Fuels, chemicals and materials from the oceans and aquatic sources 9781119117179, 1119117178, 9781119117186, 1119117186, 9781119117193

Table of contents :
Content: Overview of ocean and aquatic sources for the production of chemicals and materials --
Production and conversion of green macroalgae (Ulva spp.) --
A new wave of research interest in marine macroalgae for chemicals and fuels : challenges and potentials --
Kappaphycus alvarezii : a potential sustainable resource for fertilizers and fuels --
Microalgae bioproduction : feeds, foods, nutraceuticals, and polymers --
Innovations in crustacean processing : bio-production of chitin and its derivatives --
Recent progress on the utilization of chitin/chitosan for chemicals and materials --
Characterization and utilization of waste streams from mollusc aquaculture and fishing industries --
Fish processing waste streams as a feedstock for fuels.

Citation preview

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Wiley Series in Renewable Resources Series Editor Christian V. Stevens – Faculty of Bioscience Engineering, Ghent University, Ghent, Belgium

Titles in the Series Wood Modification – Chemical, Thermal and Other Processes Callum A. S. Hill Renewables-Based Technology – Sustainability Assessment Jo Dewulf & Herman Van Langenhove Introduction to Chemicals from Biomass James H. Clark & Fabien E.I. Deswarte Biofuels Wim Soetaert & Erick Vandamme Handbook of Natural Colorants Thomas Bechtold & Rita Mussak Surfactants from Renewable Resources Mikael Kjellin & Ingegard Johansson Industrial Application of Natural Fibres – Structure, Properties and Technical Applications Jorg Mussig Thermochemical Processing of Biomass – Conversion into Fuels, Chemicals and Power Robert C. Brown Biorefinery Co-Products: Phytochemicals, Primary Metabolites and Value-Added Biomass Processing Chantal Bergeron, Danielle Julie Carrier & Shri Ramaswamy Aqueous Pretreatment of Plant Biomass for Biological and Chemical Conversion to Fuels and Chemicals Charles E. Wyman Bio-Based Plastics: Materials and Applications Stephan Kabasci Introduction to Wood and Natural Fiber Composites Douglas Stokke, Qinglin Wu & Guangping Han Cellulosic Energy Cropping Systems Douglas L. Karlen Introduction to Chemicals from Biomass, Second Edition James Clark & Fabien Deswarte Lignin and Lignans as Renewable Raw Materials: Chemistry, Technology and Applications Francisco G. Calvo-Flores, Jose A. Dobado, Joaquin Isac-Garcia & Francisco J. Martin-Martinez Cellulose Nanocrystals: Properties, Production and Applications Wadood Hamad

Forthcoming Titles Biorefinery of Inorganics: Recovering Mineral Nutrients from Biomass and Organic Waste Erik Meers & Gerard Velthof Bio-Based Solvents Francois Jerome & Rafael Luque Nanoporous Catalysts for Biomass Conversion Feng-Shou Xiao & Liang Wang The Chemical Biology of Plant Biostimulants Danny Geelen Biobased Packaging: Material, Environmental and Economic Aspects Mohd Sapuan Salit & Muhammed Lamin Sanyang Thermochemical Processing of Biomass: Conversion into Fuels, Chemicals and Power 2e Robert C. Brown

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources Edited by

FRANCESCA M. KERTON Department of Chemistry, Memorial University of Newfoundland, Canada

NING YAN Department of Chemical and Biomolecular Engineering, National University of Singapore, Singapore

This edition first published 2017 © 2017 John Wiley & Sons Ltd 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. The right of Francesca M. Kerton and Ning Yan to be identified as the authors of the editorial material in this work has been asserted in accordance with law. Registered Offices John Wiley & Sons, Inc., 111 River Street, Hoboken, NJ 07030, USA John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK Editorial Office 9600 Garsington Road, Oxford, OX4 2DQ, UK For details of our global editorial offices, customer services, and more information about Wiley products visit us at www.wiley.com. Wiley also publishes its books in a variety of electronic formats and by print-on-demand. Some content that appears in standard print versions of this book may not be available in other formats. Limit of Liability/Disclaimer of Warranty In view of ongoing research, equipment modifications, changes in governmental regulations, and the constant flow of information relating to the use of experimental reagents, equipment, and devices, the reader is urged to review and evaluate the information provided in the package insert or instructions for each chemical, piece of equipment, reagent, or device for, among other things, any changes in the instructions or indication of usage and for added warnings and precautions. 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. 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. 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. Library of Congress Cataloging-in-Publication Data applied for Hardback 9781119117162

Cover design by Wiley Cover images: (Top) © LEONELLO CALVETTI/Gettyimages; (Bottom Left) Ingram Publishing / Alamy Stock Photo Set in 10/13pt TimesLTStd by SPi Global, Chennai, India

10 9 8 7 6 5 4 3 2 1

Contents List of Contributors Series Preface Preface 1

2

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials Francesca M. Kerton and Ning Yan 1.1 Introduction 1.2 Shellfish-Based Biomass 1.2.1 Crustacean Shells 1.2.2 Mollusc Shells 1.3 Finfish-Based Biomass 1.4 Plant-Based Biomass 1.5 Summary and Outlook References Production and Conversion of Green Macroalgae (Ulva spp.) Shuntaro Tsubaki, Wenrong Zhu and Masanori Hiraoka 2.1 Production of Ulva Biomass 2.1.1 Land-Based Tank Culture in K̄ochi 2.1.2 Improvement for More Intensive Culture 2.2 Conversion of Ulva Biomass 2.2.1 Microwave-Assisted Hydrothermal Reaction of Biomass 2.2.2 Microwave-Assisted Conversion of Ulva Biomass 2.3 Conclusions References

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1 1 3 3 7 9 12 13 14 19 19 20 25 27 28 29 36 36

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A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels: Challenges and Potentials Ravi S. Baghel, Vaibhav A. Mantri and C.R.K. Reddy 3.1 Introduction 3.2 Macroalgal Feedstock for Chemicals 3.3 Marine Macroalgae as a Biorefinery Feedstock 3.4 Marine Macroalgal Biomass as an Energy Feedstock 3.4.1 Bioethanol 3.4.2 Biodiesel 3.4.3 Biobutanol 3.4.4 Bio-oil 3.5 Advances in Cultivation Technology 3.6 Marine Algal Cultivation for CO2 Sequestration 3.7 Opportunities, Challenges and Conclusions References

4

Kappaphycus alvarezii: A Potential Sustainable Resource for Fertilizers and Fuels Dibyendu Mondal and Kamalesh Prasad 4.1 Introduction 4.2 Composition and Processing of Kappaphycus alvarezii 4.3 Simultaneous Production of Liquid Fertilizer (κ-Sap) and κ-Carrageenan from Fresh Kappaphycus alvarezii Seaweed 4.4 κ-Sap as Potential Plant Stimulant 4.5 Manipulation of κ-Sap for Sustainable Biomass Intensification of Maize 4.6 Bioethanol Production from Kappaphycus alvarezii 4.6.1 Pretreatment of Freshly Harvested Biomass 4.6.2 Hydrolysis of the Dry Biomass to Obtain Fermentable Sugars 4.6.3 Pretreatment of Hydrolysate to Reduce the Concentration of Fermentation Inhibitory Components 4.6.4 Enzymatic Fermentation of the Hydrolysate to Yield Ethanol 4.6.5 Purification of Ethanol from Fermentation Broth 4.7 Fuel Intermediates and Useful Chemical from Kappaphycus alvarezii 4.8 Environmental Impact of Fuel and Fertilizers Production from Kappaphycus alvarezii 4.9 Conclusion and Future Prospect Acknowledgement References

43 43 44 45 46 47 48 48 55 55 56 57 58

65 65 66 68 69 71 72 74 74 74 76 77 77 79 79 79 80

Contents

5

6

7

Microalgae Bioproduction – Feeds, Foods, Nutraceuticals, and Polymers Clifford R. Merz and Kevan L. Main 5.1 Introduction 5.2 Microalgae and Bioproduction Methods 5.2.1 Microalgae Groups Considered 5.2.2 Bioproduction of Microalgae – Methods 5.3 Microalgae Feedstock Products and Coproducts 5.3.1 Microalgae as Animal Feed 5.3.2 Microalgae as a Human Food Source 5.3.3 Microalgae in Nutraceuticals 5.3.4 Biopolymers from Microalgae 5.4 Conclusion – The Path Forward Acknowledgments References Innovations in Crustacean Processing: Bioproduction of Chitin and Its Derivatives Heather Manuel 6.1 Introduction 6.2 Innovations in Crustacean Processing 6.2.1 Conventional Processing Technologies 6.2.2 Innovations in Crustacean Processing 6.3 Utilization of Marine By-Products 6.3.1 Processing Technologies for Crustacean By-Products 6.3.2 A Biorefinery Approach for Value-Chain Optimization of Crustacean Biomass Waste 6.4 Bioproduction of Chitin and Its Derivatives 6.4.1 Background 6.4.2 Isolation and Extraction of Chitin and Chitosan 6.4.3 Non-chemical Structural Modifications of Chitin and Chitosan 6.5 Conclusions References Recent Progress in the Utilization of Chitin/Chitosan for Chemicals and Materials Bin Li and Xindong Mu 7.1 Structure, Source and Properties of Chitin/Chitosan 7.2 Isolation and Purification of Chitin/Chitosan 7.3 Derivatives of Chitin/Chitosan 7.4 Utilization of Chitin/Chitosan for Chemicals and Materials 7.4.1 Utilization of Chitin/Chitosan for Chemicals

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83 83 85 85 86 94 94 95 96 98 102 103 103

113 113 115 115 122 128 129 130 132 132 134 139 141 143

151 151 153 155 156 156

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7.5

8

9

7.4.2 Utilization of Chitin/Chitosan for Materials Closing Remark and Perspectives References

Characterization and Utilization of Waste Streams from Mollusc Aquaculture and Fishing Industries Jennifer N. Murphy and Francesca M. Kerton 8.1 Introduction 8.2 Processing and Characterization of Mollusc Shells 8.2.1 Processing Technologies 8.2.2 Characterization of Shells 8.3 Applications of Mollusc Shells 8.3.1 Soil Amendment 8.3.2 Treatment of Metal Contamination and Acid Mine Drainage 8.3.3 Phosphate Removal and Water Purification 8.3.4 Building Materials 8.3.5 Mollusc-Derived Calcium Oxide in Catalysis 8.4 Conclusions References Fish Processing Waste Streams as a Feedstock for Fuels Kelly Hawboldt and Ibraheem Adeoti 9.1 Introduction 9.2 Fish Processing By-Product 9.3 Chemical and Physical Properties of Crude Fish Oil 9.3.1 Chemical Composition of Crude Fish Oil 9.4 Oil Recovery Processes and Parameters 9.4.1 Physical/Thermal Separation Processes 9.4.2 Chemical Extraction Processes 9.4.3 Biological/Chemical Hydrolysis and Fermentation 9.4.4 Purification 9.4.5 Preservation of Feedstock and the Recovered Oil 9.5 Fuel Properties of Crude and Refined Fish Oils 9.5.1 Rheological Properties 9.5.2 Chemical Properties Affecting Fuel Quality 9.5.3 Thermal Properties 9.5.4 Other Fuel Properties 9.6 Performance of Crude Fish Oil as a Fuel 9.7 Upgrading Marine Crude Bio-Oil 9.7.1 Types of Refined Fish Oil Products 9.7.2 Transesterification 9.7.3 Pyrolysis

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189 189 192 192 195 199 201 202 208 212 219 224 225 229 229 230 231 233 236 236 238 244 245 246 247 247 248 249 250 251 251 252 255 258

Contents

9.8

9.9 9.10 9.11

Index

9.7.4 Microemulsification 9.7.5 Alternative Processes Emission Comparison for Bio-Oils 9.8.1 Crude Fish Oil 9.8.2 Fish Biodiesel 9.8.3 Biogas from Fish Waste 9.8.4 Fish Biofuels from Other Processes Comparison of Crude Oil and Refined Oil Performance as a Fuel Comparison of Fish Biofuels Summary References

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258 259 259 261 262 263 264 265 268 268 269 277

List of Contributors Ibraheem Adeoti Department of Process Engineering, Memorial University of Newfoundland, Canada Ravi S. Baghel Marine Biotechnology and Ecology Division, CSIR – Central Salt & Marine Chemicals Research Institute, India; Academy of Scientific and Innovative Research (AcSIR), Central Salt & Marine Chemicals Research Institute, India Kelly Hawboldt Department of Process Engineering, Memorial University of Newfoundland, Canada Usa Marine Biological Institute, K¯ochi University, Japan

Masanori Hiraoka Francesca M. Kerton foundland, Canada

Department of Chemistry, Memorial University of New-

Bin Li CAS Key Laboratory of Bio-Based Material, Qingdao Institute of Bioenergy and Bioprocess Technology, Chinese Academy of Sciences, China Kevan L. Main Marine & Freshwater Aquaculture Research Program, Mote Marine Laboratory, USA Vaibhav A. Mantri Marine Biotechnology and Ecology Division, CSIR – Central Salt & Marine Chemicals Research Institute, India; Academy of Scientific and Innovative Research (AcSIR), Central Salt & Marine Chemicals Research Institute, India Heather Manuel Centre for Aquaculture and Seafood Development, Fisheries and Marine Institute of Memorial University of Newfoundland, Canada Clifford R. Merz

University of South Florida, College of Marine Science, USA

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List of Contributors

Dibyendu Mondal Natural Products & Green Chemistry Division, CSIR – Central Salt and Marine Chemicals Research Institute, India; Department of Chemistry, CICECO-Aveiro Institute of Materials, University of Aveiro, Portugal Xindong Mu CAS Key Laboratory of Bio-Based Material, Qingdao Institute of Bioenergy and Bioprocess Technology, Chinese Academy of Sciences, China Jennifer N. Murphy Department of Chemistry, Memorial University of Newfoundland, Canada Kamalesh Prasad Natural Products & Green Chemistry Division, CSIR – Central Salt and Marine Chemicals Research Institute, India; AcSIR – Central Salt & Marine Chemicals Research Institute, India C.R.K. Reddy Marine Biotechnology and Ecology Division, CSIR – Central Salt & Marine Chemicals Research Institute, India; Academy of Scientific and Innovative Research (AcSIR), Central Salt & Marine Chemicals Research Institute, India Shuntaro Tsubaki Department of Applied Chemistry, Graduate School of Science and Engineering Tokyo Institute of Technology, Japan Ning Yan Department of Chemical and Biomolecular Engineering, National University of Singapore, Singapore Wenrong Zhu

Graduate School of Kuroshio Science, K¯ochi University, Japan

Series Preface Renewable resources, their use, and modification are involved in a multitude of important processes with a major influence on our everyday lives. Applications can be found in the energy sector, paints and coatings, and the chemical, pharmaceutical, and textile industry, to name but a few. The area interconnects several scientific disciplines (agriculture, biochemistry, chemistry, technology, environmental sciences, forestry, etc.), which makes it very difficult to have an expert view on the complicated interaction. Therefore, the idea to create a series of scientific books that will focus on specific topics concerning renewable resources has been very opportune and can help to clarify some of the underlying connections in this area. In a very fast-changing world, trends are not only characteristic for fashion and political standpoints; science is also not free from hypes and buzzwords. The use of renewable resources is again more important nowadays; however, it is not part of a hype or a fashion. As the lively discussions among scientists continue about how many years we will still be able to use fossil fuels—opinions ranging from 50 to 500 years—they do agree that the reserve is limited and that it is essential not only to search for new energy carriers but also for new material sources. In this respect, renewable resources are a crucial area in the search for alternatives for fossil-based raw materials and energy. In the field of energy supply, biomass and renewable-based resources will be part of the solution alongside other alternatives such as solar energy, wind energy, hydraulic power, hydrogen technology, and nuclear energy. In the field of material sciences, the impact of renewable resources will probably be even bigger. Integral utilization of crops and the use of waste streams in certain industries will grow in importance, leading to a more sustainable way of producing materials. Although our society was much more (almost exclusively) based on renewable resources centuries ago, this disappeared in the Western world in the nineteenth century. Now it is time to focus again on this field of research. However, it

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should not mean a “retour á la nature,” but it should be a multidisciplinary effort on a highly technological level to perform research toward new opportunities, to develop new crops and products from renewable resources. This will be essential to guarantee a level of comfort for a growing number of people living on our planet. It is “the” challenge for the coming generations of scientists to develop more sustainable ways to create prosperity and to fight poverty and hunger in the world. A global approach is certainly favored. This challenge can only be dealt with if scientists are attracted to this area and are recognized for their efforts in this interdisciplinary field. It is, therefore, also essential that consumers recognize the fate of renewable resources in a number of products. Furthermore, scientists do need to communicate and discuss the relevance of their work. The use and modification of renewable resources may not follow the path of the genetic engineering concept in view of consumer acceptance in Europe. Related to this aspect, the series will certainly help to increase the visibility of the importance of renewable resources. Being convinced of the value of the renewables approach for the industrial world, as well as for developing countries, I was myself delighted to collaborate on this series of books focusing on different aspects of renewable resources. I hope that readers become aware of the complexity, the interaction and interconnections, and the challenges of this field and that they will help to communicate on the importance of renewable resources. I certainly want to thank the people of Wiley’s Chichester office, especially David Hughes, Jenny Cossham, and Lyn Roberts, in seeing the need for such a series of books on renewable resources, for initiating and supporting it, and for helping to carry the project to the end. Last, but not least, I want to thank my family, especially my wife Hilde and children Paulien and Pieter-Jan, for their patience and for giving me the time to work on the series when other activities seemed to be more inviting. Christian V. Stevens Faculty of Bioscience Engineering Ghent University, Belgium Series Editor “Renewable Resources” June 2005

Preface This book provides a holistic view on fuels, chemicals and materials from renewable sources in the oceans and other aquatic media. To our knowledge, it is the first of its kind to cover water-based biomass—both plants and animals—for value-added applications beyond food, despite the fact that there are previously published books focused on more specialized sources (such as algae). The concept of biorefinery, referring to processes that convert biomass into fuels, chemicals and materials, has received wide awareness and acknowledgement in the new century. The first-generation biorefinery uses sugar- or starch-rich crops, associated with the issue of food security, while the second-generation biorefinery is based on cellulosic materials. In both cases, however, land scarcity sometimes becomes a limiting factor. In this context, oceans and other aquatic media, which account for over 2.5 times more area than land on Earth, appear to be a complementary source of feedstocks for biorefineries. We realized the underestimated potential of a great variety of water-based biomass resources years back. Together with other researchers around the world, we have strived to extend the concept of biorefinery to include diversified biomass resources from the ocean. For instance, F. Kerton and coworkers proposed and practiced the concept of ‘Marine Biorefinery’, taking fishery by-products and transforming them into a range of value-added products in Newfoundland, Canada, while N. Yan proposed and practiced the concept of ‘Shell Biorefinery’, in which waste crustacean shells are fractionated into three major components and further upgraded for a range of applications. To date, research and development toward this sort of biorefinery are still in a nascent state, with most work only demonstrated in the lab while large-scale commercial productions may be years ahead. However, there is a consensus being reached around the globe that the valorization of ocean biomass could nicely complement the existing biorefinery, to help avoid the compromise of food security and land use for human beings. Ocean biomass also features unique components and

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structures enabling the production of high-value chemicals and materials that are difficult to be obtained from other biomass resources or fossil fuels. These promising aspects made us come together and organize this book, covering various aspects of ocean-biomass-based biorefinery. The book is structured in the following manner: Chapter 1 provides an overview of ocean and aquatic sources for chemicals and materials. Chapters 2–4 describe the production, harvesting and conversion of marine macroalgae into fuels and other compounds. Chapter 5 switches to the topic of microalgae, reviewing its transformation into feeds, foods, nutraceuticals and polymers. Chapters 6 and 7 are focused on crustacean shells, with the Chapter 6 providing recent developments in fractionation of shells into chitin, while Chapter 7 summarizes a broad range of applications of chitin in chemical production and materials science. The final two chapters (Chapters 8 and 9) describe the utilization of waste streams from mollusc and finfish industries, respectively. This book should be able to serve as a valuable reference for academic and industrial professionals in research and development sectors in renewable fuels, chemicals and materials. Most chapters are written at an introductory level but with sufficient details to serve both undergraduate and graduate students majoring in chemistry, chemical engineering, marine sciences and biotechnology and beyond. Finally, the editors would like to express their gratitude to all the chapter authors for their invaluable time and contribution to the book and our colleagues, students and family for their patience while we worked on it. Francesca M. Kerton Memorial University of Newfoundland, St. John’s, Canada Ning Yan National University of Singapore, Singapore October 2016

1 Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials Francesca M. Kerton1 and Ning Yan2 1 Department 2 Department

1.1

of Chemistry, Memorial University of Newfoundland, Canada of Chemical and Biomolecular Engineering, National University of Singapore, Singapore

Introduction

The Earth is a watery planet—about 71% of its surface is covered by water [1]. Among all liquid water resources, less than 1% is freshwater, and over 99% is salty seawater. Freshwater in lakes and rivers, despite being in a very small percentage, has shaped our civilizations since the beginning of humankind. On the other hand, people’s perspective towards the ocean has been changing over time. In the old days, the oceans served for trade, adventure and discovery, as it set different civilizations apart. At present, the oceans are widely regarded as one of Earth’s most valuable natural resources for food, various minerals, crude oil and natural gas. As there is an increasing concern regarding sustainability, human beings currently strive for a paradigm shift of obtaining resources from renewable feedstocks Fuels, Chemicals and Materials from the Oceans and Aquatic Sources, First Edition. Edited by Francesca M. Kerton and Ning Yan. © 2017 John Wiley & Sons Ltd. Published 2017 by John Wiley & Sons Ltd.

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Figure 1.1 Overview of the animal and plant resources from the ocean and other aquatic sources: microalgae, macroalgae, fish, crustaceans and molluscs.

instead of non-renewable, depleting ones. More than 150,000 animals and 100,000 plants can be found in the oceans, all of which are renewable organic species. Sea plants can be divided into microalgae and macroalgae, whereas sea animals can be broadly categorized into three main types, namely fish, crustaceans and molluscs (Figure 1.1). Unfortunately, the huge potential of the oceans and other aquatic sources to provide renewable organic carbon, hydrogen, nitrogen and other elements as starting materials for chemicals and materials appears to be underestimated. Indeed, according to the data from Web of Science in 2015, of the total relevant papers on renewable feedstocks, only 2.3% were concerned with algae or oceanic biorefinery [2]. In fact, compared to conventional land-based biomass, aquatic (in particular, oceanic) biomass has several advantages [3]. First of all, a majority of seaweeds and fishery waste are not consumed as human food, and as such, there are no ethical issues of compromised food supply due to chemical and material production. At the same time, the development of ocean-based biorefinery can release the land area constrains, which are a serious problem in some countries such as Japan and Singapore. Many areas in the world are short of fertile soil for the generation of land-based biomass, and through the development of ocean-sourced feedstocks, people in these regions would utilize renewable materials without costly land-based agriculture. Last but not least, certain oceanic biomass species have intrinsic advantages over land-based resources, such as faster growth rate, less demanding growth conditions, more enriching components and so on. People have achieved remarkable success in harnessing land-based biomass—starch, woody biomass and vegetable oils—for fuels and chemicals. A landmark event was the opening of the world largest cellulose bioethanol

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials

3

refinery plant with an annual productivity of 30 million gallons by DuPont in November 2015 [4]. Woody biomass, consisting primarily of cellulose, hemicellulose and lignin, enters the biorefinery to be separated and further converted into a wide scope of valuable products [5, 6]. We could anticipate similar concepts towards valorization of aquatic-source-based biomass feedstocks. In the aquatic biomass refinery, ‘wastes’ could be fractionated through an array of processes into different components and further transformed into end products via physical, chemical and biological treatments. Once these objectives are met, new opportunities for building waste industries from ocean-based feedstock will arise. To achieve that, strong supports from research institutes, governments, organizations, companies and the public are integral. In particular, groundbreaking fundamental research from researchers worldwide is crucially required to conquer the technical barriers for integrated, value-added applications of oceanic biomass. In this chapter, we aim to provide an overview of various feedstocks from ocean and other aquatic sources, including sea-plant-based biomass, finfish-based biomass and shellfish-based biomass. The chemical component, current production scale, utilization and potential application and/or upgrading of each of these are summarized in separate sections.

1.2 1.2.1

Shellfish-Based Biomass Crustacean Shells

Global shellfish production, such as crabs, shrimps and lobsters, reached around 12 million tons in 2014 [7]. With such massive production, and due to the significant shell content (e.g. the shell of a crab can account for 60% of its weight), tremendous amounts of waste are generated from these crustacean species every year. As an estimation, astonishing 6–8 million tons of waste from crustaceans are produced annually [8]. Long before the modern era, shells were used as currency and regarded as a symbol of wealth. Later on, they were gradually substituted with other materials and became useless. Nowadays, there has been essentially no satisfactory solution to utilize the crustacean shells. Raw shells, such as dried shrimp shell or crab shell powder, have very low monetary value. Newport International, a seafood company partnering with co-packing plants in many Southeast Asian countries, including Indonesia, Vietnam, Thailand and Philippines, sells the by-product of dried shrimp shells at merely US$ 100–120 per ton. The price is commensurable with wheat straws and corn stover, which are agricultural wastes typically sold at US$ 50–90 per ton [9]. Due to the very low profitability, a vast majority of waste shells are disposed or landfilled without use. In developing countries that lack regulations, waste shells are often directly discarded, posing environmental concern. In developed countries, disposal can be costly—for instance, as high as US$ 150 per ton can be charged in Australia and Canada.

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Crustacean shells constitute 15–40% chitin, 20–40% protein and 20–50% calcium carbonate [10]. With several million tons of shells generated worldwide each year, the huge potential value of such shells is currently wasted. It is crucial to reconsider how to utilize such an abundant and cheap renewable resource, rather than continue treating it as waste. Further details on the processing of crustacean shells and utilization of chitin and chitosan can be found in Chapters 6 and 7 of this book. The protein in shells is a good nutrient for animal feed. For example, the protein from Penaeus shrimp shell is a complete protein food as it contains all the essential amino acids. The ratio of essential amino acids to total amino acids is 0.4; the nutrient value is comparable with that of soybean meals [11]. The market demand for protein meal continues to increase due to the rapid growth in livestock breeding. If all the protein from crustacean waste shells from Southeast Asia is extracted as animal feed, an annual market value of over US$ 100 million could be expected even based on the most conservative estimation [12]. Calcium carbonate is widely applied in construction, pharmaceutical, agricultural and paper industries. Current production of calcium carbonate mainly comes from geological sources such as marble and chalk. Ground calcium carbonate, being the major product, has a market price based on a particle size, which ranges from US$ 60–66 per ton for coarse particles to US$ 230–280 per ton for fine particles [13]. Ultrafine particles can reach an astonishing US$ 14,000 per ton. Provided that the calcium carbonate from crustacean shells can only be made into coarse particles, an annual market value of up to US$ 45 million could be estimated from Southeast Asian countries. Due to its bio-origin, calcium carbonate from waste shells is superior to that from marble and limestone for applications involving human consumption, such as calcium carbonate tablets. The last major component, chitin, is a linear polymer of β(1→4)-linked 2-acetamido-2-deoxy-d-glucopyranose [14]. The structure of chitin is similar to that of cellulose, but chitin has an amide or an amine group instead of a hydroxyl group on the C2 carbon in the repeating unit. Aside from being one of the major components in crustacean shells, chitin is widely present in the exoskeleton of insects, fungi and plankton, making it the second most abundant biopolymer around the world, with approximately 100 billion tons produced per year [15]. Chitin and chitosan (the water-soluble derivative) have been identified as useful functional polymers in several niche applications, including cosmetics, water treatment and biomedicals [16]. However, the current utilization of chitin neither matches its abundance nor fully harnesses its structural uniqueness. Chitin serves as a major renewable feedstock that simultaneously offers organic carbon and organic nitrogen elements. While a consensus has been reached on the importance of renewable organic carbon, not much has been emphasized on renewable organic nitrogen resources. The necessity is not obvious—after all, nitrogen is the dominant fraction in the Earth’s atmosphere. However, nitrogen

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials

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gas has to be converted into ammonia prior to application or further transformations. Ammonia synthesis is undesirable for the low efficiency that this single process accounts for 2–3% global energy consumption [17]. In addition, three moles of hydrogen gas, which is currently produced from fossil fuels, are consumed for every mole of nitrogen gas. The chemical industry cannot claim to be sustainable without addressing the sustainability issue of the nitrogen source in its products. Chitin appears to be more suitable for the production of some nitrogencontaining compounds. The major elements required—organic carbon, nitrogen and oxygen—are already in place. Chitin is also enriched with functional groups, thus requiring fewer derivatization steps when used as a raw material compared with fossil fuels. Effective valorization of chitin into chemicals may represent a ‘Game-Changing Innovation’ by bringing substantial benefits for both the economy and environment. Valorization of shells from crustacean species is not easy. First and foremost, fractionation is needed for further physical, chemical or biochemical processing. However, the current commercialized route to fractionate crustacean shells is associated with serious environmental and economic issues. Two key steps in the process include the removal of protein from the shell by sodium hydroxide solution and the digestion of calcium carbonate by hydrochloric acid. If chitosan is the final product, an additional step of treating chitin with 40% concentrated sodium hydroxide solution is required. The entire process is destructive, wasteful and expensive—protein and calcium carbonate fractions are currently destroyed and never recovered; sodium hydroxide and hydrochloric acid are highly corrosive and hazardous; production of 1 ton chitosan from crustacean shells needs more than 10 tons water. All these factors lead to negative environmental impacts and high capital costs. As a result, the price of good-quality chitin is as high as US$ 200 per kilogram, although the starting material is not costly. Due to the high price, global industrial use of chitin is estimated to be only 10,000 tons annually [15]. The lack of competitive pricing of chitin in the market, in turn, limits its production scale, forming a ‘high cost/low demand/low production’ pattern. Economically and ecologically unfavourable, chitin production plants are absent in many developed countries and only exist on a small scale in countries such as Thailand and Indonesia. There are considerable challenges in the post-fractionation steps as well. While the utilization of calcium carbonate and proteins is comparatively easier, the transformation of chitin to value-added, nitrogen-containing chemicals is a critical problem. We envisage that the major obstacles in valorizing chitin to be similar to those in woody biomass valorization. Unlike fossil fuels, biomass feedstocks such as chitin and cellulose are highly functionalized, oxygen-enriched polymers. Side reactions occur easily, leading to the formation of a variety of complicated compounds under severe reaction conditions. In addition, natural chitin is a highly crystallized biomass that impedes accessibility of reagents to the polymer

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

chains. Finally, it is often challenging to separate these bio-based products from the reaction system in a cost-effective way. To establish a new profitable industry with crustacean shell waste, creative solutions have to be developed for both upstream and downstream sectors. In the upstream sector, the key issue is enabling manufacturing competitiveness to lessen production cost and environmental impact; in the downstream sector, the key issue is establishing economic sustainability via integrated, value-added applications of each component. A revolutionary fractionation method to separate chitin, calcium carbonate and proteins is highly desirable (Figure 1.2). For an ideal protocol, the following characteristics should apply: (i) all three major components are processed into separate fractions; (ii) strongly corrosive or hazardous reagents are avoided; and (iii) waste generation is minimized. Fortunately, new technologies that partially satisfy these criteria are emerging. For example, lactic acid fermentation has been developed for chitin production both on a lab scale and a pilot-plant scale [18]. The process adopts blended bacteria to simultaneously consume proteins and decompose calcium carbonate. Protein hydrolysate and calcium lactate can be recovered after chitin separation. Another method is to utilize specific ionic liquids, which can remarkably dissolve carbohydrate polymers and thus extract chitin from waste shells [19]. In this way, the produced chitin has high molecular weight and is thus suitable for processing into fibres and films. In addition to these, we propose exploring the possibility of shell fractionation via physical methods more intensively. Ball mill and steam explosion may be effective in separating the major components in the shells. Finally, a process combining chemical force and mechanical force might prove to

CaCO3 Fractionation

Shell waste

Constructions Papermaking Pharmaceuticals 6 4 O HO

OH 5 O

3 2 HN

1 O 7 8

n

Nitrogencontaining chemicals

Textiles

Chitin Food additives

Protein

Fertilizers

Animal feeds

Figure 1.2 The concept of “waste-shell refinery” for various useful chemicals and materials (diagram based on the concept presented in Ref. [8]). (Source: Data taken from Yan and Chen 2015 [8].)

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials

7

be advantageous, since synergistic effects may lead to unprecedented performance. To exemplify a scenario, combined use of ball mill and a small amount of acid catalyst leads to a complete degradation of wood without extra heating. A similar strategy could be applicable to shells, enabling a highly effective, solvent-free approach for fractionation. In the downstream sector, diversified utilization of each component is essential. While calcium carbonate and proteins can find direct applications, there is an untapped potential towards chitin utilization. The transformation of chitin into functional polymers and a series of value-added chemicals is a promising direction. There have been decades of research on chitin conversion to polymer derivatives for distinct applications. Conversion of chitin into small nitrogen-containing chemicals is also developing very fast, although it is still at a very early stage. A nitrogen-containing furan derivative was obtained directly from chitin via boric-acid-catalyzed depolymerization and dehydration [20, 21]. Recently, a number of other value-added chemicals have also been produced from chitin or chitin monomer on the lab scale [22–25]. Future investigation should be focused on the following: (i) exploration of new routes from chitin to other potentially related chemicals; (ii) enhanced product yield via improved catalysis and/or chitin pretreatment; and (iii) facile separation of products, such as membrane-based pervaporation technique. 1.2.2

Mollusc Shells

As described in Chapter 8 of this book, in 2013, over 18 million tons of molluscs were harvested, which amounted to 11% of the world’s fisheries, and they were mainly produced via aquaculture [7]. In addition to being a valuable source of protein in our diets, molluscs have the benefits of being able to reach maturity in only 2–3 years and are filter feeders, so they do not need to be fed by the farmer. Waste materials, which could be valorized, are produced at a number of stages during harvesting and processing this food. For example, some molluscs will be dead when harvested, die during harvesting or are damaged during the harvesting process (e.g. cracked shells). These wastes can be used to supply biorenewable calcium carbonate and possibly protein product streams (Figure 1.3). The protein can be used in a similar way to that obtained from crustacean shells as described earlier, that is, as a feed or fertilizer. More recently, there has been interest in the use of mussel protein as a nutritional supplement because it contains components that have the potential to treat obesity [26]. Therefore, we expect enhanced interest in mollusc production in the coming years, which could lead to more waste being produced. Furthermore, if the mollusc is processed before being marketed as a food (e.g. canned)—the meat will be removed from the shell, but the shell will often still contain residual protein (i.e. the adductor muscle of the mollusc). The waste streams from mollusc production, if not handled correctly, can be a biohazard, and if the waste is not used, it must be disposed of at specialist landfill sites with

8

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Shell

Protein

Cosmetics pharmaceuticals

Fish feed

Water/soil treatment acid mine drainage

Flavourings food supplements

Building materials composites catalyst support

Amino acids (chemical building blocks)

Figure 1.3 Potential products and applications of materials from a mollusc-based waste utilization process.

associated tipping costs of approximately US$ 150 per ton. This leads to an added incentive for farmers to explore alternative waste disposal/processing options as described as follows. There are two main processes that have been explored for cleaning mollusc shells in order to eliminate the biohazard risk and produce value-added materials. The first process involves burning off residual protein and organic matter by heating the shells to 500 ∘ C. This has been explored on a pilot-plant scale in the region of Galicia, Spain [27]. Various challenges were encountered during this scale-up, for example, SO2 and NO2 emissions when oil was used in the heating process. Furthermore, in such a process, the protein stream is wasted. From a sustainability perspective, biocatalytic (enzymatic) cleaning processes are perhaps more promising. Enzymatic proteolysis of finfish and crustacean by-products has been studied extensively, but there are few examples of its application to mollusc and mollusc shell processing. In most examples, only a small amount of the catalytic protease enzyme is needed, and temperatures are not much higher than room temperature (40–70 ∘ C) [28]. As with most enzymatic processes, the pH of the process must be monitored in order to prevent deactivation of the enzyme. The protein hydrolysate stream produced via such hydrolysis reactions has potential uses in flavourings or supplements within the food industry. Further hydrolysis and separations could yield amino acids, which could be used as chemical building blocks to build up more complex structures such as bioactive compounds (pharmaceuticals).

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials

9

Although less extensively studied compared to crustacean waste streams described in Section 1.2.1, applications of mollusc shells in a range of areas have been proposed and investigated on a lab scale. The calcium-carbonate-rich shells produced by molluscs have the potential to become high-value, low-volume products (e.g. cosmetics or medicine) or low-value, high-volume products (e.g. soil amendment or building materials). Applications of mollusc shells include the following: feed additives for poultry, soil amendment, treatment for acid mine drainage, water purification, additive for building materials (e.g. concrete) and lime (calcium oxide) production. Additonally, it is worth noting that the structure of biogenic calcium carbonate [29, 30] is significantly different from that of quarried calcium carbonate, and this may lead to additional value to end users. For example, in Thailand, a range of different shells were studied as cost-effective replacements for Portland cement in the production of plastering cement [31]. It is also worth noting that molluscs can play an important role in integrated multi-trophic aquaculture (IMTA) that is being explored worldwide as a method of sustainable farming. In such approaches, different organisms are grown/farmed in close proximity in an attempt to mimic natural aquatic ecosystems and prevent pollution of the oceans nearby. For example, a pilot project has been studied in the Bay of Fundy, New Brunswick, Canada [32]. Seaweed (Kelp), mussels and Atlantic salmon were grown nearby, uneaten salmon feed and faeces provided nutrients to the mussels and excess dissolved nitrogen and phosphorus produced during the salmon farming were taken up by the seaweed. It is hoped that such practices will prevent hypernutrification and eutrophication of coastal waters around finfish farms. If IMTA becomes a well-established way to reduce environmental impacts of finfish farming, levels of mollusc production will likely increase as the aquaculture industry continues to grow to meet the needs of a growing global population [7].

1.3

Finfish-Based Biomass

About 580 species of fish, including finfish and shellfish, are farmed worldwide, and production from wild capture and aquaculture exceeded 160 million tons in 2013, with aquaculture contributing 70 million tons [7]. The most important farmed finfish species are carp, tilapia, catfish and salmon. The amount and type of fish farmed often depend on the location and climate. For example, the major producers of farmed salmon are Chile, Norway, Scotland and Canada. Finfish farming can occur in cages at sea or on land. Research is ongoing towards development of farming sites further out at sea, as, at present, most of them are normally situated in sheltered, coastal locations. However, there is interest in developing multi-platform sites where IMTA (described earlier) can be pursued and monitored alongside off-shore wind-farm platforms so that food and power would be generated at a single site and costs of manpower and transport could be shared.

10

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources Hatchery/ nursey

Smolts (young fish) Grow-out cages

Feed

Harvest

Primary processing

Food distributor

Biofuel fertilizer supplements pet food

Bioprocessing

Secondary processing

Fish meal and fish oil

Figure 1.4

Stages in the production of farmed finfish and potential end products.

The typical stages in finfish farming are outlined in Figure 1.4. Upon harvesting, fish frequently undergo primary processing where they are decapitated and gutted and often fileted. The fish may then undergo secondary processing such as smoking or salting. During processing, significant quantities of waste are produced: heads, viscera (guts), belly flaps, frames (bones), skin, gills and blood water [33]. Approximately 70 million tons of fish per year are processed worldwide by filleting, freezing, canning or curing, and these activities generate 30–50% waste, that is, 21–35 million tons of waste [7]. It should also be noted that biological waste material is also produced during the farming process, such as morts (dead fish) are found in cages and must be removed in order to prevent disease. Furthermore, in some cases, disease outbreaks or severe weather (low or high temperatures) can cause spikes in fish deaths. In most cases, this waste (processing waste and morts) currently goes to landfill and, in some cases, is used to produce fertilizer. However, a major problem with this waste stream is that it is a biohazard, and this risk is heightened if the material is stored without undergoing some form of pretreatment (e.g. pasteurization). In order for the valorization potential of the waste to be maximized, it would be ideal for bioprocessing to occur as soon after harvesting and primary processing as possible. Some of the bioactive components within the fish waste stream are temperature and time sensitive (i.e. they decompose/degrade when heated or stored). If bioprocessing of properly stored and preserved material is performed, a broad range of high-value products could potentially be obtained, including amino acids, bioactive peptides, enzymes, [34] collagen, hydroxyapatite, calcium carbonate and ω-3 fatty acids. However, if the material degrades, the by-products of fish processing will only have a relatively low value, such as fertilizer, animal feed, heating oil and biogas from silage. In many cases, lessons can be learnt from waste management strategies and biosecurity containment approaches from other locations around the world. For

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials

11

example, in the field of salmon aquaculture, Canadians are exploring strategies previously used successfully in Norway and Scotland. From 2015 onwards, the Newfoundland Aquaculture Industry Association (NAIA), in cooperation with their member salmon-farming companies, has been investigating the industrial application and biosecurity benefits of ensiling waste from fish farms [35]. Appropriate on-farm salmon silage systems from Norway and Scotland were studied. Fish silage is ‘a liquid product made from whole fish or parts of fish that are liquefied by the action of enzymes in the fish in the presence of an added acid. The enzymes break down fish proteins into smaller soluble units, and the acid helps to speed up their activity while preventing bacterial spoilage’ [36]. In the NAIA systems, formic acid is being used, and the resulting silage is being shipped to a dairy farm to be used as a feedstock in an anaerobic digester, which is used to provide power on the farm. We think that there are many opportunities to enhance waste management strategies by combining efforts across several industries: in this case, agriculture (a dairy farm) and aquaculture (salmon farms in the region), but such efforts could involve other areas with biological waste streams such as food processing industries, forestry and municipal organic waste streams. Fish silage can be used as a fuel in an anaerobic digester to produce energy in the form of biogas. However, as described in Chapter 9 of this book, there is significant potential in isolating oils and generating liquid fuels from fish waste streams [37]. Some fish species are naturally oilier than others, with those fish with darker flesh pigments generally having a higher lipid content (e.g. salmon, trout and Arctic char). A number of processes have been used to obtain fish oil from fishery waste including the fishmeal process, supercritical carbon dioxide extraction and fermentation processes [37–39]. Innovations in this area have included the use of mobile fishmeal process plants that can travel around a region with fishing activities (wild capture or aquaculture) and perform the processing on-site without the need to build a dedicated fishmeal plant at each site. Once the fish oil is isolated, it can be purified or upgraded in a number of ways. Many of these methods have become well-established in the field of processing vegetable/land-based biomass and include transesterification to produce biodiesel and pyrolysis to produce bio-oil [40]. Studies to apply such methods to ocean or aquatic sourced biomass are still ongoing. There is significant room for progress and improvements, given the different chemical nature of the feedstocks. Furthermore, in many cases, it is not economically viable to process fishery waste on-site immediately, and this leads to by-product decomposition. In the case of lipids, this involves oxidation of the unsaturated bonds and means that they are no longer suitable in high-value products (e.g. ω-3 fatty acid food supplements). Therefore, the use of the oil component of the waste stream will often involve transformation into a fuel if the fish by-product stream is to be fully utilized.

12

1.4

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Plant-Based Biomass

Plants in our oceans can be divided into two categories: microalgae and macroalgae. Microalgae typically contain a greater proportion of natural oils for potentially producing fuels compared to macroalgae. Therefore, they have been extensively studied as a potential way of producing renewable fuels [41–43]. The US Department of Energy identified a number of attractive properties that microalgae possess that make them a preferred feedstock for producing biofuels. They are highly productive and can offer large biomass yields per area cultivated. They can be grown using a range of different sources of water and are not limited to being grown using freshwater, which is in limited supplies in many areas. Furthermore, microalgae can be used to recycle carbon from processes that produce CO2 -rich emissions such as power-generating stations that combust fossil fuels. If microalgae are used to produce oil for fuel, a by-product stream (rich in protein and carbohydrate) will also be obtained. Researchers are now studying the potential of whole utilization of microalgae (e.g. production of β-chitin, pigment and other by-products in addition to oil) [44] and new methods of growing microalgae efficiently in order to bring an algae-based biorefinery to fruition [45]. Further details on microalgae bioproduction and potential products are described in Chapter 5 of this book. Macroalgae or seaweed can also be used as a feedstock for chemicals, fuels and materials. They are generally subdivided into three types based on pigments—red, brown and green algae. In various locations around the world, due to eutrophic ocean conditions with enriched nitrogen and other nutrient levels, blooms of algae, known as green or golden tides, have occurred [46]. In addition to cultivating and harvesting seaweed for either food or other uses, it is important that the overgrown seaweed in these blooms can be processed and dealt with in a sustainable and productive manner. It is worth noting that algae have already been exploited industrially as they are the only sources of certain sulfated polysaccharides, which possess valuable gelation properties, for example, agar, carrageenan and alginic acid [47]. However, they contain secondary metabolites as well, which may be of high economic value as a secondary commercial product if the algae are processed in the correct way in order to maintain the metabolite’s structure and function. The United Nations’ Food and Agriculture Organization noted that commercial harvesting of macroalgae has now reached 25 million tons per annum and is valued at US$ 7.4 billion [7]. Furthermore, many have highlighted, as with microalgae, the potential of marine algae to act as a carbon sink and sequester carbon dioxide from the atmosphere. In order to make a marine biorefinery economically viable, it is important to produce several low-volume, high-value materials in addition to lower cost biopolymers, fuels and food stuffs. In this regard, macroalgae seem ideal feedstocks for biorefineries. In a recent case study, the red seaweed Kappaphycus alvarezii was studied in detail [48]. This seaweed is already cultivated in Asia

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials

13

as a source of carrageenan. It has a daily growth rate of 3–6%, and it sequesters 17–20 tons of CO2 per hectare per year. In addition to carrageenan, the seaweed contains 9–18 wt% dietary fibre, 6–15 wt% protein, 0.4–1.0 wt% lipid and 10–25 wt% minerals. Mondal et al. showed that, by crushing and centrifuging fresh K. alvarezii, two materials were obtained—a potassium chloride rich juice and carbohydrate-rich granules. The juice can be used to produce a fertilizer and also feed into other steps in the integrated biomass fractionation process. The granules can be extracted with seawater to give carrageenan for the hydrocolloid industry and residual cellulose as a by-product, which can be used as a solid fuel or processed further. As with land-based biomass, there are many options available for processing and upgrading macroalgae, and many have been explored and are discussed in more detail in later chapters of this book. A review was also recently published on the use of macroalgae in the production of energy and fuels [49]. In this area, microwave treatments seem particularly promising, given the large amounts of water within freshly harvested algae. Tsubaki et al. have shown that catalytic hydrolysis of algae into reducing sugars can be effectively performed under microwave irradiation, and this process and others are described in more detail in Chapter 2 [50]. It should also be noted that ton-scale microwave processing units are now being used industrially and may be valuable tools in developing a marine biorefinery using algae or other bio-feedstocks described earlier. As reported in a recent news article, one of the pioneers in the microwave field sees significant growth and future potential—‘in the same way that the rise of Tesla changed the car industry, he sees microwaves transforming chemical manufacturing’ [51]. As with all harvestable materials, there is always concern that crops will fail or become damaged in some way. Therefore, interest has grown in cultivating algae (both micro- and macro-) on land under more controlled conditions than is possible in the open ocean. In Japan, a land-based culturing system for the production of the edible seaweed Ulva prolifera has been developed, and this is described in more detail in Chapter 2 of this book [52]. Similar approaches and methods should be developed for other classes of algae. Furthermore, species can be developed through selective breeding programs to be enhanced in a particular secondary metabolite that is desirable from either a nutritional or a chemical feedstock point of view.

1.5

Summary and Outlook

The use of ocean-sourced biomass to produce chemicals, fuels and materials presents a cornucopia of opportunities. Advances are being made around the world in terms of sustainable development. However, as a source of renewable materials, the oceans are often overlooked. Furthermore, with increasing regulations regarding greenhouse gas emissions and taxation on carbon, the ability of algae to grow quickly and sequester carbon dioxide whilst affording a valuable

14

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

renewable feedstock is going to garner significant attention in the coming years. Therefore, in order to make progress, academia, industry and government need to work together, and collaborations across disciplines are needed. The global human population continues to grow, and the United Nations has highlighted aquaculture activities as an important way to make sufficient food (especially protein) in a sustainable way. The growth in this field during the past 10 years and into the future has been termed the ‘blue revolution’. In the food processing industries, it is impossible to avoid the production of waste streams, and 30–60% of the weight of shellfish or finfish is discarded in this way. The waste material represents a biohazard, and in many countries, legislation exists to prevent dumping at sea or placing it in a standard landfill site. The cost of waste disposal is significant (US$ 150 per ton), and this means that the aquaculture field is ripe for innovation in this regard, and waste materials should ideally be thought of as by-product streams for valorization. In this way, it is possible to produce a broad array of co-products including organic fertilizers, fuels, new composite materials and bioactive enzymes and proteins. The outlook for the establishment of ocean biorefineries is promising, and entrepreneurs are taking notice of these opportunities for establishing a paradigm shift in how we consider aquaculture and the fisheries. For example, in Newfoundland, a small-scale marine biorefinery has been set up, which is taking fishery by-products and transforming them into a range of value-added products [53]. We expect to see further economic and sustainable development towards valorization of ocean biomass in the following decades. The remaining chapters of this book describe some of the exciting discoveries to date in this field.

References 1. “Ocean”, Wikipedia: The Free Encyclopedia. Wikimedia Foundation, Inc., https://en .wikipedia.org/wiki/Ocean (accessed October 10, 2016). 2. Data from Web of Science for the Year 2015, Topic = Biomass OR Renewable, AND feedstock. 3. Kerton, F.M., Liu, Y., Omari, K.W. and Hawboldt, K. (2013) Green chemistry and the ocean-based biorefinery. Green Chemistry, 15 (4), 860–871. 4. Biofuels Digest, November 21, 2015, http://www.biofuelsdigest.com/bdigest/2015/11/01/ (accessed October 10, 2016). 5. Dodds, D.R. and Gross, R.A. (2007) Chemicals from biomass. Science, 318 (5854), 1250–1251. 6. Somerville, C., Youngs, H., Taylor, C. et al. (2010) Feedstocks for lignocellulosic biofuels. Science, 329 (5993), 790–792. 7. Food and Agriculture Organization of the United Nations. The State of World Fisheries and Aquaculture, 2014. 8. Yan, N. and Chen, X. (2015) Don’t waste seafood waste. Nature, 524 (7564), 155–157. 9. Pricing for Hay and Straw, Farming Online, http://www.farming.co.uk/prices/baled_hay_ straw/ (accessed October 10, 2016).

Overview of Ocean and Aquatic Sources for the Production of Chemicals and Materials

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10. Rinaudo, M. (2006) Chitin and chitosan: properties and applications. Progress in Polymer Science, 31 (7), 603–632. 11. Ibrahim, H.M., Salama, M.F. and El-Banna, H.A. (1999) Shrimp’s waste: chemical composition, nutritional value and utilization. Food, 43 (6), 418–423. 12. Pricing for Soybean Meal Futures, World Bank, http://www.indexmundi.com/ commodities/?commodity=soybean-meal (accessed October 10, 2016). 13. Indicative Chemical Prices A-Z, ICIS, Division of Reed Business Information, RELX Group, http://www.icis.com/chemicals/channel-info-chemicals-a-z/ (accessed October 10, 2016). 14. Chen, X. and Yan, N. (2014) Novel catalytic systems to convert chitin and lignin into valuable chemicals. Catalysis Surveys from Asia, 18 (4), 164–176. 15. Kim, S.-K. (2011) Chitin, Chitosan, Oligosaccharides and Their Derivatives: Biological Activities and Applications, CRC Press, Boca Raton, FL. 16. Pillai, C.K.S., Paul, W. and Sharma, C.P. (2009) Chitin and chitosan polymers: chemistry, solubility and fiber formation. Progress in Polymer Science, 34 (7), 641–678. 17. International Efficiency Technology Database, http://ietd.iipnetwork.org/content/ammonia# key-data (accessed October 10, 2016). 18. Cira, L.A., Huerta, S., Hall, G.M. and Shirai, K. (2002) Pilot scale lactic acid fermentation of shrimp wastes for chitin recovery. Process Biochemistry, 37 (12), 1359–1366. 19. Qin, Y., Lu, X., Sun, N. and Rogers, R.D. (2010) Dissolution or extraction of crustacean shells using ionic liquids to obtain high molecular weight purified chitin and direct production of chitin films and fibers. Green Chemistry, 12 (6), 968–971. 20. Chen, X., Chew, S.L., Kerton, F.M. and Yan, N. (2014) Direct conversion of chitin into a N-containing furan derivative. Green Chemistry, 16 (4), 2204–2212. 21. Chen, X., Liu, Y., Kerton, F.M. and Yan, N. (2015) Conversion of chitin and N-acetyl-d-glucosamine into a N-containing furan derivative in ionic liquids. RSC Advances, 5 (26), 20073–20080. 22. Bobbink, F.D., Zhang, J., Pierson, Y. et al. (2015) Conversion of chitin derived N-acetyl-d-glucosamine (NAG) into polyols over transition metal catalysts and hydrogen in water. Green Chemistry, 17 (2), 1024–1031. 23. Pierson, Y., Chen, X., Bobbink, F.D. et al. (2014) Acid-catalyzed chitin liquefaction in ethylene glycol. ACS Sustainable Chemistry & Engineering, 2 (8), 2081–2089. 24. Chen, X., Gao, Y., Wang, L. et al. (2015) Effect of treatment methods on chitin structure and its transformation into nitrogen-containing chemicals. ChemPlusChem, 80 (10), 1565–1572. 25. Omari, K.W., Besaw, J.E. and Kerton, F.M. (2012) Hydrolysis of chitosan into levulinic acid and 5-hydroxymethylfurfural in water under microwave irradiation. Green Chemistry, 14 (5), 1480–1487. 26. Cheema, S. and Gangadaran, S. (2016) Blue Mussels as Nutraceuticals to Target Obesity and to Improve Cardiovascular Health, Aquaculture Canada and Cold Harvest Conference, St. John’s, Canada. 27. Barros, M.C., Bello, P.M., Bao, M. and Torrado, J.J. (2009) From waste to commodity: transforming shells into high purity calcium carbonate. Journal of Cleaner Production, 17 (3), 400–407. 28. Silva, V.M., Park, K.J. and Hubinger, M.D. (2010) Optimization of the enzymatic hydrolysis of mussel meat. Journal of Food Science, 75 (1), C36–C42. 29. Pokroy, B., Fitch, A.N. and Zolotoyabko, E. (2007) Structure of biogenic aragonite (CaCO3 ). Crystal Growth & Design, 7 (9), 1580–1583.

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30. Li, X.D., Chang, W.C., Chao, Y.J. et al. (2004) Nanoscale structural and mechanical characterization of a natural nanocomposite material: the shell of red abalone. Nano Letters, 4 (4), 613–617. 31. Lertwattanaruk, P., Makul, N. and Siripattarapravat, C. (2012) Utilization of ground waste seashells in cement mortars for masonry and plastering. Journal of Environmental Management, 111 (Nov 2012), 133–141. 32. Nguyen, T. and Williams, T. (2013) Background Paper: Aquaculture in Canada, Publication No. 2013-12-E, Ottawa, Canada, Library of Parliament, http://www.lop.parl.gc.ca/content/ lop/ResearchPublications/2013-12-e.htm (accessed October 10 2016). 33. Blier, P.U. and Le Francois, N.R. (2010) Valorization of aquaculture by-products, in Finfish Aquaculture Diversification (eds N.R. Le Francois, M. Jobling, C. Carter and P.U. Blier), CAB International, Quebec, pp. 546–555. 34. Shahidi, F. and Kamil, J.Y.V.A. (2001) Enzymes from fish and aquatic invertebrates and their application in the food industry. Trends in Food Science and Technology, 12 (12), 435–464. 35. Green, D. (2016) The Newfoundland Aquaculture Industry Association Ensiling Demonstration Project, Aquaculture Canada and Cold Harvest Conference, St. John’s, Canada. 36. Tatterson, I. N. and Windsor, M. L. (1974) “Fish Silage”, Ministry of Agriculture, Fisheries and Food, Torry Research Station, Torry Advisory Note No. 64, Aberdeen, UK http://www .fao.org/wairdocs/tan/x5937e/x5937e00.htm#Contents (accessed October 10, 2016). 37. Adeoti, I.A. and Hawboldt, K. (2014) A review of lipid extraction from fish processing by-product for use as a biofuel. Biomass and Bioenergy, 63 (Apr. 2014), 330–343. 38. Rai, A.K., Swapna, H.C., Bhaskar, N. et al. (2010) Effect of fermentation ensilaging on recovery of oil from fresh water fish viscera. Enzyme and Microbial Technology, 46 (1), 9–13. 39. Liaset, B., Julshamn, K. and Espe, M. (2003) Chemical composition and theoretical evaluation of the produced fractions from enzymic hydrolysis of salmon frames with Protamex™. Process Biochemistry, 38 (12), 1747–1759. 40. Balat, M. (2008) Global trends on the processing of biofuels. International Green Energy, 5 (3), 212–238. 41. Sheehan, J., Dunahay, T., Benemann, J. et al. (1998) A look back at the U.S. Department of Energy’s Aquatic Species Program: Biodiesel from Algae. NREL/TP-580-24190. 42. Chisti, Y. (2010) Fuels from microalgae. Biofuels, 1 (2), 233–235. 43. Ferrell, J. and Sarisky-Reed, V. (2010) National Algal Biofuels Technology Roadmap. U.S. Department of Energy, Office of Energy Efficiency and Renewable Energy Biomass Program. http://biomass.energy.gov and http://energy.gov/eere/bioenergy/algal-biofuels (accessed October 10, 2016). 44. Merz, C. and Main, K. (2014) Microalgae (diatom) production – The aquaculture and biofuel nexus. Oceans’14 MTS/IEEE Conference proceedings – IEEE Xplore, St. Johns, Newfoundland, Canada. DOI: 10.1109/OCEANS.2014.7003242. 45. Chisti, Y. (2012) Raceways-based production of algal crude oil, in Microalga Biotechnology: Potential and Production (eds C. Posten and C. Walter), de Gruyter, Berlin, pp. 113–146. 46. Smetacek, V. and Zingone, A. (2013) Green and golden seaweed tides on the rise. Nature, 504 (5 Dec 2013), 84–88. 47. Bixler, H.J. and Porse, H. (2011) A decade of change in the seaweed hydrocolloids industry. Journal of Applied Phycology, 23 (3), 321–335. 48. Mondal, D., Sharma, M., Maiti, P. et al. (2013) Fuel intermediates, agricultural nutrients and pure water from Kappaphycus alvarezii seaweed. RSC Advances, 3 (39), 17989–17997.

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49. Ghadiryanfar, M., Rosentrater, K.A., Keyhani, A. and Omid, M. (2016) A review of macroalgae production, with potential applications in biofuels and bioenergy. Renewable and Sustainable Energy Reviews, 54 (Feb 2016), 473–481. 50. Tsubaki, S., Oono, K., Hiraoka, M. et al. (2014) Hydrolysis of green-tide forming Ulva spp. by microwave irradiation with polyoxometalate cluster. Green Chemistry, 16 (4), 2227–2233. 51. Tremblay, J.-F. (2016) Microwaving by the ton: could industrial-scale microwaves transform chemical manufacturing? Chemical & Engineering News, 94 (36), 24–25. 52. Hiraoka, M. (2012) Intensive biomass production by green seaweed Ulva. Journal of the Japan Institute of Energy, 91 (11), 1154–1160. 53. Shell-Ex Bio-Marine, Marine Bio-refinery, Newfoundland, Canada, http://www.shell-ex .com/partners.html (accessed October 10, 2016).

2 Production and Conversion of Green Macroalgae (Ulva spp.) Shuntaro Tsubaki1 , Wenrong Zhu2 and Masanori Hiraoka3 1

2.1

Department of Applied Chemistry, Graduate School of Science and Engineering Tokyo Institute of Technology, Japan 2 Graduate School of Kuroshio Science, K¯ ochi University, Japan 3 Usa Marine Biological Institute, K¯ochi University, Japan

Production of Ulva Biomass

Currently, production of seaweed biomass is frequently proposed as a useful way to both fix CO2 and produce a feedstock for bioproducts and bioenergy. Stable production of the seaweed biomass is required for industrial commercial-scale utilization. The green tide of seaweeds are also a good biomass feedstock [1]. However, the seaweed biomass provided by natural harvests or traditional ocean-based cultivation methods fluctuates seasonally and yearly due to variable environmental factors such as temperature, salinity, and nutrient limitation. In addition, recent climate change seems to be influencing abundance and distribution of seaweed populations [2]. Harvests from natural populations of green macroalga Ulva spp. are insufficient to meet their demand in Eastern Asia, although they are commonly utilized as food stuff in this region [3]. Land-based culture of seaweed is an important option for their stable production. We designed a tank culture system for

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources, First Edition. Edited by Francesca M. Kerton and Ning Yan. © 2017 John Wiley & Sons Ltd. Published 2017 by John Wiley & Sons Ltd.

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

macroalgal biomass production; based on this system, a commercial-scale plant for the production of Ulva biomass for human food was first built in Muroto, K¯ochi Prefecture, Japan, in 2004 [4]. The plant has been operating within balanced budget financing since 2009 [5]. In this section, we introduce the tank culture system and the ongoing study for more efficient mass production. 2.1.1

Land-Based Tank Culture in K¯ochi

In comparison with traditional ocean-based culture methods, land-based tank culture systems provide the opportunity to realize a stable macroalgal mass production by control of culture conditions including temperature, seawater exchange, and nutrient supply. It can solve the problems of unstable harvests, contamination of other organisms, and hard labors in the traditional ocean-based cultivation. However, investment in plant and equipment and operating cost are required. To maintain commercial interest, high biomass productivity per tank unit and reduction of energy input are indispensable. We developed three main methods for enhancing the biomass production efficiency in the tank culture system as described as follows. 2.1.1.1

“Germling Cluster” Method

Unattached culture as a free-floating form has several advantages, because harvesting is easy, less laborious, and capital-intensive, and the density of the cultures can be easily controlled at optimal levels [6]. To date, several seaweed species have been grown using this approach. Some red seaweeds such as Gracilaria tikvahiae and Chondrus crispus can vegetatively propagate through fragmentation of thalli [6, 7]. However, fragmented thalli of many species except sterile strains often reproductively mature, release spores, and consequently fall into decay and vanish. Especially in Ulva spp., such reproductive maturation frequently occurs in the fragmented thalli, making it difficult to keep their yield sustainable [7, 8]. Another serious problem, repeated vegetative propagation for a long period allows epiphytes to attach to the cultured thalli. Eventually, the growth rate of the epiphytized thalli largely declines [7]. A new “germling cluster” method was proposed for dense tank culture of seaweed in free-floating form [9]. As shown in Figure 2.1, the procedure for the production of germling clusters in the laboratory is explained as follows. 1. First, a single Ulva strain is isolated by unialgal technique using phototactic behavior of its zoids [10] and cultured in a glass vessel within an incubator. 2. The well-developed thallus of more than 10 cm length is prepared for supply of a large number of zoids.

Production and Conversion of Green Macroalgae (Ulva spp.)

21

Densely concentrated zoid suspension Light

3

4

5 2 6 1

Outdoor tank cultivation 7

8 Figure 2.1 Culture process for the production of Ulva germling clusters in the laboratory. See text for details.

3. Thallus fragments of 1–2 mm length are excised from the apical part of the well-developed thallus and transferred to a Petri dish containing seawater medium. The dish is placed at 20 ∘ C under cool white fluorescent light (100 μmol photons/m2 /s) for the required time with a 12 h:12 h, light:dark cycle. 4. Under these culture conditions, thallus fragments can synchronously produce zoids within 2–4 days. Zoids released from mature fragments are collected by their phototactic response and concentrated. 5. A large number of zoids concentrated densely are placed in a Petri dish, adjusted to a density of >104 zoids per 1 mL medium, and cultured under the condition mentioned earlier. After a few weeks, germlings grow at a

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

high density on the bottom of the dish and attach to one another to form aggregations in the form of a green mat. 6. The aggregations are scraped off the dish with a spatula and torn into numerous small clusters of germlings. 7. These germling clusters are cultured with aeration, drifting freely with the current in the vessel. 8. When they attain a length of 1 mm or more, they are ready for mass cultivation in outdoor tanks and maintained as a stock in glass bottles. In order to supply the clonal germling clusters continuously, a few subsets of the stock of germling clusters are set to the first process again. By repeating these culture processes, a large stock of clonal germling clusters can be produced. This method is characterized by (i) the use of germlings or young thalli with higher growth and lower sporulation activities compared to the aged ones; (ii) continuous supply of the germlings, of which each batch is all harvested at once without multiple harvests from the same batch, avoiding a long-term culture; (iii) permanently repeated clone culture of a targeted strain. This method removes the risk of decay by reproductive maturation, avoids heavy prolification of epiphytes, and provides genetically uniform biomass. 2.1.1.2

Deep Seawater

Temperature control is required for stable seaweed cultivation all year round. However, great energy input for cooling in summer and warming in winter is necessary to maintain constant water temperature for optimal growth of seaweed. To reduce the cost, we used deep seawater (DSW). The DSW is pumped up from a depth of over 300 m to the plant using pipelines positioned 2 km off the east coast of Muroto Cape. It has three main characteristic properties useful for seaweed cultivation: its temperature is constant around 13 ∘ C throughout the year; it contains extremely few contaminants including epiphytic organisms; and it has a high concentration of inorganic nutrients (NO3 -N; 27.0 ± 2.00 μM, PO4 -P; 1.86 ± 0.18 μM) [11]. By a continuous supply of the DSW to the culture tanks of seaweeds, these properties become profitable for cost cutting for maintaining stable temperature, prevention of contamination of epiphytic and herbivorous organisms, and reduction of additional fertilizers. Actually, without addition of fertilizers, only continuous supply of unfiltered DSW to the tanks at an exchange rate of 3 volumes per day successfully led to good Ulva growth, of which daily growth rate (DGR = [(Wt /W0 )1/t − 1] × 100, W0 = initial weight, Wt = weight after t days) was 43% on an average throughout the year (Table 2.1) [12]. 2.1.1.3

Multistep Tank System

In a multistep tank system, the tanks are placed in multistep series to save energy input to the system. Figure 2.2 shows the growth curves of Ulva cultured in a 1-ton

Production and Conversion of Green Macroalgae (Ulva spp.)

23

Table 2.1 Monthly final wet weight and daily growth rate (DGR) after 1-week culture from an initial wet weight of 100 g and temperature ranges in a 1-ton tank using deep seawater (DSW) for Ulva prolifera. Month

Final wet weight (g)

DGR (%)

Temperature (∘ C)

Jan. Feb. Mar. Apr. May Jun. Jul. Aug. Sep. Oct. Nov. Dec. Average

540 1100 1260 2620 1250 1330 1300 1400 940 1710 920 860 1270

27 41 43 60 43 45 44 46 38 50 37 36 43

8.4–12.2 11.1–15.0 10.4–14.1 13.0–16.5 14.6–15.6 14.8–16.9 15.9–18.8 16.0–23.7 16.1–20.4 16.2–21.2 11.0–14.5 9.2–15.0

Source: Hiraoka et al. 2004 [12]. Reproduced with permission of Japanese Journal of Phycology.

100,000 1t 7t

Wet weight (g)

10,000

1000

100 0

5 Days

Figure 2.2 Growth curves of Ulva prolifera cultured using 1- and 7-ton tanks. (Source: Hiraoka et al. 2004 [12]. Reproduced with permission of The Japanese Journal of Phycology.)

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

cylindrical tank (transparent polycarbonate, 1.5 m upper diameter, 0.8 m height) and a 7-ton rectangular tank (1.5 m × 5.5 m × 0.9 m) [12]. An initial 100 g-wet of Ulva increased exponentially in weight, but the growth declined as the biomass density in the tank increased and finally almost ceased. Subsequently, the same Ulva plants were transplanted to the larger 7-ton tank and rapidly grew again and showed a similar growth curve as that in the 1-ton tank. The results demonstrated that the initial mass set at about 100 g-wet per 1-ton tank volume increased by more than 10 times after 1-week culture. Figure 2.3 shows a single unit of the multistep tank system designed on the basis of those experimental data and the actual plant under operation. A set of the unit has four step tanks. An initial 10 g-wet of Ulva germling clusters prepared by laboratory culture is placed in the first outdoor 100-L tank. Every week, the cultured Ulva clusters are transferred to the 1ton × 10 1 kg 100 g

Laboratory

7ton × 10 10 kg

× 6 sets 100 L

1 ton 100 kg

stock

10 g

100 g

1 kg

Figure 2.3 A multistep tank system and commercial-scale plant for mariculture of edible Ulva ¯ actually operated in Muroto, Kochi Prefecture, Japan.

Production and Conversion of Green Macroalgae (Ulva spp.)

25

sequenced tanks with approximately 10 times larger volume. Finally, a total of 100 kg-wet biomass is harvested from the single unit every week. Six sets of the unit are provided and are set to run on consecutive days, producing 100 kg-wet of Ulva every day for 6 days, and a holiday is set every week. The annual biomass production is 30 tons in an approximately 500-ton tank volume, to which 1500 tons of DSW are supplied per day. Harvested materials from which surface seawater are removed by centrifugation are dried in cool-wind driers. The dried materials decrease to approximately 10% of the value of original wet ones in weight. The plant provides 3 tons of dry green laver per year for use as a topping for many foods, in soups, and as a confectionery coating. 2.1.2

Improvement for More Intensive Culture

In order to utilize the algal biomass as a feedstock for bioproducts, its price must be lower than its price when it is used as a food. Namely, a more intensive culture is necessary. We have been trying to look for strains with high growth rate and to prompt algal growth by addition of nutrients. 2.1.2.1

Selection of Strain

Compared with the growth rate of Ulva prolifera (cf. Table 2.1), we have isolated some Ulva strains with a very high growth of more than 100% in DGR or 0.69 per day of relative growth rate [RGR = (ln Wt − ln W0 )/t], which expressed continuously accelerating growth of algae during an exponential phase [13]. Until now, the best growing strain is Ulva meridionalis described as a new species in 2011 [14]. Originally, some fishermen cultivating U. prolifera had noticed occasional Ulva blooming in summer in the Yoshino River estuary, Tokushima Prefecture, Japan, although U. prolifera populations normally occur in winter. Therefore, we investigated this peculiar Ulva, which has a thin and branching morphology similar to that of U. prolifera. A strain of U. meridionalis was first isolated from the summer Ulva bloom and named E16 [15]. This strain grows well at high temperatures >25 ∘ C. In a preliminary experiment in an outdoor semicylindrical tank with an upper square area of 1 m × 1 m as shown in Figure 2.4, it showed RGR 0.87 day−1 in the initial exponential phase and a high productivity of approximately 60 g-dry/m2 /day when stock biomass in the tank was over 500 g-wet (Figure 2.5). 2.1.2.2

Nutrient Supply

Nitrogen is the most frequently limiting nutrient to seaweed growth, followed by phosphorus and occasionally Fe [16]. There are many reports on the effect of dissolved nitrogen and phosphorus on the growth of Ulva [17, 18]. However, few studies have examined the effect of Fe levels. We have found that additional Fe(II) accelerates the growth rate of U. prolifera cultured in an outdoor tank continuously

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Sea water input

Over flow

Figure 2.4 An experimental outdoor tank with an upper square area of 1 m × 1 m, continuously supplied with surface seawater in which nitrate and phosphate were added as adjusted to the same concentration as in DSW.

High increment phase

3000 Wet weight (g)

Exponential phase

10,000 Logarithm of wet weight (g)

4000

2000

1000

0

1000

100

10

1 0

2

4

6

8

10

Culture time (day)

0

2 4 6 8 Culture time (day)

(a)

(b)

10

Figure 2.5 Growth curve of Ulva meridionalis in the experimental outdoor tank (a) and its logarithmical representation (b).

Production and Conversion of Green Macroalgae (Ulva spp.)

27

supplied with DSW. When the Fe(II) concentration was increased from 0.07 ppb in untreated DSW to 0.3 ppb in the tank by adding a Fe elution material, the DGR of 71% in the control sample during the summer season rose by 100% [19]. Although the effect of additional Fe on U. meridionalis growth has not been completely examined, it is expected to lead to a higher productivity in U. meridionalis.

2.2

Conversion of Ulva Biomass

Chemical components of algae are less persistent than lignocellulose since they do not contain lignin and cuticle; however, the high water content and unique chemical composition of Ulva biomass make it difficult for effective production of biochemicals. The chemical compositions of U. meridionalis (filamentous type) and Ulva ohnoi (floating foliose type) are listed in Table 2.2. Ulva biomass is predominantly composed of carbohydrates such as cellulose (structural polysaccharide), starch (storage polysaccharide), and matrix polysaccharide called ulvan. The monosaccharide composition of the neutral sugars is predominantly composed of glucose, which mainly originates from starch and cellulose (Table 2.2). The contents of cellulose and starch in Ulva biomass are relatively smaller than those in lignocellulosic biomass. Rhamnose contents for the two species of Ulva studied are 25.9% and 34.2%, which originates from ulvan. Ulvan is generally made up of repeating units of rhamnose and glucuronic acid with sulfation at C-3 of rhamnose moiety [21]. Xylose, glucose, and iduronic acid are sometimes contained in ulvan as minor constituents depending on the species of Ulvales. Protein contents of U. meridionalis and U. ohnoi were relatively higher than those in terrestrial plants, attaining 24.7% and 13.1%, respectively. Table 2.2

Chemical composition of Ulva meridionalis and Ulva ohnoi.

Components

Ulva meridionalis

Chemical composition (%) Cellulose 8.6 Starch 13.0 Neutral sugar 44.9 Uronic acid 11.6 Protein 24.7 Ash 8.6 Lipid 6.5 Monosaccharide composition (%) Rhamnose 25.9 Galactose 11.0 Glucose 56.2 Xylose 4.8 Mannose 2.1

Ulva ohnoi 19.7 4.2 34.5 13.4 13.1 24.6 12.6 34.2 0.7 52.3 12.7 0.1

Source: Tsubaki et al. 2014 [20]. Reproduced with permission of Royal Society of Chemistry.

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Recently, growing numbers of studies have been focused on the production of biofuels and biochemicals from Ulva biomass due to their high potential as biomass feedstock [22–24]. The Ulva biomass can be converted into biogas [25–28], bio-oils [29–36], bioethanol [37–41], biochemicals such as lactic acid, acetone, butanol [42, 43], biochar [44], and biodiesel [45]. These products can be obtained through biological (e.g., methane, ethanol) and physicochemical (e.g., pyrolysis, microwave, hydrothermal, ultrasonication) treatments as well as chemical degradations using, for example, ionic liquid [46] and H-ZSM5 catalyst [47]. Ulva biomass is also interesting for producing value-added functional chemicals because seaweeds contain a variety of biologically active compounds such as sulfated polysaccharides, pigments, and proteins [48, 49]. Ulvan is one of the expected valuable chemicals in Ulva biomass [21, 50] and available for food application as well as medical application due to its antitumor, antioxidant, antimicrobial effects [51–55]. Degradation of ulvan further produces rhamnose and sulfated oligosaccharides [56, 57]. We have been using microwave irradiation and hydrothermal treatments to convert Ulva biomass into valuable chemicals such as ulvan and rhamnose. Microwave irradiation is capable of rapid treatment of biomass due to its high heating efficiency. Hydrothermal treatment is another important technology for treating wet biomass such as algae and food wastes [58]. Hydrothermal reactions are recognized as environmentally benign biomass treatments because water at high temperature and pressure enables hydrolysis of biomass without addition of hazardous acid catalysts due to the high ion product (KW = [H+ ][OH]) of water at high temperature. In addition, energy-consuming drying processes can be skipped by directly applying hydrothermal treatments to the wet biomass. Since microwaves can directly heat water molecules, wet algal biomass is very compatible with hydrothermal treatment using microwave energy. This section, therefore, summarizes our recent progress in the microwave-assisted hydrothermal treatments of Ulva biomass. 2.2.1

Microwave-Assisted Hydrothermal Reaction of Biomass

Microwaves are electromagnetic irradiation at frequencies between 300 MHz and 300 GHz. Microwave ovens have been widely used for cooking in domestic kitchens as well as industrial heating processes. Microwaves at 2.45 GHz constitute the Industry, Science, and Medical (ISM) band (microwave frequencies available for ISM applications) mostly used for heating devices. Microwave heating originates from the molecular motions of dipoles and ionic substances in an oscillating electromagnetic field (Figure 2.6) [59]. These heating mechanisms lead to the advantages of microwave heating processes as listed as follows: 1. Internal heating without direct contact between the heat source and the sample 2. Rapid heating rate 3. Relatively high heat efficiency

Production and Conversion of Green Macroalgae (Ulva spp.)

29

δ– δ+ Dipole rotation Figure 2.6

Ionic conduction

Mechanisms of microwave heating of aqueous solution.

4. Selective heating of highly microwave-susceptible materials 5. Easy temperature control 6. Easy design of compact and continuous systems Microwave irradiation has been applied for facilitating chemical reactions [60, 61], material processing [62], environmental engineering [63] as well as biorefinery processes such as pretreatment of lignocellulosic biomass [64, 65]. Microwave irradiation suppresses the decomposition of sugars in the conversion process of sugar-rich biomass by avoiding wall effects under hydrothermal conditions [66, 67]. This phenomenon was suggested to be due to the higher selectivity of microwaves for heating water molecules in the reaction medium. Therefore, it is effective for extracting or hydrolyzing polysaccharides for the production of useful sugars from biomass such as corn fiber, starch, and cassava pulp [68–70]. In addition, microwave irradiation permits high heating efficiency for electrolyte solutions due to the ionic conduction mechanism (Figure 2.6) [59]. In our previous studies, addition of electrolytes in the hydrothermal treatments of di- and polysaccharides was effective in reducing the reaction severity by improving the frequency factor [20, 71]. The ionic conduction mechanism was successfully applied to the hydrolysis of high-salt food waste (e.g., stones of pickled fruits) with significant reduction in the reaction temperature and molecular weight of the extracted components [72]. Based on the previous results, we are extending microwave-assisted hydrothermal reactions for extraction and hydrolysis of Ulva biomass components (Figure 2.7). Ulva biomass can produce value-added chemicals such as ulvan and rhamnose for food and medical applications, as well as starch and cellulose for bioenergy applications. In the following sections, therefore, we describe microwave-assisted extraction and hydrolysis under hydrothermal condition for efficient conversion of Ulva biomass. 2.2.2 2.2.2.1

Microwave-Assisted Conversion of Ulva Biomass Microwave-Assisted Extraction of Ulvan from Ulva Biomass

Hydrothermal extractions of ulvan from U. meridionalis and U. ohnoi were conducted as shown in Figure 2.8 [73]. Namely, the raw algae sample was

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Starch HO HO

Cellulose

OH O

HO OH O

OH O

OH O HO

Extraction

10–20%

O OH O

OH O HO

OH O HO O O HO HO

OH OH O O HO O O HO OH HO n

OH O

OH OH

OH O

10–15% Bioenergy application

Phycocolloids

Rare sugars

30–50%

Ulvan

Fast-growing Ulva spp. Extraction

+

Rhamnose OH

–

Na OOC O HO

H 3C O HO OH

O

Hydrolysis

O

H3C HO HO

OH

O OH

Food and medical application Figure 2.7 Schematic diagram of fractionation of chemical components of fast-growing Ulva biomass.

Raw algae Microwave-assisted extraction (100–180°C) Centrifugation

Extraction residue

Extract Ethanol precipitation freeze drying Sulfated phycocolloids

Figure 2.8 The schematic diagram of microwave-assisted extraction of ulvan.

suspended in water in a high-pressure tolerant Teflon vessel and then microwaved using a multimode microwave reactor START-D (Milestone Inc.). The reaction temperature was controlled with proportional, integral, and differential (PID) controller equipped with a thermocouple thermometer. Extraction was conducted at 100–180 ∘ C for 10 min with 4 min of heating-up time, and then, the reactor was cooled in an ice bath. The extract was obtained by centrifuging three times. The crude sulfated ulvan was separated by ethanol precipitation and freeze-dried. Figure 2.9a shows the yields of ulvan from U. meridionalis and U. ohnoi, and Figure 2.9b shows the viscosity of the ulvan under the same conditions. The 30–40% of crude ulvan could be extracted within 14 min of microwave irradiation [73]. Ulvan of U. ohnoi exhibited higher viscosity compared to U. meridionalis,

Production and Conversion of Green Macroalgae (Ulva spp.)

31

5

100 80

Ulva ohnoi

Viscosity η (mPa S)

Polysaccharide yield (%)

Ulva meridionalis

60 40

Figure 2.9 ulvan.

3 2 1

20 0

4

0 100

120

140

160

180

100

120

140

160

Extraction temperature (°C)

Extraction temperature (°C)

(a)

(b)

180

Effects of microwave-assisted extraction for (a) yields of ulvan and (b) viscosity of

Table 2.3 Advantages of the microwave-assisted extraction of ulvan. Microwave-assisted extraction

Conventional extraction External heating

Extraction time Extraction temperature Extraction agent

Direct and internal heating 10–30 min 100–150 ∘ C None

Sugar stability

High

Purification

None

Heating property

≤3 h 90–100 ∘ C Chelating agent (e.g., ammonium oxalate, sodium hydroxide) Unexpected decomposition by the wall effect Dialysis, neutralization

and this gradually decreased with increases in the temperature due to degradation of the ulvan. Since ulvans exhibits unique rheological properties, they could be utilized as ingredients in food and beverages [21]. The biological effects of ulvans further add higher values to these unutilized seaweeds [48, 50]. Table 2.3 summarizes the advantages of the microwave-assisted extraction compared to the conventional methods. Microwave-assisted extraction drastically reduces the duration of extraction since microwave permits very high heating rate by direct heating of the mixture of Ulva biomass and water. Especially, the ionic conduction of the electrolyte components in Ulva biomass can improve microwave susceptibility of the extraction system. The extraction under hydrothermal conditions can produce ulvan without the addition of chelating agents; therefore, the process does not require time-consuming purification processes such as dialysis. In addition, degradation of sugars could be suppressed by microwave treatment since the extraction process is complete within several tens of minutes.

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2.2.2.2

Microwave-Assisted Hydrolysis of Ulva Biomass

Hydrolysis of ulvan generates oligosaccharides, neutral sugars, and uronic acids [20]. Rhamnose is one of the promising high-value-added monosaccharides, which is available for food ingredients [48]. The hydrolysis of U. meridionalis was, therefore, conducted by using microwave irradiation with the addition of polyoxometalate cluster catalysts (POMs). POMs were used as efficient and less corrosive acid catalysts and alternatives to the conventionally used mineral acids (e.g., H2 SO4 , HCl). Since POMs exhibit very strong acidity and oxidation power [74], they have been applied for hydrolysis of crystalline cellulose [75], delignification process for the paper industry [76], and oxidation of cellulose and hemicellulose [77]. We have previously reported that POMs effectively absorb microwave energy especially when compared with more generally used strong ion-exchange resin catalysts, and this reduces the microwave energy consumption required for the reaction (Figure 2.10) [78]. The catalyses of three types of POMs (phosphotungstic, silicotungstic, and phosphomolybdic acids) were investigated for the hydrolysis of U. meridionalis. Microwave irradiation was conducted at temperatures between 120 and 220 ∘ C for 10 min using a multimode microwave reactor as described in Section 2.2.2.1. The yield of reducing sugar generated after microwave reaction was determined by the dinitrosalicylic acid (DNS) method. Production of rhamnose was facilitated at lower temperatures below 160 ∘ C, attaining around 35–50% of the total neutral sugar. Generation of uronic acid was also facilitated at the same temperature, indicating that selective hydrolysis of ulvan occurs at mild temperatures. On the other hand, the production of glucose became prominent at higher reaction temperatures due to hydrolysis of starch and cellulose. Phosphotungstic and silicotungstic acids produced higher reducing sugar yields compared to H2 SO4 and HCl (Figure 2.11). Increasing the concentration of 120

160 °C 10 min

Microwave energy consumption (kJ)

100 80 60 40 20 0

H2O

PW

SiW

PMo

Dowex Amberlyst

POM Figure 2.10 Microwave susceptibility of polyoxometalate clusters and strong ion-exchange resins. (Source: Tsubaki et al. 2013 [78]. Reproduced with permission of Elsevier.)

Production and Conversion of Green Macroalgae (Ulva spp.)

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350

Sugar yield (mg/g)

300 250 200 150 100 50 0 PW

SiW

PMo

HCl

H2SO4

POM Figure 2.11 Sugar yield after hydrolysis of Ulva meridionalis using polyoxometalate clusters, hydrochloric acid, and sulfuric acid at 50 mM. (Source: Tsubaki et al. 2014 [20]. Reproduced with permission of Royal Society of Chemistry.)

phosphotungstic acid increased the reducing sugar yield and attained a maximum value of 34 wt% at 50 mM. Effects of microwave heating were, then, compared with external heating using an induction oven (SSN-400) with an identical thermal history. Microwave heating produced higher yields of neutral sugars and uronic acids compared to the induction oven, and this was most likely due to the higher stability of sugars by avoiding the wall effect (Figure 2.12). The results suggested that the combination of microwave irradiation and POMs 500

Sugar yield (mg/g)

400 300 200 100 0 Microwave heating Uronic acid

Induction heating Neutral sugar

Figure 2.12 The yields of uronic acid and neutral sugar from Ulva meridionalis after microwave and induction heating treatments. (Source: Tsubaki et al. 2014 [20]. Reproduced with permission of Royal Society of Chemistry.)

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

synergistically facilitated hydrolysis of Ulva biomass to convert them into monosaccharides abundantly containing rhamnose. 2.2.2.3

Microwave Susceptibility of Ulvan

Analyses of the dielectric property of the biomass substrates are very important for understanding the microwave susceptibility of the biomass–water mixture and designing efficient microwave processes [79]. The dielectric property of ulvan in water was, therefore, measured to reveal the microwave heating mechanism of ulvan related to their chemical structure, especially focusing on the sulfate groups. First, sulfate groups of ulvan were removed by a solvolytic reaction using a mixed solution of dimethylsulfoxide, pyridine, and methanol (Figure 2.13a). The degree of desulfation was analyzed by using FT-IR spectroscopy. The band for the stretching vibration of the sulfate groups at 1250 cm−1 significantly decreased after solvolysis, indicating the efficient removal of sulfate groups from ulvan (Figure 2.13b). The dielectric properties of native and desulfated ulvan were measured using the coaxial probe method and vector network analyzer in a frequency range between 100 MHz and 10 GHz. The dielectric properties of ulvan were expressed using equation 2.1. (2.1) 𝜀∗ = 𝜀′ − j𝜀′′ where the real and imaginary parts represent relative permittivity (𝜀′ ) and dielectric loss (𝜀′′ ), respectively. Dielectric loss tangent (tan 𝛿) at 2.45 GHz was further calculated using equation 2.2. 𝜀′′ (2.2) tan 𝛿 = ′ 𝜀 The imaginary part (𝜀′′ ) of the complex dielectric constant is shown in Figure 2.14. The native ulvan exhibited U-shaped dielectric spectra at both 40 and 80 ∘ C. The 𝜀′′ at lower frequency was due to conduction loss of electrolytes. Dusulfation of ulvan Ulvan Ion exchange (Dowex 50 x8, H+ form) H+ from phycocolloids DMSO ∙ Pyridine ∙ methanol (100 °C, 2 h) Dialysis ∙ freeze drying

Absorbance

Phycocolloids

Native

Desulfated 4000 3500 3000 2500 2000 1500 1000 400

Desulfated phycocolloids

(a)

Wavenumber (cm–1) (b)

Figure 2.13 Desulfation of ulvan from Ulva meridionalis. (a) Desulfation procedure of ulvan. (b) IR spectra of native and desulfated ulvans. (Source: Tsubaki et al. 2014. Reproduced with permission of Elsevier.)

Production and Conversion of Green Macroalgae (Ulva spp.)

200

H+-form Native Desulfated Water

100

80 °C H+-form Native Desulfated Water

150 Loss factor

150 Loss factor

200

40 °C

50

35

100 50

0 0.1

0 0.1

1.0 10 Microwave frequency (GHZ) (a)

1.0 10 Microwave frequency (GHZ) (b)

Figure 2.14 Dielectric spectra of ulvan in water (0.5 wt%) at a frequency range of 100 MHz– 10 GHz at (a) 40 ∘ C and (b) 80 ∘ C. (Source: Tsubaki et al. 2014. Reproduced with permission of Elsevier.)

Dielectric loss tangent (tan δ) – Conductivity (σ)

Dielectric loss tangent (tan δ) – pH 0.5

0.4

0.4

0.3

0.3

tan δ

tan δ

0.5

0.2

0.1

0.1 0

0.2

0 0

1

2 3 σ (S/m) (a)

4

5

0

2

4

6

8

10

pH (b)

Figure 2.15 The correlations of dielectric loss tangent of algal polysaccharides (carrageenans and alginates) and (a) conductivity and (b) pH. (Source: Tsubaki et al. 2015. Reproduced with permission of Elsevier.)

On the other hand, the 𝜀′′ at higher frequency was due to dielectric loss of water. Increases in the temperature from 40 to 80 ∘ C increased 𝜀′′ at lower frequencies because of the elevated ionic conduction of the electrolytes. Desulfation of ulvan led to a decrease in the conduction loss, suggesting that the sulfated groups were associated with ionic conduction. The counter cation was, then, changed from Na+ -form (native) to H+ -form by passing through a suitable cation-exchange resin (DOWEX 50 × 8). The H+ -form ulvan exhibited significantly higher conduction

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loss compared to the Na+ -form. This phenomenon could be due to the high ionic conduction of H+ by a proton relay mechanism. Similarly to ulvan, carrageen and sodium alginates from red and brown algae also exhibited significant conduction loss at a lower frequency [80]. The correlation among dielectric loss tangent (tan 𝛿) at 2.45 GHz, conductivity, and pH is shown in Figure 2.15. Tan 𝛿 of carrageen and sodium alginates exhibited strong correlation with the conductivity of the solution. The prominent increase in the conduction loss was observed at pH values below 2. The results demonstrated that the ionic conduction of acidic groups and their counter cations were strongly associated with the dielectric property of the algal polysaccharide–water system.

2.3

Conclusions

Fast-growing Ulva biomass is one of the most promising groups of algal biomass for fixing CO2 and producing biochemicals and biofuels. This chapter reviewed our recent advances in the production and conversion technologies for efficient utilization of Ulva biomass. The tank culture system of Ulva biomass permits constant production of biomass at a high rate through a whole year. At the same time, efficient conversion of Ulva biomass can be achieved by a combination of microwave and hydrothermal treatments to produce value-added biochemicals such as ulvan and rhamnose in an environmentally benign way. In addition, starch and cellulose in Ulva biomass can be utilized for bioenergy applications through ethanol fermentation or hydrothermal liquefaction treatments. The continuous-flow-type microwave system [65] will further enable scaling-up of the biorefinery process with higher efficiency in the future. A combination of production and conversion processes of Ulva biomass will be suitable for territorially distributed biorefinery systems in rural areas by providing a new bio-based industry.

References 1. Smetacek, V. and Zingone, A. (2013) Green and golden seaweed tides on the rise. Nature, 504, 84–88. 2. Tanaka, K., Taino, S., Haraguchi, H. et al. (2012) Warming off southwestern Japan linked to distributional shifts of subtidal canopy-forming seaweeds. Ecology and Evolution, 2 (11), 2854–2865. 3. Critchley, A.T. and Ohno, M. (eds) (1993) Seaweed Cultivation and Marine Ranching, Kanagawa International Fisheries Training Center, Japan International Cooperation Agency, Yokosuka. 4. Hiraoka, M. and Oka, N. (2006) Mariculture of seaweeds based on the new “germling cluster method” and utilizing deep seawater in Japan, in Advances in Seaweed Cultivation and Utilization in Asia (eds P. Siew-Moi, A.T. Critchley and P.O. Ang Jr.,), University of Malaya Maritime Research Centre, Kuala Lumpur, pp. 53–59.

Production and Conversion of Green Macroalgae (Ulva spp.)

37

5. Hiraoka, M. (2012) Intensive biomass production by green seaweed Ulva. Journal of the Japan Institute of Energy, 91, 1154–1160. 6. Schramm, W. (1991) Cultivation of unattached seaweeds, in Seaweed Resources in Europe: Uses and Potential (eds M.D. Guiry and G. Blunden), John Wiley & Sons, Ltd, Chichester, pp. 379–408. 7. Hanisak, M.D. (1987) Cultivation of Gracilaria and other macroalgae in Florida for energy production, in Seaweed Cultivation for Renewable Resource, Developments in Aquaculture and Fisheries Science, vol. 16 (eds K.T. Bird and P.H. Benson), Elsevier Science Publishers, Amsterdam, pp. 191–218. 8. DeBusk, T.A., Blakeslee, M. and Ryther, J.H. (1986) Studies on the outdoor cultivation of Ulva lactuca L. Botanica Marina, 29 (5), 381–386. 9. Hiraoka, M. and Oka, N. (2008) Tank cultivation of Ulva prolifera in deep seawater using a new “germling cluster” method. Journal of Applied Phycology, 20, 97–102. 10. Kawai, H., Monomura, T. and Okuda, K. (2005) Isolation and purification techniques for macroalgae, in Algal Culturing Techniques (ed. R.A. Andersen), Elsevier Academic Press, Burlington, pp. 133–143. 11. Sumida, T., Tamura, E. and Kawakita, H. (2001) Characteristics of Muroto deep seawater. Bulletin of the Society of Sea Water Science, Japan, 55, 158–165. 12. Hiraoka, M., Ohno, M., Dan, A. et al. (2004) Utilization of deep seawater for the mariculture of seaweeds in Japan. The Japanese Journal of Phycology, 52 (Supplement), 215–219. 13. Fogg, G.E. and Thake, B. (1987) Algal Cultures and Phycoplankton Ecology, 3rd edn, The University of Wisconsin Press, Ltd, London. 14. Horimoto, R., Masakiyo, Y., Ichihara, K. et al. (2011) Enteromorpha-like Ulva (Ulvophyceae, Chlorophyta) growing in the Todoroki River, Ishigaki Island, Japan, with special reference to Ulva meridionalis Horimoto et Shimada, sp. nov. Bulletin of the National Museum of Nature and Science. Series B, Botany, 37 (4), 155–167. 15. Shimada, S., Yokoyama, N., Arai, S. et al. (2008) Phylogeography of the genus Ulva (Ulvophyceae, Chlorophyta), with special reference to the Japanese freshwater and brackish taxa. Journal of Applied Phycology, 20, 979–989. 16. Hurd, C.L., Harrison, P.J., Bischof, K. et al. (eds) (2014) Seaweed Ecology and Physiology, 2nd edn, Cambridge University Press, Cambridge. 17. Björnsäter, B.R. and Wheeler, P.A. (1990) Effect of nitrogen and phosphorus supply on growth and tissue composition of Ulva fenestrata and Enteromorpha intestinalis (Ulvales, Chlorophyta). Journal of Phycology, 26, 603–611. 18. Luo, M.B., Liu, F. and Xu, Z.L. (2012) Growth and nutrient uptake capacity of two co-occurring species, Ulva prolifera and Ulva linza. Aquatic Botany, 100, 18–24. 19. Oka, N., Sumida, T., Hiraoka, M. et al. (2009) Effects of Fe(II) for tank cultivation of Ulva prolifera in deep seawater. Algal Resources, 1, 63–66. 20. Tsubaki, S., Oono, K., Hiraoka, M. et al. (2014) Hydrolysis of green-tide forming Ulva spp. by microwave irradiation with polyoxometalate cluster. Green Chemistry, 16, 2227–2233. 21. Lahaye, M. and Robic, A. (2007) Structure and functional properties of ulvan, a polysaccharide from green seaweeds. Biomacromolecules, 8, 1765–1774. 22. Wei, N., Quaterman, J. and Jin, Y.-S. (2012) Marine macroalgae: an untapped resource for producing fuels and chemicals. Trends in Biotechnology, 31, 70–77. 23. Coelho, M.S., Barbosa, F.G. and de Souza, M.R.A.Z. (2014) The scientometric research on macroalgal biomass as a source of biofuel feedstock. Algal Research, 6, 132–138.

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24. Herrmann, C., FitzGerald, J., O’Shea, R. et al. (2015) Ensiling of seaweed for a seaweed biofuel industry. Bioresource Technology, 196, 301–313. 25. Matsui, T. and Koike, Y. (2010) Methane fermentation of a mixture of seaweed and milk at a pilot-scale plant. Journal of Bioscience and Bioengineering, 110, 558–563. 26. Bruhn, A., Dahl, J., Nielsen, H.B. et al. (2011) Bioenergy potential of Ulva lactuca: biomass yield, methane production and combustion. Bioresource Technology, 102, 2595–2604. 27. Allen, E., Browne, J., Hynes, S. et al. (2013) The potential of algae blooms to produce renewable gaseous fuel. Waste Management, 33, 2425–2433. 28. Marquez, G.P.B., Takeuchi, H. and Hasegawa, T. (2015) Biogas production of biologically and chemically-pretreated seaweed, Ulva spp., under different conditions: freshwater and thalassic. Journal of the Japan Institute of Energy, 94, 1066–1073. 29. Zhuang, Y., Guo, J., Chen, L. et al. (2012) Microwave-assisted direct liquefaction of Ulva prolifera for bio-oil production by acid catalysis. Bioresource Technology, 116, 133–139. 30. Liu, J., Zhunag, Y., Li, Y. et al. (2013) Optimizing the conditions for the microwave-assisted direct liquefaction of Ulva prolifera for bio-oil production using response surface methodology. Energy, 60, 69–76. 31. Neveux, N., Yuen, A.K.L., Jazrawi, C. et al. (2014) Biocrude yield and productivity from the hydrothermal liquefaction of marine and freshwater green macroalgae. Bioresource Technology, 155, 334–341. 32. Li, J., Wang, G., Chen, M. et al. (2014) Deoxy-liquefaction of three different species of macroalgae to high-quality liquid oil. Bioresource Technology, 169, 110–118. 33. Singh, R., Balagurumurthy, B. and Bhaskar, T. (2015a) Hydrothermal liquefaction of macro algae: effect of feedstock composition. Fuel, 146, 69–74. 34. Singh, R., Bhaskar, T. and Balagurumurthy, B. (2015b) Effect of solvent on the hydrothermal liquefaction of macro algae Ulva fasciata. Process Safety and Environmental Protection, 93, 154–160. 35. Rojas-Pérez, A., Diaz-Diesta, D., Frias-Flores, C.B. et al. (2015) Catalytic effect of ultrananocrystalline Fe3 O4 on algal bio-crude production via HTL process. Nanoscale, 7, 17664–17671. 36. Graz, Y., Bostyn, S., Escot Bocanegra, P. et al. (2016) Hydrothermal conversion of Ulva macro algae in supercritical water. The Journal of Supercritical Fluids, 107, 182–188. 37. Isa, A., Mishima, Y., Takimura, O. et al. (2009) Preliminary study on ethanol production by using macro green algae. Journal of the Japan Institute of Energy, 88, 912–917. 38. Kim, N.J., Li, H., Jung, K. et al. (2011) Ethanol production from marine algal hydrolysates using Eschericha coli KO11. Bioresource Technology, 102, 7466–7469. 39. Yanagida, M., Kawai, S. and Murata, K. (2013) Strategies for the production of high concentrations of bioethanol from seaweeds. Production of high concentrations of bioethanol from seaweeds. Bioengineered, 4, 224–235. 40. Korzen, L., Pulidiani, I.M., Israel, A. et al. (2015) Single step production of bioethanol from the seaweed Ulva rigida using sonication. RSC Advances, 5, 16223. 41. Trivedi, N., Reddy, C.R.K., Radulovich, R. et al. (2015) Solid state fermentation (SSF)-derived cellulase for saccharification of the green seaweed Ulva for bioethanol production. Algal Research, 9, 48–54. 42. Hwang, H.J., Kim, S.M., Chang, J.H. et al. (2012) Lactic acid production from seaweed hydrolysate of Enteromorpha prolifera (Chlorophyta). Journal of Applied Phycology, 24, 935–940.

Production and Conversion of Green Macroalgae (Ulva spp.)

39

43. van der Wal, H., Sperber, B.L., Houweling-Tan, B. et al. (2013) Production of acetone, butanol, and ethanol from biomass of the green seaweed Ulva lactuca. Bioresource Technology, 128, 431–437. 44. David, A.R. and de Nys, R. (2016) The effects of feedstock pre-treatment and pyrolysis temperature on the production of biochar from the green seaweed Ulva. Journal of Environmental Management, 169, 253–260. 45. Suganya, T., Kasirajan, R. and Renganathan, S. (2014) Ultrasound-enhanced rapid in situ transesterification of marine macroalgae Enteromorpha compressa for biodiesel production. Bioresource Technology, 156, 283–290. 46. Pezora-Conte, R., Leyton, A., Anugwom, I. et al. (2015) Decomposition of the green alga Ulva rigida in ionic liquids: closing the mass balance. Algal Research, 12, 262–273. 47. Lorenzetti, C., Conti, R., Fabbri, D. et al. (2016) A comparative study on the catalytic effect of H-ZSM5 on upgrading of pyrolysis vapors derived from lignocellulosic and proteinaceous biomass. Fuel, 166, 446–452. 48. Holdt, S.L. and Kraan, S. (2011) Bioactive compounds in seaweed; functional food applications and legislation. Journal of Applied Phycology, 23, 543–597. 49. Jiao, G., Yu, G., Zhang, J. et al. (2011) Chemical structure and bioactivities of sulfated polysaccharides from marine algae. Marine Drugs, 9, 196–223. 50. Alves, A., Sousa, R.A. and Reis, R.L. (2013) A practical perspective on ulvan extracted from green algae. Journal of Applied Phycology, 25, 407–424. 51. Peasura, N., Laohakunjit, N., Kerdchoechuen, O. et al. (2015) Characteristics and antioxidant of Ulva intestinalis sulphated polysaccharides extracted with different solvents. International Journal of Biological Macromolecules, 81, 912–919. 52. Dussault, D., Vu, K.D., Vanach, T. et al. (2016) Antimicrobial effects of marine algal extracts and cyanobacterial pure compounds against five foodborne pathogens. Food Chemistry, 199, 114–118. 53. Shao, P., Shao, J., Jiang, Y. et al. (2016) Influences of Ulva fasciata polysaccharide on the rheology and stabilization of cinnamaldehyde emulsions. Carbohydrate Polymers, 135, 27–34. 54. Tabarsa, M., Lee, S.-J. and You, S.G. (2012) Structural analysis of immunostimulating sulfated polysaccharides from Ulva pertusa. Carbohydrate Research, 361, 141–147. 55. Ahmed, O.M. and Ahmed, R.R. (2014) Anti-proliferative and apoptotic efficacies of ulvan polysaccharides against different types of carcinoma cells in vitro and in vivo. Journal of Cancer Science & Therapy, 6, 202–208. 56. Costa, C., Alves, A., Pinto, P.R. et al. (2012) Characterization of ulvan extracts to assess the effect of different steps in the extraction procedure. Carbohydrate Polymers, 88, 537–546. 57. Coste, O., Malta, E.J., López, J.C. et al. (2015) Production of sulfated oligosaccharides from the seaweed Ulva sp. using a new ulvan-degrading enzymatic bacterial crude extract. Algal Research, 10, 224–231. 58. Biller, P. and Ross, A.B. (2012) Hydrothermal processing of algal biomass for the production of biofuels and chemicals. Biofuels, 3, 603–623. 59. Gabriel, C., Gabriel, S., Grant, E. et al. (1998) Dielectric parameters relevant to microwave dielectric heating. Chemical Society Reviews, 27, 213–223. 60. Kappe, C.O., Dallinger, D. and Murphee, S.S. (eds) (2009) Practical Microwave Synthesis for Organic Chemists, Wiley-VCH Verlag GmbH & Co., KGaA, Weinheim. 61. Loupy, A. (2006) Microwaves in Organic Synthesis, Wiley-VCH Verlag GmbH & Co., KGaA, Weinheim.

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62. Bykov, Y.V., Rybakov, K.I. and Semenov, V.E. (2002) High-temperature microwave processing of materials. Journal of Physics D: Applied Physics, 34, R55–R75. 63. Jones, D.A., Lelyveld, S.D., Mavrofidis, S.W. et al. (2002) Microwave heating applications in environmental engineering – a review. Resources, Conservation and Recycling, 34, 75–90. 64. Azuma, J., Tanaka, F. and Koshijima, T. (1984) Microwave irradiation of lignocellulosic materials I. Enzymatic susceptibility of microwave-irradiated woody plants. Mokuzai Gakkaishi, 30, 501–509. 65. Tsubaki, S., Azuma, J., Yoshimura, T. et al. (2016) Microwave-induced biomass fractionation, in Biomass Fractionation Technologies for a Lignocellulosic Feedstock Based Biorefinery, 1st edn edn (ed. S. Mussatto), Elsevier, Amsterdam, pp. 103–126. 66. Tsubaki, S., Oono, K., Onda, A. et al. (2012) Microwave-assisted hydrothermal hydrolysis of cellobiose and effects of additions of halide salts. Bioresource Technology, 123, 703–706. 67. Tsubaki, S., Oono, K., Onda, A. et al. (2013) Comparative decomposition kinetics of neutral monosaccharides by microwave and induction heating treatments. Carbohydrate Research, 375, 1–4. 68. Yoshida, T., Tsubaki, S., Teramoto, Y. et al. (2010) Optimization of microwave-assisted extraction of carbohydrates from industrial waste of corn starch production using response surface methodology. Bioresource Technology, 101, 7820–7826. 69. Matsumoto, A., Tsubaki, S., Sakamoto, M. et al. (2011) A novel saccharification method of starch using microwave irradiation with addition of activated carbon. Bioresource Technology, 102, 3985–3988. 70. Hermiati, E., Azuma, J., Tsubaki, S. et al. (2012) Improvement of microwave-assisted hydrolysis of cassava pulp and tapioca flour by addition of activated carbon. Carbohydrate Polymers, 87, 939–942. 71. Tsubaki, S., Oono, S., Onda, A. et al. (2016) Effects of ionic conduction on hydrothermal hydrolysis of corn starch and crystalline cellulose induced by microwave irradiation. Carbohydrate Polymers, 10, 594–599. 72. Tsubaki, S., Sakamoto, M. and Azuma, J. (2010) Microwave-assisted extraction of phenolic compounds from tea residues under microwave autohydrolytic condition. Food Chemistry, 123, 1255–1258. 73. Tsubaki, S., Oono, K., Hiraoka, M. et al. (2016) Microwave-assisted hydrothermal extraction of sulfated polysaccharides from Ulva spp. and Monostroma latissimum. Food Chemistry, 210, 311–316. 74. Kozhevnikov, I.V. (1998) Catalysis by heteropoly acids and multicomponent polyoxometalates in liquid-phase reactions. Chemical Reviews, 98, 171–198. 75. Shimidzu, K.I., Furukawa, H., Kobayashi, N. et al. (2009) Effects of Brønsted and Lewis acidities on activity and selectivity of heteropoly acid-based catalysts for hydrolysis of cellobiose and cellulose. Green Chemistry, 11, 1627–1632. 76. Weinstock, I.A., Atalla, R.H., Reiner, R.S. et al. (1997) A new environmentally benign technology for transforming wood pulp into paper, engineering polyoxometalates as catalysts for multiple processes. Journal of Molecular Catalysis A: Chemical, 116, 59–84. 77. Sarma, B.B. and Neumann, R. (2014) Polyoxometalate-mediated electron transfer-oxygen transfer oxidation of cellulose and hemicellulose to synthesis gas. Nature Communications, 5, 4621. 78. Tsubaki, S., Oono, K., Ueda, T. et al. (2013) Microwave-assisted hydrolysis of polysaccharides over polyoxometalate clusters. Bioresource Technology, 144, 67–73.

Production and Conversion of Green Macroalgae (Ulva spp.)

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79. Tsubaki, S., Hiraoka, M., Hadano, S. et al. (2014) Functional group dependent dielectric properties of sulfated hydrocolloids extracted from green macroalgal biomass. Carbohydrate Polymers, 107, 192–197. 80. Tsubaki, S., Hiraoka, M., Hadano, S. et al. (2015) Effects of acidic functional groups on dielectric properties of sodium alginates and carrageenans in water. Carbohydrate Polymers, 115, 78–87.

3 A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels: Challenges and Potentials Ravi S. Baghel1,2 , Vaibhav A. Mantri1,2 and C.R.K. Reddy1,2 1 Marine

Biotechnology and Ecology Division, CSIR – Central Salt & Marine Chemicals Research Institute, India 2 Academy of Scientific and Innovative Research (AcSIR), Central Salt & Marine Chemicals Research Institute, India

3.1

Introduction

While the world is eagerly looking at leveraging the potential of oceans for food and energy security, marine macroalgae gain significant importance due to their ubiquitous occurrence, abundant availability and high productivity together with sustainable farming technologies. They have been a part of human life from time immemorial, and their utilization during prehistoric civilizations from various places was established from archaeological evidence gathered across many countries including Costa Rica, Japan, China and Egypt [1]. Macroalgae – the term coined for representing heterogeneous artificial group – are lower members of the plant kingdom having polyphyletic origin and different evolutionary Fuels, Chemicals and Materials from the Oceans and Aquatic Sources, First Edition. Edited by Francesca M. Kerton and Ning Yan. © 2017 John Wiley & Sons Ltd. Published 2017 by John Wiley & Sons Ltd.

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lines. They are conventionally classified into three divisions primarily based on pigment composition – Rhodophyta (red algae), Phaeophyta (brown algae) and Chlorophyta (green algae) – and partly derived from reserve storage product. The two most important classes of marine red algae are the Bangiophyceae and the Florideophyceae, which represent 161 and 6224 species respectively, while over 1800 species of brown algae are reported in Phaeophyceae, the green algae are represented by Chlorophyceae and Ulvophyceae enumerating 1610 and 3046 species, respectively (cf. [2]). Marine macroalgae have developed specific metabolic pathways in order to survive in harsh saline environments where fluctuations are recurrent due to tidal rhythms. The vast diversity of algal taxa provides ample opportunities for bioprospection of this valuable resource. Further, they have been the only sources of certain structural sulfated polysaccharides such as agar, agarose, carrageenan, alginate and ulvan that have been industrially exploited for many years. Their primary and secondary metabolites are also industrially lucrative and have niche markets in speciality chemicals such as pigments and flavouring agents. Besides industrial utilization, they also play a crucial role in sustaining the productivity of marine ecosystems. For example, the dense underwater canopy of kelp offers breeding and nursery ground for juvenile fish and other commercially valuable aquatic organisms, thus providing ecosystem service. The past decade witnessed a steady rise in scientific, industrial and consumer demand for developing commodity products from marine macroalgae, opening up several business opportunities. Although still produced on a modest scale – when compared to global aquaculture industry – the level of commercial harvesting of marine macroalgae has achieved a new milestone with 25 million tonnes per year production (95.6% accounts to farming) with a market value of over US$ 7.4 billion [3]. Being a photosynthetic organism, by converting light energy and CO2 into a storage product, commercial farming of marine algae acts as a carbon sink. This chapter provides an overview of recent advancements made in the utilization of marine macroalgae for food, chemicals and fuels along with challenges and potentials.

3.2

Macroalgal Feedstock for Chemicals

Marine macroalgae are utilized worldwide for a variety of chemicals, namely polysaccharides, pigments, minerals, lipids, proteins, amino acids and phenolic compounds. However, storage of structural carbohydrates accounts for their major industrially exploited products. These are colloidal chemicals and generally form an intermediate state between a solution and a suspension. Of these, agar, carrageenan and alginates assume high commercial significance in the global hydrocolloid market due to their excellent gelling, stabilizing and emulsifying characteristics along with high viscosity properties [4]. The processed food industry is the primary market for marine algal hydrocolloids, whereas personal

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care, cosmetics and pharmaceuticals are the emerging areas which could potentially offer better profit margins. Phycocolloid-yielding marine algal farming has almost doubled its market value over the past decade, and currently, marine algal hydrocolloid production is a US$ 1 billion industry [4]. The methods used in this area have been constantly improved with respect to yield and gel strength using either alkali, acid or enzyme treatment to extract native polysaccharides from marine macroalgae [5–8]. These techniques are more economical, efficient and greener than conventional batch processing, due to reduced processing time, conserving water and reactants. Nevertheless, it is interesting to note that only between 13.27% and 27.89% marine algal biomass is being industrially used for the production of polysaccharides [4]. The majority of algae harvested worldwide are used directly as food, as described in Chapter 2, and a smaller quantity as fertilizer. The drawback of currently existing technologies for polysaccharide isolation lies in not utilizing the feedstock to its full potential, which affords scope for developing applications to use the spent or unutilized fraction. Methods enabling multiple-product recovery from marine algal biomass for economic gain need immediate attention.

3.3

Marine Macroalgae as a Biorefinery Feedstock

Efficient integration of technologies is a key determinant for the sustainable utilization of biological resources. The biorefinery concept is analogous to petrochemical refineries and thus takes advantage of the differences in biomass composition to maximize the value derived from the same. With the production and use of biofuels gaining worldwide attention, the concept of obtaining multiple products through a biorefinery has gained momentum. The production of several low-volume, high-value chemicals ensures economic viability to biofuel production. Marine macroalgae are considered to be excellent candidates for biofuel production due to their prolific growth rates, ease of farming and ability to sequester carbon due to higher efficiency of photosynthesis [9]. These inherent advantages make them a preferred choice compared to first- and second-generation biofuel biomass. Although significant technical advancements have been made in biofuel production from this marine renewable resource at an R&D level, their commercial production is yet to start [10]. The diverse biochemical components of marine macroalgae (such as pigments, lipid, mineral, protein, cellulose and phycocollides) have recently given impetus to biomass valorization in which these can be segregated sequentially along with fuel, which is termed the ‘integrated biorefinery approach’. Almost all studies till date used the entire algal biomass for biofuel production, which overlooks the well-established hydrocolloid market and value therein. Therefore, utilizing marine macroalgal feedstock for the recovery of different high-value commodity chemicals in an integrated manner and conversion of only a selective fraction into biofuels are desirable.

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The first attempt at obtaining more than one value-added product from marine macroalgae Kappaphycus alvarezii led to the integrated production of algal sap [plant biostimulant] and granules [rich in carrageenan] [11]. A subsequent innovation has demonstrated the feasibility of obtaining algal sap and ethanol from fresh biomass of K. alvarezii [12]. The ethanol has also been produced by biorefining of Gracilaria verrucosa cellulose along with agar [13]. Niu et al. [14] attempted at segregated pigment and agar production from Gracilaria lemaneiformis. Additionally, phycobiliproteins and bio-oil have been produced in an integrated manner from Gracilaria gracilis [15]. However, in the majority of cases, at the most two products were obtained from marine algal biomass, thus leaving the opportunity for further improvement towards economic benefits of this sort of marine biorefining. A stand-alone integrated scheme for the production of 5-hydroxymethyl furfural (HMF) and potassium sulfate (K2 SO4 ) and pure water has been developed from fresh biomass of K. alvarezii [16]. The agarophytic biomass, including 15 taxa, was characterized to quantify pigments, lipids, minerals, protein agar and cellulose from the biorefinery context [17]. In yet another attempt, a tractable integrated process has been developed for sequential extraction of pigments, lipid, agar, minerals and energy-dense substrate (cellulose) from Gelidiella acerosa, Gelidium pusillum and Gracilaria dura. The distinct advantage of this process over direct extraction is the improved quality of agar (1.5–3-fold increase in gel strength) without alkali and acid pretreatment of the sample coupled with elimination of residue and the reduction in chemical usage in cellulose extraction by up to 85% [18, 19]. In a related work, a method of sequential recovery of four economically important fractions – mineral-rich liquid extract (MRLE), lipid, ulvan, and cellulose from Ulva fasciata – was reported recently [20]. The sequential concentration of residual biomass by successive extraction steps reduces reagent demand in the downstream extraction and processing of fractions. These yields were comparable to those obtained by direct processing of the individual components from primary biomass. The integration of ethanol production and chemical feedstock recovery from macroalgal biomass could substantially enhance the sustainability of marine biomass use. The biorefinery is gaining momentum, but to push this futuristic idea forward and make it widely employed is a long-term exercise. There is still a long way to go, developing new commodity products and high-density molecules will be the priority in the coming years. A diagrammatic representation of obtaining different value-added products from marine macroalgal feedstocks is given in Figure 3.1.

3.4

Marine Macroalgal Biomass as an Energy Feedstock

The ever-increasing demands for fossil fuels, coupled with environmental concerns pertaining to greenhouse-gas emissions due to fossil fuels have led to considerable interest among researchers to develop viable and sustainable process for

A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels

47

Recovery of edible salt

Solar evaporation

Confectionary

Processed food Pigments

Milling

Recycling of water

Salt Pans Milling Animal feed Processing Bio compost

Washing Agriculture application

Marine algae

Liquid plant growth enhancer

Washed biomass Agriculture application Biofuel Bio refinary

Lipids

Solid residue

Nutraceuticals

Spent biomass Spent biomass

Hydrocolloid Molecular biology Fermentation applications

Biofuel

Cellulose

Fermentation

Biofuel

Microbial growth media

Plant tissue culture

Figure 3.1

Schematic representation of integrated method of utilization of marine macroalgae.

the production of renewable fuels based on biological material. Biofuels comprise fuel produced through contemporary biological processes and include bioethanol, biodiesel, biogas, bio-oil and biobutanol. The United States and Brazil are the top biofuel producers, and over 95% of it is being produced through edible oil globally [9]. The implications of using edible crops for production of fuel are intensely deliberated. Marine macroalgae are considered potential candidates due to several advantages over terrestrial biomass, such as high photosynthetic efficiency coupled with fast growth; no dependency on agricultural inputs such as land, fertilizer, pesticides and irrigation; easy depolymerization due to the absence of lignin and abundance of fermentable polysaccharides. Among the biofuels, bioethanol is the most popular and widely used as an alternative to fossil fuels, while other forms, namely biogas and bio-oil, need further scientific attention to assess their commercial viability. 3.4.1

Bioethanol

The most common biofuel in use is ethanol with a market share of 93 billion litres [21]. The global demand is expected to increase 3.4-fold by 2035 [22]. It has been produced from several taxa of macroalga by converting their polysaccharides

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into reducing sugars through either acidic hydrolysis or an enzymatic route. Macroalgae contain an array of carbohydrates such as cellulose, laminarin, mannitol, carrageenan and agar. The sugars thus produced need appropriate microorganisms for their fermentation. The low lignin content of macroalgal biomass is a major advantage with respect to saccharification over lignocellulosic biomass. Nevertheless, substrate specificity is a limiting factor for enzymatic hydrolysis due to the several types of hydrocolloids one can expect from algae [23]. Successful bioethanol production has been achieved from K. alvarezii [12], Gelidium amansii [24], Saccharina sp. [25], Laminaria japonica [25] and U. fasciata [26]. Moreover, construction of an engineered microbial platform for direct bioethanol production using either alginate alone [27] or whole biomass of Saccharina japonica [28] has also been ascertained. Table 3.1 summarizes the various pretreatment and conditions employed during enzymatic hydrolysis, saccharification yield, choice of microorganisms used for fermentation and yield achieved during bioethanol production from several brown, red and green algae. 3.4.2

Biodiesel

Biodiesel chemically comprises high-value mono-alkyl esters of long-chain fatty acids. Its use is beneficial to the environment since it is sourced from renewable biomass and has lower emission rates compared to common petroleum diesel. The methods used to obtain biodiesel include pyrolysis, transesterification, microemulsion and supercritical processing of vegetable, animal fats or other plant-based oils. In the case of macroalgae, first converted lipids and free fatty acids are extracted and then, in the second stage, converted to biodiesel using one of the suitable methods listed earlier. In an alternative method, glucose was obtained from Cladophora glomerata which was further converted to free fatty acids to achieve biodiesel production [60]. Xu et al. [61] used L. japonica as a carbon source for oleaginous yeast to produce biodiesel. However, lower content of lipids and non-compliance of yeast to utilize a wide variety of marine macroalgae as a carbon source hinder further developments in this field. 3.4.3

Biobutanol

Marine macroalgal biomass can be converted to butanol through acetone–butanol (AB) fermentation using solventogenic anaerobic Clostridium strains [62]. Clostridium strains can convert a wide variety of sugars (hexoses and pentoses) into acetone, butanol and ethanol (ABE) by anaerobic fermentation. A few reports have demonstrated the production of butanol from macroalgal biomass [57]. However, Clostridium sp. did not effectively utilize some of the glucose-based polysaccharides such as mannitol from brown algae, which caused slow reaction rates and low productivity of organic acids and total solvents [62].

Pretreatment and/ or acid hydrolysis

Sargassum sagamianum

Laminaria japonica

Laminaria japonica

Alaria crassifolia (Kjellman)

n.a.

Enzymatic hydrolysis

Laminarinase 0.1 U/100 g, pH 6, 32 ∘ C, 25% load (wet base) Subject to 0.5 mL 2% Sulfuric acid, 121 ∘ C, 30 min, 25% Meicelase, 50 ∘ C, pH 5.5, 120 h. load Hydrolysates of both steps were merged. 0.1% Sulfuric acid, Cellulase (45 FPU/g) 1.0 h, 121 ∘ C and cellobiose (55 CBU/g), pH 4.8, 50 ∘ C, 2.0% load, 48 h Celluclast 1.5 L, 100 mM HCl, 121 ∘ C, 15 min, 10% viscozyme L, 1%, pH 5.5, 50 ∘ C, 24 h load Thermal n.a. liquefaction, 150 ∘ C, 15 MPa, 15 min, 10% load

Brown marine macroalgae Laminaria n.a. hyperborea Laminaria pH 6, 23 ∘ C, 30 min, hyperborea 25% load (wet base)

Taxa

Table 3.1 Marine macroalgal bioethanol, an appraisal.

Saccharomyces cerevisiae (IAM 4178)

Saccharomyces cerevisiae

Escherichia coli KO11, SSF Pichia stipitis (CBS 7126)

28.4% Glucose and 21.3% galactose

27.8% Glucose

8.1% (w/w)

30.1% (w/w)

n.a.

Pichia angophorae (CBS 5830) Saccharomyces cerevisiae (ethanol red), SSF

Microbial organism used for fermentation

n.a.

Saccharification yield (w/w)

0.4 g ethanol/g sugars 0.3–0.35 g ethanol/g sugar

(continued)

Yeon et al. [34]

Kim et al. [33]

23–29

10

Ge et al. [32]

Yanagisawa et al. [31]

Horn et al. [29] Adams et al. [30]

11.30

22

55.0 g/L

0.143 L/kg

2.84

n.a.

Normalized References yield (%)

0.43 g/g Substrate 0.45 (v/v)

Yield

Saccharomyces cerevisiae

Saccharomyces cerevisiae (DK 410362), SSF

25.50%

29.09% Glucan

0.06% Sulfuric acid, 170 ∘ C, 15 min, 5% load

72% Sulfuric acid, 30 ∘ C, 60 min + 4% sulfuric acid, 120 ∘ C, 40 min

Saccharina japonica

Laminaria digitata and S. latissima

Sargassum spp.

Cellulase 50 FPU/g, cellulase 250 FPU/g, U/g, pH 4.8, 50 ∘ C, 10% load, 100 h Cellulase 15 FPU/g-glucan, β-glucosidase 70 pNPGU/g-glucan, pH 4.8, 40 ∘ C, 3% load, 48 h ® Cellic CTec2 n.a.

n.a.

Escherichia coli (ATCC8739) with synthetic pathway, SSF Pichia angophorae KCTC 17574, SSF

n.a.

14.3

4.90 g/L

n.a.

2.79

2.79 g/L

7.7

7.7 g/L

69.1% of Total carbohydrate in biomass

Bacillus sp. JS-1, 1 g dcw/L. pH 7, 30 ∘ C, 7.5 d

40 mM Sulfuric acid, with Termamyl 120 L, 121 ∘ C, 60 min, 10% solid load 4% Sulfuric acid, 115 ∘ C, 90 min, 10% load

Saccharina japonica

28.1

4.7% (v/v)

n.a.

n.a.

Laminaria japonica

13.2

167 mL per kg biomass

Pichia angophorae, SSF

0.5 U Trichoderma laminarinase, 24 ∘ C n.a.

2 M HCl, 30 min, 5% load

Laminaria digitata

n.a.

Normalized yield (%)

Yield

Saccharification Microbial organism yield (w/w) used for fermentation

Enzymatic hydrolysis

Pretreatment and/ or acid hydrolysis

(Continued)

Taxa

Table 3.1

Manns et al. [40]

Lee et al. [39]

Borines et al. [38]

Jang et al. [37]

Wargacki et al. [36]

Adams et al. [35]

References

4% Sulfuric acid, 30 min Minor pretreated by milling 0.1–1% H2 SO4 , autoclaved at 121 ∘ C for 15 min

4% Sulfuric acid, 120 ∘ C, 60 min 900 mM Sulfuric acid, 100 ∘ C, 60 min, 5% solid load, repeatedly 200 mM Sulfuric acid, 130 ∘ C, 15 min, 10% solid load 1.5% Sulfuric acid, 140 ∘ C, 15% solid load, 60 min 1% Sulfuric acid, 60 min, 33.3% load (w/w) 5.0% Sodium hydroxide, 80 ∘ C, 120 min, 5% load

Gelidium amansii Kappaphycus alvarezii

Gracilaria verrucosa

Kappaphycus alvarezii

Gelidium amansii

Kappaphycus alvarezii

2% Sulfuric acid, 120 ∘ C, 30 min

Gracilaria salicornia

Red marine macroalgae

Saccharina japonica Laminaria digitata Sargassum spp. (spent biomass)

35 g/L Galactose, 8 g/L glucose 81.62 g/L

n.a.

Cellulase 20 FPU/g, β-glucosidase 60 U/g, pH 5.0, 50 ∘ C, 10% load

Cellulase 45 FPU/g

37.82 g/L

30.5% Reducing sugar

n.a.

None

17.4 g Glucose/kg fresh algae 68.58% Glucose 21.6% After five times

80.6% Glucose 81.67 mg/g

Celluclast 1.5 L (%, v/w), 50 ∘ C for 48 h n.a.

MP biomedicals cellulase, 5 g/L, pH 5.0, 26 h n.a.

n.a.

n.a.

Brettanomyces custersii (KCTC18154P) Saccharomyces cerevisiae (CBS1782), SSF Saccharomyces cerevisiae (HAU strain)

Saccharomyces cerevisiae

Saccharomyces cerevisiae (NCIM 3523)

n.a.

E. coli KO11

Saccharomyces cerevisiae Saccharomyces cerevisiae

n.a.

12.80

3.30

39.2

19.5

14.89

3.3 g/L

27.6 g/L

65 g/L

14.89 g/L

n.a.

95.37

n.a.

7.91

1.61

1.61 g/L

79.1 g/kg Dry biomass

77.7

n.a.

77.7 % g/L

37.9% Oil

(continued)

Kumar et al. [13]

Hargreaves et al. [48]

Park et al. [47]

Meinita et al. [46]

Jeong et al. [45] Khambhaty et al. [12]

Wang et al. [44]

Choi et al. [41] Hou et al. [42] Sudhakar et al. [43]

Gelidium amansii

Gracilaria dura

Gelidium pusillum

Gelidiella acerosa

Gelidium amansii

n.a.

Ionic liquid + AIL, 120 ∘ C, 30 min 4% DOWEX™ DR-G8, 120 ∘ C, 30 min, 10% load

Gelidium amansii Eucheuma cottonii

Cellulase 15 FPU/g, 𝛽-glucosidase 52 CBU/g, pH 4.8, 50 ∘ C, 2% load Autoclaving at Cellulase (Celluclast 121 ∘ C for 60 min 1.5 L) and 𝛽-glucosidase (Novozyme 188) n.a. Cellulase 22086 (Novozyme, Denmark) 2% v/v, (pH 4.8), 45 ∘ C, 48 h n.a. Cellulase 22086 (Novozyme, Denmark) 2% v/v, (pH 4.8), 45 ∘ C, 48 h n.a. Cellulase 22086 (Novozyme, Denmark) 2% v/v, (pH 4.8), 45 ∘ C, 48 h 50 mM Sulfuric acid, Mixture of Celluclast 121 ∘ C, 90 min L and Viscozyme 1.5L with 16 unit/m

Enzymatic hydrolysis

Pretreatment and/ or acid hydrolysis

(Continued)

Taxa

Table 3.1

n.a.

n.a.

n.a.

n.a.

0.42 g/g Sugar

0.416 g/g Sugar

0.41 g/g Sugar

18.8 g/L

Saccharomyces cerevisiae (MTCC no. 180) Saccharomyces cerevisiae (MTCC no. 180) Saccharomyces cerevisiae (KCTC 1126)

n.a.

25.7 mg/L

Saccharomyces cerevisiae (MTCC no. 180)

42.4

Around 75% theoretical yield

0.92 g Reducing sugar/g cellulose 0.93 g Reducing sugar/g cellulose 0.91 g Reducing sugar/g cellulose 64.7%

n.a.

n.a.

Normalized yield (%)

Saccharomyces cerevisiae (KCTC 7906)

Saccharomyces cerevisiae (YSC2, type II)

n.a.

Yield

90.7%

81% Galactose 99.8% Glucose

Saccharification Microbial organism yield (w/w) used for fermentation

Sunwoo et al. [52]

Baghel et al. [19]

Baghel et al. [19]

Baghel et al. [19]

Kim et al. [51]

Malihan et al. [49] Tan and Lee [50]

References

Cellulase 22086 (Novozyme, Denmark) 2% v/v, (pH 4.8), 45 ∘ C, 48 h

n.a.

90 mM Sulfuric acid, 121 ∘ C, 90 min

90 mM Sulfuric acid, 121 ∘ C, 90 min

180 mM H2 SO4 at 140 ∘ C for 5 min

n.a.

0.1–1% H2 SO4 , autoclaved at 121 ∘ C for 15 min

Kappaphycus alvarezii

Eucheuma denticulatum

Kappaphycus alvarezii

Gracilaria corticata

Gracilaria corticata

Cellulase 30 FPU/g, pH 4.8, 50 ∘ C, 24 h

Viscozyme L, 2%, 5% load, 36 h

150 ∘ C, 15 MPa, 15 min, 10% load

n.a.

Ulva pertusa

Ulva fasciata

Green marine macroalgae

Mixture of Celluclast L and Viscozyme 1.5 L with 16 unit/m Mixture of Celluclast L and Viscozyme 1.5 L with 16 unit/m Mixture of Celluclast L and Viscozyme 1.5 L with 16 unit/m n.a.

70 mM Sulfuric acid, 121 ∘ C, 90 min

Gracilaria verrucosa

184.4 mg/g

98.59% Glucose

0.269 g reducing sugar/g residual biomass 129.85 mg/g

66.7%

53.1%

53.4%

74.6%

Saccharomyces cerevisiae (ATCC 24858) Saccharomyces cerevisiae (MTCC No. 180)

Saccharomyces cerevisiae

Saccharomyces cerevisiae (KCTC 1126) Saccharomyces cerevisiae (KCTC 1126) Saccharomyces cerevisiae (KCTC 1126) Kluyveromyces marxianus (KCTC7150) Saccharomyces cerevisiae (MTCC no. 180)

12.4

8.3

0.45 g/g Sugar

3.02

12.4 g/L

3.02 g/L

n.a.

0.47 g/g sugar

n.a.

13.0 g/L

n.a.

n.a.

14.5 g/L

11.2 g/L

n.a.

19.1 g/L

(continued)

Trivedi et al. [55]

Choi et al. [54]

Sudhakar et al. [43]

Baghel et al. [18]

Ra et al. [53]

Sunwoo et al. [52]

Sunwoo et al. [52]

Sunwoo et al. [52]

n.a.

Ulva rigida

Source: Reprinted from Jiang et al. [59] with permission from Elsevier.

100 mL Amyloglucosidase (300 units per mL), 40 mL 𝛼-amylase (250 units per mL), and 0.1 g cellulase (0.3 units per mg); 37 ∘ C with shaking at 150 rpm for 24 h

14.4–19.6%

Cladosporium sphaerospermum Saccharomyces cerevisiae

93.81%

n.a.

Ulva fasciata

Clostridium beijerinckii

75–93%

Sulfuric acid, pH 2, 150 ∘ C, 10 min, 10% load n.a.

Ulva lactuca

Saccharomyces cerevisiae (ATCC 96581), SSF

n.a.

Ball milling

Chaetomorpha linum

Saccharification Microbial organism yield (w/w) used for fermentation

Prehydrolysis, Novozyme 188 and Celluclast 1.5 L, 15 FPU/g, 50 ∘ C, 24 h n.a.

Enzymatic hydrolysis

Pretreatment and/ or acid hydrolysis

(Continued)

Taxa

Table 3.1

40

0.33–0.34 g/g n.a. Sugar

0.47 g/g sugar n.a.

40%

Korzen et al. [58]

Trivedi et al. [26]

van der Wal et al. [57]

Schultz-Jensen et al. [56]

Normalized References yield (%)

Ethanol/100 g 18 DM

Yield

A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels

3.4.4

55

Bio-oil

Research on bio-oil production from macroalgae is in its nascent stage and is the most promising. Bio-oil is obtained from algal biomass through pyrolysis. There are three types of pyrolysis that could convert biomass into bio-oil, biochar and gas. The significant difference among the pyrolysis types is the residence time of the solid phase within the reactor and correlated energy transfer coupled with temperature distribution. There are a few reports which demonstrate the production of bio-oil from marine macroalga in the range of 16–47% dry weight, such as Enteromorpha clathrata and Sargassum natans [63]; Laminaria digitata and Fucus serratus [64]; Laminaria saccharina [65]; Undaria pinnatifida, L. japonica and Pyropia tenera [66]. The density of bio-oil is nearly equal to that of petroleum oil. However, the presence of high levels of nitrogen in marine macroalgal biomass in comparison with lignocellulosic biomass as well as a large percentage of ash poses an impediment in bio-oil conversion and its commercial development.

3.5

Advances in Cultivation Technology

Global marine macroalgal production has increased exponentially during the last 50 years, with commercial activities centred in Southeast Asia accounting for over 99.6% of the total marine macroalgal production. This escalated industrial demand has outstripped the supply, triggering dramatic growth through aquaculture. This is an important trend in terms of the sustainability because only about 792,383 tons were harvested from wild which corresponds to only 4% of all biomass being utilized [67]. Biomass production through farming required considerable improvements in existing technology and new farming techniques. Certain improvements have been made in farming techniques, especially off-shore farming, coupled with improvements in cultivar through selective breeding and genetic improvement for yield increment and other traits of economic value, including Saccharina [68], Undaria and Porphyra [69, 70]. The high-temperature-tolerant cultivar of Porphyra haitanensis has been developed through hybridization which can withstand 5 ∘ C elevated temperature compared to wild-type. This improved strain is being cultivated extensively in China [71]. The study of phycopathology is essential as several recent reports described disease outbreak in the Philippines, Malaysia, Zanzibar, Japan, China and Korea. The studies are mostly centred on how induced defence mechanisms in marine macroalgae work [72]. Territorial grazing by herbivorous fishes and epiphytic infestation are yet other issues in commercial large-scale farming. Although much research has been carried out on epiphytic infestations to understand the cause and quality control of product and so on [73], understanding of grazing phenomenon needs more attention. The epiphytes compete with macroalgae for space, nutrients, light and so on, causing detachment of the host tissue. While epiphytes and grazers occur together, grazers might be useful to control the

56

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

epiphytic infestation as observed in G. gracilis from South Africa. Mechanization in harvesting of biomass from nets has been achieved using winches and cranes mounted on large boats or barges to remove either the complete planting setup or just the desired new growth from lines and nets [74]. Attention has also been turned to the use of polyculture practices in marine macroalgal farming. Integrated Multi-Trophic Aquaculture (IMTA) provides an excellent opportunity to integrate farming of organic extractors (e.g. shellfish/fish) and inorganic extractors (marine macroalgae) in fed aquaculture systems. Macroalgae have been widely integrated in various IMTA systems around the world, and their utility have been extensively reviewed [75]. The impediments encountered in near-shore cultivation could be overcome by developing efficient and innovative designs for offshore farming. The concept of a multi-use platform has been gaining momentum. For example, in Europe, the massive expansion of wind farms in offshore areas of the North Sea triggered the idea of a combination of wind turbines with installations of extensive marine algal aquaculture [76]. These sites provide an ideal opportunity for devising and implementing a multiple-use concept as the infrastructure for regular service support is readily available. Nevertheless, focus of such systems should be on improving designs that can withstand hydrodynamic forces experienced in open sea and on improving cultivation techniques to avoid dislodgement of algal seeds.

3.6

Marine Algal Cultivation for CO2 Sequestration

The world is facing the serious challenge of global warming owing to emissions of greenhouse gases including CO2 . The concept of combining bioenergy with carbon capture and storage might provide a workable solution to energy production with net negative emissions of CO2 . Marine macroalgae have high productivity rates due to efficient photosynthesis with estimated gross primary productivity of approximately 1600 g C/m2 /y [77]. Species such as Ulva, Porphyra, Macrocystis, Sargassum, Laminaria, Palmaria, Fucus and Ascophyllum have high capacity of CO2 assimilation per gram fresh weight [78–81]. There are several reports that emphasized the marine macroalgae cultivation for efficient CO2 utilization [82, 83]. The CO2 sequestration statistics given by Hughes et al. [84] follows three basic components: 1. Production of marine macroalgae: 1 ha of sea front is considered to yield 200 tons wet weight (w.w./ha), which corresponds to approximately 20 tons dry weight, of which 6 tons/ha is carbon. Approximately 40% of net primary productivity is lost as dissolved organic carbon; therefore, 2.4 tons of carbon is released into the water column per hectare during cultivation per year. About 15% of this is refractory which is resistant to microbiological degradation and will join the oceanic carbon pool. This is effectively sequestered, totalling 0.36 tons of carbon, while the remaining is respired back through microbial loop.

A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels

57

2. Biomass conversion: The anaerobic biomass conversion facility is considered by the authors as the best data are available for this system. The industrial digestion of brown marine algae yielded 11 m3 methane per tonne w.w. macroalgae. Hence, for a hectare, it would correspond to 220 m3 of methane equating to 110 kg of carbon as methane and given a 60/40% methane/CO2 mix in the biogas produced, 60 kg carbon as CO2 . The remaining carbon is retained in the digestate or in the liquor that is produced. 3. The common application for this digestate is for soil amendment. A portion of it will be respired back to CO2 by microbial degradation, and some of which resists degradation will effectively be stored in the soil. For example, Kelp species are reported to have 7% dry biomass as refractory fibre; thus, 0.42 tC would be stored as refractory organic material in the soil. Within this model, for every hectare of the macroalgae that is used to produce biofuels, a total of 0.78 tC is stored either into the marine refractory dissolved organic carbon pool or into the soil refractory organic carbon pool, and 161 GJ of energy is generated (using the conversion ratio of 55.7 kJ/g methane). Although this model has certain limitations, it is evident that marine macroalgal cultivation for biofuel production can be a carbon negative.

3.7

Opportunities, Challenges and Conclusions

Although there are several Southeast Asian countries involved in the commercialization of marine macroalgae or algae-derived products, there is much more effort required at a global scale to make this sector more popular. Marine macroalgae have a potential to become widely used health-promoting foods due to the array of primary and secondary metabolites within them. The metabolome analysis techniques developed recently [85] might aid in elucidating biochemical regulation, leading to metabolite profiling for potent new functional ingredients/foods. There is exponential growth expected in the marine algal sector in the coming years, due to the demand for raw material foods, feed, fertilizer and speciality chemicals in cosmetics, nutraceuticals and pharmaceuticals [72]. Although chemical extractions of valuable compounds have used routine industrial protocols for the last several years, new emerging enzyme-based techniques are bound to make impacts by enabling recovery of value-added compounds which might not be recoverable when conventional chemical extraction methods are used. Research in nanoscience might also help in targeted extraction of certain valuable compounds of high value. Marine macroalgae-based liquid extracts are fast emerging as potential plant growth stimulants occupying substantial market share in countries where agriculture is predominant. The successful expansion of the marine macroalgal market to multiple product profiles requires equally competitive protocols to be developed for their farming (as discussed in Chapter 2). The domestication of new cultivar and expansion of farming to new locations are real challenges, while one needs to also consider genetic improvement of cultivar

58

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

through classical breeding. Although, off-shore waters provide vast areas for marine algal farming, commercial success can only be achieved if engineering inputs are combined with implementation of the concept of multiple use of infrastructure. The co-cultivation or IMTA shall enable the use of resources and infrastructure judicially to exploit their benefits to their full potential, while capitalizing on environmental mitigation of excess nutrients. Therefore, large-scale open-ocean farming could be implemented for the production of next-generation biofuels. Furthermore, marine macroalgae have unique advantages in displaying high productivity and, at the same time, potential to avoid competition for arable land. The biorefinery model would be the most successful as shown by several short-term studies, only if the production of low-volume, high-value co-products can be achieved. The sequential extraction of multiple value-added products using innovative green methodologies is crucial for the recovery of solvents and other chemicals without compromising on yield. The proper integration of production could make it a profitable business venture which would not be possible by targeting only one or two products even after significant optimization. The production of biofuel strategies can be changed based on the kind of product one is looking at, such as fermentation, anaerobic digestion, pyrolysis and gasification. The integration of production of pigments, lipids, phycocolloids, and liquid fertilizers has been achieved successfully [19]; however, further integration of high-value compounds for maximizing feedstock is essential. It is anticipated that demand for these compounds will be easily satisfied which in turn would influence the price of biofuel. To further achieve this, biorefineries need to be located close to the off-shore regions where marine macroalgae are cultivated and partial processing of biomass has to be done immediately in order to reduce labour and transportation costs. Large-scale commercial cultivation also opens up new opportunities for CO2 sequestration. Extensive techno-economic analytical studies have shown that the energy potential of marine macroalgae compares very favourably to other terrestrial sources of biomass. The utilization of marine macroalgae for food, chemicals and fuels is profitable, if a biorefinery concept is adopted. Although, currently, no commercial marine macroalgal-based biorefinery is operational, several countries, namely the United States, Australia, China, Japan, South Korea, the United Kingdom, Italy, India and the Philippines have announced plans to invest heavily in research and development on algal-based biofuel. The amalgamation of genetic engineering, ocean engineering, and chemical engineering will certainly pave way for advancement of utilization of macroaglae for food, chemicals and fuels.

References 1. Dillehay, T.D., Ramírez, C., Pino, M. et al. (2008) Monte verde: seaweed, food, medicine, and the peopling of South America. Science, 320, 784–786. 2. Rioux, L.E. and Turgeon, S.L. (2015) Seaweed carbohydrates, in Seaweed Sustainability – Food and Non-food Applications (eds B. Tiwari and D. Troy), Elsevier, Amsterdam, pp. 141–192.

A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels

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3. FAO (2014) The State of World Fisheries and Aquaculture: Opportunities and Challenges, Rome. 4. Bixler, H.J. and Porse, H. (2011) A decade of change in the seaweed hydrocolloids industry. Journal of Applied Phycology, 23, 321–335. 5. Hernandez-Carmona, G., Freile-Pelegrin, Y. and Garibay, E.H. (2013) Conventional and alternative technologies for the extraction of algal polysaccharides, in Functional Ingredients from Algae for Foods and Nutraceuticals, Woodhead Publishing Series in Food Science, Technology and Nutrition: Number 256 (ed. H. Dominguez), Woodhead Publishing, pp. 475–516. 6. Pomin, V.H. (2012) Fucanomics and galactanomics: current status in drug discovery, mechanisms of action and role of the well-defined structures. Biochimica et Biophysica Acta, 1820, 1971–1979. 7. Sharma, M., Chaudhary, J.P., Mondal, D. et al. (2015) A green and sustainable approach to utilize bio-ionic liquids for the selective precipitation of high purity agarose from an agarophyte extract. Green Chemistry, 17, 2867–2873. 8. Shukla, M.K., Kumar, M., Prasad, K. et al. (2011) Partial characterization of sulfohydrolase from Gracilaria dura and evaluation of its potential application in improvement of the agar quality. Carbohydrate Polymers, 85, 157–163. 9. Bharathiraja, B., Jayamuthunagai, J., Chakravarthy, M. et al. (2015) Algae: promising future feedstock for biofuels, in Algae and Environmental Sustainability (eds B. Singh et al.), Volume 7 of the Series Developments in Applied Phycology, Elsevier, pp. 1–8. 10. van Hal, J.W., Huijgen, W.J.J. and Lopez-Contreras, A.M. (2014) Opportunities and challenges for seaweed in the biobased economy. Trends in Biotechnology, 32, 231–233. 11. Eswaran, K., Ghosh, P.K., Siddhanta, A.K. et al. (2002) Integrated method for production of carrageenan and liquid fertilizer from fresh seaweeds. US Patent 6893479, filed Aug 19, 2002 and issued May 17, 2005. 12. Khambhaty, Y., Mody, K., Gandhi, M.R. et al. (2012) Kappaphycus alvarezii as a source of bioethanol. Bioresource Technology, 103, 180–185. 13. Kumar, S., Gupta, R., Kumar, G. et al. (2013) Bioethanol production from Gracilaria verrucosa, a red alga, in a bio-refinery approach. Bioresource Technology, 135, 150–156. 14. Niu, J., Xu, M., Wang, G. et al. (2013) Comprehensive extraction of agar and R-phycoerythrin from Gracilaria lemaneiformis (Bangiales, Rhodophyta). Indian Journal of Geo-Marine Science, 42 (1), 21–28. 15. Francavilla, M., Manara, P., Kamaterou, P. et al. (2015) Cascade approach of red macroalgae Gracilaria gracilis sustainable valorization by extraction of phycobiliproteins and pyrolysis of residue. Bioresource Technology, 184, 305–313. 16. Mondal, D., Sharma, M., Maiti, P. et al. (2013) Fuel intermediates, agricultural nutrients and pure water from Kappaphycus alvarezii seaweed. RSC Advances, 3, 17989–17997. 17. Baghel, R.S., Reddy, C.R.K. and Bhavanath, J. (2014) Characterization of agarophytic seaweeds from the biorefinery context. Bioresource technology, 159, 280–285. 18. Baghel, R.S., Trivedi, N. and Reddy, C.R.K. (2016) A simple process for recovery of a stream of products from marine macroalgal biomass. Bioresource Technology, 203, 160–165. 19. Baghel, R.S., Trivedi, N., Gupta, V. et al. (2015) Biorefining of marine macroalgal biomass for production of biofuel and commodity chemicals. Green Chemistry, 17, 2436–2443. 20. Trivedi, N., Baghel, R.S., Gupta, V. et al. (2016) An integrated process for the extraction of fuel and chemicals from marine macroalgal biomass. Scientific Reports, 6, 30728. 21. Renewable Fuels Association, Ethanol Industry Outlook (2015), https://ethanolrfa.3cdn.net/ c5088b8e8e6b427bb3_cwm626ws2.pdf (accessed 30th August, 2016).

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

22. International Energy Agency (2012) World Energy Outlook. 23. Choi, D., Simb, H.S., Piao, Y.L. et al. (2009) Sugar production from raw seaweed using the enzyme method. Journal of Industrial and Engineering Chemistry, 15, 12–15. 24. Meinita, M.D.N., Marhaeni, B., Winanto, T. et al. (2013) Comparison of agarophytes (Gelidium, Gracilaria, and Gracilariopsis) as potential resources for bioethanol production. Journal of Applied Phycology, 25, 1957–1961. 25. Lee, J.Y., Kim, Y.S., Um, B.H. et al. (2013) Pretreatment of Laminaria japonica for bioethanol production with extremely low acid concentration. Renewable Energy, 54, 196–200. 26. Trivedi, N., Reddy, C.R.K., Radulovich, R. et al. (2015) Solid state fermentation (SSF)-derived cellulase for saccharification of the green seaweed Ulva for bioethanol production. Algal Research, 9, 48–54. 27. Takeda, H., Yoneyama, F., Kawai, S. et al. (2011) Bioethanol production from marine biomass alginate by metabolically engineered bacteria. Energy & Environmental Science, 4 (7), 2575–2581. 28. Enquist-Newman, M., Faust, A.M.E., Bravo, D.D. et al. (2014) Efficient ethanol production from brown macroalgae sugars by a synthetic yeast platform. Nature, 505 (7482), 239–243. 29. Horn, S.J., Aasen, I.M. and Østgaard, K. (2000) Ethanol production from seaweed extract. Journal of Industrial Microbiology and Biotechnology, 25, 249–254. 30. Adams, J., Gallagher, J. and Donnison, I. (2009) Fermentation study on Saccharina latissima for bioethanol production considering variable pre-treatments. Journal of Applied Phycology, 21, 569–574. 31. Yanagisawa, M., Nakamura, K., Ariga, O. and Nakasaki, K. (2011) Production of high concentrations of bioethanol from seaweeds that contain easily hydrolyzable polysaccharides. Process Biochemistry, 46, 2111–2116. 32. Ge, L., Wang, P. and Mou, H. (2011) Study on saccharification techniques of seaweed wastes for the transformation of ethanol. Renewable Energy, 36, 84–89. 33. Kim, N.J., Li, H., Jung, K. et al. (2011) Ethanol production from marine algal hydrolysates using Escherichia coli KO11. Bioresource Technology, 102, 7466–7469. 34. Yeon, J.H., Lee, S.E., Choi, W.Y. et al. (2011) Repeated-batch operation of surface-aerated fermentor for bioethanol production from the hydrolysate of seaweed Sargassum sagamianum. Journal of Microbiology and Biotechnology, 21, 323–331. 35. Adams, J.M.M., Toop, T.A., Donnison, I.S. et al. (2011) Seasonal variation in Laminaria digitata and its impact on biochemical conversion routes to biofuels. Bioresource Technology, 102, 9976–9984. 36. Wargacki, A.J., Leonard, E., Win, M.N. et al. (2012) An engineered microbial platform for direct biofuel production from brown macroalgae. Science, 335, 308–313. 37. Jang, J.S., Cho, Y., Jeong, G.T. et al. (2012) Optimization of saccharification and ethanol production by simultaneous saccharification and fermentation (SSF) from seaweed, Saccharina japonica. Bioprocess and Biosystems Engineering, 35, 11–18. 38. Borines, M.G., de Leon, R.L. and Cuello, J.L. (2013) Bioethanol production from the macroalgae Sargassum spp. Bioresource Technology, 138, 22–29. 39. Lee, J.Y., Li, P., Lee, J.Y. et al. (2013) Ethanol production from Saccharina japonica using an optimized extremely low acid pretreatment followed by simultaneous saccharification and fermentation. Bioresource Technology, 127, 119–125.

A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels

61

40. Manns, D., Deutschle, A.L., Saake, B. et al. (2014) Methodology for quantitative determination of the carbohydrate composition of brown seaweeds (Laminariaceae). RSC Advances, 4, 25736–25746. 41. Choi, J., Choi, J.W., Suh, D.J. et al. (2014) Production of brown algae pyrolysis oils for liquid biofuels depending on the chemical pretreatment methods. Energy Conversion and Management, 86, 371–378. 42. Hou, X.R., Hansen, J.H. and Bjerre, A.B. (2015) Integrated bioethanol and protein production from brown seaweed Laminaria digitata. Bioresource Technology, 197, 310–317. 43. Sudhakar, M.P., Merlyn, R., Arunkumar, K. and Perumal, K. (2016) Characterization, pretreatment and saccharification of spent seaweed biomass for bioethanol production using baker’s yeast. Biomass & Bioenergy, 90, 148–154. 44. Wang, X., Liu, X. and Wang, G. (2011) Two-stage hydrolysis of invasive algal feedstock for ethanol Fermentation. Journal of Integrative Plant Biology, 53, 246–252. 45. Jeong, T.S., Kim, Y.S. and Oh, K.K. (2011) Two-stage acid saccharification of fractionated Gelidium amansii minimizing the sugar decomposition. Bioresource Technology, 102, 10529–10534. 46. Meinita, M., Kang, J.Y., Jeong, G.T. et al. (2012) Bioethanol production from the acid hydrolysate of the carrageenophyte Kappaphycus alvarezii (cottonii). Journal of Applied Phycology, 24, 857–862. 47. Park, J.H., Hong, J.Y., Jang, H.C. et al. (2012) Use of Gelidium amansii as a promising resource for bioethanol: a practical approach for continuous dilute-acid hydrolysis and fermentation. Bioresource Technology, 108, 83–88. 48. Hargreaves, P.I., Barcelos, C.A., da Costa, A.C.A. et al. (2013) Production of ethanol 3G from Kappaphycus alvarezii: evaluation of different process strategies. Bioresource Technology, 134, 257–263. 49. Malihan, L.B., Nisola, G.M., Mittal, N. et al. (2014) Blended ionic liquid systems for macroalgae pretreatment. Renewable Energy, 66, 596–604. 50. Tan, I.S. and Lee, K.T. (2015) Solid acid catalysts pretreatment and enzymatic hydrolysis of macroalgae cellulosic residue for the production of bioethanol. Carbohydrate Polymer, 124, 311–321. 51. Kim, H.M., Wi, S.G., Jung, S., Song, Y. and Bae, H.-J., (2015) Efficient approach for bioethanol production from red seaweed Gelidium amansii. Bioresource Technology, 175, 128–134. 52. Sunwoo, I.Y., Ra, C.H., Jeong, G.T. and Kim, S.K. (2016) Evaluation of ethanol production and bioadsorption of heavy metals by various red seaweeds. Bioprocess and Biosystems Engineering, 39, 915–923. 53. Ra, C.H., Nguyen, T.H., Jeong, G.T. and Kim, S.K. (2016) Evaluation of hyper thermal acid hydrolysis of Kappaphycus alvarezii for enhanced bioethanol production. Bioresource Technology, 209, 66–72. 54. Choi, W.Y., Han, J.G., Lee, C.G. et al. (2012) Bioethanol production from Ulva pertusa Kjellman by high-temperature liquefaction. Chemical and Biochemical Engineering Quarterly, 26, 15–21. 55. Trivedi, N., Gupta, V., Reddy, C.R.K. et al. (2013) Enzymatic hydrolysis and production of bioethanol from common macrophytic green alga Ulva fasciata Delile. Bioresource Technology, 150, 106–112.

62

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

56. Schultz-Jensen, N., Thygesen, A., Leipold, F. et al. (2013) Pretreatment of the macroalgae Chaetomorpha linum for the production of bioethanol – comparison of five pretreatment technologies. Bioresource Technology, 140, 36–42. 57. van der Wal, H., Sperber, B.L.H.M., Houweling-Tan, B. et al. (2013) Production of acetone, butanol, and ethanol from biomass of the green seaweed Ulva lactuca. Bioresource Technology, 128, 431–437. 58. Korzen, L., Abelson, A. and Israel, A. (2016) Growth, protein and carbohydrate contents in Ulva rigida and Gracilaria bursa-pastoris integrated with an offshore fish farm. Journal of Applied Phycology, 28, 1835–1845. 59. Jiang, R., Ingle, K.N. and Golberg, A. (2016) Macroalgae (seaweed) for liquid transportation biofuel production: what is next? Algal Research, 14, 48–57. 60. Chen, H., Zhou, D., Luo, G. et al. (2015) Macroalgae for biofuels production: progress and perspectives. Renewable and Sustainable Energy Reviews, 47, 427–437. 61. Xu, X., Kim, J.Y., Oh, Y.R. and Park, J.M. (2014) Production of biodiesel from carbon sources of macroalgae, Laminaria japonica. Bioresource Technology, 169, 455–461. 62. Huesemann, M.H., Kuo, L.-J., Urquhart, L. et al. (2012) Acetone-butanol fermentation of marine macroalgae. Bioresource Technology, 108, 305–309. 63. Wang, S., Wang, Q., Jiang, X. et al. (2013) Compositional analysis of bio-oil derived from pyrolysis of seaweed. Energy Conversion and Management, 68, 273–280. 64. Yanik, J., Stahl, R., Troeger, N. et al. (2013) Pyrolysis of algal biomass. Journal of Analytical and Applied Pyrolysis., 103, 134–141. 65. Anastasakis, K. and Ross, A.B. (2011) Hydrothermal liquefaction of the brown macro-alga Laminaria saccharina: effect of reaction conditions on product distribution and composition. Bioresource Technology, 102, 4876–4883. 66. Bae, Y.J., Ryu, C., Jeon, J.K. et al. (2011) The characteristics of bio-oil produced from the pyrolysis of three marine macroalgae. Bioresource Technology, 102, 3512–3520. 67. White, W.L. and Wilson, P. (2015) World seaweed utilization, in Seaweed Sustainability — Food and Non-Food Applications (eds B.K. Tiwari and D.J. Troy), Elsevier, Amsterdam (The Netherlands), pp. 7–25. 68. Yan, L., Jianzhou, C., Xueyan, S. et al. (2006) The application of DNA molecular marker technique in heritable breeding of Laminaria. Transactions of Oceanology and Limnology, 1 (107), 75–81. 69. Chaoyuan, W. and Guangheng, L. (1987) Progres in the genetics and breeding of economic seaweeds in China. Hydrobiologia, 151 (1), 57–61. 70. Dai, J., Zhang, Q. and Bao, Z. (1993) Genetic breeding and seedling raising experiments with Porphyra protoplasts. Aquaculture, 111 (1-4), 139–145. 71. Yan, X.H., Zhang, S.J. and Huang, L.B. (2009) Induction and isolation of pigmentation mutants of Porphyra yezoensis Ueda (Bangiales, Rhodophyta) by 60 Co-𝛾 ray irradiation. Oceanologia et Limnologia Sinica, 40, 56–61. 72. Hafting, J.T., Cornish, M.L., Deveau, A. et al. (2015) Marine algae: gathered resource to global food industry, in The Algae World, Springer, Netherlands, pp. 403–427. 73. Vairappan, C.S., Kamada, T., Lee, W.W. et al. (2013) Anti-inflammatory activity of halogenated secondary metabolites of Laurencia snackeyi (Weber-van Bosse) Masuda in LPS-stimulated RAW 264.7 macrophages. Journal of Applied Phycology, 25 (6), 1805–1813.

A New Wave of Research Interest in Marine Macroalgae for Chemicals and Fuels

63

74. Radulovich, R., Neori, A., Valderrama, D. et al. (2015) Farming of seaweeds, in Seaweed Sustainability — Food and Non-Food Applications (eds B.K. Tiwari and D.J. Troy), Elsevier, Amsterdam (The Netherlands), pp. 27–53. 75. Chopin, T., MacDonald, B., Robinson, S. et al. (2013) The Canadian Integrated Multi-Trophic Aquaculture Network (CIMTAN)—a network for a new ERA of ecosystem responsible aquaculture. Fisheries, 38 (7), 297–308. 76. Buck, B.H. and Krause, G. (2012) Integration of aquaculture and renewable energy systems, in Encyclopaedia of Sustainability Science and Technology (ed. R.A. Meyers), Springer, New York, pp. 511–533. 77. Hughes, A.D., Black, K.D., Campbell, I. et al. (2012) Does seaweed offer a solution for bioenergy with biological carbon capture and storage? Greenhouse Gases: Science and Technology, 2, 402–407. 78. Chung, I., Beardall, J., Mehta, S. et al. (2011) Using marine macroalgae for carbon sequestration: a critical appraisal. Journal of Applied Phycology, 23, 877–886. 79. Gao, K. and McKinley, K.R. (1994) Use of macroalgae for marine biomass production and CO2 remediation—a review. Journal of Applied Phycology, 6, 45–60. 80. Jackson, G.A. (1987) Modelling the growth and harvest yield of the giant kelp Macrocystis pyrifera. Marine Biology, 95, 611–624. 81. Muraoka, D. (2004) Seaweed resources as a source of carbon fixation. Bulletin of Fisheries Research Agency, 1, 59–63. 82. Erlania and Radiarta, I.N. (2015) The use of seaweeds aquaculture for carbon sequestration: a strategy for climate change mitigation. Journal of Geodesy and Geomatics Engineering, 2, 109–115. 83. Kaladharan, P., Veena, S. and Vivekanandan, E. (2009) Carbon sequestration by a few marine algae: observation and projection. The Marine Biological Association of India, 51 (1), 107–110. 84. Hughes, A.D., Kelly, M.S., Black, K.D. et al. (2012) Biogas from Macroalgae: is it time to revisit the idea? Biotechnology for Biofuels, 5, 86. 85. Gupta, V., Thakur, R.S., Reddy, C.R.K. et al. (2013) Central metabolic processes of marine macrophytic algae revealed from NMR based metabolome analysis. RSC Advances, 3 (19), 7037–7047.

4 Kappaphycus alvarezii: A Potential Sustainable Resource for Fertilizers and Fuels Dibyendu Mondal1,2 and Kamalesh Prasad1,3 1 Natural

Products & Green Chemistry Division, CSIR – Central Salt and Marine Chemicals Research Institute, India 2 Department of Chemistry, CICECO-Aveiro Institute of Materials, University of Aveiro, Portugal 3 AcSIR – Central Salt & Marine Chemicals Research Institute, India

4.1

Introduction

Sustainable developments targeting production of fuels, chemicals and functional materials from biomass are imperative to mitigate the threats of depletion of fossil resources [1]. Among several bioresources, seaweeds are emerging as one of the most promising bioresources for such research owing to their advantages over other biomass counterparts [2–4]. Recent data showed that the market of seaweed industry has grown beyond $6 billion, with an annual consumption of more than 8 million tonnes [5]. Moreover, seaweeds are rich in high-quality ingredients such as hydrocolloids, bioactive polysaccharides, fatty acids, vitamins,

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources, First Edition. Edited by Francesca M. Kerton and Ning Yan. © 2017 John Wiley & Sons Ltd. Published 2017 by John Wiley & Sons Ltd.

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pigments, sugars and proteins which make them as an unconventional source for value-added products having direct commercial utility [4]. Therefore, seaweed could yield value-added products such as nutraceuticals, colorants, antioxidants, water-soluble biopolymers, cellulose and flavours. These products have direct applications in food, pharmaceutical, cosmetics, biomaterials and fuel industries [5]. However, this much utilization accounts for only 350 [11]. The common practice for processing of K. alvarezii centres on the extraction of carrageenan. For this purpose, the freshly harvested seaweed biomass is washed, ground and then sundried. Then the dried biomass undergoes chemical and thermal pretreatment before further processing. Several methods such as a freeze–thaw method, a KCl precipitation method and a solvent precipitation method (Figure 4.2) have been reported for the extraction of carrageenan having different gel strengths and yields [14].

Red seaweed (fresh) Washing, grinding and sun-drying Dry seaweed mass Alkali treatment (10% KOH aqueous solution) Alkali treated seaweed Autoclave in water (1:20 w/v) at 110°C for 90 min. Grinding and centrifugation

Seaweed residue

Carrageenan rich seaweed extractive

Frozen at −15°C for 12 h

1.0−1.5% KC1 solution precipitation Stirring until cooling

Thawing the gel Drying

Precipitation with alcohol Separation

Washing and freeze−thaw

Repeated washing with alcohol

Drying of carrageenan

Drying

Purified carrageenan

Figure 4.2 Different extraction procedures for κ-carrageenan from Kappaphycus alvarezii.

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Kappaphycus alvarezii Powder soaked in DES 1h

Isopropyl alcohol

85 °C

Centrifugation

κ-Carrageenan rich bottom layer

κ-Carrageenan

Kappaphycus alvarezii Extract in DES Figure 4.3 Schematic of κ-carrageenan production using a deep eutectic solvent. (Source: Das et al. 2016 [15]. Reproduced with permission of Elsevier.)

Considering the time-consuming extraction methods discussed earlier and in order to find better alternative solvents to extract κ-carrageenan, deep eutectic solvents (DESs) were used to extract carrageenan from K. alvarezii (Figure 4.3) [15]. DESs are analogues of ionic liquids and can be synthesized by the reaction of a hydrogen bond acceptor (such as quaternary ammonium salts) and hydrogen bond donors (such as urea, ethylene glycol, glycerol, etc.) [16]. As can be seen from Table 4.1, three different DESs and their hydrated counterparts were employed for the extraction of carrageenan, and the extraction efficiency of these solvents was compared with those obtained using a conventional method [15]. It was found that the physicochemical as well as rheological properties of κ-carrageenan obtained using DESs as solvents were at par to those obtained using a conventional method and were superior in quality when compared to κ-carrageenan obtained using water as solvent. Considering the tedious nature of the extraction method employed in conventional extraction process, the DESs can be considered as suitable alternative solvents for the facile extraction of the polysaccharide directly from the seaweed. However, among the hydrated and non-hydrated DESs, the hydrated ones were found to be more effective in comparison to their non-hydrated counterparts. DESs afforded a high yield of carrageenan, and their use allows facile extraction that may help towards sustainable valorization of seaweed for fuels and chemicals.

4.3

Simultaneous Production of Liquid Fertilizer (𝛋-Sap) and 𝛋-Carrageenan from Fresh Kappaphycus alvarezii Seaweed

It is well known that the major portion (>80% of the total weight) of freshly harvested seaweed is water. K. alvarezii specifically contains only 4–10% dry matter, and the remaining is water. K. alvarezii is known as a source of refined

Kappaphycus alvarezii: A Potential Sustainable Resource for Fertilizers and Fuels

69

Table 4.1 Effect of different deep eutectic solvents on yield and viscosity of κ-carrageenan. S. no.

Solvent system/method

Yield% (±S.D.)

Viscosity (cP) of solvent at 25 ∘ C

1 2

Choline chloride–urea 1:2 Choline chloride–ethylene glycol 1:2 Choline chloride–glycerol 1:2 10% Hydrated choline chloride–urea 1:2 10% Hydrated choline chloride–ethylene glycol 1:2 10% Hydrated choline chloride–glycerol 1:2 Water Conventional methoda

37.60 ± 1.20 50.66 ± 1.78

289.10 35.49

30.93 ± 0.90 53.64 ± 1.25

266.60 34.49

46.02 ± 2.30

16.50

60.25 ± 1.10

52.64

46.87 ± 2.00 36.58 ± 1.90

0.90 n.a.

3 4 5 6 7 8

Source: Das et al. 2016 [15]. Reproduced with permission of Elsevier. n.a., Not applicable. a Craigie and Leigh [17].

and semi-refined κ-carrageenans, and its extract can be used as a foliar spray to increase flowering and growth of a number of crops [18]. As described by Chapman and Chapman [19], conventionally, all seaweeds, including K. Alvarezii, are dried at their harvesting place and baled for shipment to processing plants. Among the drying techniques, sun-drying remains the most cost-effective one, even though oil-fired mechanical dryers are used to a limited extent. The dry seaweed is used mainly for the production of phycocolloids. However, some cottage industries are also producing seaweed fertilizer such as in certain coastal villages of India, the freshly harvested seaweed is boiled in earthen pots to make the liquid extract utilized as fertilizer and the solid residue is either discarded or sometimes used as manure [19]. Prior to drying to yield dry seaweed or making extract to obtain fertilizer, in 2005, Eswaran et al. invented an integrated method for the simultaneous production of sap (κ-Sap) and solid residue to utilize the fresh biomass of K. alvarezii [20] to a maximum extent. The process includes simple crushing to release sap followed by centrifuging to separate the sap and solid residue called seaweed granules. The sap is useful as a potent liquid plant stimulant while the residue is a superior raw material for the extraction of semi-refined or refined κ-carrageenan, thereby maximizing the value of the seaweed. Moreover, the requirement of time and area for drying of the seaweed to obtain the raw material for κ-carrageenan production can be eliminated, and the process also reduces the cost of transporting and storing this raw material because of its lesser bulk and easier handling due to its free-flowing granular nature.

4.4

𝛋-Sap as Potential Plant Stimulant

It is well known that seaweed extracts can be used as foliar spray to enhance plant productivity [19], but among many of them, the pure κ-Sap derived from K. alvarezii is unique as a plant growth stimulant. A pronounced increment of

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Table 4.2 Details of multi-institutional crop trials carried out on maize using Kappaphycus alvarezii sap. S. no.

Variety

University/ institute

Location of trial

Maximum grain yield obtained (kg/ha)a

Increase in grain yield over and above RDFb (%)

1 2

Disha 3502 Rajkumar

West Bengal West Bengal

3595 3170

48.4 17.8

3 4

900M Gold DA 61 A

Bihar Meghalaya

7290 5265a

35.2 16.6

5

Hema (NAH 1137)

Karnataka

7695

15.8

6 7

PEHM-2 HQPM1

Rajasthan Haryana

4599 5786

15.7 11.5

8

NK 6240

Tamil Nadu

7242

13.8

9

HQPM1

BCKV, Nadia PSB, Shantiniketan RAU, Samastipur ICAR NEH Region, Umiam GKVK, Bangalore MPUAT, Udaipur CCHAU, Hisar, Karnal TNAU, Coimbatore BAU, Ranchi

Jharkhand

4092

21.3

Source: Mondal et al. 2015 [22]. Reproduced with permission of Springer. Percent increases over and above RDF are statistically significant at 5% level of significance. a Average yield over two seasons. b RDF, recommended dose of fertilizers.

yields of many crops as well as plant productivity has been found after application of the κ-Sap as a foliar spray at 2.5–10% (v/v, dilution with water) level [21]. The results achieved with soybean under rain-fed conditions have been reported recently, where a 46% increase in yield was observed upon application of the sap at 12.5% concentration (Table 4.2) [23]. κ-Sap has been applied to enhance the yields of many crops such as wheat, brinjal, potato, capsicum, peanut, chilli and okra by 10–33% over and above control (water spray) treatment. Sugar cane and other C4 crops also responded well to the pristine κ-Sap [22]. A recent study was carried out as part of a pan India multi-institutional multi-crop project to evaluate the efficacy of the κ-Sap wherein about 40 agricultural institutes participated in field trials. As can be seen from the Table 4.2, the grain yield of maize was enhanced significantly upon application of κ-Sap at 5% (v/v) dilution with water [22]. A study was conducted to elucidate the composition of the κ-Sap which is beneficial for the plant productivity. Table 4.3 summarizes the data for elemental composition and plant growth promoter present in the κ-Sap. Besides a large amount of potassium, the sap is rich in several others macro- and micronutrients. The significant presence of several plant growth regulators (Indole-3-acetic acid (IAA), kinetin, zeatin and GA3 ) and quaternary ammonium compounds (choline and glycine betaine) in the κ-Sap which are beneficial for plant growth and productivity was also noted.

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Table 4.3 Composition of Kappaphycus alvarezii sap. Element

Concentration

Growth promoters

Concentration

K Na Mg Ca N P Fe Mn Zn B Cu

21 ± 0.72 g/L 1.3 ± 0.11 g/L 2.1 ± 0.12 g/L 1.7 ± 0.05 g/L 85.0 ± 7.0 mg/L 29.0 ± 2.5 mg/L 10.0 ± 4.0 mg/L 2.8 ± 0.15 mg/L 7.7 ± 1.15 mg/L 2.8 ± 0.24 mg/L 300 μg/L

Indole-3-acetic acid Kinetin Zeatin Gibberellic acid (GA3 ) Glycine betaine Choline Betaine aldehyde

21.11 mg/L 9.21 mg/L 18.62 mg/L 25.72 mg/L 78.47 mg/L 60.71 mg/L Detected

Source: Mondal et al. 2015 [22]. Reproduced with permission of Springer.

4.5

Manipulation of 𝛋-Sap for Sustainable Biomass Intensification of Maize

Enhancements in food production and biomass for energy are challenging goals for humanity. Being able to achieve them with low carbon and water footprints is also of critical importance. Liquid seaweed fertilizers are reported to have profound effects on the productivity of crops [24]. Seaweeds as a source of plant nutrients are also attractive, considering the overall life-cycle analysis. Since seaweed fertilizers are reportedly low in nutrients such as nitrogen and phosphorus, their performance is augmented through nutrient supplementation, for example, through addition of protein hydrolysate or blending of different types of extracts [25]. Performance enhancement through simplification of composition is not known. Moreover, as stated earlier, the κ-Sap is a complex concoction of several known and yet to be identified components and thus difficult to identify the most critical constituents responsible for productivity enhancement. On the other hand, Weiss and Ori explored the evidence of cross talk between cytokinins and gibberellins [26]. Such interactions among different hormones present in the κ-Sap were studied by simplification of its composition through selective removal of one or more hormones. Ethyl acetate treatment was found to be effective for the removal of GA3 (GA3 free κ-Sap), and diethyl ether was employed to remove indole-3-acetic acid (IAA free κ-Sap) from the pristine κ-Sap [19, 22]. These κ-Sap variants were applied as foliar spray (5% v/v dilution with water) together with pristine κ-Sap. Among all, application of GA3 -free κ-Sap resulted in significant enhancement in the photosynthetic rate accompanied by improvement of biomass yield without any compromise on the grain yield advantage bestowed by pristine κ-Sap (Figure 4.4) [22]. The result was reproducible for three consecutive seasons. The enhanced biomass production was achieved without any additional inputs of fertilizer and irrigation

72

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources Macro and micronutrients, IAA, GA3, kinetin, zeatin, GB, choline and others

25–35% Grain yield enhancement

Liquid SAP

Foliar s pray

Zea mays

Pristine κ-sap

Macro and micronutrients, IAA, kinetin, zeatin, GB, choline and others

− GA3 Elimination of gibberellin

No corn stover yield enchancement

25–35% Grain yield enhancement

Liquid SAP

Foliar s pray

GA3 free κ-sap

Zea mays 18–30% Corn stover yield enhancement

Figure 4.4 Effect of pure κ-Sap and GA3 free κ-Sap on yield and biomass production of maize. (Source: Mondal et al. 2015 [22]. Reproduced with permission of Springer.)

water, underscoring the importance of the finding in the context of sustainability. This novel finding has profound implications from the perspectives of environment and biomass energy (Figure 4.5) [22].

4.6

Bioethanol Production from Kappaphycus alvarezii

Large-scale production of bioethanol from biomass can be suitable alternatives of fossil fuels for sustainable mobility. In the past decades, many research endeavours were devoted to the ethanol production from a variety of bioresources [27]. However, production of third-generation bioethanol from unconventional seaweed biomass has started very recently [28]. Many types of seaweeds showed potential for the production of ethanol, but the use of K. alvarezii as a source of bioethanol is scant in the literature [9, 29–31]. Presence of low lignin content and high levels of easily hydrolysable polysaccharides in this seaweed makes it a suitable feedstock for the production of ethanol via fermentation. Since seaweed sugars are not simple and easily fermentable, thus pretreatment of the biomass is required to obtain fermentable sugars before fermentation. Moreover, the hydrolysate requires further processing before enzymatic fermentation to reduce its salt content and other

Kappaphycus alvarezii: A Potential Sustainable Resource for Fertilizers and Fuels

Grain Root biomass Above ground vegetative biomass

CO2 sequestered (g/plant)

400

300

72.1

73

85.0 22.6

88.3

92.8 19.4

18.5

25.9

200 317 247

242

267

Control

Pristine sap

100

0 Gibberellin free sap

IAA free sap

Treatments (a) 6000 Grain

5000

Root biomass Above ground vegetative biomass

1.27E+03

Energy (KJ/plant)

364

1.23E+03

1.31E+03

4000

306

1.07E+03 418 3000

2000

298

2.87E+03

3.18E+03

4.23E+03

3.59E+03

Gibberellin free sap

IAA free sap

1000

0 Control

Pristine sap

Treatments (b) Figure 4.5 Effect of κ-Sap variants (control, pristine κ-Sap and GA3 free κ-Sap) on (a) CO2 sequestration by maize and (b) energy content of maize plants. (Source: Mondal et al. 2015 [22]. Reproduced with permission of Springer.)

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

contaminants which have inhibitory effects on fermentation. Saccharomyces cerevisiae is a commonly used microorganism for ethanol production from K. alvarezii [9, 29, 30]. In short, the process of ethanol production from K. alvarezii can be divided into five steps as follows. 4.6.1

Pretreatment of Freshly Harvested Biomass

In most cases, the freshly harvested seaweed is washed and allowed to dry and powdered before hydrolysis, and thereby, only a single product was envisaged. However, in a recent report, the biorefinery concept was employed to yield liquid fertilizer and seaweed granules from fresh K. alvarezii seaweed. Subsequently, these granules were utilized as feedstocks for ethanol production [9]. 4.6.2

Hydrolysis of the Dry Biomass to Obtain Fermentable Sugars

The dry seaweed biomass of K. alvarezii contains κ-carrageenan (Figure 4.1) as its main component. This phycocolloid yields the fermentable sugar d-galactose upon hydrolysis. Both chemical and enzymatic hydrolysis protocols are reported in the literature, but acid hydrolysis was the most preferable because of its low cost. Meinita et al. have studied the effect of hydrochloric acid and sulfuric acid treatment for the hydrolysis of K. alvarezii dry powder to yield maximum reducing sugars (Figure 4.6) [30]. The study showed hydrolysis of K. alvarezii dry powder (100 g/L) using 0.2 M sulfuric acid at 130 ∘ C for 15 min which yielded a maximum reducing sugar concentration of 30.5 g/L, out of which galactose was 25.6 g/L. Recently, Khambhaty et al. have reported a concentration of about 70 g/L for the reducing sugars, from the seaweed granules obtained after recovering the sap [8]. The saccharification process used 0.9 N sulfuric acid at 100 ∘ C for 1 h. In a more recent study by Hargreaves et al., a hydrolysate with a maximum galactose concentration of 81.62 g/L (51% based on the d-galactose content in K. alvarezii) was obtained through dilute acid treatment (1% v/v H2 SO4 ) at 121 ∘ C for 1 h [29]. The same authors also studied the enzymatic hydrolysis of residual cellulosic biomass obtained after acid hydrolysis of K. alvarezii seaweed to maximize the overall yield of fermentable sugars. 4.6.3

Pretreatment of Hydrolysate to Reduce the Concentration of Fermentation Inhibitory Components

Although the acid hydrolysis technique is a cost-effective one, it also produces some other components such as 5-hydroxymethyl furfural (HMF) and levulinic acid (LA) during acid hydrolysis of K. alvarezii seaweed [29–31]. Such compounds are known to have inhibitory effects on microorganisms and thereby inhibit the fermentation process. Thus, it is important to remove such metabolic inhibitors

Kappaphycus alvarezii: A Potential Sustainable Resource for Fertilizers and Fuels

1.0

Galactose (g/L)

(a)

Galactose by H2SO4 Galactose by HCI Glucose by H2SO4 Glucose by HCI

6 5

0.8 0.6

4 0.4

3 2

Glucose (g/L)

8 7

75

0.2

1

14

14

12

12

10

10

8

8 6

6 Total sugar by H2SO4 Total sugar by HCI Reducing sugar by H2SO4 Reducing sugar by HCI

4 2

4 2 0

0 (c)

HMF (g/L)

2.5 2.0

HMF by H2SO4 HMF by HCI Levulinic acid by H2SO4 Levulinic acid by HCI

2.5 2.0

1.5

1.5

1.0

1.0

0.5

0.5

0.0 0.0

Reducing sugar (g/L)

0.0 16

(b)

Levulinic acid (g/L)

Total sugar (g/L)

0 16

0.0 0.2

0.4 0.6 Acid catalyst (M)

0.8

1.0

Figure 4.6 Effect of catalyst concentration on sugars and by-products in acid hydrolysis of K. alvarezii dry powder. (a) Glucose and galactose. (b) Reducing sugar and total sugar. (c) By-products (HMF and levulinic acid). (Source: Meinita et al. 2012 [30]. Reproduced with permission of Springer.)

from the hydrolysate prior to fermentation. In a recent study, removal of HMF from acid hydrolysate was studied through adsorption using activated charcoal [29]. It was observed that the concentration of HMF was decreased from 35 to 1.5 g/L upon increasing the concentration of activated carbon from 0% to 25% (w/v) with no significant variation of galactose concentration. On the other hand, Meinita et al. reported HMF removal from the hydrolysate of K. alvarezii with a significant loss of galactose (about 43.1% when 5% (w/v) of activated charcoal was used) [30]. Although the source of activated carbon was not specified in both the

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

studies [29, 30]; however, some cases of activated carbon from different sources, for example, coconut husk versus wood, can have significantly different adsorption properties, and this might be the reason for differences. Acid hydrolysis of K. alvarezii not only de-polymerizes the polysaccharide (Figure 4.1), it also releases sulfate groups and its counter ion to yield d-galactose as the fermentable sugar. Therefore, the concentration of total dissolved solids in the hydrolysate increases which also hampers the fermentation process due to high salt concentrations. Recently, Khambhaty et al. studied an electrodialysis (ED) process to remove the ionic constituents (91.5% removal) from the hydrolysate while retaining the sugar concentration relatively intact [8]. 4.6.4

Enzymatic Fermentation of the Hydrolysate to Yield Ethanol

The most common organism utilized for the production bioethanol from K. alvarezii is S. cerevisiae. Meinita et al. used commercial brewer’s yeast, freeze-dried S. cerevisiae for bioethanol production through fermentation of the hydrolysate of K. alvarezii. They also studied the ethanol production rate with time and found that in the early phase of production, the rate was slow but rapidly increased after 12 h with a maximum conversion achieved after 24 h of fermentation [30]. The overall yield of bioethanol by this process was 41% of the theoretical yield, whereas under similar conditions, pure galactose gave an ethanol yield of 63% of the theoretical value. The lower yield in the case of K. alvarezii hydrolysate was attributed to the presence of hydrolysate by-products which inhibited the fermentation process. Table 4.4 provides an overview of the ethanol productivities from K. alvarezii reported by different groups. Table 4.4

Yield and reaction parameters for the production of bioethanol from K. alvarezii. Saccharification

Condition of fermentation

Kappaphycus alvarezii granules obtained after recovery of sap Kappaphycus alvarezii dry powder

0.9 N H2 SO4 at 120 ∘ C for 60 min

Saccharomyces 92.3% cerevisiae, NCIM 3455, SHF, 96 h

15.4

[9]

1.7 g/L

1.31

[30]

38 g/L

29.28

[29]

0.2 M H2 SO4 at 130 ∘ C for 15 min Kappaphycus 1% H2 SO4 at alvarezii dry powder 121 ∘ C for 60 min

Saccharomyces cerevisiae, SHF, 24 h Saccharomyces cerevisiae, CBS1782, SSF, 12 h

Yield as reported

References Yield normalized to g EtOH/g dry seaweed (%)

Feedstocks

Kappaphycus alvarezii: A Potential Sustainable Resource for Fertilizers and Fuels

4.6.5

77

Purification of Ethanol from Fermentation Broth

After the fermentation process, a typical fermentation broth contains 6–12% ethanol while the remaining is mostly water besides other contaminants. Distillation is the most common purification process for the recovery of ethanol [32]. However, due to the formation of an azeotropic mixture, a significant amount of energy is required for distillation. Various azeotrope separation components such as benzene, hexane and diethyl ether are used to separate the azeotrope mixture to enhance the ethanol recovery. In large-scale plants, recovery of bioethanol through distillation columns varies in the range of 95–99.6% [33].

Fuel Intermediates and Useful Chemical from Kappaphycus alvarezii

4.7

In the course of saccharification to improve ethanol yields from the K. alvarezii, HMF and LA were formed as by-products which are known to have inhibitory effect on fermentation. Many studies have explored their effective removal in order to enhance the fermentation process [29–31]. However, instead of suppressing these impurities, there may be greater merit in producing these compounds more selectively instead of ethanol since biofuels obtained via HMF and LA pathway have better quality compared to ethanol in terms of energy density. In this direction, several research efforts have been made [33]. Kim et al. studied the production of HMF and LA derivative as biofuels from Gelidium amansii, a red algae [34]. The process utilized an ionic liquid supported solid acid catalyst in the presence of CrCl2 (Scheme 4.1). The reaction was performed in the presence of ethanol to yield ether and ester derivatives of HMF and LA, respectively. Regardless of these technical advances, production of biofuel alone from seaweed is not advantageous in the context of its high carbon footprint and economy of the overall process. However, the economics of HMF production could be more attractive if the process was integrated with simultaneous generation of other value-added products having direct practical applications. K. alvarezii seaweed is ideally suited for such an approach because of its unique composition (Figure 4.1). Recently, gainful utilization of this fresh seaweed in a biorefinery O O

O OH

OH

O O

O OH

CrCI2/ [Emim]CI O O OH

O n

SO3H CH3CH2OH

O

+ O

O 30% (Isolated) EMF/LAEE = 5:2

Scheme 4.1 Scheme for the production of biofuel from Gelidium amansii. (Source: Kim et al. 2010 [34]. Reproduced with permission of ChemSusChem.)

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

Figure 4.7 Integrated process for the production of fuel intermediates, fertilizers and freshwater from fresh K. alvarezii seaweed. (Source: Mondal et al. 2013 [11]. Reprodued with permission of Royal Soceity of Chemistry.)

approach was explored by Mondal et al., and a self-sustainable integrated scheme was proposed for the concomitant production of HMF, LA, formic acid, potash fertilizers, potable water and fortified sap as liquid fertilizers [11]. In the process, heating the seaweed in the presence of 0.48 M Mg(HSO4 )2 at 105 ∘ C for 1 h was found to be optimum to yield HMF with 61% carbon conversion efficiency from the κ-carrageenan present in the seaweed. The remaining carbon was mainly galactose which was further converted to LA and formic acid. The overall carbon conversion efficiency combining HMF, LA and formic acid was reported to be 83%. The integrated scheme is shown in Figure 4.7 [11]. The processing of 1 ton of granule would yield 0.18 ton HMF, 0.056 ton LA, 0.020 ton FA, 0.27 ton K2 SO4 and 5.77 m3 pure water. The process energy requirement for the overall process of Figure 4.7 was proposed to be met from additional supplies of granule through combustion of this biomass which additionally yielded 0.74 ton glaserite fertilizer and the required amount of sulfuric acid for Mg(HSO4 )2 preparation. In a very recent study, Lee et al., for the production of platform chemicals (HMF and LA) and sugar (galactose and glucose) from K. alvarezii, investigated the use of a dilute-acid-catalyzed hydrothermal reaction [35]. The study showed that yield and selectivity of

Kappaphycus alvarezii: A Potential Sustainable Resource for Fertilizers and Fuels

79

the different products could be monitored preferentially by changing the acid concentration, reaction temperature and reaction time. Large-scale production platform chemicals and sugars from K. alvarezii will have potential in biofuels and chemical industries.

4.8

Environmental Impact of Fuel and Fertilizers Production from Kappaphycus alvarezii

Farming and harvesting of seaweed have some environmental advantages and disadvantages as well. Cultivation of seaweeds does not require any nutrition, pesticides and land, and sometimes, seaweed farming improves the water quality in the area where it is grown. Seaweed also helps to fix dissolved nitrogen and phosphorus in the sea [36]. Commercial level of cultivation of K. alvarezii has been started along the south-east coast of India, and the coastal communities at large have benefited from this seaweed farming in terms of their socio-economic development because the sap extracted from K. alvarezii has been marketed as a potent plant stimulant. However, whether the production method is environmentally sustainable or not still needs to be addressed. Ghosh et al. carried out the overall life-cycle assessment for the sap production from this seaweed and showed that the production of 1 kL of seaweed extract, at factory gate, from the fresh biomass of K. alvarezii grown onshore under open-sea conditions has a low carbon footprint of 118.6 kg CO2 equivalent [37]. No study so far has attempted to assess the environmental sustainability and energy requirement for the production of bioethanol from K. alvarezii. However, Khambhaty et al. envisaged that the downstream processing of seaweed granules into bioethanol was energy-intensive [8]. However, co-production of several other valuable chemicals and fertilizers in a biorefinery approach would minimize the overall economy of the process.

4.9

Conclusion and Future Prospect

It has been observed from the reports discussed earlier that K. alvarezii is a suitable precursor for the production of a number of chemicals and fuels. Considering the economic and environmental advantages of the seaweed for the production of chemicals of practical application, there is a possibility for the development of a number of downstream processing industries. To sustain the supply of the seaweed for large-scale commercial exploitation, a viable method for the cultivation of the seaweed must be developed.

Acknowledgement This is CSIR-CSMCRI Communication No. 046/2016. KP thanks the Council of Scientific and Industrial Research (CSIR, New Delhi) for the Award of

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

CSIR-Young Scientist Awardees Project and overall financial support. DM thanks CSIR for senior research fellowship. The Centralized Instrument Facility of the Institute is acknowledged for providing the instrumental facilities for carrying out the research work.

References 1. Farrell, A.E., Plevin, R.J., Turner, B.T. et al. (2006) Ethanol can contribute to energy and environmental goals. Science, 311, 506–508. 2. Wi, S.G., Kim, H.J., Mahadevan, S.A. et al. (2009) The potential value of the seaweed Ceylon moss (Gelidium amansii) as an alternative bioenergy resource. Bioresource Technology, 100, 6658–6660. 3. Raymundo-Pinero, E., Cadek, M. and Béguin, F. (2009) Tuning carbon materials for supercapacitors by direct pyrolysis of seaweeds. Advanced Functional Materials, 19, 1032–1039. 4. Lee, K.Y. and Mooney, D.J. (2012) Alginate: properties and biomedical applications. Progress in Polymer Science, 37, 106–126. 5. McHugh, D.J. (2003) A Guide to the Seaweed Industry, FAO Fisheries Technical Paper. No. 441, FAO, Rome. http://www.fao.org/3/a-y4765e.pdf. 6. Rowbotham, J.S., Dwyer, P.W., Greenwell, H.C. and Theodorou, M.K. (2012) Thermochemical processing of macroalgae: a late bloomer in the development of third-generation biofuels? Biofuels, 3, 441–461. 7. Rhein-Knudsen, N., Ale, M.T. and Meyer, A.S. (2015) Seaweed hydrocolloid production: an update on enzyme assisted extraction and modification technologies. Marine Drugs, 13, 3340–3359. 8. Khambhaty, Y., Mody, K., Gandhi, M.R. et al. (2012) Kappaphycus alvarezii as a source of bioethanol. Bioresource Technology, 103, 180–185. 9. Aresta, M., Dibenedetto, A. and Dumeignil, F. (2012) Biorefinery from Biomass to Chemicals and Fuels, De Gruyter, Berlin, Germany. 10. Lewis, J.G., Stanley, N.F. and Guist, G.G. (1990) in Algae and Human Affairs (eds C.A. Lembi and J.R. Waaland), Cambridge University Press, Cambridge, p. 218. 11. Mondal, D., Sharma, M., Maiti, P. et al. (2013) Fuel intermediates, agricultural nutrients and pure water from Kappaphycus alvarezii seaweed. RSC Advances, 3, 17989–17997. 12. Li, L., Ni, R., Shao, Y. and Mao, S. (2014) Carrageenan and its applications in drug delivery. Carbohydrate Polymers, 103, 1–11. 13. Kumar, K.S., Ganesan, K. and Subba Rao, P.V. (2015) Seasonal variation in nutritional composition of Kappaphycus alvarezii (Doty) Doty—an edible seaweed. Journal of Food Science and Technology, 52, 2751–2760. 14. Prajapati, V.D., Maheriya, P.M., Jani, G.K. and Solanki, H.K. (2014) Carrageenan: a natural seaweed polysaccharide and its applications. Carbohydrate Polymers, 105, 97–112. 15. Das, A.K., Sharma, M., Mondal, D. and Prasad, K. (2016) Deep eutectic solvents as efficient solvent system for the extraction of-carrageenan from Kappaphycus alvarezii. Carbohydrate Polymers, 136, 930–935. 16. Abbott, A.P., Capper, G., Davies, D.L. et al. (2003) Novel solvent properties of choline chloride/urea mixtures. Chemical Communications, 2003, 70–71. 17. Craigie, J.S. and Leigh, C. (1978) in Handbook of phycological methods (eds J.A. Hellebust and J.S. Craigie), Cambridge Univ. Press, Cambridge, pp. 109–131.

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18. Christopher, S.R. and Michael, G.B. (1998) Method for extracting semi-refined carrageenan from seaweed. Patent no. US5801240, 6 March. 19. Chapman, V.J. and Chapman, D.J. (1980) Seaweeds and Their Uses, Chapman and Hall, London & New York, Chapter 2, pp. 30–61. 20. Eswaran, K., Ghosh, P.K., Siddhanta, A.K., Patolia, J.S., Periyasamy, C., Mehta, A.S., Mody, K.H., Ramavat, B.K., Prasad, K., Rajyaguru, M.R., Kulandaivel, S., Reddy, C.R.K., Pandya, J.B. and Tewari, A. (2005) Integrated method for production of carrageenan and liquid fertilizer from fresh seaweeds. U.S. Patent 6893479, May 17. 21. Prasad, K., Das, A.K., Oza, M.D. et al. (2010) Detection and quantification of some plant growth regulators in a seaweed-based foliar spray employing a mass spectrometric technique sans chromatographic separation. Journal of Agriculture & Food Chemistry, 58, 4594–4601. 22. Mondal, D., Ghosh, A., Prasad, K. et al. (2015) Elimination of gibberellin from Kappaphycus alvarezii seaweed sap foliar spray enhances corn stover production without compromising the grain yield advantage. Plant Growth Regulation, 75, 657–666. 23. Rathore, S.S., Chaudhary, D.R., Boricha, G.N. et al. (2009) Effect of seaweed extract on the growth, yield and nutrient uptake of soyabean (Glycine max) under rainfed conditions. South African Journal of Botany, 75, 351–355. 24. Craigie, J.S. (2011) Seaweed extract stimuli in plant science and agriculture. Journal of Applied. Phycology, 23, 371–393. 25. Miao, Z., Yu, X., Wang, Y., Miao, H. and Tang, K. (2007) Seaweed amino acid rare-earth compound foliage fertilizer and its preparing method, CN20061117577 20061026. 26. Weiss, D. and Ori, N. (2007) Mechanisms of cross talk between gibberellin and other hormones. Plant Physiology, 144, 1240–1246. 27. Sanchez, O.J. and Cardona, C.A. (2008) Trends in biotechnological production of fuel ethanol from different feedstocks. Bioresource Technology, 99, 5270–5295. 28. Jiang, R., Ingle, K.N. and Golberg, A. (2016) Macroalgae (seaweed) for liquid transportation biofuel production: what is next? Algal Research, 14, 48–57. 29. Hargreaves, P.I., Barcelos, C.A., da Costa, A.C.A. and Pereira, N. Jr., (2013) Production of ethanol 3G from Kappaphycus alvarezii: evaluation of different process strategies. Bioresource Technology, 134, 257–263. 30. Meinita, M., Kang, J.-Y., Jeong, G.-T. et al. (2012) Bioethanol production fromthe acid hydrolysate of the carrageenophyte Kappaphycus alvarezii (cottonii). Journal of Applied Phycology, 24, 857–862. 31. Dyah, M., Meinita, N., Hong, Y.-K. and Jeong, G.-T. (2012) Comparison of sulfuric and hydrochloric acids as catalysts in hydrolysis of Kappaphycus alvarezii (cottonii). Bioprocess & Biosystem Engineering, 35, 123–128. 32. Balat, M., Balat, H. and Öz, C. (2008) Progress in bioethanol processing. Progress in Energy and Combustion Science, 34, 551–573. 33. Chheda, J.N., Leshkova, Y.R. and Dumesic, J.A. (2007) Production of 5-hydroxymethyl furfural and furfural by dehydration of biomass-derived mono- and poly-saccharides. Green Chemistry, 9, 342–350. 34. Kim, B., Jeong, J., Shin, S. et al. (2010) Facile single-step conversion of macroalgal polymeric carbohydrates into biofuels. ChemSusChem, 3, 1273–1275. 35. Lee, S.-B., Kim, S.-K., Hong, Y.-K. and Jeong, G.-T. (2016) Optimization of the production of platform chemicals and sugars from the red macroalga, Kappaphycus alvarezii. Algal Research, 13, 303–310.

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36. Ghadiryanfar, M., Rosentrater, K.A., Keyhani, A. and Omid, M. (2016) A review of macroalgae production, with potential applications in biofuels and bioenergy. Renewable and Sustainable Energy Reviews, 54, 473–481. 37. Ghosh, A., Vijay, A.K.G. and Seth, A. (2015) Life cycle impact assessment of seaweed based biostimulant production from onshore cultivated Kappaphycus alvarezii (Doty) Doty ex Silva—Is it environmentally sustainable? Algal Research, 12, 513–521.

5 Microalgae Bioproduction – Feeds, Foods, Nutraceuticals, and Polymers Clifford R. Merz1 and Kevan L. Main2 1 University

2

5.1

of South Florida, College of Marine Science, USA Marine & Freshwater Aquaculture Research Program, Mote Marine Laboratory, USA

Introduction

Consideration of using aquatic organisms for biofuel feedstock applications has been discussed since the mid-twentieth century and for nutritional needs since the early 1900s. These aquatic organisms typically fall into the categories of microalgae and macroalgae (also known as seaweed). Of the two, microalgae produce more of the desirable types of natural oils required for renewable biofuels [1], aquaculture feeds for protein sustainability, human nutraceutical uses, as well as additional material and chemical coproduct capabilities. For this reason, we have tailored this chapter’s discussion to focus primarily on bioproduction of microalgae. In comparison with other forms of renewable energy, such as wind, tidal, wave, solar, and salinity gradients [2–4], which primarily target electrical energy production, liquid biofuels allow solar/chemical energy to be stored and used in existing engines and transport infrastructure [5]. There has been a significant Fuels, Chemicals and Materials from the Oceans and Aquatic Sources, First Edition. Edited by Francesca M. Kerton and Ning Yan. © 2017 John Wiley & Sons Ltd. Published 2017 by John Wiley & Sons Ltd.

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amount of literature published on the production of oil from microalgae. Algal oil (also referred to as single-cell oils (SCO)) and its conversion to useable biofuels has been demonstrated repeatedly [6–8], but commercial production remains elusive. According to the US Department of Energy’s May 2010 National Algal Biofuels Technology Roadmap [9], (micro)algae is the preferred feedstock for high energy density, fungible, liquid transportation fuels. Following are several of the attractive aspects provided by microalgae for fuel production: • Algal productivity can offer high biomass yields per acre of cultivation. • Algae cultivation strategies can minimize or avoid competition with arable land and nutrients used for conventional agriculture. • Algae can utilize wastewater, reclaimed water, and saline water, thereby reducing competition for limited freshwater supplies. • Algae can recycle carbon from CO2 -rich flue emissions from stationary sources, including power plants and other industrial emitters. • Algal biomass is compatible with the integrated biorefinery vision of producing a variety of fuels and valuable coproducts. In a recent paper, Merz and Main [10] explored various water–energy–food nexus synergies between the aquaculture and biofuel sectors with respect to microalgae (diatom) production, harvesting and processing technologies. We explored possible links to coproduct development and where improvements in manufacturing and scalability could provide the push to further develop and solidify mutual commercial algal production technologies. Using an interdisciplinary systemwide approach, markets for jointly developed main products and coproducts were examined. For example, in the biofuel sector, the main product is the algae oil used to produce biofuel, with the remaining algae protein and carbohydrate meal being used to offset and improve the overall bottom-line production costs through the coproduct application as livestock feeds, whereas in the aquaculture sector, the main product is farmed fish, with algal meal used as a nutritional protein resource for the fish, thereby reducing the dependence on dwindling wild capture fisheries. Any residual oils that are not needed to meet the fish nutritional requirements will provide another coproduct to offset and improve the overall bottom-line production costs through coproduct applications, such as algal-oil-rich ω-3 long carbon [C] chain (≥C20)–polyunsaturated fatty acids (LC-PUFA) used in human nutraceutical supplements. Improvements in microalgae production technology will benefit both of these large and important industry sectors, in addition to other sectors such as the emerging biopolymers sector. This chapter continues to advance a multi-faceted synergistic nexus approach by presenting a survey focused on microalgae production with respect to specific major feedstock products, coproducts, and applications. A background review of traditional production methods and novel methods to grow microalgae in the

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marine environment is presented in Section 5.2. A discussion of major feedstock products and coproducts is provided in Section 5.3, including feeds, foods, nutraceuticals, and biopolymers. Finally, the chapter content is summarized along with directions for future work.

5.2

Microalgae and Bioproduction Methods

Mass production of microalgae is accomplished in a variety of production units that are located outdoors (in ponds, tanks or raceways, tubes, bags, or photobioreactors (PBRs)) or indoors, inside greenhouses or buildings (in tubes, tanks, bags, PBRs). Often, outdoor axenic microalgae cultures are started with cultures that are initially grown indoors, where predators can be excluded and nutrients and environmental conditions can be controlled, and then they are moved to outdoor tanks or raceways for mass production. Production of microalgae requires nitrogen and phosphorus, in an approximate ratio of 6:1, respectively, by weight, [11]; diatoms also require silicate. Trace minerals (iron, copper, zinc, cobalt, manganese, and molybdenum) and vitamins (especially B12 , thiamine, and sometimes biotin) are also needed in most axenic cultures. Other additives, such as chelating agents, may also be needed to culture certain species of algae [12, 13]. Guillard’s (F/2) medium [14] is the standard for making stock solutions to grow single-species marine algae cultures; it includes the major nutrients, trace elements, and vitamins. Due to cost restrictions, pond culture of microalgae is usually supplied with agricultural-grade fertilizer [15]. The economic feasibility of any microalgae bioproduction enterprise will vary greatly depending on the following: • • • • •

Algal type and species selected Location of the algal cultivation facility Production method and scale selected for use Source, quality, and cost of the input water and nutrients used Target market and price of the feedstock and coproducts produced. The following subsections discuss these topics in more detail.

5.2.1

Microalgae Groups Considered

Microalgae are the most primitive form of plants and are important for life on Earth, accounting for the production of nearly half of the atmospheric oxygen and serving as the base of the food web. These unicellular species exist individually or in chains or groups depending on the species, and their sizes can range from a few micrometers (μm) to a few hundreds of micrometers. In addition, microscopic microalgae, typically found distributed throughout the water column in freshwater, brackish, and marine environments, are specially adapted to environments

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dominated by viscous forces [16]. Taxonomically, microalgae are usually found grouped in the following four major classes [1]: 1. Diatoms (Bacillariophyceae) – Well over 100,000 species are known to exist [17] in marine, brackish, and freshwater environments and are estimated to be responsible for 20% of global carbon fixation, making them the dominant primary producers in the ocean [18, 19]. Diatoms contain polymerized silica in their frustules (cell walls) and store carbon in the form of natural oils (especially under silicon [Si] limitation [20]) or as a polymer of carbohydrates known as chrysolaminarin. 2. Green algae (Chlorophyceae) – Highly abundant, especially in freshwater. The main storage compound is starch, although oils can be produced under certain conditions. 3. Blue-green algae (Cyanophyceae) – Found in a variety of environments, the nearly 2000 species are closer to bacteria in structure and organization and play an important role in fixing nitrogen from the atmosphere. 4. Golden algae (Chrysophycea) – Approximately 1000 known species, this group is similar to diatoms, producing natural oils and carbohydrates as storage compounds. The majority of the organisms collected and studied by the biofuels sector to date fall into the top two classes – the diatoms and the green algae. However, the biodiversity of microalgae is enormous, and they represent an almost untapped resource. Most of these microalgae species produce unique products such as vitamins, minerals, proteins, pigments, carbohydrates, carotenoids, antioxidants, polyunsaturated fatty acids, enzymes, polymers, peptides, toxins, and sterols [21, 22]. 5.2.2

Bioproduction of Microalgae – Methods

Most algal production systems to date have focused on natural growth conditions using phototrophic organisms because of their ability to use readily accessible solar energy and CO2 (atmospheric and/or flue gas emissions) and nutrients from the aquatic habitats to create biomass. In specific cases, heterotrophic organisms (grown without sunlight and supplied alternate organic compounds, such as acetate, glucose, or crude glycerol (a by-product from the biodiesel transesterification process [23]) as the carbon source) have been utilized for targeted production of biomass feedstock and selected coproducts/metabolites. A challenge for the mass production of any biological system is maintaining adequate levels of production, while securing sustainable and cost-effective supplies of nutrients (nitrogen [N], phosphorous [P]), C, and Si (for diatoms) to maintain continuous growth [24]. Concentrations of cells in phytoplankton microalgal culture media are generally higher than those found in nature; therefore, they must be enriched with nutrients to make up for any deficiencies found

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in the water source used. When macronutrients are not limited, the Redfield molar element ratio C:N:P is 106:16:1 for most phytoplankton. Diatoms need, among other nutrients, silicic acid to create biogenic silica for their frustules (cell walls). As a result, the Redfield–Brzezinski nutrient ratio was proposed for diatoms and stated to be C:Si:N:P = 106:15:16:1 [25]. Micronutrients consist of various trace metals and vitamins, such as B1 (thiamin), B12 (cyanocobalamin), and biotin. Microalgae are also capable of utilizing nutrients, such as N and P, from a variety of alternate wastewater sources (e.g., agricultural runoff, concentrated animal feed operations, and industrial and municipal wastewater), thus providing an additional option for sustainable bioremediation of wastewater for environmental and economic benefits [26]. Aside from nutrient quality and quantity, significant parameters important in regulating microalgal growth include the following: illumination (in phototrophic growth conditions) intensity and duration, pH, turbulence, salinity, and temperature. The growth dynamics of an axenic culture, versus a culture containing more than one organism, is characterized by the following five phases [27, 28]: 1. Initial induction phase: Characterized by little increase in cell density. The length of this phase can be relatively long if the algal culture is transferred from a plate or shortened if the liquid culture is inoculated with microalgae already in phase II growth. 2. Rapid growth rate phase: Characterized by a rapid increase in the cell density as a function of time in accordance with logarithmic equation 5.1. Ct = C0 emt

(5.1)

where Ct and C0 are the cell concentrations at time t and 0, respectively, and m is the specific species growth rate. 3. Declining growth rate phase: Characterized by a reduction in cell division when physical/chemical factors begin to limit growth. 4. Production (stationary) phase: Characterized by a relatively constant cell density when the growth rate and limiting factors are balanced. 5. Final death or “crash” phase: Characterized by a deterioration of the significant nutrients and parameters important in regulating growth to a level incapable of sustained growth, resulting in a rapid decrease in cell density and eventual culture collapse. During the initial inoculation and early growth period, the cell density of the desired microalgae species is low and concentration of nutrients is high, so caution must be exercised to prevent introduction of any contaminant with a faster growth rate capable of outgrowing the culture. Successful production of any microalgal culture is its maintenance in the upper regions of the exponential growth phase. Unless precautions are taken to ensure the quality of microalgae produced (i.e., see Section 5.2.2.1, perfusion culture discussion), the nutritional value of the microalgae will begin to degrade after the culture moves beyond

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phase 3 because of reduced digestibility, deficient composition, and possible production of toxic metabolites [27]. There are a wide variety of systems currently in use for microalgal production that are run in batch, continuous, or semicontinuous (fed-batch) modes. Large-scale microalgae cultivation occurs in open systems (ponds, raceways, and lagoons) and indoor/outdoor closed systems (tanks, PBRs, or bags). Small-scale, closed cultivation methods using biopharmaceutical fermenters, membrane bioreactors, and perfusion technologies have been used for heterotrophic algal growth and specific single-use by-products. Recently, open integrated multitrophic aquaculture systems, based on the principals originally used in China’s early polyculture pond systems, are gaining prominence. The following sections discuss production methods typically in use, under development, or under re-evaluation for the bioproduction of microalgae along with the corresponding production scales of each. 5.2.2.1

Batch, Continuous, Fed-batch Semicontinuous, and Perfusion Culture Methods

In the batch culture method, a single inoculation of microalgal cells are placed into a growth chamber containing fertilized culture (e.g., filtered seawater, treated wastewater) followed by a specific growth period with final harvesting when the algal population reaches its maximum or near-maximum density (Phase 2/3). Depending upon the ultimate production scale, batch culture systems are often used because of their simplicity, repeatability, and flexibility to rapidly changing species and remedy culture shortcomings. In large production batch operations, the culture volume is grown in a stepwise progression toward increasingly larger culture volumes until the largest tank size is reached and the culture volume has achieved a maximum density and is harvested. Although repeatable and fairly straightforward, large production batch systems entail multiple repetitions of the first three growth phases, leading to increases in time and labor to harvest, clean, sterilize, refill, and inoculate each culture. In addition, caution must be exercised with regard to timing the individual harvests, as well as preventing culture contamination during initial inoculation and the early growth period of each repetition; otherwise, the harvest cell quality may be less predictable than in continuous systems [27, 28]. In the continuous culture method, a continuous stream(s) of fertilized culture medium is fed into a growth chamber, while excess effluent (containing, among others, microalgae, coproducts/metabolites, and residuals) is simultaneously removed and collected and harvested [29]. By maintaining an equal volumetric flow rate into and out of the growth chamber, a steady-state culture volume is established; thereby enabling the culture maintenance to remain very close to the phase 2/3 maximum growth rate. Several different methods for continuous

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culture systems are currently used, including the turbidostat and chemostat culture systems [27, 28]: • Turbidostat culture system: An automated operation in which cell density in the growth chamber is maintained at a preset (fixed) level via dilution of the culture with additional fresh medium. • Chemostat culture system: An automated operation in which cell growth rate in the growth chamber is maintained at a constant preset (fixed) level via addition of fresh medium possessing a limiting vital nutrient (e.g., N or Si). Advantages of the continuous culture system approach include single or modular tank configurations capable of producing a product with a more predictable and repeatable quality, as well as a production system suited to technological control and automation, which in turn increases the overall system reliability and bottom-line economics. Disadvantages include difficulty in maintaining a sterile culture over a prolonged time, and as such, with the exception of a few specific processes (e.g., single-cell protein production, certain beer production, and municipal waste treatment), continuous cultures have not been widely adopted by the industry [29]. Fed-batch is a commonly used technique in the fermentation industry. A fed-batch semicontinuous culture method is an indoor or outdoor batch system in which the required nutrient stream(s) is fed intermittently or continuously; following partial, periodic harvests, the system is immediately refilled with fresh culture. This technique prolongs the use of large tank cultures, allowing the culture to be grown, partially harvested, and regrown [29]. Advantages include greater yields of microalgae than with the batch method, for a given tank size, due to reduced turnaround time [27, 28]. Disadvantages include unpredictable duration due to an increase in microalgal competitors, predators and/or contaminates, and metabolites, which eventually causes the culture to crash. Perfusion culture methods are derived from biopharmaceutical cell-culture techniques. This method has received renewed interest in microalgal bioproduction, because of its potential for higher cell densities and productivity in a relatively small-size bioreactor when compared to batch and fed-batch culture systems. The perfusion culture method is essentially a fed-batch semicontinuous turbidostat/chemostat process where necessary nutrient streams(s) are fed into the growth chamber/bioreactor, during cell growth and production phases, at dilution rates exceeding cellular growth rate. However, it differs by the addition of an in-line cell retention (i.e., reinoculation) device(s) that provides a connection between the bioreactor and retention device where algal cells can be retained and the spent medium (cell-free) is removed from the bioreactor. The goal is to eliminate substrate limitations and inhibit or remove by-products in high-cell-density cultures. This system is needed because when there is a high cell density, the accumulation of toxic metabolites will begin to limit further cell growth. Many cell retention devices have been tried with

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varying degrees of success, hampered historically by logistical and validation complexity, scalability limitations, or lack of adoption by industry. Recently designed perfusion culture systems are targeting larger operational scales (i.e., biorefinery) with the promise of higher productivity and lower failure rates using cell retention devices, such as multiport settling tanks, spin filter, or emerging alternating tangential flow (ATF) membrane technology [30–34]. Additional work is needed before this technology will be more widely applied in microalgae production. 5.2.2.2

Indoor/Outdoor and Open/Closed Bioproduction Systems

Extensive experience exists in the operation and engineering of open tanks, ponds, and raceways for microalgae production; large outdoor ponds have been successfully used to produce microalgae for centuries. Pond bottoms are either lined or earthen, with culture depths typically ranging from 0.25 to 1 m. Open raceway ponds for mass culture of microalgae have been used since the 1950s. A typical open-air raceway pond is made of a recirculation loop channel that is typically about 20–50 cm deep. Mixing and circulation are produced by a continuously revolving paddlewheel. Flow is guided around bends by baffles placed in the flow channel. During the day, cultures are fed continuously in front of the paddlewheel where the flow begins. Broth is harvested behind the paddlewheel, upon completion of the circulation loop. Indoor/outdoor PBRs, such as open raceway systems, are typically land-based; however, PBRs are closed systems. The microalgal broth (culture and growth medium) is continually circulated from a reservoir, through the closed array tubes, and back to a reservoir, thus, reducing the evaporative water losses and assisting the ecosystem balance through efficient mixing. Conceptually, PBRs consist of an array of transparent rigid plastic or glass tubes that contain the algal culture and allow light (natural or artificial) to be captured and delivered to the microalgal broth. The tubes are generally 0.1 m or less in diameter, sized to minimize temperature increases and maximize light penetration into the dense broth of a highly productive biomass. Outdoor ponds and raceways are perceived to be less expensive than PBRs because they cost less to build and operate; however, they typically occupy large amounts of land and produce less biomass when compared with PBRs [35]). Monocultures from indoor systems are often used to inoculate the PBRs and outdoor systems, but monocultures are difficult to maintain outdoors due to contamination by local algal species, predatory organisms, such as chytrid parasites [36], and rotifer grazers [37, 38]. Exceptions to these contamination problems include a few fast-growing algal species or those capable of living in extreme conditions, such as high salinity or pH. In addition, high evaporation rates in arid environments (such as the southwestern United States where evaporation in open algae ponds can reach up to 2 m of water annually [34, 39])

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and significant environmental variations (e.g., temperature, humidity, sunlight) can lead to unpredictable culture crashes or poor batch-to-batch consistencies. PBR advantages include successful production of large quantities of single-species microalgal biomass for prolonged durations [40–42], along with higher surface-to-volume ratios that can support higher volumetric cell densities in the circulating biomass. Challenges to this production method include the need for temperature monitoring and maintenance without evaporative cooling, periodic cleaning of biofilm formation and scaling, and general fluid pumping monitoring and maintenance needs, which can increase long-term facility operation and maintenance costs. Regardless of the microalgal production system, recovery and drying of microalgae biomass from the algal broth are necessary for most coproduct bioproduction enterprises. Biomass is recovered from the broth by various means, such as centrifugation followed by drying. Recovery cost is often a significant portion of the overall production cost. Biomass recovery costs from PBR cultured broth is a fraction of the recovery cost for broth produced in open ponds or raceways, which can lead to an improvement in the overall production cost. This is because typical algal biomass produced in PBRs is nearly 30 times the biomass (average open-system production of 0.14 g/L compared to 4 g/L in closed systems) obtained in open raceways [35]. Thus, in comparison with raceway broth, a much smaller volume of PBR broth needs to be processed to obtain an equal quantity of biomass. 5.2.2.3

Novel Bioproduction Systems Under Development or Re-evaluation

5.2.2.3.1 Integrated Aquaculture System (IAS) In an integrated aquaculture system (IAS), different types of organisms (i.e., fish and plants) are grown in the same water system and utilize the resources (i.e., nutrients) or services (i.e., filtered water) of the other organisms being produced. Microalgae production can be produced in an IAS if a recirculating aquaculture system (i.e., fish farm) is linked to a microalgae production system with the filtered nutrients (i.e., N, P) produced by the fish and the clean water produced by the microalgae being returned to the fish tanks in a batch-fed semicontinuous perfusion culture process. IAS technology was developed based on the principals originally used in China’s early polyculture pond systems [43], where fed species are grown together with extractive species that utilize waste produced by the fed species. Both fed and extractive species components have commercial value, and the overall sustainability of the production system is improved. In a fish/microalgae/clam IAS, several species of microalgae were produced and evaluated as a food source for bivalves (European little-neck clams [44]). Some microalgae species grow better in IAS correlated with factors such as (i) nitrogen concentration, (ii) form of nitrogen, and (iii) N:P ratio in the fish effluent. Another approach is to grow microalgae using wastewater from the fish system and to harvest and process the microalgae

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as a food resource for the fish. The algae grown in the IAS can secure sustainable supplies of nutrients from the fish wastewater. However, constraints to large-scale production of microalgae in IAS include identifying algal species that can adapt to the fish effluent characteristics (pH, temperature, salinity, nutrient levels, etc.). 5.2.2.3.2 Dialysis Membrane Reactors Dialysis-based membrane bioreactors possess a unique in situ separation capability lacking in other bioreactors. This capability is a combination of cell retention and selective inhibitory metabolic product removal using passive filtration. The method dates back to 1896 [45], and a resurgent re-evaluation holds promise for algae cultivation [34]. Dialysis membranes have been used extensively in numerous reactor applications for bacterial cultivation, production, and harvesting in laboratory and batch fermenter–continuous reservoir systems [46, 47]. Commonly cultured axenic microalgae species cultivated in dialysis cultures include Scenedesmus obliquus, Phaeodactylum tricornutum, Skeletonema costatum, Microcystis aeruginosa, Chlorella pyrenoidosa, Thalassiosira pseudonana, Thalassiosira fluviatilis, and Cyclotella cryptica [34, 47–53]. Early studies have shown that algal cell density achieved via dialysis culture exceeded that of suspended cultures in natural environments [49] and wastewater [51, 54]. The dialysis membrane reactor consists of a tubular, semipermeable membrane containing the microalgae within and allows for transport of nutrients and growth factors into and metabolic products out of the reactor through pores in the membrane wall. Nutrient transport occurs passively via diffusion-driven concentration gradients, thus requiring minimal energy. Those who have tried dialysis cultures at pilot scale have faced several challenges: • Reducing the cost of the dialysis bags. • The tiny pores in the bags can be easily clogged due to fouling by the cultured algae and/or by the bath solution that the bag is immersed in. Pore clogging quickly retards diffusion in some systems and limits the ability to recycle the bags. • Transport of nutrients to algae within the dialysis bag is limited by the rate of diffusion through the pores in the bag. For any given dimensions, the larger the bag gets, the smaller the surface-to-volume ratio. That problem can be partially overcome if the surrounding liquid and the internal contents of the bag are constantly mixed. The only way to really overcome that limitation is to increase the surface-to-volume ratio (i.e., using a very long and thin tube as the dialysis bag). Solving these and other related issues will require new approaches as well as consideration of other microalgal species that may work better than those (i.e., Scenedesmus) already evaluated to date. The benefits of dialysis membrane reactors are the ability to protect the desired strain from biological contaminants

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(such as predators, competitors, and diseases), the ability to continually refresh the culture medium, and the prospect of producing a concentrated culture in a large reservoir (such as a nutrient-rich lake or ocean). If microalgae cultivation is integrated with wastewater treatment processes, denser cultures may achieve better nutrient removal efficiency [55] and require a smaller reactor footprint to achieve similar yields. Denser cultures require less energy and fewer inputs to dewater, which also improves the overall bottom-line process economics. 5.2.2.3.3 Offshore Membrane Enclosures for Growing Algae (OMEGA) An innovative proposed approach to naturally dewatering the algal broth in a nonterrestrial-based PBR system can be found in a 2009–2013 project developed, patented, and licensed by NASA and supported by the California Energy Commission entitled “Offshore Membrane Enclosures for Growing Algae” (OMEGA). OMEGA is a system for cultivating microalgae using wastewater contained in large, floating PBRs deployed in marine environments offshore coastal communities with adjacent wastewater outfalls [56]. The flow-through system consists of tubular PBRs made of flexible, transparent, linear low-density polyethylene. These PBRs contain freshwater microalgae growing in the wastewater that use energy from the sun, carbon dioxide and nutrients from the wastewater to produce biomass that can be converted into biofuels as well as other useful products (i.e., fertilizer). The floating OMEGA system, used in either batch or continuous-flow mode, employs forward osmosis (FO) across tubular membrane walls to passively concentrate nutrients and dewater the microalgae culture broth [57], with the salinity gradient between the inner low-salinity wastewater and the outer high-salinity seawater providing the osmotic gradient for driving water transport across the semipermeable membrane walls and into the seawater. The use of floating membrane bags in protected ocean bays surrounded by a real-time coastal monitoring system [58] has several additional advantages [59]: • Structural support is provided by the buoyancy of the surrounding seawater. • Surrounding seawater provides a heat sink to prevent overheating, which can be an issue with terrestrial-based PBRs [60]. • Surface-wave motion helps to mix entrained microalgal broth. • Access to local wind/water/wave conditions will assist in maintaining operation and logistical support. Additional information on the evolving OMEGA system can be obtained from the OMEGA Global Initiative (OGI), a nonprofit 501(c) (3) corporation founded to identify OMEGA sites, mobilize local OMEGA developers, mitigate developmental risks, and facilitate successful OMEGA implementation, operation, and global commercialization [61].

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Microalgae Feedstock Products and Coproducts

In order to be digested by humans and other nonruminant animals, microalgae must first be processed to break down the algal cell wall and, thus, allow digestive enzymes to make the protein accessible. Nutritional analyses have shown that the protein in microalgae is well balanced (as determined by the WHO/FAO [62]) and that the nutritional/amino acid profile is comparable and even superior to other conventional sources of protein (i.e., egg and soybean protein) [63, 64]. The nutritional profile of microalgae can be shifted based on the following: (i) addition of specific nutrients to support growth, (ii) addition of enrichments to the algal production system prior to harvest, or (iii) exposure to nutritional stress, which can change the nutritional profile. For example, late logarithmically phase 2/3 microalgae typically contain 30–40% protein, 10–20% lipid, and 5–15% carbohydrate [65–67]. However, when cultured through stationary phase 4, this can change dramatically; for example, in a nitrate-limiting condition, the carbohydrate % can grow to double that of the protein present [68, 69]. Applications of microalgae as a feedstock and other coproducts are explored in the following subsections. Feedstock products include the use of microalgae as a food source for aquacultured fish and invertebrates and as a nutritious food and nutraceutical resource for humans. Microalgae can also be used to generate valuable coproducts such as bioplastics and biopolymers. These topics are addressed in the following subsections. 5.3.1

Microalgae as Animal Feed

Large-scale production of microalgae is of great importance to the commercial culture of specific life stages of bivalves (larvae and adults), crustaceans (larvae), zooplankton (which are the primary food for larval fishes and crustaceans), and, to a lesser degree, finfish (some larvae and adults) [70]. As primary producers, microalgae form the base of the trophic pyramid and are a critical link in the food chain of many aquatic animal species. Marine microalgae can be farmed on land and in the ocean and provide a highly nutritious source of protein and biomolecules for aquatic and terrestrial livestock without impacting our limited freshwater resources. Many of the beneficial biomolecules, naturally provided by microalgae, are not produced by animals and need to be incorporated into animal and human diets [71]. The genera of algae most commonly cultured as feeds for aquaculture species are Chlorella, Chaetoceros, Isochrysis, Nannochloropsis, Dunaliella, and Tetraselmis [14, 72, 73]. In addition, the marine diatom C. cryptica has been investigated as a potential heterotrophically grown aquaculture feedstock [74, 75]. There has been much interest in incorporating microalgae as a substitute for fishmeal and soybean protein in animal feed (poultry, cattle, and fish feeds) [63, 76–78]. For example, Spirulina has been used as a protein replacement or feed

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supplement in shrimp, silver sea bream, abalone, and poultry diets [79–81]; in shrimp farming, it has been shown to provide a significant cost/performance ratio. In 1910, Allen and Nelson achieved the mass production of microalgae (Chlorella) as a feed source for invertebrates [82]; however, Bruce [83] reported the methods to isolate and maintain single-species microalgae cultures of Isochrysis galbana and Pyramimonas grossii to feed oyster larvae. Dr Fujinaga in Japan pioneered two very different techniques to produce microalgae as feed for marine shrimp [84]. In the first technique, Fujinaga isolated the desired microalgae feed species from natural seawater and then expanded production of that species with the addition of nutrients, light, and oxygen to produce a dense single-species or axenic culture. In 1946, Fujinaga developed a second technique, termed bloom induction, where he added inorganic fertilizer to seawater in a pond, which resulted in a multispecies algal bloom that supported populations of zooplankton as feed for later stages of shrimp farming [85]. 5.3.2

Microalgae as a Human Food Source

Many types of microalgae have a high content of nutritious protein and are a valuable source of vitamins (especially B1 , B2 , C, and nicotinic acid) and β-carotene. Production costs for microalgae are still too high to compete with conventional protein sources [63]. Efforts are underway to develop methods to incorporate dried microalgae into known food items (bread, pasta, etc.) and to reduce the production cost [86]. Some of the roadblocks to increased consumption of microalgae as a human food source include the powder-like consistency, dark green color, and slightly fishy smell. Additionally, the Protein Advisory Group of the United Nations recommends limiting daily consumption of microalgae to 20 g of algae per day or 0.3 g of algae per kilogram of body weight because of potential toxic reactions to the nucleic acids (purines) in some species of microalgae, which cannot be broken down by the human metabolism [64]. At the same time, other species of microalgae such as the filamentous blue-green microalgae Spirulina have satisfied the food and nutritional supplement needs of differing cultures and regions for centuries. Spirulina was traditionally harvested from the lakes in Mexico and Africa for consumption by native Indians in Mexico and for incorporation into traditional foods in Africa [87]. Spirulina has also become very popular in the human health food industry in the United States and in Asia. About 15 different microalgal species in the genera Spirulina and Arthrospira are referred to as Spirulina [81]. The most commonly cultured species is Arthrospira platensis, which grows naturally in highly alkaline waters of lakes in warm regions. As a food, Spirulina has high levels of protein (55–70% dry weight), and the composition of biomolecules is well balanced (including all essential amino acids), and has high concentrations of PUFAs (i.e., EPA, DHA, ARA, LA). It also contains vitamins (B1 , B2 , B3 , B6 , B9 , B12 , C, D, and E), is a prolific source of potassium [81, 88], and has antioxidant, anti-inflammatory, and

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antibacterial properties [89]. The levels of β-carotene are very high in Spirulina, so it is commonly used as a supplement to reduce the risk of cancer. Spirulina also has a unique property, not yet found in other species of microalgae, where it has been shown to detoxify or chelate toxic minerals, such as arsenic [90, 91]. Batch or continuous-mode commercial production of Spirulina traditionally occurs in large open ponds or smaller raceway ponds in highly alkaline waters (pH > 9.2) in order to prevent the growth of predators [92, 93]. In Hawaii and California, intensive, high-density culture of Spirulina for the health food market is being done in shallow concrete or lined earthen D-ended raceways or in round ponds with paddlewheels that move and mix the algae in the system. Microalgae powder is currently being incorporated into some health foods and into vitamin capsules. The most common products on the market are Chlorella and Spirulina. However, new studies have shown that Nannochloropsis, Dunaliella, and Scenedesmus contain high levels of protein and are rich in the desired fatty acids (i.e., ARA and EPA) and antioxidants. These microalgae species, especially Nannochloropsis, have great potential for use in human health as a nutritional supplement [94]. 5.3.3

Microalgae in Nutraceuticals

The following sections discuss several of the high-value nutraceuticals being produced from microalgae, namely polyunsaturated fatty acids (PUFAs) and antioxidants. Besides nutraceuticals, microalgae represent a valuable source of almost all essential vitamins (e.g., A, B1 , B2 , B6 , B12 , C, E, nicotinate, biotin, folic acid, and pantothenic acid) [95, 96]). 5.3.3.1

Polyunsaturated Fatty Acids (PUFAs) in Microalgae

Merz and Main [10] discussed that a significant, but declining, proportion of world capture fisheries for nonfood production is processed into fishmeal (mainly for high-protein feed) and fish oil (as a feed additive in aquaculture and also for human nutritional supplements [97, 98]. While fish oil has become famous for its “essential fatty acids” such as eicosapentaenoic acid (EPA; C20:5n-3) and docosahexaenoic acid (DHA; C22:6n-3) ω-3 LC-PUFA content, fish do not actually produce ω-3 fatty acids, rather they accumulate their LC-PUFA ω-3 reserves by directly consuming microalgae or consuming forage fish that have accumulated ω-3 fatty acids via direct consumption of microalgae [10]. These ω-3 fatty acids can be obtained in the human diet directly from consuming the fish or the microalgae that produce them. Because the demand for purified EPA and DHA is exceeding the availability of the current common source of fish oil, alternative sources such as microalgae are being examined. Microalgae tend to store their energy source in lipid form as the culture ages with the PUFA content increasing until the culture approaches the late growth or early stationary phase of

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growth and then decreases gradually at the late stationary and death phases [99]. Microalgae mainly accumulate the following fatty acids: (C): 14:0, 14:1, 16:0, 16:1, 18:0, 18:1, 18:2, 18:3, 20:4, 20:5 (EPA), 22:5, 22:6 (DHA) [100–102]. DHA is the most abundant fatty acid found in the human brain. It is also essential for proper infant visual and neurological development and is an important component of human breast milk [103]. Because of the other components contained within fish oil, the DHA recovery and purification process can be both involved and expensive. Many species have been examined for DHA production; however, the heterotrophic dinoflagellate Crypthecodinium cohnii is most often selected for nutraceutical DHA production because of its unique ability to produce relatively high lipid accumulations (>20%), primarily as DHA (30–50%), with no other single PUFA present above 1%, which reduces follow-on recovery and purification costs [104, 105]. Industrial C. cohnii commercial fermentations are usually two-stage carbon-fed batch processes, with the first being an active growth phase under nutrient excess conditions, followed by a lipid accumulation phase under nutrient deficient conditions [106, 107]. EPA plays an important role in the regulation of biological functions and prevention and treatment of a number of human diseases such as heart and inflammatory diseases. EPA has been found in a wide variety of marine algae; however, only a few microalgal species have demonstrated industrial nutraceutical EPA production potentials [108]. P. tricornutum is a high-EPA-producing algal species with EPA comprising 30–40% of its total fatty acids when grown under optimum culture conditions [109]. Monodus species are photoautotrophic algae that can produce high levels of EPA, but the dependence on light results in low cell densities, making them unfavorable species for use in the industrial production of EPA [88]. Researchers identified heterotrophic strains of the marine diatom Nitzschia (especially N. alba and N. laevis) as a good producer of EPA where the oil content can be as high as 50% of the dry weight with EPA comprising 4–5% of the oil [110]. N. laevis can be grown heterotrophically using organic C sources such as glucose or glutamate [111] with the cellular EPA content higher than that in photoautotrophic growth [112–114]. Wen and Chen [30] discussed the development of a perfusion culture manufacturing process modified to permit cell bleeding during operation, allowing for continuous cell harvesting while removing inhibitory compounds during N. laevis cultivation. 5.3.3.2

Antioxidants in Microalgae

Chlorophyll, phycobiliproteins, and carotenoids are among the most significant light incidence pigments found in microalgae. Of these, the antioxidant functionality of carotenoids to interact and neutralize free radicals plays a significant role in the overall human health through the reduction of related cell damage. Astaxanthin obtained from the freshwater green flagellate microalgae Haematococcus pluvialis is the most potent commercially produced antioxidant available. 𝛽-carotene

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obtained from the halophilic, unicellular green algae Dunaliella salina also has the capabilities of being a powerful antioxidant as well as a vitamin A precursor [96]. Additionally, fucoxanthin, the major carotenoid of diatoms, is being investigated for its possible antioxidant and anti-inflammatory capabilities [115]. H. pluvialis is a freshwater microalgal species of Chlorophyta from the family of Haematococcaceae. H. pluvialis can be grown photoautotrophically, as well as mixotrophically or heterotrophically, using organic C sources. Commercial processes generally begin in closed PBRs followed by a carotenoid-accumulation step in the resting cells under reduced N and P conditions in open raceway ponds. Typical growth cycles can take several weeks [116–118] with Astaxanthin accumulations up to 6% of dry weight [119] when grown under optimum solar irradiance and less than that when grown mixotrophically or heterotrophically. D. salina is a marine photoautotroph capable of utilizing alternate C sources such as glycerol and acetate [120, 121]. The average concentration of carotenoids in most algae ranges between 0.1% and 0.2%, but Dunaliella will produce up to 14% of its dry weight in 𝛽-carotene when grown optimally under warm temperature conditions with maximum solar irradiance and hypersaline water [119]. Batch or continuous-mode commercial production of D. salina traditionally occurs in extremely large open ponds or smaller raceway ponds with salinity values greater than 30% NaCl (i.e., approximately 10 times the seawater salinity) in order to prevent the growth of predators [122] and contaminating species [92]. Closed-system PBR cultivation has been investigated [123]. Other algae strains used for 𝛽-carotene production include Spirulina platensis and Chlorella [88]. 5.3.4

Biopolymers from Microalgae

Existing uses and potential applications for microalgal bioplastics and biopolymers run the gamut from biosensors, biomedical and surgical dressings, agriculture, industrial membrane solvent/solute transport processes, water treatment, food, seed treatment, antimicrobials, and formation of biodegradable films [124, 125]. Interest in microalgal based polymers has evolved along with microalgae-based biofuels as alternative revenue sources to offset bottom-line production costs. For example, after oil extraction, the remaining biomass of some microalgal species can contain 30% cellulose, which can be used as a cellulose-based bioplastic feedstock. The genera of microalgae that have been used in cellulose biopolymer investigations include Phormidium, Chlorella, Chlamydomonas, Scenedesmus, Stichococcus, Anabaena, and Porphyridium [126]. At present, there is limited use of these on a large industrial scale, with commercialization success focused on smaller niche markets, such as the use of chitin and chitosan in medicine [127]. These biopolymers are also produced by some microalgae in addition to their more well-known production by crustaceans, which are discussed elsewhere in this book. The following sections of this chapter focus on chitin and chitin-derived derivatives such as chitosan, chitin

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oligosaccharides (COS), and glucosamine in the context of sustainable and interdisciplinary Water–Food–Energy Nexus challenges and applications. 5.3.4.1

Chitin and Chitosan: Abundant Biomaterial with High Applicability

Chitin and chitosan biopolymers possess a unique structure, multidimensional properties, and highly sophisticated functions. Chitin is the second most abundant polymer on Earth, after cellulose, and the most abundant natural polymer in the ocean, serving as an enormous reservoir of organic carbon and nitrogen [128]. As a natural structural biopolymer, chitin has a role analogous to that of collagen in the higher animals and cellulose in terrestrial plants [125, 129–132]. Chitin differs from cellulose in that within the glucose unit, one hydroxyl group (—OH) is replaced at the C-2 position with an acetamide group, allowing for increased hydrogen bonding between adjacent polymers and increasing the strength of the chitin–polymer matrix. Few biological polymers have such a high content of primary amines, and these amines confer important functional properties, which ultimately can be exploited for biomaterial fabrication, among other uses. Chitin and its derivatives have been used as natural flocculants and coagulants in microalgae-based systems as part of their solid/liquid separation and biomass dewatering for biofuel production, wastewater treatment, and nutrient recovery [133]. However, the continuing trend is toward producing high-value products for cosmetics and biotechnological uses in the medical field because of chitin’s inherent biodegradable, biocompatible, and nontoxic nature [134, 135]. Chitin contains acetylated units of N-acetyl-d-glucosamine (GlcNAc) (see Figure 5.1) and was first described in 1811 by Braconnot [136] as part of his research on mushrooms. Chitin is insoluble in water and common organic solvents, and because of its insolubility, chitin is usually converted to chitosan [137]. Chitosan, discovered in 1859 by Rouget [138], is obtained by the partial deacetylation (i.e., acetyl group removal from a molecule) of chitin in concentrated

Figure 5.1

Schematic of α-chitin structure.

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sodium hydroxide solution, which results in variable amounts of acetylated units of N-acetyl-d-glucosamine (GlcNAc) and deacetylated units of d-glucosamine (GlcN). It is important to note that the term “chitosan” does not refer to a single well-defined structure, and the functional properties of chitosan (i.e., such as thickening, film formation, and antimicrobial activity) are related to its molecular weight, degree of N-acetylation (DA), and sequence (i.e., whether the acetylated residues are distributed along the backbone in a random or blocky manner). Commercial chemical conversion of chitin to chitosan is typically accomplished by N-deacetylation (removing an acetate moiety from chitin) through harsh acid and base treatments to demineralize and deproteinize or through enzymatic hydrolysis in the presence of chitin deacetylase [139–141]. 5.3.4.2

Uses of Chitin Polymorphic Forms and 𝛃-Chitin

Chitin exists in three different crystalline structure/polymorphic forms, referred to as α-, β-, and γ-chitin [142–146]. α-Chitin is the most common polymorphic form found in commercial chitins, usually isolated from large amounts of marine crustacean (e.g., crab and shrimp shell) biowaste available as a by-product of food processing. β-Chitin is both rare and unique, being found naturally in Loligo squid pens, the nanofibrils (fibers) of some microalgae (diatoms), the tubes of pogonophores (giant undersea tube worms), and vestimentiferans. Among the few living organisms that can serve as a source of β-chitin, only diatom microalgae have the potential of being able to be cultivated in large and scalable quantities for industrial bioproduction. Diatoms are found throughout the world’s oceans and freshwater systems; however, only Thalassiosira and Cyclotella are known to produce β-chitin fibrils. Euryhaline, nontoxic, centric diatoms T. fluviatilis (phototrophic) and C. cryptica (heterotrophic) produce β-chitin nanofibrils [115, 129, 147]. These nanofibrils (fibers), also referred to in the literature as mucilage or slime threads, originate from the marginal and central pores in the silica frustule and serve to reduce the sinking velocity via an increased form resistance. Walsby and Xypolita [148] reported a doubling of the sinking rate of T. fluviatilis and C. cryptica cells after chitinase digestion of the chitin fibers, demonstrating their importance as a flotation mechanism [149]). In addition, it was reported that both diatom species produce chitin fibers under both CO2 - and NO3 -limiting conditions, suggesting a considerable amount of effort is expended in generating these pure β-chitin fibers in terms of energy and resources [147, 149]. Nanofibril analysis revealed them to be naturally free of protein and entirely composed of pure β-(1→4)-linked N-acetyl glucosamine [147, 149–153] with β-chitin comprising 31–38% of the cellular material (including the silica) of this diatom [150]. The unit-cell structural arrangement of the β-chitin crystal is shown in Figure 5.2. The structure is seen as a series of hydrogen-bonded sheets, parallel in orientation with successive chains of the same sense held together by weak

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Figure 5.2 β-Chitin schematic of structure and hydrogen bonding (dashed lines).

intrasheet intermolecular hydrogen bonding of the amide groups [154]. This contrasts that of α-chitin, whose unit-cell form consists of two antiparallel polymer chains held together by strong intra- and intersheet hydrogen bonds. Because of the weaker intrasheet hydrogen bonding, β-chitin can accept intercalated water molecules within its lattice and yields crystallosolvates [155–157], and reversible hydration–dehydration and stable hydrate formation are facilitated. The swelling of β-chitin in water is strongly anisotropic without modifications of the β-chitin sheets that are maintained by the strong N—H· · ·O C intermolecular hydrogen bonds [156] and can lead to unique transport properties in chitosan membranes produced from them. In addition, β-chitin is more amenable to N-deacetylation compared to α-chitin, which in turn should facilitate conversion to chitosan and potentially lead to a new, easier, and “greener” way to develop chitin chemistry and fabrication of β-chitin-based biopolymers. The anisotropic swelling of β-chitin in water can have significant impact on water and/or ion transport through membranes, which is a key factor in many industrial processes and applications. Controllable ion transport, water and water vapor transmission combined with chitosan’s inherent antimicrobial/antifouling properties present potential positive opportunities in many membrane-based applications, especially with respect to industrial membrane-based systems such as electroacidification, water desalinization, water treatment/purification, and emerging technologies such as salinity gradient power (SGP) energy generation [2, 158], and FO. Additional technological development is needed to produce and evaluate the suitability of the Thalassiosira and Cyclotella microalgal-produced β-chitin and chitosan membranes for these applications. 5.3.4.3

Chitosan and Its Derivatives: Oligosaccharides (COS) and Glucosamine

Chitosan is produced by the deacetylation of chitin, broken down by either acid hydrolysis or enzymatic hydrolysis into COS (also known as chitosan oligosaccharides) and the monomeric sugar glucosamine. These hydrolyzed products of chitosan are readily soluble in water because of their shorter chain

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lengths, low molecular weight, and free amino acid groups found in deacetylated units of d-glucosamine (GlcN) [159]. Chitosan is a versatile, natural, biopolymer of generally poor solubility but possesses desirable physical properties of low toxicity along with being both highly biocompatible and biodegradable [160, 161]. This versatility has generated much interest since chitosan-degraded products have potential applications in biomedicine, agriculture, nutrition, and biotechnology [162]. Because of the free amino groups, chitosan has a high affinity to absorb metal cations leading to its interest as a natural chelating biopolymer [163, 164] and as a candidate for absorbents to remove toxic heavy metals in industrial wastewater [165]. Water-soluble bioactive COS and glucosamine research activities have focused on a wide variety of health, food, cosmeceutical, and nutritional applications. Chemical hydrolysis techniques are not typically considered for use as bioactive materials because of the potentially toxic nature of the products used in the process. However, COSs prepared using chitosanases (chitosan-hydrolyzing enzymes that hydrolyze chitosan but not chitin [166]) are finding greater use and acceptance [167]. One promising application of chitosanases is the conversion of fishery/marine crustacean chitinous biowaste/by-products into COS [168]. Liang et al. [169] reported successful β-chitin (squid pen)-based COS development of a food colorant absorber using chitosanase (Bacillus cereus TKU033). In addition, glucosamine has been marketed as a natural nutraceutical product for the treatment of osteoarthritis and osteoarthritic pain. Additional technological development is needed to produce and evaluate the suitability of the Thalassiosira and Cyclotella microalgal-produced β-chitin for these applications.

5.4

Conclusion – The Path Forward

The demand for suitable biomass for biofuels, aquafeed/feeds for food production, and for nutraceutical production alongside biopolymeric-based industrial technologies, such as desalination and SGP, has fueled continued interest in microalgae bioproduction. Microalgae are a good candidate because of their variations in mixotrophic growth requirements, composition, higher biomass production, faster growth, and scalability. Besides the phototrophic capabilities, the potential exists for heterotrophic (without sunlight) growth of some microalgal species for additional opportunity for biomass production using alternative and sometimes waste stream carbon sources, such as glycerin (a biodiesel by-product of the transesterification process) or CO2 flue gas as their sole carbon and energy source. Continued research into optimizing overall biomass growth and emerging bioproduction techniques, such as perfusion cell-culture manufacturing methodologies and offshore cage/membrane enclosures, can positively affect ancillary coproducts, with some of the resulting metabolites such as chitosan-derived COS, lipid-derived ω-3 nutraceuticals and feeds for animal/marine/human feedstock production, reducing the overall total cost of the microalga biomass production operation.

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Acknowledgments The authors would like to thank several anonymous reviewers for their comments, which improved the quality of this manuscript.

References 1. Sheehan, J., Dunahay, T., Benemann, J., et al. (1998) A look back at the U.S. Department of Energy’s Aquatic Species Program: Biodiesel from Algae. NREL/TP-580-24190. 2. Merz, C., Moreno, W., Barger, M. and Stephen, M. (2012) Salinity gradient power (SGP): a developmental roadmap covering existing generation technologies and recent investigative results into the feasibility of bipolar membrane-based salinity gradient power generation. Innovation and Technology – Proceedings of the National Academy of Inventors, 14 (3/4), 249–275. doi: 10.3727/ 194982412X13500042168857 3. Merz, C. (2013) Consideration of a variable frequency energy conversion system for marine and onshore wind energy extraction. Marine Technology Society Journal, 47 (4), 218–225. doi: 10.4031/MTSJ.47.4.11 4. Weisberg, R., Liu, Y., Merz, C. et al. (2012) A critique of alternative power generation for Florida by mechanical and solar means. Marine Technology Society Journal, 46 (5), 12–23. doi: 10.4031/MTSJ.46.5.1 5. Scott, S., Davey, M., Dennis, J. et al. (2010) Biodiesel from algae: Challenges and prospects. Current Opinion in Biotechnology, 21, 277–286. 6. Chisti, Y. (2010) Fuels from microalgae. Biofuels, 1, 233–235. 7. Chisti, Y. and Yan, J. (2011) Algal biofuels – a status report. Applied Energy, 88, 3277–3279. 8. Chisti, Y. (2012) Raceways-based production of algal crude oil, in Microalga Biotechnology: Potential and Production (eds C. Posten and C. Walter), de Gruyter, Berlin, pp. 113–146. 9. Ferrell, J., Sarisky-Reed, V. (2010) National Algal Biofuels Technology Roadmap. U.S. Department of Energy, Office of Energy Efficiency and Renewable Energy Biomass Program. http://biomass.energy.gov. 10. Merz, C., Main, K. (2014) Microalgae (diatom) production – The aquaculture and biofuel nexus. Oceans’14 MTS/IEEE Conference Proceedings – IEEE Xplore, St. John’s, Newfoundland, Canada. DOI:10.1109/OCEANS.2014.7003242. 11. Valero, S., Tresguerras, T. and Abuin, A. (1981) Large-scale outdoor algal production for rearing seed oysters and clams to juvenile stage, in Nursery Culturing of Bivalve Mollusks. Proceedings of the International Workshop on Nursery Culturing of Bivalve Mollusks (eds C. Claus, D. Pauw and D. Jaspers), Spec Publ. Eur. Mariculture Society, Ghent, Belgium, pp. 117–139. 12. Ukeles, R. (1976) Cultivation of plants: unicellular plants, in Marine Ecology. III (Part 1) (ed. O. Kinne), John Wiley and Sons, New York, pp. 367–466. 13. Fabregas, J., Toribio, L., Abalde, J. et al. (1987) Approach to biomass production of the marine microalga Tetraselmis suecica (Kylin) using common garden fertilizer and soil extract as cheap nutrient supply in batch cultures. Aquacultural Engineering, 6, 141–150. 14. Guillard, R. (1975) Culture of phytoplankton for feeding marine invertebrates, in Culture of Marine Invertebrate Animals (eds W. Smith and H. Chanley), Plenum Press, New York, pp. 29–60.

104

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

15. Gonzalez-Rodriguez, E. and Maestrini, S. (1984) The use of some agricultural fertilizers for the mass production of marine algae. Aquaculture, 36, 245–256. 16. Thurman, H.V. (1997) Introductory Oceanography, Prentice Hall College, New Jersey, USA. ISBN 0-13-262072-3. 17. Norton, T., Melkonian, M. and Anderson, R. (1996) Algal biodiversity. Phycologia, 35, 308–326. 18. Trequer, P., Nelson, D.M., Van Bennekom, A.J. et al. (1995) The silica balance in the world ocean: a reestimate. Science, 268, 375–379. 19. Field, C., Behrenfeld, M., Randerson, J. and Falkowski, P. (1998) Primary production of the biosphere, integrating terrestrial and oceanic components. Science, 281, 237–240. 20. Hildebrand, M., Davis, A., Smith, S. et al. (2012) The Place of Diatoms in the Biofuels Industry. Biofuels, 3 (2), 221–240. doi: 10.4155/BFS.11.157 21. Morineau-Thomas, O., Jaouen, P. and Legentilhomme, P. (2002) the role of exopolysaccharides in fouling phenomenon during ultrafiltration of microalgae (Chlorella sp. and Porphyridium purpureum): advantages of a swirling decay flow. Bioprocess and Biosystems Engineering, 25, 35–42. 22. Cardozo, K., Thais, G., Marcelo, P. et al. (2006) Metabolites from algae with economic impact. Comparative Biochemistry and Physiology Part C: Toxicology & Pharmacology (Elsevier Inc.), 146 (1-2), 60–78. doi: 10.1016/j.cbpc.2006.05.00 23. Liang, Y., Sarkany, N. and Cui, Y. (2009) Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic, and mixotrophic growth conditions. Biotechnology Letters, 31, 1043–1049. DOI: 10.1007/s10529-009-9975-7. 24. Yang, J., Xu, M., Zhang, X. et al. (2011) Life-cycle analysis on biodiesel production from microalgae: water footprint and nutrients balance. Bioresource Technology, 102, 159–165. 25. Brzezinski, M. (1985) The Si:C:N ratio of marine diatoms: interspecific variability and the effort of some environmental variables. Journal of Phycology, 21, 347–357. 26. Shilton, A., Powell, N., Mara, D. and Craggs, R. (2008) Solar-powered aeration and disinfection, anaerobic co-digestion, biological CO2 scrubbing and biofuel production: the energy and carbon management opportunities of waste stabilization ponds. Water Science and Technology, 58, 253–258. 27. Coutteau, P. (1996) Micro-algae, in Manual on the Production and Use of Live Food for Aquaculture (eds P. Lavens and P. Sorgeloos) FAO Fisheries Technical Paper. No. 361, Chapter 2, FAO, Rome, pp. 7–48. 28. Barsanti, L. and Gualtieri, P. (2010) Algae: Anatomy, Biochemistry, and Biotechnology, 2nd edn, CRC Press Taylor & Francis Group, pp. 252–257. 29. Lim, H., Shin, H. (2013) Fed-Batch Cultures: Principles and Applications of Semi-Batch Reactors. Cambridge University Press. ISBN: 9780521513364. 30. Wen, Z. and Chen, F. (2002) Perfusion culture of the diatom Nitzschia laevis for ultra-high yield of eicosapentaenoic acid. Process Biochemistry, 38, 523–529. 31. Crowley, J., Wubben, M., Coco Martin, J. (2005) Process for cell culturing by continuous perfusion and alternating tangential flow. US Patent 2008131934, filing date Mar 4, 2005 and issued June 5, 2008. 32. Shevitz, J. (2000) Fluid filtration system. US Patent 6544424, filing date Mar 17, 2000 and issued April 8, 2003. 33. Pollock, J., Ho, S. and Farid, S. (2013) Fed-batch and perfusion culture processes: economic, environmental, and operational feasibility under uncertainty. Biotechnology and Bioengineering, 110 (1), 206–219.

Microalgae Bioproduction – Feeds, Foods, Nutraceuticals, and Polymers

105

34. Drexler, I. and Yeh, D. (2014) Membrane applications for microalgae cultivation and harvesting: a review. Reviews in Environmental Science and Biotechnology. DOI:10.1007/s11157-014-9350-6. 35. Chisti, Y. (2007) Biodiesel from microalgae. Biotechnology Advances, 25 (3), 294–306. 36. Hoffman, Y., Aflalo, C., Zarka, A. et al. (2008) Isolation and characterization of a novel chytrid species (phylum Blastocladiomycota), parasitic on the green alga Haematococcus. Mycological Research, 112, 70–81. 37. Hirayama, K. and Ogawa, S. (1972) Fundamental studies on the physiology of the rotifer for its mass culture. I. Filter feeding of rotifer. Bulletin of the Japanese Society of Scientific Fisheries, 38, 1207–1214. 38. Hwang, T., Park, S., Oh, Y. et al. (2013) Harvesting of Chlorella sp. KR-1 using a cross-flow membrane filtration system equipped with an anti-fouling membrane. Bioresource Technology, 139, 379–382. 39. Hanson, R.L. (1991) Evapotranspiration and droughts, in National Water Summary 1988–1989 – Hydrologic Events and Floods and Droughts, vol. 2375 (eds R. Paulson, E. Chase, R. Roberts and D. Moody), U.S. Geological Survey Water Supply Paper, pp. 99–104. 40. Molina, G., Acién Fernández, F., García, C. and Chisti, Y. (1999) Photobioreactors: light regime, mass transfer, and scale up. Journal of Biotechnology, 70, 231–247. 41. Tredici, M. (1999) Bioreactors, photo, in Encyclopedia of Bioprocess Technology: Fermentation, Biocatalysis and Bioseparation (eds M. Flickinger and S. Drew), Wiley, pp. 395–419. 42. Carvalho, A., Meireles, L. and Malcata, F. (2006) Mi**croalgal reactors: a review of enclosed system designs and performances. Biotechnology Progress, 22, 1490–1506. 43. Neori, A., Chopin, T., Troell, M. et al. (2004) Integrated aquaculture: rationale, evolution and state of the art emphasizing seaweed biofiltration in modern Mariculture. Aquaculture, 231, 361–391. 44. Borges, M., Silva, P., Moreira, L. and Soares, R. (2005) Integration of consumer-targeted microalgal production with marine fish effluent biofiltration – a strategy for mariculture sustainability. Journal of Applied Phycology, 17, 187–197. 45. Metchnikoff, E., Roux, E. and Salimbeni, T. (1896) Toxine et antitoxine cholerique. Annales de l’Institut Pasteur, T, 10, 257–282. 46. Schultz, J. and Gerhardt, P. (1969) Dialysis culture of microorganisms: design, theory, and results. Bacteriological Reviews, 32, 1–47. 47. Marsot, P., Cembella, A. and Houle, L. (1991) Growth kinetics and nitrogen-nutrition of the marine diatom Phaeodactylum tricornutum in continuous dialysis culture. Journal of Applied Phycology, 3, 1–10. 48. Trainor, F. (1965) A study of unialgal cultures of Scenedesmus incubated in nature and in the laboratory. Canadian Journal of Botany, 43, 701–706. 49. Jensen, A., Rystad, B. and Skoglund, L. (1972) The use of dialysis culture in phytoplankton studies. Journal of Experimental Marine Biology and Ecology, 8, 241–248. 50. Jensen, A. and Rystad, B. (1973) Semi-continuous monitoring of the capacity of sea water for supporting growth of phytoplankton. Journal of Experimental Marine Biology and Ecology, 11, 275–285. 51. Dor, I. (1975) High density dialysis culture of algae on sewage. Water Research, 9, 251–254.

106

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

52. Cooper, S., Battat, A., Marsot, P. and Sylvestre, M. (1983) Production of antibacterial activities by two Bacillariophyceae grown in dialysis culture. Canadian Journal of Microbiology, 29, 338–341. 53. Marsot, P., Cembella, A. and Houle, L. (1991) Growth kinetics and nitrogen-nutrition of the marine diatom Phaeodactylum tricornutum in continuous dialysis culture. Journal of Applied Phycology, 3, 1–10. 54. Blais, C., Fournier, R. and Marsot, P. (1984) Continuous microalgal culture using swine manure dialysate as a nutrient source. Aquacultural Engineering, 3, 275–287. 55. Xu, M., Bernards, M. and Hu, Z. (2014) Algae-facilitated chemical phosphorous removal during high-density Chlorella emersonii cultivation in a membrane bioreactor. Bioresource Technology, 153, 383–387. 56. Wiley, P., Harris, L., Reinsch, S. et al. (2013) Microalgae cultivation using offshore membrane enclosures for growing algae (OMEGA). Journal of Sustainable Bioenergy Systems, 3, 18–32. 57. Buckwalter, P., Embaye, T., Gormly, S. and Trent, J. (2013) Dewatering microalgae by forward osmosis. Desalination, 312, 19–22. 58. Merz, C. (2001) An overview of the coastal ocean monitoring and prediction system (COMPS). Proceedings of MTS/IEEE OCEANS’011, MTS 0-933957-28-9, DOI:10.1109/OCEANS.2001.968281. 59. Harris, L., Tozzi, S., Wiley, P. et al. (2013) Potential impact of biofouling on the photobioreactors of the offshore membrane enclosures for growing algae (OMEGA) system. Bioresource Technology, 144, 420–428. 60. Carvalho, A., Luis, A., Meireless, A. and Malcata, F. (2006) Microalga reactors: a review of enclosed system designs and performances. Biotechnology Progress, 22, 1490–1506. 61. TEDx Academy (2015). Jonathan D. Trent, OMEGA Global Initiative and NASA-AMES Research Center, OMEGA: An Energy, Water, Food Life-support System for Spaceship Earth [Cited March 17, 2016]. http://omegaglobal.org/upcoming-talk-at-the-carnegieinstitution-for-science/. 62. FAO/WHO (1973) Energy and protein requirement. Report of a Joint FAO/WHO ad hoc Expert Committee Vol. 52. Geneva. 63. Becker, E. (2007) Micro-algae as a source of protein. Biotechnology Advances, 25, 207–210. 64. Becker, E. (2004) Microalgae in human and animal nutrition, in Handbook of Microalgal Culture: Biotechnology and Applied Phycology (ed. A. Richmond), Blackwell Science, Oxford, pp. 312–351. 65. Brown, M., Jeffery, S., Volkman, J. and Dunstan, G. (1997) Nutritional properties of microalgae for mariculture. Aquaculture, 151, 315–331. 66. Renaud, S., Thinh, L. and Parry, D. (1999) The gross composition and fatty acid composition of 18 species of tropical Australian microalgae for possible use in mariculture. Aquaculture, 155, 207–221. 67. Brown, M. (2002) Nutritional value of microalgae for Aquaculture, in Advances en Nutricion Acuicola VI (eds L. Cruz-Suarez, D. Ricque-Marie, M. Tapia-Salazar et al.), Memorias del VI Simposium Internacional de Nutricion Acuicola. 3 al 6 de September, Cancun, Quintana Roo, Mexico. 68. Harrison, P., Thompson, P. and Calderwood, G. (1990) Effects of nutrient and light limitation on the biochemical composition of phytoplankton. Journal of Applied Phycology, 2, 45–56.

Microalgae Bioproduction – Feeds, Foods, Nutraceuticals, and Polymers

107

69. Brown, M., Garland, C., Jeffrey, S. et al. (1993) The gross and amino acid compositions of batch and semi-continuous cultures of Isochrysis sp. (clone of T. ISO), Pavlova lutheri and Nannochloropsis oculata. Journal of Applied Phycology, 5, 285–296. 70. Fulks, W. and Main, K. (1991) Rotifer and Microalgae Culture Systems – Proceedings of a U.S.–Asia Workshop. The Oceanic Institute, Honolulu, Hawaii. ISBN 0-9617016-2-5. 71. Yaakob, Z., Ali, E., Zainal, A. et al. (2014) An overview: biomolecules from microalgae for animal feed and aquaculture. Journal of Biological Research-Thessaloniki, 21, 6. doi: 10.1186/2241-5793-21-6 72. Persoone, G. and Claus, C. (1980) Mass culture of algae: a bottleneck in the nursery culturing of mollusks, in Algae Biomass (eds G. Shelef and C. Soeder), Elsevier/North-Holland Biomedical Press, New York, pp. 265–285. 73. Liao, I., Su, H. and Lin, J. (1983) Larval foods for penaeid prawns, in CRC Handbook of Mariculture: Crustacean Aquaculture (ed. J. McVey), CRC Press Inc., Boca Raton, Florida, pp. 43–69. 74. Pahl, S., Lewis, D., Chen, F. and King, K. (2010a) Heterotrophic growth and nutritional aspects of the diatom Cyclotella cryptica (Bacillariophyceae): effect of some environmental factors. Journal of Bioscience and Bioengineering, 109 (3), 235–239. 75. Pahl, S., Lewis, D., Chen, F. and King, K. (2010b) Growth dynamics and the proximate biochemical composition and fatty acid profile of the heterotrophically grown diatom Cyclotella cryptica. Journal of Applied Phycology, 22, 165–171. 76. Vonshak, A. (1997) Spirulina platensis (Arthrospira): Physiology, Cell Biology and Biotechnology. Basingstoke, Hants, London, UK, Taylor and Francis. 77. Qiao, H., Wang, H., Song, Z. et al. (2014) Effects of dietary fish oil replacement by microalgae raw materials on growth performance, body composition and fatty acid profile of juvenile olive flounder, Paralichthys olivaceus. Aquaculture Nutrition, 20 (6), 646–653. 78. Taelman, S., De Meester, S., Wim Van Dijk, W. et al. (2015) Environmental sustainability analysis of a protein-rich livestock feed ingredient, in the Netherlands: microalgae production versus soybean import. Resources, Conservation and Recycling, 101, 61–72. 79. El-Sayed, A. (1994) Evaluation of soybean meal, spirulina meal and chicken offal meal as protein sources for silver sea bream (Rhabdosargus sarba) fingerlings. Aquaculture, 127, 169–176. 80. Britz, P. (1996) The suitability of selected protein sources for inclusion in formulated diets for the South African abalone, Haliotis midae. Aquaculture, 140, 63–73. 81. Habib, M., Parvin, M., Huntington, T. and Hasan, M. (2008) A Review on Culture, Production and Use of Spirulina as Food for Humans and Feeds for Domestic Animals and Fish, Food and Agriculture Organization of the United Nations, Rome. 82. Ryther, J. and Goldman, J. (1975) Microbes as food in mariculture. Annual Reviews of Microbiology, 29, 429–433. 83. Bruce, J., Knight, M. and Parke, M. (1940) The rearing of oyster larvae on an algal diet. Journal of the Marine Biological Association U.K., 24, 337–374. 84. De Pauw, N. and Pruder, G. (1986) Use and production of microalgae as food in aquaculture: practices, problems and research needs, in Realism in Aquaculture: Achievements, Constraints, Perspectives (eds M. Bilio, H. Rosenthal and C. Sinderman), European Aquaculture Society, Bredene, pp. 77–106. 85. Yang, W.T. (1975) A manual for Large-Tank Culture of Penaeid Shrimp to the Postlarval Stages, University of Miami Sea Grant, Coral Gables, Florida. 94 pp.

108

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

86. Acién Fernández, F., Fernandez Sevilla, J. and Molina Grima, E. (2013) Photobioreactors for the production of microalgae. Reviews in Environmental Science and Biotechnology, 12, 131–151. 87. Vaz, B., Moreira, J., Morais, M. and Costa, J. (2016) Microalgae as a new source of bioactive compounds in food supplements. Current Opinion in Food Science, 7, 73–77. 88. Oilgae (2016). Omega – 3Fatty Scids [Cited March 25, 2016]. http://www.oilgae.com/non_ fuel_products/omega_3fattyacids.html. 89. Liu, J., Hou, C., Lee, S. et al. (2011) Antioxidant effects and UVB protective activity of Spirulina products fermented with lactic acid bacteria. Process Biochemistry, 46, 1405–1410. 90. Maeda, S. and Sakaguchi, T. (1990) Accumulation and detoxification of toxic metal elements by algae, in Introduction to Applied Phycology (ed. I. Akatsuka), SPB Academic Publishing, The Hague, The Netherlands, pp. 109–136. 91. Okamura, H. and Aoyama, I. (1994) Interactive toxic effects and distribution of heavy metals in phytoplankton. Environmental Toxicology and Water Quality, 9, 7–15. 92. Borowitzka, L. and Borowitzka, M. (1989) Industrial production: methods and economics, in Algal and Cyanobacterial Biotechnology (eds R. Cresswell, T. Rees and N. Shah), Longman Scientific, London, pp. 294–316. 93. Chen, F. (1996) High cell density culture of microalgae in heterotrophic growth. Trends in Biotechnology (TIBTECH), 14, 421–426. 94. Kent, M., Welladsen, H.M., Mangott, A. and Li, Y. (2015) Nutritional evaluation of Australian microalgae as potential human health supplements. PLoS ONE, 10 (2), e0118985. DOI:10.1371/journal.pone.0118985. 95. Richmond, A. (2004) Handbook of Microalgal Culture: Biotechnology and Applied Phycology, 1st edn, Wiley-Blackwell. ISBN: 978-0-632-05953-9. 96. Iersel, S., Flammini, A. (2010). Alga-based biofuels: applications and co-products. FAO Environmental and Natural Resource Service Series. No. 44. Rome, FAO. 117 pp. 97. FAO United Nations (2014) The State of the World Fisheries and Aquaculture: Opportunities and Challenges, FAO, Rome. 98. Ruiz-Lopez, N., Haslam, R., Napier, J. and Sayanova, O. (2014) Successful high-level accumulation of fish oil omega-3 long chain polyunsaturated fatty acids in a transgenic oilseed crop. The Plant Journal, 77, 198–208. DOI:10.1111/tpj.12378. 99. Yongmanitchai, W. and Ward, O. (1989) Omega-3 fatty-acids – alternative sources of production. Process Biochemistry, 24, 117–125. 100. Vazhappilly, R. and Chen, F. (1998) Eicosapentaenoic acid and docosahexaenoic acid production potential of microalgae and their heterotrophic growth. Journal of the American Oil Chemists’ Society, 75, 393–397. 101. de-Bashan, L., Bashan, Y., Moreno, M. et al. (2002) Increased pigment and lipid content, lipid variety, and cell and population size of the microalgae Chlorella spp. when co-immobilized in alginate beads with microalgae-growth-promoting bacterium Azospirillum brasilense. Canadian Journal of Microbiology, 48, 514–521. 102. Perez-Garcia, O., Escalante, F., de-Bashan, L. and Bashan, Y. (2011) Heterotrophic cultures of microalgae: metabolism and potential products. Water Research, 45, 11–36. 103. Nettleton, J. (1993) Are omega-3 fatty acids essential nutrients for fetal and infant development? Journal of the American Dietetic Association, 93, 58–64. 104. Henderson, R., Leftley, J. and Sargent, J. (1988) Lipid composition and biosynthesis in the marine dinoflagellate Crypthecodinium cohnii. Phytochemistry, 27, 1679–1683.

Microalgae Bioproduction – Feeds, Foods, Nutraceuticals, and Polymers

109

105. Swaaf, M., Rijk, T., Eggink, G. and Sijtsma, L. (1999) Optimisation of docosahexaenoic acid production in batch cultivations by Crypthecodinium cohnii. Journal of Biotechnology, 70, 185–192. 106. Wynn, J., Behrens, P., Sundararajan, A. et al. (2005) Production of single cell oils by dinoflagellates, in Single Cell Oils (eds Z. Cohn and C. Ratledge), AOCS Press, Illinois, pp. 86–98. 107. Mendes, A., Reis, A., Vasconcelos, R. et al. (2009) Crypthecodinium cohnii with emphasis on DHA production: a review. Journal of Applied Phycology, 21, 199–214. 108. Barclay, W., Meager, K. and Abril, J. (1994) Heterotrophic production of long chain omega-3 fatty acids utilizing algae and algae-like microorganisms. Journal of Applied Phycology, 6, 123–129. 109. Yongmanitchai, W. and Ward, O. (1992) Growth and eicosapentaenoic acid production by Phaeodactylum tricornutum in batch and continuous culture systems. Journal of the American Oil Chemists’ Society, 69, 584–590. 110. Kyle, D., Gladue, R. (1991) Eicosapentaenoic acids and methods for their production. US PCT Patent WO 1991014427 A1, filing date March 20, 1991 and issued October 3, 1991. 111. Lewin, J. and Hellebust, J. (1978) Utilization of glutamate and glucose for heterotrophic growth by marine pennate diatom Nitzschia laevis. Marine Biology, 47, 1–7. 112. Tan, C. and Johns, M. (1996) Screening of diatoms for heterotrophic eicosapentaenoic acid production. Journal of Applied Phycology, 8, 59–64. 113. Wen, Z. and Chen, F. (2000) Production potential of eicosapentaenoic acid by the diatom Nitzschia laevis. Biotechnology Letters, 22, 727–33. 114. Wen, Z. and Chen, F. (2003) Heterotrophic production of eicosapentaenoic acid by microalgae. Biotechnology Advances, 21, 273–294. 115. Gugi, B., Costaouec, T., Burel, C. et al. (2015) Diatom-specific oligosaccharide and polysaccharide structures help to unravel biosynthetic capabilities in diatoms. Marine Drugs, 13 (9), 5993–6018. DOI:10.3390/md13095993. 116. Boussiba, S., Vonshak, A., Cohen, Z., Richmond, A. (1997) Procedure for large-scale production of astaxanthin from haematococcus. US PCT Patent 9,728,274, filing date Jan 30, 1997 and issued August 7, 1997. 117. Olaizola, M. (2000) Commercial production of astaxanthin from Haematococcus pluvialis using 25,000 liter outdoor photobioreactors. Journal of Applied Phycology, 12, 499–506. 118. Cysewski, G. and Lorenz, R. (2004) Industrial production of microalgal cell-mass and secondary products – species of high potential: haematococcus, in Microalgal Culture: Biotechnology and Applied Phycology (ed. A. Richmond), Blackwell Science, Oxford, pp. 281–288. 119. Barowitzka, M. (2010) Carotenoid production using microorganisms, in Single Cell Oils: Microbial and Algal Oils, 2nd edn (eds Z. Cohn and C. Ratledge), AOCS Press, Urbana, Illinois, USA, pp. 225–240. 120. Suarez, G., Romero, T. and Borowitzka, M. (1998) Cultivo de la Microalga (Dunaliella salina) en Medio Organico. Anais do IV Congresso Latino-American, II Reuniao Iberio-Americana, VII Reuniao Brasileira de Ficologia, in Sociedada Ficologica de America Latina e Caribe & Sociatade Brasiliera de Ficologia (eds E. de Paula, M. Corediro-Marino, D. Santos et al.), Caxambu, pp. 371–382. 121. Mojaat, M., Pruvost, J., Foucault, A. and Legrand, J. (2008) Effect of organic carbon sources and Fe2+ ions on growth and β-carotene accumulation by Dunaliella salina. Biochemical Engineering Journal, 39, 177–184.

110

Fuels, Chemicals and Materials from the Oceans and Aquatic Sources

122. Post, F., Borowitzka, L., Borowitzka, M. et al. (1983) The protozoa of a Western Australian hypersaline lagoon. Hydrobiologia, 105, 95–113. 123. Garcia-Gonzalez, M., Moreno, J., Manzano, J. et al. (2005) Production of Dunaliella salina biomass rich in 9-cis-β-carotene and lutein in a closed tubular photobioreactor. Journal of Biotechnology, 115, 81–90. 124. Shahidi, F., Arachchi, J. and Jeon, Y. (1999) Food applications of chitin and chitosans. Trends in Food Science and Technology, 10, 37–51. 125. Pillai, C., Paul, W. and Sharma, C. (2009) Chitin and chitosan polymers: chemistry, solubility and fiber formation. Progress in Polymer Science, 34, 641–678. 126. Oilgae (2016b) Biopolymers and Bioplastics [Cited March 18, 2016]. http://www.oilgae .com/non_fuel_products/biopolymers.html. 127. Kerton, F., Liu, Y., Murphy, J., Hawboldt, K. (2014) Renewable resources from the oceans. Oceans’14 MTS/IEEE Conference proceedings – IEEE Xplore, St. John’s, Newfoundland, Canada. DOI:10.1109/OCEANS.2014.7002983. 128. Durkin, C., Mock, T. and Armbrust, V. (2009) Chitin in diatoms and its association with the cell wall. Eukaryotic Cell, 8 (7), 1038–1050. 129. Muzzarelli, R. (1977) Chitin, Pergamon Press, Oxford. 130. Muzzarelli, R., Jeuniaux, C. and Gooday, G. (1986) Chitin in Nature and Technology, Plenum Publishing Corporation, New York, NY, USA. 131. Roberts, G. (1992) Chitin Chemistry, Macmillan Press, London. 132. Mayer, G. and Sarikaya, M. (2002) Rigid biological composite materials: structural examples for biomimetic design. Experimental Mechanics, 42, 395–403. 133. Billuri, M., Bonner, J., Fuller, C. and Islam, M. (2015) Impact of natural cationic polymers on charge and clarification of microalgae suspensions. Environmental Engineering Science, 32 (3), 212–221. DOI:10.1089/ees.2014.0301. 134. Jang, M., Kong, B., Jeong, Y. et al. (2004) Physicochemical characterization of α-chitin, β-chitin and 𝛾-chitin separated from natural resources. Journal of Polymer Science: Part A: Polymer Chemistry, 42, 3423–3432. 135. Jumaa, M., Furkert, F. and Muller, B. (2002) A new lipid emulsion formulation with high antimicrobial efficacy using chitosan. European Journal of Pharmaceuticals and Biopharmaceutics, 53, 115–123. 136. Braconnot, H. (1811) Sur la Nature des Champignons. Annales de chimie et de physique, 79, 265–304. 137. Blair, H. and Ho, T. (1980) Studies in the adsorption and diffusion of ions in chitosan. Journal of Chemical Technology and Biotechnology, 31, 6–10. 138. Rouget, C. (1859) Des substances amylacees dans le tissue des animaux, specialement les Articules (Chitine). Comptes Rendus, 48, 792–795. 139. Suh, H. and Matthew, H. (2000) Application of chitosan-based polysaccharide biomaterials in cartilage tissue engineering: a review. Biomaterials, 21 (24), 2589–2598. 140. Ravi, K.M. (2000) A review of chitin and chitosan applications. Reactive and Functional Polymers, 46, 1–27. 141. Shukla, S., Mishra, A., Arotiba, O. and Mamba, B. (2013) Chitosan-based nanomaterials: a state-of-the-art review. International Journal of Biological Macromolecules, 59, 46–58. 142. Lotmar, W. and Picken, L. (1950) A new crystallographic modification of chitin and its distribution. Experientia, 6 (2), 58–59. 143. Rudall, K. (1963) The chitin/protein complexes of insect cuticles. Advanced Insect Physiology., 1, 257–313.

Microalgae Bioproduction – Feeds, Foods, Nutraceuticals, and Polymers

111

144. Blackwell, J. (1973) Chitin, in Biopolymers (eds A. Walton and J. Blackwell), Academic Press, New York, pp. 474–489. 145. Rudall, K. and Kenchington, W. (1973) The chitin system. Biological Reviews, 40, 597–636. 146. Paralikar, K. and Balasubramanya, R. (1984) Electron diffraction study of alpha-chitin. Journal of Polymer Science, Polymer Letters Edition, 22, 543–546. 147. McLachlan, J. and Craigie, J. (1966) Chitin fibers in Cyclotella cryptica and growth of C. cryptica and Thalassiosira fluviatilis, in Some Contemporary Studies in Marine Science (ed. H. Barnes), George Allen and Unwin Ltd., London, England, pp. 511–517. 148. Walsby, A. and Xypolita, A. (1977) The form resistance of chitan fibres attached to the cells of Thalassiosira fluviatilis Hustedt. British Phycology Journal, 12, 215–223. 149. Morin, L., Smucker, R. and Herth, W. (1986) Effects of two chitin synthesis inhibitors on Thalassiosira fluviatilis and Cyclotella cryptica. FEMS Microbiology Letters, 37, 263–268. 150. McLachlan, J., McInnes, A. and Falk, M. (1965) Studies on chitin-poly-nacetylglucosamine fibers of diatom Thalassiosira fluviatius Hustedt. 1. Production and isolation of chitin fibers. Canadian Journal of Botany, 43, 707–713. 151. Falk, M., Smith, D., McLachlan, J. and McInnes, A. (1966) Studies on chitan (β-(1→4)-linked 2-acetamido-2-deoxy-β-d-glucan) fibers of the diatom Thalassiosira fluviatilis Hustedt. II. Proton magnetic resonance, infrared, and X-ray studies. Canadian Journal of Chemistry, 44, 2269–2281. 152. Blackwell, J., Parker, K. and Rudall, K. (1967) Letter to the editor: chitin fibers of the diatoms Thalassiosira fluviatilis and Cyclotella cryptica. Journal of the Marine Biology Association U. K., 28, 383–385. DOI:10.1016/S0022-2836(67)80018-4. 153. Dweltz, N. and Colvin, J. (1968) Studies on chitan (β-(1→4)-linked 2-acetamido-2-deoxyβ-d-glucan) fibers of the diatom Thalassiosira Fluviatilis Hustedt. III. The structure of chitan from X-ray diffraction and electron microscope observations. Canadian Journal of Chemistry, 46, 1513–1521. 154. Minke, R. and Blackwell, J. (1978) The structure of α-chitin. Journal of Molecular Biology, 120, 167–181. 155. Gardner, K. and Blackwell, J. (1975) Refinement of the structure of β-chitin. Biopolymers, 14, 1581–1595. 156. Saito, Y., Okano, T., Gaill, F. et al. (2000) Structural data on the intra-crystalline swelling of β-chitin. International Journal of Biological Macromolecules, 4 (4), 981–986. 157. Saito, Y., Kumagai, H., Wada, M. and Kuga, S. (2002) Thermally reversible hydration of β-chitin. Biomacromolecules, 3 (3), 407–410. 158. Merz, C. (2008) Investigation and evaluation of a bi-polar membrane based seawater concentration cell and its suitability as a low power energy source for energy harvesting/MEMS devices. Doctoral dissertation. University of South Florida. 159. Jeon, Y., Park, P. and Kim, S. (2001) Antimicrobial effect of chitooligosaccharides produced by bioreactor. Carbohydrate Polymers, 44, 71–76. 160. Kim, S. and Ye, J. (2011) Continuous production of chitooligosaccharides by enzymatic hydrolysis, in Chitin, Chitosan, Oligosaccharides and Their Derivatives (ed. S. Kim), CRC Press, Florida, pp. 47–51. 161. Ahmed, A., Taha, R., Mohajer, S. et al. (2012) Preparation, properties, and biological applications of water soluble chitin oligosaccharides from marine organisms. Russian Journal of Marine Biology, 38 (4), 351–358.

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162. Songsiriritthigul, C., Pantoom, S., Aguda, A. et al. (2008) Crystal structures of Vibrio harveyi chitinase a complexed with chitooligosaccharides: implication for the catalytic mechanism. Journal of Structural Biology, 162, 491–499. 163. Muzzarelli, R. and Tubertini, O. (1968) Chitin and chitosan as chromatographic supports and adsorbents for collection of metal ions from organic and aqueous solutions and sea water. Talanta, 16, 1571–1579. 164. Roberts, G. (1992b) Chitin Chemistry, Macmillan, London, pp. 206–229. 165. Kurita, K. (2006) Chitin and chitosan: functional biopolymers from marine crustaceans. Marine Biotechnology, 8, 203–226. DOI:10.1007/s10126-005-0097-5. 166. Kim, S. and Rajapakse, N. (2005) Enzymatic production and biological activities of chitosan oligosaccharides (COS): a review. Carbohydrate Polymers, 62 (4), 357–368. 167. Ngo, D., Kim, M. and Kim, S. (2008) Chitin oligosaccharides inhibit oxidative stress in live cells. Carbohydrate Polymers, 74, 228–234. 168. Thadathil, N. and Velappan, S. (2014) Recent developments in chitosanase research and its biotechnological applications: a review. Food Chemistry, 150, 392–399. 169. Liang, T., Huang, C., Dzung, N. and Wang, S. (2015) Squid pen chitin chitooligomers as food colorants absorbers. Marine Drugs, 13, 681–696DOI:10.3390/md13010681.

6 Innovations in Crustacean Processing: Bioproduction of Chitin and Its Derivatives Heather Manuel Centre for Aquaculture and Seafood Development, Fisheries and Marine Institute of Memorial University of Newfoundland, Canada

6.1

Introduction

The global seafood sector is perhaps the most complex of all food sectors. It is based on more species (about 1000 commercial species) and a wider range of processing technologies compared to any other food sector [1]. Total world fisheries production from wild capture and aquaculture sources exceeded 160 million tonnes (Mt) in 2013, with wild capture fisheries accounting for 92 Mt and aquaculture contributing 70 Mt [2]. Wild capture fisheries production has been relatively stable for the past 30 years at approximately 90 Mt, whereas aquaculture production has increased by 8–9% in the same time period [3]. Eighteen countries produce more than 75% of the world capture fisheries production with the top three producers being China (>13 Mt), Indonesia (5.4 Mt) and the United States (5.1 Mt) [4]. The Food and Agriculture Organization (FAO) has reported that 28.8% of fish stocks in 2011 were overfished, meaning that they were fished at biologically unsustainable levels and require strict management plans to Fuels, Chemicals and Materials from the Oceans and Aquatic Sources, First Edition. Edited by Francesca M. Kerton and Ning Yan. © 2017 John Wiley & Sons Ltd. Published 2017 by John Wiley & Sons Ltd.

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rebuild stocks to levels where they are biologically sustainable [4]. In 2012, more than 136 Mt (86%) of the world fish production was utilized for direct human consumption, 16.3 Mt (10%) was reduced to fishmeal and fish oil while the remaining 5.4 Mt (4%) went to other non-food uses such as bait, aquaculture feed, animal feed and pharmaceutical uses [4]. While the seafood sector contributes significantly to the world requirement for protein, it also has a reputation as being the most wasteful food sector. Approximately 70 Mt of fish is processed by filleting, freezing, canning or curing, and these activities generate 30–50% waste [4]. In 2009, approximately 46% of the landed volume (by weight) of all seafood produced in Canada was discarded as waste [5]. Similarly, in 2007, it was estimated that about 47% of South Korea’s seafood harvest by weight was by-product. According to the US Department of Agriculture, over half of the world production of fish ends up as by-product (or waste) [1]. The tuna canning industry generates up to 65% solid waste, and fillet production of farmed salmon generates 45–50% waste, including belly flaps, heads, bones, frames, viscera, skin, gills and dark muscle [4]. Of particular relevance to this chapter, crustacean processing generates anywhere from 35 to 70% waste. Shrimp destined for cooked and peeled products result in 60–70% waste, and snow crab produce about 35% waste when processed into individually quick frozen (IQF) cooked sections. Fish processing discards contain high-quality proteins, lipids, ω-3 fatty acids, micronutrients (e.g. vitamins A, D, riboflavin and niacin) and minerals (e.g. iron, zinc, selenium and iodine) [4]. Crustacean processing discards contain valuable products including proteins, lipids, astaxanthin, organic acids, essential amino acids, chitin and calcium [6]. Today, consumers are much more aware and better educated about the health and environmental impacts of the use of chemicals in aquaculture and seafood production. This increased awareness, coupled with a desire for environmental sustainability, has led many consumers to search for more natural products produced without chemical additives and harvested using sustainable practices. With respect to value-added products such as nutraceutical and natural health products which can be extracted from marine processing by-products, consumers want high-quality products with high biological activity [1, 7, 8]. In this regard, processing efficiencies are of particular importance. Seafood processors must be responsive and flexible to industry and consumer needs. They must be able to enhance the quality of and add value to their products while adhering to buyer and market specifications for environmentally sustainable products. As global demand for fish protein and sustainable seafood increases, the seafood sector can no longer afford the processing inefficiencies of the past which generated more than 50% waste (by-product). Proper biowaste management will be critical to meet regulatory requirements and the growing ecological and environmental concerns of the local and international markets [7]. With respect to traditional crustacean processing, large volumes of biomass waste are generated which often end up in landfills or are dumped at sea. There

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has been an increased research interest in the twenty-first century focusing on the extraction of high-value speciality products (e.g. nutraceuticals, pharmaceuticals and speciality chemicals) from food processing wastes using biotechnology. While the processing of such biomass waste does produce high-value products, this often requires more processing steps with extra costs. To be economically sustainable, all inputs need to be converted to useful products with minimum processing steps. Conventional food processing technologies can be applied to some extent; however, they do have limitations (e.g. overheating, high-energy use and loss of end-product functionality), and therefore, innovative technologies must be developed [8–10]. The scope of this chapter is to highlight and review some of these innovative technologies and how they can be implemented through a biorefinery approach to optimize the value chain of crustacean biomass with a particular focus on bioproduction of chitin, chitosan and chito-oligosaccharides from shrimp and crab.

6.2

Innovations in Crustacean Processing

6.2.1 6.2.1.1

Conventional Processing Technologies Cold-Water Shrimp (Pandalus borealis)

Atlantic Canada, Greenland, Norway, Denmark, Iceland, Faroe Islands and Russia are major producers of cold-water shrimp Pandalus borealis – the primary cold-water shrimp resource in the North Atlantic (Figure 6.1). Northern shrimp are typically harvested from near-shore or offshore harvest sites using commercial fishing trawlers. Offshore shrimp are processed at sea onboard factory freezer trawlers (FFTs). The shrimp are washed in seawater, and then, they are blast frozen at −40 ∘ C, shell-on. Offshore shrimp processed in this manner are referred to as

Figure 6.1 Northern cold-water shrimp, Pandalus borealis. (Courtesy of: Centre for Aquaculture and Seafood Development (CASD), Marine Institute, Memorial University of Newfoundland (MUN).)

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Figure 6.2 Shrimp in trawl positioned above holding pound. (Courtesy of: CASD, Marine Institute, MUN.)

‘industrial shrimp’ and are sold shell-on raw frozen to the consumer. Alternatively, the shrimp may be cooked shell-on prior to freezing and sold as shell-on cooked frozen [11]. Inshore shrimp are not processed at sea. They are processed fresh onshore. In Atlantic Canada, shrimp are harvested using trawl technology. When the trawl is retrieved from the ocean floor, it is hoisted high above the deck with the cod end positioned above the holding pound (Figure 6.2). The cod end is opened, and the shrimp are released onto the deck (Figure 6.3) and transferred to bagging stations either above the deck or below in the fish hold. Shrimp are loaded by hand into buckets lined with mesh bags and filled to a weight of 26 lb. The shrimp bags are placed in the fish hold where they are directly iced until they reach shore. Upon landing, the shrimp are unloaded at the processing plant’s receiving dock where they are placed in a maturation solution containing phosphates/sulfites for 24–36 h prior to processing. The maturation process aids peeling (i.e. removal of shell and head material) as inshore shrimp are typically processed into cooked and peeled products and sold as frozen meat. Following maturation, the shrimp undergo immersion cooking at 98 ∘ C for 1–2 min (total cooking time may be 10–30 min depending on the cooking process), or continuous steam cooking at 100 ∘ C, followed by cooling (immersion bath or cold-water spray), automatic peeling and grading, glazing, individually quick freezing, and packaging. Shrimp processed in this manner are referred to as ‘cooked and peeled’ (Figures 6.4 and 6.5). 6.2.1.1.1 Onboard Handling In the shrimp fishery in Atlantic Canada, the catch has traditionally been stored and handled in polyethylene mesh bags filled to a weight of 26 lbs [12]. This practice can be detrimental to the quality of landed shrimp when it is considered that the shrimp in the bottom of the fish hold are damaged due to the weight of the shrimp and ice on the top, and an aluminium pound board is placed on top of the last layer

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Figure 6.3 Shrimp held in pound. (Courtesy of: CASD, Marine Institute, MUN.)

Cold water shrimp processing (cooked and peeled shrimp) Near shore shrimp

Caught in trawl

Fish hold

Off loading

Boxed/bagged direct iced Cooling

Cooking

Pre-cleaning

98°C, 1–2 min

Peeling and grading

Cleaning and separating

Maturation 24–36 h @ 4° C

Meats

Glazing

30–35% yield

Inspection and packaging

Figure 6.4

IQF

Process flow for cooked and peeled shrimp.

of shrimp. This causes crushing of the shrimp in the lower layers and results in greater yield loss due to breakage and drip loss, as well as autolytic deterioration due to the release of digestive enzymes. In addition, inshore vessels are at sea fishing up to 3 days. Therefore, the first day’s catch is already 3 days old before it

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Figure 6.5 Commercial processing of cooked and peeled shrimp. (Courtesy of: SOF, Marine Institute, MUN.)

is landed and offloaded at the processing plant. The 3-day-old shrimp will then be placed in the maturation solution for up to 2 days prior to production. Thus, the shrimp may be 5 days old by the time they are actually processed into the final cooked and peeled products. The age range of the landed shrimp coupled with crushing damage due to storage and handling practices has resulted in inconsistent product quality [12]. Boxing of shrimp is a method that has traditionally been used by the Scandinavian shrimp fishing fleet to hold their catch while at sea. Boxing involves mixing shrimp and ice in a specific ratio and storing in 380 L insulated tubs/boxes. The vessels used in the Scandinavian fishery are larger than those used in Atlantic Canada, and they are designed with an open-hold concept to facilitate boxing of the product. The shrimp products exported by these countries are seen by the marketplace as superior products to Atlantic Canadian products. The higher quality is thought to be linked to handling procedures that include boxing of the raw material instead of bagging [12]. A study comparing boxing and bagging of shrimp catches conducted in 2003–2004 concluded that boxing of shrimp catches provides improvements over bagging in terms of the landed quality of shrimp [12]. Improvements were noted in the finished product colour (Figure 6.6). There was also a reduction in the amount of broken shrimp, and freshness was maintained over a longer period of time. Other advantages of boxing were related to reductions in handling issues, less time to store the product, and reduced time to offload the vessel. However,

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(b)

Figure 6.6 Boxed shrimp after processing (a); and bagged shrimp after processing (b). (Courtesy of: CASD, Marine Institute, MUN.)

for the Atlantic Canadian fleet, to switch from mesh bags to insulated tubs will require a high initial investment. Mesh bags cost about $0.35 each, whereas the insulated boxes cost about $300 each, and each vessel would need 160–200 boxes. Vessels under 55 ft will have difficulty carrying more than 100 boxes. Therefore, modifications to the fish hold would be required in order to accommodate the insulated boxes, or smaller vessels will have to limit the amount of product they will be able to carry per trip which would not be economically feasible. Anecdotal information suggests that since the release of the study results, about 50% of the NL shrimp fishing fleet has adopted a modified boxing method using fish tote pans, the other 50% are still using the mesh bag system. 6.2.1.1.2 Maturation Process After landing and offloading, shrimp are placed in a maturation solution for a period of 24–36 h. The maturation solution is maintained at 0–4 ∘ C and consists of a 1–10% solution of polyphosphates in which the shrimp are immersed. The solution is usually recycled and periodically refreshed with polyphosphate addition and eventually discarded every 24–36 h. This technique increases the meat yield from 20% to 30% as it makes the removal of shells easier during the mechanical peeling process [13–15]. Katch 150, for example, is a speciality blend of polyphosphates designed for use in the process of deshelling shrimp [16]. Katch 150 provides increased yields while improving quality, colour and texture. Phosphates do not

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generally penetrate the shell and are only functional prior to cooking. In addition to providing a better yield, phosphates have been shown to increase consumer acceptability of cooked and peeled shrimp [13]. 6.2.1.1.3 Cooking and Blanching Equipment Commercial shrimp cooking is an important processing step which has a direct impact on product yield and quality (i.e. colour, flavour and texture). There are two main types of commercial shrimp cooking equipment currently in use: continuous-belt cookers and batch-immersion cookers. KM Fish Machinery offers both continuous and batch cookers in a variety of sizes [17]. Mepaco offers continuous cookers designed to use direct/indirect steam, oil/water immersion or thermal screw configuration [18]. Innotec’s Impingement Flash Cooker uses a patented water cooking technique to cook shrimp at relatively low temperatures. The process can be controlled using multiple adjustable-temperature zones and a specially designed cooking belt. Innotec describes the technology as a gentle way of cooking, resulting in improved quality, better yield and optimum food safety [19]. 6.2.1.1.4 Automatic Peelers and Graders A typical shrimp peeling system for cold-water shrimp consists of peelers that are automatic bulk fed. The peelers are a series of inclined rollers arranged such that a smaller roller is placed between two adjacent larger rollers to form a peeling ‘nip’. The peeling rollers operate via a drive mechanism which rotates the larger rollers in alternating rotational directions. The larger rollers nip the shrimp shell, thus pulling it away from the meat. From there, the shrimp meat is cleaned, and any remaining shell and unwanted material are separated as a waste material from the cooked and peeled meat. The final product is then IQF, inspected and graded prior to final packaging. 6.2.1.2

Snow Crab (Chionoecetes opilio)

Canada, the world’s largest producer of snow crab, Chionoecetes opilio (Figure 6.7), accounts for approximately 2/3 of the global supply. About 75% of Canada’s snow crab exports go to the United States, with the rest being destined for China and Japan [20]. Snow crab are harvested using conical pots (Figure 6.8). After the pots are hauled, the crab are placed in the vessel’s fish hold and are either directly iced or placed in a refrigerated seawater (RSW) system. When the vessel lands, the crab are offloaded, iced and transported to the processing plant where they are sorted and graded according to size and liveliness condition. The crab are then placed in a warmwater bath prior to butchering. During butchering, the carapace, gut, viscera, liver and gills are removed. The remaining crab sections are cleaned and bled in

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Figure 6.7 Snow crab (Chionoecetes opilio) in pans for stowage in fish hold. (Courtesy of: CASD, Marine Institute, MUN.)

Figure 6.8 Conical crab pot. (Courtesy of: CASD, Marine Institute, MUN.)

an ice-water bath or slurry system. The cleaned sections are then cooked either by immersion in boiling water at 98 ∘ C for ∼12 min or via continuous-steam cooking to an internal temperature of 71 ∘ C. Following the cooking process, the sections are cooled to 4 ∘ C by cold-water immersion or a cold-water spray. From there, the sections are brine dipped and frozen at −30 ∘ C. The frozen sections are then packaged and sold as shell-on clusters/sections or may be further processed and sold as meat combo packs, snap-and-eat clusters/sections, claws (cap-off) or split clusters/sections [21] (Figure 6.9).

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Figure 6.9 Cooked crab sections in master carton. (Courtesy of: CASD, Marine Institute, MUN.)

6.2.1.3

Crustacean Biomass Waste

In 2014, the total commercial fish landings in Canada were reported to be 832,414 Mt, which included 329,821 Mt of crustaceans (i.e. lobster, shrimp, crab) [22]. In 2011, the CASD estimated that 46% of total Canadian landings are unutilized (or wasted) and that 25% of the national fish waste comprised of crustacean processing discards [5]. Therefore, of an estimated 382,910 Mt fish waste generated in Canada in 2014, 208,103 Mt of this was attributed to crustacean discards representing 54% of the total crustacean landings. A similar situation likely exists in other regions of the world with fishing industries. Most of this unutilized raw material is dumped at sea or in landfills with significant disposal costs to industry, lost economic opportunity and has a negative impact on the environment. During the production of cooked and peeled cold-water shrimp, up to 70% of the landed weight will end up as processing waste consisting of the shell, head, viscera and protein which is removed during the peeling, washing and separating steps (Figure 6.10). The production of IQF-cooked snow crab sections generates ∼35% waste comprised of the carapace (shell), gut, viscera and protein (Figure 6.11). Shrimp and crab processing waste contains high-value products including chitin, astaxanthin, calcium and protein which could be recovered for the production of high-end nutraceutical and pharmaceutical products. This is further explored in Section 6.3. 6.2.2

Innovations in Crustacean Processing

The role of technology in seafood processing has evolved rapidly over the past decade to support innovation, productivity, waste reduction, waste recovery and

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Figure 6.10

Figure 6.11 MUN.)

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Shrimp shell waste collected from shrimp peelers.

Crab shell waste from butchering process. (Courtesy of: CASD, Marine Institute,

utilization, increase shelf life, improve food safety and facilitate exports [1]. In recent years, there have been many advances made with respect to processing technologies specifically for crustaceans. These innovations have been developed with the goal of achieving quality and yield improvements, processing efficiencies and cost reductions and offsetting decreases in the labour force. In developed countries, the ageing workforce together with low-cost competing products from Asia and the difficulty of attracting and keeping skilled processing line workers further complicates the seafood processing industry. This has

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sparked a trend in developed countries towards improving processing technologies and processing automation in a traditionally labour-intensive sector. In order to be competitive in today’s market, seafood processors have to produce high value-low cost products [1]. 6.2.2.1

Ice Slurry Systems

For crustacean processing, chilling technologies are critical throughout the production process to ensure product quality and minimize deterioration. Traditionally, flake-ice and RSW systems have been employed for rapid chilling to decrease the final product temperature to just below 0 ∘ C. However, due to the highly perishable nature of crustaceans, they must be rapidly chilled to subzero temperatures immediately after harvesting/butchering to prevent spoilage. Newer chilling systems have recently enabled the storage of seafood at subzero temperatures through the addition of salts or other compounds to ice–water mixtures which are usually referred to as ‘ice slurry systems’. Such systems have been receiving increasing attention for the storage and preservation of aquatic foods due to their faster chilling rate in comparison with traditional flake-ice or RSW and reduced physical damage to the product. The spherical ice particles of an ice slurry mixture induce less damage to aquatic food tissue compared to traditional flake-ice particles [23]. Other major benefits of ice slurry systems are covered in detail by Pineiro et al. [23]. Ice slurry is a homogeneous mixture of small ice particles and carrier liquid which can be pumped. The liquid can be pure freshwater or a binary solution consisting of water and a freezing-point depressant (e.g. sodium chloride, ethylene glycol, ethanol or propylene glycol). The size, shape and smoothness of the ice particles are important characteristics for the slurry to be an effective coolant. Kauffeld et al. [24] provide a detailed discussion on the characteristics of ice particles in ice slurry and their effect on cooling and suggest that globular ice particles make better ice slurry compared to dendritic ice particles. Most applications of ice slurry are for indirect-contact cooling. However, it is used for direct-contact cooling in fish processing applications. The use of ice slurry systems for shellfish was first reported by Chinivasagam et al. [25], who studied the spoilage patterns of five Australian prawn species. Their results indicated that storage in slurry ice increased the shelf life of prawns from 10–17 days to more than 20 days and decreased the development of volatile compounds in comparison to prawns stored in flake ice. In 2002, Huidobro et al. [26] evaluated the effect of slurry ice and flake ice on the quality of shrimp stored onboard, with the main focus on shell appearance (i.e. brightness). This study indicated that storage in ice slurry caused the development of dull colour in shrimp shell and should only be used if the final product is sold shell-off. This dull colour may indicate that the

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water-soluble astaxanthin pigment normally present in shrimp is leached into the ice slurry mixture. This may have an impact on the quality and yield of potential bioactive compounds that could be extracted from the shells. The choice of ice slurry system must be chosen carefully based on the intended cooling application as different systems produce different types of ice particles which ultimately affects the effectiveness of cooling. There are more than 700 ice slurry systems in use in the fishing industry worldwide with Iceland, Japan and Norway among the top three users of the technology [24]. In Canada, ice slurry systems have been customized for installation onboard shrimp and crab vessels to improve quality and yield of the catch and to decrease operating costs. 6.2.2.2

Automated Crab-Butchering Machines

Crab butchering is a labour-intensive process that requires skilled workers who remove the mandibles and the carapace from the crab prior to separating it into two sections from which the gills and viscera are subsequently removed. Due to a decreasing labour supply, snow crab processors have been seeking more effective, automated mechanical processes to complete this step in their production process. In 2010, a Newfoundland-based company, Quinlan Brothers Ltd (QBL), with support from the Department of Fisheries and Aquaculture (DFA), the Canadian Centre for Fisheries Innovation (CCFI), National Research Council (NRC) and Centre for Aquaculture and Seafood Development (CASD), developed a viable prototype of an automated crab-butchering machine [27]. QBL has since commercialized this technology and has incorporated it into its crab processing line. In 2014, the Baader Group launched its new and revised automated crab-butchering machine called the BAADER 2801 (replaces the CB801) [28]. After 2 years of research and development and working closely with snow crab processors to automate the manual butchering process, Baader now offers an automated solution that incorporates new electronics, new butchering methods, a smaller footprint and more emphasis on hygiene and safety [28]. 6.2.2.3

CoolSteam® Cooking

The CoolSteam® Technology developed by Laitram Machinery (Figure 6.12) uses a forced convection method in which a low-temperature mixture of air and steam is constantly circulated inside the cooking chamber with uniform and efficient heat distribution. This provides a more consistent cook. Shrimp and crab processors in North America have been replacing older immersion cookers with the CoolSteam cooking technology due to a number of advantages this technology provides over immersion cooking, namely improved quality, improved yield, cost and energy savings imparted by lower cooking temperatures and more efficient heat distribution.

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Figure 6.12

6.2.2.4

Laitram CoolSteam® demo cooker. (Courtesy of: CASD, Marine Institute, MUN.)

High-Pressure Processing

High-pressure processing (HPP) of foods was first commercialized in Japan in 1992 as a means of microbial inactivation in jams and fruit juices. According to Smelt et al. [29], high pressure induces effects that result in vegetative cell death, including the following: (i) unfolding of globular proteins [30]; (ii) membrane damage (e.g. detachment and inactivation of membrane proteins) [31]; (iii) disintegration of ribosomes [32]; (iv) intracellular pH changes [33]. The technology has since been applied to a range of foods such as deli meats, bacon, guacamole, salsa, fish and shellfish [34]. Unlike thermally processed foods, HPP-treated foods retain the appearance, flavour, texture and nutritional qualities of the unprocessed product [29, 34, 35]. More recently, the technology has been applied commercially to bivalves (oysters) and crustaceans (lobster) to aid in raw meat removal from the shell. Industrially HPP systems (Figure 6.13) consist of either a vertical or a horizontal HP vessel and an external pressure-generating device such as a single-acting hydraulically driven pump [36]. HPP involves the application of high hydrostatic pressure to packaged foods or whole raw shellfish (i.e. in the shell). The pre-packaged food or whole raw shellfish is placed into a carrier

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Figure 6.13

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NC Hiperbaric Wave 6600 55L horizontal high-pressure processing system.

which is automatically loaded into the HP vessel, and the vessel plugs are closed. Water is pumped into the vessel until the desired maximum pressure is reached (pressurization). Most HP vessels operate up to a maximum pressure of 50,000–87,000 psi. The pressure is maintained for the desired dwell time (usually 1–2 min). Following the HPP cycle, the pressure is gradually released (decompression), and the carrier is automatically ejected from the vessel. With respect to processing shellfish, there are two main objectives of applying HPP technology: (i) inactivation of Vibrio species (Gram-negative bacteria) in oysters; (ii) achievement of clean separation of meat from the shell to facilitate shucking (oysters, mussels) and meat extraction (lobster, crab). In the case of oysters, bands are placed around the shells prior to HPP to avoid loss of meat and prevent recontamination of the meat post HPP due to opening of the shells. Because the food product is completely surrounded by water during HPP, all molecules are subjected to the same amount of pressure at exactly the same time due to the isostatic principle of pressure transmission [37, 38], and consequently, the product form/shape is maintained. HPP technology provides clean separation of meat from the shell and facilitates a new approach to crab meat extraction (Figure 6.14). This offers potential

Figure 6.14

Raw HPP of snow crab meat. (Courtesy of: CASD, Marine Institute, MUN.)

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to open up new markets and dramatically increase the value of crab products. Commercializing HPP technology involves developing new technology for crab meat extraction and incorporating both the HPP and meat extraction technologies into a highly automated production system. Meat extraction is now done manually around the world, despite many efforts to find a better solution. For that reason, it is done mostly in low-wage countries. The use of HPP enables easier meat extraction and potentially a greater degree of automation for the extraction process. Research is currently ongoing with respect to meat extraction automation for high-pressure-processed snow crab at the Memorial University of Newfoundland through the CCFI and the CASD. Results from this work are anticipated to be available by December 2017.

6.3

Utilization of Marine By-Products

Many failed attempts have been made to commercialize high-value speciality products from crustacean by-products mainly due to the lack of a suitable strategy to optimize the biomass value chain. Most efforts have focused only on the recovery technologies without identifying the specific targeted applications of the final product. Other problems arise due to the following reasons: the raw materials are treated as waste rather than as input materials; the extraction process is very costly with too many processing steps; the processing methodology has no flexibility and cannot be easily adapted to account for biological variations in the biomass waste; traditional manufacturers are not ready to be the early adopters of the new technologies [8–10, 39]. Rustad et al. [39] proposed a three-step strategy for the utilization of marine by-products which, if properly implemented, would address the limitations identified earlier. The proposed strategy includes the following: 1. Development of technologies to take care of the by-products 2. Development of simple technologies to produce bulk products for further refining 3. Development of technologies to take care of the valuable components. As previously mentioned, one of the main limitations in using marine biomass waste as input feedstock for the extraction of additional value-chain products is how they are treated and handled. In order for marine biomass waste to be further utilized to extract high-value products suitable for human consumption, or use in biomedical applications, a number of criteria must be considered. First and foremost, high-quality by-products are required. Secondly, the yield of the desired products must also be high. Controlled and standardized processes that are flexible enough to handle biological variations, produce stable, safe and high-quality products and are cost-effective must be developed. The end products must have documented proof of bioactive, nutritional and functional properties [8–10, 39].

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In this section, we explore the processing technologies that can be applied to implement the proposed strategy with a focus on value-chain optimization of crustacean biomass waste using a biorefinery approach. In this scenario, the biomass waste becomes the input feedstock while the end-use products and their applications form the value chain [9]. By extending the value chain in this manner, all input materials can be converted to useful products with minimum processing. 6.3.1

Processing Technologies for Crustacean By-Products

As discussed in Section 6.2.2, a number of processing innovations have been developed to improve yield and quality of crustacean products. These technologies, while initially designed and implemented to improve yield and quality of traditional value-chain products (e.g. cooked and peeled shrimp, cooked snow crab sections), may also help improve the quality of the leftover unutilized materials (i.e. shell, heads, viscera, liver) which, in theory, could be used as the input material for the development of additional value-chain products (e.g. nutraceuticals, pharmaceuticals). In order for this strategy to succeed, however, these unutilized materials must be treated and handled as input materials rather than as waste streams. This will require modifications to existing processing lines and educating processors about how to take care of the unutilized raw materials so that the value chain can be extended beyond traditional products. Ice Slurry – It may not be best choice for shrimp if recovery of the pigment is the goal; however, for chitin extraction where pigment recovery is not the main objective, ice slurry may aid in decolourization of the shell, thereby minimizing the need for a chemical bleaching step in the process. For crab shell, ice slurry may delay oxidation of the shells and adhering meat and prevent the black/blue discolouration which often develops in crab shell waste due to the presence of polyphenol oxidase [40]. Automated Crab Butchering Machines – Currently, these are not well designed to collect the carapace, viscera, gills and liver that are removed during this step. These machines could be redesigned with a collection chute and conveyor system to rapidly flume these nutrient-rich materials into an appropriate holding/storage container until they can be further processed into higher value products such as chitin, bioactive peptides and protein hydrolysates. Cookers and Blanchers – New cooking technologies have been designed to cook crustaceans at lower temperatures and shorter time periods with less water. The impact of milder cooking could translate to higher quality shell and protein materials which could help transform these traditional waste streams into input materials for the recovery for value-added products. Peelers – The remaining shrimp shell waste that accumulates at the peeling step represents a valuable unutilized resource for the extraction of high-value bioactive compounds. Typical shrimp plants are not designed to properly collect and store this raw material for further utilization, and hence, it ends up in landfills or is

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dumped at sea. As with the crab-butchering technology, shrimp peelers could be redesigned with a collection chute and conveyor system to rapidly flume the shell materials into an appropriate holding/storage container until they can be further processed. HPP – The shell materials resulting from HPP of snow crab and lobster represent unutilized raw materials that could potentially be used for value-chain optimization for the extraction of high-value nutraceuticals, pharmaceuticals and so on. HPP rarely affects the primary structure of low-molecular-weight molecules such as peptides, lipids and vitamins due to the very low compressibility of covalent bonds at pressures below 2 GPa (∼290,000 psi) [38, 41–43], and therefore, the same may be true for other bioactive compounds contained in the shell, such as astaxanthin and chitin. It is also possible that the quality of these compounds may be higher in HPP shell than in traditional thermally processed shells, resulting in higher value products with potentially better yields. Finally, HPP could be applied as an additional safety measure for value-added products extracted from crustacean waste streams to inactivate spoilage/pathogenic microorganisms and enzymes and extend shelf life. 6.3.2

A Biorefinery Approach for Value-Chain Optimization of Crustacean Biomass Waste

The term biorefinery first appeared in the scientific literature in 2001, but it was not until 2007 when the biorefinery concept began to achieve more significance in scientific publications [44]. According to the International Energy Association Bioenergy [45], biorefining involves the processing of biomass in a sustainable manner to obtain marketable bio-based products (food, feed, chemicals) and bioenergy (biofuels, power, heat). Similarly, Cherubini [46] describes biorefining as the integration of biomass transformation processes and equipment for the production of fuels, energy and chemicals. Gonzalez-Delgado and Kafarov [44] define biorefining as the processing of sustainable biomass to obtain energy, biofuels and high-value products. The general biorefining concept uses a wide range of technologies to separate biomass into its principal constituents (e.g. carbohydrates, proteins, fats) which can be further transformed into value-added products and biofuels. The concept is similar to that of oil refineries which fractionate complex mixtures of crude oil feedstock to obtain multiple products (e.g. petroleum, diesel, gasoline, kerosene, lubricants, tar, etc.) [44]. Most biorefinery applications have focused on the production of biofuels from plant-based feedstock from both food (e.g. barley, corn, soybean wheat) and non-food crops (e.g. wood fuel, Camelina). In recent years, significant research efforts have focused on the development of biorefineries for land-based feedstocks from agricultural and forestry wastes, yet little attention has been paid to ocean-based feedstocks [44, 47–49]. Kerton et al. [48, 49] have suggested that a biorefinery could be developed using ocean-sourced

Innovations in Crustacean Processing: Bioproduction of Chitin and Its Derivatives

Harvesters

Seafood processors

Biomass feedstock

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Biorefinery

Biotechnology industry

Chitin

• Chitosan • Chito-

oligosaccharides

Proteins Raw shirmp

Cooked and peeled shrimp

Shrimp shells and heads Biomass transform -ation

Live crab

Cooked crab sections

Crab carapace Lipids

Pigments Calcium carbonate

• Amino acid • Bioactive peptides • Aqua feed

Value added products • Fatty acids • Omega-3 • Astaxanthin • Aqua feed • Calcium

Increasing $ value / increasing cost of production Figure 6.15 crab.

Biorefinery approach for value-chain optimization of cold-water shrimp and snow

feedstocks from finfish and shellfish waste particularly in rural, coastal areas from Southeast Asia to the Eastern Seaboard of the United States and Atlantic Canada. A proposed biorefinery approach for value-chain optimization of cold-water shrimp and snow crab is presented in Figure 6.15. In this model, the waste from traditional processing of cooked and peeled shrimp or cooked snow crab sections becomes the feedstock for the biorefinery which separates the unutilized shell components into their main constituents: chitin, proteins, pigments (astaxanthin) and lipids. These additional value-chain products can be further processed into higher value products such as chitosans, calcium carbonate, bioactive peptides, amino acids and fatty acids. Traditional transformation processes, however, require the application of harsh chemical treatments which pose a threat to the environment and, in the case of biomedical applications, could render these bioproducts unsuitable. According to Kerton et al. [48], the application of green chemistry may be a means of reducing the environmental impact in the valourization of crustacean waste. New extraction, oxidation, deproteination and demineralization methods, such as biological methods, will be necessary, for example, to mitigate the negative environmental impacts of more traditional chitin/chitosan production processes. It should also be noted that while more value can theoretically be extracted from shrimp and crab biomass under this model, the cost of innovative and multiple extraction processes could be prohibitive. Thus, not only are green technologies

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important, but sustainable cost-effective production methodologies must be developed for the model to be economically feasible.

6.4 6.4.1

Bioproduction of Chitin and Its Derivatives Background

Chitin was discovered in 1811 by French scientist Henri Braconnot, who named the material fungine [50]. Odier found the same substance in 1823 and called it chitine [51]. However, it was not until the 1950s that a sustained interest in chitin research, including its derivatives, developed. Since then, four broad areas of chitin research have developed including the following: (i) isolation from shellfish and other sources; (ii) structural studies and properties; (iii) chemical derivatization; (iv) applications for chitin, chitosan and their derivatives [52]. Chitin (Figure 6.16) is a linear amino polysaccharide composed of β-(1–4)-linked N-acetyl-d-glucosamine units which may be de-N-acetylated up to 60–80% [53–55] (to produce chitosan (Figure 6.16)). Chitin contains 6–7% nitrogen, whereas chitosan contains 7–9.5% nitrogen [53, 56]. The source of chitin affects its crystallinity, purity, polymer chain arrangement and its properties [55]. There are three polymorphic forms of chitin: α-, β- and γ-chitin. The α-form is the most abundant and is found in crab and shrimp shells. β-Chitin is found in molluscs such as squid, and γ-chitin has been isolated from the stomach lining of squid and cuttlefish [56, 57]. In α-chitin, the chains are arranged in an antiparallel fashion which promotes strong hydrogen binding between the chains, resulting in a very tight and compact crystalline structure which requires harsh methods for extraction [56–58]. β-Chitin is arranged in parallel sheets and is held together by weaker intermolecular forces and can be easily, but irreversibly, transformed into α-chitin by steam. The γ-form is a combination of α- and β-chitin containing two parallel (𝛽) and one antiparallel (𝛼) strands [56, 57], which can also be converted to α-chitin by treatment with lithium thiocynate [59]. A critical evaluation of potential sources of chitin and chitosan concluded that shrimp, prawn and crab waste were the principal sources of this biopolymer and would remain so for the immediate future. Together, chitin and chitosan have gained an outstanding reputation with numerous applications in the fields of water engineering, cosmetics, paper engineering, textile engineering, food engineering, agriculture, photography, chromatographic separations, medical and pharmaceutical in the recent decades [55, 60, 61]. Chitin and chitosan are generally non-toxic, non-soluble in water and most organic solvents. Chitin is the second most abundant natural polymer and is formed from N-acetyl-d-glucosamine. It is the structural component of crustaceans, insects, arthropods, fungi and yeast. Shrimp and crab shell waste are the main commercial sources of chitin (i.e. α-chitin). Due to its highly crystalline structure and strong hydrogen bonds, chitin is not readily dissolved in common solvents. Therefore, it

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Chitin OH

OH

O

HO HO

O

OH

O

O

HO

NH

NH

O

O

OH

HO NH

n

O

O CH3

CH3

CH3

Chitosan OH HO HO

OH

O

O

O HO

NH2 Figure 6.16

NH2

OH O

O HO

n

OH

NH2

Deacetylation of chitin.

is often converted to its more N-deacetylated derivative, chitosan, which can be water-soluble. Traditional methods of chitin extraction are chemically intensive processes using large volumes and high concentrations of HCl, NaOH and ethanol. Long exposure of chitin to HCl can significantly modify its physicochemical properties; cleavage reactions can decrease the molecular weight, and significant deacetylation of the chitin can occur, both of which negatively affect the fundamental properties of the chitin isolated [62, 63]. Chitosans are characterized mainly by viscosity and degree of deacetylation (DDA). The control over these two parameters allows the production of a wide range of chitosans which can be used in medical, pharmaceutical, cosmetic, nutraceutical and industrial fields [61, 64]. The traditional method of chitosan production uses a high concentration and large volumes of caustic (up to 70% w/v) to which chitin is added followed by heating at 95–110 ∘ C for up to 2 h [48]. This is a chemically intensive and environmentally hazardous process. Chemical extraction methods may also produce toxic compounds, rendering the chitosan unsuitable for biomedical applications, and is a source of environmental pollution [57, 65]. Due to the limitations associated with traditional chemical extraction methods for chitin and chitosan, there have been significant efforts directed towards the use of more eco-friendly methods such as enzymatic methods, lactic acid fermentation (LAF), combined biological and chemical methods, as well as combined use of

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waste materials. In this section, we explore selected bioproduction methods for chitin, chitosan and their derivatives. 6.4.2

Isolation and Extraction of Chitin and Chitosan

Most traditional isolation methods of chitin from shrimp and crab shells involve three main processing steps following initial particle size reduction which include the following: (i) deproteination – removal of protein using strong alkali and heat treatment (e.g. 1–2% w/v KOH, 90 ∘ C for 2 h); (ii) demineralization – removal of minerals, mainly calcium carbonate, by treatment with strong acid (e.g. 5–7% w/v HCl for 2 h) and (iii) decolouration – removal of pigment using a bleaching/oxidizing agent (e.g. hydrogen peroxide, ethanol, sodium hypochlorite) to obtain a colourless product [57, 66, 67]. This process may be carried out on fresh or dried shells, and the demineralization and deproteination steps may be carried out in reverse order if pigment recovery is not a concern [67, 68]. Chitosan is produced by the deacetylation of chitin (Figure 6.16) using highly concentrated NaOH or KOH solution (40–50% w/v) at high temperature (>100 ∘ C) for 2 h. Chitosan is differentiated from chitin on the basis of the DDA or nitrogen content. The term chitosan is preferred when chitin reaches a DDA above 60% or a nitrogen content greater than 7% by weight [69, 70]. 6.4.2.1

Enzymatic Extraction

There has been limited industrial use of enzymatic methods for chitin extraction due to the high costs associated with such methods on the industrial scale [57]. However, enzymatic hydrolysis has generated significant research interest because it produces a higher grade chitin and subsequently higher grade chitosan compared to chemical methods. Commercially available proteolytic enzymes (Table 6.1) such as Alcalase (EC 3.4.21.62), chymotrypsin (EC 3.4.21.2) and papain (EC 4.3.22.2) have been used to remove protein and extract chitin from shellfish waste [71, 72]. Synowiecki and Al-Khateeb [86] extracted chitin from shrimp (Crangon crangon) waste using Alcalase to achieve 89.0% deproteination. The chitin thus obtained contained a residual protein content of 4.4–7.9%, which is about twice as high as that of the commercial product treated with NaOH [87]. Bacterial release of chitin was also achieved using proteases isolated from Pseudomonas maltophilia or Bacillus sp. TKU004 for deproteination [75, 76]. Gagne and Simpson [74] showed that residual protein levels in shrimp waste following deproteination with chymotrypsin and papain were 1.3% and 2.8%, respectively. However, a high enzyme-to-waste ratio (E/W) of 0.7% for chymotrypsin and 1% for papain was required for maximum deproteination. A study conducted by Jo et al. [77] comparing the effectiveness of deproteination of various commercial proteases found that Delvolase exhibited the highest DP for crab shell waste which, when treated with 1% Delvolase, reached

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Table 6.1 Sources of protease and chitin deacetylase enzymes for deproteination and deacetylation of chitin.

Protease enzymes Alcalase (EC 3.4.21.62) Chymotrypsin (EC 3.4.21.2) Papain (EC 4.3.22.2 Protease Delvolase A21 crude protease Chitin deacetylase enzymes CDA (EC 3.5.1.41)

Source

References

Commercial Commercial Commercial Pseudomonas maltophilia or Bacillus sp. TKU004 Commercial Bacillus mojavensis

[71–73] [71, 72, 74] [71, 72, 74] [75, 76]

Mucor rouxii Absidia coerulea Vibrio cholera Absidia corymbifera DY-9 Mortierella sp. DY-52 Saccharomyces cerevisiae Colletotrichum lindemuthianum ATCC 56676

[79] [6] [80] [81, 82] [83] [84] [85]

[77] [78]

85% DP within 1 day. However, complete removal of the residual protein associated with the chitin was not achieved using Delvolase. Gildberg and Stenberg [73] used Alcalase (2.4 1 FG) to deproteinate Northern shrimp (P. borealis) waste with the goal of obtaining a high-quality protein hydrolysate and producing chitosan from the resulting press cake. Their process showed that Alcalase treatment allowed about 70% of the total amino-N to be recovered without affecting the yield or quality of the chitosan subsequently produced. Younes et al. [78] used non-commercial Bacillus mojavensis A21 crude protease to obtain 88% deproteinization of shrimp (Metapenaeus monoceros) shells. The optimum process conditions included the following: an enzyme/substrate ratio of 7.75 U/mg; temperature of 60 ∘ C; incubation time of 6 h. The solid fraction was chemically demineralized and subsequently converted to chitosan by alkali treatment, yielding a chitosan with a low degree of acetylation (4%) and high antimicrobial activities. While enzymatic hydrolysis has been shown to produce a higher grade chitin compared to chemical methods, at least on a lab scale, enzymatic conversion to chitosan remains a challenge. For example, the conversion of chitin to chitosan has been studied on a laboratory scale using various sources of chitin deacetylase (CDA, EC 3.5.1.41) (Table 6.1). CDAs result in a chitosan that has a more regular pattern of acetylation (PA) compared to a chitosan produced using hot NaOH. CDAs recognize a specific pattern of four GlcNAc units in chitin of which one undergoes deacetylation [81]. CDA was first found in extracts from the fungus Mucor rouxii [79], but since then, several different fungal CDAs have been

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discovered [81]. However, the CDAs studied thus far have been ineffective in deacetylating natural insoluble crystalline chitin and only results in a 1% increase in the DDA [81, 82, 88]. Prakash et al. [6] used fungal CDA isolated from Absidia coerulea to deacetylate chitin. Their results suggested that even with surplus chitin substrate and high enzyme activity, only a small percentage of the substrate is transformed to chitosan. Win and Stevens [88] used various physical (heating, sonication, grinding) and chemical (derivatization, interaction with saccharides) pretreatments to try and increase the accessibility of the acetyl groups of crystalline chitin to fungal CDA; however, these treatments were ineffective. 6.4.2.2

Lactic Acid Fermentation

Chitin extraction using LAF has been performed on a lab scale; however, it is not yet used commercially due to the cost of lactic acid production (pennotec.com) [89]. An advantage of LAF over traditional chemical extraction methods is that it allows for the recovery of chitin, proteins and astaxanthin [67, 86, 90–92]. Lactic acid bacteria (LAB) typically grow between 10 and 50 ∘ C and require a carbon and a nitrogen source. At the beginning of the growth cycle, lactic acid is produced by the breakdown of a carbon substrate such as glucose or sucrose. The lactic acid results in an acidic pH, and this environment has been shown to effect the demineralization of shrimp shells, due to solubilization of calcium, but to varying degrees depending on the fermentation conditions. Deproteination is achieved by the enzymatic action of LAB on shell proteins, free ammonia released during shell solubilization and eventually catabolism of amino acids [90, 93]. Cira et al. [90] reported that the optimum LAF parameters using a Lactobacillus spp. strain B2 (isolated form shellfish waste) were 10% (w/w) sucrose with an innoculum level of 5% (v/w) which promoted high acidification (i.e. pH decreased from 7.5 to 4.6 and 0.53 mmol/g TTA) at 36 ∘ C, achieving 85% demineralization and 87.6% deproteinization after 6 days of incubation. In comparison, however, Adour et al. [93] reported low levels of demineralization (60%) and deproteinization (20.6%) of white shrimp shell when fermented with Lactobacillus helveticus at 30 ∘ C using 300 g/L glucose substrate and an innoculation level of 10% (v/v). Reasons cited for the low levels of demineralization and deproteinization, respectively, included the following: the initial alkalinity of the culture medium (8.5–9.0) likely due to the ammonia release from shells during their preparation (1.5–2 g/L) may have interfered with acidification of the culture medium; the presence of free ammonia as a second nitrogen source. Greene et al. [62] recently evaluated lactic acid demineralization of green crab shells. They found that contrary to their expectation, the rate of demineralization did not increase on a linear scale with increasing concentration of lactic acid. Instead, the rate of demineralization had a positive correlation with the conductivity of the solution, which measures mobility of the H+ ion in solution. Because

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lactic acid is very viscous, the more concentrated the solution, the lower the mobility of the H+ ion and, therefore, the lower the rate of demineralization as the H+ ion has limited ability to attack the shell. The use of undiluted (11.4 M) lactic acid resulted in only 5% removal of calcium carbonate after 180 min at room temperature, compared with 61% removal with 1.14 M and 53% removal with 2.28 M lactic acid. Other factors affecting the rate of lactic acid demineralization include temperature, shell-to-acid ratio and reaction time [62, 94, 95]. Greene et al. [62] reported that under ideal conditions, 90–95% of the calcium carbonate and other minerals can be removed from green crab shells in 90 min, but the remaining 5–10% took more than 350–400 min. This was attributed to the ‘shrinking core model’ in which the demineralization reaction slows over time because the reactive sites of the calcium carbonate (and other minerals) move deeper into the interior of the shell [62, 94]. 6.4.2.3

Integrated Extraction Methods

Neither enzymatic, nor LAF, alone has proven to be effective for the deproteination and demineralization of crustacean shells. To improve the recovery of chitin and reduce the amount of alkali and acid required, enzyme treatments combined with chemical treatments and LAF in combination with chemical treatments, both have been studied as an alternative to chemical extraction for chitin recovery [6, 62, 73, 90, 96–98]. Greene et al. [62], for example, did not obtain high rates of demineralization using lactic acid alone. However, by combining lactic acid (1:4 acid-to-water ratio) in a 1:1 molar ratio with 5.00 M HCl, demineralization was improved, and the amount of HCl required was reduced in comparison to using only HCl. The combined acid mixture resulted in 77.8% weight loss compared with only 37.5% using lactic acid alone. Gildberg and Stenberg [73] combined the use of a commercial protease, Alcalase (2.4 1 FG), to remove proteins from Northern shrimp shells (P. borealis) followed by chemical demineralization, deproteination and deacetylation of the press cake to produce chitosan. Using this method, 70% of the total amino-N was recovered as protein hydrolysate compared with less than 15% in the conventional chitosan production process. It was also possible to recover the astaxanthin pigment using this method. In another experiment, chitin obtained from LAF was further purified after pigment extraction, using acid and alkali treatments [90]. This method reduced the amount of chemicals required for chitin purification by 50–77% with respect to conditions previously reported in the literature [67, 99]. Other innovative approaches have included combining waste materials such as green crab shells and lactic acid from milk processing [62] and snow crab shells with lactic acid from rye grass fermentation [89]. The intent of this approach is to improve the economics of both chitin extraction and lactic acid production by combining them in a single biorefinery process. In this method, lactic acid dissolves shell calcium carbonate which buffers the fermentation reaction and increases the

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lactic acid yield. Adour et al. [93] investigated the feasibility of recovering chitin from white shrimp shells by LAF using date juice waste as a substrate. Date juice has a high mineral content which seems to have interfered with demineralization, but improved the proteolytic activity of L. helveticus, resulting in nearly complete deproteination of the white shrimp shells. 6.4.2.4

Physical Chemoenzymatic Chitin Extraction

Physical methods using particle size reduction are promising as alternative means of chitin extraction; however, obtaining high yields of purified chitin at low cost has proved challenging. It has been suggested that a combined approach using physical, chemical and enzymatic methods may provide a cost-effective and environmentally friendly compromise to the traditional chemically intensive chitin extraction process applied to crustacean shells [57]. In 2012–2014, the author and her research team at the CASD tested this hypothesis by developing a 240 kg batch pilot-scale demonstration system (Figure 6.17) which was based on a modified chemoenzymatic process with novel mechanical pretreatment for the production of chitin from shrimp and crab shell waste. The physical chemoenzymatic process included a series of mechanical pretreatment steps for particle size reduction, enzymatic removal of protein using commercially available protease enzymes, followed by mild acid treatment for demineralization which required 80% less HCl compared to the traditional process [100]. Although the pilot-scale physical chemoenzymatic chitin extraction process developed by the CASD effectively eliminated the use of chemicals in the deproteination step and significantly reduced the chemical usage in the demineralization step, the depigmentation step remains a chemically intensive process.

Rotary sieve

Control panel

Chemical dosing pump

Demineralization tank

Deproteinization tank

Wastewater collection tank

Figure 6.17

Chitin pilot processing line. (Courtesy of: CASD, Marine Institute, MUN.)

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Table 6.2 Biomedical, pharmaceutical and biotechnological applications of chitosan and chitosan derivatives. Potential applications

Principle properties/ characteristics

References

Surgical sutures Dental implants Artificial skin Rebuilding of bone, bone-filling agent Corneal contact lenses Time release drugs for animals and humans Encapsulating material

Biocompatible Biodegradable Renewable Film forming

[55] [55] [55] [55, 101]

Hydrating agent Non-toxic, biological tolerance

[55] [55, 101]

Hydrolysed by lyzosyme Wound-healing properties Efficient against bacteria, viruses, fungi Biocompatible, antimicrobial, blood clotting, wound healing Non-toxic, biocompatibility, cationic, chelating ability with DNA Antimicrobial, low toxicity, biodegradable, cationic–electrostatic interactions, porous structure, gel-forming properties, high affinity for in vivo macromolecules

[55]

Wound dressing Gene delivery Tissue engineering and wound healing

[101] [102, 103] [103, 104]

In addition, the CASD chitin pilot processing line generated large volumes of wastewater laden with astaxanthin and protein [100]. Ideally, pigment and protein recovery would be incorporated into the extraction process, or green oxidation catalysts could be used for pigment removal [48]. 6.4.3

Non-chemical Structural Modifications of Chitin and Chitosan

Biomedical, pharmaceutical and biotechnological applications (Table 6.2) are the strongest high-value growth markets for chitin, chitosan and their derivatives. The main developments in this field have included their use in wound dressings, controlled drug release, hair care and as a bone-filling agent produced from hydroxyapatite–chitin–chitosan composite [101]. Chitosan has unique physical, chemical and biological properties that make it commercially attractive; however, commercial processing to produce a consistent high-quality product has proven to be difficult and expensive [9]. Superior quality chitosan should have a DDA of 70–90% [6, 105], with a minimum of 78% DDA required for medical-grade chitosan [100]. Chitin has a very stable crystalline structure, and achieving the high DDA necessary to produce biomedical grade chitosan requires a long incubation period, or a multi-stage process, in a harsh chemical environment at a high temperature. This type of heterogeneous process

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Fuels, Chemicals and Materials from the Oceans and Aquatic Sources Table 6.3 Quality specification for medical-grade chitosan [100]. Parameter

Medical-grade chitosan

Appearance Protein (%) Total ash (%) Moisture (%) Viscosity (mPa s) DDA (%) Heavy metals (ppm) Insolubles (% (w/w)) Bacterial endotoxins (EU/g) Microbial enumeration (CFU/g)

White to off-white/beige powder