and a rise in food prices has shifted the focus towards next-generation or secondgeneration biofuels [4]. Lignocellulosic biomass or agricultural residue are mainly used as a substrate for production of second-generation biofuels. Although they are cost-effective, the challenges associated with the land and water requirements for the cultivation of second-generation biofuels limits their usage [5]. In the coming years, the emergence of algae as a source of bioethanol is considered a sustainable approach to deal with the limitations of first- and second-generation biofuels. Due to the exceptional ability of algae to grow on land unsuitable for food production using wastewater or brackish water, the interest in developing algae as a resource of bioethanol has exploded in the present era. Additionally, the cultivation of algae for bioethanol production is more beneficial in terms of yield contrast to cultivation of other feedstocks (Table 5.1). Compared to land plants, the energy requirement for the production of supporting tissue is less in algae. Also, algae take up the nutrients over their entire surface, which results in increased bioethanol yield by algae [6].

Potential of Algal Biomass as Feedstock

169

Table 5.1 Comparison of algae with other feedstocks. (Adapted from Adams et al. [7]).

Feedstock

Average world yield (kg/ha/ year)

Dry weight of hydrolysable carbohydrates (kg/ha/year)

Potential ethanol volume (L/ ha/year)

Wheat (grain)

2800

1560

1010

Maize (kernel)

4815

3100

2010

Sugarbeet

47070

8825

5150

Sugarcane

68260

11600

6756

Macroalgae

730000

40150

23400

5.2

Microalgae and Macroalgae as Bioethanol Feedstock

The term algae comprises a large group of photosynthetic organisms representing the oldest life-forms which is responsible for almost 50% of photosynthesis on Earth [8]. Algae possess chlorophyll as a primary photosynthetic pigment and lack roots, stems, and leaves. The sterile covering of cells around the reproductive cells is absent in algae [9]. Algae are ubiquitous in distribution and can grow in different habitats including fresh water, salt water, wet soils, pools and deserts, etc. Most of the algae is photoautotrophic while some can grow under heterotrophic conditions. The phototrophic algae have an inherent ability to convert sunlight and carbon dioxide into nutrients such as carbohydrates. Besides photoautotrophy, certain algae are heterotrophic that are capable of growing in the dark, which eliminates the dependency on the light source [10]. The algae is categorized into microalgae (or phytoplankton or microphytes) and macroalgae (or seaweeds) on the basis of their morphology and size. As the name suggests, microalgae refers to the small photosynthetic organisms, most of which are unicellular and are made of 28-30% protein, 2-40% lipids/ oils and 4-57% carbohydrates by weight [11]. The microalgae are classified under four main groups namely, Cyanophyceae (blue-green algae), Chlorophyceae (green algae), Bacillariophyceae (including the diatoms) and Chrysophyceae (including golden algae) [12]. Apart from being exploited for production of biodiesel and bioethanol, certain other valueadded products such as omega-3 fatty acids, pharmaceutical products, biogas, biohydrogen are also produced from microalgae [13–15]. Chlorella

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vulgaris, Scenedesmus obliquus, Dunaliella tertiolecta and Microcystis aeruginosa are some of the microalgae which are efficiently used as feedstocks for conversion to bioethanol [16–19]. Microalgae is known to have the capability to accumulate high amount of lipids. This serves microalgae as an indispensable candidate for production of biodiesel. However, the biomass left after production of biodiesel contains carbohydrates which can further be used for the production of bioethanol. Kumar et al. [20] obtained 0.158 g of bioethanol from the lipid extracted residual biomass. The development of such a cyclic system promotes the sustainable production of biofuels in a biorefinery. Another category of algae, macroalgae, are large aquatic photosynthetic plants consisting of leaf-like thallus. Red, brown and green algae are the important classes of macroalgae. Macroalgae contains approximately 70-90% water, high protein content (app. 10%) and varying levels of carbohydrates [21]. As compared to microalgae, they have low lipid content. Lack of lignin or comparatively much less lignin in the cellulosic structures of macroalgae makes it separable from the lignocellulosic biomass. Research in the field of using macroalgae for bioethanol production is gaining impetus due to its ability to produce carbohydrate rich biomass from inexpensive raw materials [22, 23]. Various macroalgae, for instance, Sacchoriza, Alaria, Sargassum fulvellum, Ulva lactuca and Spirogyra are used for bioethanol production [11, 24, 25]. In addition, certain red algae, e.g., Gelidium amansii, contain agar, a polymer of galactose and galactopyranose which is difficult to metabolize into ethanol. This necessitated the use of methods of saccharification for unlocking galactose from agar and to further release cellulose from glucose so as to increase ethanol yields during fermentation [26]. Hence, the type of algal strain and the pretreatment method play a crucial role in terms of ethanol yield [27]. Table 5.2 summarizes the studies related to the production of bioethanol from various microalgae and macroalgae along with the pretreatment method applied to algal biomass. Most of the studies explored the potential of bioethanol production using microalgae belonging to the Chlorophyceae (green algae) algal group. The microalgae Chlorella vulgaris can accumulate 37% (dry weight) of starch and is thus considered to be the most promising feedstock [28]. Another microalgae, Chlamydomonas reinhardtii, showed rapid growth but it contains starch granules in the cell which necessitated the pretreatment of algal biomass for releasing the fermentable sugars and other nutrients for yeast fermentation [29, 30]. Likewise, macroalgae also serves as a raw material for bioethanol production. The macroalgal cell wall is composed of a varying amount of carbohydrate. The green algae contains 25-50% carbohydrate content whereas red and brown algal cell wall

Green

Green

Green

Brown

Green

Green

Chlamydomonas reinhardtii UTEX 90

Chlorococcum humicola

Schizocytrium sp.

Chlamydomonas reinhardtii

Scenedesmus abundans PKUAC 12

Algae type

Dunaliella

Microalgae

Algae used

Acid pretreatment with 3% H2SO4 followed by cellulase

12N H2SO4

44 10.3

Saccharomyces cerevisiae

5.5

Escherichia coli KO11 Saccharomyces cerevisiae

48

Saccharomyces cerevisiae

H2SO4 pretreatment

Hydrothermal degradation and enzymatic hydrolysis

23.5

Saccharomyces cerevisiae

Enzymatic treatment using α-amylase of B. licheniformis origin and amyloglucosidase from Aspergillus niger, pH 5.6, temperature 37°C

Ethanol yield (%)

3.4

Fermenting organism

Saccharomyces cerevisiae IAM 4140

Enzymatic hydrolysis with glucoamylase

Pretreatment method

Table 5.2 Studies of bioethanol production from microalgal and macroalgal biomass.

(Continued)

[34]

[30]

[33]

[32]

[29]

[31]

Reference

Potential of Algal Biomass as Feedstock 171

HCl-catalyzed saccharification at 121°C for 15 min

Virus infection and enzymatic hydrolysis

Green

Green

Green

Green

Green

Yellow-green

Dunaliella tertiolecta

Chlorella variabilis NC64A

Chlorella vulgaris FSP-E

Scenedesmus obliquus

Chlorella vulgaris

Tribonema sp.

Acid hydrolysis with 3% H2SO4 at 121°C for 45 min

Sonication and milling pretreatments accompanied by enzymatic hydrolysis

Dilute acid hydrolysis with 2% H2SO4

Dilute acidic hydrolysis with 1% H2SO4 followed by enzymatic hydrolysis

Pretreatment method

Algae type

Algae used

Saccharomyces cerevisiae

Saccharomyces cerevisiae KCTC 7906

Zymomonas mobilis ATCC29191

Zymomonas mobilis

Escherichia coli KO11

Saccharomyces cerevisiae YPH500

Fermenting organism

Table 5.2 Studies of bioethanol production from microalgal and macroalgal biomass. (Continued)

56.1

89

21.3

87.5 (theoretical yield)

32

14

Ethanol yield (%)

(Continued)

[40]

[39]

[38]

[37]

[36]

[35]

Reference

172 Liquid Biofuel Production

Green

Blue-green

Halophilic

Green

Chlorella sp. KR-1

Spirulina

Dunaltella sp.

Chlorella vulgaris

Brown

Red

Laminaria hyperborea

Gracilaria salicornia

Macroalgae

Algae type

Algae used

Pichia angophorae Escherichia coli

Acid hydrolysis with 2% H2SO4 at 120°C and for 30 min followed by enzymatic hydrolysis with cellulase

Saccharomyces cerevisiae

Saccharomyces cerevisiae

Saccharomyces cerevisiae

Saccharomyces cerevisiae

Fermenting organism

Disruption and washing in water at pH 2 and 65°C

Microwave irradiation with 1M HCl at 100°C for 60 min and hydrothermal treatment with 1M HCl at 120°C for 60 min

Acid pretreatment with H2SO4

Acid pretreatment with 1-2% H2SO4

Simple enzymatic and chemical treatment using Pectinex at pH 5.5 and 45°C and 0.3 N HCl at 121°C for 15 min

Pretreatment method

Table 5.2 Studies of bioethanol production from microalgal and macroalgal biomass. (Continued)

7.9

0.86

13.2

24.2

0.85-1

16

Ethanol yield (%)

(Continued)

[46]

[45]

[44]

[43]

[42]

[41]

Reference

Potential of Algal Biomass as Feedstock 173

Algae type

Brown

Brown

Green

Brown

Red

Red

Red

Algae used

Laminaria japonica

Laminaria digitata

Spirogyra

Alaria crassifolia

Gelidium elegans

Gelidium corneum

Kappaphycus alvarezii

0.9 N H2SO4 at 100°C for 1 h

0.5–1% oxalic acid at 121°C for 30 and 60 min

Acid hydrolysis with 0.2% H2SO4 accompanied by enzymatic hydrolysis with Meicelase pretreatment in 50°C for 120 h

Saccharomyces cerevisiae (NCIM 3523)

Saccharomyces cerevisiae

Saccharomyces cerevisiae IAM 4178

15.4

5.8

5.5

>3

(Continued)

[50]

[49]

[48]

[48]

Saccharomyces cerevisiae IAM 4178

[47]

[11]

Reference

Acid hydrolysis with 0.2% H2SO4 accompanied by enzymatic hydrolysis with Meicelase pretreatment in 50°C for 120 h

13.2

16.1

Ethanol yield (%)

[25]

Pichia angophorae

Recombinant Escherichia coli KO11

Fermenting organism

Chemical pretreatment with 1% NaOH for 2 hrs

pH 4 with 2 M and 0.2 M HCl

Acid hydrolysis followed by simultaneous enzyme treatment

Pretreatment method

Table 5.2 Studies of bioethanol production from microalgal and macroalgal biomass. (Continued)

174 Liquid Biofuel Production

Algae type

Red

Brown

Green

Green

Red

Red

Red

Brown

Algae used

Gracilaria verrucosa

Sargassum

Chaetomorpha linum

Ulva fasciata Delile

Gracilaria sp.

Gelidium amansii

Eucheuma cottonii

Saccharina japonica

Unpretreated

Defluviitalea phaphyphila Alg1

Saccharomyces cerevisiae

Acid hydrolysis with 3% H2SO4

25

2.49

1.6

23.6

Saccharomyces cerevisiae Pichia stipitis KCTC 7228

45

44

65

43

Ethanol yield (%)

Saccharomyces cerevisiae MTCC No. 180

Saccharomyces cerevisiae ATCC 96581

Saccharomyces cerevisiae

Saccharomyces cerevisiae

Fermenting organism

Thermal acid hydrolysis with 91 mM H2SO4 and enzyme saccharification

Sequential acid pretreatment with H2SO4 and enzymatic hydrolysis with cellulase

Enzymatic hydrolysis using cellulase 22119

Ball milling

3.4–4.6% H2SO4 concentration, 115°C and 1.50 h with enzymatic hydrolysis

Enzymatic hydrolysis with cellulase and β-glucosidase

Pretreatment method

Table 5.2 Studies of bioethanol production from microalgal and macroalgal biomass. (Continued)

[58]

[57]

[56]

[55]

[54]

[53]

[52]

[51]

Reference

Potential of Algal Biomass as Feedstock 175

176

Liquid Biofuel Production

constitute 30-60% and 30-50% of carbohydrates, respectively, which makes them an efficient raw material for bioethanol production [59]. The algal groups Laminaria and Alaria, however, contains complex carbohydrates such as laminarian and mannitol in their structure; hence the pretreatment of seaweeds is the paramount need for the sustainable production of bioethanol [58]. The details of the entire processes involved for the conversion of algal biomass into ethanol is given in the following sections.

5.3 Process Involved for Production of Bioethanol from Algae Bioethanol production from algae involves a series of steps which is depicted in Figure 5.1. At first, the starch accumulating algae is cultivated in aqua culture environments using open ponds or closed photobioreactors (PBR) with sunlight as a source of energy. The municipal wastewater or brackish water contains sufficient quantities of nutrients, hence it served as a culture medium for the algal growth. The algae are harvested and sugars present in the algae are then fermented by micro-organisms. Since most of the sugars are present in the form of structural and storage carbohydrates, pretreatment is needed to rupture the algal cell wall. The pretreatment of algal biomass is done by mechanical methods (e.g., bead beating, ultrasonic disintegrators, microwave, sonication, etc.) or with enzymatic dissolution

Nutrients + Water

Algal cultivation Open/Photobioreactor

Biomass separation/ Harvesting

Biomass pretreatment (Physical/Chemical)

Glucose fermentation (Yeast/Bacteria)

Ethanol

Figure 5.1 Schematic view of bioethanol production from algae.

Distillation

Potential of Algal Biomass as Feedstock

177

of cell wall [60]. After, pretreatment, the carbohydrates are released from the intracellular medium of algae [61]. The released carbohydrates then need to be hydrolyzed into simple sugars prior to fermentation, which is carried out through a process termed saccharification. The saccharification is done using acid or enzymatic (alpha- and glucoamylase) hydrolysis. In the succeeding steps, the sugars are fermented into ethanol. The saccharification and fermentation is done either simultaneously or in different steps depending on the type of algal strain used. The fermentation requires yeast for fermenting the decaying biomass into the fermentation solution. Aside from yeast, certain genetically manipulated bacteria known to possess the fermentation potential can also be used. Sugar pathway is used for fermenting algal biomass into ethanol. Finally, the ethanol is separated from the fermenting solution by distillation.

5.4 Algal Biomass Cultivation The choice of reactor plays a critical role in influencing the productivity of algal biomass. Certain features such as light supply, gas-liquid mass transfer, risk of contamination, cost effectiveness, easy control ability, and minimal land requirement for cultivation should be taken into consideration while selecting the cultivation system for algae [62]. Hence, the cultivation system should be designed in such a way so as to promote commercially viable and economically feasible production of algae. Among the above-mentioned characteristics, efficient light source and good circulation devices are the most important design criteria. Usually, light-emitting diode (LED), with the advantages of long-life expectancy and energy-saving ability, is used as a light source in the cultivation system. On the other hand, a good choice of circulation system ensures the uniform distribution of cells and liquid broth, proficient CO2 mass transfer efficiency and minimum heat generation within the system. Two types of reactors, namely open systems and closed systems, are used for the cultivation of algae. Table 5.3 lists the type of cultivation systems used for algal mass production. The intrinsic properties of algae along with local climatic conditions also influence the selection of a particular system [63].

5.4.1 Open Pond Systems Open pond systems, also termed raceway pond method, is the most widely used system. Typical raceway ponds are constructed of a closed loop with

Capacity

–

3L

2.5 L

100 L

2.5 L

1L

2.5 L

1L

1L

20 L

Cultivation system

Open ponds

Fermentor

Photobioreactor

Bag photobioreactor

Fermentor with surface aeration

Air lift reactor

Glass photobioreator

Glass photobioreactor

Photobioreactor

Column photobioreactor

Tribonema sp.

Scenedesmus obliquus

Chlorella vulgaris FSP-E

Arthrospira platensis SAG 21.99

Scenedesmus obliquus

Sargassum sagamianum

Chlorococcum humicola

Chlamydomonas reinhardtii UTEX 90

Laminaria hyperborea

Chlorella sp.

Algal species

Table 5.3 List of cultivation systems used for algae.

Microalgae

Microalgae

Microalgae

Microalgae

Microalgae

Macroalgae

Microalgae

Microalgae

Macroalgae

Microalgae

Classification

[40]

[38]

[37]

[65]

[17]

[24]

[64]

[29]

[45]

[63]

Reference

178 Liquid Biofuel Production

Potential of Algal Biomass as Feedstock

179

oval-shaped recirculation channels. The ponds are open to air with water depth of 0.2 to 0.5 m and are equipped with a paddle wheel for proper homogenization of culture. Raceway ponds are built by materials such as glass, concrete or membrane [61]. The ponds are operated in continuous mode with constant feeding of culture in front of the paddle wheel while algae is harvested behind the paddle wheel after completion of the circulation loop. The main benefit of raceway ponds is easy operation and low production cost. However, being open to the environment, they have the chance of contamination by other algae and microorganisms. Only highly selective algal species such as Chlorella (grows in nutrient-rich media), Spirulina (grows well in high alkalinity) and Dunaliella (grows at high salinity) can grow in open air cultures [66, 67]. Further, raceway ponds are commercially unfavourable due to the problems of light limitation, nutrient limitation, temperature fluctuations and ineffectual homogenization [61].

5.4.2 Closed Photobioreactors (PBR) The efficacy of closed PBR is determined by integration of capture, transportation, distribution and utilization of light by microalgae through photosynthesis [68]. The closed PBRs are superior to raceway ponds in terms of photosynthetic efficiency and biomass productivity. The materials used for the construction of PBR include glass, plexiglass, polyvinyl chloride (PVC), acrylic-PVC and polyethylene [69]. They are provided with sterilized gas filters for necessary gas exchange, so the chances of contamination are minimum in closed PBRs. They are designed in such a way as to maximize the surface/volume ratio. The flat plate and tubular PBR are primarily used for industrial production of algal biomass as they offer large surface/volume ratio. The better homogeneity of culture and mass transfer in these reactors increases the biofixation of CO2. The production of excess O2 limits its application [70]. Besides, the shortcoming of high equipment cost, closed PBRs have several advantages over open systems [71]: 1) Minimum chances of contamination and facilitates axenic algal cultivation; 2) Greater control over culture conditions such as light, pH, temperature, CO2 concentration; 3) Less CO2 loss; 4) Higher cell concentrations and 5)  Prevention of water evaporation. Different designs of closed PBRs are used for mass cultivation of algal species, the pros and cons of which are mentioned in Table 5.4.

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Table 5.4 Advantages and limitations of different closed PBRs. (Adapted from Ugwu et al. [72].) Type of closed photobioreactor

Advantages

Limitations

Vertical column PBR

High mass transfer, good mixing with low shear stress, low energy consumption, high potentials for scalability, easy to sterilize, readily tempered, good for immobilization of algae, reduced photoinhibition and photo-oxidation.

Small illumination surface area, construction requires sophisticated materials, stress to algal cultures, decrease of illumination surface area upon scale-up.

Flat plate PBR

Large illumination surface area, suitable for outdoor cultures, good for immobilization of algae, good light path, good biomass productivities, relatively cheap, easy to clean up, readily tempered, low oxygen buildup.

Scale-up requires many compartments and support materials, difficulty in controlling culture temperature, some degree of wall growth, possibility of hydrodynamic stress to some algal strains.

Horizontal tubular PBR

Large illumination surface area, suitable for outdoor cultures, fairly good biomass productivities, relatively cheap.

Gradients of pH, dissolved oxygen and CO2 along the tubes, fouling, some degree of wall growth, requires large land space.

5.5 Pretreatment of Algal Biomass The algal cell wall contains polysaccharide such as cellulose which must be converted into monomeric sugars such as glucose, fructose, etc. Hence, successful lysing of algal cells is the prerequisite for bioethanol production which require the selection of an appropriate pretreatment method to ensure the release of fermentable sugars. Pretreatment is carried out by various physical, chemical and biological methods.

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5.5.1 Physical Pretreatment The physical pretreatment implies various methods such as mechanical comminution, microwave treatment, bead beating and ultrasonication for the lysis of algal cell wall. Combination of methods such as chipping, milling and grinding are used for mechanical comminution of algal biomass for the reduction of cellulose crystallinity [73]. The purpose of this approach is to decrease the biomass particle size with the view to attain large surface area [64]. For large-scale application of mechanical comminution, energy consumed is higher than the theoretical energy present in biomass which makes it an unattractive method for biomass pretreatment [74]. Furthermore, another mechanical method, bead beating, is applied to the algal biomass. In this method, minute glass or ceramic beads spinning on high speed are collided with the algal biomass. The shear forces in this method are lower when compared to high-pressure techniques of cell disruption. Size of the beads is an important factor that needs to be considered. The rate of cell disruption is faster at the higher volume ratio of beads to cell suspension [75]. This method is applicable in the laboratory as well as industrial scale [76, 77]. Certain industries make use of microwave treatment and ultrasonication for physical pretreatment. The application of microwave irradiation enables the faster release of fermentable sugars from the algae [44]. Microwave technology reduces the cost associated with dewatering and extracting of dry algal biomass, hence is more economical than other physical methods. In microwave treatment, high-frequency waves are applied for degradation of algal biomass. The contact between the polar material (water) and rapidly oscillating electric field (produced by microwaves) produces heat due to the frictional forces resulting from inter- and intra-molecular movements [78]. Due to heat production, formation of water vapour takes place within the cell which eventually rupture the cell. This produces an electroporation effect that leads to opening of cell membrane and release of monosaccharides [79]. The microwaves when used with other pretreatment methods such as in conjunction with chemical can increase the yield of ethanol [44]. Ultrasonication is another pretreatment method where ultrasonic disintegrators disrupt the cells through generation of intense sonic pressure waves. The repetitive compression and rarefaction of waves causes the formation of cavities or microbubbles. The bubbles then collapse violently in a process called cavitation and generate mechanical shear force which break the cell wall and membrane [80]. During processing, the ultrasonic disintegrators produce extensive heat which is avoided by placing the algal sample in an ice-cold atmosphere [75]. The ultrasonication of algae Scenedesmus obliquus

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YSW15 resulted in release of carbohydrates to the cell surface and/or within the periplasmic membrane from the interior of cells [81].

5.5.2 Chemical Pretreatment The acid hydrolysis exposed the internal structures of the biomass and increased the accessible surface area for enzymatic hydrolysis [52]. Generally, dilute and concentrated acids are used for carrying out the acid hydrolysis of algal biomass. Sulphuric acids, hydrochloric acids, nitric acids and phosphoric acids are examples of concentrated acids used for pretreatment. Though highly efficient in hydrolysis of cellulose, the use of concentrated acids causes corrosion of equipments [82]. The toxicity and hazardous nature has shifted the research focus to dilute acids. The acid concentration and hydrolysis time are the important parameters to be considered during the pretreatment. Sulfuric acid below 4 wt% concentrations are effective for the successful pretreatment of biomass. Borines et al. [52] reported the optimum acid concentration in the range between 2.51-4.01% for the macroalgae Sargassum sp. at optimum temperature of 115°C and hydrolysis time of 1.12-1.5 h. Hence, milder temperatures are preferable pretreatment of algal biomass when compared to terrestrial biomass which is carried out under extreme temperature of 165-210°C due to the presence of lignin and high hemicellulose content [83]. Though dilute acid pretreatments are effective in enhancing cellulose hydrolysis, the conversion of glucose (released from cellulose) into hydromethylfurfural (fermentation inhibitors or toxic compound) takes place under severe pretreatment conditions [84, 27]. These toxic compounds inhibit cell growth, which results in decreased ethanol production [85]. The other chemical method that is commonly employed is the use of alkali for pretreatment. These processes use lower temperatures and pressure than other treatment methods. The chemical agents such as sodium, potassium, calcium and ammonium hydroxides are used for alkaline pretreatment, among which sodium hydroxide is most widely used [86, 87]. The alkaline pretreatment approach was explored for the first time for carrying out the hydrolysis of microalgal biomass from the species Chlorococcum infusionum using 0.75% NaOH at 120°C for 30 min [32]. The NaOH break the intermolecular bond of hemicelluloses with other polymeric substances. In alkaline processes, the degradation of sugars is less compared to acid pretreatment. Most of the caustic salts used for pretreatment can be recovered or regenerated. But, like dilute acid pretreatments, the formation of inhibitory by-products viz. furfural, hydroxymethylfurfural and formic acid also occur during alkali pretreatment processes, resulting in reduced productivity [88].

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5.5.3 Biological Pretreatment The biological pretreatment method for degradation of biomass for releasing fermentable sugars relies on the use of microbes and enzymes. Though slower and more expensive than other pretreatment methods, the biological pretreatment requires less energy and is capable of disrupting the cell wall completely without formation of any inhibitory by-products. However, the hydrolysis rate is low in this process when compared to other treatment processes, which can be enhanced by prior pretreatment of algal biomass using physical and chemical pretreatment methods [89]. In the biological pretreatment method, fungi such as brown, white and soft rot are used for the degradation of lignin and hemicellulose [90]. The brown rots can degrade cellulose while white and soft rots attack both cellulose and lignin [73]. The long process time and requirement of continuous monitoring of the growth of micro-organisms however, limits its utilization on commercial scale [91]. The enzymatic hydrolysis employ the use of cellulase enzyme produced by bacteria and fungi for the degradation of cellulose into glucose. Clostridium, Bacillus, Streptomyces, Cellulomonas and Microbispora are some of the examples of bacteria possessing cellulase enzyme whereas fungi species producing cellulase enzyme include Trichoderma, Fusarium, Penicillium, Schizophillum sp. etc. [59]. Three types of cellulase enzymes, namely endoglucanase, exoglucanase and beta glucosidase are known to degrade cellulose. The endonuclease randomly cleave the region where cellulose fibers have low crystallinity. The exonuclease attacks the cellulose units from the ends of the exposed chains produced by endonuclease. Beta glucosidase hydrolyze the exocellulase products into glucose [59]. The maximum enzyme hydrolysis activity is found at optimum conditions of temperature, pH and time period. The extraction of glucose from algae Chlamydomonas reinhardtii has been done at optimum enzymatic conditions of 45°C, pH 4.6 and 60 min [92].

5.6 Fermentation of Algal Hydrolysate During fermentation, different micro-organisms and bacteria convert fermentable sugars into ethanol [93]. The selection of an ideal microorganism depends on certain factors: ethanol tolerance, osmotic tolerance, broad substrate utilization efficiency, ability to withstand high temperature, and resistance for inhibitors present in the hydrolysate and genetic stability [94]. The fermentation processes followed for fermentation of algal hydrolysate includes [84, 95]: (1) separate hydrolysis and fermentation (SHF);

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(2) simultaneous saccharification and fermentation (SSF). The fermentation method used is assessed in terms of cell growth, reducing sugar consumption and bioethanol production profiles [35]. Generally, SHF and SSF are employed in the fermentation of algal hydrolysate. In SHF, enzyme production, biomass hydrolysis, hexose and pentose fermentations are carried out in separate reactors [89]. The hydrolysis pretreatment can convert the substrate into glucose in the first reactor. In the second reactor, fermenting bacteria or yeast is inoculated which transform the glucose into ethanol. The yeast Saccharomyces cerevisiae is the foremost player in this approach. The downside of this process is the inhibition of cellulases which occurs as a result of accumulation of glucose and cellobiose during hydrolysis [96]. Also, the usage of separate reactors make this technology a costlier affair. SSF is undertaken in order to tackle with the limitations of SSH. Both the saccharification and fermentation can be performed simultaneously in the same reactor in SSF. The pioneer study in this field was done by Takagi et al. [97]. The problem of end product inhibition is overcome with this approach, hence high yield of ethanol is obtained. Coupling the two processes also reduces the capital cost. However, the temperature optima for enzyme and yeast may differ and cause various effects on the growth of micro-organisms. An array of factors including pH, temperature, nutrient levels, concentration of toxic substances together with process variables such as the use of batch or continuous reactors influence the yield of bioethanol [98]. Both the temperature and pH produces a significant effect on the bioethanol yield which further depend upon the optimum reaction condition of the micro organism carrying out the fermentation process. The yeast Saccharomyces cerevisiae requires an optimum temperature of 40°C and pH of 4.5 for the fermentation of macroalgae Sargassum sp. whereas optimum pH of 5 and temperature of 50°C is needed for extracting ethanol from Gracilaria verrucosa biomass [52, 99]. Another yeast Saccharomyces bayanus can carry out the fermentation of algae Chlorococum sp. at an optimum temperature of 30°C [27]. As far as process variables are concerned, hydrolysate of Gelidium amansii produced more than 50 times increased bioethanol yields in continuous reactors when compared to batch reactors. In batch reactors, inhibitory compounds accumulate that leads to decrease in bioethanol yield [100].

5.7 Distillation The ethanol obtained after the fermentation reaction is purified using distillation, which is still the most widely used method for ethanol purification

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185

irrespective of higher energy consumption [101]. The main elements of the distillation unit include: 1) feed (ethanol to be purified), 2) energy source (generally steam) 3) overhead, 4) bottom product and 5) condenser [2]. Distillation removes water and other impurities from the fermented product thereby purifying the ethanol up to 95%. However, some modification in the distillation system is the need of the hour in order to obtain high purity ethanol with less consumption of energy. Based on the volatility of components, two phases are formed. The more volatile components will be in vapor rich region and the less volatile in the liquid rich region. At the completion of the process, the end product is drawn off from the system and blended with the petrol [102].

5.8 Manipulation of Algal Biomass Even though commercial algal cultivation is not a new approach, we are still far from utilizing its full processing capabilities. The algae manipulation through metabolic engineering and genetic methods provide a ray of hope for improving the algal processing capabilities. These approaches have been applied to algae during the past few years with the view to increase the carbon dioxide fixation and carbohydrate productivity for efficient production of bioethanol [103]. In metabolic engineering, the biochemical composition of algae is altered by changing the culture conditions. One such strategy is through macroelement (nitrogen, sulfur or phosphorus) limitation that enhances the starch accumulation in the algae. For example, when microalgae are allowed to grow under the conditions of nitrogen depletion, they start transforming the proteins or peptides to lipid or carbohydrates as energy reserve components [5]. The metabolic engineering is successfully applied to the algal genera Chlorococcum, Tetraselmis, Anthrospira platensis, Synechococcus, Scenedesmus and Dunaliella [18, 20, 32, 65, 104, 105]. As a substitute to metabolic engineering, genetic transformation allows direct control over the algae cellular machinery through introduction of transgenes [106]. Though the algae have high genetic diversity, cyanobacteria have received extensive interest in the field of genetic manipulation. This is attributed to the short life cycle and the transforming ability of cyanobacteria. Synechococcus sp. and Synechocystis sp. are the examples of species which are genetically transformed with powerful molecular toolboxes [107, 108]. Table 5.5 lists the cyanobacteria that have been engineered for the production of bioethanol. The algal photosynthesis mainly depends on the Calvin cycle in which ribulose-1,5-biphosphate combines with CO2 and produces two molecules

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Table 5.5 Genetically transformed cyanobacteria species. Gene expressed using genetic transformation

Citation

Algae

Strain

Promoter used

Synechococcus sp.

PCC 7942

rbcLS

pdc, adh II

[107]

Synechococcus elongatus

PCC 7942

Ptrc

PduP

[109]

PCC 7942

rbcL

ictB, ecaA, acsAB

[110]

PCC 6803

psbA2

pdc, adh II

[108]

PCC 6803

Prbc

pdc, adh2

[111]

Synechocystis sp.

of 3-phosphoglyceric acid (3-PGA) which is further used for the synthesis of glucose and other metabolites [102]. Through genetic engineering, ethanol producing genes, i.e., pyruvate decarboxylase (PDC) and alcohol dehydrogenase (ADH) are introduced into the algae for redirecting 3-PGA to ethanol. The rationale behind this is to increase the conversion of fixed carbon to ethanol and to improve the growth conditions of micro-organisms [112]. Besides algae, the fermenting micro-organism E.coli, Zymomonas mobilis and Saccharomyces cerevisiae are also genetically modified. The genetic modification of these micro-organisms mainly increases their tolerance to ethanol. In addition, it also improves the yield of ethanol and increases the ability of yeasts to utilize different sugars such as hexoses and pentoses for bioethanol synthesis [113].

5.9

Pros and Cons of Bioethanol Production from Algae

The algae have certain features which make them sustainable and environmentally sound feedstock. These features are: 1) Algae have simple cell division cycle enabling them to complete their life cycle in a short time, which results in quick production of high biomass yield; 2) Algae can grow in brackish water, salt water and wastewater and do not compete with food crops; 3) Algae have the ability to remove chemical and organic contaminants and heavy metals so they are used in conjunction with wastewater treatment processes [114]. The microalgae Scenedesmus obliquus can significantly remove nitrogen and phosphorus

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187

from urban wastewater [115]; 4) Algae have the inherent ability to survive under stressful conditions. When subjected to physical stress or nutrient limitation environment, they accumulate more carbohydrates and hence increase the bioethanol yield; 5) Certain species of algae, Scenedesmus obliquus and Chlorella kessleri are known to mitigate CO2 from the power plants [116]. Besides the advantages, algae have certain shortcomings. The principal limitations of algae as a fuel includes, 1)  Microalgal farming is costly and complicated compared to conventional agricultural practices because of the small size of microalgal cells; 2) Many algal species can produce toxins ranging from simple ammonia to physiologically active polypeptides and polysaccharides at different stages of their life cycle [117]; 3) Seaweeds being complex in their chemical composition require few microbial strains for carrying out the fermentation of seaweed biomass to ethanol, which is indeed a costly process.

5.10 Conclusions The production of bioethanol from algae is considered to be an effectual approach for renewable bioethanol production. Several microalgal and macroalgal species have been explored for the efficient production of green fuel bioethanol. The pretreatment methods, however, affect the algal biomass productivity. Hence, it is the indispensable criterion to optimize the pretreatment methods and grow the algae at optimal environmental conditions. In view of this, environmental conditions such as light intensity, temperature, CO2 supply, nutrient uptake and algal harvesting require special attention. Moreover, the selection of an appropriate cultivation system for algal growth plays a pivotal role for mass production of algal biomass. In addition, the genetic manipulation of the selected algal strains for increasing their survival in adverse environmental conditions, together with the genetic engineering of fermenting micro-organisms for improving fermentation of algal biomass, are essential for effective bioethanol production.

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64. Harun, R., Liu, B., Danquah, M.K., Analysis of process configurations for bioethanol production from microalgal biomass, in: Progress in Biomass and Bioenergy production, pp. 395–408, InTech Open, Europe, 2011b. 65. Markou, G., Angelidaki, I., Nerantzis, E., Georgakakis, D., Bioethanol production by carbohydrate-enriched biomass of Arthrospira (Spirulina) platensis. Energies, 6, 3937–3950, 2013. 66. Soong, P., Production and development of Chlorella and Spirulina in Taiwan, in: Algae Biomass, A. Vonshak (Ed.), pp. 97–113, Elsevier: North Holland Biomedical Press, Amsterdam, 1980. 67. Belay, A., Mass culture of Spirulina outdoors—The Earthrise farms experience, in: Spirulina platensis (Arthrospira): Physiology, Cell-Biology and Biotechnology, A. Vonshak (Ed.), pp. 131–158, Taylor and Francis, London, 1997. 68. Zijffers, J.W., Janssen, M., Tramper, J., Wijffels, R.H., Design process of an area-efficient photobioreactor. Mar. Biotechnol., 10, 404–415, 2008. 69. Wang, B., Lan, C., Horsman, M., Closed photobioreactors for production of microalgal biomasses. Biotechnol. Adv., 30, 904–912, 2012. 70. Ho, S.H., Chen, C.Y., Lee, D.J., Chang, J.S., Perspectives on microalgal CO2emission mitigation systems: A review. Biotechnol. Adv., 29, 189–198, 2011. 71. Singh, R.N. and Sharma, S., Development of suitable photobioreactor for algae production—A review. Ren. Sustain. Energy Rev., 16, 2347–2353, 2012. 72. Ugwu, C.U., Aoyagi, H., Uchiyama, H., Photobioreactors for mass cultivation of algae. Bioresour. Technol., 99, 4021–4028, 2008. 73. Kumar, P., Barrett, D.M., Delwiche, M., Stroeve, P., Methods for pretreatment of lignocellulosic biomass for efficient hydrolysis and biofuel production. Ind. Eng. Chem. Res., 48, 3713–3729, 2009. 74. Hendriks, A.T.W.M. and Zeeman, G., Pretreatments to enhance the digestibility of lignocellulosic biomass. Bioresour. Technol., 100, 10–18, 2009. 75. Hopkins, T.R., Physical and chemical cell disruption for the recovery of intracellular proteins, in: Purification and Analysis of Recombinant Proteins., pp. 57–84, CRC Press, New York, 1991. 76. Lee, S.J., Kim, S.B., Kim, J.E., Kwon, G.S., Yoon, B.D., Oh, H.M., Effects of harvesting method and growth stage on the flocculation of the green alga Botryococcus braunii. Lett. Appl. Microbiol., 27, 14–18, 1998. 77. Geciova, J., Bury, D., Jelen, P., Methods for disruption of microbial cells for potential use in the dairy industry—A review. Int. Dairy J., 12, 541–553, 2002. 78. Naghdi, F.G., Gonzalez, L.M.G., Chan, W., Schenk, P.M., Progress on lipid extraction from wet algal biomass for biodiesel production. Microb. Biotechnol., 9, 718–726, 2016. 79. Amarni, F. and Kadi, H., Kinetic study of microwave-assisted solvent extraction of oil from olive cake using hexane: Comparison with the conventional extraction. Innov. Food Sci. Emerg. Technol., 11, 322–327, 2010. 80. Khanal, S.K., Grewell, D., Sung, S., Van Leeuwen, J., Ultrasound applications in wastewater sludge pretreatment: A review. Crit. Rev. Environ. Sci. Technol., 37, 277–313, 2007.

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6 Development of Acid-Base-Enzyme Pretreatment and Hydrolysis of Palm Oil Mill Effluent for Bioethanol Production Nibedita Deb, Md. Zahangir Alam*, Maan Fahmi Rashid Al-khatib and Amal Elgharbawy Bioenvironmental Engineering Research Centre (BERC), Department of Biotechnology Engineering, Faculty of Engineering, International Islamic University Malaysia, Kuala Lumpur, Malaysia

Abstract Nowadays, palm oil mill effluent (POME) is the main water stream pollutant in Malaysia. This paper reviews the development in the methods of bioethanol production from POME by acid-base-enzyme pretreatment and hydrolysis processes. The methods currently used to treat POME are inefficient in terms of either cost or environmental preservation. Several techniques of pretreatment of POME are discussed, where the structure of POME is broken down and made more accessible to the cellulase enzymes. Generally, hydrolysis process is used as a mild acid-base-enzyme for monomeric sugar production from POME. The acid-base-enzyme consists of H2SO4, NaOH and cellulase enzymes to run the hydrolysis process. In this paper, different strategies are also described for hydrolysis processes, including concentrate acid, dilute acid and enzymatic hydrolysis. In addition, the production of bioethanol by fermentation of sugars obtained from the hydrolysis process is discussed and it points out some key properties that should be targeted for cost-effective and innovative pretreatment processes. Keywords: POME characterization, pretreatment, enzymatic hydrolysis, bioethanol

*Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (197–218) © 2019 Scrivener Publishing LLC

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6.1 Introduction The palm oil industry is one of the largest agricultural industries in Malaysia with average crude palm oil production of more than 20.35 million per annum [1]. However, production of such a huge amount of crude palm oil has produced an even larger amount of palm oil mill effluent (POME) [2]. POME is an extremely polluting wastewater with high biochemical oxygen demand (BOD) which is in the range around 21,500-28,500 mg/l and chemical oxygen demand (COD) which is in the range of about 44,300–70,900 mg/l that can cause naturally pollution of water resources and severe pollution to the atmosphere [3]. Besides that, POME was identified as a potential source to produce renewable bioenergies such as biomethane and biohydrogen through anaerobic digestion [4]. Hence, a combination of wastewater treatment and renewable bioenergies production would be an extra advantage in the palm oil industry. This approach should be applied directly to ensure sustainability according to the world’s sustainability concept and that palm oil is generated in an environmentally friendly manner [5]. POME is the wastewater discharged from the sterilization process, crude oil clarification process and cracked mixture separation process [6]. POME generated a large amount of ethanol and methane gas from anaerobic process. This process has 21 times the global warming potential (GWP) compared to the additional gasses. However, it might be solved by utilizing it as fuel for power generation and cogeneration. Although it has a large amount of potential, actuality of established technologies, convenience of knowledge and reasons, biomass recycling use is still in a very early stage in Malaysia and especially for bioethanol it is in a preliminary process [7]. The POME characteristics depend on the quality of the palm oil production processes and raw material in palm oil mills. The crude palm oil (COP) extraction from FFB needs massive amounts of water. It has been estimated that approximately 5-7.5 tonnes of water are essential for creating crude palm oil which is around 1 tonne, and more than 50% of the water ends up as POME [8]. Three major processing operations responsible for producing the POME have been categorized. Sterilization of FFB, shell hydrocyclone contributes, hydrocyclone separation of a cracked mixture of kernel and simplification of the extracted CPO account for about 4%, 36%, and 60% respectively of the POME in the mills. It has been estimated that in Malaysia about 53 million tonnes of POME are being produced every year based on palm oil production in 2005 (14.8 million tonnes). It has been estimated that about 0.5-0.75 tonnes of POME will be discharged from mill for every tonne of fresh fruit bunch [9].

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Wastewater treatment facility is among the most essential subsidiaries in the palm oil mill system [10]. This is because the facility is to treat POME that produces a large volume during the production of CPO. Due to the chemical and physical properties of POME, the anaerobic treatment is the most efficient system used for the wastewater plant in the early stage. The existing systems fulfill the necessity of the palm oil mill operators to carefully release the treated POME. That treatment process discharges one of the greenhouse gases (GHG), CH4 in the air as the result of anaerobic digestion of POME. Usually, POME composition is based on the season, raw matter quality and the individual processes being conducted at any given time. Commonly, POME is low in pH, which is the range approximately 4-5, as a result of the organic acids generated in the process of fermentation. It also contains a huge amount of total solids and oil and grease which are almost 40,500 mg/l and 4,000 mg/l. POME contains dissolved elements such as carbohydrate, nitrogenous compounds, high concentration of protein, lipids and minerals, which can be converted into valuable materials by the microbial processes [11]. The effluents from palm oil mill can cause considerable environmental problems, if discharged untreated [12]. For that reason, the challenge of converting POME into an eco-friendly waste needs an efficient treatment and effective disposal technique. POME is produced in large amounts annually. Studies have been conducted to treat this waste in a more economical way than what is being used currently. Having a high lignocellulosic content and carbohydrates, producing ethanol from POME is one of the best ways to treat it. In order to ferment POME to ethanol, hydrolysis of the lignocellulosic material has to be done. Hydrolysis are commonly done using concentrated acid, dilute acid or enzymes. Concentrated acid gives high yields, but requires large amounts of acid; this poses problems when it comes to neutralization and sugar recovery, in addition to it being harmful to the environment. Dilute acid reduces acid consumption, but the drawback here is the potential of degrading the product because of the high temperatures used. Enzymatic hydrolysis on the other hand is simply too expensive. Ligocellulose is a major constituent of the oil palm tree and hence of POME. Ligocellulose contains lignin, hemicellulose and cellulose, all of which have a high biotechnological importance due to high energy content. If prepared appropriately they can be a good substrate for the growth of microorganism which gives off products of high value. For example, we can say ethanol, which is a by-product of microbial fermentation of starch. The main sources of energy nowadays are coal, petroleum crude oil and natural gas, and the consumption of these fossil fuels grows annually [13].

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Only recently have people come to comprehend the crucial need to develop alternative sources of energy [14]. This is mainly due to factors like diminishing oil reserves in the land, the lack of technology to retrieve oil from ocean bed reserves, the concern over global climate change and the constantly increasing fuel price. As a result, people have slowly started to move from the scarce, unclean fossil fuels to renewable, environmentally friendly biofuels. Bioethanol is not a fossil fuel; it is actually used as a renewable energy source extracted from biomass product via fermentation of starches [15]. Normally, day by day the consumption of fuel is increasing, and the result of such use of fossil fuels can be a greater scarcity of energy sources. Bioethanol is a solution for this phenomenon. Mostly, a blend of bioethanol and gasoline can be used in the traditional gasoline engine to reduce the pressure on gasoline. The biofuels are also a form of chemical organic carbon. But bioethanol emits less carbon than fossil fuels [16]. Moreover, the bio conversion process to generate ethanol does not produce hazardous material [17]. Consumption of bioethanol as a new technology is increasing day by day and it has been researched and developed as well. Generally bioethanol is produced from the fermentation of sugar by using yeast as a microbial element. The extracts are derived from crops such as corn, maize, sugarcane, sorghum, cassava and palm oil. This fermentation process is a relatively steady process, so it is important for industrial use to make it faster. This paper examines bioethanol as produced by the acid-base-enzyme pretreatment and hydrolysis of POME, because it is cheaply and readily available in large amounts in the country. The high costs of the enzyme problem are solved by using enzymes that are locally produced from POME [18]. If bioethanol production from POME is applied on a large scale, it has the potential of solving the economic and environmental problems associated with this wastewater [19].

6.2 Biomass Energy According to Overend [18], biomass is the vegetable matter produced by the action of solar energy on plant photosynthesis, which builds plant cell walls and other components such as seeds and fruits. Bio energy is the energy resulting from the use of biomass as a feedstock to produce various types of energy like heat, electricity, liquid and solid fuels. The basic concept of biomass as a source of energy involves the capture of energy from

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201

insulation along with carbon dioxide from the atmosphere in the biomass by fixing the gaseous CO2 into organic carbon.

CO2+H2O+light+Cholophyll → (CH2O) +O2

(1)

Where, CH2O is the primary organic product. For every gram mole of carbon fixed 470 kJ (112 kcal) is absorbed. This shows the great potential of biomass as an energy sources. This biomass is then harvested and converted to biofuels or synfuels or otherwise used to directly produce heat by combustion. Alternatively, the biomass can either be harvested for foodstuff, feed, fiber and useful materials or it can be left to take its natural life cycle. The waste produced by consumption or the decomposing matter will be converted after a long time to fossil fuels. The disposed waste can also be converted directly to synfuels. Another pathway for this biomass is to grow natural hydrocarbons; certain species are rich in hydrocarbons like the rubber tree. This is a good way to store big amounts of energy into fixed carbon [8]. People became aware of the serious issues associated with the use of fossil fuels by the mid-90s, with the biggest concern being global warming. Emissions resulting from the combustion of fossil fuels like CO2, CH4 and NOx have the potential of trapping solar radiation reflected from the earth’s surface. With the increasing concentration of these gases in the atmosphere, a significant increase in the ambient temperature has been recorded worldwide. This increase in temperature can cause warming of the upper layers of the oceans and an increase in sea level, a shift of the agricultural zones and a complex effect on polar ice caps which are not well understood yet [8]. Another environmental issue associated with the combustion of fossil fuels is acid rain which results from SOx that are emitted upon the combustion of sulfur oxide containing fuels. Acid rain has the potential to damage materials and buildings, in addition to affecting the growth of biomass.

6.3 Palm Oil Mill Effluent (POME) The palm oil industry is the economic backbone of Malaysia. In 2012, about 18.9 million tonnes of crude palm oil were produced and 17.5 million tonnes were exported [20]. Regularly, large amounts of POME are produced in the process of producing palm oil. In Figure 2.1, generation of POME is detailed by the flow diagram.

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Sterilization

Stripping

Sterilizer condensate

POME

Mulching

Empty fruit bunches Fibre for boiler fuel

Digestion Nuts

Nut cracker

Clarification tank Hydrocyclone

Sludge

Oil Kernel

Separator

POME

Shell for boiler fuel

Centrifuge purification

Vacuum Oil POME Storage

Figure 6.1 Flow diagram of a palm oil mill [21].

In the beginning of the palm oil industry, the rivers were capable of absorbing the pollutants without major effects being noticed. However, given the size of the industry nowadays, the oxygen depleting potential of POME is 100 times that of domestic sewage. The fresh fruit bunches (FFB) are harvested and transported to the palm oil mill for oil extraction. Each FFB contains hundreds of fruits containing a pericarp that carries the oil inside. The oil extraction in the mill starts with steam sterilization of the FFB at conditions of 3 bars, 140°C for 75-90°min. This prevents free fatty acid formation due to enzyme action, facilitates stripping and prepares the fruit for subsequent processing. The condensate of sterilization steam is a major part of the wastewater [22]. Then, a rotary drum-stripper is used to detach the fruits from the fruit bunches. The fruits are passed through the bar screen of the stripper and are collected and discharged into a digester where they get mashed by the rotating arms. In this step, the oil bearing

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cells are broken, and twin-screw presses are used to press out the oil under high pressure. A way to improve oil flow here is the addition of hot water, which can be separated later in the separation and purification system [22]. Figure 6.1 shows a simplified process flow diagram to produce palm oil. The crude palm oil (CPO) from the screw presses has 35-45% palm oil, 45-55% water and fibrous materials. The oil is separated in a clarification tank and then passed through a high-speed centrifuge and a vacuum dryer. In the screw presses, a cake is produced that consists of moisture, oily fiber and nuts. The fiber and nuts are separated and the former is used as boiler fuel, while the latter is processed further to separate kernels and shells. The discharge from this process forms the source of the wastewater [16]. The wet palm oil milling process is the most common way of extracting palm oil from fresh fruit bunches (FFB), typically in Malaysia. It involves several stages in which a huge amount of water and steam are required for washing and sterilizing. Thus, this has resulted in a huge amount of wastewater generated from palm oil mill or better known as POME.

6.4 Pome Characterization POME is derived from three types in palm oil production; clarification that gives off 60%, sterilization 36% and hydrocyclone 4% [11]. POME contains cell walls organelles, short fibers, and carbohydrates like cellulose, hemicellulose and lignocellulosic materials. The raw POME is acidic and hot. It is characterized by high amounts of total suspended solids, BOD, COD and oil and grease. Table 6.1 shows the characteristics, metal and chemical composition of POME, respectively.

6.5 Pretreatment Enzymatic hydrolysis has the potential to become the primary means of hydrolysis of biomass to produce bioethanol, one of the main factors for this being the high theoretical yield [8]. That being said, further research is yet to be done to realize this yield and this is mainly due to the physicochemical, structural and compositional factors of lignocellulosic materials. It is here where pretreatment comes into play. Lignin and hemicellulose shield the cellulose, making it hard for the cellulase to access the cellulose efficiently [23]. Add to that the crystalline structure of cellulose, which makes the access of cellulase to the cellulose fibers inefficient. Therefore, the pretreatment aims at removing lignin and hemicelluloses

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Table 6.1 Characteristics of raw POME. Parameter

Average

Metal/chemical

Average

pH

4.7

Phosphorous

180

Oil and grease (mg/l)

4000

Potassium

2270

BOD (mg/l)

25000

Magnesium

615

COD (mg/l)

50000

Calcium

439

TSS (mg/l)

40500

Boron

7.6

TDS (mg/l)

50100

Iron

46.5

TVS (mg/l)

38500

Manganese

2.0

Total sugar (g/l)

3.00

Copper

0.89

Reducing Sugar (g/l)

11.26

Zinc

2.3

Cellulose (%)

39.56

Protein (%DM)

11.0

Hemicellulose (%)

23.33

Fiber (%DM)

7.9

Lignin (%)

25.02

NDF

28.2

Ammonical Nitrogen (mg/l)

35

Ether Extract (%DM)

29.2

Total Nitrogen (mg/l)

750

Ash (%DM)

8.7

Source: Alam et al. [24]; Wong et al. [25]; Fanimo et al. [10]; and Brown et al. [13].

wrapping the cellulose, shaking the crystalline structure to give cellulase more access, and finally making the substrate more porous. In order to achieve those aims efficiently, several parameters should be observed like sugar yield from pretreatment or further hydrolysis, the disruption or degradation of the product, the formation of inhibitory products and finally the cost [26]. Pretreatment of lignocellulosic material has been done using many different methods like physical-physiochemical methods, chemical methods and biological methods. It has been suggested that each raw material requires a different pretreatment; therefore, a general procedure cannot be established.

6.5.1 Physical and Physicochemical Pretreatment Physical methods include mechanical commination and pyrolysis [27]. In the former, waste material is chipped, grinded and milled to reduce cellulose crystallinity, whereas in the latter, mild acid hydrolysis has been found

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to produce reducing sugars from cellulose with 80-85% conversion [28]. In physicochemical methods, steam explosion, afirmonia fiber explosion and CO explosion are utilized. In steam explosion, the biomass are chipped and exposed to high pressured steam, and then the pressure is reduced. This sudden change in pressure causes explosive decompression. And because high temperatures are used, hemicellulose and lignin are degraded. This method is quite efficient in terms of yield. However, its use is limited by factors like xylan fraction destruction, production of inhibitory products and incomplete disruption of lignin bonds to the cellulose and hemicellulose.

6.5.2 Chemical Pretreatment Several chemicals are used to pretreat lignocellulosic materials. Ozone is a good candidate for lignin degradation. It has been improved the sawdust hydrolysis yield up to 57% following an ozone retreatment. Ozone pretreatment is very efficient in terms of yield, it operates under room temperature and pressure and it does not produce toxic compounds. The only disadvantage of this process is high cost resulting from high ozone amounts used. Another chemical pretreatment involves the use of bases; e.g., ammonia and sodium hydroxide. Bases work by saponification of ester bonds between xylan and other hemicelluloses and lignin. Consequently, the lignin structure is broken and the crystallinity and polymenzation are reduced. In addition to increasing the surface area alkaline treatment can function at ambient temperature and pressure, but required very long retention times. Another limitation for alkaline pretreatment use is the production of irrecoverable salts that are precipitated in the product. The most popular pretreatment uses acids like H2SO4 and HCL. Essentially, acid pretreatment can be divided into concentrated and dilute acid hydrolysis [29]. In concentrated acid hydrolysis, high acid concentrations require a corrosion resistant material, which adds to the cost of hydrolysis. And to make the process more cost-effective, acid recovery is required [30]. Dilute acid hydrolysis on the other hand is a more favorable option since it is effective and not very costly. It efficiently hydrolyzes hemicellulose especially xylan to release xylose and other sugars [31]. In terms of process economy, this is advantageous since xylan accounts for up to one-third of the carbohydrates content in many lignicellulosic materials. Acids like H2SO4, HCL and H3PO4 have been used in dilute acid pretreatment, the most widely used H2SO4. An effective factor in this process is temperature; it divides the hydrolysis into two general categories:

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high temperature (above 160°C) for low solid substrate loadings, and low temperature (below 160°C) for high solid substrate loadings. In pretreated POME solids using varying concentrations of H2SO4, the optimum concentration was found to be 0.5% (v/v). When fungal enzymes are used, acid pretreatment is favored since those enzymes operate at low pH values [32].

6.5.3 Biological Pretreatment Biological pretreatment uses microorganisms that produce enzymes to extensively degrade lignin from the substrate. The most widely used microorganisms are brown, white and soft-rot fungi [33]. The main advantages of this pretreatment are that it is environmentally friendly with minimum waste production, and low energy consumption. This comes at the expense of very slow hydrolysis [26].

6.6 Hydrolysis Although the previously used methods for treatment of POME are satisfactorily efficient, they give rise to some problems like requiring large land areas to construct the treatment plant, not being very cost effective and emitting by-products that are harmful to the environment like methane. Therefore, the palm oil industry is yet to find a balance between maintaining economic growth and preserving the environment. The optimum approach is to achieve the converting of POME to a useful end product using an environmentally friendly method, and towards the production of bioethanol using microbial conversion. The first step in the production of bioethanol is breaking the POME into simple sugars that can be processed by yeast. Hence, the most important factor in the process is the hydrolysis of the raw POME. Hydrolysis is the process in which glucosidic bonds the links the glucose molecules together in the large cellulose polymer are cleared by the addition of a water molecule [34].

6.6.1 Concentrated Acid Hydrolysis The hydrolysis of lignocellulosic materials through concentrated sulphuric acid or hydrochloric acid to produce fermentable sugars was first discovered by Braconnot in 1819 [35]. Concentrated acid hydrolysis is a powerful

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method; it gives high yields of sugars. Another advantage is that it can operate at low temperatures [36]. That being said, this method utilizes very high concentrations of acids, which makes it highly corrosive. Hence, expensive alloys or non-metallic materials are required. Another factor that makes this method ineffective is that the acid recovery process is energy demanding [34]. Moreover, the neutralization process produces large amounts of gypsum. In addition, stringent environmental laws limit the application of such concentrated acids. Therefore, the potential commercial interest in this process has been greatly reduced [36].

6.6.2 Dilute Acid Hydrolysis Dilute acid hydrolysis is the most commonly used chemical hydrolysis method. This method is usually carried out in a batch process with retention time of a few minutes. The main advantage of this process is the reduced acid use. However, this comes at the expense of energy use since high temperatures are required to reach optimum yields. According to [13], high temperatures cause oligosaccharides from lignocelluloses to decompose, which reduces the simple sugar yield to 55-60%. The decomposition also releases microbial toxins like acetic acid and furfural. Furthermore, like concentrated acid hydrolysis, this method requires corrosion resistant equipment.

6.6.3 Base Hydrolysis Base pretreatments increase cellulose digestibility and they are more effective for lignin solubilisation than cellulose and hemicellulose solubilization by acid or hydrothermal processes. This procedure can be worked at surrounding temperature and times going from seconds to days. Moreover, probable loss of fermentable sugars and creation of unexpected compounds must be thought about to advance the pretreatment conditions [37]. More suitable, alkaline pretreatments are potassium, calcium and ammonium hydroxides. On the other hand, lime pretreatment increases the crystallinity index by removing amorphous substances like lignin. Compared to NaOH or KOH pretreatments, lime required less safety and is cost effective. It is also easily separated from hydrolysate by reaction with CO2. Furthermore, no furfural or 5-hydroxymethylfurfural (HMF) was detected in hydrolyzates obtained with alkaline peroxide pretreatment, which favours the fermentation step in an ethanol production process [19].

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6.6.4 Enzymatic Hydrolysis Enzymatic hydrolysis of biomass are the most commonly used saccharification methods. The utilization of celluloses and hemicelluloses present in the feedstock is enhanced by this method [38]. The enzymes utilized function are a series of steps to break down the polymeric cellulose and hemicellulose by hydrolyzing the glycosidic bonds, leaving monomeric sugars ready for fermentation [38]. The bond cleavage is finished with the synergistic activity of endoglucanases and exoglucanases [13]. Following this reaction, soluble intermediates like elluloligosaccharides and cellobiose are hydrolyzed to produce glucose by β-glucosidase. Amongst the hydrolysis processes, enzymatic hydrolysis gives the highest sugar yield [13]. The cellulase enzymes used are highly specific, and they do not degrade the pentoses produced during prehydrolysis [19]. The biggest challenge that is yet to be overcome is the actual yield of sugars compared to the theoretical one. It has been demonstrated that the actual yield could be less than 20% of the theoretical. This problem has triggered a lot of research aiming at developing pretreatment methods than can maximize the sugar yield from the hydrolysis. And although the quantities of enzymes required compared to the final production of fuel can be ineffective in terms of cost, methods can be developed to produce low-cost enzymes, in addition to the advantage pretreatment brings. Furthermore, compared to the other hydrolysis processes, enzymatic hydrolysis has a lower utility cost because of the moderate conditions of pH and temperature required for the enzymes to function efficiently. This reduces the expenses of energy provision, corrosion resistant materials and maintenance incurred by the other hydrolysis methods [38].

6.6.5 Cellulase Enzymes Hydrolysis Cellulase is a suite of enzymes, produced by bacteria, fungi and protozoans, which work collectively to hydrolyze cellulose [39]. In catalyzing cellulolysis, at least three types of cellulases are involved. In this type of enzymes, endoglucanase (EC 3.2.1.4), this type works on the cellulose fiber to create free chain-ends by attacking regions of low crystallinity. However, Exoglucanase (EC 3.2.1.9.1) process works on the previously created free chain-ends and removes cellobiose units for further simplification. On other the hand, B-glucosidase (EC 3.2.1.21), which finally produces glucose from cellobiose hydrolysis [40].

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Cellulose enzymes catalyze the reaction in which water is added to glucan chains to release the monomers; glucose molecules. The overall reaction can be expressed by the following Equation 2:

(C6H12O5)n + nH2O → nC6H12O6

(2)

Looking at the stoichiometry of the reaction, 180 mass units of monomeric sugar are produced for each 162 mass units of glucan and 18 mass units of water. That accounts for 11.1% sugar gain. An advantage of cellulase enzymes is their high specificity which assures that the final product will always be glucose, despite the presence of cellobiose or cellotriose as an intermediate as was demonstrated. Similarly, hemicellulose can be hydrolyzed by water addition reaction catalyzed by enzymes known collectively as hemicellulases. In hemicellulose, the hexose sugars glucose, galactose and mannose follow the same stoichiometry as cellulose. However, the pentosesarabinan and xylan follow a different stoichiometry than can be described by the following Equation 3:

(C5H8O4)n + nH2O → nC5H10O5

(3)

Whereby 132 mass units of sugar form 150 mass units, making the gain equal 13.6%. Herhicellulases are generally classified under cellulases. The hydrolysis of hemicellulases is important because it gives cellulases access to cellulose. Pretreatment plays an important role in hydrolyzing hemicellulose since it can be done at mild conditions. In other words, the pretreatment of hemicellulose supplemented by cellulase action provides a good treatment that helps improve cellulose hydrolysis.

6.7 Fermentation Process Fermentation is the process toward getting energy from the oxidation of biological compounds, like carbohydrates and utilizing an endogenous electron acceptor, which is normally a biological compound. Whereas, the electrons are given to an exogenous electron acceptor for respiration like oxygen, by means of an electron transport chain. Fermentation does not really need to be completed in an anaerobic field. For example, with the existence of copious oxygen, yeast cells extremely raise fermentation to oxidative phosphorylation, providing sugars are quickly accessible for utilization. Sugars are the most widely recognized substrates of fermentation and characteristic examples of fermentation items are hydrogen, ethanol and lactic acid. Besides, more exotic compound was created by fermentation like acetone and butyric

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acid. Yeast does fermentation in the generation of ethanol in wines, beers and other mixed beverages, together with the creation of substantial amounts of carbon dioxide. Fermentation happens in mammalian muscle during times of exceptional practice where oxygen supply gets to be restricted, causing the making of lactic acid. On the other hand, the carbon-based materials which are animal and plant waste, are converted into energy sources by biological processes such as microorganisms and enzyme [2, 41]. Different lignocellulose bioethanol conversion process has allowed the selection and capacity of feedstock that can be bioconverted to increase gradually. Currently, materials in the feedstock derived from animal and plant waste include municipal solid waste (MSW), sewage sludge, construction materials, tires fabric, paper [9] and POME [42].

6.8 Bioethanol Generally, the process of bioethanol production consists of two types: fermentation ethanol and synthetic ethanol. Fermentation ethanol is created from biomass through hydrolysis and sugar fermentation. The biomass is pretreated by acid hydrolysis followed by fermentation. In the process of biomass to bioethanol, acid-based enzymes are utilized to catalyze this reaction. In contrast, fermentation can be a series of the biocatalyzed reaction which converts simple sugar to ethanol. This reaction is also aided by yeast that fed on simple sugars. The main advantages of the enzyme are produced by the local [2]. Another similar study as the combined with acid pretreatment and enzymatic hydrolysis has also provided development of ethanol content from durian seed [41]. The authors also acknowledged that acid pretreatment contributed a better form of sugar for the yeast to consume the substrate than the one without it. The better form of glucose is essential since Saccharomyces cerevisiae is only capable to ferment glucose, another form of sugars are capable to be fermented by other types of microorganisms [2, 41]. By combining both acid pretreatment and enzymatic hydrolysis, it is normal to have better glucose yield and ethanol produced due to the combination of both favorable characteristics, compared to the one with only acid pretreatment. From biomass conversion of the bioethanol, mainly from empty fruit bunch (EFB) and raw POME applications a simple previously minimal treatment of waste from palm industry. As a result of EFB and raw POME, the competition with food-based simple materials can be reduced, therefore protecting the cost of food product [43]. However, bioethanol is synthesized from lignin, hemicellulose and cellulose, which create from the different bases of biomass. NaOH pretreated sample was hydrolyzed with H2So4

Hydrolysis of Palm Oil Mill Effluent

Raw POME

Acid

Treated POME

Pre-treatment

Hydrolysis

Base

Enzyme

211

Hydrolysate Ethanol Recovery Bioethanol

Fermentation Inoculum

Figure 6.2 Block diagram of bioethanol production.

hydrolyzed with a different set of acid concentration. The fermentation with saccharomyces cerevisae was carried out in a single reactor for anaerobic stage. The authors also recommended the ethanol concentration did not show a significant effect of the inoculums concentration. On the other hand, the yeast concentration is increased dramatically the fermentation duration is decreased. But the concentration of the glucose is increased gradually the ethanol concentration is also increased. Figure 6.2 shows that the block diagram of the bioethanol production from POME. Forestry and agricultural waste have the potential of becoming the first source of bioethanol production since it does not compete with human food material, and it solves the problem of deforestation and land erosion due to planting and cultivating crops for the sole purpose of energy production. Currently, ethanol is mainly produced from the fermentation of sugar and starch [44]. Moreover, bioethanol is used to produce a wide variety of chemicals used in pharmaceuticals, cosmetics, medicine and industry.

6.8.1 Lignocellulosic Bioethanol Forestry and agricultural waste have the potential of becoming the first source of bioethanol production since it does not compete with the human food material, and it solves the problem of deforestation and land erosion· due to planting and cultivating crops for the sole purpose of energy production [45]. Currently, ethanol is mainly produced from the fermentation of sugar and starch. Bioethanol is a high-octane, water-free alcohol. It is a colorless liquid

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with a boiling temperature of 78°C and a freezing temperature of -112°C. It has a mild characteristic odor. The use of ethanol is a fuel for internal combustion engines is not new; it has been blended with petroleum fuels to produce a motor fuel extender for many years. Also, due to its high-octane number, ethanol is used as an additive to enhance octane (reduce engine knocking). Moreover, ethanol helps gasoline burn more completely to reduce harmful emissions by supplying it with dissolved oxygen [8]. Moreover, bioethanol is used to produce a wide variety of chemicals used in pharmaceuticals, cosmetics, medicine and industry. Examples of those chemicals are: acetaldehyde, acetic acid, isopropyl alcohol, vinasse and potassium sulfate.

6.8.2 Bioethanol Production by Fermentation of Sugars Fermentation involves enzymatically catalyzed chemical reactions that yield energy, usually carried out by certain microorganisms under anaerobic conditions. The substrate for those reactions is an organic compound that acts as the electron acceptor. In ethanol production, monomeric hexoses (glucose, mannose and galactose) along with pentoses (xylose and arabinose) will be fermented by facultative anaerobes to yield ethanol and carbon dioxide in equimolar amounts. The reaction can be represented by the following equations (4) and (5):

(C6H10O5)n + nH2O → nC6H12O6

(4)

n(C6H12O6) → 2n CH3 CH2OH + 2nCO2

(5)

The microorganisms used for ethanol production are yeasts. Yeasts are nonmotile cells that propagate asexually by budding. All yeasts belong to three groups of higher fungi: Ascomycetes, Basidiomycetes and Fungi Imperfecti. Saccharomyces cerevisiae belongs to the Ascomycetes. Given its high conversion rate, its ethanol tolerance and its wide public acceptance, Saccharomyces cerevisiae are the most commonly used yeast for alcoholic fermentation [46]. Sugars are needed to be present in monomeric form for Saccharomyces cerevisiae to be able to ferment them as it does not ferment polymers. With ethanol and carbon dioxide being the major products, glycerol, acetic acid, lactic acid, succinic acid and fusel (a mixture of amyl alcohol and isoamyl alcohol) are produced in minor amounts. The fermentation process is exothermal; 900 kJ are released for every liter of produced ethanol [8]. Saccharomyces cerevisiae has high bioethanol yield; according to Jessen, [47] minimum of 1.9 mol ethanol/mol hexose will be produced. The production of ethanol from lignocellulosic material is not as

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easy as the established production from corn or sugar cane. This is mainly due to the structure of lignocelluloses. Hemicellulose hydrolyzates contain xylose and arabinose. Saccharomyces cerevisiae cannot ferment five carbon sugars [48] and cannot ferment arabinose except in the presence of rich media [49]. Genetically engineered Saccharomyces cerevisiae that harbors genes for xylose fermentation and arabinose metabolism has been developed to achieve maximum conversion rate.

6.8.3 Bioethanol Determined by GC/MS from POME Hydrolysate The fermentation of hydrolyzed POME based media bioconverted by the Saccharomyces cerevisiae was utilized as a crude bioethanol sample to be extracted by GC/MS. The crude bioethanol sample was first run through the centrifugation of the centrifuge tube to separate different insoluble suspended solid and matters from the sample. The infiltrate of this step was used as the feed sample by GC/MS. The bioethanol sample was purified by using acetone and exclusion chromatography. After that bioethanol contained was determined by chromatography. The chromatographic profile for bioethanol is shown in Figure 6.3. The analytical results showed that Abundance

1.665

1.1e+07 1e+07 9000000 8000000 7000000 6000000

2.112

5000000 4000000 3000000 2000000 1000000 0 Time–>

5.00

10.00

Figure 6.3 GC/MS total ion chromatograms (TIC) of chemical constituents of the bioethanol extracts of Saccharomyces cerevisiae.

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it was possible to extract the bioethanol with the column of DBWAX, as it was seen that different bioethanol content was recovered at all, because bioethanol was bound to the resin of GC/MS from Figure 6.3. The analytical result revealed that the element was 11.6585% v/v of bioethanol (FCCCD sample run 4). There are many reports of using these columns of bioethanol content [50, 51].

6.9 Conclusion Several pretreatment techniques for POME are described and generally studied to improve bioethanol production. All these techniques make the POME accessible to enzymatic responses. This knowledge is useful to reach an integrated and efficient biomass conversion method to bioethanol. Among the several techniques, chemical pretreatment is presently the most effective, which reach very high yields and include enzyme the most favorable technologies for industrial uses.

6.10 Acknowledgment The authors are grateful to the Department of Biotechnology Engineering for providing the lab facilities and the Research Management Centre, IIUM, for providing the funds.

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7 Technological Barriers in Biobutanol Production Arpita Prasad, Shivani Thakur, Swati Sharma, Shivani Saxena and Vijay Kumar Garlapati* Department of Biotechnology and Bioinformatics, Jaypee University of Information Technology, Waknaghat, HP-173234, India

Abstract The depletion of fossil fuels coupled with ever-increasing pollution has driven people’s attention towards renewable energy sources such as biodiesel and bioalcohols. Among biofuels, biobutanol has emerged as a potential biofuel as a substitute for gasoline. The production of biobutanol comprises different tedious tasks and is hindered by a number of obstacles. Traditionally, biobutanol production by traditional ABE fermentation utilizing Clostridium acetobutylicum species suffers from limitations such as low butanol yields, solvent toxicity issues in the solventogenic phase to bacterial cell wall and cost of pretreatment of lignocellulosic biomass. Hence, the present review puts forth the discussion on existing limitations along with feasible solutions such as metabolic engineering approaches and utilization of solvent-tolerant microbial strains for an enhanced biobutanol production. Keywords: Biobutanol, ABE (acetone-butanol-ethanol) fermentation, oxo-synthesis, aldol condensation strain modification

7.1 Introduction Biofuel is the energy derived from animal material (animal manure, animal fats) and renewable plants (algae, lignin) with the potential of mitigating the upcoming issues of fossil fuel depletion and existing greenhouse gas emissions. Owing to the positive attributes of renewable biofuels, *Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (219–236) © 2019 Scrivener Publishing LLC

219

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overwhelming research is going on towards production technologies of biodiesel, bioethanol and biobutanol. The economics of biofuels production is mostly dependent on the cost of the fermentation substrate. The biomass feedstock cost and biomass availability are the two main decisive factors in establishing a production technology for alternative fuels [1]. Currently, bioethanol and biodiesel are used in the transportation sector. Biobutanol is one of the option, investigated recently and emerging as a potential source of biodiesel and bioethanol. The sources for biobutanol production mainly comprise lignocellulosic waste biomass and non-food agricultural products, which is considered a second-generation biofuel system. Although the biotechnological production of biobutanol is a more complex process than bioethanol, the high energy content associated with the lesser water miscibility, vapour pressure and corrosiveness are the favorable factors towards the ongoing biobutanol production. Moreover, the potential of Clostridium bacteria to ferment various substrates such as municipal wastes is also an added advantage to the biobutanol production. Different engineered strains also came into the race of efficient biobutanol production [2]. Different industries are putting maximum effort into biobutanol production on an industrial scale by focusing mainly on novel alternative methods for existing traditional ABE (Acetone, Butanol and Ethanol) fermentation. Gevo and Butamax are the two leading technology developers of butanol. Gevo commenced with the world’s first commercial-scale biobutanol production with a capacity of 18 MGPY and targeted to produce 50,000 to 100,000 gallons per month of isobutanol in upcoming years. Butamax in conjunction with Fagen Inc. set up a large-scale commercial production of biobutanol via retrofit of ethanol plants through the patented Butamax technology. India-based Laxmi Organic, Industries, in collaboration with Green Biologics, England, built a commercial-scale plant of 1,000 metric tons of butanol per year and began production from 2010 onwards. The facility would use sugarcane as a feedstock, and a combination of thermophilic organisms and thermostable enzymes to break the biomass down into butanol. Cobalt Biofuels in Mountain View, California, USA, had raised $25 million in equity to continue pursuing its goal of commercializing biobutanol production (http://www.biofuelstp.eu/butanol.html).

7.2 Production Technologies of Biobutanol Butanol could be produced biologically as well as chemically. In chemical processes it is produced through oxo synthesis (through syngas reacts

Technological Barriers 221 with propylene) and aldol condensation. In biological process, anaerobic bacteria-based fermentation approach is used to produce acetone and ethanol. The advantages and disadvantages of biological and physicochemical methods for biobutanol production are tabulated in Table 7.1. Cellulose and hemicelluloses also serve as substrates for biobutanol production with the aid of Clostridium sp. and other cellulolytic enzymes. Various pretreatment techniques such as utilization of alkaline peroxidases, steam explosion, hydrothermal techniques, and organic acids could be used to utilize the lignocellulosic substrate effectively. After pretreatment, detoxification is the subsequent step which is done through the utilization of activated charcoal, overliming, electrodialysis, and membrane extraction based detoxification methods. The pretreated and detoxified Table 7.1 Advantages and disadvantages of biological and physico-chemical method for the production of biobutanol. Method

Advantages

Disadvantages

Biological method

• Renewable source of fuel (feedstock)

• High feedstock cost significantly increases operating cost • Low butanol titre increase recovery cost. Low titres also increases sugar loading and water usage • Solvent recovery using distillation is energy intensive and expensive • Low butanol yield increases feedstock cost.

Physico-chemical

• Requires only one step for producing n-butanol from ethanol. • Relatively high yield

• Catalysts used in the process are costly

The sequence steps for formation of acetyl-CoA and its utilization for further fermentation intermediates are depicted in Figure 7.1

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AMYLASE

2 NADH, 2ATP 2NAD, 2ADP +PI

(1) (2)

GLUCOSE

(2)

2 PYRUVATE (3)

2CoA 2Co2

2 ACETYL CoA (4)

ACETOACETYL CoA

2NADH

2 ACETYL P

ATP

2NAD+

ACETOACETATE

BUTYRYL CoA

2 ACETALDEHYDE

ADP+PI

2 ACETATE ADP+PI ATP

BUTYRLDEHYDE

BUTYRL P

BUTANOL

BUTYRATE

ETHANOL

ACETONE

Figure 7.1 Biobutanol production from starch.

substrate is fermented finally through the utilization of microbial strains which convert polysaccharides to different sugars which are subsequently converted to pyruvate by glycolysis. The formed pyruvate is converted to acetyl-CoA by pyruvate dehydrogenase complex (PDC), which in turn act as a common originator of all fermentation intermediate [1]. The fermentation process is divided into two phases, namely acidogenesis and solventogenesis. In acidogenesis, glucose in substrate feed stream is converted to butyric acid and acetic acid by the action of Clostridium tyrobutyricum. The product streams are then circulated and passed through a series of heat exchangers, where they are sterilized at 250°F and then cooled back to 98.6°F before entering into the next phase of fermentation, solventogenesis [3]. During solventogenesis, the cells enter into the stationary phase where acids are converting to solvents by Clostridia acetobutylicum. The product stream obtained is then pumped to a centrifuge where separated solids are sent for Dried Distillers Grain (DDGS) for drying and liquids are sent to the separations process [4]. Butanol must be recovered from the fermentation broth by processes such as adsorption, using immobilization techniques with the help of membrane reactors and gas stripping. An overview of the biobutanol production from lignocellulosics is shown in Figure 7.2.

Technological Barriers 223 Bagasse, Barley straw, Wheat straw, Corn stover, Switch grass, Corn core Lignocellulosic Feedstock

Pretreatment

Detoxification

Dilute Sulphuric acid, Alkaline Peroxidase, Steam explosion pretreatment, hydrothermal pretreatment, organic acid pretreatment

Activated charcoal, over liming, Electrolysis, membrane extraction etc.

Fermentation

Batch Fermentation, Fed Batch fermentation, Continuous fermentation

Recovery Distillation, Gas Stripping, pervaporation, liquid-liquid extraction, Adsorption etc.

BUTANOL

Figure 7.2 Butanol production from lignocellulosic feedstocks.

7.3 Lignocellulosic Materials for Bio-Butanol Production Lignocellulosic materials used for biobutanol production face many challenges such as cost of the raw material, pretreatment and hydrolysis strategies, low butanol tolerance of fermenting strain which will affect the yield and productivity as well as downstream processing of biobutanol. There exists another barrier also which hinders its production, the inconsistency in biomass availability throughout the year. Lignin, ash, protein and waxes are also present in trace amounts whereas relative proportions of cellulose, hemicellulose, lignin are critical factors in the determination of optimum energy conversion route but other contents can lead to a diminution of theoretical butanol yield [5]. Among different problems encountered during utilization of cellulosic or lignocellulosic material for hydrolysates production, chemical by-products generation results in ceasing of cell growth as well as fermentation. Biologically these lignocellulosic materials are difficult to hydrolyze. Moreover, the significant amount of waste produced during the hydrolytic process also adds to the economy of the process [6].

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Recent research has shown that the fermentation of any polysaccharides needs to use an additional nutritional supplement. Lee et al. [7], reported the use of different supplements such as KH2PO4, K2HPO4, ammonium acetate, para-aminobenzoic acid, thiamin, biotin, MgSO4 · 7H2O, MnSO4 · H2O, FeSO4 · 7H2O, NaCl, and yeast extract for biobutanol production. Pretreatment is a critical and limiting step which has a predominant effect on overall biobutanol production thus has to be optimized. While using wheat bran as a substrate for biobutanol production by C. beijerinckii species, Liu and Qureshi [8], used a combination of pretreatments where wheat bran was treated with sulfuric acid at high temperature followed by neutralization with calcium hydroxide. This procedure raised the cost of the butanol produced considerably, but the solution to this problem found by using a cheap raw material for production which decreased the cost. Combination of pretreatments like 1M HCl with high temperature for 2 hours or enzymatic hydrolysis (using α-amylase and β-amylase) was utilized for cassava flour and resulted in 23.98 and 13.78 g.L-1 biobutanol production using enzymatic and acid hydrolysis, respectively [9]. In another study, biobutanol (12.0 g/L) was produced from wheat straw using enzyme mix of cellulose, β-glucosidase and xylanase with the process conditions of pH 5.0, 45°C for 72 h and 80 rpm. To hydrolyze the raw materials, use of other alternative mechanical and physicochemical technologies was also reported and those alternatives include microwave-assisted pretreatment processes, steam explosion, ozonolysis, oxidative delignification and pulsed-electric-field. Qureshi et al. [10], believed that barley straw could be used for butanol production. However, the barley straw showed the presence of few inhibitors which interfered with the production yield of butanol and hence pretreatment step using lime (called as over liming) was carried out to achieve an efficient fermentation. Consequently, this pretreatment step resulted in higher butanol production when compared to the yield with glucose as a substrate. Al-Shorgani et al. [11], also reported the formation of inhibitors during the acid pretreatment of cellulosic raw material (rice bran and de-oiled rice bran). Similarly, other studies utilized over liming pretreatment and subsequent extraction of inhibitors with a nonionic polymeric adsorbent. These procedures remarkably improved the biobutanol production and yield. Qureshi et al. [6], concluded that the formation of fermentation inhibitors after hydrolysis of cellulosic raw material is substrate and pretreatment dependent. Thus, it is necessary for a specific study to be carried out for each substrate and treatment. Several authors are of the opinion that the industrial feasibility of biobutanol production can only increase if a low-cost substrate can be employed [12, 13]. Diversification of

Technological Barriers 225 Glucose

Acetyl-CoA Acetoacetyl-CoA synthase

Amino Acids

Thiolase

Piruvate Acetoacetyl-CoA 3-hydroxybutyryl CoA dehydrogenase NADPH

acetolactate synthase

Acetoacetyl-CoA reductase NADPH

3S

2 Acetolactate

3R

Hydroxybutyryl-CoA R-enoyl-CoA dehyrdatase

Crotonase

Crotonyl-CoA Trans 2-enoyl CoA reductase NAD(P)H

Butyryl CoA dehydrogenase FADH2

ketoacid reducto isomerase

2,3-Dihydroxy-Isovalerate dihydroxy acid dehydrogenase

alfa-Keto-isovalerate alfa-Keto-isovalerate

Butyryl CoA

keto-isovalerate dehydogenase

Butyryl aldehyde dehydrogenase NAD(P)H

Isobutyraldehide

Butyraldehyde

Alcohol dehydrogenase

Butanol dehydrogenase NAD(P)H

Isobutanol

Butanol

(a)

(b)

Figure 7.3 (a) Metabolic representation of CoA-dependent pathway for production butanol, (b) Isobutanol synthesis pathway.

substrates and the use of regional crops such as molasses, starch or cellulose for butanol production is one of the approaches to tackle the high-cost of the fermented substrate. The metabolic representation of butanol (through CoA-dependent pathway) and isobutanol production is depicted under Figure 7.3.

7.4 Natural Producers of Biobutanol Clostridium sp. is the primary/natural producers involved in the production of biobutanol through the CoA-dependent pathway. The various species utilized for the production include C. acetobutylicum, C.  saccharoperbutylacetonicum, C. beijerinckii, C. saccharoacetobutylicum, C. aurantibutyricum, C. cadaveris, C. sporogenes, C. pasteurianum, and C. tetanomorphum [14].

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In Clostridia sp., acetyl-CoA produced from various carbon sources such as lactose and sugars fermentation through acidogenesis and solventogenesis. Acidogenesis produce acetic and butyric acid whereas solventogenesis produce acetone, butanol and ethanol. To make the process more economical, it is necessary to find a suitable method which shifts the metabolism from acidogenesis to solventogenesis. The primary barrier in the biobutanol production attributed to the adverse effects of the compounds or products obtained after solventogenesis on the microbial cell membrane. Butanol manifests chaotropic effects on the bacterial cell membrane due to which even its concentration as little as 2%, compromises bacterial survival. Therefore it becomes a prime concern to get rid of the toxicity of the solvent products. Deviation of metabolic intermediates from biosynthesis of aliphatic amino acid in yeast is also one of the natural metabolic pathways for biobutanol production. In some yeast species, fusel alcohols are one of the fermentation by-products [15]. In the Ehrlich pathway, keto acids are decarboxylated to produce aldehydes which in turn to alcohols. Since keto acids are the amino-acids precursors such as n-propanol, isobutanol and n-butanol are the precursors of isoleucine, valine and non-valine, respectively. Since isobutanol has a better octane number, therefore, it is preferred over n-butanol for industrial use [16]. The industrial application of isobutanol is hampered by the meager intrinsic production in yeast, but this route of biobutanol production diverts the PROTEINS AMINO ACIDS

Serine hydroxymethyl transferase

SERINE

GLYCINE Glycine oxidase

(1)

Serine deaminase

GLYOCYLATE CYCLE

GLYOXYLATE Butyryl - CoA (2)

Malate Synthase

CoA

PYRUVATE

Multistep conversion

β -ETHYLMALATE β- Isopropyl malate

(3)

NADH, CO2 (5) α -Ketoacid isomerase

α-ISOKETOVALERATE

α -KETOVALERATE NAD+, CO2 (4)

BUTANOL

(6) NAD+, CO2 Pyruvate decarboxylase & alcohol dehydrogenase

ISOBUTANOL

Figure 7.4 Production of butanol and isobutanol by using glycine as a substrate.

Technological Barriers 227 various pathway intermediates towards amino acid biosynthesis which usually is not possible naturally [8]. The steps involved in production of butanol and isobutanol are summarized in Figure 7.4.

7.5 Main Obstacles in the Biobutanol Production The problems associated with biobutanol production were increased cost of feedstock, low butanol titer which further increases butanol recovery and downstream processing, reduced sugar loadings and increased water usage which increases the capital expense. High water usage is not sustainable which further increases the cost of production and it is energy intensive and relatively expensive, low butanol tolerance of the microbes as this much amount of alcohol destroys their cell wall [17].

7.5.1 Approaches to Overcome the Obstacles To overcome the obstacles, metabolic engineering is one of the alternatives that could adapt for modified strains by overexpressing the butanol production genes such as BCS operon related genes and add, bdh. Overexpression of grosESL gene lead to improved strain tolerance and increase in butanol titer. Recently, Global transcription machinery engineering (gTME) is another promising approach to enhance biobutanol production. If there is any alteration in the transcription factor, there is a scope for gTME to change the metabolic strength and direction. The gTME system has been proved as an efficient solution to improve substrate utilization and product tolerance [18].

7.6 Engineered Pathways towards a Better Solventogenic Producer 7.6.1 Engineered Pathways in Bacteria Mutagenesis is gradually becoming a method of choice to improve C. acetobutylicum. Several attempts are made in this direction since mutagenesis has helped in improving yield, tolerance to butanol, and sugar source utilization. To increase the efficiency of production, many species of Clostridium have been engineered so that they become capable of utilizing some other carbon sources (liquefied corn flour, glycerol and a mixture

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of hydrogen and carbon monoxide) [19]. Acetone has to be removed to improve fuel alcohols in the ABE process which can be done by inactivating adc gene which codes for acetoacetate decarboxylase necessary for acetone synthesis [20]. Metabolic engineering is another approach which is used to obtain isopropanol from acetone. A mixture of isopropanol, butanol and ethanol (IBE) produced through engineering of C. acetobutylicum by overexpressing the dehydrogenase gene of C. beijerinckii in C. acetobutylicum. The modified strain is capable to produce more than 99% of fuel alcohol with a negligible amount of acetone. Metabolic engineering helps in improving metabolic fluxes of wild strain to improve yield [21]. The metabolic engineered solventogenic Clostridia is able to ferment starchy and molasses towards biobutanol production. Gaseous substrates also serve as carbon substrates for engineered acetogenic Clostridia strain; hence there is no competence with the nutritional feedstock as substrate. The major genes involved in the butanol synthesis pathway in C. acetobutylicum are thlA, hbd, crt, bcd, adhE, and bdhA, which codes for the thiolase, 3-hydroxybutyryl-CoA dehydrogenase, crotonase, butyryl-CoA dehydrogenase, butanol/butyraldehyde dehydrogenase, and butanol dehydrogenase enzymes, respectively. The Clostridium butanol pathway genes were introduced into other fast-growing bacteria such as E. coli, the engineered bacteria shown butanol toxicity tolerance and is able to metabolize alternative substrates [22]. Pseudomonas putida and Bacillus subtilis are another bacterial species which can serve as hosts with butanol toxicity tolerance through efflux pumps. Lactobacillus brevis, which has a high tolerance to butanol, and can digest C5 and C6 substrates, has also been used. Another major hindrance to butanol production is the intrinsic kinetic characteristics and cofactor specify of all enzymes that occur naturally in the bacterial pathway. Synthetic pathways are a solution to this limitation which can be made by combining enzymes from different organisms into a synthetic butanol pathway expressed in E. coli. To manifest this, the modified strain of E. coli is transfected with the vectors carrying genes of two enzymes: 2-keto-acid decarboxylase of low substrate specificity along with an alcohol dehydrogenase. This manipulation resulted in high yields of isobutanol [23]. The 2-keto-acid pathway of Corynebacterium glutamicum has also been engineered, taking advantage of the high amino-acid production characteristic of the bacteria. The pretreatment of lignocellulosic materials is another costly affair which can overcome by using Clostridium cellulolyticum which can naturally digest lignocellulose. Gaida et al. [24], reported for the first time the metabolic engineering of C. cellulolyticum where the bacterium was engineered with the CoA-dependent pathway

Technological Barriers 229 to produce n-butanol directly from crystalline cellulose using NADH as a cofactor. When natural producers of butanol are intended to use, then the best strategies are those based on the selection of strains, fermentation conditions and best product recovery techniques that may avoid toxicity or enhance the fuel concentration by avoiding the production of acetone or isopropanol. The introduction of the metabolic pathway for ABE fermentation in other bacteria possesses several advantages such as nonrequirement of anaerobic fermentations with fast-growing or butanol toxicity tolerance abilities. A two-fold increase in the concentration of fuel has been observed in comparison to natural strains of Clostridium acetobutylicum when the non-natural producer bacteria engineered with the 2-keto-acid pathway. A metabolic approach is a promising area in biobutanol production to attain higher yields [25].

7.6.2 Engineered Pathways in Yeast Although bacterial systems such as E. coli have been known to show more efficient systems for biobutanol production, the engineered Sacchromyces cerevisae system has also been proved as quite an efficient system to produce biobutanol. Production of biobutanol through yeast systems have some positive attributes such as adaptability in various application sectors and have greater reproducibility. Moreover, the bacterial-based biobutanol system suffered from disadvantages like the requirement of strictly anaerobic conditions, complex downstream processing, narrow pH requirements and infection prone due to phages and viruses. However, these limitations can be overcome by utilizing yeast-based systems especially S. cerevisaebased system. S. cerevisiae have the enzyme machinery to synthesize isobutanol by a two-compartment Ehrlich pathway. In this pathway, keto-isovalerate, which is an intermediary product of valine synthesis in mitochondria, is catabolised into isobutanol. Genes ILV2, ILV5, ILV3, encode for the enzymes acetolactyl synthase, ketoacid reductasoisomerase, and dihydroxy acid dehydratase respectively, are majorly involved in isobutanol synthesis. Overexpression of these genes leads to enhanced biobutanol production. The above genes mentioned above are present in mitochondria, so this system can replace in cytosol which is done by overexpression of cytosolic form encoded by ILV2, ILV5, ILV3, ARO10 with deletion of mitochondrial ILV2 associated genes. To increase dihydroxyacid dehydratase encoded by ILV3 or to improve the cofactor associated with it, many additional genetic manipulations were researched to enhance other by-products besides

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ethanol. With the combination of these strategies, more than 80% of the maximum isobutanol theoretical yield achieved. Codon usage linked with cytosolic pathway than using from associated mitochondrial systems has too proved to be a useful technique. In one of the alternatives, the catabolic pathway of amino acids which resulted from protein hydrolysis was utilized [26]. Many researchers have reported the use of glycine as the substrate for the synthesis of butanol and isobutanol in S. cerevisiae through glyoxylate pathway. Various genetic engineering approaches have been utilized to change the metabolic activities in S. cerevisiae. These include reconstruction of the 1-butanol biosynthetic pathway through increased flux towards cytosolic acetyl-CoA by means of the transformation with a plasmid expressing the genes (encode for ADH2 (alcohol dehydrogenase), ALD6 (acetaldehyde dehydrogenase), ACS1/ACS2 (acetyl-CoA synthetase), and ERG10 (acetyl-CoA acetyltransferase) enzymes). An endogenous 1-butanol pathway in S. cerevisiae which was dependent on catabolism of threonine, was proposed, discovered, characterized and engineered by Si et al. [27]. Various strategies have been introduced to achieve the higher 1-butanol titer in S.cerevisiae which include the overexpression of the Ehrlich pathway enzymes, and single gene deletion adh1delta (which cause a deficiency of alcohol dehydrogenase) was proposed [27]. Matsuda et al. [28], reported enhanced biobutanol yield by utilizing S.  cerevisiae-based metabolic engineering approach. The strains engineered in such a way that they lacked the genes of pyruvate dehydrogenase complex LPD1 to reduce the competition between pyruvate supply for isobutanol and acetyl Co-A biosynthesis in mitochondria. The genes of the enzyme transhydrogenase-like shunts (converts NADH to NADPH) were overexpressed to resolve the cofactor imbalance. Endogenously, α-ketobutyrate is a key intermediate in n-butanol production pathway. Usually, α-ketobutyrate is synthesized from catabolism of threonine and alternately it can be synthesized from acetyl Co-A via Citramalate synthase (Cim A) (Figure 7.5). Through this approach, a maximum theoretical n-butanol yield of 411 mg/g glucose was achieved. The maximum theoretical n-butanol yield reported a value of 411 mg/g glucose through this approach. Further research has been done to improve the n-butanol production by cloning and overexpressing the CimA genes from Methanococcus jannaschii, Leptospira interrogans, and Geobacter sulfurreducens with previously confirmed α-ketobutyrate utilizing genes (mLEU1, mLEU4, mLEU2, and LEU5). The strain LI with overexpressed Cim A from L. interrogans, showed high improved n-butanol titer of 349 mg/L. The yield obtained was far higher than the strain THRm solely

Technological Barriers 231 PYC1/2 Oxaloacetate

AAT1 HOM3/2/6 THR4/1 Threonine

Glucose

Pyruvate

CimA

Pyruvate

Acetyl-CoA (R)-citramalate LEU1/2

ILV1

α-ketobutyrate Acetyl-CoA+H2O CoA Degradation

LEU4 2-ethyl-malate

2-ethyl-malate

LEU1/2

LEU1/2

α-ketovalerate Replenishment LEU5 CoA

Mitochondrial membrane

α-ketovalerate KDCs ADHs n-butanol

Figure 7.5 The synergistic pathway for n-Butanol production via the endogenous threonine pathway and an introduced CimA mediated pathway (Atsumi et al. [29]). Single and double arrows represent single and multiple enzymatic steps, respectively; red arrows represent heterologous pathways; overexpressed genes are marked with red colour. KDCs: keto-acid decarboxylases; ADHs: alcohol dehydrogenases.

overexpressing the threonine pathway. The strain LI, uses two metabolic pathways to synthesize butanol, i.e., the endogenous threonine pathway and the introduced citramalate pathway. This synergistic path leads to a maximum theoretical yield of 411 mg/g glucose n-butanol [30].

7.7 In-Situ Butanol Recovery Integrated with Batch and Fed-Batch Fermentation There were many problems and disadvantages associated with the traditional fed-batch system. One of such problems was increased solvent toxicity during the biphasic batch butanol fermentation. In-situ recovery process integrated with fed-batch culture is the ideal setup to overcome the

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solvent toxicity using silicone and oleyl alcohol as pre-extraction solvents along with the utilization of silicon membrane [31]. The butanol extraction process through diffusion was carried out using silicone membrane which limits the diffusion of acetone and acids that subsequently results in higher biobutanol yield. Gas stripping was another batch process method where a semi-synthetic medium with embedded lactose which was entirely fermented by C. Acetobutylicum. The integration of gas stripping setup with the liquid-liquid extraction process results in higher utilization of lactose which led to the enhanced biobutanol production [32]. The lower yields of butanol production through traditional fermentation process were mainly due to the accumulation of butanol in the fermentation broth. Butanol removal from the fermentation broth and its separation was a costlier process. To avoid such problems addition of butyrate as a precursor to the system is the ideal approach to trigger the metabolic pathway towards the butanol production and further promising results were attained through the integration with in-situ butanol removal via vacuum membrane distillation [33] which alleviates the butanol toxicity issues. This integration is a very effective method to enhance the butanol yield with higher economic feasibility with easier downstream processing. Another approach for enhanced biobutanol titers was opting for adsorbent based fermentation along with renewable carrier [34]. Alkali-treated steam explodes straw showed as a suitable carrier for adsorbent fermentation of biobutanol. The adsorption of ABE solvent on substrate facilitates the increased bacterial concentration alleviation of the end product inhibition with improved biobutanol production features.

7.8 Future Prospects The future of the industrial process for the production of biobutanol can improve by utilization of novel omics-based approaches and sophisticated downstream processes. Clostridium acetobutylicum is the most intensively studied solvent-producing species involved in biobutanol production. A thorough investigation of existing metabolic pathway towards biobutanol production helps in pinpointing the responsible genes, and its overexpression in different hosts. Advances in continuous culture technology, integrated fermentation processes, in situ product removal and improved downstream processing can also provide new approaches to improve the substrate utilization that also provides a future direction of economic biobutanol production by reducing butanol toxicity and process stream volumes towards the enhanced bioreactor performance.

Technological Barriers 233

7.9 Conclusions Biobutanol is also a superior biofuel and in a very long term has been shown to meet the demands for the next-generation biofuels. Production of biofuels is even considered as a useful means to slow down carbon dioxide emissions; this is also a green industry with ecological benefits to humankind, and that could also contribute to decreasing the present concerns over global climate change. Production of biobutanol through Clostridium sp. in higher volumes is a viable strategy to compete with the chemical-based butanol production. Recent advances in bioethanol plants could be costeffectively retro-fitted for biobutanol production requiring relatively minor changes to fermentation. The lower titers associated with the biobutanol production with the Clostridium sp. can resolve by utilizing the novel engineering, metabolic approaches coupled with integrated recovery processes.

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10. Qureshi, N., Cotta, M.A., Saha, B.C., Bioconversion of barley straw and corn stover to butanol (a biofuel) in integrated fermentation and simultaneous product recovery bioreactors. Food Bioprod. Process., 92, 298–308, 2014. 11. Al-Shorgani, N.K., Kalil, M.S., Yusoff, W.M., Biobutanol production from rice bran and de-oiled rice bran by Clostridium saccharoperbutylacetonicum N1-4. Bioprocess. Biosyst. Eng., 35, 817–826, 2012. 12. Desai, S.H., Rabinovitch-Deere, C.A., Tashiro, Y., Atsumi, S., Isobutanol production from cellobiose in Escherichia coli. Appl. Microbiol. Biotechnol., 98, 3727–3, 2014. 13. Lin, P.P., Rabe, K.S., Takasumi, J.L., Kadisch, M., Arnold, F.H., Liao, J.C., Isobutanol production at elevated temperatures in thermophilic Geobacillus thermoglucosidasius. Metab. Eng., 24, 1–8, 2014. 14. He, H., Lui H, H., Gan, Y.R., Genetic modification of critical enzymes and involved genes in butanol biosynthesis from biomass. Biotechnol. Adv., 471, 1, 2010. 15. Hazelwood, L.A., Walsh, M.C., Pronk, J.T., Daran, J.M., Involvement of vacuolar sequestration and active transport in tolerance of Saccharomyces cerevisiae to hop iso-α-acids. Appl. Environ. Microbiol., 76, 318–328, 2009. 16. Branduardi, P., Longo, V., Berterame, N., Rossi, G., Porro, D., A novel pathway to produce butanol and isobutanol in Saccharomyces cerevisiae. Biotechnol. Biofuels., 6, 68, 2013. 17. García, V., Päkkilä, J., Ojamo, H., Muurinen, E., Keiski, R.L., Challenges in biobutanol production: How to improve the efficiency?. Renew Sust. Energ. Rev., 15, 964–980, 2011. 18. Bennett, G.N., Rudolph, F.B., The central metabolic pathway from acetyl‐ CoA to butyryl‐CoA in Clostridium acetobutylicum. FEMS Microbiol. Rev., 17, 241–249, 1995. 19. Ezeji, T.C., Qureshi, N., Blaschek, H.P., Production of acetone butanol (AB) from liquefied corn starch, a commercial substrate, using Clostridium beijerinckii coupled with product recovery by gas stripping. J. Ind. Microbiol. Biotechnol., 34, 771–777, 2007. 20. Kopke, M., Held, C., Hujer, S., Liesegang, H., Wiezer, A., Wollherr, A., Ehrenreich, A., Liebl, W., Gottschalk, G., Durre, P., Clostridium ljungdahlii represents a microbial production platform based on syngas. Proc. Nat. Acad. Sci., 107, 13087–13092, 2010. 21. Jang, Y.S., Malaviya, A., Lee, J., Im, J.A., Lee, S.Y., Lee, J., Eom, M.H., Cho, J.H., Seung Do, Y., Metabolic engineering of Clostridium acetobutylicum for the enhanced production of isopropanol‐butanol‐ethanol fuel mixture. Biotechnol. Prog., 29, 1083–1088, 2013. 22. Berezina, O.V., Zakharova, N.V., Brandt, A., Yarotsky, S.V., Schwarz, W.H., Zverlov, V.V., Reconstructing the clostridial n-butanol metabolic pathway in Lactobacillus brevis. Appl. Microbiol. Biotechnol., 87, 635–646, 2010. 23. Smith, K.M., Cho, K.M., Liao, J.C., Engineering Corynebacterium glutamicum for isobutanol production. Appl. Microbiol. Biotechnol., 87, 1045–1055, 2010.

Technological Barriers 235 24. Gaida, S.M., Liedtke, A., Jentges, A.H.W., Engels, B., Jennewein, S., Metabolic engineering of Clostridium cellulolyticum for the production of n-butanol from crystalline cellulose. Microb. Cell Fact., 15, 6, 2016. 25. Atsumi, S. and Liao, J.C., Metabolic engineering for advanced biofuels production from Escherichia coli. Curr. Opin. Biotechnol., 19, 414–419, 2008. 26. Brat, D., Weber, C., Lorenzen, W., Bode, H., Boles, E., Cytosolic re-localization and optimization of valine synthesis and catabolism enables increased isobutanol production with the yeast Saccharomyces cerevisiae. Biotechnol. Biofuels., 5, 65, 2012. 27. Si, T., Luo, Y., Xiao, H., Zhao, H., Utilizing an endogenous pathway for 1-butanol production in Saccharomyces cerevisiae. Metab. Eng., 22, 60–68, 2014. 28. Matsuda, F., Ishii, J., Kondo, T., Ida, K., Tezuka, H., Kondo, A., Increased isobutanol production in Saccharomyces cerevisiae by eliminating competing pathways and resolving cofactor imbalance. Microb. Cell Fact., 12, 119, 2013. 29. Atsumi, S. and Liao, J.C., Metabolic engineering for advanced biofuels production from Escherichia coli. Curr. Opin. Biotechnol., 19, 414–419, 2008. 30. Risso, C., Van Dien, S.J., Orloff, A., Lovley, D.R., Coppi, M.V., Elucidation of an alternate isoleucine biosynthesis pathway in Geobacter sulfurreducens. J. Bacteriol., 190, 2266–2274, 2008. 31. Qureshi, N., Maddox, I. S., Friedl, A., Application of continuous substrate feeding to the ABE fermentation: Relief of product inhibition using extraction, perstraction, stripping, and pervaporation. Biotechnol. Prog., 8, 382–390, 1992. 32. Maddox, I.S., Qureshi, N., Roberts-Thomson, K., Production of acetonebutanol-ethanol from concentrated substrate using clostridium acetobutylicum in an integrated fermentation-product removal process. Process Biochem., 30, 209–15, 1995. 33. Chiam, C.K. and Sarbatly, R., Vacuum membrane distillation processes for aqueous solution treatment—A review. Chem. Eng. Processing: Process Intens., 74, 27–54, 2013. 34. Xue, C., Zhao, X., Liu, C.G., Chen, L.J., Bai, F.W., Prospective and development of butanol as an advanced biofuel. Biotechnol. Adv., 31, 1575–1584, 2013.

8 Biobutanol: Research Breakthrough for its Commercial Interest Sandip B. Bankar1*, Pranhita R. Nimbalkar2, Manisha A. Khedkar2 and Prakash V. Chavan2 1

Department of Bioproducts and Biosystems, School of Chemical Engineering, Aalto University, Aalto, Finland 2 Department of Chemical Engineering, Bharati Vidyapeeth Deemed University College of Engineering, Pune, India

Abstract Biobutanol as an advanced potent biofuel has re-gained great attention in recent years due to its eco-friendly nature and superior properties compared to other available liquid fuels (ethanol and methanol). However, the cost of biobutanol production via conventional acetone-butanol-ethanol (ABE) fermentation by Clostridium sp. is not economically competitive which in turn hampered its industrial application. This chapter exclusively describes re-commercialization efforts and highlight developments in feedstock utilization, strain engineering and fermentation process expansion, which significantly contribute in overall production cost. Additionally, advancement in hybrid separation processes for in situ butanol recovery is discussed in detail. Besides, it also sheds light on techno-economic assessment, key technical challenges and opportunities of the lignocellulosic biomass-to-bioenergy production in comparison with the first generation technologies. More importantly, this chapter would offer a new perspective in relation to existing knowledge and new opportunities for economically feasible ABE fermentation process in near future. Keywords: ABE fermentation, biobutanol, biosynthetic pathway, clostridia, feedstock, hybrid separation processes, saccharification, strain engineering

*Corresponding author: [email protected]; [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (237–284) © 2019 Scrivener Publishing LLC

237

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8.1 Introduction Energy is an indispensable component of modern society as almost everything right from home appliances to industrial processes are energy dependable. According to International Energy Outlook 2017 (IEO2017), total world energy consumption was around 575 quadrillion Btu (QBtu) in 2015 and projected to reach 736 QBtu by 2040 [1]. Mainly, about 91% of energy comes from non-renewable sources such as petroleum, coal, natural gas, and nuclear [2]. In early decades, the crude oil prices were quite low and affordable while country’s advancement has led to dramatic rise in oil prices during recent years [3]. In addition, rapid depletion of existing reservoirs along with flourishing environmental issues, clearly suggest that an urgent and decisive action needs to be taken to develop sustainable, clean, affordable and renewable energy sources. Renewable sources such as wind, solar, geothermal, tidal, hydropower, and biomass are being well thought out, all of which can equally contribute to renewable energy portfolio [3]. Though, current energy consumption largely relies on non-renewable resources, there are tremendous research and technology development efforts are on-going to build up more potential way out from renewable sources to overcome energy insecurity. Biofuel has received a great attention and can be considered as a leading candidate for renewable energy generation, particularly for liquid transportation fuel [4]. Generally, the term biofuel is used for fuels that are produced in biological way using renewable sources (e.g. biomass, mainly various agricultural residues and other related wastes). These biologically derived fuels are eco-friendly, environmentally safe and hence would be better substitute to conquer petroleum’s dependency [3]. Biofuels are classified in various generations based on type of substrate used for their production. The fuels produced directly from food (starchy) materials are categorized under ‘first generation’, while second generation biofuels are produced specifically from non-food materials such as lignocellulosic biomass (Figure 8.1). Besides, third and fourth generation fuels are produced from algae and genetically modified crops, respectively [5]. Though, current spotlight is on 2nd and 3rd generation, the 1st generation biofuels constructs an excellent model to understand and apply the bio-refinery perception. At present, major biofuels are ethanol and biodiesel, which are being commercially produced from conventional sugar and starch-based feedstocks such as sugarcane and corn in Brazil and the USA, respectively [6]. Nonetheless, butyl alcohol/butanol is another very attractive energy source available and considered as superior fuel when

Biobutanol: Research Breakthrough 239 Biofuel Generations

First Starchy and sugar based Sugar beet

Corn

Sugar cane

Wheat

Sweet sorghum

Barley

Second Lignocellulosic biomass Straw

Third Algal biomass Micro-algae

Wood chips

Macro-algae Energy crops

Figure 8.1 Different feedstocks used for biofuel production.

compared with ethanol in many regards. Hence, this chapter outlines the current status and research efforts taken to bring biobutanol close to commercialization as next generation liquid fuel.

8.2 Butanol: Next Generation Liquid Fuel As four carbon straight-chain primary alcohol, butanol is a clear, colorless liquid possessing excellent fuel properties [7]. As a result, it has been acknowledged as an advanced biofuel and attracted great attention across the world [8]. The butanol market has been forecasted to grow more rapidly in all over the world. China holds largest share of butanol market during 2014-2019 [9]. Interestingly, Asia-Pacific region covers biggest market of n-butanol, which accounts for 51.3% consumption by volume in 2014 [10]. Moreover, global n-butanol market is expected to reach $9.9 billion by 2020 [9]. Table 8.1 summarizes and compares some common properties of different alternative fuels and also shows that properties of butanol matches more closely with gasoline than other listed fuels. In more detail, butanol has low volatility in comparison with gasoline, ethanol and methanol, which makes it easier to

Gasoline 32 0.7–0.21 53.8–103 14.6 0.36 91–99 81–89 – < –60 Low Yes

Parameters

Energy density (MJ/L)

Vapor pressure (kPa) at 20°C

Vapor pressure of mixture with gasoline (kPa)

Air:Fuel ratio

Heat of vaporization (MJ/kg)

Research octane number

Motor octane number

Cetane number

Freezing temperature (°C)

Hygroscopicity

Compatibility with existing infrastructure

Yes

Low

– 89.5

–

78

96

0.43

11.2

44.1

0.53

29

n-butanol

Table 8.1 Comparison of different fuels on basis of fuel properties [5].

Yes

Low

– 108

–

112

–

–

–

46.9

1.17

29

Isobutanol

No

High

– 114

54

102

129

0.92

9.0

138

7.58

19.6

Ethanol

No

High

– 97.6

–

97–104

129–134

1.16

6.5

800

12.8

16

Methanol

240 Liquid Biofuel Production

Biobutanol: Research Breakthrough 241 handle and safer to use in any weather conditions. Additionally, it exhibits a very low vapor pressure and high flash point which confirms its safety at high temperatures. Besides, the heat of vaporization of butanol is slightly higher than that of gasoline. Hence, butanol-blended engine does not exhibit cold start problem as seen in methanol or ethanol blended gasoline [11]. It also has low solubility in water which further reduces the chances of groundwater contamination [12]. Furthermore, it is less corrosive than ethanol and thus it can be delivered through existing engine infrastructure [13]. More importantly, butanol can be blended with gasoline at any ratio of more than 10% without engine modification unlike ethanol. Butanol has several applications in various industries start from its use as chemical feedstock to solvents for chemical synthesis. Particularly, in coatings, it’s an intermediate in the production of butyl acrylate and methacrylate esters that are used in latex surface coating, enamels, and lacquers [14]. Almost half of produced butanol per year is utilized in ester synthesis. In addition, some of its derivatives are established plasticizers for plastics, rubber mixes, and dispersions [15]. Besides, butanol can serve as a solvent in the manufacturing processes for paints, natural and synthetic resins, varnishes, gums, dyes, alkaloids, vegetable oils, fats, and waxes [16]. It can also be used in textiles as a swelling agent in garment manufacturing and in pharmaceuticals as an extractant specifically for production of drugs such as antibiotics, hormones, and vitamins. Moreover, it is widely used in cosmetics for the production of eye makeup, foundations, lipsticks, nail care products and many more [14].

8.3 Routes of Butanol Production 8.3.1 Chemical Route Butanol is mainly produced via chemical synthesis. There are three major processes usually being used in the chemical butanol production namely oxo synthesis, Reppe synthesis and crotonaldehyde hydrogenation [17]. The most common chemical route is oxo process, in which reaction of propylene with carbon monoxide and hydrogen in the presence of an appropriate catalyst was performed (Equation 1). The resulting mixture of n and isobutyraldehyde is hydrogenated to corresponding n- and isobutyl alcohols and further distilled to recover butanol of desired quality [18]. In the Reppe process, reaction between propylene, carbon monoxide and water takes place in presence of catalysts at low temperature and pressure which then directly results into butanol formation (Equation 2) [17]. However, the maintenance of stringent

242

Liquid Biofuel Production

conditions during reaction is quite difficult which in turn makes the process economically non-viable. Besides, crotonaldehyde hydrogenation involves three main steps as: aldol condensation, dehydration, and hydrogenation (Equation 3). Unlike above two, this process starts with acetaldehyde as initial reactant which can be obtained through ethanol dehydrogenation. Rh, Co or Ru

(1) CH3CH=CH2 H2, CO

CH3CH2CH2CHO + CH3CHCHO H2

Ni

CH3

CH3CH2CH2CH2OH Rh, Co or Ru

(2) CH3CH=CH2

CH3CH2CH2CH2OH + CH3CHCH2OH + 2CO2 H2, CO

Aldol condensation

(3) 2CH3CHO

CH3

Dehydration

CH3CH(OH)CH2CHO

CH3CH=CHCHO + H2O H2

Hydrogenation

CH3CH2CH2CH2OH

(1) Oxo synthesis, (2) Reppe process, and (3) crotonaldehyde hydrogenation [14]. Overall, all of these processes include reactions with carbon monoxide which will have adverse effect on human health. Hence, biological route of butanol production is safer and most preferred over chemosynthesis.

8.3.2 Biological Route Butanol can also be produced through microbial fermentation. More specifically, biological route of butanol production is via acetone-butanolethanol (ABE) fermentation using Clostridium sp. [19]. A typical ABE fermentation evolved as the second largest industrial fermentation process in the world after ethanol [5]. The general steps involved in butanol production are shown in Figure 8.2. Fermentative ABE production flourished during the early 20th century. Nonetheless, due to development of petrochemical industry, ABE fermentation process had lost its competitiveness by 1960s. With the increasing butanol markets, the historical biobutanol production has re-announced

Biobutanol: Research Breakthrough 243

Pure butanol

Product value

Recovery and purification

Detoxification

Saccharification

Lignocellulosic biomass

Acetone Butanol Ethanol ABE fermentation By clostridia Genetically engineered microbes

Processes Overliming Activated charcoal Ion exchange resins

Pretreatments Physical Chemical Biological Operational cost, energy and time

Figure 8.2 Detail steps involved in biobutanol production.

its importance and more production plants have been developed globally. China takes the lead in installing 0.21 million TPA (tons per annum) of solvent production facility and planned to expand it to 1 million TPA [14]. Generally, ABE fermentation process yields acetone, butanol and ethanol in the ratio of 3:6:1 [20]. The entire fermentation process is categorized in two phases namely acidogenic and solventogenic phase. In former, the acetic acid and butyric acids are produced followed by re-assimilation into acetone, butanol and ethanol in latter stage [21]. The optimum temperature for ABE fermentation is between 30 and 40°C while initial pH of fermentation broth is in the range of 6.8-7; and it drops down to 4.5-5 during acidogenic phase [22]. The detailed ABE fermentation process is discussed in subsequent section.

8.4 Microbial ABE Production The formation of butanol in microbial fermentation was first reported by Louis Pasteur in 1861. In subsequent years, Albert Fitz worked keenly in the field of fermentation and produced butanol from glycerol with the help of two bacteria [18]. Further, some other researchers continued their

244

Liquid Biofuel Production

work and research was focused in order to produce acetone, amyl alcohol or butanol at the beginning of 20th century. Thereafter, in 1912, Chaim Weizmann isolated a bacterium strain (later named Clostridium acetobutylicum) which was able to use starch as a substrate and exhibited higher product yields of butanol and acetone in comparison to Fernbach’s process. During First World War, major attention was given on acetone as it was essential in production of cordite (smokeless gun powder) and thus butanol was just considered as by-product [14]. Later on, because of automobile industry expansion, butanol became the main product of fermentation and acted as solvent for quick drying lacquers. Thereon, several countries started butanol production using locally isolated Clostridium strains at industrial level during the period 1920-1980. Furthermore, new strains were isolated along with novel substrates and better equipment design to improve fermentation processes. Meanwhile, around 66% of butanol and 10% of acetone were still produced through ABE fermentation [23]. However, soon after the end of war, declination of fermentation industry began and even became more drastic during the 1950s. By the period of 1960, almost entire solvent production via fermentation was ceased in the United States and the United Kingdom. This cessation was thought to be due to two major reasons. First, the butanol production through petrochemical route grew rapidly and competed acutely with the fermentation process [24]. Second, the prices of molasses which was used as a major feedstock for fermentation, increased suddenly because of its simultaneous use in many other areas such as cattle feed [18]. However, as a result of oil crisis in 1970s, once again development of biofuels was resurrected and research efforts were started in the direction of physiology optimization and new downstream process incorporation.

8.4.1 Microbial Strains Butanol producing strains can mainly be divided as wild type and genetically modified [25]. Butanol can be naturally produced by members of Clostridium genus. Chaim Weizmann first isolated Clostridium acetobutylicum and discovered that it produces acetone, ethanol and butanol from starch. Clostridia are strict anaerobes, rod-shaped, spore-forming Gram positive bacteria [26]. However, only few Clostridium sp. can produce significant amount of butanol under appropriate conditions. Genetically modified strains were also developed in recent years. The four major primary butanol producers: C. acetobutylicum, C. beijerinckii, C. saccharobutylicum, and C. saccharoperbutylacetonicum [27]. Among them, C. beijerinckii

Biobutanol: Research Breakthrough 245 produces isopropanol instead of acetone along with butanol and ethanol thus responsible for forming green solvents [11].

8.4.2 Biosynthetic Pathways of Clostridia As explained earlier, ABE fermentation can be categorized into two phases: acidogenesis and solventogenesis. The biosynthetic pathways in ABE fermentation by C. acetobutylicum are as shown in Figure 8.3. The metabolic activities of Clostridial sp. are known to form end products such as butanol, acetone, ethanol, acetic acid, butyric acid, CO2, and H2 [28]. Clostridia can easily utilized a large variety of substrates right from monosaccharides such as hexose and pentose to polysaccharides including starch, Glucose ATP ADP

Fructose 6-P ATP ADP

Glyceraldehyde 3-P 4ATP

Acidogenesis

Solventogenesis

4ADP

Pyruvate ATP

Acetate

ADP

14

CoA

Acetyl-P

Pi

CoA 2CO2

1

NAD+

NADH

Acetyl-CoA

13

Acetylaldehyde

11 Acetyl CoA 2

NADH NAD+

Ethanol

12

Acetate

15

CoA 16

Acetoacetate

Acetoacetyl-CoA

Acetone CO2

NADH NAD+

3 10

3-hydroxybutyryl -CoA H2O

Butyrate

4

Crotonyl-CoA NADH ATP

Butyrate

ADP

9

Pi

CoA

Butyryl-P

8

NAD+

5

Butyryl-CoA

NAD+

NADH

6

NAD+

NADH

Butylaldehyde

7

Butanol

Figure 8.3 The metabolic pathway of ABE fermentation by C. acetobutylicum. PPP: pentose phosphate pathway; EMP: Embden–Meyerhof pathway; 1: pyruvate ferrodoxin oxidoreductase; 2: thiolase; 3: 3-hydroxybutyryl-CoA dehydrogenase; 4: crotonase; 5: butyryl- CoA dehydrogenase; 6: butyraldehyde dehydrogenase; 7: butanol dehydrogenase; 8: phosphotransbutyrylase; 9: butyrate kinase; 10: butyrate CoA transferase; 11: acetaldehyde dehydrogenase; 12: ethanol dehydrogenase; 13: phosphotransacetylase; 14: acetate kinase; 15: acetoacetyl CoA acetate; 16: acetoacetate decarboxylase [5].

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Liquid Biofuel Production

xylan, and even glycerol can be used as a carbon source by few Clostridial sp. [29]. In addition, various second generation inexpensive substrates such as agricultural waste, fruit and vegetable industry waste, sago starch and others. These wastes can be hydrolyzed to yield simple monomeric sugars and are considered then used as carbon source to produce butanol by different Clostridial strains. On the other hand, it has also been reported that Clostridia can able to secrete extracellular enzymes like amylase, glucosidase and pullulanase for efficient conversion of complex polysaccharides to monosaccharides [14]. Usually, Clostridia, an anaerobe uptake sugars via the phosphoenolpyruvate dependent phosphotransferase system. Especially, these microorganisms metabolize hexose sugars by Embden–Meyerhof pathway (EMP) while pentose sugars goes through pentose phosphate pathway (PPP) to produce pyruvate. In more detail, in case of EMP, 1 mol of hexose is converted to 2 mol pyruvate with net production of 2 mol of ATP and NADH [30]. On the other hand, substrates with hemicellulose can be converted to xylose and glucose by hydrolysis. The resulted glucose normally metabolized through EMP pathway. Xylose is converted to xylulose by isomerase activity and then further into glyceraldehyde 3-phosphate and fructose 6-phosphate via subsequent phosphorylation and dissimilation, which finally enters EMP pathway for rest of conversions. The fermentation of 3 mol pentose yields 5 mol pyruvate, 5 mol ATP and 5 mol NADH [22]. Acidogenic phase: in this phase, Clostridia grow exponentially and acids are dominant products leading to drop in pH from 6.5 to around 4.5 [22]. Here, glucose is consumed by Clostridium sp. to produce pyruvate and thus glycolytic pathway is active. The produced pyruvate is further converted to acetyl-CoA, which is main precursor responsible for acid and solvent formation. As can be seen in Figure 8.3, phosphate acetyl transferase and acetate kinase are two key enzymes responsible for acetate production using acetyl-CoA as substrate. However, butyrate formation can be driven by phosphate butyltransferase and butyrate kinase using butyryl-CoA as reactant. Likewise, series of enzymes play a crucial role in forming necessary intermediates which directly or indirectly helps in solvent production. Concomitant with each acid production, one energy molecule (ATP) is generated [31]. The acids produced in this phase are known to permeate through cell membrane which triggers solventogenic shift. Solventogenic phase: as soon as cells sense hostile environment (low pH), phase shift occurs from acidogenesis to solventogenesis [32]. Generally, in this phase no growth was observed while produced acids are re-assimilated into their respective solvents viz. acetone and butanol [33]. This activity may require additional conditions which differ from species to species. For example, C. saccharoperbutylacetonicum N1-4 can able to produce higher

Biobutanol: Research Breakthrough 247 butanol levels only in presence of sufficient amount of glucose in fermentation medium. Further, the sequential conversion forms acetaldehyde and butyraldehyde which can be used for acetone, ethanol and butanol formation [32]. The reduction of butryl-CoA to butanol is mediated by two of dehydrogenases namely butyraldehyde and butanol dehydrogenase [34]. These enzymes are cofactor dependent and hence give their maximum activity in presence of coenzyme NADH and/or NADPH [35]. Literature report is available explaining the dominancy of each one during entire fermentation period [36]. At low pH, NADH-dependent butanol dehydrogenase is more active rather than NADPH-dependent [34]. Besides, ethanol production is quite independent and occurs under certain adverse medium conditions.

8.5 Feedstocks Used in ABE Fermentation Process The major aspect of ABE process is selection of feedstock as it contributes around 60-70% in final product cost and thus affecting economics of biobutanol production [37]. Selection can be done by considering important parameters such as availability, cost of feedstock and its holocellulose content (cellulose and hemicellulose). The possible substitute which fits in aforementioned factors is ‘biomass’. In addition, utilization of biomass as renewable feedstock can generate employment in various sectors and provide incentive to improve rural transportation infrastructure, which would facilitate agricultural and economic development. Besides, huge production of biomass, if made profitable enough can attract farmers to take up the job ensuring continual supply of biomass and good returns to the farmers, thereby providing a boost to agribusiness and rural empowerment [38]. Biomass is the fourth largest source of energy after coal, petroleum, and natural gas [5]. In early days, food based substrates such as corn and starchy materials have been extensively used for biofuel production. However, prolong utilization of such materials will ultimately results into food versus fuel crisis and may be unviable for long term operation. Therefore, renewable, less expensive and readily available sustainable substrates such as forest, agricultural and municipal wastes (second generation feedstocks) are center of attraction in recent years [39]. Until now, various lignocellulosic materials have been utilized for ABE production such as barley straw [40, 41], switchgrass and wheat straw [42], spruce chips [43], corn stover [44], pineapple peel waste, cauliflower waste [45, 46], press mud, pea pod waste [47, 48] and many more. Table 8.2 summarizes the variety of raw materials used by researchers during biobutanol production.

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Liquid Biofuel Production

Table 8.2 Different feedstocks used for butanol production. Feedstock

References

Agricultural residues: Barley straw, Corn stover, Rice straw, Sorghum straw, Wheat straw, Bagasse, Switchgrass, Sweet sorghum bagasse, Corn fiber, Wheat bran, Rice bran and others

[49, 15, 25, 38]

Forest residues: Black locust, Hybrid poplar, Eucalyptus, Spruce, Pine, Yellow poplar, Miscanthus, Alfalfa

[25, 43, 49, 50, 51]

Solid waste: Processed paper, Food waste, Solid cattle manure, Poultry waste, Plastics, Domestic organic waste, Distillers’ dry grain solubles

[49, 52]

Together with lignocellulosic biomass, researchers are also trying to utilize algae as potential substrate for biofuel production. Algae are aquatic photosynthetic microorganisms which can easily and rapidly grow on saline water, coastal seawater, and municipal wastewater or on land incompatible for agriculture and farming [53, 54]. Utilization of algae as feedstock is advantageous in many ways viz. algae generate a high dry weight biomass yield, it has no lignin and low hemicellulose levels, resulting in an increased hydrolysis efficiency, higher fermentation yields and thus reduced cost of product [55]. It also has ability to double their biomass in 2-5 days when compared with other feedstocks harvested once or twice a year. Costa and de Morais [56] used some of microalgal and macroalgal sp. namely Botryococcus, Chlorella and Chlamydomonas, Spirulina platensis for biodiesel and bioethanol production, respectively. Besides, algal biomass has also been used for biohydrogen and biomethane production. However, their utilization in biobutanol production is still at laboratory stage and need more studies to confirm its feasibility at large scale.

8.6 Saccharification and Detoxification Processes Pretreatment and detoxification steps are equally important in overall ABE process, along with fermentation [57]. Lignocellulosic material constitutes cellulose, hemicellulose, and lignin forming a complex structure [38]. To avail holocellulose for its further conversion into simple monomers, usually pretreatment processes are employed. Presence of lignin adds up

Feedstock

Sugarcane bagasse

Wheat straw

Wheat straw

Corn fiber

Switchgrass

Aspen

Wood flour

Hybrid poplar chips

Wheat and rye straw

Pretreatment

Acidic

Alkaline

Steam explosion

Liquid hot water

Ammonia fibre explosion

CO2 explosion

Ionic liquids

Organosolv

Ozonolysis

Ozonated wheat and rye straw under room condition

180°C, 60 min, 1.25% H2SO4 and 60% ethanol

1-ethyl-3-methylimidazolium acetate

Supercritical CO2 at 3100 psi, 112–165°C for 10–60 min

Aqueous ammonia hydroxide 30%, 5 or 10 days

160°C for 20 min

190°C for 10 min, 0.2% H2SO4

[78]

[77]

About 82% of cellulose recovered Yields of up to 88.6 and 57%, respectively

[76]

[75]

[74]

[73]

[72]

[71]

[70]

References

About 40% of lignin removed

Yield of 84.7% sugars

40–50% delignification, Hemicellulose content decreased by 50%

Yield of 74% arabinose and 54% xylose

Recovery of glucose 102% and xylose 96%

8.6% monomeric sugars

24.6 g/L total sugars

2% H2SO4, 122°C, 24 min 2.15% H2O2 (v/v), pH 11.5, 35°C for 24 h

Yield

Pretreatment conditions

Table 8.3 Effect of different pretreatments on variety of feedstocks.

Biobutanol: Research Breakthrough 249

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Liquid Biofuel Production

complexity in process and thus reduces the thermo-chemical yield [58]. Hence, pretreatment is an essential and fundamental step to disrupt lignin structure for successful hydrolysis and downstream operations [59]. At present, many pretreatment technologies have been developed such as acid, alkali, steam explosion, organosolv, biological pretreatment, ionic liquids and others [60]. Among different pretreatments studied, steam explosion has been increasingly considered as one of the cost effective and efficient technologies in biorefinery process [61–63]. Generally, it breaks the rigid structure of biomass to loosen cellulose which can be further used for enzymatic hydrolysis. In respect of this, various researchers studied steam explosion treatment by incorporating mineral acids, organic acids, and Lewis acids viz. sulfuric, hydrochloric, nitric, phosphoric, maleic, and oxalic acids along with ferric chloride and zinc chloride [64, 65] in order to get higher sugar release. Additionally, alkalis such as sodium hydroxide, potassium hydroxide, and ammonium hydroxide have also been used earlier [66–69]. The appropriate pretreatment method is expected to maximize sugar release and minimize the generation of fermentation inhibitors. More importantly, the final sugar yield depends not only on biomass characteristics but also on their interaction with pretreatment conditions. Therefore, setting up pretrement variables is a crucial step in bioprocess. The pretreatment processes can be performed individually and/or in combinations depending on type of feedstock used. Table 8.3 enlist different pretreatment methods available in literature. During hydrolysis process, certain inhibitors such as furan derivatives (furfural and hydroxymethyl furfural), phenolics and weak acids are formed which are toxic to growing microorganisms thereby affecting fermentation process [79–81]. Hence, detoxification of hydrolysates is usually preferred step before fermentation to remove the inhibitors [69]. Different detoxification methods are available in literature viz. over-liming using calcium hydroxide, adsorption by using activated charcoal and ion exchange resins [82, 83]. Each method has its own characteristics and impacts on total sugar.

8.7 Strain Engineering and Developments in Butanol Production Although butanol is known to be an ideal alternative to fossil fuels, the biobutanol production through traditional route is still not economically competitive due to low butanol yield and ineluctable formation of side products [14]. The low butanol yield at the end of fermentation is directly

Biobutanol: Research Breakthrough 251 linked with butanol toxicity to growing host cells. In this view, various attempts have been made to modify host cell (Clostridia) in order to generate robust strains. The resulted engineered host is expected to offer maximum butanol yield, increased selectivity for butanol production along with improved strain tolerance. In addition, metabolic engineering tactics have also been implemented to tentatively engineer non-host cells such as Escherichia coli and Saccharomyces cerevisiae to produce butanol as the desirable end product [84]. Figure 8.4 shows the important technologies of strain development for C. acetobutylicum. A variety of genetic tools and methodologies for Clostridium sp. with depiction of genome sequencing, gene transfer, gene knockout, over-expression, host-vector systems, and advanced genome editing technologies are reviewed and implemented [85]. However, the

(a)

Homologous recombination

(b)

Antisense RNA DNA

Plasmid

mRNA Integration mRNA asRNA

RNA polymerase

Hot cell

(c)

Mobile group II intron

(d)

CRISPR-CAS tool Cas9

Inton RNA Inton Target DNA

gRNA

ItrA LtrA

KO gene

Figure 8.4 Strain development technologies for C. acetobutylicum.

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genetic progression is still lags behind due to difficulty and complexity in transferring related pathways to host bacteria. Initially, Green et al. [86] disrupted butyrate and acetate formation in the metabolic pathway of C. acetobutylicum through integrational plasmid technology which resulted in improved butanol production. Furthermore, antisense RNA technology was also used to determine role of genes related to sporulation and central metabolic pathway [87]. In allele-coupled exchange approach, Heap et al. [88] integrated over 40 kb of lambda DNA into the chromosome of pyrE mutant of C. acetobutylicum to avoid intrinsic instability of an insertional ClosTron mutant. Another possible approach would be targeting key genes for byproduct formation in biosynthetic pathway of C. acetobutylicum in order to divert carbon flux towards butanol and eradicating the unwanted products. Particularly, three genes namely adc, ctfA and ctfB are responsible for conversion of acetoacetyl-CoA to acetone. Cooksley et al. [89] reported no acetone formation in the ctfA and ctfB knockout mutants. Similarly, disruption of several other genes viz. ack, pta, ptb, and buk were undertaken to evaluate the net effect on butanol formation [89–91]. The strain robustness is of great importance in ABE process development especially because of fermentation inhibition by butanol and inhibitory components from the hydrolysate of lignocellulosic biomass. The heat shock proteins (HSPs) are actively up-regulated under various stress conditions [92]. Hence, Mann et al. [93] showed over-expression of HSPs such as GroESL, GrpE and HtpG could facilitate butanol tolerance and adaptation of Clostridial strains. The engineered C. acetobutylicum with over-expression of the bi-cistronic groESL operon exhibited a higher survive rate in 2% (v/v) butanol and a higher butanol production rate signifying that regulation of HSPs may be valuable for the development of more robust phenotypes. Overall, the deep understanding of molecular mechanism through system biology is essential for effective engineering of C. acetobutylicum using modern genome editing toolkits. CRISPR-Cas9 (Clustered regularly interspaced short palindromic) has been explored as a leading-edge tool for genome editing as it provides various attractive features such as high efficiency, wide targeting site distribution, strong adaptability and convenient to use [84]. Especially, most of its applications are in eukaryotic cells ranging from fungi to human cells while very less reports are available in case of bacteria. In recent years, Wang et al. [94] employed this technology in C. beijerinckii NCIMB 8052 for the clean markerless chromosomal deletion of a 262 bp DNA fragment in the spo0A gene. Similarly, Cas9n-based genome editing’s were performed in several Clostridial sp. by some of researchers and reported the

Biobutanol: Research Breakthrough 253 poor transformation efficiency indicating invaluable technology for the Clostridium related research and development [95–101]. Another potential approach is the expression of key genes responsible for butanol formation in non-host cells and thus forcing them to form desired product. In sight of this, Steen et al. [102] introduced the alcohol dehydrogenase gene (adhE2 from C. beijerinckii) and butyryl-CoA dehydrogenase gene (ccr from Streptomyces coelicolor) into S. cerevisiae and meanwhile over-expressed the native thiolase and hydroxy-butyryl-CoA dehydrogenase genes. The resultant strain produced 2.5 mg/L of butanol using galactose as substrate under semi-anaerobic conditions. Branduardi et al. [103] engineered S. cerevisiae in respect of butanol production and achieved 92 mg/L butanol using glycine as a substrate. Besides, some other microbes have also been engineered for butanol production such as Bacillus subtilis [104], Lactobacillus brevis [105], and Pseudomonas putida [104]. However, none of these recombinant strains could produce more than 1 g/L of butanol in a batch fermentation process thus pointing that additional efforts are still require in improving butanol titer. Metabolic engineering also has significant potential to improve solvent yield and productivity by altering metabolic flux towards product of interest. In this approach, mainly enzymes involved in biosynthetic pathways are triggered in order to induce changes in metabolism. Researchers have studied numerous cofactors/pigments/electron carriers/precursors which are responsible for enhancing butanol titer along with B:A ratio [36, 106–110].

8.8 Bioreactor Operations ABE fermentation process can be operated with batch, fed-batch and continuous mode depending on production capacity. Batch and fed-batch modes are more suitable and preferred at small scale operations [8]. Usually, batch fermentation has been widely practiced in industry though it possesses several disadvantages such as long lag-phase, solvent inhibition, and low productivity. However, batch operation is still popular especially because of easy operation and well-developed technology. Fed-batch fermentation is mainly performed in laboratories since it aims to alleviate substrate inhibition [5]. This mode is majorly used to increase product titer by consequently feeding concentrated medium. To avoid end product inhibition, fed-batch operation is recommended to be applied with in-situ product removal technique. On the other hand, continuous fermentation is best choice among others for butanol production at large scale as it gives high productivity with minimum downtime and lag-phase.

acorn

45g/L glycerol + 33.75g/L glucose

Wood pulp fibers as support matrix and sugar mixture as substrate

Sugarcane baggase as matrix

Wood pulp as matrix and sugar mixtures as substrate

Oil palm empty fruit bunch

Yadav et al. 2014 [116]

Survase et al. 2012 [117]

Bankar et al. 2012 [114]

Bankar et al. 2013 [118]

Ibrahim et al. 2015 [119]

Substrate

Heidari et al. 2016 [115]

Author and year of publication

C. acetobutylicum ATCC 824, 96 hrs

C. acetobutylicum DSM 792

C. acetobutylicum B5313

C. acetobutylicum DSM 792

C. acetobutylicum KF158795, 72 hrs

C. acetobutylicum NRRL B-591, 36 hrs

Microrganism and fermentation time

Table 8.4 Literature based on fermentation processes.

Simultaneous saccharification and fermentation

Two stage immobilized column reactor system integrated with liquid– liquid extraction

Two stage chemostat system integrated with liquid–liquid extraction

Continuous

Fed-Batch

Batch

Type of fermentation

Lignin affected sugar consumption

–

20.30

4.45

–

Cell loss

Strain inhibition

Tannic acid from acorn causes inhibition

Problems associated

25.32

12.64

21.86

15.5

ABE (g/L)

254 Liquid Biofuel Production

Biobutanol: Research Breakthrough 255 Researchers have attempted to produce biobutanol through ABE fermentation which is operated under different fermentation modes. Particularly, Isar et al. [111] carried out batch fermentation using Jatropha seed cake as a substrate by C. beijerinckii for 72 h and reported total solvents of 18.6 g/L. Tashiro et al. [112] incorporated butyric acid along with glucose in fedbatch mode and reported 72% higher productivity compared to conventional batch process. Furthermore, Survase et al. [113] initially fermented spent liquor from spruce and reported 8.79 g/L of total solvents at batch mode and later at continuous mode achieved 12 g/L of total solvents with maximum solvent productivity of 4.86 g/L.h. Although continuous fermentation maximizes the solvent productivity, strain degeneration is common problem encounters in continuous process lasted for longer time [8]. This in turn results into reduction in ABE production or even no solvent formation. To overcome this concern, different reactor configurations in continuous system are studied viz. continuous stirred tank reactor with or without immobilization material, packed bed reactor and fluidized bed reactor. In continuous process, cell wash out is also the main concern which can be resolved by either cell recycling or incorporating cell immobilization concept. Bankar et al. [114] studied continuous two-stage ABE fermentation with in situ product removal by immobilizing Clostridium acetobutylicum B 5313 on sugarcane bagasse and achieved highest solvent production of 25.32 g/L with maximum sugar utilization (83.21%). Furthermore, some more reports on fermentation modes are presented in Table 8.4.

8.9 Butanol Separation Techniques Biobutanol production is facing numerous challenges because of butanol toxicity to growing Clostridia that leads to the end product inhibition [120]. Hence, there is an instilling need to find an efficient recovery system which will continuously remove butanol during fermentation. However, butanol separation at low concentration is quite challenging due to high boiling point (117°C) which lead to azeotrope formation thereby affecting butanol economics [121]. Usually, butanol can be traditionally recovered by distillation, liquid liquid extraction (LLE), and adsorption. Nonetheless, distillation is energy-intensive process as it requires 12 tons of steam to recover 1 ton of ABE and thus has been dismissed considering economic viability [5, 122]. Additionally, it is estimated that around 79.5 MJ/kg of energy required to recover butanol, which is much higher than energy content of butanol (36 MJ/kg).

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A variety of alternative methods are available such as gas stripping, pervaporation and perstraction [11, 123]. Ideally, the recovery process should exhibit long term stability, high selectivity, and faster removal rate. Further, it should be simple to perform and have minimum energy requirement so as to make process economically viable. A recent review article by Jiménez-Bonilla and Wang [124] provide perspective on the spectrum of separation methods. Besides, they also explained some of the challenges and limitations of various downstream technologies in terms of recovery, yield, and energy consumption. Moreover, several researchers have different opinions about each method which are discussed subsequently.

8.9.1 Extraction LLE is a conventional method that has been used in commercial industries like chemical and pharmaceutical. It is used to extract target solute from aqueous phase by contacting one or more extractants. Interestingly, LLE has great potential in butanol separation due to hydrophobic nature of butanol. In general, there are two basic categories for separation of butanol from fermentation broth viz. extractive fermentation or in situ extraction wherein solvent extraction module was integrated with the bioreactor and butanol can be extracted continuously, leaving all other components such as nutrients and cells in fermentation broth (Figure 8.5). In situ extraction helps to reduce the end product inhibition which in turn enhances final product yield and productivity. Moreover, the selection of extractant is most crucial parameter in case of in situ extraction and contributes in recovery performance. The primary characteristics of desired extractant are as stated; nontoxic to microorganism, high selectivity, high distribution coefficient, no emulsion formation, high stability, and

CO2

Organic phase

Inlet

Organic phase

ABE

Aqueous phase Extractant

Extractant make up

Bioreactor

Figure 8.5 Block diagram for liquid-liquid extraction of butanol separation.

Biobutanol: Research Breakthrough 257 low solubility in aqueous solution. Additionally, extractant should have other key features such as significant density difference, low viscosity, autoclavability, suitable volatility, and commercial availability at low cost [125]. Besides, ex situ extraction is usually performed at the end of fermentation wherein extractant is mixed with fermentation broth followed by product separation. The two parameters namely distribution coefficient and selectivity are majorly considered in extraction study. The distribution coefficient of component ‘i’ is defined as the ratio of concentration of component ‘i’ in organic phase to the concentration in aqueous phase at extraction equilibrium and equation is as follows [121]:

KDi

Ci , OP Ci , AP

(1)

where, the subscript OP and AP represents the organic and aqueous phases, respectively. ‘i’ can be butanol, acetone, ethanol or water. The distribution coefficient (KD) of component ‘i’ can also be expressed by following equation:

K' Di

Wi , OP Wi , AP

(2)

where, W is weight percent (wt %) or mass fraction (g/g) of solute. The relationship between KD and K’D can be derived as follows:

KDi

OP

K 'Di

(3)

AP

where, ρ is density of organic phase (OP) or aqueous phase (AP). Assuming the density of organic and aqueous phase is being close to the density of extractant and water, respectively. Equation (3) can be simplified as:

KDi

Ex

H 2O

K 'Di

(4)

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Liquid Biofuel Production

If the density of extractant is very close to that of water, then KD and K’D is equal for a given extraction system. The selectivity (Si) for the solute ‘i’ (to be extracted e.g. butanol) over water is defined by the ratio of the distribution coefficient of the component ‘i’ to the distribution coefficient of water:

Si

KDi KD , H 2O

(5)

Many extractants have been investigated by researchers in respect of efficient butanol separation. These includes; oleyl alcohol [126], decanol [114], bio-based lactate esters, biodiesel [127], methylated crude palm oil, iso-octane [128], heptanol, octanol [129], 4-n-butylphenol, and methoxy (methoxymethoxy) methane [130]. Unfortunately, the extractants with high distribution coefficient for butanol are toxic to microorganism while non-toxic one’s have low distribution coefficient. Among various extractants studied, decanol shows the highest distribution coefficient but toxic to the microorganism. Indeed, oleyl alcohol is most widely studied due to its non-toxic nature but having low distribution coefficient [131]. Roffler et al. [132] used mixed extractant namely benzyl benzoate and oleyl alcohol for solvent separation and reported enhanced productivity by 60%. The enhanced butanol production was achieved using 20% decanol in oleyl alcohol integrated with two stage continuous fermentation [114, 118]. González-Peñas et al. [133] screened 16 compounds from different chemical families and used in extraction study with due emphasis on certain biological parameters. Among the tested extractants, 2-butyl-1-octanol showed the best extracting characteristics with the highest partition coefficient (6.76), highest selectivity (644), and biocompatibility with C. acetobutylicum. Further, the traditional fermentation by C. acetobutylicum ATCC 824 produced 21.6 g/L ABE while extractive (2-butyl-1-octanol as extractant) fermentation produced 36.6 g/L ABE [134]. The novel dual extraction for ABE fermentation has been reported by Kurkijärvi and Lehtonen [129] using 9-11 carbon alcohols in conjugation with alkanes of the same size and showed the highest distribution coefficient of butanol to be 7.17 for decanol. Ionic liquids (ILs) have recently gained much attention as alternatives to organic solvents for LLE. They are potent extractants for removal of butanol from dilute solution but main drawbacks are their viscosity and third phase formation which restricts their application for in situ extraction [133]. Several researchers showed some ILs have high distribution

Biobutanol: Research Breakthrough 259 coefficient and selectivities in between 25-300 [135–137]. The butanol distribution coefficients of ILs are highly dependent on the hydrophobicity of anions followed by the hydrophobicity of cations. Moreover, Ha et al. [138] investigated the extraction behavior of different imidazolium-based ionic liquids for butanol extraction. Domańska and Wlazło [135] estimated the experimental limiting activity coefficient for 65 solutes over the six temperatures (from 318.15 K to 368.15 K) and further used to calculate the thermodynamic properties. The activity coefficient data was also used to calculate the selectivity and capacity of butanol/water separation in ILs. Kubiczek et al. [139] studied the liquid-liquid equilibrium of n-butanol from water using ILs as solvent and simulated the single and multistage extraction model using non-random two liquid (NRTL) equations. Kubiczek and Kamiński [137] reviewed the current research concerning the application of different ILs in LLE. Furthermore, Domańska and Wlazło [135] presented ternary liquid-liquid equilibrium data for mixture of ionic liquid + butanol + water at 298.15 K and monitored the effect of ILs in n-butanol extraction. Additionally, the effect of cations using experimental and NRTL model was revealed to correlate the experimental tie-lines. The ternary compositions were predicted with good precision using COSMO-RS. Besides, there are number of parameters such as minimum solvent requirement, extraction efficiencies, extraction factors, number of theoretical stages, and mass transfer characteristic that need to be studied in order to develop continuous LLE system for better recovery of desired product.

8.9.2 Gas Stripping Gas stripping is a physical separation process that allows the selective separation of ABE solvents by continuously bubbling gas into fermentation broth where the gas stream acts as a separating agent. Further, the process is driven by vapor-liquid equilibrium [140, 141]. The ABE solvents from stripped gas are recovered by condensation in a cold trap or condenser [142]. The exit gas is again circulated back to bioreactor and process of removal continued until the fermentation ends. A schematic diagram of the gas stripping is as shown in Figure 8.6. Ennis et al. [143] demonstrated that gas stripping is one of the most efficient techniques for in situ butanol removal during ABE fermentation. Generally, ABE fermentation is escorted via generation of gases (CO2 and H2) which can be used as a separating agent for butanol recovery. Furthermore, gas stripping proves to be a reliable method because of its versatility, simplicity,

260

Liquid Biofuel Production CO2, H2, ABE

Feed

Bioreactor Condenser

Make-up gas

ABE Gas recycle

Figure 8.6 Integrated fed batch fermentation with gas stripping process.

low cost for equipment investment, and no harm to growing cells [121]. However, several researchers studied key parameters viz. gas flow rate, bubble size, agitation speed and product concentration that need to be taken into account for effective separation [40, 97, 144, 145]. Besides, product recovery is also dependable on type of gases used for stripping [97]. Rochón et al. [142] proposed the model for integrated process of gas stripping using MATLAB software and reported best fit of experimental data with model predictions. Moreover, Li et al. [97] reviewed the gas stripping simulation and explained that in first order kinetics, stripping rate of solute is proportional to its concentration in the bulk phase. Overall, increased productivity and substrate utilization can be achieved by gas stripping [125]. The highest butanol recovery using double gas trap or a double stripping process has been reported to be 515.3 g/L [8], 175.6-420.4 g/L [123], and 441.7 g/L [146]. This process can also be operated in combination with other recovery systems such as membrane separation processes [125].

8.9.3 Pervaporation Pervaporation is a separation process where ideal membrane should be selectively permeable to the solvent when it get contacted with fermentation broth and permeate is removed subsequently by evaporation on the other side connected to a vacuum system at low pressure (Figure 8.7). The  mechanism involved in transport of solute is selective sorption,

Biobutanol: Research Breakthrough 261 Feed

CO2 8 7 5

6 1

2

3

4

Figure 8.7 Block diagram of integrated pervaporation with ABE fermentation 1: Fermenter; 2: Ultra Filtration (UF) membrane; 3: Hydrophobic PV condenser; 4: ABE solution; 5: Vaccum pump; 6: ABE enriched permeate, 7: Cell; 8: Retentate.

diffusion through membrane, and desorption into vapour phase [147]. Membrane pervaporation is cost effective, selective, energy efficient, commercially competitive, and gives cleaner end product without use of entrainer [148]. However, the drawbacks are membrane fouling, clogging, risk associated with reliability of the process, and biocompatibility [121]. The separation using pervaporation are influenced by various important factors such as membrane swelling, concentration polarization, permeate membrane interaction, molecular mass, size, shape, and flux (low fluxes reduces the efficiency). The effectiveness of technique is determined by two parameters: separation factor (measure of selective removal of volatiles) and membrane flux (the rate at which an organic/volatile passes through the membrane/m2 membrane area). The separation factor is given as follows:

WBuOH / WH 2O

P

WBuOH / WH 2O

F

(6)

where, WBuOH and WH2O are concentrations of butanol and water (wt%) in permeate (P) and feed (F), respectively. Separation factor (β) and membrane flux depends on feed temperature, feed composition, membrane material, thickness of the membrane, and vacuum or sweep gas partial pressure. Besides, silicone membrane has been well explored for butanol separation as it is commercially available at low cost, and offers easy manipulation [149]. Additionally, other membranes have been investigated including polypropylene, polystyrene-b-polydimethylsiloxane-bpolystyrene, oleyl alcohol liquid membrane on a polypropylene support, and PDMS-supported ionic liquid membranes [136].

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Liquid Biofuel Production

Recently, the utilization of composite membranes has been popular which showed improved selectivity and flux performance. These are as follows; zeolite-mixed PDMS, silicalite-PDMS/polyacrylonitrile membrane, silicalite-silicone composite membrane, PDMS/ceramic composite, and carbon nanotube filled PDMS [11]. Qureshi and Blaschek [149] had given comprehensive compilation of various membranes and their properties for recovery of butanol from fermentation broth/model solution. Further, the selection of membrane affects the selectivity and diffusion rate thereby deciding the final product concentration in permeate. Indeed, decrease in membrane flux is also one of the problems associated with membrane separation. Flux improvement can be made through the use of higher temperature feed stream but it requires the additional step of microorganism removal. In addition, Bharathiraja et al. [11] reviewed the effect of ultrasound irradiation on pervaporation (using silicone tubing as membrane) and reported improvement in butanol concentration from 203 to 221 g/L in permeate. Fadeev et al. [150] also showed high selectivity and permeate flux using PDMS and PTMSP membranes but later observed decrement in flux due to membrane fouling. Even silicates PDMS membranes were combined to check its performance [125, 151].

8.9.4 Perstraction Perstraction is membrane assisted technique basically combines the principles of LLE with pervaporation in one unit operation. Perstraction allow the selective removal of volatile solvent from the fermentation broth with the help of membrane. In this process, membrane is placed between feed and extractant side wherein butanol get diffused through the membrane and then extracted by extractant (permeate side is organic phase). The nutrients and other components such as acetic or butyric acid retains in the aqueous phase itself [152, 153] (Figure 8.8). There is no direct contact between two immiscible phases and thus this technique overcomes several limitations of LLE process. Another notable trait of this system is independent control over flow rate of broth and extractant that further support butanol diffusion preferentially across the membrane. Certainly, butanol diffusion rate depends entirely on the membrane which may acts as physical barrier. Therefore, the membrane should be selected in such way that it should facilitate the butanol movement toward organic phase. The membrane can either be hydrophilic or hydrophobic with an arrangement of loop of hollow fiber membranes in series [11]. Total mass transfer is determined by the sum of mass transfer coefficient in the aqueous phase, extractant phase, and also the mass transfer

Biobutanol: Research Breakthrough 263

ABE analysis

CO2 Feed

Ionic liquid

Feed

Figure 8.8 Block diagram of perstraction process integrated with ABE fermentation.

coefficient of the membrane. The overall mass transfer resistance can be reduced by optimizing membrane properties and changing the process flow parameter so that to maintain constant velocities through fiber. Tanaka et al. [154] used perstraction technique for butanol removal and reported enhanced butanol productivity (78.6 g/L.h). Qureshi and Maddox [155] extracted butanol by perstraction wherein silicone membrane and oleyl alcohol (as an extractant) were used. Chen et al. [156] reported minimum energy consumption and zero water flux across the membrane during perstraction process. Shukla et al. [157] stated gas generation in the shell side of the fermentor during perstraction that can be resolved by increasing the number of fiber tubes in the membrane. Merlet et al. [158] implemented hydrophobic ILs as extractant during perstraction and reported effective butanol separation using [omim][Tf2N] with low flux (4.3×10-3 kg h-1 m-2).

8.9.5 Adsorption This is one of the most energy efficient traditional approaches that include adsorption of volatile compound on to the surface of resins. It was first investigated for in situ product recovery in ABE fermentation to relieve butanol inhibition. Mainly, the cell free permeate is required for adsorption and that can be obtained by UF. Subsequently, permeate enter the adsorption column containing the adsorbent to adsorb butanol from aqueous solution and remaining liquid is recycled to the fermenter. The use of UF membrane prior to adsorption helps to avoid the possible fouling of adsorbent and cells loss which further retains high cell concentration in the fermenter (Figure 8.9).

264

Liquid Biofuel Production ABE exhausted phase Gases Feed ABE rich phase

Cell

Bioreactor UF membrane

Adsorption/desorption column

Figure 8.9 Integrated adsorption process for ABE separation.

Literature reports show that hydrophobic resins are most commonly utilized for butanol separation from aqueous solution based on rule of ‘like attract like’. Different type of adsorbent have been explored which includes molecular sieve, activated carbon, zeolites, composite such as calixarene-based adsorbents and polymer resins such as XAD-4, XAD-16, polyvinylpyridine, poly(styrene-co-divinylbenzene)-derived resins Dowex Optipore L-493 and SD-2, Dowex M43, and Diaion HP-20 [159–163]. Researchers used partition coefficient (Ks/w) as major factor in preliminary screening of adsorbents. However, it is applicable only when the solute concentration is well below saturation, some adsorbents behave close to linearity while others do not. Another approach could be estimation of loading capacity (LBuOH) in order to calculate maximum binding of the solute at saturation of adsorbent. Adsorption was commonly studied using Langmuir and Freundlich isotherm. Langmuir model is extensively used in adsorption process [164–166]. The Langmuir model (Equation 7) is applicable when following assumptions are met: adsorption should be a monolayer; solution behavior is ideal; adsorption sites have the same affinity; adsorbed molecules are localized; there are no lateral interactions, and adsorbed molecules are in dynamic equilibrium [164].

q

q max BCeq 1 BCeq

(7)

where, q is the adsorption capacity, qmax is the maximum adsorption capacity, B is the Langmuir constant, Ceq is the equilibrium solute concentration

Biobutanol: Research Breakthrough 265 in liquid phase. qmax is equivalent to loading capacity at saturation, and the Langmuir constant (B) is analogous to Ks/w. B describes the affinity of the adsorbent and the adsorbate. B and qmax can be obtained from the mathematic linearization of Langmuir model. Another model, Freundlich isotherm empirical model usually fits the adsorption behavior better than Langmuir without complex calculations [124]. The model is expressed in Equation (8) 1/n q Kf Ceq

(8)

where, Kf and n are Freundlich constants. The equation does not indicate a finite uptake capacity and thus it is functional in the low-to-medium concentration ranges. If n = 1, the expression becomes linear since Kf = Kw/s. So, n is related to the deviation from this ideal behavior caused by the heterogeneity of the surface adsorption sites. When 1/n is close to zero, the surface is highly heterogeneous as reported by Jiménez-Bonilla and Wang [124]. Therefore, Kf is an improved Kw/s and represents the quantity of adsorbate in the solid required to maintain at one unit for the concentration in the solution (mmol/L). Consequently, Kf is also related to the adsorption capacity [167]. From Freundlich model analysis, activated carbon showed the highest Kf , followed by other adsorbents such as KA-I and finally Optipore L 493 and SD2 while the relatively low Kf was demonstrated for Diaion HP20, HP2MG and Hytrel 8206 [168]. Other adsorption models are also available like Brunauer, Emmett and Teller (BET) isotherm and can usually fit better for the experimental data [124]. However, they are not widely used due to complexity in representing their constants. A recent study by Xue et al. [165] revealed very high qmax for active carbon (Norit Row 0.8, AC F600 and AC F400) in the range of 150-280 mg/g of butanol. Further, Farzaneh et al. [164] compared the qmax of zeolites, silicalite, and polystyrene adsorbents and showed the same trend as LBuOH. The two commercial zeolites ZSM-5 with high Si/Al ratio, CBV28014, and CBV901 as adsorbents were examined by Saravanan et al. [169]. Oudshoorn et al. [170] conducted desorption of butanol from resin and showed desorption heat required for CBV901 to be 1080 J/g of butanol.

8.9.6 Hybrid Separation Process Different separation technologies discussed in previous sections have certain pros and cons when operated individually as no process or unit operation is 100% efficient (171, 172). Hence, the attention has been brought to

266

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hybrid technology in which fermentation process can be integrated with more than one recovery technique. More importantly, a focus has been given to improve the efficiency of each process involved in hybrid separation systems. Although, innovative hybrid systems possesses a leap in energy efficiency compared to distillation, the specific energy requirement is still considerably higher than 10% of the energy content of butanol [173]. Groot et al. [174] demonstrated that around 3.67 MJ/kg of energy would be required to recover butanol when combination of beer stripper and a three-phase distillation system was employed. On the other hand, Matsumura et al. [175] reported that the hybrid combination of pervaporation and distillation was more efficient (7.4 MJ/kg of energy consumption) as compared to simple distillation (79.5 MJ/kg). Similarly, a combination of silicate adsorption and distillation was used which required 9.0 MJ/kg of energy to recover butanol from 0.5 wt% ABE solution [176]. Oudshoorn et al. [177] stated a hybrid system wherein adsorption is coupled with pervaporation that even consumes less energy for efficient recovery. Interestingly, an innovative two-stage gas stripping process with a fibrous bed bioreactor was developed by Xue et al. [8] to achieve higher concentration of butanol. Moreover, Butamax - the butanol processing industry patented the butanol recovery process in which extraction using oleyl alcohol is coupled with gas stripping, a hybridized method for efficient in situ product recovery [178]. Cost reduction in case of hybrid separation can be done by selection of suitable recovery technique that would be more efficient and operate at low cost. Besides, the 2% increase in volumetric productivity could reduce approximately 20% of capital expenditure and significant reduction in operation cost [179]. Liu et al. [180] and Van der Merwe et al. [181] suggested the flow diagram with alternative product concentration techniques to distillation. It would be beneficial to do a similar analysis for the various hybrid separation processes that will helps to decide best suited recovery process in an industrial point of view.

8.10 Techno-Economic Assessment ABE fermentation is not technically and economically viable due to various unsolved challenges such as severe product (especially butanol) inhibition during fermentation. This leads to low product concentration (< 3 wt%) and thus low yield and productivity thereby increasing the downstream process costs [121, 182, 183]. To minimize entire production cost, unconventional feedstocks such as pineapple waste, cauliflower waste, peas pod,

Biobutanol: Research Breakthrough 267 and sweet sorghum juice are utilized for biobutanol production. Generally, the overall process is simulated using a mass and energy balance which is then used to calculate the economic parameters. These parameters can be used to evaluate the commercial viability of any production process. The absolute and theoretical limits to the process economics are taken into account during techno-economic analysis. The theoretical mass and energy yield were calculated on the basis of product ratio and energy combustions in ABE fermentation [184]. The estimated mass balance for ABE fermentation process is about 0.11 kg n-butanol per kg of corn which is substantially low as compared to ethanol fermentation. Besides, ethanol titer reaches about 15% while butanol reaches to 2%. Hence there is an extensive need to improve the butanol yield by economic way, to foresee its future commercialization [5]. The economics of biobutanol production was calculated by Baral and Shah [185]. The net butanol production cost was 1.7-fold higher ($1.5/L) while the total byproduct credit assigned was 2-fold less ($0.29/L butanol). These variations may be due to feedstock price, different ABE fermentation systems and product yields, neglecting credit of ethanol, and analysis year. Further, estimated butanol production costs were $1.8/L and $1.5/L without and with byproduct credits. However, key influencing parameters for butanol cost are butanol recovery followed by sugar utilization in the fermentation, feedstock cost, corn stover to sugar conversion rate, and heat recovery. Furthermore, optimizing these sensitive operating parameters could reduce the butanol production cost to $0.6/L that is competitive with current gasoline prices. Therefore, further research and development efforts in the ABE fermentation are needed to compete with other liquid fuels. Kumar et al. [186] estimated butanol production cost to be $0.48/L using corn stover as carbon source which is comparatively lower. Malmierca et al. [187] projected butanol production cost to be 1.09 €/kg with an investment of 186 M€ for 27 kt/y of butanol for fermentation process integrated with pervaporation. Another influencing parameter for the economic assessment of the ABE fermentation process is investment cost. The initial large capital investment has a huge impact on the overall economics of the process. In addition, investment cost is directly proportional to the total cost both as depreciation and for financing (interest and repayments). Hence, major reductions in the investment cost and/or delaying investments especially until the production starts will help to improve the process economics [5]. Simulation software’s such as Aspen plus, CHEMCAD can be used to perform the material and energy balance and to estimate production and investment cost [187]. It was also used to calculate the fixed (labor and supplies) and variable (raw materials) operating costs of the plant.

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The overall production cost for non-existing or noncommercial biofuel processes (bioethanol, biobutanol) are less readily available and largely detailed with techno-economic model evaluations. An identical economic assumption reveals that, the product yield on substrate of corn ethanol is almost twice compared to corn butanol. However acetone, ethanol, and hydrogen co-products in corn butanol process contribute more significantly to the net production cost as compared to the corn ethanol process [5]. This result in an overall production cost of butanol to be $0.52/L compared to $0.404/L of ethanol produced from corn [188]. Besides, the total project investment is also doubled for the corn butanol process as compared to the corn ethanol. This is because of the low feedstock yield, and a more complicated separation of ABE mixtures. The economic feasibility of ABE fermentation can be turned up via improvement in butanol tolerance of strain and selective in situ solvent removal. The use of the two-stage continuous solvent-producing cultures immobilized on biomass achieves higher solvent productivities with improved substrate consumption and reduced the solvent toxicity problem [118]. These processes can further enhance the productivity and other economic viability aspects of the process and reduces the fermentation time [189]. The economics of butanol fermentation depends on the product market which is another effectual factor. Based on market statistics, it is expected that after adapting to biobutanol as alternate liquid fuel, the market demands will be high thus influencing product economics.

8.11 Current Status and Future Prospective Biobutanol has been a potent alternative liquid biofuel. Therefore, sustainable biochemical production of butanol is vital for reducing the environmental and social impacts caused due to forthcoming depletion of fossil fuels. In recent years, significant efforts have been made in commercial production of biobutanol or retrofitting same existing bioethanol plants for butanol production [178]. A comparative evaluation among current butanol industries is listed in Table 8.5. Incidentally, to attain profitable butanol production, requires a multidisciplinary approach which can coordinate the most recent advances from different sectors such as environmental science, biochemistry, microbiology, genetic engineering, chemical engineering, process technology and market economics. Tremendous research has taken place in finding economic alternate substrates, metabolic engineering of Clostridial strains, efficient methods of in-situ product removal, and design of fermentation bioreactors.

Genetically optimized yeast Engineered yeast or E.coli

Corn

Non-food feedstock

Sugar, starch, molasses, corn by products and cellulosic feedstocks

Forestry, agricultural Waste

C5/C6 sugars and lignocellulose

Corn

Cathay Industrial Biotech

Cobalt Technologies

Green Biologics Ltd.

Syntec Biofuel

Butalco

Butamax Advanced Technologies

–

Solventogenic Clostridium

Non-GMO Clostridium

–

Modified strain of E. coli

Corn, sugar and beets

Gevo

Microorganism

Feedstock

Butanol industry

–

–

110 gal/t

1.3–1.9 Times more butanol per weight of raw material

1.5million GPY

21million GPY

1million gallon per year (GPY)

Estimated production capacity

Table 8.5 Comparative evaluation and a sneak peek into current butanol industries.

–

Improving existing industrial production strains

Catalyst development for converting cellulosic biomass to butanol using thermo-chemical process

ABE fermentation; metabolic engineering of Clostridium and Geobacillus

–

Anaerobic fermentation

Gevo Integrated Fermentation Technology

Approach

Biobutanol: Research Breakthrough 269

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However, most of these studies are at laboratory scale and any change in one or more steps creates new challenge to meet sustainable production. Basic principles of chemical and biochemical engineering can be used to simulate ABE process with respect to design, optimization, and scale-up. The mathematical models can explore the mechanism of the process with key influencing parameters. Re-commercialization of biobutanol has been linked to the utilization of low-cost feedstock by hyper-butanol producing and high solvent tolerant microbial strains. A hybrid separation along with fermentation can be best suited option to achieve efficient continuous substrate utilization at industrial scale in the near future.

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9 Potential and Prospects of Biobutanol Production from Agricultural Residues Shuvashish Behera1*, Koushalya S2, Sachin Kumar1 and Jafar Ali B M2 1

Biochemical Conversion Division, Sardar Swaran Singh National Institute of Bio- Energy, Kapurthala, India 2 Centre for Green Energy Technology, Pondicherry University, Pondicherry, India

Abstract During the past few decades, there has been a remarkable increase in research on biological butanol production using different waste biomass. Waste agricultural residue is the potential candidate among waste biomass for butanol production and the leading option among alternatives to petroleum-derived transportation fuels due to its potential sustainability. The production of butanol through bacterial acetone-butanol-ethanol (ABE) fermentation has generated considerable research interest. Mostly, two bacterial species i.e., Clostridium beijerinckii, C. acetobutylicum have been identified as the potential butanol producers. The novel fermentation processes are being developed through advanced biotechnology and process engineering for converting the abundantly available agricultural biomass to butanol. However, the performance of butanol fermentation process using clostridial species is limited due to high cost of production, low inhibitor tolerance, strict anaerobic in nature and bacterial sluggish growth. The present chapter is focused on the bacterial advances in conversion of agricultural feedstocks to biobutanol through fermentation technologies with a focus on pathway engineering in bacteria to enhance solvent and inhibitor tolerance. Keywords: Biobutanol, agricultural residues, Clostridium, ABE

*Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (285–318) © 2019 Scrivener Publishing LLC

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9.1 Introduction Energy crisis and resource scarcity are the leading concerns in sustainable economic development around the world due to rapid outgrowth of population and urbanization. The major energy resources consumed by mankind are fossil fuel (coal, oil and natural gas) which is the decomposed residues of buried dead organisms over many centuries [1]. Environmental risks over fossil fuel have been increased as it emits greenhouse gases (GHGs) predominantly carbon dioxide [2–4]. The overconsumption of fossil fuel encounters depletion in its availability, increased oil prices and global climate changes that made the research focus on renewable sources. Recent trends include improved alternative technologies with renewable energy such as wind, geothermal, solar and bioenergy [5–7]. Bioenergy is a green energy technology derived from renewable feedstock (biomass or agricultural residues) of biological origin that imparts a low level of carbon footprints compared to conventional technology [8,  9]. Biofuel obtained from cellulosic biomass of non-edible source is more encouraged to avoid the reduction in food security under biorefinery concept such as biodiesel, bioethanol, biobutanol and biohydrogen [5, 10]. Relative to conventional butanol, biobutanol can be produced sustainably through fermentation; it is a clean fuel and qualifies under the renewable fuel standard [3]. Biobutanol production through acetone-butanol-ethanol (ABE) fermentation using some microbial species has regained much attention recently; it has certain applications as a solvent for the production of antibiotics, vitamins, hormones, inorganic synthesis, chemical intermediate, processing of paint thinner and hydraulic and brake fluids [11–15]. It is completely miscible with common organic solvents but sparingly soluble in water; moreover it can be used as gasoline without any engine modification [3, 11]. Agricultural residues such as rice straw, barley straw bagasse and other forestry residues represent a tremendous source of readily available biomass for biofuel production [16, 17]. These biomasses are rich in carbohydrate which can be utilized for the production of solvents under strict anaerobic condition using desired microbes [18]. In general, Butanologens are unable to degrade the agricultural biomass to monomeric sugars. Due to the close association of cellulose and hemicellulose with lignin in the plant cell wall, pretreatment is necessary to make these carbohydrates available for enzymatic hydrolysis and fermentation [19]. Hence, agricultural biomass requires pre-treatment and subsequent saccharification for release of fermentable sugars prior to acetone-butanol-ethanol (ABE) fermentation

Potential of Biobutanol Production 287 and then to solvents [11, 20–22]. Several modes of ABE fermentation processes such as batch, fed-batch and continuous have been implicated by the researchers for the biobutanol production which released acetate, butyrate, ethanol, acetone as the byproduct [21, 23, 24]. There are certain major challenges in butanol production, i.e., anaerobic nature, slow growth rate of microorganisms, availability of compatible feedstocks, rapid shifts of pH, sensitivity to acetic acid and butyric acid, low butanol titer, solvent tolerance, and product inhibition [25–27]. Therefore, the current chapter aims to discuss the utilization of various agricultural waste biomass for biobutanol production, development of process technologies through the involvement of strict anaerobic bacteria and challenges associated with the ABE fermentation.

9.2 Agricultural Residues In general, agricultural residues are the organic materials that are produced as a by-product from harvesting and processing of agricultural crops and serve as a potential alternative to conventional substrates for biofuel [16]. They can be classified into crop and agro-industrial residues [28]. Crop residues are plant residues that remain in the crop field after harvesting, whereas agro-industrial residues are the by-products of the post-harvest process like cleaning, sieving and milling. Availability of crop residues is very low since difficulty in collection which has other application as fertilizers or animal feed. Agro-industrial residues used as a captive energy source due to its large collection at processing site and also involve low handling cost and transportation [29]. It also prevents the usage of high cost substrates which impact food security [21, 24]. The agricultural residues differ in their cell wall composition as given in Table 9.1, consisting of cellulose, hemicellulose and lignin which are broken down into monomers through biochemical conversion. Both the domestic and industrial sector widely involve use of biomass derived from agricultural residues such as husks, straws, shells or bagasse, etc., for useful energy production [30–32]. The vital sector involves production of liquid biofuels like bioethanol, biodiesel, biobutanol or other solvents through fermentation using microbes [27]. In specific, butanol can be produced through anaerobic fermentation of various carbohydrates such as sucrose, cellulose, glucose, fructose, mannose, lactose, dextrin, starch, xylose and arabinose using microbes [8]. For instance, Lopez-Contreras et al. [33] utilized carbohydrate obtained from domestic organic waste as the substrate

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Table 9.1 Agricultural residues with its cell wall composition (% of dry material). Agricultural residues

Components

Raw material Source

Cellulose

Hemicellulose Lignin Others Reference

Husk

Rice

21.5

23.1

14.6

–

[31]

Straw

Rice

43.4

22.9

17.2

1.4

[34]

Sugarcane 40.8

30.8

25.8

2.6

[35]

Wheat

33

33

20

5.7

[36]

Sorghum

40.4

35.5

3.9

0.2

[37]

Sugarcane 42.8

25.9

22.1

1.4

[38]

Bagasse

for butanol production by C. acetobutylicum ATCC 824 and butanol production of 3 g l-1 and productivity of 0.03 g l-1h-1 was obtained from 100 g of domestic waste. This was further improved to 4.2 g l-1 and 0.09 g l-1 h-1 respectively, when substrate was hydrolyzed with enzyme such as cellulose and β-glucosidase. Similarly, various agriculture residues used for the solvent production has been described as below.

9.2.1 Husk Husk is the external covering of seeds such as rice, nuts, or oats, etc., that mainly contributed as a biofuel producer. Specifically, rice husk was found in abundance as the by-product from the rice-mill industries and these unused leftovers were burnt in open air which may cause environmental pollution. It can be converted to efficient fuels by employing different processing technology. However, it is unaccustomed as animal fodder due to its low digestibility, abnormal size, high silica content and low bulk density [39]. The cell wall of rick husk was composed of certain polymers resembling 21.5% of cellulose, 23.1% hemicellulose, 14.6% lignin that are degraded and fermented to yield biofuel [31]. Based on this aspect, a kinetic investigation on hydrolysis of rice husk was made with different concentrations of sulphuric acid catalyst for producing sugars [31]. Subsequently, there is evidence of producing approximately 0.426 g of sugars which yields 0.21 g of ethanol per 1g of dry rice husk through fermentation [40]. Besides, coffee husk gets aggregated in ample amount during its processing site which may cause environmental pollution with its direct disposal. It could be effectively used to yield good quality of liquid fuels such as 69% bioethanol from 100g of coffee husk powder [41]. Interestingly, hazelnut

Potential of Biobutanol Production 289 husk residue has been found to be appropriate for producing solid fuels with pelleting technology [42]. In addition, oat husks were also regarded as agricultural substrate for fermentation process due to its uniformity in composition and availability. It can be saccharified through alkaline pretreatment for producing monomers to yield biofuels [43]. Utilization of husk for biofuel production can be more encouraged in near future.

9.2.2 Straw Straw is a dry stalk of plants after the grain has been removed from crops such as barley, rice, oats, and rye [44]. There are mainly three straws that can be applied for the bio-chemical conversion technology due to their excess availability from the processing site, such as rice, wheat and barley, respectively. Wheat straw was found in abundance and possesses more cellulose content [16] whereas rice straw has more volatile content which can contribute to yield biofuel with high heating value [32] and barley straw follows a similar process as wheat straw does and accounted for minor research on it [45]. The fermentation technology using these raw straws has more chance of encountering certain inhibitory effects, and its preventive measures were discussed in this section.

9.2.2.1 Wheat Straw Wheat straw is a novel substrate for ABE fermentation and can serve as a low-cost feedstock for biobutanol production which tends to have higher yield residues than barley [16, 21, 24]. Its lignin content is comparable to woods and contains high cellulose content among all other agricultural residues [16]. In a study, simultaneous hydrolysis and fermentation of wheat straw produced butanol productivity of 0.27 g l-1 h-1 that was close to the result obtained through control experiment using glucose substrate (0.30 g l-1 h-1) [46]. Furthermore, improved technologies which involved pretreatment of wheat straw with 1% (v/v) sulphuric acid followed by enzymatic hydrolysis and fermentation using Clostridium beijerinckii P260 resulted in 12.0 g l-1 of butanol with 0.29 g l-1 h-1 productivity and 0.29 yield without any inhibitor formation which was more than the conventional substrate, glucose [47]. These studies demonstrated that hydrolyzates of agricultural residues undergo fermentation for butanol production with no inhibitory effects to microbes. In other investigations, ABE production from wheat straw using Clostridium beijerinckii P260 was successful through five combinations of pretreatments of substrate and fermentation processes [21].

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The combination includes pretreatment of wheat straw prior to fermentation, separate hydrolysis and fermentation (SHF) of wheat straw without removal of sediments, SHF without agitation, SHF with sugar addition and SHF with agitation by gas stripping, i.e., simultaneous hydrolysis of wheat straw followed by fermentation with agitation from gas stripping. Among which SHF with agitation by gas stripping resulted in higher productivity as 0.30 g l-1 h-1 in batch reactor [21] and it was increased to 0.36 g l-1 h-1 in fed batch reactor through addition of sugar during the fermentation process [24]. The performance of the fermentation process can further be improved through desalting and detoxifying of wheat straw that reduce the inhibitory effects [48]. Recent research on economic evaluation in conversion of wheat straw to butanol showed the use of membrane recovery process such as membrane pervaporation for butanol removal from ABE fermentation broth which was more recommended to reduce butanol producing cost compared to distillation recovery process [49].

9.2.2.2 Rice Straw The surplus availability of rice straw or cereal straw has promoted the research to focus on biofuel production. It contains low concentration of lignin with volatile matter which was higher than wood and much higher than coal as well as fixed carbon content is low that makes it more suitable towards bio-chemical conversion [50]. Soni et al. [50] used mixed microbial culture such as cellulolytic fungi Trichoderma reesei and Aspergillus wentii for the biological treatment of rice straw to release sugar which further involves C. saccharoperbutylacetonicum for batch fermentation process to ABE yield of 0.28 with productivity of 0.15 g l-1 h-1. In another study, biobutanol production using rice straw was done through alkaline pretreatment, enzymatic hydrolysis followed by fermentation processes like separate hydrolysis and fermentation (SHF) and combination of SHF with simultaneous saccharification fermentation (SSF). In SHF, butanol productivity was 1.41 g l-1 d-1 whereas SHF-SSF process resulted in maximum butanol productivity of 0.86 g l-1 d-1. On the other hand, rice straw as feedstock could be utilized to obtain reasonable butanol production [18]. Additionally, detoxified rice straw hydrolysate supplemented with yeast extract and calcium carbonate using C. sporogenes BE01 resulted in butanol yield of 4.46 g l-1 and productivity of 0.05 g l-1 h-1 [32]. An investigation on pretreatment of rice straw was evaluated with concentrated phosphoric acid and alkaline NaOH treatment prior to enzymatic hydrolysis followed by fermentation using C. acetobutylicum. Alkali treatment of rice straw resulted in 45.2 g of butanol from 163.5 g of glucose as well as concentrated phosphoric acid yields 44.2 g

Potential of Biobutanol Production 291 of butanol from 192.3 g glucose [51]. Amiri et al. [52] got a better result with organosolv pretreatment of rice straw for efficient butanol production without necessity of detoxification step and got 10.5 g l-1 of ABE concentration.

9.2.2.3 Barley Straw Barley straw follows a pretreatment process similar to wheat straw and subsequently undergoes fermentation to produce liquid fuels [45]. In a study, the solvent from pretreated barley straw using dilute sulphuric acid was fermented using Clostridium bejierinckii P260 that resulted in ABE yield about 0.33 with production of 7.09 g l-1 and productivity of 0.10 g l-1 h-1 which was less than using glucose. The low titre was due to toxicity of barley straw hydrolysate to the culture which has been reduced through pretreatment with lime. The treated barley straw achieved high ABE concentration, ABE yield and productivity of 26.62 g l-1, 0.43 g g-1and 0.39 g l-1 h-1, respectively compared to conventional and untreated barley straw [45]. Similarly, concentrated sugar solutions obtained from barley straw was fermented using C. beijerinckii P260 by employing simultaneous product recovery which resulted in 47.20 g l−1 of ABE production with a productivity and yield of 0.60 g l−1 h−1 and 0.42, respectively [53]. The pretreatment process of these straw results in production of additional chemical byproducts such as salts, furfural, acetic acids, ferulic acids, glucuronic acids, phenolic compounds and r-coumaric acids that inhibit the growth of cells and fermentation [54]. The several approaches were employed for reducing these inhibitory effects of hydrolysates such as dilution of hydrolysate before undergoing fermentation, removal of inhibitors using overliming, adsorbent resin or molecular sieve and genetically developing inhibitor tolerant strains [55].

9.2.3 Bagasse Bagasse is a fibrous agricultural residue with various applications such as enzyme production and biofuel production due to its large carbohydrate content which can be obtained from sugarcane [36, 38, 56], sorghum [37] and cassava [57]. Sugarcane bagasse is one of the largest lignocellulosic agricultural by-product and fibrous residue of cane stalks left over by the sugar industries [58]. The sugarcane bagasse consists of 42.8% cellulose, 25.9% hemicelluloses and 22.1% lignin, which make it more suitable for biofuel production [36, 38, 56]. Cassava bagasse is a fibrous residue comprising about 50% of starch on its dry weight [57] and has several advantages over wheat and rice straw

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since it possesses low ash content [59]. But it is unattractive as an animal feed due to its poor protein content. In comparison to sugarcane bagasse, it does not require pretreatment and supports biochemical conversion using microbes [58]. Furthermore, Sorghum bagasse is another solid residue obtained after sugar extraction from sweet sorghum stalks with polymer composition of cellulose 40.4%, hemicelluloses 35.5% and lignin 3.9% [37]. The biochemical conversion of bagasse can be economically beneficial in development of biofuel sector [58, 59]. The biofuel production from agricultural residues follows clean technology and has negligible impact on the environment compared to conventional technology.

9.3 ABE Fermentation 9.3.1 Butanolgenic Microorganisms ABE producing bacterial strains are available in large number, dominated by Clostridia which are rod-shaped, spore-forming, gram-positive and obligate anaerobic bacteria, belonging to the Firmicutes. These bacteria were isolated from various sources such as living plants materials, roots of nitrogen-fixing legumes and agricultural soil [60]. ABE- (acetone, butanol and ethanol) producing Clostridial strains are generally classified into four distinct groups based on their biochemical and genetic characteristics. Among the solventogenic Clostridia, C. acetobutylicum and C. beijerinckii are the most widely studied butanol producing strains. ABE fermentation by C. acetobutylicum and C. beijerinckii has two phases, acidogenesis and solventogenesis [11, 61]. Clostridium acetobutylicum is an anaerobic, saccharolytic and proteolytic bacterium that has been isolated from a number of environments such as soils from decomposed waste, sugarcane field, pulse and tuber crops. The bacterium produces endospores which allows for long-term survival in the environment even in the presence of oxygen. It exists in the biologically inactive spore stage in soils except when vegetative growth is stimulated by anaerobiosis and other favorable growth conditions [62]. Individual vegetative cells of Clostridium acetobutylicum are straight rod-shaped ranging in size of 0.5-1.5 × 1.5-6 μm. They are Gram positive, typically strictly anaerobes (oxygen free), hetero fermentative and spore-forming, motile by peritrichous flagella. The optimum growth temperature is 36°C, and biotin and 4-aminobenzoate are usually required as growth factors [63]. During sporulation, cells swell markedly and store granulose, a polysaccharide-based material that serves as a carbon and energy source during

Potential of Biobutanol Production 293 solventogenesis and allows long-term survival in the environment even in the presence of oxygen and spores are oval and subterminal. Besides C. acetobutylicum, three other species, C. beijerinckii, C. saccharoperbutylacetonicum and C. saccharobutylicum were also identified as butanol producers and they gave high butanol production and yields [11]. Some other Clostridial strains including C. carboxidivorans, C. butylicum, C. aurantibutyricum and C. pasteurianum were also investigated [64, 65]. Almost all strains can use a variety of hexose and pentose sugars as substrates. Most strains can use disaccharides such as sucrose and cellobiose and even polysaccharides such as starch. C. beijerinckii produces solvents in approximately the same ratio as C. acetobutylicum, but isopropanol is produced in place of acetone, while C. aurantibutyricum produces both acetone and isopropanol in addition to butanol [2]. C. tetanomorphum is a newly isolated species which produces almost equimolar amounts of butanol and ethanol but no other solvents [2]. C. thermocellum is an anaerobic, rod-shaped, Gram positive thermophile that is capable of producing butanol directly from cellulose. Despite its relatively recent rise to popularity in the literature, it was first isolated in 1926 in an attempt to identify novel organisms capable of degrading cellulose [66]. C. thermocellum could grow at temperatures between 50 and 68°C, and demonstrated this growth on cellulose, cellobiose, xylose, and hemicelluloses. It also produces the major fermentation products, consisting primarily of carbon dioxide and hydrogen gases, formic, acetic, lactic, and succinic acids, and ethanol [66]. Clostridium saccharoperbutylacetonicum is one of the most important acetone-butanol-ethanol (ABE) generating industrial microorganisms and one of the few bacteria containing choline in its cell wall [67]. It consists of straight, short and long rods with rounded ends, measuring 0-6  μm. Rods occur singly, or occasionally in pairs, and are motile by means of peritrichous flagella. They are initially phase-dark and Grampositive, becoming Gram-negative in older cultures. Towards the end of exponential growth, the rod-shaped cells usually begin to accumulate granulose. The species is mesophilic, optimum growth for solvent production occurring between 25 and 35°C; the optimum pH range is between pH 5 and 6 [61]. Clostridium cellulolyticum strain is a gram-positive, rod-shaped, anaerobic, mesophillic cellulolytic bacterium. C. acetobutylicum is able to convert a variety of sugars, polysaccharides and oligosaccharides to organic acids and solvents. Clostridium acetobutylicum is a solventogenic bacteria which utilizes sugar substrates to initially anaerobically produce butyric and acetic acids, CO2 and H2 gases during the exponential growth. When an acid

294

Liquid Biofuel Production

concentration (>1.5 g l-1 butyric acid) and pH (~4) threshold is reached, in the stationary phase, clostridia switches to Acetone, Butanol, Ethanol (ABE) solvent production [68]. The bacterium was first isolated from compost containing decayed grass. Studies have confirmed that this microbe is not a part of any ruminant system. The ability of Clostridium cellulolyticum to degrade cellulose is an active area of research. Two products of interest from the cellulose degradation by this organism are ethanol and hydrogen, which can be used as an alternative source of energy. C. acetobutylicum has a long history of safe use in the industrial production of acetone and butanol in fermentation systems using maize mash, molasses, or other feedstocks. Jones and Woods (1986) have thoroughly documented its history of use for solvent production. Mesophilic C. cellulolyticum synthesizes cellulosome, which is an extracellular multi-enzymatic complex, and degrades cellulose to glucose and soluble cellodextrins (mainly cellobiose) with the use of this cellulosome, where the β-1,4-glycosidic bonds of the cellulose polymer are hydrolyzed by general acid catalysis. C. cellulolyticum produces cell-free cellulosomes, and it has been proposed that the degree of cell-association, cellulose adhesion, and the composition of 31 cellulosomes are regulated by the substrate characteristics; for instance, in contrast to cellulose-grown cells, about 90% of the xylanase activity is not cellassociated when C. cellulolyticum is cultivated on xylan, and the composition of cellulosomes obtained from xylan and cellulose-grown cells are different [69]. C. kluyvera is an anaerobic strain that can produce high cellulase-free and thermo-alkali-stable xylanase. The xylanase could hydrolyze untreated lignocellulosics to reducing sugars. A coculture of Kluyvera sp. strain OM3 and Clostridium sp. strain BOH3 could directly convert xylan to 1.2 g l-1 butanol. This xylanase showed maximum activities at 70°C and pH 8.0 [70]. C. tyrobutyricum is an anaerobic bacterium which produces lactic acid, butyric acid and acetic acid as its main fermentation products from sugars, while also producing hydrogen and carbon dioxide as gaseous by-products. That efficiently produces butyric acid and is considered a promising host for anaerobic production of bulk chemicals. Due to limited knowledge on the genetic and metabolic characteristics of this strain, however, little progress has been made in metabolic engineering of this strain. The results of genomic analyses suggested that C. tyrobutyricum produces butyrate from butyrylcoenzyme A (butyryl-CoA) through acetate reassimilation by CoA transferase, differently from Clostridium acetobutylicum, which uses the phosphotransbutyrylase-butyrate kinase pathway C. tyrobutyricum has become the organism of choice because it converts both five- and

Potential of Biobutanol Production 295 six-carbon sugars to butyric, acetic and lactic acids. Elevated pH (6.3) is favorable for the production of butyric acid [71]. One disadvantage of Clostridia is a degeneration phenomenon. They gradually lose the ability to produce solvent during the cultivation. In C. acetobutylicum, this degeneration phenomenon is due to the loss of megaplasmid pSOL1 [11]. Another problem is that butanol is highly toxic to Clostridia itself. 0.10.15M butanol can cause 50% inhibition of both cell growth and sugar uptake rate by negatively affecting the ATPase activity [2]. This solvent toxicity highly inhibits the butanol yields of clostridia. Several reports suggest that C. beijerinckii might have greater potential for the industrial production of solvents than C. acetobutylicum since the former has a wider substrate range and pH optimum for growth and solvent formation [72]. Upcoming research focused to boost the butanol production by altering the strain genetically. In another approach, the enzyme responsible for the production has been introduced into yeast or E. coli sp. for making it butanol producer. Genetically engineered yeast and E. coli produced minimal amount of butanol which failed to meet the affordable demand whereas clostridium had conquered this shortcoming. Solvent producing genes such as hbd, crt, adhE1, adhE2 and bdhB were isolated from ABE producing bacteria for application through genetical transformation to other organisms like Esherichia coli [73, 74], Pseudomonas putida [75], Bacillus subtilis [75], Lactobacillus brevis [76] and Saccharomyces cerevisiae [77]. Yeast, such as Saccharomyces cerevisiae holds significant advantage over fermentation which has been treated with excess supply of substrate in the form of acetyl CoA or NADH in order to decrease the ethanol formation which consequently increases the butanol production in minimal range [78].

9.3.2 Fermentation ABE fermentation strictly employs anaerobic microbes (obligate anaerobes) for the production of acetone, n-butanol, and ethanol from carbohydrates such as glucose. The ratio of solvents produced by ABE is 3:6:1, Acetone: Butanol: Ethanol, respectively [16]. It uses solventogenic bacteria of family clostridia such as Clostridium bejierinckii [79], C. acetobutylicum, C. sporogenes, etc. ABE fermentation is drastically affected by varied operating parameters such as incubation temperature, pH, microbial cell inoculums and culture medium [80]. It needs desired optimization for enhancing the process efficacy that can be acquired by response surface methodology (RSM) [4]. Most commonly used processes for butanol fermentation are SHF, SSF, SSCF and CBP. The schematic description, pros and cons of the fermentation process using agricultural residues has been depicted in Table 9.2.

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Liquid Biofuel Production

Table 9.2 The comparison among different fermentation processes of agricultural residues. Fermentation methods

Description

Pros

Cons

SHF

• Hydrolysis of cellulose to glucose by cellulase enzyme. • Glucose to butanol using microbes.

Good yield

• Need of separate enzyme hydrolysis and fermentation due to inhibition of cellulase by glucose produced. • High equipment cost

SSF

Low equipment • Cellulose cost hydrolysis and fermentation in single step

SSCF

• Merging of cellulase production and cellulose hydrolysis as single step followed by fermentation

Overcome the • Not much shortcomings efficiently of SHF and SSF practiced

CBP

• All the three stages completed in single step

• Low cost • Economically attractive

• Temperature variation in two processes makes this single step method impossible. • Hydrolysis takes more time.

• Unavailability of perfect CBP microbes

SHF-Separate hydrolysis and fermentation; SSF-Simulataneous saccharification and fermentation; SSCF: Simultaneous saccharification and fermentation; CBP-Consolidated Bioprocessing.

Potential of Biobutanol Production 297 Among the various lignocellulosic substrates used for butanol fermentation, agricultural residues seem to be the most interesting [16, 45, 82]. The process to obtain fermentable sugars from these lignocellulosic materials is very difficult and involves several steps [83]. It requires physical, chemical, or biological pretreatment or their combinational treatments, before being utilized as fermentation substrates. Additionally, these treatments result in a hydrolysate that contains a mixture of different kinds of sugars as well as associated compounds. However, it is noteworthy that the solventogenic clostridia can utilize a wide range of carbohydrates, in particular both pentose and hexose [84]. The graphical representation of ABE fermentation using agricultural residues has been depicted in Figure 9.1. Also Butanol production from various sugar and biomass sources under different fermentation conditions has been shown in Table 9.3. Cellulose Hexose CH2OH H C C HOH HO C

Lignin

H

Pentose O H H

CH2OH

C

C OH OH

H

O

C

H H C OH

C HO CH 2OH C H

Hemicellulose Agricultural residues

Pretreated with acid/alkali

Lignin

Saccharification ENZYMES USED: Cellulase β- Glucosidase Endoglucanase Xylanase

Anaerobic microbes (Clostridium sp.) SOLVENT FERMENATATION PHASE

ACID FERMENATATION PHASE Acetate

Butyrate

BIPHASIC FERMENTATION

Acetyl- Co

Ethanol

Acetoacetyl- Co A

Acetone

Butyryl- Co A

Butanol

Figure 9.1 ABE fermentation for butanol production from agricultural residues.

Rice straw 0.49

Rice straw 0.51

SHF

SHF-SSF

β-glucosidase from CL3 strain(0.3 U/ ml) & Endo-glucanase from TCW1 strain (0.56 U/ml)

Rice straw 0.05

SSF Acid cellulase used at 30 FPUs.g-1 (dry substarte) concentration

Clostridium sporogenes BE01

Clostridial sp. TCW1

Sugarcane 0.19 bagasse

SSF

2.92

2.93

3.43

6.4

References [81]

Cocktail of cellulases used with two fermentaion mode.

(Continued)

[18]

Detoxification [32] of hydrolysate

Comparison of two processes SSF & SHF.

Butanol Butanol conc. Special yield (g/g) (g/l) character

Cellulase

Substrate

Clostridium bejierinckii NCIMB 8052

Butanologenic microorganisms Saccharification Fermentation

Table 9.3 Butanol production from various sugar and biomass sources under different fermentation condition. (Continued)

298 Liquid Biofuel Production

Clostridial sp. TCW1

β-glucosidase from CL3 strain(0.3 U/ ml) & Endo-glucanase from TCW1 strain (0.56 U/ml)

Bagasse

Bagasse

SHF-SSF

Substrate

SHF

Butanologenic microorganisms Saccharification Fermentation

0.52

0.37

2.29

1.95

References

(Continued)

[18] High capacity of sugar production than rice straw & Cocktail of cellulases with two fermentation mode.

Butanol Butanol conc. Special yield (g/g) (g/l) character

Table 9.3 Butanol production from various sugar and biomass sources under different fermentation condition. (Continued)

Potential of Biobutanol Production 299

Clostridium bejierinckii P260

0.41

0.44

Wheat straw

SSF-Batch

SSF-Fed-Batch Wheat straw

0.6 ml of each hydrolytic enzymes Cellulose, β-glucosidase and xylanase added

0.6 ml of each hydrolytic enzymes Cellulose, β-glucosidase and xylanase added

0.43

Barley straw

102.96 for 286h

21.42

26.64

[45]

References

(Continued)

Regular feed [24] to reactor will increase ABE yield by 16%.

[21] SSF with agitation by gas stripping was preferred.

Lime treated Barley straw yields more.

Butanol Butanol conc. Special yield (g/g) (g/l) character

SHF

Substrate

Three enzymes of 6 mlL-1 each added: Cellulose, β-glucosidase and xylanase

Butanologenic microorganisms Saccharification Fermentation

Table 9.3 Butanol production from various sugar and biomass sources under different fermentation condition. (Continued)

300 Liquid Biofuel Production

SHF

SHF

Cellulase: 5 IU

Cellulase: 5 IU

0.35

3.0

3.5

Hydrolysate treaments were performed.

Hydrolysate treaments were performed.

Butanol Butanol conc. Special yield (g/g) (g/l) character

Rice straw 0.32

Bagasse

Substrate

[50]

[50]

References

FPUs- filter paper units; U- unit of enzymatic activity; SHF- separate hydrolysis and fermentation; SSF simultaneous saccharification and fermentation.

C. saccharoper butylacetonicum

Butanologenic microorganisms Saccharification Fermentation

Table 9.3 Butanol production from various sugar and biomass sources under different fermentation condition. (Continued)

Potential of Biobutanol Production 301

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The performance of butanol fermentation process using clostridial sp. is severely limited by various factors, such as high cost of fermentation substrate, substrate inhibition, low butanol concentration due to low solvent tolerance, sluggish growth and low cell density achievable during solventogenic fermentation. These limitations result in low butanol productivity with low yield due to the production of additional by-products, high downstream processing cost for butanol recovery due to the formation of mixed solvents and low butanol concentration [82]. During hydrolysis process, some inhibitors that could inhibit fermentation process may be generated due to the complex components of lignocelluloses (celluloses, hemicelluloses and lignin) [85]. There are three main groups: furan derivatives (Furfural and 5-hydroxymethyl furfural (HMF)), weak acids (glucuronic acid, formic acid, acetic acid and levulinic acid) and phenolics (p-coumaric acid, ferulic acid, vanillin, hydroquinone, and syringaldehyde). Furan derivatives come mainly from degradation of monosaccharide molecules, while phenolics are generated by lignin degradation or monosaccharide degradation during acid hydrolysis. HMF’s further breakdown could form some weak acids like formic acid and levulinic acid [85–87]. Their toxicities vary with different mechanisms. Furfural and HMF can even simulate the growth of C. beijerinckii BA101 [88]. Ferulic acid and p-coumaric acid are the most toxic compounds to most solventogenic clostridia. The growth of C. beijerinckii BA101 can be totally inhibited when their concentration reaches 1.0g/L. Ferulic and p-coumaric acid can damage the hydrophobic sites of bacterial cells and increase membrane permeability, which causes leakage of cellular contents. Syringaldehyde do not inhibit the growth of cells but decrease the production of ABE dramatically by affecting in the glycolytic pathway and alcohol dehydrogenase enzymes. Evaporation, usually under vacuum, can reduce the volatiles in the hydrolysate, such as furfural, formic acid, acetic acid and vanillin [89]. But other non-volatile inhibitors can also be concentrated during evaporation. Therefore, the detoxification effects of evaporation largely depend on the types of inhibitors in the hydrolysate. The concentration of green liquor wood extracts resulted in increased inhibition to E. coli KO11 due to the concentration of sodium [90]. The pH adjustment is often called overliming because Ca (OH)2 is used in the process which is the most widely studied in the detoxification of hydrolysate [89]. The pH is usually raised to ~10 by some base, usually Ca (OH)2, and then reduced to the fermentation level. Although the mechanism of overliming is still unclear, it is already an effective way to detoxify hydrolysate, especially spruce hydrolysate [91]. pH equal to 10 tends to be a optimized condition for Ca(OH)2. Lower pH gives little improvement in detoxification and higher pH increases the degradation of lignin and induces higher concentration of phenolic inhibitors. For

Potential of Biobutanol Production 303 corn stover hydrolysate, the detoxification of NH3·H2O adjustment was more effective than Ca(OH)2 [92].

9.3.3 ABE Pathway In batch fermentation, solvent producing clostridia during its initial growth phase (acidogenisis) produces carbon dioxide, hydrogen, acetate and butyrate which results to decrease in pH of the culture medium. While the culture attains the stationary growth phase, cells undergo shift from acid to solvent production (solventogenic phase). At this phase, there is an increase in pH of the culture medium due to continued consumption of sugar and resulting in reformation of acid. In fermentation, conversion of carbohydrate to acids, solvents and other gases occurs using carbohydrate hexose and pentose. Hexose sugars are metabolized to pyruvate through Embden-Meyerhof (glycolysis) pathway whereas pentose sugars initially undergo pentose-phosphate pathway yielding pentose-5-phosphate and through transketolase-transaldolase sequence which results in fructose-6-phosphate which enters glycolysis. This pentose sugar conversion was metabolized by solvent producing clostridia [93]. The pyruvate obtained through glycolysis was cleaved to acetyl CoA and carbon dioxide in presence of coenzyme A (CoA). This acetyl CoA considered as precursor for both acid and solvent production in the fermentation pathway. It seems that acetyl-CoA is mostly used to form butyryl-CoA, based on the fact that the conversion of acetyl-CoA to butyryl-CoA exhibits enhanced thermodynamic stability. A high concentration of acetyl-CoA is needed to make this reaction go smoothly, and the quantity of acetyl-CoA plays an important role in determining the ratio of C3 and C4 products to C2 products. This shift induction is controlled either by the decrease in pH (2 g L-1) [93, 94]. In addition to acetyl-CoA, CO2 and reduced-ferredoxin products are also formed during the reaction. In many Clostridium species the primary role of NADH ferredoxin-oxidoreductase, which requires acetyl CoA as an activator, is NAD+ regeneration by reduced ferredoxin-oxidoreductase production to produce NADPH by biosynthesis. Under appropriate NADPH conditions, the reduced ferredoxin is able to transfer electrons to iron and permitting the use of protons as a final electron acceptor, resulting in the production of molecular hydrogen. Ferrodoxin is oxidised during this step and hydrogen gas is released from the cell [95]. Acetyl-CoA is a branch point between acid- versus solvent-production in C. acetobutylicum and it can be different, depending on the clostridial

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species. It can be converted into a mixture of ethanol, acetate and or butyrate. The ratio in which these products are formed depends on the amount of H2 involved. Extra H2 evolution relieves the fermentation redox and allows the direct conversion of part of the acetyl-CoA via acetyl phosphate into acetate [95, 96].

9.3.3.1 Acid Producing Phase Most of the Clostridium based ABE fermentation pathway involves mainly two phases, i.e., acid producing phase and solvent producing phase as shown in Figure 9.1. There are three intermediates such as acetyl-CoA, acetoacetyl-CoA, and butyryl-CoA responsible for the production of acids and solvents. During this acidogenesis, acetate and butyrate are produced from acetyl CoA and butyryl-CoA respectively in presence of enzymes such as phosphate acetyltransferase and acetate kinase mediating acetate formation; phosphate butyrltransferase and butyrate kinase for butyrate production [64].

9.3.3.2 Solvent Producing Phase Acid producing phase switches to solvent production where acetyl-CoA, acetoacetyl-CoA and butyryl-CoA yields ethanol, acetone and butanol, respectively. Butanol production occurs generally in the presence of reducing enzymes such as butyaldehyde dehydrogenase and butanol dehydrogenase [97]. The products of the ABE pathway were acetate, butyrate, acetone, butanol and ethanol which make toxic essence towards the bacterial strain. Among which butanol and butyrate were at higher concentration causes product inhibition that can be prevented by processing fermentation with continuous culture. In Clostridia fermentation, the sporulation occurs concomitantly with the solventogenesis. Sporulation makes the bacterial cells enter a dormant state where they lose the ability to produce solvents. It is likely that there is a relationship between sporulation and solventogenesis, given that many early molecular events connected with sporulation appear in the initiation of solventogenesis [98]. If this relationship is revealed, it may be possible to produce more solvents, including butanol, by preventing the clostridia from forming spores. The induction mechanisms for solventogenesis and sporulation in C. acetobutylicum have several features in common, so attempts have been made to elucidate the relationship between sporulation and solventogenesis.

Potential of Biobutanol Production 305

Growth phase

Acidogenic phase

Solventogenic phase & sporulation

Figure 9.2 Different stages of bacteria during ABE fermentation.

Solventogenesis may stimulate the bacteria to form spores, since solvent formation creates adverse circumstances for cell growth. Sporulation makes the bacterial cell enter a dormant state, which helps the cell to live through the adverse circumstances [96]. Various sigma factor genes from C. acetobutylicum were cloned and characterized in order to elucidate the molecular mechanism leading to the initiation of sporulation and solventogenesis. Sigma factors are required for RNA polymerases to initiate transcription by recognizing specific promoter sequences. Therefore, they play a central role in controlling mRNA transcription. SpoOA, transcription factor can control sporulation, the development of competence for DNA uptake, and many other stationary phase associated processes [96]. Different stages of Costridium bacteria including acidigenesis, solventogenesis and sporulation have been shown in Figure 9.2.

9.4 Challenges There are certain major challenges in butanol production, i.e., anaerobic nature, slow growth rate of microorganisms, availability of compatible feedstocks, rapid shifts of pH, sensitivity to acetic acid and butyric acid, low butanol titer, solvent tolerance, and product inhibition [25–27]. The importance of some of the changes has been described below.

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9.4.1 Strict Anaerobic Nature The most challenging research involves studies of strictly anaerobic microorganisms since it needs anaerobic (oxygen free) environment for its activities, enrichment, and for obtaining pure culture. It is well known that anaerobes exist on earth with complex communities that play a vital role in the carbon, nitrogen and sulphur cycles. Oxygen has a critical effect on the Clostridia growth but the growth inhibition mechanism remains unknown. Butanol producer species carried out a fermentative energy metabolism, where oxygen is not required for ATP production. Moreover, they exhibit a more or less pronounced sensitivity towards oxygen. The presence of oxygen at concentration less than 20 ppm for 30 second in the culture will upset metabolism activity of the bacteria and resulted in low solvent productivity. Peroxide repressor (PerR) gene, responsible to regulate genes encoding detoxification component, was identified as the reason of low aerotolerance of C. acetobutylicum [95, 99]. It was shown that deletion of a peroxide repressor (PerR)-homologous protein in C. acetobutylicum resulted in prolonged aerotolerance, limited growth under aerobic conditions, higher resistance to H2O2, and rapid consumption of oxygen. Several genes were identified as putative members of the clostridial PerR regulon, including the heat shock protein Hsp21, a multifunctional reverse rubrerythrin that is proposed to play a crucial role in oxidative stress defense. This work generated new opportunities to further enhance clostridial aerotolerance [100]. In clostridial metabolic pathway, the pyruvate resulting from glycolysis is cleaved by pyruvate-ferredoxin oxidoreductase (PFOR) in presence of coenzyme-A to yield carbon dioxide and acetyl-CoA with concomitant conversion of oxidized ferredoxin to its reduced form. The pyruvate-ferredoxin oxidoreductase of clostridia is a very unstable enzyme and very sensitive to the presence of oxygen. Upon the exposure to pure oxygen, the enzyme was 50% inactivated within an hour, while inactivation was not observed under a nitrogen atmosphere after 24 h [97]. In bacteria, the formation or consumption of hydrogen is catalyzed by hydrogenases. They are classified according to the transition metal cofactors (i.e., NiFe, Fe, and FeFe hydrogenases) associated with the protein [101]. The clostridial [FeFe]-hydrogenases lack other transition metals. The active site consists of four iron–sulfur clusters in addition to an H-cluster and it often catalyzes the reduction of protons to yield hydrogen at high turnover numbers. These kinds of hydrogenases are very sensitive to carbon monoxide and oxygen [97]. Various species of Clostridium have desired abilities such as Clostridium butyricum resumes its growth after oxygen consumption without causing

Potential of Biobutanol Production 307 any damage. Clostridium glycolicum is an acidogenic bacterium that can grow in presence of oxygen with oxygen consuming ability under 6 percent headspace oxygen in static culture [102]. Similarly, C. magnum could grow in non-reduced liquid culture in presence of oxygen under 1 percent headspace oxygen [103]. These reports suggest that clostridia should possess certain systems for metabolizing oxygen and tolerating active oxygen. However, the existence of these systems has not been well investigated. In addition, Bacillus has the ability to consume O2 and maintain strict anaerobic conditions for butanol production in a mixed culture system. A mixed culture of C. butylicum TISTR 1032 and B. subtilis WD 161 without anaerobic treatment was able to increase ABE production from cassava starch by 6.5 fold relative to a pure culture of TISTR 1032 strain [104]. It is interesting to note that in a pure culture of C. acetobutylicum ATCC 824, anaerobic treatment along with the addition of a reducing agent and N2 flushing resulted in a butanol concentration of only 12.3 g l-1, which was lower than the concentration of 14.9 g l-1 obtained by mixed culture using C. acetobutylicum ATCC 824 and B. subtilis DSM 4451 without anaerobic treatment [14].

9.4.2 Tolerance to Solvent ABE fermentation yields butanol with the involvement of two significant phases where the first phase was characterized by rapid growth of cells as well as formation of acetic and butyric acids in the medium which lowers the pH. These acids with its high concentration equilibrate the pH between internal cell environment and growing medium. The second phase commences after encountering low pH of the medium at below 5.0, where the accumulated acids enter the cell membrane in undissociated form and further converted to solvents [105]. This evidence proves that solvent production is not due to low external pH but result of internal pH drop. In specific, the maximum butanol concentration that could be attained from fermentation is about 20 g l-1 [106]. It represents a critical bottleneck associated with poor ability of clostridia sp. to tolerate high solvent concentration. Currently, higher butanol production suffers from longer fermentation time, low titre, low yield, low productivity and high recovery cost due its toxicity [107, 108]. This may also result in cessation of metabolic activities of anaerobes and imparts inhibitory effects to the process. Mechanism of product inhibition in anaerobes is not well known and it is reported that butanol tends to inhibit transport of sugar in Clostridium sp. The deleterious action of these solvents is due to its ability to accumulate in the cytoplasmic membrane, tampering with its structure and preventing the cell from performing essential functions such as dissipation of pH and electrical potential, normal flow

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of ions, proteins, lipids and endogenous metabolites, and inhibiting membrane protein functions. Ultimately these actions lead to cell lysis and death. Thus, this problem should be taken into consideration in order to obtain high-yield butanol via ABE fermentation [107, 109]. These problems should be taken into consideration to reduce butanol toxicity for improving its titre. Many solutions have been developed such as selection of solvent tolerant microbes and improvement of in-situ product recovery technology with simultaneous removal of product from fermentation broth using membrane and gas stripping techniques to prevent product inhibition [109]. These methods result in higher butanol production from fermentation process other than batch type where culture is not assisted with removal of product. Additionally, solvent tolerant strains have been developed using chemical mutagenesis, fermentation with continuous culture and serial enrichment methods [110]. It involves overexpression of genes such as molecular pumps, chaperones and genes expressing in sporulation, fatty acid synthesis and transcriptional regulators [111]. Genetically engineering the bacterial strains for making resistant to solvent is difficult and it has been enhanced through enrichment of mutants by serial transfers. Hybrid also evolved from protoplast fusion between E. coli and Lactobacillus brevis for enhancing the solvent tolerance from 1 to 2% (v/v) [110]. However, efforts towards improvement of the tolerance of the strains to butanol using genetic techniques have resulted in limited success to date [112, 113].

9.4.3 Sensitivity of Acids In ABE fermentation, the acids produced during initial phase were utilized for solvent production using Clostridial sp. However, the mechanism of shifting from acid to solvent forming phase in ABE fermentation was still not clear. The favorable acids produced during this process were acetic and butyric acids that have been ultimately used in solvent production [114]. Other acids such as lactate and formate also encountered by anaerobic microbes and regulation of all these acids in the culture media are of great industrial importance. Uncertain levels of butyric and acetic acid concentration are used for the induction of butanol formation [115]. Requirement of the exact concentration levels of acids for solvent production still remains controversial due to the complicated metabolic mechanisms. In specific, butyric acid production alters the lipid composition and membrane fluidity of bacteria [116]. These changes may be due to addition of butanol during the exponential phase or stationary phase with

Potential of Biobutanol Production 309 conversion of butyric acid to butanol takes place where the cells may be more sensitive to these acids. In a report, the acetic acid produced by the C. thermoaceticum cells diffuses across the cell membrane in its undissociated form and acidified the cytoplasm by passing proton across the membrane consequently altering the pH. As per the early reports, the effect of weak acids on butanol production was investigated in Clostridium sp. and showed that formic acid has fatal inhibitory reaction towards the solvent production in C. acetobutylicum whereas C. bejierinckii has strong tolerance against these acids. However, C. bejierinckii is more preferred when the lignocellulosic hydrolysate contains acetic and formic acids [88]. The effects associated with the acid production during the fermentation process can be controlled by pH maintenance and employing suitable methods for complete reutilization of acids to solvent production and value-added products.

9.4.4 Shifting of pH The pH of the medium plays a vital role in ABE fermentation during acidogenesis in which pH drops due to the rapid formation of acetic and butyric acids. On the other hand, solventogenesis is initiated when the pH reaches a critical point, beyond which acids are reassimilated and solvents were produced. The low pH is a prerequisite for the production of solvent [115]. The control over pH of the bacteria can be achieved through maintenance of internal pH in anaerobes which depends on the function of enzyme ATPase. The initial pH of the fermentation medium is an important factor in butanol production which significantly affects to both the process and the enzyme activities which decreases the butanol production. Jiang et al. [117] found optimum pH of 4.8 using C. acetobutylicum ATCC84 for enhanced butanol production. On the other hand, optimum pH between 6 and 6.2 was suitable for butanol production using C. acetobutylicum YM1. It was concluded that uncontrolled pH condition with initial pH of 6.0 ± 0.2 resulted in higher butanol concentration with high butanol titre (13.5 ± 1.42 g.l-1) [115]. The higher solvent production can be achieved with low and control pH. Therefore, the more research development must be encouraged to have efficient solvent production with improved technology.

9.5 Future Prospects and Conclusions ABE fermentation for biobutanol production using agricultural residues is a clean way of technology that has less impact on the environment compared to conventional technology. Although the technology has some

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challenges such as low productivity and yield due to product inhibition and heterofermentation, that can be overcome genetically through certain technologies such as genetic manipulation and mutagenesis. Product inhibition can also be controlled by using continuous culture other than batch which ultimately reduces cost of butanol recovery. Pretreatment of biomass/substrate and further saccharification using purchased enzymes generally makes the process costly which can be managed by producing own crude enzyme from different microbial sources and through birefinery approach. Improved detoxification, efficient microbial strains, optimization of process, increasing usage of renewable source would decrease the cost of butanol production. Future research may focus more on bacterial tolerance capacity through adaptation and genetic modification process to improve butanol titer. Ultimately, it is expected that a sustainable and cost-effective process for butanol production will be realized in the near future.

Acknowledgments The authors acknowledge the grant support from Science & Engineering Research Board, New Delhi, Govt. of India (File No. YSS/2015/000295). The authors also thank Sardar Swaran Singh National Institute of BioEnergy, Kapurthala, India as the host Institution for providing laboratory space to complete this work.

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50. Soni, B.K., Das, K., Ghose, T.K., Bioconversion of agro-wastes into acetone butanol. Biotechnol. Lett., 4, 19–22, 1982. 51. Moradi, F., Amiri, H., Soleimanian-Zad, S., Ehsani, M.R., Karimi, K., Improvement of acetone, butanol and ethanol production from rice straw by acid and alkaline pretreatments. Fuel, 112, 8–13, 2013. 52. Amiri, H., Karimi, K., Zilouei, H., Organosolv pretreatment of rice straw for efficient acetone, butanol, and ethanol production. Bioresour. Technol., 152, 450–456, 2014. 53. Qureshi, N., Cotta, M.A., Saha, B.C., Bioconversion of barley straw and corn stover to butanol (a biofuel) in integrated fermentation and simultaneous product recovery bioreactors. Food Bioprod. Process., 92, 298–308, 2014. 54. Ezeji, T.C., Qureshi, N., Blaschek, H.P., Butanol production from agricultural residues: Impact of degradation products on Clostridium beijerinckii growth and butanol fermentation. Biotechnol. Bioeng., 97, 1460–1469, 2007b. 55. Qureshi, N., Ebener, J., Ezeji, T.C., Dien, B., Cotta, M.A., Blaschek, H.P., Butanol production by Clostridium beijerinckii BA101. Part I: Use of acid and enzyme hydrolysed corn fiber. Bioresour. Technol., 99, 5915–5922, 2008d. 56. Silva, V.F., Arruda, P.V., Felipe, M.G., Gonçalves, A.R., Rocha, G.J., Fermentation of cellulosic hydrolysates obtained by enzymatic saccharification of sugarcane bagasse pretreated by hydrothermal processing. J. Ind. Microbiol. Biotechnol., 38, 809–817, 2011. 57. Carta, F.S., Soccol, C.R., Ramos, L.P., Fontana, J.D., Production of fumaric acid by fermentation of enzymatic hydrolysates derived from cassava bagasse. Bioresour. Technol., 68, 23–28, 1999. 58. Pandey, A., Soccol, C.R., Nigam, P., Soccol, V.T., Biotechnological potential of agro-industrial residues. I: Sugarcane bagasse. Bioresour. Technol., 74, 69–80, 2000a. 59. Pandey, A., Soccol, C.R., Nigam, P., Soccol, V.T., Vandenberghe, L.P.S., Mohan, R., Biotechnological potential of agro-industrial residues. II: Cassava bagasse. Bioresour. Technol., 74, 81–87, 2000b. 60. McCutchan, W.N. and Hickey, R.J., The butanol acetone fermentations. Ind. Ferment., 1, 347–388, 1954. 61. Keis, S., Shaheen, R., Jones, D.T., Emended descriptions of Clostridium acetobutylicum and Clostridium beijerinckii, and descriptions of Clostridium saccharoperbutylacetonicum sp. nov. and Clostridium saccharobutylicum sp. nov. Int. J. Syst. Bacteriol., 51, 2095–2103, 2001. 62. Calam, C.T., Isolation of C. acetobutylicum strains producing butanol and acetone. Biotechnol. Lett., 2, 111–116, 1980. 63. Dolly, M., Spitia, S., Silva, E., Schwarz, W.H., Isolation of mesophilic solvent producing strains from Colombian sources: Physiological characterization, solvent production and polysaccharide hydrolysis. J. Biotechnol., 79, 117–126, 2000. 64. Bruant, G., Levesque, M.J., Peter, C., Guiot, S.R., Masson, L., Genomic analysis of carbon monoxide utilization and butanol production by Clostridium carboxidivorans strain P7. PloS One, 5, 1–12, 2010.

Potential of Biobutanol Production 315 65. Ahn, J.H., Sang, B.I., Um, Y., Butanol production from thin stillage using Clostridium pasteurianum. Bioresour. Technol., 102, 4934–4937, 2011. 66. Akinosho, H., Yee, K., Close, D., Ragauskas, A., The emergence of Clostridium thermocellum as a high utility candidate for consolidated bioprocessing applications. Front. Chem., 2, 66, 2014. 67. Del Cerro, C., Felpeto-Santero, C., Rojas, A., Tortajada, M., Ramon, D., Garcia, J.L., Genome sequence of the butanol hyperproducer Clostridium saccharoperbutylacetonicum N1-4. Genome Announc., 1, 2, e00070–13, 2013. 68. Hartmanis, M.G. and Gatenbeck, S., Intermediary metabolism in Clostridium acetobutylicum: Levels of enzymes involved in the formation of acetate and butyrate. Appl. Environ. Microbiol., 47, 1277–1283, 1984. 69. Desvaux, M., Clostridium cellulolyticum: Model organism of mesophilic cellulolytic clostridia. FEMS Microbiol. Rev., 29, 741–764, 2005. 70. Bhatia, S.K., Kim, S.Y., Yoon, J.J., Yang, Y.H., Current status and strategies for second generation biofuel production using microbial systems. Energy Convers. Manage., 148, 1142–1156, 2017. 71. Lee, J., Jang, Y.S., Han, M.J., Lee, S.Y., Deciphering Clostridium tyrobutyricum metabolism based on the whole-genome sequence and proteome analysis. MBio, 7, 3, e00743–16, 2016. 72. Ezeji, T.C., Qureshi, N., Blaschek, H.P., Butanol fermentation research: Upstream and downstream manipulations. Chem. Rec., 4, 305–314, 2004. 73. Inui, M., Suda, M., Kimura, S., Yasuda, K., Suzuki, H., Toda, H., Yamamoto, S., Okino, S., Suzuki, N., Yukawa, H., Expression of Clostridium acetobutylicum butanol synthetic genes in Escherichia coli. Appl. Microbiol. Biotechnol., 77, 1305–1316, 2008. 74. Atsumi, S., Cann, A.F., Connor, M.R., Shen, C.R., Smith, K.M., Brynildsen, M.P., Chou, K.J.Y., Hanai, T., Liao, J.C., Metabolic engineering of Escherichia coli for 1-butanol production. Metabolic Eng., 10, 305–311, 2008. 75. Nielsen, D.R., Leonard, E., Yoon, S.H., Tseng, H.C., Yuan, C., Prather, K.L.J., Engineering alternative butanol production platforms in heterologous bacteria. Metabolic Eng., 11, 262–273, 2009. 76. Berezina, O.V., Zakharova, N.V., Brandt, A., Yarotsky, S.V., Schwarz, W.H., Zverlov, V.V., Reconstructing the Clostridial n-butanol metabolic pathway in Lactobacillus brevis. Appl. Microbiol. Biotechnol., 87, 635–646, 2010. 77. Steen, E.J., Chan, R., Prasad, N., Myers, S., Petzold, C.J., Redding, A., Ouellet, M., Keasling, J.D., Metabolic engineering of Saccharomyces cerevisiae for the production of n-butanol. Microb. Cell Fact., 7, 36, 2008. 78. Schadeweg, V. and Boles, E., n-Butanol production in Saccharomyces cerevisiae is limited by the availability of coenzyme A and cytosolic acetyl –CoA. Biotechnol. Biofuels, 9, 44, 2016. 79. Zhang, W.L., Liu, Z.Y., Liu, Z., Li, F.L., Butanol production from corncob residue using Clostridium beijerinckii NCIMB 8052. Lett. Appl. Microbiol., 55, 240–246, 2012.

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80. Kajal, G.S., Kaushal, L.L., Rekha, R.G., Ranade, D.R., Optimization for butanol production using Plackett-Burman Design coupled with Central Composite Design by Clostridium beijerenckii strain CHTa isolated from distillery waste manure. Biochem. Tech., 7, 1063–1068, 2016. 81. Su, H., Liu, G., He, M., Tan, F., A biorefining process: Sequential, combinational lignocellulose pretreatment procedure for improving biobutanol production from sugarcane bagasse. Bioprocess Technol., 187, 149–160, 2015. 82. Jang, Y.S., Malaviya, A., Cho, C., Lee, J., Lee, S.Y., Butanol production from renewable biomass by clostridia. Bioresour. Technol., 123, 653–663, 2012. 83. Blanch, H.W., Simmons, B.A., Klein-Marcuschamer, D., Biomass deconstruction to sugars. Biotechnol. J., 6, 1086–1102, 2011. 84. Tracy, B.P., Jones, S.W., Fast, A.G., Indurthi, D.C., Papoutsakis, E.T., Clostridia: The importance of their exceptional substrate and metabolite diversity for biofuel and biorefinery applications. Curr. Opin. Biotechnol., 23, 364–381, 2012. 85. Jonsson, L.J., Alriksson, B., Nilvebrant, N.O., Bioconversion of lignocellulose: Inhibitors and detoxification. Biotechnol. Biofuels, 6, 16, 2013. 86. Jonsson, L.J. and Martin, C., Pretreatment of lignocellulose: Formation of inhibitory by-products and strategies for minimizing their effects. Bioresour. Technol., 199, 103–112, 2016. 87. Kumar, A.K. and Sharma, S., Recent updates on different methods of pretreatment of lignocellulosic feedstocks: A review. Bioresour. Bioprocess, 4, 7, 2017. 88. Cho, D.H., Shin, S.J., Kim, Y.H., Effects of acetic and formic acid on ABE production by Clostridium acetobutylicum and Clostridium beijerinckii. Biotechnol. Bioprocess Eng., 17, 270–275, 2012. 89. Coz, A., Llano, T., Cifrian, E., Viguri, J., Maican, E., Sixta, H., Physicochemical alternatives in lignocellulosic materials in relation to the kind of component for fermenting purposes. Materials (Basel), 9, 574, 2016. 90. Walton, S., Heiningen, A.V., Walsum, G.P.V., Inhibition effects on fermentation of hardwood extracted hemicelluloses by acetic acid and sodium. Bioresour. Technol., 101, 1935–1940, 2009. 91. Persson, P., Larsson, S., Jonsson, N.J., Nilvebrant, N.O., Sivik, B., Munteanu, F., Thorneby, L., Gorton, L., Supercritical fluid extraction of a lignocellulosic hydrolysate of spruce for detoxification and to facilitate analysis of inhibitors. Biotechnol. Bioeng., 79, 694–700, 2002. 92. Jennings, E.W. and Schell, D.J., Conditioning of dilute-acid pretreated corn stover hydrolysate liquors by treatment with lime or ammonium hydroxide to improve conversion of sugars to ethanol. Bioresour. Technol., 102, 1240– 1245, 2011. 93. Aristilde, L., Metabolite labelling reveals hierarchies in Clostridium acetobutylicum that selectively channel carbons from sugar mixtures towards biofuel precursors. Microb. Biotechnol., 10, 162–174, 2017.

Potential of Biobutanol Production 317 94. Kudahettige-Nilsson, R.L., Helmerius, J., Nilsson, R.T., Sjoblom, M., Hodge, D.B., Rova, U., Biobutanol production by Clostridium acetobutylicum using xylose recovered from birch Kraft black liquor. Bioresour. Technol., 176, 71–79, 2015. 95. Papoutsakis, E.T., Engineering solventogenic bacteria. Curr. Opin. Biotechnol., 19, 420–429, 2008. 96. Zheng, Y.N., Li, L., Xian, M.M.Y., Yang, J., Xu, X., He, D., Problems with the microbial production of butanol. J. Ind. Microbiol. Biotechnol., 36, 1127– 1138, 2009. 97. Gheshlaghi, R., Scharer, J.M., Moo-Young, M., Chou, C.P., Metabolic pathways of clostridia for producing butanol. Biotechnol. Adv., 27, 764–781, 2009. 98. Al-Hinai, M.A., Jones, S.W., Papoutsakis, E.T., The Clostridium sporulation programs: Diversity and preservation of endospore differentiation. Microbiol. Mol. Biol. Rev., 79, 19–37, 2015. 99. Hillmann, F., Doring, C., Riebe, O., Ehrenreich, A., Fischer, R.J., Bahl, H., The role of perR in O2-affected gene expression of Clostridium acetobutylicum. J. Bacteriol., 191, 6082–6093, 2009. 100. Girbal, L., Mortier-Barriere, I., Raynaud, F., Rouanet, C., Croux, C., Soucaille, P., Development of a sensitive gene expression reporter system and an inducible promoter-repressor system for Clostridium acetobutylicum. Appl. Environ. Microbiol., 69, 4985–4988, 2003. 101. Vignais, P.M., Billoud, B., Meyer, J., Classification and phylogeny of hydrogenases. FEMS Microbiol. Rev., 25, 455–501, 2001. 102. Kusel, K., Karnholz, A., Trinkwalter, T., Devereux, R., Acker, G., Drake, H.L., Physiological ecology of Clostridium glycolicum RD-1, an aerotolerant acetogen isolated from sea grass roots. Appl. Environ. Microbiol., 67, 4734–4741, 2001. 103. Karnholz, A., Kuse, K., Gossner, A., Schramm, A., Drake, H.L., Tolerance and metabolic response of acetogenic bacteria toward oxygen. Appl. Environ. Microbiol., 68, 1005–1009, 2002. 104. Tran, H.T.M., Cheirsilp, B., Hodgson, B., Umsakul, K., Potential use of Bacillus subtilis in a co-culture with Clostridium butylicum for acetonebutanolethanol production from cassava starch. Biochem. Eng. J., 48, 260–267, 2010. 105. Ibrahim, M.F., Linggang, S., Jenol, M.A., Yee, P.L., Abd-Aziz, S., Effect of buffering system on Acetone-Butanol-Ethanol fermentation by Clostridium acetobutylicum ATCC 824 using pretreated oil palm empty fruit bunch. Bioresources, 10, 3890–3907, 2015. 106. Patakova, P., Kolek, J., Sedlar, K., Koscova, P., Branska, B., Kupkova, K., Paulova, L., Provaznik, I., Comparative analysis of high butanol tolerance and production in clostridia. Biotechnol. Adv., 36, 721–738, 2017. 107. Knoshaug, E.P. and Zhang, M., Butanol tolerance in a selection of microorganisms. Appl. Biochem. Biotechnol., 153, 13–20, 2009.

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108. Pfromm, P.H., Boadu, V.A., Nelson, R., Vadlani, P., Madl, R., Bio-butanol vs. bio-ethanol: A technical and economic assessment for corn and switchgrass fermented by yeast or Clostridium acetobutylicum. Biomass Bioenergy, 34, 515–524, 2010. 109. Mariano, A.P., Qureshi, N., Ezeji, T.C., Bioproduction of butanol in bioreactors: New insights from simultaneous in situ butanol recovery to eliminate product toxicity. Biotechnol. Bioeng., 108, 1757–1765, 2011. 110. Winkler, J., Rehmann, M., Kao, K.C., Novel Escherichia coli hybrids with enhanced butanol tolerance. Biotechnol. Lett., 32, 915–920, 2010. 111. Tomas, C., Beamish, J., Papoutsakis, E.T., Transcription analysis of butanol stress and tolerance in Clostridium acetobutylicum. Bacteriology, 186, 2006– 2018, 2004. 112. Lopez-Contreras, A.M., Kuit, W., Siemerink, M.A.J., Kengen, S.W.M., Springer, J., Claassen, P.A.M., Production of longer-chain alcohols from lignocellulosic biomass: Butanol, isopropanol and 2,3-butanediol. In: K. Waldron (eds) Bioalcohol. Woodhead Publishing, Cambridge (UK), pp. 415–460, 2010. 113. Lutke-Eversloh, T., Bahl, H., Metabolic engineering of Clostridium acetobutylicum: Recent advances to improve butanol production. Curr. Opin. Biotechnol., 22, 1–14, 2011. 114. Tashiro, Y., Takeda, K., Kobayashi, G., Sonomoto, K., Ishizaki, A., Yoshino, S., High butanol production by Clostridium saccharoperbutylacetonicum N1-4 in fed-batch culture with pH-stat continuous butyric acid and glucose feeding method. J. Biosci. Bioeng., 98, 263–268, 2004. 115. Al-Shorgani, N.K., Kalil, M.S., Yusoff, W.M.W., Hamid, A.A., Impact of pH and butyric acid on butanol production during batch fermentation using a new local isolate of Clostridium acetobutylicum YM1. Saudi J. Biol. Sci., 25, 339–348, 2018. 116. Kolek, J., Patakova, P., Melzoch, K., Sigler, K., Rezanka, T., Changes in membrane plasmalogens of Clostridium pasteurianum during butanol fermentation as determined by lipidomic analysis. PloS One, 10, e0122058, 2015. 117. Jiang, W., Wen, Z., Mianbin, W., Hong, L., Yang, J., Jianping, L., Yijun, L., Yang, L., Peilin, C., The Effect of pH control on Acetone–Butanol–Ethanol fermentation by Clostridium acetobutylicum ATCC 824 with xylose and D-glucose and D-xylose mixture. Chin. J. Chem. Eng., 22, 937–942, 2014.

10 State of Art Strategies for Biodiesel Production: Bioengineering Approaches Irem Deniz1*, Bahar Aslanbay2 and Esra Imamoglu2 1

Bioengineering Department, Engineering Faculty, Manisa Celal Bayar University, Muradiye/Manisa, Turkey 2 Bioengineering Department, Engineering Faculty, Ege University, Bornova/Izmir, Turkey

Abstract Sustainable and renewable energy demand is a universal challenge since the depletion of petroleum fuels, the increase in population globally, the accumulation of greenhouse gases and the concerns of climate change. These challenges along with the nutritional problems have given rise to the exploitation of thirdgeneration biodiesel production from microalgae. In this chapter, the novel trends in biodiesel production in bioengineering perspective are overviewed. Microalgal strains that have been conventionally used and new approaches towards metabolic engineering of algae are presented together with their advantages and bottlenecks. Moreover, new photobioreactor (PBR) types and configurations will be discussed in terms of design approaches, hydrodynamic parameters and kinetics. The chapter ends with the concept of the future possibilities of microalgal biodiesel production to meet the recent progress made in the field. Keywords: Microalgal biorefineries, microalgae cultivation, lipid extraction, microalgal oil, metabolic engineering, photobioreactor, kinetics

10.1 Introduction In the spectrum of microalgae-based-biofuels, biodiesel has been the most studied biofuel in the literature with around 417,000 studies, followed by

*Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (319–350) © 2019 Scrivener Publishing LLC

319

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303,000 on microalgae-based hydrogen, 273,000 on ethanol, and 107,000 on biogas [1]. However, feasible and practical commercial biofuel based on photoautotrophic production has not yet been installed. This is a result of the fact that the produced biomass and oil concentrations are very low and not sufficient, and also, the harvesting step of microalgae requires high energy consumptions. Nevertheless, conventional biodiesel production is supplied by transesterification of oil from agroindustrial crops which constitutes major problems due to the restriction of arable land for food production, deforestation and the environmental issues corresponding to the fertilizer consumption and water requirements. According to the bottleneck limitations of these first- and second-generation biofuel productions, microalgal biodiesel presents a viable alternative source of energy to replace or supplement the fossil fuels. Therefore, the need for new strategies and innovative approaches for biodiesel production from microalgae are crucial as microalgae provide a sustainable and renewable method for biofuel production. In this chapter the studies on novel microalgal biodiesel production methods are briefly overviewed. Different bioreactor designs with their kinetic properties and bioengineered microalgal strains are presented in terms of a potential resource for future uses for biodiesel production.

10.2 Biodiesel and Microalgal Biorefineries In recent years, interest in alternative energy sources has become increasingly important due to the environmentally harmful effects of petroleumbased fuels. Biodiesel is one of the ideal alternative fuels in terms of its several features: renewability, sustainability, efficiency, clean-burning, safety and cost-effectiveness [2]. Biodiesel is the mono-alkyl esters of long chain fatty acids from different sources such as vegetable oils, animal fats, waste cooking oils or microalgal biomass. Its formation process is called transesterification where the triglycerides and alcohol undergo a chemical reaction in the presence of an acid, base or enzymatic catalyst. At the end of the reaction, two products leave from the process as esters which are called biodiesel and glycerin and are a valuable by-product for the chemical industry [3, 4]. There are many feedstocks for the production of biodiesel divided into three groups as first, second and third generations. The choice of appropriate feedstock for production is determined according to economic issues, climate, availability, agricultural and geographical conditions [5,  6]. The most common raw materials are edible oils which are classified as first

Strategies for Biodiesel Production 321 generation derived from feedstocks like rice, soybean, coconut, olive, palm, corn, etc. However, these sources have increased problems about the using of the feedstocks for biodiesel synthesis instead of food production, which cause competition with the edible oil market, and the rise of the price of food [7–8]. In order to avoid the food vs. fuel debate and reduce food costs, second-generation feedstocks are recommended. They are produced from non-edible oils, including agricultural wastes and seed oils of lignocellulosic materials. Although it has been shown that biodiesel from non-edible oils have benefits for food industry there are as well some concerns about its use in terms of feedstock availability and potential negative effects on carbon balances and biodiversity [9]. Consequently, novel feedstocks as third generation have become the focus of biodiesel production. These feedstocks include waste frying oil, microalgal biomass, animal fat, pyrolysis oil, etc. Especially, microalgae have attracted increasing attention among these sources [10].

10.2.1

Microalgae

Microalgae are prokaryotic or eukaryotic photosynthetic microorganisms that use sunlight through the process to capture CO2 either from water or from air and release O2. They do not need a specific environment to grow; they can inhabit different type of water such as freshwater, seawater or wastewater. Also, some type of microalgae can survive in harsh conditions like extreme temperature values, acidic or alkaline pH, light, CO2 level and metal concentration due to their simple multicellular structure and adaption mechanism [11, 12]. The four most important classes of microalgae in terms of abundance in nature are the diatoms (Bacillariophyceae), the green algae (Chlorophyceae), the golden algae (Chrysophyceae) and the cyanobacteria (Cyanophyceae) which are also referred to as microalgae. These microorganisms have been widely used for many applications including human and animal nutrition, cosmetics, pharmaceuticals, CO2 capture, bioenergy production and wastewater treatment [13, 14]. In terms of bioenergy production, biodiesel is one of the most promising and should be examined in detail.

10.2.2

Microalgae and Biodiesel

Microalgae have some notable superiorities for biodiesel production. One of these advantages is high photosynthetic activity providing an inherent capacity as carbon-fixation group and oxygen production. They can be cultivated easily in different types of systems as open

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ponds and closed photobioreactors under controlled conditions. Cultivations result in high biomass formation in a short time and with some small modifications this yield can be further increased. As an example, Sabia et al. [15] reported that the increase of CO2 concentration resulted in higher biomass amount and pigment content per cell for Thalassiosira pseudonana microalgae. Also, lipid content doubled when air was enriched with 1–5% CO2. In another study, auxin hormone significantly enhanced microalgal growth and fatty acid methy esters accumulation, and has a potential for application in developing efficient microalgal cultivation [16]. Total costs of all production steps including cultivation, harvesting and extraction are low compared to the other biodiesel feedstocks. Considering the whole cultivation outputs, biodiesel production can be combined with different processes to provide a biorefinery approach and to make the overall cultivation economically feasible. Most researchers are focusing on novel biorefinery concept, mainly wastewater treatment with microalgae to remove nutrients such as nitrogen and phosphorus. In order to examine the biorefinery approach, a study was conducted to evaluate the capability of Arthrospira platensis cultivated in dairy farm wastewater for biodiesel production and wastewater treatment. The biomass of Arthrospira platensis was found to be 4.98 g L−1 and produced 30.23 wt% lipids to dry biomass cultivated in wastewater. Additionally, COD level and nutrient concentration reduced >98% within 4–5 days of culture [17]. In another study, Prieto et al. [18] developed a process that firstly, includes production of astaxanthin from Haematococcus pluvialis. After removing astaxanthin from biomass, the residue progressed through lipid extraction, transesterification and biodiesel production. The rest of glycerol content was recovered following transesterification process to minimize waste amount. To obtain a total biorefinery approach, PHB, biofertilizer and biogas products were obtained as co-products during this operation. Some other biorefinery approach is co-production of value-added products like animal feed, ethanol, methane and different chemicals. After the oil extraction for biodiesel production, microalgal residues can be processed with different technologies with the aim of zero waste [19]. Furthermore, microalgae do not compete for resources with food production as first-generation feedstocks and energy demand can be coverable with large-scale microalgae productions. From the point of biodiesel efficiency, microalgae have a strong capacity to produce lipids that contain large proportions of fatty acid triglycerides. Most microalgae species have the ability of lipid accumulation exceeding 70-80% of their dry cell weight which are mostly neutral lipids with a lower

Strategies for Biodiesel Production 323 degree of unsaturation. This makes microalgal lipids a potential replacement for fossil fuels [20].

10.2.3

Selection of Microalgal Strain for Biodiesel Production

It is noteworthy to mention that selection of the suitable algae strain is the first and most important part of biodiesel production. It is estimated that there are around 70,000 species of microalgae but only a small number, approximately 30,000, are identified and analyzed. Among these species, selected strains should have some characteristics to obtain high biodiesel yield. The strain with higher biomass productivity and high lipid contents is desired for an effective production [21, 22]. Since the biodiesel efficiency is mostly releated to fatty acid amount, researchers have focused on lipid content of different microalgae strains to find ideal sources. Also, high lipid productivity is a key desirable point of species for biodiesel production. Table 10.1 shows both lipid contents and productivities of some common microalgae strains. The lipid contents of microalgae vary with species and strains as well as culture conditions between the value of 2 to 80% in most recent studies [23–48]. As shown in Table 10.1, some species – mostly green microalgae – (Botyrococcus braunii, Chlorella, Dunaliella salina, Graesiella, Isochrysis galbana, Phaeodactylum tricornutum, Nannochloropsis oculata, Scenedesmus sp.) have much higher lipid content up to 79.89% dw. When comparing lipid productivity (mgL-1day-1) of different strains, green microalgae also have higher value than other strains and it is advantageous to select these species for biodiesel production. Chlorella sp. is one of the most common algae strains to produce biodiesel for several reasons. Primarily, these species have lipid amount of up to 57.3% and triacylglycerols are the main components among lipid content (see Table 10.1). Subsequently, growth properties are also an important parameter and Chlorella has high biomass productivity with the ability of surviving in different conditions. Also, the invasive behavior of this strain against other species is an advantage for large-scale biodiesel production [13, 49]. In addition to lipid content, fatty acid profile is another crucial point which is related to the length and degree of saturation based on four key points iodine value, oxidation stability, cetane number, and the cold filter plugging point. Fatty acids contain saturated and unsaturated groups that have 12–22 carbon atoms. The ideal mixture of fatty acids has been suggested to be C16:1, C18:1 and C14:0 in the ratio 5:4:1 [50, 51]. In this sense, Chlorella sp. is suitable for biodiesel production because of its fatty acid profile which includes palmitic acid (C16:0), palmitoleic acid (C16:1), stearic

Lipid content (% dw)

37.2

2.8-12.3

13.3- 44.3

30-45

11.55

22.84-28.23

32.1-36.5

35.5

8.93-19.68

30.34-54.56

14-57.3

13.5-40.9

20-40

Strain

Amphora sp.

Ankistrodesmus gracilis

Asterarcys quadricellulare

Botryococcus braunii

Chaetoceros calcitrans

Chaetoceros muelleri

Chlamydomonas sp.

Chlorella minutissima

Chlorella pyrenidose

Chlorella sorokiniana

Chlorella sp.

Chlorella vulgaris

Chlorococcum sp

Table 10.1 Lipid contents of different microalgae strains.

0.8-9

4.79-9.56

10-130

9.09-13.5

7.73-73.81

17.3

20–40

–

0.5

70-84

–

–

10.4

Lipid productivity (mgL-1day-1)

[34]

[33]

[29]

[47]

[31]

[30]

[29]

[28]

[27]

[26]

[25]

[24]

[23]

Reference

(Continued)

324 Liquid Biofuel Production

Lipid content (% dw)

30.8

16.51-50.5

21.4

19.8-47.8

61.16-79.89

9-24.62

32.38 - 46.28

20-25

12.6-41.4

34.36

14-50.6

27.12-52.13

29.1

26.4

Strain

Chrysotile carterae

Coccomyxa subellipsoidea

Crypthecodinium cohnii

Desmodesmus sp.

Dunaliella salina

Euglena gracilis

Graesiella sp.

Haematococcus pluvialis

Micractinium sp

Monoraphidium dybowskii

Mychonastesafer

Nannochloropsis oculata

Nannochloropsis sp.

Navicula phyllepta

114

10.2

25-146

50.7

40.01-54.39

20-80

–

64.8

–

21.95-104.4

40-100

100-600

45.85-232.37

9.9

Lipid productivity (mgL-1day-1)

Table 10.1 Lipid contents of different microalgae strains. (Continued)

[44]

[23]

[43]

[42]

[41]

[29]

[40]

[39]

[38]

[37]

[29]

[36]

[35]

[23]

Reference

(Continued)

Strategies for Biodiesel Production 325

Lipid content (% dw)

27.66

16.3-42.48

27.2-36.2

35.97–47.39

5.55

32.9

12-23.3

36

Strain

Nitzschia sp.

Phaeodactylum tricornutum

Scenedesmus abundans

Scenedesmus sp.

Skeletonema sp.

Tetraselmis suecica

Thalassiosira sp

Tisochrysis lutea

7.8

2.2-10.4

13.6

0.46

15.53

5.3 – 17.1

128.43

0.19

Lipid productivity (mgL-1day-1)

Table 10.1 Lipid contents of different microalgae strains. (Continued)

[23]

[48]

[23]

[27]

[32]

[46]

[45]

[27]

Reference

326 Liquid Biofuel Production

Strategies for Biodiesel Production 327 acid (C18:0), oleic acid (C18:1) and linoleic acid C(18:2) which has a nutritional value [52]. In an advanced stage, various simple modifications can be introduced to the strain or cultivation systems to increase the lipid accumulation. Applying nutrient stress or manipulating environmental conditions like temperature, pH, light intentsity and salinity are some examples of these alterations to enhancing lipid content. Several studies reported that green microalgae species tend to starch accumulation under optimum conditions due to their metabolic activities. However, the cells begin to synthesize higher amounts of triacylglycerols (TAGs) when they are exposed to stress conditions [53]. Wei and Huang [54] conducted a comprehensive study based on the effects of multi-factor and multi-level stresses on cultivation of Nannochloropsis oculata to obtain high lipid content. With this aim, cells were exposed to the multi-factor collaborative stress condition as high irradiation, nitrogen deficiency and elevated iron supplementation. Total lipid content (% DW) showed an increasing trend, from 28.52% to 49.14% on day 18; likewise the TAG proportion (% total lipid) increased from 8.85% to 78.16%. In another study, magnesium aminoclay nanoparticles were used to enhance lipid production of mixothropicChlorella sp. and maximum microalgal lipid productivity with nanoparticles was estimated to be 410 mg fatty acid methyl esters/L/d, which was 25% higher than control group [55]. A common approach to increase lipid production is changing salt concentration in culture medium. It was reported that the lipid productivity of 19.66 mgL-1d-1 occurred in the medium with increasing NaCl concentration, about 2.16 times as high as that in the control group for Chlorella sorokiniana cultivation [32]. Performing cultivation under green LED light, addition of different compounds such as cupric ion, phytohormones, crom and cadmium to the medium, photoperiod changes, nutrient deprivations are some type of manipulations that have been used for an efficient lipid synthesize from different species [56–59].

10.2.4

Microalgae Cultivation

After determination of the appropriate microalgae strain, their cultivation has been carried out with different production systems which are classified mainly into two groups as open ponds and closed photobioreactors (PBR). Open ponds are the most extensive cultivation systems, comprise of artificial lakes, shallow ponds, tanks, circular stirred and paddle wheel raceway ponds. The main advantage of these systems is that sunlight as the source of light energy is provided directly from the surface of ponds.

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Liquid Biofuel Production

Also, they are easy to use and maintain with their lower construction and operation costs [60]. Notwithstanding these advantages, there are numerous limitations associated with their large-scale operations. These systems require large areas of land and their operation performance can be negatively affected by environmental conditions. Increase in temperature value leads to high evaporation rates and consequently changes in salinity of culture medium [61]. The greatest drawback for open systems is culture contamination by other microorganisms. It prevents growth of desired microalgae and can cause mutation of cells. Other limitations in these systems are poor utilization of light by microalgae cells, low mass transfer and loss or diffusion of CO2 into the atmosphere [62]. There are several studies that have focused on investigation of lipid productivity in open ponds for biodiesel formation. Wang et al. [39] explored CO2 fixation rates, growth, lipid contents, fatty acid profiles, and lipid productivities of Graesiella sp. microalgae in an open raceway pond. It was shown that CO2 utilization efficiency and pH was in a positive correlation and CO2 fixation by microalgae was provided with open system to obtain an efficient lipid production. These studies about open cultivation ponds claim that low biomass and product yield is obtained in these systems. Recently, many researches have generally been performed to raise lipid productivity from microalgae. As an example, Acutodesmus obliquus microalga was cultivated in an open pond for enhancing biodiesel production, and to optimize process macro-element composition was evaluated. The results achieved in open pond cultivation using optimized medium were approximately 2 times higher than in the control medium in terms of lipid productivity [63]. To overcome limitations of open ponds, closed photobioreactors are recommended for large-scale microalgae cultivation. Closed PBR is a device made up of different transparent materials such as glass, polyethylene, plexiglass, polycarbonate or polyvinyl chloride (PVC) with novel construction designs. It consists of an internal or external illumination that its intensity affects the microalgae growth to provide a controlled bioconversion of CO2 into biomass and bioproducts [60]. PBRs may include different types of stirrers like blades or mixing can be provided with aeration from diffusers. Their control in terms of important factors such as temperature, pH, mixing, dissolved gas concentration, evaporation loss, light intensity and period is much easier than open systems; thereby, biomass yield can be increased according to target product [64]. The main advantage of PBRs is protection of culture from contamination by unwanted microorganisms. Closed photobioreactors include several types such as vertical/horizontal tubular, flat plate, airlift, bubble

Strategies for Biodiesel Production 329 column, membrane or hybrid, big-bag or plastic foil, floating type PBRs [62, 65]. Despite their advantages, closed PBRs also have some issues mostly about high construction and operation cost. The scale-up process in closed systems is complicated compared to the open systems. Novel photobioreactor designs are discussed in subsequent sections (see Sections 10.4 and 10.5).

10.2.5

Harvesting and Lipid Extraction

When the cultivation period is finished, biomass should harvest with an efficient and cost-effective method. This process is one of the most critical and complicated stages for a sustainable biodiesel production because when the whole cultivation process is considered, harvesting and dewatering have the highest operational cost due to energy consumption. An optimal harvesting technique should be determined according to cell morphology, size and medium density, as well as its energy consumption, which should be as low as possible. During the process, cell content must be protected until extraction. Microalgae harvesting methods include gravity sedimentation, flocculation, centrifugation, filtration, flotation, electrophoresis techniques or combining of several methods [22, 62, 66]. The most common commercial harvesting method is centrifugation, but its energy consumption is too high and it is only useful for valuable end products. Abomohra et al. [67], revealed a study to evaluate a chemical flocculation method to harvest S. obliquus grown on municipal wastewater as an alternative harvesting technique to the centrifugation. At the end of the study, among the inorganic flocculants ferric sulphate showed maximum relative flocculation efficiency of 99.5% and flocculation enhanced biodiesel yield by 40.9% over centrifugation. In a similar study, the effect of different harvesting methods on biomass composition in terms of lipid content and fatty acid composition was evaluated. Alkaline flocculation was reported an excellent alternative for primary harvesting algal biomass without an effect on lipid extraction efficiency of Phaeodactylum tricornutum [68]. In short, recent studies have focused on improvement of flocculation technique to provide an efficient microalgae harvesting [69–71]. Following biomass (5–15% dry weight) harvesting, the concentrated microalgae are transfered for drying where excess water is removed. The drying process must be quickly carried out since algal biomass has a risk of degradation induced by internal enzymes in a short time, especially in a hot environment. Frequently used drying techniques are freeze

330

Liquid Biofuel Production

drying, spray-drying, drum-drying, vacuum evaporation and sundrying [13, 22]. Recent studies have focused on lipid extraction from wet biomass because of excessive energy consumption of the dewatering process. This treatment consists of disruption of the algal cell walls in the medium where the microalgae is cultivated in. It decreases total cost as elimination of energy requirement; however, the yield is much lower than obtained in dry biomass. Ansari et al. [72], reported a similar result in a previous study that aimed to evaluate the extraction of lipids from dry and wet biomass of Scenedesmus obliquus. Lipid yield in wet biomass was found less than the dry biomass in this study. So, lipid extraction from wet biomass needs detailed research and improvement to achieve an industrial extraction process for biodiesel production from microalgae. According to previous studies, a pretreatment or cell disruption stage prior to wet extraction may be an efficient development that helps to increase lipid accessibility and improve mass transfer. A common technique for pretreatment is applying electric fields, such as the pulsed electric field, as promising for intracellular compounds extraction from wet biomass [73]. Silve et al. [74] found that pulsed electric fieldtreatment has hopeful features to raise lipid yield from wet biomass and in terms of biorefinery approach since process performs under average temperature values and waste amount is relatively low compared to the direct extraction. Microalgae-based biodiesel supply chain continues with the cell disruption and oil extraction stage which aims to damage cell walls to provide releasing of intracellular content. Various methods can be applied depending on the microalgae morphology and lipid properties. They are basically divided into two groups as mechanical and non-mechanical techniques. Mechanical methods include high pressure homogenization, bead milling, ultrasonication, autoclave, hydrodynamic cavitation, microwave, pulsed electric field and freeze-drying where utilization of chemicals and contamination of the leftover biomass are avoided so it provides an ecofriendly approach [75]. These processes show high energy consumption as a result of shear forces, electrical pulses, waves or heat. However, product recovery yield is excessive and scale-up is much easier than other methods. On the other hand, non-mechanical disruption methods comprise a high amount of chemicals with low energy requirement. Osmotic shock, use of organic solvents, supercritical fluid extraction, acid, base and enzyme reactions are some of the non-mechanical disruption methods [76]. The most commonly used method is solvent extraction, which contains several types of chemicals like ethanol, methanol, hexane, chloroform, diethyl ether, etc. The use of different organic solvents have some

Strategies for Biodiesel Production 331 limitations since they are highly flammable and toxic, and recovering the solvent is too difficult. In recent studies, the supercritical extraction method, which is nontoxic, easy to recover and applicable at low temperatures, is recommended as an alternative to solvent extraction. Aliev and Abdulagatov [77] revealed that applying various extraction techniques as pure supercritical-CO2, supercritical-CO2 with acetone as co-solvent and solvent extraction for microalgae have no big difference on the total extract yield and fraction of fatty acids. However, extraction with supercritical-CO2 has some advantages, as already mentioned, in comparing with solvent extraction. Thus, it has been recommended for fatty acid extraction from microalgae. There are similar studies that indicate the use of supercritical extraction for an efficient lipid recovery [78, 79].

10.2.6

Conversion of Microalgal Oil to Biodiesel

After the extraction of lipids from microalgae, transesterification process, which is a chemical reaction that converts microalgal oils (triacylglycerols) into fatty acid methyl esters (FAMEs), should be performed to produce biodiesel since the viscosity of extracted microalgal oil is too high to use in biodiesel production process. Transesterification includes a catalyst; as acid, base or enzyme, and a mono-alcohol which shift the reaction in order to obtain FAME and glycerol (Figure 10.1). This operation is divided into three groups as catalytic, non-catalytic and in-situ where the extraction and transesterification processes occur simultaneously transesterifications according to the type of catalysts [62, 80]. Prabakaran et al. [81], showed that the enzymatic transesterification with lipase that isolated from Pseudomonas sp. has a strong potential for biodiesel production because at the end of the reaction a high purity FAME are formed and glycerol can be removed easily. In another study a novel catalyst, sea shell, was employed as catalyst and lipid from Aphanothece halophytica microalga was effectively converted to FAME using cost-effective and eco-friendly white clam shell catalyst [82].

CH-OCOR2 CH2-OCOR3 Triglyceride (oil)

R1-COOCH3

CH2-OH

CH2-OCOR1 +

3HOCH3 Methanol (alcohol)

catalyst

CH-OH CH2-OH (Glycerol)

R2-COOCH3

+

R3-COOCH3 Methyl Esters (Biodiesel)

Figure 10.1 Transesterification of microalgal oil to biodiesel [83].

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Liquid Biofuel Production

10.3 Metabolic Engineering Approaches for Biodiesel Production Metabolic engineering strategy aims to increase the valuable compound content of microorganisms with some modifications on metabolic pathways in cells. The first and foremost part of this approach is understanding metabolic pathway and signal network for product formation to trigger the target metabolite production. Through the developing genome databases and previous studies, lipid biosynthesis pathways were characterized for microalgae. An uncomplicated metabolic pathway for a green microalgae which is valid for many types of species is shown in Figure 10.2. The Figure represents a simple progress for fatty acid (FA), triacylglycerol (TAG), and polyunsaturated fatty acid (PUFA) biosynthesis. Briefly, the FA synthesis is originated from pyruvate and specificially begins with the conversion of acetyl CoA to malonyl CoA, catalyzed by acetyl CoA carboxylase enzyme (ACCase). At the end of this pathway free fatty acids are synthesized which induce the formation of triacylglycerols (TAGs) primarily in the

Pyruvate PDC Acetyl-CoA TAG

ACCase*

DGAT*

Malonyl-CoA

LPAT

MAT

LPAAT*

Malonyl-ACP

G3P + GPAT* Acyl-CoA LACS FAS complex TE*

Free Fatty acids

PUFA Multiple Desaturases* & Elongases

Figure 10.2 Representation of biosynthetic pathways for fatty acids (FAs), triacylglycerols (TAGs) and polyunsaturated FAs in green microalgae. PDC, pyruvate dehydrogenase complex; ACCase, acetyl-CoA carboxylase; MAT, malonyl-CoA/ACP transacylase; ACP, acyl-carrier protein; LACS, long-chain acyl-CoA synthetase; FAS, FA synthase; TE, fatty acyl-ACP thioesterase; GPAT, glycerol-3-phosphate acyltransferase; LPAAT, lysophosphatidic acid acyltransferase; LPAT, lyso-phosphatidylcholine acyltransferase; DGAT, diacylglycerol acyltransferase; CoA, coenzyme A; G3P, glycerate-3-phosphate; TAG, triacylglycerol; PUFA, polyunsaturated FA [84].

Strategies for Biodiesel Production 333 chloroplast of cells. Also, in a different pathway starting from FA, PUFA synthesis was carried out [84]. A novel research area called “Omics” technology is a promising technology; its purpose is understanding the behavior of biological systems in detail and improvement of cellular content, indirectly. The “Omics” biology is performed predicting the role among cellular compounds which includes transcriptome, proteome, and metabolome and their interactions in the cells. Owing to this technology the identification of major enzymeencoding genes can be provided, and the metabolic pathways involved in the biosynthesis and degradation of metabolites can be modified [85]. After all, lipid synthesis can be improved by identifying the important component in this pathway and manipulating them with metabolic engineering tools. The metabolic engineering approach includes some strategies to achieve high lipid formation, such as flux balance analysis, improving photosynthetic efficiency, increasing the availability of precursor molecules such as acetyl-CoA, applying enzymes toward lipid synthesis, inducing overexpression of major genes, downregulating the catabolism of fatty acids by inhibiting β-oxidation or lipase hydrolysis and transcription factor engineering [86]. Several researchers investigated the metabolic engineering designs for biodiesel production [87–100]. Recent metabolic engineering applications that involve different strategies for enhancing lipid production are summarized in Table 10.2. Scientific studies basically focused on determination of the complete genome sequence of microalgae and some of these projects have been completed, including those for Chalimydomonas reinhardtii, Phaeodactylum tricornutum, Thalassiosira pseudonana, Cyanidioschyzon merolae, Ostreococcus lucimarinus and Ostreococcus tauri [101]. Among these species Chlamydomonas reinhardtii is the most studied microalga that becomes the model organism for biofuel production as its genome has been sequenced, and there is an extensive molecular tool set available for this specie. As also shown in Table 10.2, one possible and common strategy for lipid accumulation in microalgae is by overexpressing the key enzymes involved in fatty acid synthesis. Xue et al. [94], reported that overexpression of glucose-6 phosphate dehydrogenase lead to up regulation of some metabolisms especially pentose phosphate pathyway (PPP) and glycolysis. This up-regulation caused more carbon flow to fatty acid synthesis, thus lipid accumulation increased. Suppression of lipid catabolism is another strategy for enhancing lipid accumulation in microalgae. This method is based on knock-down of a multifunctional lipase/phospholipase/acyltransferase enzyme for

Type of modification

Mutagenesis with inhibitors: the herbicide quizalofop-P-ethyl, and chemical mutagen, ethyl methanesulfonate (EMS)

Regulation of rfpgene expression by CRISPRi

Overexpression of the acyl-ACP thioesterase from Dunaliella tertiolecta in C. reinhardtii

Expression of plant lauric acid-biased TE (C12TE) and MCFA-specific ketoacyl-ACP synthase (KASIV)

NsPDK knockdown via RNA interference strategy

Heterologous expression of AtWRI1 transcription factor

Microalgae used

Chlorella sp

Chlamydomonas reinhardtii

Chlamydomonas reinhardtii

Dunaliella tertiolecta

Nannochloropsis salina

Nannochloropsis salina

Enhancement of neutral lipid and FAME production

Enhancement of total fatty acids and faster TAG accumulation

(Continued)

[90]

[89]

[88]

[92]

Increased total lipid 15% DCW, representing a 56% improvement from the wild-type 7 and 4 fold increase in lauric acid and myristic acid accumulation

[87]

[99]

Reference

%74.4 higher lipid content than wild-type

59 and 53% higher lipid content and lipid productivity than the wild type

Outcome

Table 10.2 Metabolic engineering applications to enhance lipid accumulation.

334 Liquid Biofuel Production

Type of modification

Inhibition TOR kinase enzyme with AZD-8055 inhibitor

Overexpression of the PtLDP1 gene

Overexpression of glucose-6 phosphate dehydrogenase (G6PD)

Artifcial miRNA inhibition of phosphoenol pyruvate carboxylase

Overexpression of DGAT2

Overexpression of NoD12 under the control of the stress inducible promoter

Overexpression of DGAT2

Expressing a quadruple-genes construct (GPAT – LPAAT – PAP DGAT) in Kennedy pathway

Microalgae used

Phaeodactylum tricornutum

Phaeodactylum tricornutum

Phaeodactylum tricornutum

Chlamydomonas reinhardtii

Nannochloropsis oceanica

Nannochloropsis oceanica

Thalassiosira pseudonana

Chlorella sp.

6% (wt) of TAG and 60% (wt) of total lipid content

1.52 to 1.95 fold higher level of TAG than the wild-type

Increasing long-chain PUFAs and TAG production

Increased neutral lipid content by 69% in engineered microalgae

Inhibiton of transcription of CrPEPCs and signifcant increase of total fatty acids

2.7 fold increase in neutral lipid content compared to wild type

Increasing total lipid content significantly compared to wild type

Promoting the accumulation of neutral lipids and enhancement of TAG productivity

Outcome

Table 10.2 Metabolic engineering applications to enhance lipid accumulation. (Continued)

[98]

[97]

[96]

[100]

[93]

[94]

[93]

[91]

Reference

Strategies for Biodiesel Production 335

336

Liquid Biofuel Production

improvement of the accumulation of fatty acids and enhancement of lipid content [21]. This method was used in Nannochloropsis salina and total fatty acid accumulation was enhanced with acceleration of TAG formation [89]. This method has some disadvantages such as negative effects on proliferation and biomass productivity in some species due to interruption of energy delivering system. A novel strategy for both lipid production from microalgae and other metabolic engineering studies is the clustered regularly interspaced short palindromic repeat (CRISPR)/Cas9 system, which is the easiest and most efficient genome editing technology. This complex is a ribonucleoprotein compound consisting of a bacteria-derived DNA endonuclease is called Cas proteins which contains specific domains that cleave opposite strands of a DNA sequence and a small processed CRISPR RNAs [102]. The main advantage of CRISPR/Cas9 system is that several different sequences can be targeted with a single enzyme so a small amount of enzymes will be sufficient to obtain desirable results. However, research about CRISPR/Cas9 is still limited so there are many concerns to apply this system for valuable compound production [103]. Kao and Ng [87] studied the attemption to apply CRISPRi on gene regulation in microalgae and in the study it was revealed that dzCas9/sgRNA system suppressed the expression of exogenous genes in Chlamydomonas reinhardtii with down-regulation of the genes. For enhanced lipid production, the downregulated strain generated the highest lipid content and productivity that are 74.4% and 94.2% higher than that of the wild-type. Besides the above-mentioned methods, there are other metabolic techniques such as blocking metabolic pathways that lead to the accumulation of other energy-rich storage compounds and improved genetically engineered microalgae to increase the quality of the lipids. Consequently, the metabolic engineering approach for enhancement lipid (TAG) accumulation is a promising area that meets the energy demand of the world in the long term. However, these studies are still in the research stage since the lack of enough information causes several concerns and risks. The most serious risk is transfer of genes from genetically modified microalgae to other organisms within the invaded ecosystem. Also, results of metabolic studies may be unpredictable because there is not sufficient research for microalgae species. Despite these limitations, the metabolic engineering approach still represents one of the best options to obtain high biodiesel yield. The optimization researches should be carried out to overcome these problems and to develop an efficient biodiesel production method with a low-cost and sustainable approach.

Strategies for Biodiesel Production 337

10.4 Novel Photobioreactor Designs for Biodiesel Production Higher productivity and yield become more and more important regarding the increasing capacity of the biodiesel market. Thus, as a bioengineered approach, designing novel PBRs shows an ascending tendency among producers. In order to achieve a high volumetric productivity, it is crucial to ensure that the light capture of microalgae is appropriate and sufficient. In addition, geometry, biological and hydrodynamic parameters (turbulence, gas exchange, and nutrient requirements) inside the photobioreactor affect biomass and product concentration. In order to achieve internal illumination, Hincapie and Stuart [104] used fiber optics to distribute photons inside the culture media in a 28-L-air-lift-photobioreactor for biomass production. Their design was established to induce light/dark cycles to the microalgae and it was reported that this design could significantly reduce costs and increase durability. In another study which was also built to enhance the lit absorbance, an internal illumination using LED lights and Plexiglas End-Lighten rods were implanted to a novel photobioreactor [105]. Mixing is another critical issue in PBRs. It is necessary for an appropriate heat and mass transfer. Airlift PBRs are generally preferred as a result of their low energy consumption, adequate mixing and sufficient gas–liquid mass transfer [106]. In order to improve performance and mass transfer in airlift PBRs, different baffle types were investigated. For example, Li et al. [107] demonstrated that double waved baffles, as a vertical-baffle type, enhanced the O2 transfer rate in comparison to conventional baffles due to the increased average turbulent kinetic energy. In another paper, Chen et al. [108], investigated a flat-plate airlift PBR with two types of baffles (conventional flat and waved) in terms of their functioning for CO2 mass transfer. According to the simulation studies it was resulted that with the increase of downcomer-to-riser cross-section area ratio (Ad/Ar) the volumetric mass transfer coefficient (kLa) increased first and then decreased for single baffle while for double baffles, kLa decreased with the increase of Ad/Ar. In addition to those, in the same study it was shown that the waved baffle had higher kLa values at lower Ad/Ar than flat baffles. The effect of baffles in microalgal productions is not limited to PBRs. Open ponds are also investigated for different types of baffles to improve mass transfer rate. Mendoza et al. [109] studied the usage of a carbonation column for CO2 mass transfer into microalgal culture ponds and the efficiency of mass transfer rates were increased to 106 g/h, 172 g/h, 27 g/h, and 39 g/h, for the

338

Liquid Biofuel Production

paddlewheel, sump, straight and curved channel sections, respectively. In another research, a novel oscillating gas aerator integrated with an oscillating baffle was used to increase oxygen mass transfer by producing smaller aeration bubbles in a raceway pond [110]. It was concluded that with this new design, the approximate diameter of bubbles and their production durations decreased with decreased aeration gas rate and orifice diameter, and increased water velocity. To summarize the study, it was shown that the mass transfer coefficient was enhanced by 15% and mixing time was reduced by 32% resulting in a 19% higher microalgal biomass yield when the oscillating gas aerator and oscillating baffle are used in a raceway pond.

10.5 Advanced Photobioreactor Configurations and Kinetics Microalgal biomass kinetics and oil productions are basically affected by various factors. Lighting, nutrient supply and bulk content, bioreactor configuration and geometry (mixing and aeration type), mass transfer, temperature and pH are considered the main factors. In this regard, researchers are developing new configurations to enhance microalgal conversion and growth rate. Yang et al. [111], used a water-circulating column photobioreactor to decrease mixing time for growing microalgal biomass and it was concluded that the distance between the aeration system - solution surface and the diameter of the sparger should be optimized to decrease the energy consumption. In another study, Kim [112] designed a façade where microalgae was used to provide the energy requirements of a building. In the study it was shown that, with 3 g biodiesel production in a day per a façade, the façade system can supply 1100 gallons of biodiesel to the whole building in a year. Tubuler PBR systems were also investigated in terms of biodiesel production as tubuler PBRs are considered to have a superior type for microalgal biomass production. Recently, a novel concentric double tubes with radial aerated pores along the length direction of inner tube was developed and the biomass productivity increased by 43.6% compared to conventional axial tubes [113]. It was also found that novel aeration style could provide efficient mixing and higher biomass concentrations. In a previous study, for an air-lift PBR a new device and a method were used in order to increase specific growth rate of a microalgae where Chlorella sp. was used as a model, and it was reported that the specific growth rate constant of the novel system was 0.011 h-1 which is higher than conventional production [104]. In another study aiming to culture microalgae with

Strategies for Biodiesel Production 339 higher efficiencies, a novel vertical multi-column airlift photobioreactor was developed with a series of vertically arranged parallel columns [114]. Accordingly, when using Chlorella pyrenoidosa as a model microalgae, biomass production was reached 1.56 g L-1. Cost effectiveness is another crucial parameter for industrial processes. Hincapie and Stuart [104] designed a floating horizontal photobioreactor which was assumed to be inexpensive and scalable. The effectiveness was shown using a model microalgae, Nannochloris atomus Butcher CCAP 251/4A, in a 65-L prototype bioreactor and it was reported that a biomass concentration of 4 g L-1 and a productivity of 12.9 g m-2 d-1 were succeeded with a closed PBR under artificial illumination of 435 μmol m-2 s-1. In an open sytem, they reported to have achieved an average biomass productivity of 18.2 g m-2 d-1 without any contamination during 165 days. Prior to research the effects of an industrial PBR, Ojo et al. [115] investigated the effects in a miniature PBR which represents a potential platform technology for the biomass production with a high productivity level. In the study a novel orbitally shaken twin-well miniature PBR was developed in optimal conditions and it was discovered that kLa values were enhanced up to 80 h−1 by Chlorella sorokiniana at 20 μmol m−2 s−1 illumination. The highest biomass production and productivity (9.2 g L−1 and 2.5 g L−1 d−1, respectively) were achieved where 5% CO2 with a light intensity of 380 μmol m−2 s−1 was used at 300 rpm. Mixing is a critical issue and a configuration parameter in PBRs. Novel flat-plate photobioreactors (PBRs) with special agitators were designed and the results of fluid dynamics were compared with the control reactor without mixer and it was concluded that the PBRs with novel agitators can effectively increase liquid velocity and the algal growth rates of Chlorella pyrenoidosa where a biomass concentrations up to 42.9% (1.3 g L-1) higher than control was achieved [116]. In the study, the results of the fluid dynamics showed that the correlation analysis of microalgal growth rate with the characteristics of mixing and light regime can be considered as a key factors affecting microalgal growth rate. In another study where different agitators were used, radial velocity of fluid and light/dark cycles within a flat-plate photobioreactor was investigated using computational fluid dynamics [117]. The study showed that a maximum biomass concentration of 0.89 g L-1 could be achieved using archetype mixers, whereas in the control PBR only 0.67 g L-1 biomass could be optimized. Moreover, fluid velocity along the light attenuation and light/ dark cycles was also enhanced according to the results of computational fluid dynamics.

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10.6 Conclusions Microalgal biodiesel production is considered as a promising methodology for future applications with several benefits, such as renewability, sustainability, clean-burning, safety and cost-effectivenes issues. Therefore, new microalgal sources using molecular engineering techniques have been developed. Nevertheless, researchers are still searching for a better method for a higher productivity with a higher stability to be used in industrial processes. Recently, novel techniques with novel PBR configurations have been developed in order to increase the usage of microalgal biorefineries, especially biodiesel. Lower productivity and higher specifications with higher qualifications are still the main bottlenecks and limitations for large-scale applications, despite their benefits. With further analysis, along with more studies to show the novel advantages of microalgal biodiesel, industrial applications will be higher in the future.

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102. Lenka, S.K., Carbonaro, N., Park, R., Miller, S.M., Thorpe, I., Li, Y., Current advances in molecular, biochemical, and computational modeling analysis of microalgal triacylglycerol biosynthesis. Biotechnol. Adv., 34, 5, 1046–1063, 2016. 103. Wang, Q., Lu, Y., Xin, Y., Wei, L., Huang, S., Xu, J., Genome editing of model oleaginous microalgae Nannochloropsis spp. by CRISPR/Cas9. Plant J., 88, 6, 1071–1081, 2016. 104. Hincapie, E. and Stuart, B.J., Design, construction, and validation of an internally lit air-lift photobioreactor for growing algae. Front. Energy Res., 2, 65, 2015. 105. Pozza, C., Schmuck, S., Mietzel, T., A novel photobioreactor with internal illumination using Plexiglas rods to spread the light and LED as a source of light for wastewater treatment using microalgae. Proceedings of the IWA Congress on Water Climate and Energy, pp. 1305–18052012, 2012. 106. Rengel, A., Zoughaib, A., Dron, D., Clodic, D., Hydrodynamic study of an internal airlift reactor for microalgae culture. Appl. Microbiol. Biotechnol., 93, 1, 117–129, 2012. 107. Li, Y., Qi, X., Liu, T., Chen, Y., Investigation on hydrodynamic behaviors in a wave-baffled panel photobioreactor [J]. Chin. J. Process Eng., 5, 005, 2010. 108. Chen, Z., Jiang, Z., Zhang, X., Zhang, J., Numerical and experimental study on the CO2 gas–liquid mass transfer in flat-plate airlift photobioreactor with different baffles. Biochem. Eng. J., 106, 129–138, 2016. 109. Mendoza, J., Granados, M., de Godos, I., Acién, F., Molina, E., Heaven, S., Banks, C., Oxygen transfer and evolution in microalgal culture in open raceways. Bioresour. Technol., 137, 188–195, 2013. 110. Yang, Z., Cheng, J., Lin, R., Zhou, J., Cen, K., Improving microalgal growth with reduced diameters of aeration bubbles and enhanced mass transfer of solution in an oscillating flow field. Bioresour. Technol., 211, 429–434, 2016a. 111. Yang, Z., Cheng, J., Yang, W., Zhou, J., Cen, K., Developing a watercirculating column photobioreactor for microalgal growth with low energy consumption. Bioresour. Technol., 221, 492–497, 2016b. 112. Kim, K.H., Beyond Green: Growing Algae Facade. Proceedings of the ARCC Conference Repository, Charlotte, NC, USA, 12–15 February 2014, 2014. 113. Yang, J., Cui, X., Feng, Y., Jing, G., Kang, L., Luo, M., Experimental study on microalgae cultivation in novel photobioreactor of concentric double tubes with aeration pores along tube length direction. Int. J. Green Energy, 14, 15, 1269–1276, 2017. 114. Huang, J., Ying, J., Fan, F., Yang, Q., Wang, J., Li, Y., Development of a novel multi-column airlift photobioreactor with easy scalability by means of computational fluid dynamics simulations and experiments. Bioresour. Technol., 222, 399–407, 2016. 115. Ojo, E.O., Auta, H., Baganz, F., Lye, G.J., Design and parallelisation of a miniature photobioreactor platform for microalgal culture evaluation and optimisation. Biochem. Eng. J., 103, 93–102, 2015.

Strategies for Biodiesel Production 349 116. Huang, J., Li, Y., Wan, M., Yan, Y., Feng, F., Qu, X., Wang, J., Shen, G., Li, W., Fan, J., Novel flat-plate photobioreactors for microalgae cultivation with special mixers to promote mixing along the light gradient. Bioresour. Technol., 159, 8–16, 2014. 117. Huang, J., Feng, F., Wan, M., Ying, J., Li, Y., Qu, X., Pan, R., Shen, G., Li, W., Improving performance of flat-plate photobioreactors by installation of novel internal mixers optimized with computational fluid dynamics. Bioresour. Technol., 182, 151–159, 2015.

11 Bio-Oil Production from Algal Feedstock Naveen Dwivedi1* and Shubha Dwivedi2 1

Associate Professor and Head, Department of Biotechnology, S. D. College of Engineering and Technology, Muzaffarnagar, India 2 Assistant Professor, Department of Biotechnology, Meerut Institute of Engineering and Technology Meerut, India

Abstract For many decades, the majority of the population has relied on fossil fuel as a source of energy. We are totally dependent on its various usages in our daily life. Due to never-ending demand and limited storage of this exhaustible source of energy, researchers have been encouraged to find an alternate source of energy. The technology of the formation of biofuel is not new and over the last several years many researchers have worked on the production of biofuel from sugars, grain, kitchen waste, lignocellulosic material, etc. In recent years the whole momentum is focused on the production of biofuel from algal species. It constitutes the fourth generation of biofuel production from algal blooms and high solar efficiency cultivation. This chapter describes the various technologies used for the production of bio-oil from algal biomass and the methods of the up-gradation of the biofuel. Keywords: Bio-oil, pyrolysis, biofuel, algal feedstocks, transestrificaion

11.1 Introduction The continued combustion of fossil fuels has created critical environmental concerns worldwide. Human population depends on petroleum-based fuel as a source of energy since its discovery years before the Christian (BC) era. The growing demand for this non-renewable source of energy, its depletion and cost, the full dependency of all humankind on its usage as a principal transportation fuel has encouraged researchers to find an *Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (351–372) © 2019 Scrivener Publishing LLC

351

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alternative fuel from renewable sources, capable of replacing fossil fuels. In the past 60 years, the per capita energy consumption has increased manyfold, from approximately 1 tons coal equivalent in 1951 to approximately 2.5 tons coal equivalent in recent years. More than 75% of the world energy requirement has been met through fossil fuels. However, scarcity of fossil fuels as well as their gradually decreasing quality in terms of sulphur and metal content, coupled with the stringent environmental norms, has imposed a threat to energy security in the 21st century. Thus, extensive research has been carried out in recent years on the development of clean technologies for the utilization of fossil fuels around the world. Alternative renewable energy sources such as biomass, solar energy, etc., are also being investigated widely to produce sustainable energy. It is forecasted that the consumption of fossil fuels will reach a plateau at around 2020 and renewable energy will supply up to 50% of the world energy demand by 2050 [1]. Biofuel technology is not new to mankind. Researchers have been working over decades to produce or use biodiesel. For example, Reliance Industries Limited, Jamnagar, is installing one pilot plant for production of oil from algae, and IOCL Faridabad has already begun research in this area. Apart from that, Natural Biochemicals and Foods Limited, Southern Online Bio Technologies Ltd, and Universal Biofuels Pvt. Ltd., are three Kakinada, Andhra Pradesh-based companies which have already commercialized the production of biodiesel from various renewable sources of energy. Currently, several biodiesel producers use vegetable oil from food crops such as soybean, canola, and palm to produce a transportation fuel with chemical and physical properties similar to petro diesel. However, the use of oil from food crops has created significant worldwide opposition due to the increase of food price and growing food shortage. The Food and Agriculture Organization (FAO) of the United Nations (UN) is persuading researchers to redirect biodiesel production to the use of non-edible crops. In addition, food-based crops alone cannot produce enough biodiesel to satisfy world demand. Lignocellulosic materials are the most promising feedstock as a natural and renewable resource essential to the functioning of modern industrial societies. A considerable amount of such materials are being generated as waste by-products through agricultural practices mainly from various agro-based industries. The use of lignocellulosic materials to replace fossil resources for the industrial production of fuels, chemicals, and materials is increasing. The carbohydrate composition of lignocellulose (i.e., cellulose and hemicellulose) is an abundant source of sugars. However, due to the feedstock recalcitrance, rigid and compact structure of plant cell walls, access to polysaccharides is hindered and release of fermentable sugars has become a bottleneck. Thus, to overcome

Bio-Oil Production from Algal Feedstock 353 the recalcitrant barriers, thermo chemical pretreatment with an acid catalyst is usually employed for the physical or chemical disruption of plant cell wall. After pretreatment, enzymatic hydrolysis is the preferred option to produce sugars that can be further converted into liquid fuels (e.g., ethanol) via fermentation by microbial biocatalysts. Biomass-derived fuels are promising alternates of fossil fuels. Bio-oil has been considered as an alternative energy source for fuel applications. Biooil is produced from biomass via two principal methods, first hydrothermal liquefaction and second pyrolysis. The hydrothermal liquefaction is conducted in an aqueous environment which makes it suitable for aquatic plants and wet biomass while pyrolysis needs drying of feedstock which increases the energy consumption of the process. One of the main challenges in bio-oil application is the poor quality of the bio-oil. Bio-oil has a high content of oxygen and acidity and thus cannot be used directly as liquid fuel, which requires upgrading of the bio-oil for fuel application [2, 3]. Algae have been considered as one of the most promising alternative sources for biofuel production, known as third-generation biofuels. Algae are a group of eukaryotic organisms, ranging from unicellular to multicellular forms. Microalgae are photosynthetic microorganisms which survive as individual cells or chains of cells [4]. Microalgae live in saline or fresh water environments and convert sunlight, carbon dioxideand water to algal biomass [5]. Microalgae have faster growth rates, higher oil contents, and less complex structures than macroalgae [3, 4]. Macroalgae are more difficult to grow in bioreactors; therefore they have not yet been researched as widely as microalgae.

11.1.1

Microalgae

Microalgae are single cell photoautotrophic (capable of synthesizing own food from inorganic substances using light as an energy source) or photoheterotrophic (use light energy but cannot use CO2 as source of carbon) microorganism. Photoautotrophic algal cells grow like plants through photosynthesis process, during which algal cells capture CO2 and photons, and convert them into biomass rich in lipid. More than 3000 algalstrains have been discovered and most of them live in an aquatic habitat such as sea, rivers and oceans [6]. Based on their habitat, microalgae are classified into fresh water and marine algae.

11.1.2

Classification of Microalgae

Microalgae can be classified based on their pigmentation, growth conditions, cells wall structure and flagellation. There are six phyla of algae:

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cyanobacteria, green algae, red algae, diatomaceae, Eustigmatophytes and Prymnesiophyceae. A brief description of each phylum of algae is given in Table 11.1. Algal cell may contain lipid up to 80% in mass depending on the strain, the growth medium composition and the culture conditions such as temperature, pH, carbon dioxide and photonic energy absorption [8, 9]. Table 11.2 shows a variety of microalgae, habitat and lipid content in gram lipid per 100 g dry algae. Table 11.1 Classification and description of microalgae phyla [7]. Algal phylum

Description

Green algae or Blue-green algae

Algal cells have green chloroplast that contains chlorophyll a and b. These cells have mitochondria. Some species have flagella.

Red algae or Rhodophycae

Cells have chloroplast with chlorophyll a and d, and phycobillins. They have double cell wall, but do not have centrioles and flagella.

Cyanobacteria

Class of prokaryotic cells that contain chloroplast with no chlorophyll. These are bacteria; but they are assimilated to algae due to their growth through photosynthesis process similar to microalgae. Some strains can grow in soil, marine or fresh water.

Eustigmatophytes

Class of eukaryotic algae which contain yellow-green chloroplast. They include strains growing in marine, freshwater or solid medium such as soil. Algae in this class have chloroplast containing chlorophyll a.

Prymnesiophytes or Prymnesiophycae

Class of algae in chlorophyll a-c phyletic line. Some strains of this group have one or two flagella. For example, Pavlova strain has smooth flagella of equal length.

Diatomaceae

Class of Bacillariophyceae. Diatomaceae cells have chloroplast carrying chlorophyll a and c. They have hard wall due to the presence of silica. Most of these cells can be found in fresh or salted sea. Majority of diatom species live in cold water.

Bio-Oil Production from Algal Feedstock 355 Table 11.2 Lipid content present in microalgae [10]. Algal strain

Habitat

Lipid content in g lipid per 100 g dry algae

Chlorella sp.

Freshwater

28-32

Amphidimium carteri

Marine

>20

Tetraselmis

Marine

15-23

Dunaliella primolecta

Marine

23

Isochrysis sp.

Marine

25-33

Thalassiosira pseudonana

Marine

>30

Nannochlorosis sp.

Marine

31-68

Porphyridium cruenturn

Marine

>40

Schizochytrium sp.

Marine

50-77

Monallanthus salina

-

>20

Phaeodactylum

-

20-30

Neochloris oleoabundans

-

35-54

Botryococcus braunii

-

25-75

Being photosynthetic microorganisms, microalgae require water rich in nutrient, carbon-dioxide and photonic energy for growth. Microalgae convert photonic energy, water and CO2 to sugars; then sugars are converted to macromolecules such as lipids or triacylglycerols (TAG) as shown in reactions (1.1) and (1.2) below; two-step reactions.

CO2 + water + photons Sugars

Sugars + O2 TAGs

(1.1) (1.2)

Lipids are believed to be the sustainable feedstock for biodiesel. In this process, microalgae are also sequestering the carbon from CO2.

11.1.3

Algae Growth

Algal cells multiply by cell division (mitosis). During the mitosis process, algal cells are divided into two identical daughter cells. Certain strains of

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algae such as diatoms Chaetoceros undergo gametes fusion through syngamy followed by mitosis [11]. It is believed that the syngamy process enhance algal cell enlargement in volume. Algal cells capture carbon from CO2 in an intermediate step and transform it into complex carbohydrate molecules such as lipids, which are the raw material in biodiesel production. Algal cells are either photoautotrophic or heterotrophic. Photoautotrophic cells capture carbon dioxide and photonic energy to convert into sugars (glucose), then lipid, as shown in Reactions 1.3 and 1.4.

6 CO2 + 6 H2O + 8 photons C6H12O6

C6H12O6 + 6 O2 TGA (or lipid)

(1.3) (1.4)

Heterotrophic cells, on the other hand, use other organic compounds as source of carbon and do not undergo photosynthesis. In photoautotrophic algae, the photosynthetic reaction takes place within the chloroplast. CO2, photons and water are key elements in algae growth process (photosynthetic reaction). It is important to note that algal cells can tolerate CO2 up to certain concentration. The level of tolerance depends mostly on the algal strains. Kurano and Miyachi [12] studied the impact of CO2 fixation in algal culture and showed that algal culture of certain Chlorella strains were inhibited by CO2 if its concentration exceeded 5%. However, certain Chlorella strains can reach high biomass productivity with CO2 exceeding 10%. Khan et al. [13] showed that Chlorella kessleri can reach high biomass productivity of 87 mg per liter per day with supplied CO2 exceeding 10%.

11.2 Technologies Used for the Production of Bio-Oil from Algal Biomass Bio-oil production from algae has been an area of interest and extensive research. Bio-oil is known as a promising alternative for crude oil to produce transportation fuels and extraction of valuable chemicals [14]. Biooil production from algae is different from lipid or oil extraction. Bio-oil production aims to break all bio macromolecules (i.e., lipid, protein and carbohydrate) into an organic liquid phase named bio-oil. Presently, two major types of processes for production of bio-oils from algae are flash pyrolysis and hydrothermal liquefaction (HTL).

Bio-Oil Production from Algal Feedstock 357 a. Flash Pyrolysis Flash pyrolysis involves the rapid thermal decomposition of biomass at medium to high temperatures (350–700°C) in the absence of oxygen, which results in the production of charcoal, bio-oil, and gaseous products. The main operating parameters in pyrolysis are temperature, residence time, and heating rate. Based on these parameters, pyrolysis can be categorized into three subgroups: slow pyrolysis, fast pyrolysis and flash pyrolysis. Table 11.3 shows the operating conditions and yield of products for pyrolysis processes [15, 16]. In comparison to fast and flash pyrolysis, the slow pyrolysis produces larger amounts of char; this char can be used as a solid fuel or for the production of syn-gas through gasification process. Fast and flash pyrolysis produce bio-oils in high yields of up to 80wt % dry feed which makes them favorable for bio-oil production. Pyrolysis oils have a higher oxygen content and consequently lower heating value than liquefaction oils [17, 18]. Pyrolysis requires dried feedstock which requires a prior drying step which increases the energy consumption of the entire process, although pyrolysis has a lower Table 11.3 Operating parameters and expected yields for pyrolysis processes [15, 16]. Pyrolysis types

Operating parameters

Liquid (%)

Gas (%)

Char (%)

Slow pyrolysis

Temp. (400°C), Residence time (more than 30 min), Heating rate (0.1–1°C/s)

30

35

35

Fast pyrolysis

Temp. (500°C), Residence time (about 10–20s), Heating rate (1–200°C/s)

50

30

20

Flash pyrolysis

Temp. (500°C), Residence time (about 1s), Heating rate (>1000°C/s)

75

13

2

358

Liquid Biofuel Production capital cost than liquefaction since it is conducted from 1 to 5 atm. Many pyrolysis technologies are being used commercially [19]. Pyrolysis of several algal species has been tested, including Chlorella [20, 21], Emiliania huxleyi [22], Nannochloropsis residue [23], Plocamium, Sargassum [24], Spirulina [43], Synechococcus, Tetraselmis [25] and cultivated mixed consortia [26]. Hu et al. [27] carried out his research on pyrolysis of blue-green algae blooms (BGAB) in a fixed bed reactor. The maximum oil yield of 54.97% was obtained at a temperature of 500°C, particle size lower than 0.25 mm, and nitrogen flow rate equal to 100 mL/ min. Researchers suggested that pyrolysis of algal biomass is a promising process for production of renewable fuel. Table  11.4 shows the literature data on pyrolysis-derived bio-oil from algae. Microwave-assisted pyrolysis has drawn attention because of advantages over conventional methods including fast and uniform heating. It reduces residence time and accelerates chemical reactions, which leads to energy saving [34]. Hu et al. [35] carried out the microwave-assisted pyrolysis of Chlorella vulgaris under different microwave powerlevels, different catalysts and amount of activated carbon and solid residue. They found that by increasing microwave power level, maximum temperature rising rate and pyrolysis temperature will increase. In their study, maximum bio-oil yield (35.83 wt %) was obtained under the microwave power of 1500W. It was shown that catalysts can enhance the pyrolysis of algal biomass. b. Hydrothermal liquefaction (HTL) Hydrothermal liquefaction (HTL) is defined as the reaction of biomass in water at elevated temperature (200–400°C) and high pressure (5–20 M Pa) with or without using a catalyst [14]. Hydrothermal liquefaction is also called direct liquefaction, hydrothermal upgrading, depolymerization, and solvolysis. Catalyst is generally used to enhance biooil yield. Both homogeneous and heterogeneous catalysts have been used in hydrothermal liquefaction process of algae these includes KOH, CH3COOH, Na2CO3, HCOOH, H2SO4, Ca3(PO4)2, NiO and zeolite [36]. Water is the most common solvent as it is abundant and economically

Maximum temperature of 700°C, 10°C/min

Slow pyrolysis

Slow pyrolysis

Fast pyrolysis

Fast pyrolysis

Fast pyrolysis

Microwave-assisted pyrolysis

Nannochloropsis sp. residue

Tetraselmis chui, Chlorella like, Chlorella vulgaris, Chaetocerous muelleri, Dunaliella tertiolecta, Synechococcus

Chlorella protothecoides

Chlorella vulgaris remnants

Microcystis aeruginosa, Chllorella protothecoides

Macro-algae (seaweed)

‘-’ Not reported

300–500°C, 10°C/min

Microwave-assisted pyrolysis

Chlorella sp.

200–300W

500°C, 600°C/s

500°C

400–600°C, 600°C/s

500–1250W, 462– 627°C, 20 min

300–700°C, 15 min

Slow pyrolysis

Blue-green algae blooms (BGAB)

Conditions

Process

Species

Table 11.4 Review on pyrolysis-derived bio-oil from algae.

Maximum bio-oil yield (21%)

M. aeruginosa (24%), C. protothecoides (18%)

53%

Maximum bio-oil yield (57.9%) (450°C)

Maximum liquid percentage (43%) (Tetraselmis chui, 500°C)

Maximum bio-oil yield (31.1%) (400°C)

Maximum bio-oil yield (28.6%) (750 W)

Maximum bio-oil yield (54.9%)

Yield/conversion

22.1

29

24.5

41

-

32.7

30.7

-

HHV (MJ/kg)

[33]

[32]

[31]

[30]

[29]

[23]

[28]

[27]

Ref.

Bio-Oil Production from Algal Feedstock 359

360

Liquid Biofuel Production available, although it gives a very viscous bio-oil with high oxygen content. Production of bio-oil by hydrothermal liquefaction process, there are several algal species that have been investigated including Chlorella, Botryococcus braunii, Dunaliella, Enteromorpha, Nanno- chloropsis [37], Scenedesmus, and Spirulina [38]. The primary product of hydrothermal liquefaction is bio-oil or bio-crude, and the major by-products are the bio-char, solid residue and water soluble organic compounds. The use of organic solvent instead of water has improved bio-oil quality and yield. Both pyrolysis and hydrothermal liquefaction (HTL) processes belong to the thermo-chemical technologies in which feedstock organic compounds are converted into bio-oil products. In the liquefaction, biomass is decomposed into small and unstable molecules which can be repolymerized into oily compounds [39]. Reactions such as solvolysis, depolymerization, decarboxylation, hydrogenolysis, and hydrogenation are involved during the liquefaction process. An advantage of the thermochemical process is that it is relatively simple, usually requiring only one reactor, thus having a low capital cost. Hydrothermal liquefaction is usually performed at lower temperatures (300–400°C), longer residence times (0.2– 1.0h), and relatively high operating pressure (5–20Mpa) [14]. Hydrothermal liquefaction derived bio-oil is water insoluble and has a lower oxygen content and as a result higher energy content than pyrolysis-derived oils, although the HTL bio-oils have a higher viscosity than pyrolysis biooils. Jena and Das [40] compared HTL and slow pyrolysis processes for bio-oil production from algae. They found that hydrothermal liquefaction resulted in higher bio-oil yields, lower char yields, and lower energy consumption ratio compared to pyrolysis. Furthermore, hydrothermal liquefaction was able to produce bio-oil with higher energy density and superior fuel properties such as thermal and storage stabilities compared to pyrolysis bio-oil. Hydrothermal liquefaction is at an early developmental stage, and more research is needed to understand the reaction mechanisms and kinetics [14]. Table 11.5 summarizes the various parameters for liquefaction-derived bio-oil production from algae.

Temp.(°C)

300

220–320

300

200–500

250–340

300

200–300

Species

Spirulina

Enteromorpha prolifera

Scenedesmus

Nannochloropsis

Dunaliella tertiolecta

Botryucoccus brclunii

Chlorella pyrenoidosa

8.9–10.3

-

-

-

10–12

-

10–12

Pressure (MPa)

30–120

-

5–60

60

30

30

30

Time (min)

Table 11.5 Various parameters of liquefaction-derived bio-oil from algae.

24–39.4

64

30.9–43.8

Maximum 43

24–45

Maximum 23

32.6

Oil yield (wt%)

21.3–38.5

49

33.3–37.8

Maximum 39

35–37

28–30

32–34.7

HHV (MJ/kg)

[47]

[46]

[45]

[44]

[43]

[42]

[41]

Reference

Bio-Oil Production from Algal Feedstock 361

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Liquid Biofuel Production

11.3 Properties of Bio-Oils Bio-oils are commonly dark brown, free-flowing liquids having a characteristic smoky odor. The various physical properties of bio-oils result from the chemical composition of the oils, which is considerably dissimilar from that of petroleum-derived oils. Bio-oil is a complex mixture of several hundred organic compounds, generally containing aldehydes, alcohols, acids, esters, phenols, ketones and lignin-derived oligomers. Some of these compounds are directly associated to the unwanted properties of bio-oil. The liquefied oils have much lower oxygen and moisture contents, and consequently much higher energy value, as compared to oils from fast pyrolysis. The properties of bio-oil from both processes are significantly different from heavy petroleum fuel oil. Compared with heavy petroleum fuel oil, the bio-oils have the following undesired properties for fuel applications: (1) (2) (3) (4) (5)

High corrosiveness (acidity) High oxygen content (low heating value) High viscosity High water content High ash content

These undesired properties have certainly limited the range of bio-oil application. The differences in processing conditions and feedstock result in significant differences in the product yield and product composition of bio-oils.

11.4 Uses of Bio-Oils Bio-oil can be easily stored and transported as a renewable liquid fuel. It is an alternative source for fuel oil or diesel in various static applications including engines, furnaces, boilers and turbines for electricity generation. The crude oil could serve as a raw material for the production of wood flavors, adhesives, phenol-formaldehyde-type resins, etc. Different compounds forming the bio-oils are also possible after further processing and separation. Some industrial uses of bio-oil are given below: 1. Bio-oil can be used as a transportation fuel after upgrading. 2. Combustion fuel in furnace/burner/boiler systems for heat generation.

Bio-Oil Production from Algal Feedstock 363 3. Combustion in turbines/diesel engines for power generation. 4. It can be used as liquid smoke and wood flavors. 5. Production of anhydro-sugars like levoglucosan, which has potential for the manufacturing of pharmaceuticals, surfactants, biodegradable polymers. 6. Bio-oil can be used in making adhesives, e.g., asphalt bio-binder. 7. Chemicals and resins can be produce from bio-oil (e.g., fertilizers, acids, agri-chemicals and emission control agents).

11.5 Up-Gradation of Bio-Oil to Biodiesel along with Recent Developments Bio-oils are a dark brown liquid, which is produced from thermochemical (fast pyrolysis/hydrothermal liquefaction) process; it cannot be used as transportation fuels directly. Bio-oil has several undesired properties such as high corrosiveness, high viscosity, low heating value, high oxygen content (35-40 wt % dry basis) and water content (15-25 wt %) that limit its direct application as liquid fuel. Thus, bio-oil required upgrading to improve its quality to be used as liquid fuel or chemical feedstock for bio-refinery to extract valuable chemicals. There are various procedures for bio-oil the upgrading process including physical and chemical methods given below:

11.5.1

Esterification/Alcoholysis

Esterification of free fatty acids to corresponding alkyl ester, also called alcoholysis, is one of the reactions used for biodiesel production. In this reaction, fatty acids react with alcohol at atmospheric pressure in the presence of an acid catalyst to form alkyl ester or biodiesel. Methanol is the most common alcohol used for esterification since it is the least expensive alcohol. The temperature of the reaction is lower than the boiling point of alcohol. If methanol is used, esterification is usually conducted at 60°C. Esterification can also be performed in supercritical conditions. Peng et al. [48] has compared catalytic upgrading of bio-oil in sub and supercritical ethanol and showed supercritical upgrading was performed more effectively than sub-critical upgrading process. Water is the by-product of this reaction. Since the esterification reaction is reversible, it can be conducted by reactive distillation to separate and remove water during reaction to increase reaction yield. Both heterogeneous and homogeneous catalysts can be used for esterification; however, heterogeneous catalysts have been

364

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preferred to apply in bio-oil upgrading because separation of catalyst from products is easier. Aluminum silicate [49] and HZSM-5 [48] are among the catalysts used in bio-oil upgrading. Bio-oil contains an abundant amount of fatty acids, hence esterification has been investigated by researchers as a technique for bio-oil upgrading. Table 11.6 shows the comparison of some common alcohols used in transesterification and Table 11.7 shows the comparison of two of the most common alkaline catalysts used in transesterification. Table 11.6 Advantages and disadvantages of different types of alcohols used in esterification of algal oil [50]. Alcohol

Advantages

Disadvantages

Methanol

- Less expensive. - More efficient reactant. - EPA study showed that rats can consume FAMEs with no adverse effects

- Toxic causing nerve deterioration due to prolonged exposure. - More poisonous than ethanol

Bioethanol

Environmentally safe (green chemically)

- Higher viscosity of biodiesel product. - Large scale use will require cellulose based technology

Ethanol

- Environmentally safe. - Preferred alcohol to use for cold weather operations.

- More expensive. - Heavier for biodiesel - Transesterification reaction is less forgiving.

Table 11.7 Advantages and disadvantages of NaOH & KOH catalyst in esterification [50]. Catalyst

Advantages

Disadvantages

NaOH

- Less expensive. - Useful for oil titration to check Free Fatty Acids (FFA).

- Hard to use - Does not provide a valuable by-product

KOH

- Easier to use - Does a better catalytic job than NaOH - Provides potash fertilizer as a byproduct

- Use 40% more by mass than NaOH

Bio-Oil Production from Algal Feedstock 365

11.5.2

Solvent Addition

Polar solvents such as ethyl acetate, acetone, methanol, ethanol, and furfural have been used for many years to homogenize and to reduce viscosity of biomass oils. The immediate effects of adding these polar solvents are decreased viscosity and increased heating value. The increase in heating value for bio-oils mixed with solvents occurs because the solvent has a higher heating value than that of most bio-oils. The solvent addition reduces the oil viscosity due to the following mechanisms: 1. Physical dilution without affecting the chemical reaction rates. 2. Chemical reactions between the solvent and the oil components that prevent further chain growth. 3. Reducing the reaction rate by molecular dilution or by changing the oil microstructure. Mostly the studies have directly added solvents after pyrolysis, which works well to decrease the viscosity and increase stability and heating value. Several new studies showed that reacting the oil with alcohol (e.g., ethanol) and acid catalysts (e.g., acetic acid) at mild conditions by using reactive distillation, resulted in a better bio-oil quality.

11.5.3

Emulsification

Emulsification is one of the important methods for bio-oil upgrading with other fuels. Pyrolysis oils are not miscible with hydrocarbon fuels, but with the help of surfactants they can be emulsified with diesel oil. The emulsification process is costly due to surfactant addition and high energy cost for high production of emulsions. Upgrading of bio-oil through emulsification with diesel oil has been investigated by many researchers. A process for producing stable microemulsions, with 5–30% of bio-oil in diesel, has been developed at Canmet Energy Technology Centre. Those emulsions are less corrosive and show promising ignition characteristics. Generally upgrading of bio-oil through emulsification with diesel oil is quite simple. It provides a short-term approach to the use of bio-oil in diesel engines. The emulsions showed promising ignition characteristics, but fuel properties such as heating value, cetane and corrosivity were still unsatisfied. Furthermore, this process required high energy for production. Design, production and testing of injectors and fuel pumps made from stainless steel or other materials are required.

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11.5.4

Hydrotreating/Hydro Deoxygenation

Hydrotreating is an established refinery process to reduce N, O and S atoms from oil cuts or petrochemical feedstocks. In this process, oxygen is removed as water by catalytic reaction with high pressure hydrogen. Nitrogen and sulfur are also removed as H2S and NH3 [14]. The process is typically conducted at high pressure (up to 20MPa) and moderate temperatures (300–450°C), and requires a hydrogen source. The most commonly used catalysts are NiMo-based and CoMo-based catalysts which are industrial hydrotreating catalysts for removal of nitrogen, sulfur and oxygen. Hydrogenation is another reaction that can be carried out during hydrotreating; examples of reactions (1.5) involved in hydrotreating of biooil are given below:

R −OH + H2

Hydrodeoxygenation:

R2

R1 Hydrogenation:

C H

+ H2

C H

R —H + H2O CH2 R2 + H2O R1 CH2

It is commonly supposed that the higher the hydrogen content of a fuel product, the better its quality. Thus, hydrogenation can enhance bio-oil quality during hydrotreating process. Different catalysts have been tested in hydrotreating of bio-oil including Ni–Cu/SiO2, CoMo/γ-Al2O3, Ru/C, Pd/C, Ni–Cu/Al2O3, NiMo/Al2O3, Ru/TiO2 and Ru/Al2O3.

11.5.5

Hydro-Cracking

Hydro-cracking is a catalytic cracking process supported by the presence of hydrogen gas. The process takes place at temperatures more than 350°C and relatively high pressure (up to 2000 psi) in the presence of a suitable catalyst. In hydro-cracking, hydrogenation accompanies cracking of complex organic molecules to simpler molecules. The products of the catalytic cracking process comprise coke, an aqueous phase, an organic liquid phase, and gasses. Following reaction 1.6 shows an example of hydro-cracking reactions:

Hydro-cracking:

CH2 R2 + H 2 CH2 R1

R1—CH3 + H3C—R2

Bio-Oil Production from Algal Feedstock 367 In hydro-cracking, hydrogen is used to break C–C bonds while, in hydrotreatment, H2 is utilized to break C–N, C–O, and C–S bonds. The combination of hydrotreating and hydro-cracking for bio-oil upgrading has been also investigated. In this process, bio-oil is first upgraded by hydrotreating. Subsequently, heavy components are separated from the upgraded bio-oil and referred to hydro-cracking process to break in to light components. Hydro-cracking is an effective way to break heavy molecules to light products, but it requires high temperature and high hydrogen pressure which increases process cost.

11.5.6

Zeolite Cracking

Zeolites are micro-porous alumino-silicate materials commonly used as catalysts and adsorbents. Zeolites have very high surface areas and adsorption capacity. Zeolites contain active sites, usually acid sites. Zeolite cracking is a technique for bio-oil upgrading. This process is performed in a temperature range of 350–500°C. In this process, deoxygenation accompanies cracking, and products include gases (light alkanes, CO, CO2), water-soluble organics, water, oil-soluble organics and coke. Hydrogen is not used in zeolite cracking; therefore the process is conducted at atmospheric pressure. Mortensen et al. [51] have compared zeolite cracking and hydro-deoxygenation and have come to the conclusion that zeolite cracking is not a promising route for bio-oil upgrading because of the low quality of upgraded bio-oil.

11.6 Conclusion • Bio-oils are a promising renewable energy source which have received extensive recognition around the world for their characteristics as combustion fuels used in boiler, engines or gas turbines and resources in chemical industries. But compared with heavy petroleum fuel oil, it has several undesired properties for fuel applications such as high oxygen and water contents, high viscosity, and corrosiveness. These undesired properties require upgrading of bio-oil before using it as transportation fuel. • At present, two thermochemical conversion techniques are being used for bio-oil production from algae, i.e., pyrolysis and hydrothermal liquefaction. • Pyrolysis is already commercialized, but it requires drying of feedstock which increases the energy consumption of the whole process.

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References 1. Hazelwood, I., Sipila, K., Korhonen, M., Power production from biomass III—Gasification and pyrolysis R&D&D for industry. VTT Symposium 192, Espoo, Finland, 14–15 September, p. 642, 1998. 2. Ma, Z., Custodis, V., Bokhoven, J.A., Selective deoxygenation of lignin during catalytic fast pyrolysis. Catal. Sci. Technol., 4, 766–772, 2014. 3. Mohammed, S., Bakhtiyor N., Kunio Y., A review of production and upgrading of algal bio-oil. Renewable Sustainable Energy Rev., 58, 918–930, 2016. 4. Bahadar, A. and Khan, M.B., Progress in energy from microalgae: A review. Renewable Sustainable Energy Rev., 27, 128–148, 2013. 5. Demirbas, A., Use of algae as biofuel sources. Energy Convers. Manage., 51, 2738–2749, 2010. 6. Sheehan, J.J., Terri, D., Benemann, J., Roessler, P., A look back at the U.S. department of energy’s aquatic species program-biodiesel from algae. National Renewable Energy Laboratory, 1998, NREL/TP: 580-24190. U.S. Department of Energy, 1617 Cole Boulevard. 7. Rodolfi, L., Zittelli, C.G., Bassi, N., Padovani, G., Biondi, N., Biondi, G., Tredici, M.R., Microalgae for oil: Strain selection, induction of lipid synthesis and outdoor mass cultivation in a low-cost photobioreactor. Biotechnol. Bioeng., 102, 100–112, 2009. 8. Meng, X., Yang, J., Xu, X., Zhang, L., Nie, Q., Xian, M., Biodiesel production from oleaginous microorganism. Renewable Energy, 34, 1–5, 2009. 9. Chisti, Y., Biodiesel from microalgae beat Bioethanol. Trends Biotechnol., 126, 1–6, 2008. 10. Chisti, Y., Biodiesel from microalgae. Biotechnol. Adv., 25, 294–306, 2007.

Bio-Oil Production from Algal Feedstock 369 11. Rao, S., Algal Cultures Analogues of Blooms and Applications, vol. 1, Science Publishers, Enfield, New Hampshire, 2006. 12. Kurano, N. and Miyachi, S., Microalgal studies for the 21st century. Hydrobiologia, 512, 27–32, 2004. 13. Khan, S.A., Rashmi, Hussain, M.Z., Banerjee, U.C., Prospective of biodiesel production from microalgae in India. Renewable Sustainable Energy Rev., 13, 2361–2372, 2009. 14. Xiu, S.N. and Shahbazi, A., Bio-oil production and upgrading research: A review. Renewable Sustainable Energy Rev., 16, 4406–4414, 2012. 15. Marcilla, A., Catala, L., Garcia-Quesada, J.C., Valdes, F.J., Hernandez, M.R., A review of thermochemical conversion of microalgae. Renewable Sustainable Energy Rev., 27, 11–19, 2013. 16. Brennan, L. and Owende, P., Biofuels from microalgae A review of technologies for production, processing, and extractions of biofuels and co-products. Renewable Sustainable Energy Rev., 14, 557–577, 2010. 17. Duan, P.G. and Savage, P.E., Upgrading of crude algal bio-oil in supercritical water. Bioresour. Technol., 102, 1899–906, 2011a. 18. Peterson, A.A., Vogel, F., Lachance, R.P., Froling, M., Antal, M.J., Tester, J.W., Thermochemical biofuel production ninhydrothermal media: A review of sub and supercritical water technologies. Energy Environ. Sci., 1, 32–65, 2008. 19. Huber, G.W., Iborra, S., Corma, A., Synthesis of transportation fuels from biomass: Chemistry, catalysts, and engineering. Chem. Rev., 106, 4044–4098, 2006. 20. Demirbas, A., Oily products from mosses and algae via pyrolysis. Energy Sources Part A, 28, 933–940, 2006. 21. Babich, I.V., Vander Hulst, M., Lefferts, L., Moulijn, J.A., O’ Connor, P., Seshan, K., Catalytic pyrolysis of microalgae to high quality liquid biofuels. Biomass Bioenergy, 35, 3199–3207, 2011. 22. Wu, Q.Y., Dai, J.B., Shiraiwa, Y., Sheng, G.Y., Fu, J.M., A renewable energy source hydrocarbon gases resulting from pyrolysis of the marine nanoplanktonic alga Emilianiahuxleyi. J. Appl. Phycol., 11, 137–142, 1999. 23. Pan, P., Hu, C.W., Yang, W.Y., Li, Y.S., Dong, L.L., Zhu, L.F. et al., The direct pyrolysis and catalytic pyrolysis of Nannochloropsis sp. residue for renewable bio-oils. Bioresour. Technol., 101, 4593–4599, 2010. 24. Li, D.M., Chen, L.M., Zhang, X.W., Ye, N.H., Xing, F.G., Pyrolytic characteristics and kinetic studies of three kinds of red algae. Biomass Bioenergy, 35, 1765–1772, 2011. 25. Grierson, S., Strezov, V., Shah, P., Properties of oil and char derived from slow pyrolysis of Tetraselmischui. Bioresour. Technol., 102, 8232–8240, 2011. 26. Sarkar, O., Agarwal, M., Kumar, A.N., Mohan, S.V., Retrofitting hetrotrophically cultivated algae biomass as pyrolytic feedstock for biogas, bio-char and bio-oil production encompassing biorefinery. Bioresour. Technol., 178, 132–138, 2015.

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27. Hu, Z.Q., Zheng, Y., Yan, F., Xiao, B., Liu, S.M., Bio-oil production through pyrolysis of blue-green algae blooms (BGAB): Product distribution and biooil characterization. Energy, 52, 119–125, 2013. 28. Du, Z.Y., Li, Y.C., Wang, X.Q., Wan, Y.Q., Chen, Q., Wang, C.G. et al., Microwave-assisted pyrolysis of microalgae for biofuel production. Bioresour. Technol., 102, 4890–4896, 2011. 29. Grierson, S., Strezov, V., Ellem, G., Mcgregor, R., Herbertson, J., Thermal characterisation of microalgae under slow pyrolysis conditions. J. Anal. Appl. Pyrol., 85, 118–123, 2009. 30. Miao, X.L. and Wu, Q.Y., High yield bio-oil production from fast pyrolysis by metabolic controlling of Chlorella protothecoides. J. Biotechnol., 110, 85–93, 2004. 31. Wang, K.G., Brown, R.C., Homsy, S., Martinez, L., Sidhu, S.S., Fast pyrolysis of microalgae remnants in a fluidized bed reactor for bio-oil and biochar production. Bioresour. Technol., 127, 494–499, 2013. 32. Miao, X.L., Wu, Q.Y., Yang, C.Y., Fast pyrolysis of microalgae to produce renewable fuels. J. Anal. Appl. Pyrol., 71, 855–863, 2004. 33. Budarin, V.L., Zhao, Y.Z., Gronnow, M.J., Shuttleworth, P.S., Breeden, S.W., Macquarrie, D.J. et al., Microwave-mediated pyrolysis of macro-algae. Green Chem., 13, 2330–2333, 2011. 34. Motasemi, F. and Afzal, M.T., A review on the microwave-assisted pyrolysis technique. Renewable Sustainable Energy Rev., 28, 317–330, 2013. 35. Hu, Z.F., Ma, X.Q., Chen, C.X., A study on experimental characteristic of microwave-assisted pyrolysis of microalgae. Bioresour. Technol., 107, 487–493, 2012. 36. Duan, P.G. and Savage, P.E., Hydrothermal liquefaction of a microalga with heterogeneous catalysts. Ind. Eng. Chem. Res., 50, 52–61, 2011b. 37. Biller, P. and Ross, A.B., Potential yields and properties of oil from the hydrothermal liquefaction of microalgae with different biochemical content. Bioresour. Technol., 102, 215–225, 2011. 38. Valdez, P.J., Tocco, V.J., Savage, P.E., A general kinetic model for the hydrothermal liquefaction of microalgae. Bioresour. Technol., 163, 123–127, 2014. 39. Demirbas, A., Liquefaction of biomass using glycerol. Energy Sources Part A, 30, 1120–1126, 2008. 40. Jena, U. and Das, K.C., Comparative evaluation of thermochemical liquefaction and pyrolysis for bio-oil production from microalgae. Energy Fuel, 25, 5472–5482, 2011. 41. Vardon, D.R., Sharma, B.K., Scott, J., Yu, G., Wang, Z.C., Schideman, L. et al., Chemical properties of biocrude oil from the hydrothermal liquefaction of Spirulina algae, swine manure, and digested anaerobic sludge. Bioresour. Technol., 102, 8295–8303, 2011. 42. Zhou, D., Zhang, L.A., Zhang, S.C., Fu, H.B., Chen, J.M., Hydrothermal liquefaction of macroalgae enteromorpha prolifera to bio-oil. Energy Fuel, 24, 4054–4061, 2010.

Bio-Oil Production from Algal Feedstock 371 43. Vardon, D.R., Sharma, B.K., Blazina, G.V., Rajagopalan, K., Strathmann, T.J., Thermochemical conversion of raw and defatted algal biomass via hydrothermal liquefaction and slow pyrolysis. Bioresour. Technol., 109, 178–187, 2012. 44. Brown, T.M., Duan, P.G., Savage, P.E., Hydrothermal liquefaction and gasification of Nannochloropsis sp. Energy Fuel, 24, 3639–3646, 2010. 45. Minowa, T., Yokoyama, S., Kishimoto, M., Okakura, T., Oil production from algal cells of dunaliella-tertiolecta by direct thermochemical liquefaction. Fuel, 74, 1735–1738, 1995. 46. Sawayama, S., Inoue, S., Dote, Y., Yokoyama, S.Y., Co2 fixation and oil production through microalga. Energy Convers. Manage., 36, 729–731, 1995. 47. Yang, W.C., Li, X.G., Liu, S.S., Feng, L.J., Direct hydrothermal liquefaction of undried macroalgae Enteromorpha prolifera using acid catalysts. Energy Convers. Manage., 87, 938–945, 2014. 48. Peng, J., Chen, P., Lou, H., Zheng, X.M., Catalytic upgrading of bio-oil by HZSM-5 in sub- and super-critical ethanol. Bioresour. Technol., 100, 3415– 3418, 2009. 49. Zhang, Q., Chang, J., Wang, T.J., Xu, Y., Upgrading bio-oil over different solid catalysts. Energy Fuel, 20, 2717–2720, 2006. 50. Briggs, M., Pearson, J., Farag, I.H., Biodiesel processing, in: Incorporating lessons on biodiesel into the science classroom, NH Science Teacher Association (NHSTA) Annual Conference, 2004, Philips Exeter Academy. Exeter, NH, http://www.unh.edu/p2/biodiesel. 51. Mortensen, P.M., Grunwaldt, J.D., Jensen, P.A., Knudsen, K.G., Jensen, A.D., A review of catalytic upgrading of bio-oil to engine fuels. Appl. Catal. Gen., 407, 1–19, 2011.

12 Effect of Upgrading Techniques on Fuel Properties and Composition of Bio-Oil Krushna Prasad Shadangi1 and Kaustubha Mohanty2* 1

Department of Chemical Engineering, V. S. S. University of Technology, Burla, India 2 Department of Chemical Engineering, Indian Institute of Technology Guwahati, India

Abstract Pyrolysis is one of the thermochemical methods of conversion which convert biomass into solid, liquid and gaseous fuel. The yield of liquid is comparably more than others. The yield of pyrolytic liquid varies with the feedstock types. Literature revealed that non-edible oil seeds, seed press cakes produced high yield of biooil compared to other biomasses. The low crystallinity index of seeds and seed press cake favours for low temperature degradation. Due to the presence of high extractive content, the biomass resulted in high yield of bio-oil containing more organic liquid with less aqueous liquid. The organic liquid is the oil (bio-oil) which can be used as fuel. However, the bio-oil has various disadvantages such as high viscosity, high pH, high oxygen content and low calorific value. Due to these drawbacks, it cannot be used directly as it is. Hence, upgrading of bio-oil is necessary to improve these fuel properties. Catalytic upgrading, hydrogenation and chemical upgrading are such techniques which can reduce the oxygen content and enhance the fuel quality and stability of the bio-oil. It has been observed that co-pyrolysis of biomass with waste polystyrene helped in increasing the fuel properties. Polystyrene is a common material used for various purposes to meet human demand and is non-biodegradable. Proper waste management is nowadays a big issue. If such waste materials can be used to enhance the quality of bio-oil then waste can be used to produce valuable product. Bio-oil/pyrolysis oil can be used as substitutes for conventional fuels in a wider range. Pyrolysis is considered to be one of the sustainable solutions which may be economically profitable on a large scale and minimize environmental concerns especially in terms of waste minimization. *Corresponding author: [email protected] Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (373–386) © 2019 Scrivener Publishing LLC

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Keywords: Pyrolysis, hydrodeoxygenation, hydrogenation, steam reforming, emulsification, esterification

12.1 Introduction The oil produced from pyrolysis of biomass is called bio-oil. It is a browncoloured liquid. The bio-oil can be used as a future fuel or for the production of bio-chemicals. The pyrolysis of biomass produces bio-oil, non-condensable gas and char. However, the yield of pyrolytic oil is more for fast pyrolysis compared to slow pyrolysis. The bio-oil is carbon neutral fuel since it is a product of bio-mass and the CO2 thus generated is again consumed by the plant itself during its photosynthesis. Biomass has negligible sulphur content and hence the possibility of formation of SO2 is also less from the combustion of bio-oil. The bio-chemicals can be extracted using various techniques. Distillation and liquid-liquid extraction are such methods which can be used to extract the valuable chemicals. However, such techniques are the end operations of any chemical process. The complex composition of bio-oil is the major disadvantage which not only decreases the stability of the oil but also changes the chemical composition. There are various other chemical properties which are very important if the bio-oil is considered to be used as fuel. The fuel properties differ with the chemical composition of the bio-oil. The variation in the composition with the storage of bio-oil changes the fuel properties and consequently reduced the usability as direct fuel. There are various upgrading techniques which may be followed to enhance the storage stability and the fuel properties of the bio-oil. The upgraded bio-oil can be a better alternative liquid fuel for future. With the shrinking supply of fossil fuels and increasing environmental pollution, biomass is one of the resources that can be used for fuel production. Bio-oil has the potential to protect the environment from pollution in the universe compared to fossil fuel. Hence, various upgrading techniques can be followed to enhance the fuel properties. Such upgrading techniques include in-situ catalytic pyrolysis, catalytic vapour cracking, hydrogenation, steam reforming, esterification, co-processing and emulsification. However, every technique has some advantages and disadvantages. This study provides a brief overview of the fuel properties and composition of biomass pyrolysis oil/bio-oil and various upgrading technologies to enhance the fuel properties and storage ability.

Effect of Upgrading Techniques on Bio-Oil 375

12.2 Bio-Oil and its Properties Bio-oil or pyrolytic oil has the potential to be used as a fuel for the future and be a replacement of fossil fuels such as petrol, diesel and kerosene, etc. It is not a product of thermodynamic equilibrium during pyrolysis but rather is produced with a short reaction time and rapid cooling or quenching due to the pyrolysis temperatures. It is a complex mixture of various chemical compounds and initiated from the degradation of cellulose, hemicellulose, lignin and the extractives [1]. The composition of bio-oil also varies with the pyrolysis conditions, such as temperature, pressure, types of reactor and inert gas flow rate. All the factors have an effect on the composition and the yield of pyrolytic oil. As a whole, bio-oil is a complex mixture of phenol, cresol, furans, acids, alkanes, alkenes, nitriles, esters, nitrogenated compounds, oxygenated compounds, benzene and other aromatic compounds. It is a mixture of a wide range of molecular weight chemical compounds. Apart from all these chemical compounds, bio-oil also contains water and many other water soluble chemicals. Probably due to this, bio-oil does not have good storage stability. The thermodynamic equilibrium between the chemical compositions of the bio-oil changes during storage. The bio-oil is separated into two phases such as organic phase and aqueous phase. The organic phase, also called the oil phase, can be used as a fuel. The complex nature of bio-oil affects the fuel properties. The fuel properties such as viscosity, density, water content, calorific value, pH and pour point are very important which differ with the biomass types and the pyrolysis conditions. The fuel properties of the bio-oil continuously vary with the aging time due to the chemical reactions between the chemical compounds. Moreover, bio-oil is highly viscous, acidic, less calorific value, and low pour point [2, 3]. After separation of the aqueous phase pyrolytic oil, the oil phase also contains some water in emulsion form which is very difficult to separate by simple gravity separation process. The presence of water and the oxygenated compounds decreases the calorific value of the pyrolytic oil. Bio-oil is more acidic which reduces the storage stability and the usability due to the presence of various acids [4–6]. The bio-oil is a mixture of saturated and unsaturated hydrocarbons that decides the low temperature flow ability. The presence of unsaturated hydrocarbons decreases the flow ability [2, 3]. Hence, upgrading of bio-oil is most crucial before using it as a fuel.

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12.3 Upgrading of Bio-Oil Upgrading of bio-oil techniques proceed to reduce the drawbacks of bio-oil by improving the physico-chemical properties. The upgrading of bio-oil, which increases the storage stability, is very important. The complexity of bio-oil improves after upgrading by reducing the oxygen and acid content. The oxygenated compounds break and oxygen is removed as water. Apart from this, the concentration of specific compounds increases. Such effects in the composition of bio-oil vary with the techniques followed to upgrade. Various upgrading techniques can be followed during the production of bio-oil and also after the production of bio-oil. Catalytic pyrolysis, catalytic vapour cracking and co-pyrolysis process can be performed during the thermal degradation of biomass. However, hydrogenation, steamreforming, molecular distillation, supercritical fluid extraction, esterification and emulsification methods are used after the bio-oil is produced.

12.3.1

Catalytic Pyrolysis

Catalytic pyrolysis is the process where catalyst is mixed with the biomass and the generated vapours are condensed and collected as bio-oil. This process is also known as in-situ catalytic pyrolysis process. The evidence of positive effect of CaO, Kaolin and Al2O3 catalysts in the fuel properties of the pyrolytic oil is reported elsewhere [7, 8]. It was reported that catalyst at higher ratio enhanced the pour point, calorific value, reduced the viscosity of the pyrolytic oil. The yield of aqueous pyrolytic liquid was more in catalytic pyrolysis compared to non-catalytic pyrolysis process. The variation in fuel properties and composition of pyrolytic oil with the biomass to catalyst ratio was noticed. The catalytic pyrolysis has less effect in the yield of pyrolytic liquid compared to direct thermal process. The yield of aromatic hydrocarbons in the catalytic pyrolysis was also varying with catalyst ratio and types and the feed composition. Catalytic pyrolysis was carried out using fixed bed reactor [9] or fluidized bed [10] under nitrogen flow using various catalyst such as HZSM-5 [10, 9], CaO [11] ZSM-5, Al-SBA-15, alumina [12] and Cu [13] and all these studies have reported their effects on the yield and composition of bio-oil. The types of catalyst can be selected by considering various aspects such as high deoxygenating activities [12, 14] or preferred selectivity [15], influence of temperature and catalyst-to-biomass ratio on product yields [10], etc. The investigation indicated that catalytic pyrolysis lowered the oxygen content of the bio-oils and boosted the calorific values compared to the pyrolysis process without catalyst [10, 9, 12]. Similar conclusions were also observed

Effect of Upgrading Techniques on Bio-Oil 377 from various experiments with different biomass, comprising green corncob [10], microalgae [9], herb residue [12] and waste woody biomass [13]. It was noticed that catalytic pyrolysis led to higher aromatic hydrocarbons content in bio-oil using HZSM-5 [9], alumina [12] or HZSM-5/γ-Al2O3 [15] as catalyst compared to non-catalytic pyrolysis. The yield of long chain carbon compounds [9] is higher in the non-catalytic pyrolytic oil. Catalytic pyrolysis with HZSM-5 zeolite catalyst leads in the formation of the coke, water and non-condensable gas yield by reducing the heavy oil fraction in the bio-oil as reported elsewhere [10]. The conversion of oxygen element in heavy oil into CO, CO2 and H2O in presence HZSM-5 zeolite catalyst could be the reason of reducing the heavy oil fraction in the bio-oil [10].

12.3.2

In-Situ versus Ex-Situ Catalytic Pyrolysis Process

The pyrolytic vapours generated from the raw materials is passed through a bed of catalyst and recracked; the process is known as catalytic vapour cracking. Catalytic vapour cracking is also known as ex-situ catalytic pyrolysis. Catalytic vapour cracking provided better efficiency in terms of yield and conversion with a less amount of catalyst in comparison with in-situ catalytic process [16]. Both processes enhanced the physical properties and reduced the oxygen content of the pyrolytic oil. However, in the in-situ pyrolysis process the possibility of catalytic poisoning is more. Also, the contact between the feed and the catalyst is comparatively less than the ex-situ catalytic pyrolysis process since the catalyst mixed with the feedstock do not provide intimate contact between the generated vapour and catalyst. Due to this the amount of catalyst required for better conversion and efficiency is much higher. Literature revealed that with increase in the catalyst ratio, the fuel properties such as viscosity decreased [4–8]. However, higher catalyst ratio will eventually increase the cost of production of bio-oil. In ex-situ process, proper contact between the pyrolysis vapour and the catalytic surface is achieved. Hence, the amount of catalyst required is less for the better catalytic conversion in comparison with in-situ process. One of the disadvantages of this process is that some of the catalyst may be entrained with the pyrolytic vapour if the catalyst has less particle size. To overcome this problem tablet of catalyst can be used. Literature also revealed that the use of catalyst has increased the calorific value and decreased the viscosity of the pyrolytic oil significantly. Catalytic pyrolysis raised the basicity of the pyrolytic oil compared to thermal pyrolysis. The increase in pH may be due to the conversion of acidic compounds into esters and hydrocarbons by catalytic decarboxylation reaction. The reduction of viscosity may be due to the presence of water in the pyrolytic oil [4–8]. The reduction in the viscosity of

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pyrolytic oil may be due to the formation of lighter hydrocarbons and also increases the calorific value. Catalytic pyrolysis reduces the oxygen content in the pyrolytic oil and converts the acids to esters which eventually results in higher calorific value. The formation of lighter hydrocarbons during catalytic pyrolysis decreases the distillation temperature of the bio-oil. Literature revealed that distillation temperature of the oils was found to be decreasing with the increase in catalyst loading. Hence, it could be observed that less use of catalyst is very important. Ex-situ pyrolysis may be the best process which enhances the quality and fuel properties of pyrolytic oil.

12.3.3

Hydrodeoxygenation

The process hydrodeoxygenation is generally followed after the production of pyrolytic oil or bio-oil. In this process, hydrogenation reaction takes place in presence of catalyst under high pressure of hydrogen which removes oxygen from the oil. Various oxygenated compounds such as acids, aldehydes, esters, ketones and phenols upon hydrogenation breaks and the oxygen is released in the form of water [17]. Hydrogenation of bio-oil was performed by several researchers using catalysts such as Ni Mo, Co Mo sulphide, Pt supported on mesoporous ZSM-5, Pt/ZSM-5 and Pt/ Al2O3 [17–19]. It was reported that Pt supported on mesoporous ZSM-5 has better in hydro de-oxygenation of di-benzo furan [17]. Literature also reported that various metal catalysts performed better in the dehydrogenation reaction [17–19]. In comparison with the crystalline catalyst the amorphous catalyst exhibits better hydrodeoxygenation characteristics [20, 21]. There are several advantages of amorphous catalyst such as easy preparation, high thermal stability, high activity and low cost.

12.3.4

Hydrogenation

Pyrolytic oil or bio-oil is a complex mixture of saturated and unsaturated hydrocarbon compounds. The presence of unsaturated hydrocarbon compounds in bio-oil decreases the stability, flow ability at low temperature. Hence, hydrogenation process can be followed to increase the saturated compounds by breaking the double or triple bonds present in the unsaturated hydrocarbons. The hydrogenation reaction can be achieved in presence of catalyst such as nickel, palladium, platinum catalyst, Al-SBA-15 supported palladium bifunctional catalysts [22] and bifunctional Pd catalysts [23], Al2O3 based catalysts [24, 25] and Ru/SBA-15 catalysts [26], or in absence of catalyst at very high temperature. Hydrogenation usually constitutes the addition of pairs of hydrogen atoms to a molecule, such as an alkene.

Effect of Upgrading Techniques on Bio-Oil 379 Hydrogenation is a chemical reaction between molecular hydrogen (H2) and another compound or element in presence of catalyst. Hydrogenation of biooil is followed to convert aromatics and alkenes into saturated alkanes and cycloalkanes which are less reactive and toxic. Not only alkenes and alkynes, but also aldehydes, nitriles and imines are converted to their corresponding saturated compounds such as alcohols and amines. The unstable constituents of bio-oil turned in to esters and alcohols by catalytic hydrogenation reaction due to the effect between metal sites and acid sites over respective catalysts [27, 28]. Hydrogenated fuel oil can be stored for long periods. Hydrogenation of bio-oil decreases the amount of aldehydes, organic acids and other reactive compounds; as a result the corrosiveness and acidity of the bio-oil can be minimised [29, 30]. Hydrogenation of pyrolytic oil or biooil increases the stability, decreases the viscosity and acidity.

12.3.5

Steam Reforming

Steam reforming is one of the bio-oil upgrading techniques which simultaneously produce liquid and gaseous bio fuel such as clean hydrogen. It is the major advantage of this process. Steam reforming can be carried out using a fixed bed reactor system [31] and fluidized bed reactor system [32] at high temperature usually 800–900 °C in presence of catalyst [33–37]. Along with hydrogen, carbon monoxide is also produced during the steam reforming process. Steam reforming yields high octane number gasoline ranged hydrocarbons. The water fractions of pyrolytic oil or bio-oil are the promising feed for production of hydrogen. The yield of hydrogen can be enhanced using various metals (magnesium, lanthanum, cobalt, chromium and nickel) based catalysts [38, 39]. Magnesium and lanthanum was used by Garcia et al. as support modifiers which enhanced steam adsorption [38]. However, cobalt and chromium additives alleviated the coke formation reactions. Garcia et al. chose magnesium and lanthanum as support modifiers to enhance steam adsorption that facilitates the carbon gasification, while cobalt and chromium additives were applied to alleviate the coke formation reactions, which modified the metal sites forming alloys with nickel and possibly reducing the crystallite size [38]. Takanabe et al. reported that steam reforming of pyrolytic oil over Pt/ZrO2 catalysts completely converted the acetic acid by producing hydrogen [39].

12.3.6

Emulsification

The term emulsion signifies the mixing of two immiscible liquids. Literature revealed that pyrolytic oil is a mixture of organic and aqueous phase which

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gets separated into two phases usually under gravity [2–8]. The organic rich part (bio-oil) is miscible with diesel while aqueous part is immiscible. Since the raw bio-oil has various drawbacks it cannot be used directly, but it can be blended with diesel and can be used as fuel. The bio-oil fractions which are not miscible with diesel can be emulsified using any surfactant. Emulsion is the simple and effective way of upgrading bio-oil by improving the ignition characteristics, fuel properties such as heating value, corrosivity, cetane number up to a satisfied degree. Xu et al. reported that the lubrication ability of the emulsified fuel was better compared with the conventional diesel fuel [40]. The combustion characteristics of a blend fuel of bio-oil and diesel such as combustion components distribution, ignition delay and temperature distribution in the combustor at different proportion was studied by Wang [41]. Ikura et al. reported that emulsion of biooil reduced the cetane number if the concentration of surfactant increased [12]. Jiang et al. studied the storage stability and thermal stability of bio-oil through emulsification with biodiesel and reported that during the aging, acid numbers slightly decreased with slightly increased in the molecular weight [42]. Chiaramonti et al. prepared emulsified bio-oil with diesel by the ratios of 25, 50 and 75  wt. % and observed that the emulsions were more stable compared to the original ones [43]. It was also obtained that with increase in the quantity of bio-oil the viscosity of emulsified bio-oil increased. Maximum 0.5 to 2% of addition of emulsifier provided the emulsified bio-oil of acceptable viscosity. It was also reported that the occurrence of solid char particles in the pyrolysis bio-oil could enhance its lubrication performance [40]. Zuogang et al. studied bio-oil emulsion with diesel by power ultrasound [44]. It was reported that treating time and ultrasound power has an effect on the stability of the emulsion fuels. The emulsified fuel was stable for 35 h when produced using ultrasound power of 80 W and treating time of 3 min. Qiang et al. studied the properties of emulsified fuel such as friction-reduction, anti-wear and extreme-pressure and found the properties to be better compared to diesel [45]. However, such results were not similar with the conclusion of Xu et al. since it depends on the device used for the experiment [40]. Due to the drawbacks of pyrolysis biooil, such as low heating value, high viscosity, high corrosiveness and poor stability, upgrading of bio-oil before practical application is necessary to acquire high-grade fuel.

12.3.7

Esterification

When a carboxylic acid is treated with an alcohol and an acid catalyst, an ester is formed along with water and the process is known as esterification.

Effect of Upgrading Techniques on Bio-Oil 381 In-general, bio-oil/pyrolytic oil comprises with various acids produced by the pyrolysis of biomass. The presence of such acids decreases the storage stability, calorific value of the pyrolytic oil. Esterification of such acids can produce esters and water. The formation of water shows the elimination of oxygenation and the formation of esters increases the calorific value and storage stability, decreases the viscosity [46]. The reduction of acids in the bio-oil also decreases the possibility of corrosion and reactivity of chemical compounds during the storage. The esterification can be carried out using homogeneous and heterogeneous catalyst to improve the yield and quality of bio-oil [47]. Similar observations are also reported by various researchers [48–51]. It was reported that acid catalyst has high reactivity to convert organic acids, such as formic acid, propionic acid and acetic acid, into esters effectively [52].

12.4 Conclusion Bio-oil has a great potential to be used as an alternative fuel in combustion engine, boiler and for the extraction of various valuable biochemicals. The characteristic properties of bio-oil vary with the source biomass and their composition. Hence, the oil produced from biomass is complex in nature. The complexity of the bio-oil also varies with the process conditions. Therefore, proper control of the production parameters has to be followed to produce the required composition of bio-oil. Due to various disadvantages, the bio-oil cannot be used as a fuel and stored for longer period. Various techniques have been followed to reduce the complexity and increase the stability of the bio-oil. Some of the techniques such as catalytic vapour cracking and in-situ catalytic cracking process are useful to produce upgraded bio-oil up to some extent. Apart from these two processes, hydrogenation, hydrodeoxygenation, esterification and emulsion techniques are followed after the production of bio-oil. Many works have been done on the hydrogenation, hydrodeoxygenation, esterification of bio-oil using various catalysts to upgrade it. However, more studies are required to investigate the effect of various acid, base and metal catalyst on the yield and composition of bio-oil. More studies are required on the effect of catalyst on catalytic vapour cracking which will provide upgraded bio-oil during the production process and may reduce the catalyst poisoning and decrease the cost of upgrading. Some physical treatment of bio-oil with various alkanes and alcohols need to be studied to separate the solid char present in the bio-oil.

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Index 2-keto-acid pathway, 228, 229 4-Methylanisole, 122–126 ABE fermentation, 229, 242–245, 247, 253, 255, 258, 259, 263, 266–269 Ablative pyrolysis, 100, 101, 115 Acid hydrolysis, 174, 182, 204–207, 210, 224, 302 Acid pretreatment, 173, 175, 182, 205, 206, 210, 223, 224 Acidity, 19, 147, 353, 362, 379 Acidogenesis, 222, 226, 245, 246, 292, 304, 309 Adaption mechanism, 321 Adsorbents, 264, 265, 367 Aeration, 39, 45 Agricultural wastes, 74, 76, 211, 246, 269, 287, 321 Agroindustrial crops, 320 Airlift photobioreactor, 46, 339 Airlift reactors, 46 Alcohol dehydrogenase, 186, 225, 226, 228, 230, 231, 253, 302 Alcoholysis, 363 Aldol condensation, 221, 242 Algae manipulation, 185 Algal biomass, 11, 26, 27, 30–32, 35, 37, 40–42, 48, 52, 53, 58–63, 81, 167, 170, 176, 177, 179–185, 239, 248, 329, 353, 356, 358 Algal cell wall, 170, 176, 180, 181, 330 Algal cultivation, 8, 11, 18, 40, 44, 81, 176, 179, 185 Alkaline peroxidases, 235

Alkaline pretreatment, 182, 205, 207, 289, 290 Ammonical Nitrogen, 204 Anaerobic conditions, 212, 229, 253, 307 Anaerobic digestion, 48, 50, 52, 53, 58, 61, 81, 198, 199 Anisole, 120–126 Anthrospira platensis, 185 Aromatics, 115, 116, 118, 119, 126, 379 Arthrospira platensis, 178, 322 Artificial ponds, 44 Astaxanthin, 56, 322 Bacillus subtilis, 228, 253, 295 Base hydrolysis, 207 Base pretreatment, 207 Batch Fermentation, 223, 253, 255, 260, 290, 303 Bead beating, 176, 181 Beta glucosidase, 183 Bimetallic catalysts, 144 Biobutanol, 1, 18, 49, 70, 90, 219–232, 237–270, 285–310 Biodiesel, 7, 16–20, 23, 26, 27, 29–31, 35–37, 48–50, 58–64, 73, 74, 76, 82, 145, 169, 170, 220, 238, 248, 258, 286, 319–340, 352, 355, 356, 363, 364, 380 Biodiesel production, 17, 59, 60, 62, 63, 76, 319–340, 352, 356, 363 Bioenergy production, 44, 237, 321 Bioengineering, 319

Lalit Kumar Singh and Gaurav Chaudhary (eds.) Liquid Biofuel Production, (387–394) © 2019 Scrivener Publishing LLC

387

388

Index

Bioethanol feedstock, 169 Bioethanol, 1, 2, 18, 36, 48, 49, 51, 77, 78, 167–187, 197–214, 220, 248, 268, 286, 288, 364 Biofertilizer, 7, 12, 17, 23, 26, 38, 55, 322 Biofilm, 44 Biofuel pathways, 51 Biogas, 48, 50–53, 81, 169, 320, 322 Biohydrogen, 49, 52, 169, 198, 248, 286 Biological pretreatment, 183, 206, 250, 297 Biomass pre-treatment, 105 Bio-oil, 52, 90–147, 351–368, 371–381 Bio-oil upgrading, 92, 111, 112, 126, 364, 365, 367, 376, 379 Biophotolysis, 49, 52, 53 Bioproducts, 328 Biorefinery, 4, 26, 27, 29–31, 170, 250, 286, 322, 330 BOD, 62, 198, 203, 204 Botyrococcus braunii, 323 Botryucoccus brclunii, 361 Boudouard reaction, 128, 129, 140 Brackish water, 168, 176, 186 Bubble column photobioreactors, 45, 46 Bubble column, 10, 14, 27, 45, 46, 140–142 Bubbling fluid bed, 94, 99 Butamax, 220, 266, 269 Calorific value, 106, 129, 133, 138–140, 375–378, 381 Carbohydrate-rich algae, 167 Carbon dioxide fixation, 53, 185 Carbon dioxide, 5, 7, 10, 19, 24, 38, 41, 45, 50, 51, 61, 114, 169, 185, 201, 210, 212, 286, 293, 294, 303, 306, 353–356 Carbon-fixation group, 321 Carrier gas, 100, 101, 103, 110, 120–125 Catalyst supporting, 144

Catalysts, 106, 112, 113, 115, 118, 119–124, 140, 142–148, 221, 241, 331, 358, 363–367, 376, 378, 381 Catalytic cracking, 48, 111, 114–116, 366, 381 Catalytic fast hydropyrolysis, 117–119 Catalytic pyrolysis, 115, 116, 374–378 Catalytic vapour cracking, 374, 376, 377, 381 Cell disruption, 62, 181, 330 Cellulase enzymes hydrolysis, 208 Cellulosic biomass, 269, 286 Chemical flocculation, 42, 329 Chemical hydrolysis, 207 Chemical pretreatment, 174, 182, 183, 204, 205, 353 Chemical synthesis, 92, 135, 138, 139, 241 Chlamydomonas reinhardtii, 170, 171, 178, 183 Chlorella kessleri, 187, 356 Chlorella sp, 74, 80, 173, 178, 323, 324, 327, 334, 335, 338, 355, 359 Chlorella vulgaris, 54, 56, 62, 81, 170, 172, 173, 178, 324, 358, 359 Chlorococcum, 171, 178, 182, 185, 324 Chlorophyll, 44, 169, 354 Chloroplast, 19, 333, 354, 356 Chrysophyceae, 169 CimA genes, 230 Circulating fluid bed, 94, 99 Clarification, 198, 202, 203 Closed photobioreactor systems, 40, 41 Closed photobioreactors, 41, 44, 82, 176, 179, 322, 327, 328 Clostridia, 222, 226, 228, 244–246, 251, 255, 268, 292–299, 302–308 Clostridium Phytofermentans Xy1A, 78, 79 Clostridium sp., 221, 225, 242, 244, 246, 251, 294, 297, 307, 309 CoA-dependent pathway, 225, 228 COD, 198, 203, 204, 322 Cold plasma, 92, 117–126, 147

Index 389 Combustion, 17, 48, 51, 52, 81, 90, 94, 103, 104, 107, 114, 126, 128, 136, 138, 201, 212, 267, 351, 362, 374, 380, 381 Concentrated acid hydrolysis, 205–207 Contaminants, 24, 50, 127, 135, 186 COP, 198 Co-pyrolysis, 376 CPO, 198, 203 Cyanobacteria, 37, 55, 56, 185, 186, 321, 354 Cytosolic pathway, 230 Decarboxylation, 111, 114, 360, 377 Degree of deoxygenation, 112, 114, 115, 126 Depolymerization, 358, 360 Detoxification, 221, 223, 248, 250, 291, 298, 302, 306 Devolatilisation, 128 Dewatering of algal biomass, 42 Diatomaceae, 354 Dielectric barrier discharge, 118, 120–123 Dilute acid hydrolysis, 172, 205, 207 Discharge power, 122, 123 Dissolved gas concentration, 328 Distillation, 29, 30, 49, 53, 103, 111, 114, 176, 184, 185, 221, 223, 232, 255, 266, 290, 363, 365, 374, 376, 378 Distribution coefficient, 256–259 DNA shuffling, 74, 77 Downstream processing, 48, 223, 227, 229, 232, 302 Dried Distillers Grain (DDGS), 222 Dunaliella salina, 54 Dunaliella tertiolecta, 170, 172, 359, 361 D-xylose isomerase (XI), 78, 79 Ehrlich pathway, 226, 229, 230 Embden–Meyerhof pathway, 245, 246 Emulsification, 365, 374, 376, 379, 380 End product inhibition, 184, 232, 253, 255, 256

Endoglucanase, 183, 208, 297 Entrained flow gasifier, 130–137 Enzymatic Hydrolysis, 171–175, 182, 183, 199, 203, 208, 210, 224, 250, 286, 289, 290, 353 Enzyme Pretreatment, 197, 200 Esterification, 111, 363, 364, 374, 380, 381 Eukaryotic, 80, 252, 321, 353, 354 Eustigmatophytes, 354 Exoglucanase, 183, 208 Ex-situ catalytic pyrolysis, 377 Extraction, 3, 7, 12, 18, 19, 20, 22, 24, 25, 29, 37, 40, 48, 61, 62, 75, 183, 198, 202, 221, 223, 224, 232, 254–259, 266, 292, 322, 329–331, 374, 376, 381 Façade, 338 Fast pyrolysis, 91–98, 101–104, 107, 110, 111, 116, 357, 359, 362, 363 Fatty acid methyl esters, 327, 331 Fed-batch fermentation, 231, 253 Fermentable sugars, 170, 180, 181, 183, 206, 207, 286, 297, 352 Fermentation modes, 255 FFB, 198, 202, 203 Fiber, 36, 183, 201, 203–205, 208, 262, 263, 337 First generation biofuel, 2, 36 Fischer-Tropsch synthesis, 135, 140, 141, 145 Fixed bed gasifier, 130, 135 Flash pyrolysis, 356–357 Flat panel photobioreactor, 47, 48 Flat plate photobioreactors, 41, 47, 339 Fossil fuels, 1, 6, 22, 36–38, 49, 89, 90, 168, 199–201, 250, 268, 320, 323, 351–353, 374 Fuel blend, 168 Furans, 81, 103, 375 Gas stripping, 222, 223, 232, 256, 259, 260, 266, 290, 300, 308

390

Index

Gasification, 48, 52, 58, 76, 90–92, 97, 111, 114, 126, 130, 135, 136, 138–140, 357, 379 Gasification temperature, 138–140 Gasifying agent, 91, 92, 126, 128, 130, 135, 137–139 Gelidium amansii, 170, 175, 184 Gene expression cassettes, 78, 79 Genetic transformation, 185, 186 Geobacter sulfurreducens, 230 Gevo, 220, 269 Global transcription machinery engineering (gTME), 227 Global warming, 80, 89, 168, 198, 201 Glucoamylase, 171, 177 Glycolysis, 222, 303, 306, 333 Glyoxylate pathway, 230 Gracilaria verrucosa, 175, 184 Green algae, 55, 169, 170, 321, 354, 358, 359 Green microalgae, 323, 327, 332 Greenhouse gas emission, 48, 50, 55, 114, 168, 219 Greenhouse gases, 36, 80, 168, 199, 286 Haematococcus pluvialis, 54 Heat and electricity generation, 91, 133, 135 Heating value, 52, 53, 92, 103, 104, 110, 115, 126, 289, 357, 362, 363, 365, 380 Hemicellulose, 51, 74, 90, 91, 103, 105–107, 182, 183, 199, 203–210, 213, 246–249, 286–288, 291–293, 297, 302, 352, 375 Heterogeneous catalysts, 358, 363 Heterotrophic cultivation, 81 Heterotrophic growth, 74 High value co-products from algal biomass, 35 Homogeneous catalysts, 363 Homogenization, 45, 179, 330 Horizontal tubular photobioreactors, 46, 47

Hybrid separation processes, 237, 266 Hydrocarbon, 35, 90, 92, 107, 110, 114–119, 126, 129, 138, 141, 143, 201, 365, 375–379 Hydrocarbons cracking, 138, 139 Hydro-cracking, 366, 367 Hydrocyclone, 198, 202, 203 Hydrodeoxygenation, 111, 366, 378 Hydrodynamic parameters, 337 Hydrogenation, 111, 119, 128, 144, 145, 241, 242, 360, 366, 374, 376, 378 Hydrogenolysis, 124, 145, 360 Hydrolysis, 53, 81, 107, 109, 171–175, 177, 182–184, 197, 203–211, 223, 224, 230, 246, 248, 250, 286, 288–290, 296, 301, 302, 333, 353 Hydrothermal, 106, 171, 173, 207, 221, 223, 353, 356, 358, 360, 363 Hydrothermal carbonization, 106 Hydrothermal liquefaction, 353, 356, 358, 360, 363 Hydrothermal treatment, 106, 173 Hydrotreating, 111, 112, 117, 119, 366, 367 Hydroxymethylfurfural, 91, 182, 207 Illumination, 39, 41, 47, 180, 328, 337, 339 In situ recovery, 231 Inorganic compounds, 105, 106 In-situ catalytic pyrolysis, 374, 376 In-situ recovery, 231 Ionic liquids, 249, 250, 258, 259 Isobutanol, 220, 225–230, 240 Jet fuel, 48, 50 Ketones, 103, 112, 115, 362, 378 Kinetic characteristics, 228 Lactobacillus brevis, 228, 253, 295, 308 Leptospira interrogans, 230

Index 391 Light hydrocarbon, 110, 146 Light intensity, 38, 39, 41, 47, 51, 187, 328, 339 Lignocellulose, 74, 81, 207, 210, 213, 228, 269, 302, 352 Lignocellulosic biomass, 50, 74, 75, 77, 81, 90, 105, 168, 170, 238, 239, 248, 252 Lignocellulosic Hydrolysate, 74, 77, 80–82, 309 Lignocellulosic waste biomass, 220 Lignocellulosics, 222, 294 Lipid extraction, 322, 329, 330 Liquid fuel, 73, 90, 133, 135, 139, 140, 239, 267, 288, 291, 353, 362, 363, 374 Long chain hydrocarbon, 141, 144, 146 Macroalgae, 44, 49, 51, 169, 170, 173, 178, 182, 184, 353 Mass cultivation, 47, 82, 179 Mass transfer, 45–47, 94, 105, 112, 135, 143, 147, 177, 179, 180, 259, 262, 263, 328, 330, 337, 338 Mechanical comminution, 181 Membrane reactors, 222 Metabolic engineering, 74, 77, 185, 227, 228, 230, 251, 253, 268, 269, 294, 332–336 Metallic catalysts, 143, 144 Methanococcus jannaschii, 230 Microalgal biodiesel, 73, 74, 319, 320 Microalgal cultivation, 81, 82, 322 Microalgal growth, 48, 81, 322, 339 Microalgal hydrolysate, 74 Microwave treatment, 181 Mitosis, 355 Mixotrophic cultivation, 81 Moisture content, 53, 90, 92, 106, 107, 128, 130–135, 137, 362 Molecular distillation, 376 Multicellular structure, 321 Multi-tubular fixed bed, 140–142

Nannochloropsis, 323, 325, 327, 334–336, 358, 361 Natural waters, 40, 44 NDF, 204 Neochloris oleoabundans, 81, 355 Non-catalytic fast hydropyrolysis, 117–119 Non-fermentable sugars, 74 Non-food agricultural products, 220 Non-thermal plasma, 117 Oil and grease, 199, 203, 204 Oleaginous Yeast, 73 Olefin, 140, 143, 144 Oligomers, 362 Omics, 232, 333 Omics-based approaches, 232 Open cultivation, 80, 328 Open pond production pathway, 40 Open ponds, 39, 61, 82, 176, 178, 327, 328, 337 Operating cost, 37, 44, 82, 107, 134, 138, 139, 221, 267 Operating parameters, 128, 138, 145, 267, 295, 357 Operation costs, 147, 328 Organic carbon, 41, 74, 200, 201 Overexpression, 227, 229, 230, 232, 308, 333, 334, 335 Oxidation, 61, 91, 92, 104, 126, 128, 135, 140, 141, 180, 209, 323, 333 Oxygen content, 91, 104, 112–116, 126, 357, 360, 362, 376–378 Oxygen production, 321 Paddle wheel raceway ponds, 327 Paraffin, 140, 143, 144 Penicillium, 183 Pentose phosphate pathway, 78, 79, 245, 246, 303 Pentose sugars, 74, 75, 81, 246, 293, 303 Phaeodactylum tricornutum, 323, 326, 329, 333, 335

392

Index

Phenolic compounds, 81, 105, 291 Photoautotrophic cell, 41, 356 Photoautotrophic production, 40, 320 Photoheterotrophic, 41, 353 Photosynthesis, 39, 41, 47, 57, 61, 82, 169, 179, 185, 200, 353, 354, 356, 374 Photosynthetic, 38, 41, 44, 46, 47, 50, 56, 57, 61, 80, 169, 170, 179, 248, 321, 333, 353, 355, 356 Photosynthetic microorganisms, 80, 248, 321, 353, 355 Photosynthetic pigments, 44 Physical pretreatment, 181 Physicochemical pretreatment, 204 Pichia Stipitis, 77, 175 Pigment, 38, 44, 49, 52, 53, 56, 169, 253, 322, 353 PK2 gene, 78, 79 Polar solvents, 104, 365 Polysaccharides, 37, 48, 51, 187, 222, 224, 245, 246, 293, 352 Polyunsaturated fatty acid, 37, 52, 56, 332 POME characterization, 203 Prymnesiophyceae, 354 Pseudomonas putida, 228, 253, 295 Pyrolyser, 94, 99–102, 115 Pyrolysis, 48, 52, 76, 90–107, 110–147, 204, 321, 353, 356–365, 374–380 Pyrolysis liquid, 92, 103, 116 Pyrolysis temperature, 107, 110, 358, 375 Pyruvate, 78, 79, 186, 222, 226, 230, 231, 245, 246, 303, 306, 332 Pyruvate decarboxylase, 186, 226 Pyruvate kinase, 78, 79 Raceway ponds, 40, 177, 179, 327 Recombinant yeast strain ScF2, 74, 77 Red algae, 54, 170, 354 Reduced sugar, 227 Reducing sugar, 184, 204, 205, 294 Respiration rate, 77 Rotating cone, 92, 93, 96, 98, 100

Saccharification, 170, 172, 175, 177, 184, 208, 248, 254, 286, 290, 297–301 Saccharomyces bayanus, 184 Saccharomyces cerevisae, 211 Salinity, 39, 179, 327, 328 Sargassum fulvellum, 170 Scenedesmus, 43, 57, 58, 62, 170–172, 178, 181, 185–187, 323, 326, 330, 360, 361 Screw reactor, 101 Seaweed, 44, 56, 169, 176, 187, 359 Second generation biofuel, 36, 17, 168, 220, 268, 320 Separate hydrolysis and fermentation, 183, 290, 296, 301 Sequestration, 37, 80 Shallow ponds, 327 Simultaneous saccharification and fermentation, 184, 254, 296, 301 Single reactor, 211 Slow pyrolysis, 91, 357, 359, 360, 374 Slurry bubble column, 140–142 Solubilization, 207 Solvent toxicity, 231, 268, 295 Solventogenesis, 222, 226, 245, 246, 292, 293, 304, 305, 309 Solvent-tolerant microbial strains, 219 Spent biomass, 75 Spirulina, 40, 43, 54, 56, 173, 179, 358, 360, 361 Steam explosion, 106, 108, 205, 221, 223, 224, 249, 250 Steam reforming, 91, 114, 128, 129, 138, 139, 141, 374, 376, 379 Steam to biomass ratio, 138 Sterilization, 48, 198, 202, 203 Stirred type bioreactor, 45 Strain engineering, 250 Streptomyces, 183, 253 Sugar pathway, 177 Supercritical fluid, 330, 376 Surfactants, 363, 365 Synechocystis, 185, 186

Index 393 Syngas, 52, 91, 92, 126–147, 220 Synthetic pathways, 228 TDS, 204 Technological barriers, 219 Terrestrial crops, 82 Thermal treatment, 103 Thermo-chemical, 52, 107, 250, 269, 360 Third generation bioethanol, 168 Threonine pathway, 231 Torrefaction, 106, 107, 109 Total Nitrogen, 204 Transesterification, 20, 29, 48, 49, 320, 322, 331, 364 Transport fuels, 48, 63 Trichoderma, 183, 290 Triglycerides, 320, 322 TSS, 62, 204 Tubular photobioreactor, 41, 45–47

Ultrasonication, 181, 330 Ulva lactuca, 170 Upgrading techniques, 373 Value-added products, 169, 309, 322 Van Krevelen plot, 113 Vertical tubular photobioreactor, 45 Waste biomass, 114, 220, 287 Water content, 42, 102–106, 110, 362, 363, 375 Water gas reaction, 128, 129, 138, 139 Water gas shift reaction, 128, 129, 138, 139, 140, 143–145 Yeast fermentation, 170 Yeast integrative plasmid pRS403, 78 Zeolites, 114–116, 264, 265, 367 Zymomonas mobilis, 172, 186