Enzymatic technologies for marine polysaccharides 9781138103078, 1138103071

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Content: Section 1: General view and sources of marine polysaccharides and oligosaccharides1. Marine biodiversity as a new source of promising polysaccharides: innovative polysaccharides emerging from the marine biodiversity[Sylvia Colliec-Jouault, Corinne Sinquin, Agata Zykwinska, and Christine Delbarre-Ladrat]2. Applications of marine polysaccharides in food processing[Vazhiyil Venugopal]3. The manufacture, characterization, and uses of fucoidans from macroalgae[J. H. Fitton, D. N. Stringer, S. S. Karpiniec, and A. Y. Park]4. Chemical and biological routes for the valorization of macroalgal polysaccharides[Valerie J. Rodrigues and Annamma A. Odaneth]5. Marine exopolysaccharides provide protection in extreme environments[Carla C. C. R. de Carvalho]6. Structural mechanisms involved in mild-acid hydrolysis of a defined tetrasaccharide-repeating sulfate fucan[Francisco F. Bezerra and Vitor H. Pomin]7. Biosynthesis and extrusion of ss-chitin nanofibers by diatoms[Gregory L. Rorrer]8. The mucus of marine invertebrates: Cnidarians, polychaetes, and echinoderms as case studies[L. Stabili]9. Biorefinery of unique polysaccharides from Laminaria sp., Kappaphycus sp., and Ulva sp.: structure, enzymatic hydrolysis, and bioenergy from released monosaccharides[Mark Polikovsky and Alexander Golberg]10. Fermentative production and application of marine microbial exopolysaccharides[Shweta Singh, Anjula Katoch, Rajwinder Kaur, Kulwinder Singh Sran, Bhupender Kumar, and Anirban Roy Choudhury]Section 2: Extraction techniques, structural determination, and methodologies to assess biological activities11. Marine polysaccharides: extraction techniques, structural determination, and description of their biological activities[O. Ibraheem and O. M. Babatunde]12. Fucoidan: a tool for molecular diagnosis and targeted therapy of cardiovascular diseases[Murielle Maire, Lucas Chollet, Lydia Rolland, Didier Letourneur, Cedric Chauvierre, and Frederic Chaubet]13. Marine polysaccharides as promising source of biological activities: extraction and purification technologies, structure, and activities[A. Mzibra, I. Meftah Kadmiri, and H. El Arroussi]14. Microwave-assisted conversion of marine polysaccharides[Shuntaro Tsubaki, Ayumu Onda, Tadaharu Ueda, Masanori Hiraoka, Satoshi Fujii, and Yuji Wada]15. Role of marine polysaccharides in treatment of metabolic disorders[Manigandan Venkatesan, Velusamy Arumugam, Rathinam Ayyasamy, Karthik Ramachadran, Subhapradha Namasivayam, Umamaheswari Sundaresan, Archunan Govindaraju, and Ramachandran Saravanan]Section 3: Enzymatic technologies16. Role of carbohydrate active enzymes (CAZymes) in production of marine bioactive oligosaccharides and their pharmacological applications[Md. Imran and Sanjeev C. Ghadi]17. Microbial enzymes and potential use in algal polysaccharide modifications[Daniela de Borba Gurpilhares, Lara Duraes Sette, and Adalberto Pessoa Jr.]18. Molecular modification of marine sulfated polysaccharides[Sutapa Biswas Majee, Dhruti Avlani, and Gopa Roy Biswas]19. Marine algae-degrading enzymes and their applications in marine oligosaccharide preparation[Benwei Zhu, Limin Ning, Yun Sun, and Zhong Yao]20. Enzymatic technologies of chitin and chitosan[P. V. Suresh]21. Enzymes used to produce glycosaminoglycan mimetics from marine polysaccharides[Christine Delbarre-Ladrat, Veronique Verrez-Bagnis, Sylvia Colliec-Jouault, and Agata Zykwinska]22. Production of value-added materials from alginate using alginate lyases and 4-deoxy-l-erythro-5-hexoseulose uronic acid-metabolic enzymes from alginolytic bacteria and marine gastropods[Takao Ojima, Ryuji Nishiyama, and Akira Inoue]

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Enzymatic Technologies for Marine Polysaccharides

Enzymatic Technologies for Marine Polysaccharides Edited by Antonio Trincone

CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2019 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed on acid-free paper International Standard Book Number-13 978-1-138-10307-8 (Hardback) This book contains information obtained from authentic and highly regarded sources. Reasonable efforts have been made to publish reliable data and information, but the author and publisher cannot assume responsibility for the validity of all materials or the consequences of their use. The authors and publishers have attempted to trace the copyright holders of all material reproduced in this publication and apologize to copyright holders if permission to publish in this form has not been obtained. If any copyright material has not been acknowledged, please write and let us know so we may rectify in any future reprint. Except as permitted under U.S. Copyright Law, no part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www.copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC), 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com

Contents Preface����������������������������������������������������������������������������������������������������������������������������������������������ix Introduction������������������������������������������������������������������������������������������������������������������������������������xi Acknowledgments������������������������������������������������������������������������������������������������������������������������xv Editor............................................................................................................................................ xvii Contributors��������������������������������������������������������������������������������������������������������������������������������� xix Section 1:  General view and sources of marine polysaccharides and oligosaccharides Chapter 1 Marine biodiversity as a new source of promising polysaccharides: innovative polysaccharides emerging from the marine biodiversity�����������3 Sylvia Colliec-Jouault, Corinne Sinquin, Agata Zykwinska, and Christine Delbarre-Ladrat Chapter 2 Applications of marine polysaccharides in food processing������������������������25 Vazhiyil Venugopal Chapter 3 The manufacture, characterization, and uses of fucoidans from macroalgae����������������������������������������������������������������������������������������������������� 47 J. H. Fitton, D. N. Stringer, S. S. Karpiniec, and A. Y. Park Chapter 4 Chemical and biological routes for the valorization of macroalgal polysaccharides������������������������������������������������������������������������������������������������������65 Valerie J. Rodrigues and Annamma A. Odaneth Chapter 5 Marine exopolysaccharides provide protection in extreme environments����������������������������������������������������������������������������������������������������������95 Carla C. C. R. de Carvalho Chapter 6 Structural mechanisms involved in mild-acid hydrolysis of a defined tetrasaccharide-repeating sulfate fucan������������������������������������������� 111 Francisco F. Bezerra and Vitor H. Pomin Chapter 7 Biosynthesis and extrusion of β-chitin nanofibers by diatoms�������������������������������������������������������������������������������������������������������������129 Gregory L. Rorrer

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Chapter 8 The mucus of marine invertebrates: Cnidarians, polychaetes, and echinoderms as case studies������������������������������������������������������������������������������151 L. Stabili Chapter 9 Biorefinery of unique polysaccharides from Laminaria sp., Kappaphycus sp., and Ulva sp.: structure, enzymatic hydrolysis, and bioenergy from released monosaccharides��������������������������������������������163 Mark Polikovsky and Alexander Golberg Chapter 10 Fermentative production and application of marine microbial exopolysaccharides����������������������������������������������������������������������������������������������189 Shweta Singh, Anjula Katoch, Rajwinder Kaur, Kulwinder Singh Sran, Bhupender Kumar, and Anirban Roy Choudhury Section 2: Extraction techniques, structural determination, and methodologies to assess biological activities Chapter 11 Marine polysaccharides: extraction techniques, structural determination, and description of their biological activities���������������������219 O. Ibraheem and O. M. Babatunde Chapter 12 Fucoidan: a tool for molecular diagnosis and targeted therapy of cardiovascular diseases��������������������������������������������������������������������������������������273 Murielle Maire, Lucas Chollet, Lydia Rolland, Didier Letourneur, Cédric Chauvierre, and Frédéric Chaubet Chapter 13 Marine polysaccharides as promising source of biological activities: extraction and purification technologies, structure, and activities�����������301 A. Mzibra, I. Meftah Kadmiri, and H. El Arroussi Chapter 14 Microwave-assisted conversion of marine polysaccharides�����������������������321 Shuntaro Tsubaki, Ayumu Onda, Tadaharu Ueda, Masanori Hiraoka, Satoshi Fujii, and Yuji Wada Chapter 15 Role of marine polysaccharides in treatment of metabolic disorders�������335 Manigandan Venkatesan, Velusamy Arumugam, Rathinam Ayyasamy, Karthik Ramachadran, Subhapradha Namasivayam, Umamaheswari Sundaresan, Archunan Govindaraju, and Ramachandran Saravanan Section 3:  Enzymatic technologies Chapter 16 Role of carbohydrate active enzymes (CAZymes) in production of marine bioactive oligosaccharides and their pharmacological applications�����������������������������������������������������������������������������������������������������������357 Md. Imran and Sanjeev C. Ghadi

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Chapter 17 Microbial enzymes and potential use in algal polysaccharide modifications�������������������������������������������������������������������������������������������������������� 375 Daniela de Borba Gurpilhares, Lara Durães Sette, and Adalberto Pessoa Jr. Chapter 18 Molecular modification of marine sulfated polysaccharides����������������������395 Sutapa Biswas Majee, Dhruti Avlani, and Gopa Roy Biswas Chapter 19 Marine algae–degrading enzymes and their applications in marine oligosaccharide preparation������������������������������������������������������������������������������ 417 Benwei Zhu, Limin Ning, Yun Sun, and Zhong Yao Chapter 20 Enzymatic technologies of chitin and chitosan���������������������������������������������449 P. V. Suresh Chapter 21 Enzymes used to produce glycosaminoglycan mimetics from marine polysaccharides����������������������������������������������������������������������������������������������������467 Christine Delbarre-Ladrat, Véronique Verrez-Bagnis, Sylvia Colliec-Jouault, and Agata Zykwinska Chapter 22 Production of value-added materials from alginate using alginate lyases and 4-deoxy-l-erythro-5-hexoseulose uronic acid–metabolic enzymes from alginolytic bacteria and marine gastropods�����������������������495 Takao Ojima, Ryuji Nishiyama, and Akira Inoue Index��������������������������������������������������������������������������������������������������������������������������������������������� 511

Preface Recent estimation of global biomass, based on a more accurate description of biodiversity and size of species populations, using the recent effort of global sampling like those conducted by national and international expeditions, revealed that land biomass surpasses the marine biomass by about two orders of magnitude. However, in the oceans, 70% of the tree of life is present, highlighting the biological diversity of marine environment compared to land dominated by the plant kingdom. When looking at the biomass composition, polysaccharides represent the most abundant biomacromolecules that have many biological functions, including energy storage, and play a structural role notably in living cell walls and mediate the intra- and intercellular recognition within one organism or between organisms. Therefore, because of the wide biological diversity of marine environment, marine polysaccharides show an immense structural diversity that needs to be explored. What makes marine polysaccharides different? In polysaccharides, monosaccharide units are joined one to the other by acetal linkages (glycosidic linkage) to give long chains. The wide stereochemical variability of monosaccharides and the numerous linkage possibilities offer an immense number of possible polysaccharide structures. Composition of polysaccharides biosynthesized by prokaryote and eukaryotes revealed very diverse monosaccharide variability and, sometimes, with very rare residues, notably when these macromolecules are involved in a defense pathway and/or pathogenicity. Based on known structure of marine polysaccharides—essentially extracted from micro- and macroalgae— it appears that they are often composed of rare or very unique residues. Anhydro-d/lgalactose is found only in carrageenans and agarans, and l-fucose and l-guluronic acid composing fucoidans and alginate, respectively, are very rare monosaccharides; rhamnose of ulvan or arabinopyranose of cladophoran also represents very unique residues. This short list of rare residues is probably just a taste of the carbohydrate diversity occurring in oceans. In addition, many marine polysaccharides are anionic, decorated by carboxylic function and/or sulfate ester groups. Sulfate groups, which are very common in marine polysaccharides but very rare in land organisms (e.g., glycosaminoglycan found in animals), could also be seen as a signature of marine polysaccharides. The structural diversity of marine polysaccharides in terms of monosaccharide sequences of the carbohydrate backbones as well as their decoration by organic or inorganic groups confers to this class of macromolecules a very wide range of functional properties. The well-known gelling properties of agars, carrageenans, and alginate have no equivalent and are widely exploited in food and cosmetic industries. Many biological properties of marine oligo- and polysaccharides have been identified and make them suitable to reach the cosmetic and agricultural markets. More recently, a new challenge such as the exploitation of marine biomass as source of sustainable energy to participate in the future replacement of fossil resources has emerged. Whatever the applications envisioned, the structure (e.g., composition, degree of polymerization) of the polysaccharides or the ix

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oligosaccharides must be well controlled. In this context, enzymes including glycoside hydrolases, polysaccharide lyases, sulfatases, for the most studied, are unavoidable tools to design structure of interest. Ultimately, the production of monosaccharides requires also set of enzymes. In this context, technology involving enzymatic degradation/modification of marine polysaccharides is continually in progress. This is in the context of active research in this field of marine glycoscience that this book, Enzymatic Technologies for Marine Polysaccharides, takes full place. The first section of the book, dedicated to the exploration of the marine polysaccharides diversity, is treated under various angles, including a description of the chemical complexity of these underexploited macromolecules and their current applications and new perspectives in foods, pharmaceuticals, cosmetics, and biomaterials offered by recent researches. Efficient valorization of the marine polysaccharide biomass requires a rigorous analysis of the polysaccharides structure and their biological properties. Therefore, the second section concerns the development of extraction techniques, improvement of methods aiming at the characterization of the structure and function of marine. The last section is articulated around enzymatic technologies from the discovery of novel enzymes, namely, glycoside hydrolases and polysaccharide lyases, to their production pipelines related to the fields of biorefinery, food industry, pharmaceutics, and other fine chemicals. William Helbert CERMAV, CNRS and Grenoble Alpes Université, Grenoble, France

Introduction The idea to prepare a book about the enzymatic technologies used in the processing of marine polysaccharides came after the publication of several review articles. During the preparation of these articles, I was getting more and more convinced that carbohydrate hydrolyzing enzymes of marine origin still appear today as an important topic after the old scientific attention for them, picked up from early results of research projects conducted since the 1960s, with a particular focus on the selectivity of these enzymes in their hydrolytic reactions. Often these biocatalysts have represented a crucial tool in the hand of chemists struggling with the complexity of macromolecular structures before the era of high-resolution NMR and MS facilities. The interdisciplinary nature of the current interest in marine polysaccharides and marine carbohydrate-hydrolyzing enzymes was also evident from the analysis above. A number of interesting original articles were found in active research fields related to bioprospecting and others, and cross-fertilization appeared to be very beneficial in that many disciplines are involved in these studies. The potential in bioactivity of marine polysaccharides is still considered underexploited. These molecules, including the derived oligosaccharides, are an extraordinary source of chemical diversity and literature highlights many applicative fields, including the food industry, cosmetics, biomedicine, agriculture, environmental protection, and wastewater management. The book was prepared with the idea to provide recent advances contributed by scientific expert leaders from academy and industry in that marine polysaccharides have important applications in various sectors of human interest. Sources and supply methodology are covered after depicting a general view on marine polysaccharides in Section 1. Contributions related to the complex structural determination and all modern methodologies to assess biological activities are included in Section 2, while Section 3 focuses on enzymatic processing and case studies in application fields. Marine originating enzymes for the hydrolysis and manipulation of polysaccharides are in the spotlight being the natural tools of their metabolism. All modern aspects of enzymatic technology are included in the last section for production of material with defined structures or as the core base of enzymatic processes in biotechnological industrial pipelines. The valorisation of marine biodiversity opens the book (Chapter 1) with emphasis on microorganisms offering the richest source of polysaccharides, particularly attractive as biomolecules with high-added value. The great structural diversity of marine microbial exopolysaccharides and their biological properties are discussed for future development in the pharmaceutical sector. This topic is also discussed in Chapter 10 of this section. Exopolysaccharides still represent a small fraction in the current market of polymers because of the production cost, although many developments have been made and some are still in progress for production and downstream processes. xi

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Introduction

Marine polysaccharides have interesting structural and functional properties, making them an established asset in commercial food additives. A complete picture under this angle of view is discussed in Chapter 2, specifying commercial exploitation of microalgae for their polysaccharides and other components in food applications. The great importance of fucoidan in this context is introduced here with a contribution (Chapter 3) dealing with manufacture, characterization, and uses of this molecule from macroalgae. A further chapter covers chemical modifications that are used to enhance the functional properties and applications of macroalgal polysaccharides (Chapter 4). Analysis of issues that exist in current processes is reported. A need to tackle problems with advanced chemical and biological tools would offer humankind sustainable solutions for next generations. Discussed is the whole asset of chemical and biological catalyst-mediated valorization process. The process of mild-acid hydrolysis employed for depolymerizing sulfated fucans is reviewed in Chapter 6. In particular new insights from recent results of experiments using nuclear magnetic resonance spectroscopy or computational simulations/ calculations, put the attention on the conformational and dynamical aspects of the molecule in solution. Turning to a basic science focus, an interesting report (Chapter 5) dealing with natural functions of marine exopolysaccharides is included here. The production of these polysaccharides and their role in protecting microbial organisms in marine environments are discussed as inspiration for potential application as, for example, biosorbents of heavy metals, biosurfactants for the bioremediation of oil, and moisturizer and skin protector in cosmetics. On the same basic focus science line, molecular aspects regarding the mucus of marine invertebrates are reported in Chapter 8. This knowledge may represent the basis for future investigations related to exploitation of this particular biomass as a source of bioactive compounds also exploitable in the pharmaceutical and biotechnological fields. Importance of chitin is discussed in a comprehensive perspective (Chapter 7), particularly for the biosynthetic and cellular processes involving β-chitin nanofiber formation. Key enzymes involved in diatom chitin metabolism, complementary to the overall theme of enzymatic technologies for marine polysaccharides are discussed in search for future avenues of research. Returning back to biorefinery concept a complete analysis (Chapter 9) is dedicated to applications of these polysaccharides and their monosaccharides for energy production. Unique polysaccharides and their production and processing from Laminaria sp., Kappaphycus sp., and Ulva sp. are discussed with emphasis on the fundamental role of the seaweed associated bacteria. The second section of the book collects contributions related to the efficiency of extraction techniques, the complexity of problems related to detailed structural determination and covers all modern methodologies related to biological activities of marine polysaccharides. Two overviews are offered from different perspectives. The first one, in Chapter 11, concluded that heterogeneity of marine polysaccharides in structural terms requires different extraction protocols characterized by different purity levels that still represents a challenge in that purity level is greatly influencing the type and the quality of biological effects. The second overview, in Chapter 13, is focused on the extraction and purification technologies with a broad discussion on the structure-function and chemical diversity of these molecules concluding with a description of the biological activities with emphasis on their applications in new fields such us plant biostimulants. In addition, a third overview (in Chapter 15) is focused more on the role of polysaccharides from marine environment in metabolic disorders depicting the current developments in the field of nutraceutical, cosmeceutical, and pharmacological applications. A particular aspect for fucoidan is reported in Chapter 12, where details on the use of this polymer for molecular

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diagnosis and targeted therapy of cardiovascular diseases are reported. Interestingly, in this section, the use of microwave has been investigated in Chapter 14; it summarizes practical microwave-assisted extraction and hydrolysis process of marine polysaccharides. Fundamental microwave heating mechanisms and dielectric properties of materials were also described to understand the mechanisms of extraction process under microwave irradiation. The last section of the book is dedicated to the core base of enzymatic processes in biotechnological industrial pipelines for production of material with defined structures. All modern aspects of enzymatic technology are included. In Chapter 16 the role of enzymatic technologies first in the extraction of native molecules and then in the production of marine bioactive oligosaccharides is reviewed. Progressively, Chapter 17 is a presentation of important microbial enzymes with potential use in modification of polysaccharides structures from native red, green, and brown algae, while in Chapter 18 details on molecular modification of sulfated polysaccharides of marine origin are investigated. The marine algae-degrading enzymes that emerged as potential tools for hydrolyzing the polysaccharides into oligosaccharides in food biotechnology and agriculture fields are the focus of Chapter 19, with special attention to alginate lyases, agarases, and carrageenases, but also including fucanolytic enzymes. Chapter 20 debates different enzymatic processes by specific enzymes acting on chitin and chitosan as well as documentation of potentials of enzymatic technologies for the prospective production of functional oligosaccharides/oligomers and monomers of commercial significance from these untapped marine polysaccharides. A specific contribution (Chapter 21) is dedicated to modification of the chemical structure of marine polysaccharides to produce GAG-mimetic compounds using enzymatic methods, and another (Chapter 22) is devoted to production of value-added materials from alginate using alginate lyases and others alginate-metabolic enzymes from alginolytic bacteria and marine gastropods. Antonio Trincone Istituto di Chimica Biomolecolare, CNR, Rome, Italy

Acknowledgments I would like to thank all colleagues acting as reviewers of the material published in this book. Without their experience and availability, the publication of this book would have been impossible: Oluyomi S. Adeyemi

Tetsushi Mori

Marina Basaglia

Barbara Mulloy

Giancarlo Cravotto

Thadathil Nidheesh

Alan Critchley

Nikolay Nifantiev

Mirjam Czjzek

Patrizia Pagliara

Pedro Carlos de Barros Fernandes

Angela Pennacchio

Concetta Gugliandolo

Annarita Poli

William Helbert

Harish Prashanth

Jun-ichi Kadokawa

Marguerite Rinaudo

Paola Laurienzo

Giuseppina Sandri

Vuyo Mavumengwana

Alexandra S. Silchenko

Osada Mitsumasa

Tiago Silva

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Editor Antonio Trincone has published more than 140 scientific papers (articles, reviews, and book contributions), which have appeared in leading international scientific journals. Since 1983 to the present, he has been at the Istituto di Chimica Biomolecolare belonging to Consiglio Nazionale delle Ricerche, in Naples, Italy, currently he is a Senior Researcher. He has been Professor of Organic Chemistry in charge for several years at the University of Salerno, Italy. His research activity has focused on the biomolecular aspects—many regarding stereochemistry and bioorganic chemistry—of enzymes as biocatalysts. His recent interests are in the biorefinery pipeline context using thermophilic and/or marine enzymes and in the biocatalyzed synthesis of bioactive compounds. Antonio has been the editor of a number of scientific books and various special issues in leading scientific journals, dedicated to marine biotechnology. He is Specialty Chief Editor of Marine Biotechnology (Frontiers in Marine Science), editing different research topics for this journal, and he is part of the Editorial Board of the Multidisciplinary Digital Publishing Institute (MDPI) journals Marine Drugs and Molecules. Most of his recent work is dedicated to the editorial process and review activities for scientific projects, published articles, and scientific positions.

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Contributors Velusamy Arumugam Department of Environmental Biotechnology School of Environmental Sciences Bharathidasan University Tiruchirappalli, India Dhruti Avlani Department of Pharmacy NSHM Knowledge Campus Kolkata, India Rathinam Ayyasamy Department of Animal Science Centre for Pheromone Technology Bharathidasan University Tiruchirappalli, India O. M. Babatunde Department of Chemical Sciences Biochemistry Unit Ondo State University of Science and Technology Okitipupa, Ondo State, Nigeria Francisco F. Bezerra Program of Glycobiology Institute of Medical Biochemistry Leopoldo de Meis and University Hospital Clementino Fraga Filho Federal University of Rio de Janeiro Rio de Janeiro, Brazil

Gopa Roy Biswas Department of Pharmacy NSHM Knowledge Campus Kolkata, India Carla C. C. R. de Carvalho Department of Bioengineering Institute for Bioengineering and Biosciences (iBB) Instituto Superior Técnico Universidade de Lisboa Lisbon, Portugal Frédéric Chaubet Galilée Institute, Paris 13 University Villetaneuse, France Cédric Chauvierre Inserm, U1148 LVTS, Paris Diderot University Paris 13 University Paris, France Lucas Chollet Galilée Institute, Paris 13 University, Villetaneuse and Algues & Mer, Kernigou Ouessant, France Anirban Roy Choudhury Biochemical Engineering Research and Process Development Centre (BERPDC) Institute of Microbial Technology (IMTECH) Council of Scientific and Industrial Research (CSIR) Chandigarh, India xix

xx Sylvia Colliec-Jouault Ifremer–Centre Atlantique EM3B Laboratory Nantes, France Christine Delbarre-Ladrat Ifremer–Centre Atlantique Nantes, France H. El Arroussi Moroccan Foundation for Advanced Science, Innovation and Research (MASCIR) Rabat, Morocco J. H. Fitton Marinova Pty Ltd Cambridge, Tasmania, Australia Satoshi Fujii Department of Chemical Science and Engineering School of Materials and Chemical Technology Tokyo Institute of Technology Tokyo, Japan and Department of Information and Communication Systems Engineering Okinawa National College of Technology Okinawa, Japan Sanjeev C. Ghadi Department of Biotechnology Goa University Goa, India Alexander Golberg Porter School of Environmental and Earth Sciences Tel Aviv University Tel Aviv, Israel Archunan Govindaraju Department of Animal Science Centre for Pheromone Technology Bharathidasan University Tiruchirappalli, India

Contributors Daniela de Borba Gurpilhares Faculdade de Farmácia Universidade Federal do Rio de Janeiro Rio de Janeiro, Brazil Masanori Hiraoka Usa Marine Biological Institute Kochi University Kochi, Japan O. Ibraheem Department of Biochemistry Federal University Oye-Ekiti Ekiti State, Nigeria Md. Imran Department of Biotechnology Goa University Goa, India Akira Inoue Laboratory of Marine Biotechnology and Microbiology Hokkaido University Hakodate, Hokkaido, Japan I. Meftah Kadmiri Moroccan Foundation for Advanced Science, Innovation and Research (MASCIR) Rabat, Morocco S. S. Karpiniec Marinova Pty Ltd Cambridge, Tasmania, Australia Anjula Katoch Biochemical Engineering Research and Process Development Centre (BERPDC) Institute of Microbial Technology (IMTECH) Council of Scientific and Industrial Research (CSIR) Chandigarh, India

Contributors Rajwinder Kaur Biochemical Engineering Research and Process Development Centre (BERPDC) Institute of Microbial Technology (IMTECH) Council of Scientific and Industrial Research (CSIR) Chandigarh, India Bhupender Kumar Biochemical Engineering Research and Process Development Centre (BERPDC) Institute of Microbial Technology (IMTECH) Council of Scientific and Industrial Research (CSIR) Chandigarh, India Didier Letourneur Inserm, U1148 LVTS, Paris Diderot University Paris 13 University Paris, France Murielle Maire Galilée Institute Paris 13 University Villetaneuse, France Sutapa Biswas Majee Department of Pharmacy NSHM Knowledge Campus Kolkata, India A. Mzibra Hassan II Institute of Agronomy and Veterinary Medicine (IAV) and Moroccan Foundation for Advanced Science, Innovation and Research (MASCIR) Rabat, Morocco Subhapradha Namasivayam Department of Medical Biotechnology Chettinad Academy of Research and Education Chennai, India

xxi Limin Ning College of Medicine and Life Sciences Nanjing University of Chinese Medicine Nanjing, China Ryuji Nishiyama Laboratory of Marine Biotechnology and Microbiology Hokkaido University Hakodate, Hokkaido, Japan Annamma A. Odaneth DBT-ICT Centre for Energy Biosciences Institute of Chemical Technology Mumbai, India Takao Ojima Laboratory of Marine Biotechnology and Microbiology Hokkaido University Hakodate, Hokkaido, Japan Ayumu Onda Research Laboratory of Hydrothermal Chemistry Kochi University Kochi, Japan A. Y. Park Marinova Pty Ltd Cambridge, Tasmania, Australia Adalberto Pessoa Jr. Departamento de Tecnologia Bioquímico-Farmacêutica Universidade de São Paulo São Paulo, Brazil Mark Polikovsky Porter School of Environmental and Earth Sciences Tel Aviv University Tel Aviv, Israel

xxii Vitor H. Pomin Program of Glycobiology Institute of Medical Biochemistry Leopoldo de Meis and University Hospital Clementino Fraga Filho Federal University of Rio de Janeiro Rio de Janeiro, Brazil and Department of Biomolecular Sciences and Research Institute of Pharmaceutical Sciences School of Pharmacy University of Mississippi University, Mississippi Karthik Ramachadran Department of Medical Biotechnology Chettinad Academy of Research and Education Chennai, India Valerie J. Rodrigues DBT-ICT Centre for Energy Biosciences Institute of Chemical Technology Mumbai, India Lydia Rolland Algues & Mer, Kernigou Ouessant, France Gregory L. Rorrer College of Engineering Oregon State University Corvallis, Oregon Ramachandran Saravanan Department of Medical Biotechnology Chettinad Academy of Research and Education Chennai, India Lara Durães Sette Departamento de Bioquímica e Microbiologia Instituto de Biociências Universidade Estadual Paulista Júlio de Mesquita Filho (UNESP) São Paulo, Brazil

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Contributors Shweta Singh Biochemical Engineering Research and Process Development Centre (BERPDC) Institute of Microbial Technology (IMTECH) Council of Scientific and Industrial Research (CSIR) Chandigarh, India Corinne Sinquin Ifremer–Centre Atlantique EM3B Laboratory Nantes, France Kulwinder Singh Sran Biochemical Engineering Research and Process Development Centre (BERPDC) Institute of Microbial Technology (IMTECH) Council of Scientific and Industrial Research (CSIR) Chandigarh, India Loredana Stabili Water Research Institute of the National Research Council S.S. Talassografico of Taranto Taranto, Italy D. N. Stringer Marinova Pty Ltd Cambridge, Tasmania, Australia Yun Sun College of Food Sciences and Light Industry Nanjing Tech University Nanjing, China Umamaheswari Sundaresan Department of Environmental Biotechnology School of Environmental Sciences Bharathidasan University Tiruchirappalli, India

25/03/19 4:54 PM

Contributors P. V. Suresh Department of Meat and Marine Sciences CSIR-Central Food Technological Research Institute Mysuru, India Shuntaro Tsubaki Department of Chemical Science and Engineering School of Materials and Chemical Technology Tokyo Institute of Technology Ookayama, Meguro Tokyo, Japan Tadaharu Ueda Department of Marine Resource Science Kochi University Kochi, Japan Manigandan Venkatesan Department of Medical Biotechnology Chettinad Academy of Research and Education Chennai, India Vazhiyil Venugopal Former Head, Seafood Technology Section Food Technology Division Bhabha Atomic Research Center Mumbai, India and Department of Food Science and Technology Kerala University of Fisheries and Ocean Sciences (KUFOS) Kochi, Kerala, India

xxiii Véronique Verrez-Bagnis Ifremer–Centre Atlantique EM3B Laboratory Nantes, France Yuji Wada Department of Chemical Science and Engineering School of Materials and Chemical Technology Tokyo Institute of Technology Ookayama, Meguro Tokyo, Japan Zhong Yao College of Food Sciences and Light Industry Nanjing Tech University Nanjing, China Benwei Zhu College of Food Sciences and Light Industry Nanjing Tech University Nanjing, China Agata Zykwinska Ifremer–Centre Atlantique EM3B Laboratory Nantes, France

section one

General view and sources of marine polysaccharides and oligosaccharides

chapter one

Marine biodiversity as a new source of promising polysaccharides Innovative polysaccharides emerging from the marine biodiversity Sylvia Colliec-Jouault, Corinne Sinquin, Agata Zykwinska, and Christine Delbarre-Ladrat Ifremer—EM3B laboratory, Centre Atlantique, Nantes, France

Contents 1.1 Diversity of marine ecosystems............................................................................................3 1.1.1 Deep-sea environments.............................................................................................4 1.1.2 Shallow submarine thermal springs........................................................................ 4 1.1.3 Tropical environments...............................................................................................4 1.1.4 Arctic and antarctic oceans....................................................................................... 5 1.2 Diversity of marine bioresources......................................................................................... 5 1.2.1 Marine macroresources............................................................................................. 5 1.2.2 Marine microresources..............................................................................................6 1.3 Diversity of marine polysaccharides and their biological activities............................... 6 1.3.1 Chitin/chitosan...........................................................................................................6 1.3.2 Galactans and carrageenans.....................................................................................6 1.3.3 Alginates......................................................................................................................7 1.3.4 Fucoidans.....................................................................................................................8 1.3.5 Fungal polysaccharides..............................................................................................8 1.3.6 Microalgal polysaccharides....................................................................................... 9 1.3.7 Bacterial polysaccharides........................................................................................ 10 Summary......................................................................................................................................... 16 References........................................................................................................................................ 17

1.1  Diversity of marine ecosystems Oceans and seas offer enormous possibilities for the discovery of new species that could be considered as bioactive compound providers and even producers. Oceans and seas support adapted life forms that evolve across a wide range of environmental conditions and habitats. The habitat heterogeneity found in the marine environment results in the species richness. Marine ecosystems can depend on photosynthetic or chemosynthetic primary production. The latter is thus independent of sunlight. Consequently, marine ecosystems can be extreme environments in which species survive, evolve, and adapt to unusual physical and chemical conditions with some highly unique biological interactions. 3

4

Enzymatic Technologies for Marine Polysaccharides

The progress in exploration tools [submersibles, remotely operated vehicles (ROVs), landers] has permitted access to remote habitats previously considered as biological deserts and now recognized as oasis or rain forests (Gallo et al. 2015).

1.1.1  Deep-sea environments The deep sea (>200 m deep), which is governed by chemosynthesis, is a mosaic of habitats in which communities of organisms proliferate through local production of organic matters. Symbiotic activity is observed with microorganisms that use the chemical compounds brought by fluids to produce organic matters consumed by other species. An abundant fauna dominated by a small number of species colonizes these abyssal plains where unique extremal ecosystems are located such as active hydrothermal vents, seeps, and highly diverse landscapes such as ridges, deep-water coral reefs, mud volcanoes, seamounts, and canyons (Sarrazin et al. 2014; Cuvelier et al. 2017). A high diversity of microorganisms has been discovered living in close association with higher organisms (worms, shrimps, mussels, invertebrates) as well as rocks, sediments, diluted vent fluids, or seawater columns. They are often new species of bacterial groups called extremophiles. The main groups are proteobacteria (Gram-negative) and archaea and Gram-positive bacteria with hyperthermophilic, mesophilic, and psychrophilic members and also anaerobes, aerobes, chemolithoautotrophs, chemoautotrophs, and heterotrophs. But the presence of viruses and fungi has also been described (Pettit 2011; Deming 1998).

1.1.2  Shallow submarine thermal springs Shallow marine hydrothermal vents (found at depths below 50 m) in Italy (Eolian Islands, Cape Palinuro) and in Japan (Yaeyama Archipelago) host thermophilic and mesophilic bacteria. This marine ecosystem, resembling the deep-sea hydrothermal vents, hosts micro- and macrofauna tolerant to a wide range of pH, temperature, and salinity values. Thermophilic heterotrophs and thermophilic sulfur oxidizers have been detected (Nunoura et al. 2013). Thermophilic and halophilic strains with the genera Bacillus, Vibrio, Citrobacter, and Escherichia have been described (Gugliandolo et al. 2003; Canganella et al. 2002).

1.1.3  Tropical environments The Pacific Ocean, the largest oceanic division on Earth, is a great reservoir for the discovery of new microorganisms. In New Caledonia, marine coastal biotopes offer a high endemic biodiversity providing new bacterial strains. A bank of marine bacteria has been constituted from various sites along the western coast of New Caledonia, Pseudoalteromonas strains, and a new member of the Vibrio genus has been identified (Dufourcq et al. 2014; Chalkiadakis et al. 2013). Microbial mats (kopara) present in shallow brackish and hyposalted ponds in French Polynesian atolls are composed of phototrophic and chemotrophic prokaryotes. Kopara built up in stratified layers is dominated by a large diversity of cyanobacteria, sulfate reducers, and other heterotrophic bacteria forming gelatinous deposits (Che et al. 2001). Different sites of the coastal regions of Goa in India have been explored to collect marine bacterial strains, and a Vibrio strain identified as Vibrio furnissii has been characterized (Bramhachari et al. 2007). Other types of mat ecosystems are present around the globe; they are highly dependent on the presence of light. According to their location and

Chapter one:  Marine biodiversity as a new source of promising polysaccharides

5

physicochemical features, they host their own microbial diversity, dominated by either cyanobacteria or proteobacteria, Chloroflexi, and Firmicutes (Prieto-Barajas et al. 2018).

1.1.4  Arctic and antarctic oceans In polar regions, sea ice is the major component and provides a home to a unique community dominated by microorganisms. In both arctic and antarctic oceans, bacteria are in close association with phytoplankton and are in abundance in the bottom layers of the ice. Bacteria are often attached to microalgal cells and are also associated with particulate material rich in exopolymeric substances giving transparent exopolymer particles. Psychrophilic bacteria have been characterized; they are able to grow from 0°C to 25°C; low-temperature extremophiles can produce cold-adapted enzymes as well as exopolysaccharides. The described genera found in cold environment are mainly Octadecabacter, Glaciecola, Psychrobacter, Marinobacter, Pseudoalteromonas, Shewanella, Polaribacter, and Flavobacterium (Cavicchioli et al. 2002; Brinkmeyer et al. 2003; Nichols Mancuso et al. 2005).

1.2  Diversity of marine bioresources Traditionally, the marine macroresource (e.g., mammals, invertebrates, macroalgae) has been exploited mainly to produce polysaccharides. Nowadays, advances in molecular techniques and microbial research are creating significant opportunities to exploit the biological resource of the marine microorganisms (e.g., microalgae, cyanobacteria, bacteria).

1.2.1  Marine macroresources A wide variety of polysaccharides can be extracted from marine animals and algae. Chitin is a natural polysaccharide and second most abundant polymer after cellulose produced annually in the world. The exoskeleton of marine organisms, mainly crabs and shrimps, is widely used to produce marine chitin (Laurienzo 2010). In addition, marine macroresource is an alternative source to obtain glycosaminoglycans or GAGs (e.g., chondroitin sulfate, dermatan sulfate and hyaluronan) traditionally extracted from mammalian tissues. Chondroitin sulfate and hyaluronan can be extracted from fish wastes and by-products such as cartilage or skin and fish eyes, respectively (Nandini et al. 2005; Alonso et al. 2010). Heparin, dermatan sulfates, and even fucosylated chondroitin sulfate can be isolated from ascidian tissues (chordate and tunicate) (Pavao 2014). The cell walls of marine macroalgae contain sulfated or unsulfated polysaccharides that can be considered as a rich source of heparin-like or GAG-like entities. Polysaccharides from the three major divisions of marine macroalgae (Rhodophyta, Phaeophyta, and Chlorophyta) have been largely studied to explore their potential as a cheap and safe source of new generation of heparinoids or GAG-like entities. Among the numerous algal polysaccharides, carrageenans from red seaweeds (Rhodophyta) and fucoidans from brown seaweeds (Phaeophyta) share numerous properties with GAGs (Tseng 2001; Wijesinghe and Jeon 2012). Despite the great interest in these compounds, exploitation of the polysaccharides from the macroresource can be limited; difficult physical access to certain habitats, low reproducibility due mainly to the seasonal variations, and dependence on ecological (e.g. oil spills, heavy metals, germs, viruses) and political hazards are major concerns.

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Enzymatic Technologies for Marine Polysaccharides

1.2.2  Marine microresources Microorganisms, such as fungi, microalgae, cyanobacteria, and bacteria, have many advantages over the macroresource. They can grow in laboratory under fully controlled culture conditions compatible with current safety standards (traceability, containment), different molecules can be produced by the same strain (reproducibility), growth conditions can be controlled to optimize production, and so on. Production is much more rapid (a few days for microalgae and less than 60 hours for bacteria) and is very profitable (completely automatable, high-capacity culture systems, e.g., photobioreactors and fermenters, already exist). Finally, both extraction and purification of molecules are often simpler than those from higher plants or animal organs. In the last 20 years or so, several collections of marine bacteria, microalgae, and fungi have been established in Europe and worldwide for research and development (R&D) by optimizing the production of biomolecules, particularly of enzymes and polysaccharides. Thanks to new molecular biology tools and bioinformatics, the manipulation (cloning) and modification (site-directed mutagenesis) of genetic material are possible, with the aim of creating or modifying protein features to optimize the biosynthesis or the activity of the molecule of interest. Thus, the discovery of new, especially bioactive, compounds from these “micromarine plants” is expanding (Raja et al. 2008; Rehm 2010; Schmid et al. 2015).

1.3 Diversity of marine polysaccharides and their biological activities 1.3.1 Chitin/chitosan Chitin is produced as insoluble crystalline microfibers that are structural components of either the arthropod exoskeleton or the cell walls of fungi and yeast. The main source of commercially available chitin comprises crabs and shrimps. Chitin is a linear polysaccharide composed of (1→4)-N-acetyl-β-d-glucosamine (GlcNAc) residues. Partial deacetylation of chitin under alkaline conditions or by enzymatic hydrolysis in the presence of a deacetylase leads to its soluble derivative, namely, chitosan. Because its structural features are close to those of GAGs, a family of linear anionic polysaccharides present in the extracellular matrix and the on cell surface of animal tissues, chitosan is widely explored in biomedical applications for skin, bone, and cartilage engineering (Suh and Matthew 2000; Di Martino et al. 2005; Ahsan et al. 2018). To improve its biological properties and in order to form polyelectrolyte complexes, positively charged chitosan is frequently associated with negatively charged polysaccharides, such as alginate or GAGs (e.g., heparin, chondroitin sulfate, hyaluronic acid) (Ahsan et al. 2018). Chitosan-based biomaterials structured at micro- and nanoscales have also been extensively developed for drug, protein, and gene delivery (Bugamelli et al. 1998; Erbacher et al. 1998; Koppolu et al. 2015).

1.3.2  Galactans and carrageenans Agars and carrageenans are sulfated galactan polysaccharides found in the cell matrix of Rhodophyta or red algae. Agars are extracted from Gelidium, Gracilaria, Pterocladia, and Gracilariopsis species. Their structures are composed of a backbone consisting of agarose (70%), a neutral galactan, and agaropectin, a highly charged galactan. Approximately 80% of produced agars are used in the food industry as thickening or gelling agents, and 10–20% of remaining agars are used in the pharmaceutical and biotechnological industries.

Chapter one:  Marine biodiversity as a new source of promising polysaccharides

7

Because of their low sulfate content (2–5%), their biological activities have been studied less extensively. Carrageenans are polysaccharides extracted mainly from Chondrus, Gigartina, Euchema, Hypnea, and Kappaphycus red seaweed species. Carrageenans are hydrophilic sulfated linear galactans composed of repeating units of (1→3)-α- and (1→4)-β-galactans, substituted by one (κ-carrageenan), two (ι-carrageenan), or three (λ-a-carrageenan) sulfated groups per disaccharide unit. They have a high molecular weight (HMW) and a high sulfate content, ranging between 20 and 40% w/w. Because of their high sulfate content, λ-carrageenans present GAG-like properties and more precisely heparin-like properties. As heparin, they can prolong the coagulation time and consequently have in vitro anticoagulant activity, but concentrations higher than those of heparin are required to observe the same prolongation in clotting assays, 100 µg/mL and 2 µg/mL, respectively (Carlucci et al. 1997; Barabanova et al. 2008). A lowmolecular-weight (LMW) derivative obtained from a sulfated galactan of Botryocladia occidentalis was as active as heparin in a venous thrombosis model performed in rats but showed a weak antithrombotic activity in arterial thrombosis model also performed in rats (Melo and Mourao et al. 2008; Fonseca et al. 2008). A study performed with a LMW ι-carrageenan derivative showed an inhibitory effect on both human osteosarcoma cell line proliferation by induction of apoptosis and tumor growth in an established xenograft tumor model (Jin et al. 2013). Another study reported tumor-inhibiting activities of different derivatives obtained from a λ-carrageenan from Chondrus ocellatus. These derivatives can manifest different degrees of antitumor and immunomodulation activities according to their molecular weights (Zhou et al. 2004). To summarize, the described antitumor effects of carrageenans are closely linked to their structure and physicochemical properties (Khotimchenko 2010). For more than 30 years, the antiviral activities of carrageenans have been studied against several different viruses, and their efficacy has been found to be very broad (for both enveloped and nonenveloped viruses) and closely related to both the molecular weight and the degree of sulfation of the carrageenans (Wang et al. 2012; Kalitnik et al. 2013; Bouhlal et al. 2011).

1.3.3 Alginates Cell walls of brown algae (Phaeophyceae), including Laminaria hyperborea, Laminaria digitata, Laminaria japonica, Ascophyllum nodosum, and Macrocystis pyrifera, are particularly rich in alginate. Although commercially available alginate is of algal origin, another potential source of this macromolecule may comprise bacteria, such as Azotobacter vinelandii and Pseudomonas spp., which produce alginate through a fermentation process (Clementi 1997; Rehm and Valla 1997). Alginate is a linear anionic heteropolysaccharide composed of (1→4)-α-l-guluronic acid blocks (GG) and (1→4)-β-d-mannuronic acid blocks (MM), which are intercalated by heterogeneous MG blocks. Alginates extracted from different sources differ in G and M residue contents according to the length of each block; this leads to important heterogeneity in chemical composition, directly affecting the physicochemical properties of alginate. In its monovalent salt form, especially sodium form, alginate is perfectly soluble in water, while in the presence of divalent cations (e.g. Cu2+, Zn2+, Ca2+), alginate is able to gel. The divalent cations induce chain–chain association between GG blocks, which results in junction zone formation, as proposed by the “egg-box model” (Tombs and Harding 1998). Because of its thickening and in particularly gelling properties in the presence of divalent ions, mainly Ca2+, alginate is widely used in the food industry and for biomedical applications (Sabra et al. 2001). In the biomedical domain, alginate is explored for delivery of LMW drugs and proteins (Tonnesen and Karlsen 2002;

8

Enzymatic Technologies for Marine Polysaccharides

Lee and Mooney 2012). Alginate hydrogels also provide a hydrophilic environment suitable for cell encapsulation, and such cell delivery systems are widely explored in tissue engineering (Lee and Mooney 2012). Ca2+ ions liberated from alginate in the calcium form activate platelet aggregation and reduce coagulation time (Segal et al. 1998). Thus, alginate in its calcium form is used for wound dressing elaboration, and several alginate-based dressings are already available, including CoalganTM (Brothier Laboratories), AlgiSite MTM (Smith & Nephew), and Comfeel PlusTM (Coloplast).

1.3.4 Fucoidans Fucoidans, or sulfated fucans, are fucose-containing sulfated polysaccharides found in brown algal cell wall tissue (Phaeophyceae). They are a family of compounds including fucoidin, fucoidan, ascophyllan, sargassum, and glucuronoxylofucan. They are present mainly in Fucales and Laminariales but also in several orders such as Chordariales and Ectocarpales. Fucoidans are composed mainly of sulfated fucose units, but they present a structural diversity in association with the algal species. They can be highly branched with a large proportion of both α(1→3) and α(1→4) linkages, more or less sulfated with a low content of uronic acids and xylose, rhamnose, and mannose. The biological activity of fucoidans depends on their structure, their degree of sulfation, and also their molecular weight dispersion (Berteau and Mulloy 2003; Deniaud-Bouet et al. 2017). Their anticoagulant activity is the first described biological property using partially purified extracts presenting both HMW and high polydispersity. The preparation of LMW derivatives has highlighted the mechanism of action of fucoidan in blood coagulation. The LMW fucoidans exert their anticoagulant activity by enhancing thrombin inhibition via its natural inhibitors (Nishino et al. 1991; Church et al. 1989; Colliec-Jouault et al. 1991). The antithrombotic activity of fucoidans has been widely reported; they are efficient in preventing venous thrombosis as well as arterial thrombosis with a hemorrhagic risk lower than that of LMW heparin (Colliec-Jouault et al. 2003; Durand et al. 2003). Depending on their molecular weights, fucoidans can inhibit or enhance angiogenesis induced by endothelial cells (Boisson-Vidal et al. 2007; Soeda et al. 2000). Besides their beneficial activity on thrombosis and vascular biology, fucoidans act on the inflammation and immune systems. They can inhibit some processes involved in tissue breakdown and inflammation such as induction of matrix metalloproteinases by inflammatory cytokines and complement cascade and can promote in vitro proliferation of various cells by potentiating heparin-binding growth factor activities. Fucoidans, as potent modulators of connective tissue proteolysis, are effective compounds for wound repair and tissue engineering (O’Leary et al. 2004; Senni et al. 2006, 2011). In several in vitro tumor cell lines and in vivo tumor models, antiproliferative, apoptotic, antimetastatic, and antineoplastic effects of different fucoidans have been described. Different modes of action of fucoidans can be observed: induction of apoptosis, stimulation of immune cells, antiadhesive properties blocking both adhesion and implantation of tumor cells, inhibition of tumor cell proliferation, and/or differentiation (de Jesus Raposo et al. 2015; Zhang et al. 2013; Foley et al. 2011; Liu et al. 2005; Nakano et al. 2012). To conclude, fucoidans present a great potential for medical applications (Wijesinghe and Jeon 2012).

1.3.5  Fungal polysaccharides Marine fungi are an inexhaustible reserve of bioactive secondary metabolites. Their biodiversity is related to the multitude of marine habitats, spreading out in coastal ecosystems as well as in deep-sea hydrothermal environments, in which they are found associated

Chapter one:  Marine biodiversity as a new source of promising polysaccharides

9

with marine macroorganisms such as sponges, corals, algae, and mangrove. Marine fungi are symbiotic and epiphytic eukaryotic microorganisms, and until now have been inadequately explored as microbial exopolysaccharide (EPS) producers compared to the marine prokaryotes. The fungal EPS are produced by fermentation; they can be either secreted in culture broth or tightly attached to the cell surface (Freitas et al. 2017). Few studies have been performed on the biological activity of the fungal EPS; members of the fungal genera Alternaria, Aspergillus, Fusarium, and Penicillium have been studied mainly for the structural characterization and antioxidant properties of their EPS. They produce highly branched neutral polysaccharides composed of glucose, galactose, and mannose units. The EPS secreted by the coral-associated fungus Aspergillus versicolor is a mannoglucan with a sidechain composed of mannose trisaccharides branched for every eight sugar units in the backbone. Its molecular weight is approximately 7,000 g/mol, and it possesses antioxidant properties (Chen et al. 2012). Two other fungal EPS are described for their in vitro antioxidant activities: mangrove-associated fungi Aspergillus sp. Y16 and Fusarium oxysporum. Aspergillus sp. Y16 fungus isolated from a leaf produces a galactomannan with a sidechain of specific galactofuranose units; its molecular weight is approximately 15,000 g/mol (Chen et al. 2011). The EPS of Fusarium oxysporum is a galactofuranose-containing mannoglucogalactan with a molecular weight of approximately 60,000 g/mol (Chen et al. 2015). A polysaccharide has been isolated from the marine filamentous fungus Phoma herbarum; it is a HMW polysaccharide (>2 × 106 g/mol) composed mainly of glucose with a trace amount of glucuronic acid. After a chemical sulfation, its antioxidant properties have been observed to increase (Yang et al. 2005).

1.3.6  Microalgal polysaccharides The world of microalgae remains to be explored. The estimated number of species ranges from 40,000 to 200,000 (Norton et al. 1996), and microalgae could contribute to 50% of total world photosynthetic activity (Day et al. 1999). Microalgae colonize all regions ranging from the polar ices to the deserts. They adapt to extreme environments, survive in saltwater marshes, in acid media, and even under very weak irradiance conditions. Their biodiversity, together with an exceptional adaptability, is matched by a proportional richness in molecular diversity. Microalgae can be cultured in photobioreactors with different designs and contribute to the rise of more biotechnological approaches. In fact, microalgae appear to be very easy to manipulate. Their production is an emergent market; the main exploited species are Arthrospira platensis, Aphanizomenon sp., Dunaliella salina, Chlorella vulgaris, and Porphyridium cruentum. The microalgal polysaccharides can be attached to the cell wall or secreted; they are EPS, often with a complex structure composed of several highly substituted monosaccharides. Some are homopolymers of glucose and galactose, but the majority are anionic heteropolymers with a high amount of acidic sugars and sulfate groups (Freitas et al. 2017). Microalgae constitute a still unexplored source of potential drugs with the exception of spirulan from the cyanophyte Arthrospira platensis, a sulfated polysaccharide composed of rhamnose, fucose, and glucose for which heparin-like properties have been described and consequently considered as “heparinoid” (Hayashi et al. 1996; Lee et al. 2000; Hayakawa et al. 2000). However, some other species of microalgae distributed in marine habitats (e.g., Amphora costrata, Coscinodscus nobilis, Aphanocapsa, Glocothece) contain sulfated polysaccharides potentially exploitable as drugs either in their natural form or on depolymerization and/or chemical derivatization (de Jesus Raposo et al. 2015; Raposo et al. 2013). The sulfated EPS extracted from the diatom Phaeodactylum tricornutum and the chlorophyte Chlorella stigmatophora have

10

Enzymatic Technologies for Marine Polysaccharides

both anti-inflammatory and immunomodulatory activities. Both EPS have sulfate and uronic acid percentages of around 10% w/w and 5% w/w, respectively (Guzman et al. 2003). The rhodophytes Porphyridium sp., P. cruentum, P. purpureum, and P. marinum produced a sulfated EPS composed mainly of xylose and galactose that has antiviral properties against different groups of viruses such as hepatitis B virus, herpes simplex and viruses, retroviruses (Huleihel et al. 2001; Huleihel and Arad 2001), and more recently its anti-inflammatory effect has been described in vitro (Levy-Ontman et al. 2017).

1.3.7  Bacterial polysaccharides Polysaccharides from marine prokaryotes offer a source of safe, biocompatible, biodegradable, and valuable renewable products with specific biological functions emphasized by a significant structural diversity (Table 1.1). Major recent advances in deep-sea biotechnology have allowed investigators to isolate novel microorganisms. Several EPS-producing marine strains have been studied, which led to the discovery and isolation of novel macromolecules (http://www.esf.org/publications/marine-sciences.html) (Deming 1998; Guezennec 2002; Laurienzo 2010; Finore et al. 2014). Marine bacteria from deep-sea hydrothermal vent environments, belonging to three main genera (Vibrio, Alteromonas, and Pseudoalteromonas), have demonstrated their ability to produce unusual extracellular polymers in an aerobic carbohydrate-supplemented medium (Figure 1.1).

(A)

OH HO

COONa O

O O

O

NHAc

O

HO

OH

COONa O O

HO

O HO

(B)

OH O

Glc

AcNH

OH OH O

O

n

Glc

OH O

HO HO

OH

OH

O HO

Gal

HO OH O

O HO O HO OH HO

HO HO

O OH

NaOOC

GlcA

NaOOC

O

O OH

OH GlcA O OH

O COONa O

GalA

O NaO3SO

O

Gal

OH O

HO OH

O OH Glc

n

Glc

Figure 1.1 A tetrasaccharidic repeating unit of HE800 EPS as described in 1999 by Rougeaux and coworkers (Rougeaux et al. 1999). The HE800 EPS repeating unit comprises the hyaluronan disaccharide (A). A nonasaccharidic repeating unit of GY785 EPS as described in 2004 by Roger and coworkers (Roger et al. 2004). The repeating unit is rich in uronic acid residues and bears one sulfate group (B).

Microorganism

Origin

Osidic composition (molar ratio) or repeating unit

References

Alteromonas infernus

Sulfated and acidic branched heteropolysaccharide; a nonasaccharide Diluted vent fluid repeating unit among a dense population of Riftia [SO3Na] pachyptila, Guaymas ↓ basin, Gulf of California 2

Raguenes et al. 1997b; Roger et al. 2004

Diluted hydrothermal fluid, North Fiji basin

Raguenes et al. 1996; Rougeaux et al. 1998

[→4)–β–Glcp–(1→4)–α–GalpA–(1→4)–α–Galp–(1→] 3 ↑ 1 β–Glcp–(1→6)–α–Galp–(1→4)–β–GlcpA–(1→4)–β–GlcpA 2 3 ↑ ↑ 1 1 α–Glcp α–Glcp

Alteromonas macleodii subsp. fijiensis

A hexasaccharide repeating unit →4)–β–D–Glcp–(1→4)–α–D–GalpA–(1→4)–α–D–Galp–(1→ 3 ↑ 1 α–D–GlcpA 3 ↑ 1 β–D–GlcpA 4 ↑ 1 4,6–Pyr–β–D–Manp

Alteromonas macleodii Hydrothermal vent Sulfated and acidic branched heteropolysaccharide; repeating units subsp. fijiensis biovar polychaete annelid, East from 16 to 18 monosaccharides (7 types) deepsane Pacific Rise

Cambon-Bonavita et al. 2002; Le Costaouec et al. 2012

Chapter one:  Marine biodiversity as a new source of promising polysaccharides

Table 1.1  Structural diversity of bacterial polysaccharides isolated from various marine ecosystems

(Continued)

11

12

BK-TandF-9781138103078_TEXT_TRINCONE-181207-Chp01.indd 12

Table 1.1 (Continued)  Structural diversity of bacterial polysaccharides isolated from various marine ecosystems Origin

Osidic composition (molar ratio) or repeating unit

References

Alteromonas sp. strain 1644

Hydrothermal vent polychaete annelid, East Pacific Rise Shallow marine hydrothermal vent, Vulcano Island, Italy Shallow marine hydrothermal vent, Ischia Island, Italy

Glc/Gal/GlcA/GalA et 3-O-[(R)-1-carboxyethyl]-d-GlcA

Bozzi et al. 1996

A tetrasaccharide repeating unit: Man/Glc (1/0.2)

Maugeri et al. 2002; Poli et al. 2010

2 sulfated EPS: EPS-1: α-Man and β-Glc (1.0/0.7) EPS-2: α-Man and pyruvic acid 2 sulfated acidic EPS

Manca et al. 1996

Poli et al. 2010

Glc/Gal/Fuc/Fru (1/0.07/0.04/0.02)

Kambourova et al. 2009

Man/Glc/Gal/ManN

Nicolaus et al. 2002

Gal/Man/GlcN/Ara (1.0/0.8/0.4/0.2)

Poli et al. 2010

Glc/Gal/Rib/Xyl (0.68/1.0/trace/trace)

Lee et al. 2001

Glc/Fru/GlcN/GalN (1/0.7/0.3/0.2)

Poli et al. 2007

Man/Glc and Rha (trace)

Llamas et al. 2012

Glc/Man/GalA

Mata et al. 2006

Glc/GlcA/Man/Fuc/Gal/Rha (1/0.3/0.25/0.2/0.1)

Amjres et al. 2015

Bacillus licheniformis

Bacillus thermoantarcticus Bacillus thermodenitrificans Geobacillus tepidamans V264 Geobacillus sp. 4001

Geobacillus sp. 4004

Hahella chejuensis Halomonas alkaliantarctica Halomonas almeriensis Halomonas anticariensis Halomonas stenophila

Shallow marine hydrothermal vent, Vulcano Island, Italy Geyser, Bulgary Shallow marine hydrothermal vent, Vulcano Island, Italy Shallow marine hydrothermal vent, Ischia Island, Italy Sediment, Cheju Island, South Korea Salt sediments, salt lake, Cape Russell, Antarctica Salt marsh, Almeria, Spain Salt marsh, Malaga, Spain Saline wetland, Brikcha, Morocco

27/03/19 4:03 PM

(Continued)

Enzymatic Technologies for Marine Polysaccharides

Microorganism

Microorganism

Origin

Osidic composition (molar ratio) or repeating unit

References

Halomonas ventosae

Salt marsh, Malaga, Spain Spanish hypersaline water, Malaga, Spain Spanish hypersaline water, Murcia, Spain Southern Ocean

Glc/Man/Gal

Mata et al. 2006

Glc/Man/Gal, anionic polysaccharide

Man/GlcA/GalNAc/Glc/GlcNAc/Ara/Gal/GalA/Xyl/Rha (48/10/10/9/8/6/4/2/2/1) Sulfated polysaccharide Gal/Glc/Rha/Fuc/GalA

Martínez-Cánovas et al. 2004 Martínez-Cánovas et al. 2004 Nichols Mancuso et al. 2005 Raguenes et al. 2004

Seawater, Marseille, France

Sulfated polysaccharide; a disaccharide repeating unit

Komandrova et al. 1998

Deep-sea hydrothermal vent, East Pacific Rise

Sulfated polysaccharide; an octasaccharide repeating units

Iodomarina fontislapidosi Iodomarina ramblicola Olleya marilimosa CAM030 Paracoccus zeaxantificiens subsp. payriae Pseudoalteromonas marinoglutinosa KMM232

Pseudoalteromonas sp. HYD721

Microbial mats, French Polynesia

Glc/Man/Gal, anionic polysaccharide

→3)–β–D–Manp–(1→4)–α–L–Rhap–(1→ 2  SO3H

Rougeaux et al. 1999a

→4)–β–D–Manp–(1→4)–β–D–Glcp–(1→4)–α–D–Galp–(1→4)–β–D–Glcp–(1→ 2 3 ↑ ↑ 1 1 α–L–Rhap β–D–Galp 3 ↑ 1 β–D–GlcpA 4 ↑ 1 [SO3H]→3–β–D–Manp

13

(Continued)

Chapter one:  Marine biodiversity as a new source of promising polysaccharides

Table 1.1 (Continued)  Structural diversity of bacterial polysaccharides isolated from various marine ecosystems

Origin

Osidic composition (molar ratio) or repeating unit

References

Pseudoalteromonas ruthenica SBT033 Pseudoalteromonas sp. strain CAM025 Pseudoalteromonas sp. strain CAM036 Pseudoalteromonas sp. strain SM9913 Shewanella colwelliana

Coastal regions, India

Man/Glc/Gal/Xyl, acidic sugars

Saravanan et al. 2008

Seawater in Southern Ocean Seawater in Southern Ocean Deep-sea sediment, Yellow Sea, China Eastern oyster Crassostrea virginica — Marine fouling material, Bengal Deep-sea hydrothermal vent, East Pacific Rise Coastal regions, India Tropical marine water

Acetyl, sulfate, Glc/GalA/Rha/Gal (1/0.5/0.1/0.08) Acetyl, succinyl, sulfate GalA/Glc/Man/GalNAc/Ara (1/0.8/0.84/0.36/0.13) Glc/t-Ara/t-Glc/t-Gal/Xyl/Glc/Glc (6.2/1.1/1.1/.03/0.4/0.5/0.5)

Nichols Mancuso et al. 2004 Nichols Mancuso et al. 2004 Qin et al. 2007

—

Abu et al. 1994

Glc/Rib/Man (1/0.05/0.02) Glc/AraN/RibN/Xyl

Rinker et al. 2000 Muralidharan et al. 2003 Rougeaux et al. 1999b

Thermotoga maritima Vibrio alginolyticus Vibrio diabolicus Vibrio furnissii Vibrio harveyi

[-->3)-β-GlcNAc-(1-->4)-β-GlcA-(1-->4)-β-GlcA-(1-->4)-αGalNAc(1-->] Neutral sugars (Glc/Gal) and acidic sugars Neutral sugars, acidic sugars, sulfate

Bramhachari et al. 2007 Bramhachari et al. 2006

Abbreviations: A = uronic acid; Ac = acetyl; Ara = arabinose; Fru = fructose; Fuc = fucose; Gal = galactose; Glc = glucose; Gro = glycerophosphate; Man = mannose; N = amine; Rha = rhamnose; Rib = ribose; S = sulfur; SO3 = sulfate; t = terminal; Xyl = xylose.

Enzymatic Technologies for Marine Polysaccharides

Microorganism

14

Table 1.1 (Continued)  Structural diversity of bacterial polysaccharides isolated from various marine ecosystems

Chapter one:  Marine biodiversity as a new source of promising polysaccharides

15

In particular, a very interesting new EPS-producing Vibrio has been discovered and named Vibrio diabolicus (Raguenes et al. 1997a). It is the first species of Vibrio to be isolated from an active deep-sea hydrothermal vent sample, and it can produce a very innovative EPS presenting a chemical resemblance to hyaluronic acid (HA). The structure of this EPS, named HE800 EPS, consists of a linear tetrasaccharide repeating unit: two glucuronic acid residues, one N-acetylated glucosamine residue, and one N-acetylated galactosamine residue. It is very rare to find hexosamine sugars in bacterial EPS; furthermore, it is a HMW EPS (>106 g/mol) (Rougeaux et al. 1999b). Another new bacterium was also isolated in deep-sea sediments of the Guaymas basin (Gulf of California); it is a deepsea aerobic mesophilic heterotrophic bacterium described as a new species of the genus Alteromonas, and the proposed name is Alteromonas infernus (Raguenes et al. 1997b). This bacterium can produce a water-soluble EPS, named GY785 EPS, which is a branched heteropolysaccharide whose repeating unit is a nonasaccharide: four glucose residues, two galactose residues, two glucuronic acids, and one galacturonic acid bearing one sulfate group at C2 position. The GY785 EPS is naturally slightly sulfated with a sulfur content of 3% w/w (Roger et al. 2004). The production yield of marine EPS is usually around 1 g/L (Decho 1990), and is sometimes close to 5 g/L (Raguenes et al. 1997b). Industrial development is conceivable for a production yield of approximately 10 g/L. Production costs are driven by the yield of polysaccharide, the amount and cost of carbon source, and the downstream processes needed for molecule separation. Therefore, studies on production optimization are needed even if some bacterial strains are naturally able to produce EPS at a high yield (e.g., ≥50 g/L) such as Agrobacterium sp. (curdlan), Xanthomonas campestris (xanthan), Zymomonas mobilis (levan), Alcaligenes faecalis (curdlan), and Bacillus sp. (levan) (Donot et al. 2012). Factors such as medium composition (e.g., carbon and nitrogen sources) and fermentation conditions (e.g., temperature, pH, aeration) are the variables most frequently employed to optimize the fermentation process. The nutritional conditions can also affect the production yield as well as the osidic composition of the EPS in some cases (Decho 1990; Finore et al. 2014). After fermentation, the downstream process for the recovery of EPS requires classical steps such as centrifugation to remove cells from the culture broth without lysis, and EPS isolation, extraction, or purification steps. After isolation, the polysaccharide is freeze-dried for a better conservation. Polysaccharides are highly hydrophilic, due to the presence of hydroxyl and carboxyl groups, especially when they are polyanionic, a widespread feature in marine environment, or more scarcely when they bear sulfate groups; therefore, they always conserve a content of water (DeAngelis 2012). With their polyanionic properties, the EPS can be considered such as GAG-mimetics, but this similarity can be improved by either chemical or enzymatic modifications. The ability of the native HMW form of HE800 EPS to enhance in vivo bone repair has been demonstrated in a rat model (Zanchetta et al. 2003). We assumed that native HE800 EPS could be a good candidate as a biomaterial employed to design biocompatible scaffolds, especially in association with fibrillar collagens. HE800 EPS has been identified as a good candidate for constructing skin substitutes or dermal equivalent with functional properties, especially in association with fibrillar collagens. It presents HA-like activities, and it can promote both collagen structuring in dermal equivalent and fibroblast colonization of this reconstructed tissue (Senni et al. 2013). A very recent study showed that a biomaterial made of HMW GY785 EPS incorporated in an injectable cellulose-based hydrogel-bearing siloxane group (silated hydroxypropylmethylcellulose or Si-HPMC) and transplanted into nude mice stimulated the production of a cartilage-like extracellular matrix containing high amounts of GAG and type II collagen when compared to Si-HPMC alone.

16

Enzymatic Technologies for Marine Polysaccharides

These results indicate that HMW GY785 EPS-enriched Si-HPMC is a promising hydrogel for cartilage tissue engineering (Rederstorff et al. 2017). Low-molecular-weight sulfated or oversulfated derivatives have been prepared from HE800 and GY785 EPS, respectively. LMW HE800 EPS and LMW GY785 EPS derivatives have been obtained by a free-radical depolymerization. Then a sulfation reaction has been developed using a chemical process to generate new bioactive derivatives having a molecular weight of 60oC Sodium salt soluble; K+ and Ca2+ salts insoluble Soluble Na+, Ca+, K+ salts insoluble, but swells Gels, strongest with K+ Soluble, when hot

Soluble at >60oC Sodium salt soluble; K+ and Ca2+ salts give thixotrophic dispersion Soluble Insoluble

Soluble Na+ salt soluble

Gels strongest with Ca++ Soluble with difficulty

No gelation Soluble, when hot

Insoluble

Soluble, when hot

Soluble, when hot

No Stable Yes Poor

Yes Stable No Good

Yes Stable No Good

Hot (80oC) milk Cold (20oC) milk Gelation Concentrated sugar solution Concentrated salt solution Stability Freeze–thaw pH >5 Syneresis Salt tolerance

Source: Adapted from Rudolph (2000).

Soluble Soluble, thickens

36

Enzymatic Technologies for Marine Polysaccharides

to control syneresis, generally decrease with the increasing degree of sulfation in these polymers. Carrageenan gels are usually prepared by heating the water suspension of the gel followed by cooling. However, cold gelation of κ-carrageenan in the presence of sodium ions under power ultrasound has been reported (Farahnaky et al. 2013). In common practice, κ- and ι-carrageenans are used to prepare water dessert gels, whipped toppings, instant whipped desserts, and egg-less custards and flavors (Kilinç et al. 2013; Puvanenthiran et al. 2003). Carrageenan can reduce oil absorption during frying of products such as fish sticks and poultry products. Its presence has been reported to lower the contents of lipids and cholesterol by as much as 48% and 44%, respectively, in pork patties, associated with 31% reduction in calories (Kumar and Sharma 2004). The combination of 0.1% of any of the carrageenans and 0.5% citric acid was able to inhibit browning of unpasteurized apple juice containing 0.1% sodium benzoate for up to 3 months at 3°C (Tong and Hicks 1991). κ-Carrageenan improved cooking yield and reduced expressible moisture in products such as onion rings and low-fat sausage formulations containing higher proportions of potato starch (García-García and Totosaus 2008; Glicksman 1987). Processed Euchema seaweed (PES) [also known as Philippines Natural Grade (PNG), semirefined carrageenan (SRC), refined carrageenan (ARC) or alkali modified flour (AMF)], prepared by alkali treatment of the algae, E. cottonii and E. spinosum, is a commercial food additive, which has κ-carrageenan as its major component. With appreciable water-and oil-binding capacities, PES is used as thickener, gelling agent, stabilizer, and emulsifier in processed meat, fish, and dairy products, at concentrations ranging from 8% to 10% (Glicksman 1987). Table 2.4 lists various uses of alginate and carrageenans in different food categories.

2.4.2.4  Food uses of other seaweed polysaccharides Some of the other seaweed polysaccharides used as food additives are fucoidans and laminarins (from brown algae), xylans (certain red and green algae), floridean starch (red algae), ulvans (green algae), and cellulose (in all genera). Fucoidan is a water-soluble, naturally occurring sulfated polysaccharide predominantly present in brown algae cells. Because of their antioxidant properties, fucoidan and laminarin are used to control lipid oxidation in muscle foods (Venugopal 2011a). Floridean starch exhibited low gelatinization temperature, low viscosity, high clarity, and little or no retrogradation on repetitive freeze–thaw cycles, which are useful for their applications in instant noodles and deepfrozen foods (Yu et al. 2002). It facilitates protein precipitation and hence clarifies beer (Belitz et al. 2004). Furcelleran forms thermally reversible gel at 0.2–0.5%, which is useful in puddings, cake fillings, and icings, and in processed meat products, such as spreadable meat. The gel has an advantage over pectin in marmalades since it is stable at sugar concentrations at 50–60%.

2.4.3  Polysaccharides from microalgal and marine microorganisms Polysaccharides secreted by terrestrial microorganisms are used by the food industry as stabilizers, thickeners, emulsifiers, texturizers, and gelling agents. These include xanthan gum, gellan gum, dextran, and pullulan, which have been approved by food regulatory agencies (Ramalingam et al. 2014). Marine microalagae can be rich sources of polysaccharides having potential food and other applications (Plaza et al. 2009). Because of their high viscosities over a wide range of pH, temperature, and salinity, microalgal polysaccharides can function as thickening agents (Venugopal 2016). Biopolymers from two marine Chrysophyta possess antioxidant activities (Sun et al. 2014). Similarly, an extracellular

Chapter two:  Applications of marine polysaccharides in food processing

37

Table 2.4  Some uses of alginates and carrageenans in quality improvement of some food products Applications

Alginate

Carrageenans

Water-binding agent, controls Provides freeze–thaw stability, Bakery products, including syneresis, emulsifier, enhances bread, pie fillings, cake mixes, texture, and reduced syneresis loaf volume, water absorption, to products; prevents staling jellies, marshmallow and hardening of bread crumbs and improves crumb quality; toppings, glazes, syrup, prevents staling and hardening puddings, cheese spreads, of bread crumb confectioneries Fishery products, including Enhance cooking yield, hardness Enhance cooking yield, hardness, texture and fiber surimi items, burgers, and bind strength, texture and content; antioxidant, and sausages fiber content; useful for antimicrobial activities restructured products such as shrimp substitutes Red meat items, restructured Endows food products with Increases yield, improves products, burgers thermostability and desired rigidity, and decreases consistency expressible juice; enhances storage stability, sliceability, and rigidity; antioxidant, and antimicrobial activities Vegetable products, including Thickening, gelling, stabilizing Thickening, gelling, stabilizing agents; reduces or replaces agents; alginate-pectin gels fruit preserves, sauces, pectin in jams and jellies; give better gel strength than gravies, jams, marmalades, inhibits browning of fruit individual components; canned foods juices either alone or along thickening agent in bakery with citric acid fillings, dessert gels, salad dressings, sauces, syrups, toppings, etc. Thickening, gelling, stabilizing Act as stabilizing, thickening Dairy products, including and gelling agents; agents; prevent formation of pudding, milk shakes, ice κ-carrageenan–casein large crystals in ice cream; cream, chocolate milk, interactions stabilize ice improve texture of cheese desserts, cheese spreads cream, ι-carrageenan increases spreads; PGA resists loss of viscosity; its unique suspension viscosity and sensory values of flavored soy milk; function and foaming properties useful in soft drinks, milk drinks, and as fat substitutes ice cream; it helps suspend cocoa in chocolate milk Sources: Adapted from Brownlee et al. (2005), Venugopal (2016, 2011a), and Abdul Khalil et al. (2018).

mannan from a marine bacterium exhibited high antioxidant activity having potential use as food supplement (Guo et al. 2010). Although the polysaccharides of three red microalgae, Porphyridium sp., P. aerugineum and Rhodella reticulate, differed in composition, their rheological characteristics are comparable. Mixtures of the algal polysaccharides with locust bean gum (LBG) exhibited synergism in rheological properties and syneresis. However these polysaccharides had lower gel strength than agar (Geresh and Arad 1991). There is good potential for use of EPSs from cyanobacteria, diatoms, and other marine microalagae as well as organisms from marine extreme environments as surfactants and emulsifiers (de Jesús Paniagua-Michel et al. 2014). Cyanobacterial EPSs generally show a pseudoplastic behavior, but with marked differences in both viscosity values and shear thinning

38

Enzymatic Technologies for Marine Polysaccharides

(De Philippis and Vincenzini 1998). EPS of Spirulina platensis, which can be optimally produced by a two-stage culture of the microalga, possesses many biological functions (Lee et al. 2012). The slow progress in isolation of EPSs from marine microorganisms from extreme environments is due to difficulties in their large scale cultivation, as mentioned earlier.

2.5  Polysaccharide-based edible films and coatings Edible films and coatings have provided an interesting means for controlling the quality and stability of food products. Polysaccharides are raw materials for biodegradable films. These coatings have advantages over synthetic coatings such as being edible and more environmentally friendly. In these films and coatings, polysaccharides form cohesive networks with other polymers such as proteins through covalent and/or noncovalent interactions. Glycerol and polyethylene glycol serve as plasticizers for the films, while acetic or lactic acid regulate their pH. Food items within the films exhibit enhanced shelf life as a result of antimicrobial and antioxidant activities of the polysaccharides and also enhanced permeability barrier properties of the film to oxygen, carbon dioxide, and moisture (Neito 2009; Guilbert et al. 1995). The coatings can be given by techniques such as atomized spray systems and also panning, fluidized-bed application, dipping, or spraying (Andrade et al. 2012). Benefits of polysaccharide-based films and edible coatings are summarized in Table 2.5. Carrageenan and alginate films are nontoxic, biodegradable, and biocompatible (Tavassoli-Kafrani et al. 2014). Incorporation of lysozyme to alginate film can enhance its antimicrobial properties (Amara et al. 2016). Coating with 4% (w/w) sodium alginate followed by modified atmosphere packaging extended shelf life of oysters up to 160 hours compared with 57 hours for unpackaged control (Coasta et al. 2014). Capsules of alginate can hold up to 15% (w/w) PUFA-rich fish oil. The amount of encapsulated oil remains stable during the gastric stage of human digestive system, but it is released at the intestinal phase (Bannikova et al. 2018). Alginate encapsulation also helps slow release of flavors, and other functional compounds (Ching et al. 2017). Chitosan-based films have witnessed emerging food packaging applications as antibacterial barrier. Chitosan films are highly flexible, resistant, biodegradable, and safe for consumption (Wang et al. 2018; Dutta et al. 2012). Chitosan coating alone or in combination with citric acid pretreatment was beneficial in reducing weight loss and quality changes in carrots, ensuring better color retention and superior sensory and microbial quality during refrigerated storage for 10 days (Ushkala et al. 2012). Low-dose γ-irradiation (2.5 kGy) and chitosan coating (2%) containing grape seed extract (0.1% w/w) significantly improved the sensorial quality of chicken breast meat during 21 days of storage at 4°C (Hassanzadeh et al. 2017), Reinforcement of chitosan film with cellulose of coconut fibers enhanced moisture resistance, tensile strength and thermal stability (Bhuvaneshwari et al. 2011). Chitosan-gelatin film incorporated with β-carotene-loaded Table 2.5  Benefits of polysaccharide-based films and edible coatings Extend the shelf life of foods Alleviate the problem of moisture loss in frozen muscle foods Prevent drip loss Reduce rancidity and spoilage Reduce volatile flavor loss Reduce the load of microorganisms, including pathogens Potential carriers of antioxidants (e.g., tocopherol) and/or antimicrobials (lysozyme) Reduce oil uptake during frying

Chapter two:  Applications of marine polysaccharides in food processing

39

starch nanocrystals had decreased swelling index, solubility in water, and moisture content (Hari et al 2018). Modified atmosphere storage at 27°C of mango covered with chitosan film resulted in shelf life extension of the fruits up to 18 days (Srinivasa et al. 2002). Nanocomposite films of chitosan and other marine hydrocolloids such as agar, alginate, and carrageenan can be a good carrier for nutraceuticals such as PUFA (Comunian and Favaro-Trindaade 2016; Venugopal 2009). Composite films of these polysaccharides with other gums such as low methoxypectin have better strength and barrier properties (Galus and Lenart 2013). Alginate and also chitosan are raw materials for electrospinning for the development of nanoscale microfibrous structures having a wide range of compositions, morphologies, mechanical properties, and bioactivities (Mendes et al. 2017).

2.6  Marine oligosaccharides Marine oligosaccharides are naturally produced in algae and/or by hydrolysis of polysaccharides. Oligosaccharides, including those of marine origin, can modify the viscosity and freezing point of foods, affect emulsification and gel formation, and act as humectants (Shahidi and Abuzaytoon 2005). These compounds are rarely digested by gastrointestinal enzymes. Oligosaccharides, including those from agar and alginate, possess antioxidative activities, and also have potential as prebiotics or biopreservatives, sweeteners, fiber, and humectants (Jutur et al. 2016; Trincone 2014; Patel and Goyal 2010). Chitosan oligosaccharides (COSs) are characterized by their low molecular weights, low viscosity, short chain lengths, and antioxidant and antimicrobial activities (Jung and Park 2014; Lin and Chou 2004). Because of these functionalities oligosaccharides are considered “a boon from nature’s desk” (Belorkar and Gupta 2016).

2.7  Enzymatic processes for marine polysaccharides Enzymes are traditionally used for food processing and production of food ingredients. In the realm of carbohydrates, glycosyl hydrolases (carbohydrases) comprise glucosidases, galactosidases, amylases, cellulases, agarases, chitinases, and other enzymes. These enzymes allow selective extraction seaweed ingredients at mild conditions as well as tailor-made modifications of algal polysaccharides (Rhein-Knudsen et al. 2015; Sathya and Khan 2014). Agarases, which liquefy agar by cleaving either α-l-(1,3) or β-d-(1,4) linkages, have wide applications in the biotechnological, food, and cosmetic industries (Jahromi and Barzkar 2018). Marine organisms including seafood and their processing wastes are sources of enzymes having interesting features for diverse applications (Venugopal 2016). Deacetylation of chitin by chitinase is less expensive and environmental friendly (Krolicka et al. 2018). Enzymatic processes are also available for production of COSs, employing specific enzymes such as chitosanases and nonspecific enzymes such as cellulases and lipases (Sathya and Khan 2016; Dahiya et al. 2006). Alginate lyase– mediated depolymerization gives alginate oligosaccharides (AOs) having antioxidant activity (Falkeborg et al. 2014). A novel alginate-degrading enzyme from marine bacterium Microbulbifer spp. produced disaccharide and trisaccharide from alginate, which exhibited antioxidant activities (Zhu et al. 2016). Low-molecular-weight chitosan products have been prepared using chitinase, lysozyme, and cellulase (Hamed et al. 2016; Lin et al. 2009; Kim and Rajapakse 2005). Direct production of COSs from chitin and crab shells by a combination of mechano-chemical grinding and enzymatic hydrolysis has been reported (Jung and Park 2014). Oligosaccharides can also be directly prepared from E. cottonii using cellulase and recombinant κ-carrageenase, produced by a strain

40

Enzymatic Technologies for Marine Polysaccharides

of Escherichia coli, which expresses κ-carrageenase from a marine bacterium (Duan et al. 2016). Supplementation of carbohydrases in aquafeed mitigates the adverse effects of non-starch-based polysaccharides, improves nutrient digestibility, and reduces nutrient excretion (Sinha et al. 2011). Biotechnological methods such as a sequential enzymatic acid–alkaline treatment for chitin extraction have been reported (Vázquez et al. 2017; Mao et al 2017). Enzymes isolated from cold-adapted organisms from polar region are characterized by high catalytic efficiencies at low temperatures compared with their mesophilic counterparts. These enzymes allow shorter periods for processing, protection of thermosensitive substrates or products, and prevention of undesired chemical transformations during the enzymatic reaction (Siddiqui and Cavichioli 2006).

Conclusions Marine polysaccharides, because of their natural origin, are generally safe for consumption, unlike many synthetic food additives. Chitin and chitosan fall within the lowest category of toxicological concern. Chitosan has been in use for maintenance of health; some commercial products include “Fat Absorb,” “Seaborne,” “EssentialSea,” and “EssentialSeaPlus.” Chitosan-fortified foods are available in the United States and Japan (Venugopal 2018). In Japan chitin and chitosan are approved as functional food ingredients, since they possess several required attributes of functional foods such as enhancement of immunity, delaying of ageing, and recovery from illness. Chitosan and its derivatives are finding several end users encompassing food, healthcare, and other industries, with a projected value of US $4.2 billion by 2020 (Hamed et al. 2016) (see also Global Industry Analysis, available at www.strategyr.com/Chitin_and_Chitosan_Market_ Report.asp). Algal products enjoy an estimated annual market worth about US$6 billion (Carlsson et al. 2007; Kraan 2012). Commercially important seaweed products include processed euchema seaweed (PES), MODIFILAN, and propylene glycol alginate (PGA). Agar, alginate, and carrageenan have been approved for use in foods by Codex General Standards for Food Additives (Codex Standard 192-1995), the United States Food and Drug Administration (FDA), and the European Union (Venugopal 2011a). Some concerns remain regarding toxicological safety of degraded carrageenans (Venugopal 2011, 2009). In spite of their high food and medicinal value, poor global consumption of seaweed is due to inadequate consumer awareness on their nutritional value as well as their limited availability, especially within inland cities (Rioux et al. 2017). In conclusion, there is potential for exploring polysaccharides from diverse marine sources for food product development.

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chapter three

The manufacture, characterization, and uses of fucoidans from macroalgae J. H. Fitton, D. N. Stringer, S. S. Karpiniec, and A. Y. Park Marinova Pty Ltd, Cambridge, Tasmania, Australia

Contents 3.1 Introduction........................................................................................................................... 47 3.2 Sources of fucoidan.............................................................................................................. 48 3.2.1 Brown seaweeds........................................................................................................ 48 3.2.2 Echinoderms.............................................................................................................. 48 3.3 Methods for fucoidan extraction........................................................................................ 48 3.4 Industry standards: Is it fucoidan?..................................................................................... 53 3.5 Quantifying and identifying fucoidan in biological fluids............................................ 53 3.6 Fucoidan applications..........................................................................................................54 3.6.1 Food supplements.....................................................................................................54 3.6.2 Pharmaceuticals........................................................................................................ 55 3.6.3 Biomaterials............................................................................................................... 55 3.6.4 Cosmetics................................................................................................................... 56 3.6.5 Animal applications................................................................................................. 56 3.6.6 Agricultural applications......................................................................................... 56 3.7 Safety and regulation for use in food and supplements................................................. 57 Summary......................................................................................................................................... 60 References........................................................................................................................................ 60

3.1 Introduction Fucoidans are sulfated, complex, fucose-rich polymers found in brown seaweeds and echinoderms (Fitton 2011; Fitton, Stringer, and Karpiniec 2015). Their function within brown seaweeds appears to be protective, with the highest concentrations associated with the reproductive parts of the algae. Fucoidans have structures that are speciesspecific, with properties that vary in composition, depending on the source biomass used. The polysaccharide backbone is typically composed of fucose subunits, but can also have significant contributions from galactose, xylose, arabinose, and rhamnose. Fucoidans have demonstrated effects as anticoagulants and as anti-inflammatory and anticancer agents in vivo. Fucoidans from different species are also well known as viral entry blocking agents, and are effective against a wide range of coated viruses and some bacteria. Fucoidans have well-documented effects on cancer cell cycle arrest, enzyme inhibition, thrombolytic activity, and immune modulation (Fitton 2011; Fitton, Dell’Acqua, Gardiner et al. 2015). 47

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When extracted from whole seaweed, various fucoidans are a heterodisperse mixture of similar polymers, and fucoidan extracts can contain other compounds found within the plant, depending on the extraction techniques used. The molecular weight (MW), the polydispersity, and the degree of sulfation within fucoidan extracts of one species are largely dependent on the type of extraction method applied to the initial biomass. As a general rule, when using a simple water extraction, higher temperatures and lower pH give rise to preparations with a lower average MW and less sulfation. Commercially, fucoidans have been used as an ingredient in food supplements for at least two decades. The supplements are available as capsules, tablets, beverages, and gels, and are used to support human health in a complementary setting. Fucoidans are also available and in current use as topical cosmetic ingredients. Fucoidans are not currently used in any therapeutic applications classified as “pharmaceutical.”

3.2  Sources of fucoidan 3.2.1  Brown seaweeds Brown macroalgae are the major source materials for the extraction of fucoidans. Although all brown macroalgae contain fucoidan, industrial sources require access to the large biomass “kelps,” which are available from sustainable harvesting operations. In practice, commercially available fucoidans found in food supplements and cosmetics tend to come from a small number of species: Fucus vesiculosus, Ascophyllum nodosum, Ecklonia spp., Undaria pinnatifida, Cladosiphon spp., Laminaria japonica, Macrocystis pyrifera, and Kjellmaniella crassifolia. Worldwide, there are manufacturers of bulk fucoidan in Japan, Russia, China, Korea, Vietnam, and Australia.

3.2.2 Echinoderms The fucoidans extracted from echinoderms have one major difference from those derived from macroalgae; they are linear—not branched—molecules (Berteau and Mulloy 2003). Sea cucumbers are highly valued as a whole food product, with prices 10–100-fold that of dried macroalgae, making it economically less viable to produce fucoidan extracts from them. Sea urchins also contain linear fucoidan concentrated in the egg masses (Berteau and Mulloy 2003). A comprehensive research study on sulfated glycans from invertebrates has been carried out by a Brazilian research group (Pomin 2015). To the authors’ knowledge, there are no products containing invertebrate-sourced fucoidans currently available in the food supplement sector.

3.3  Methods for fucoidan extraction Fucoidans can be extracted from the starting biomass by various processes and further purified thereafter using additional techniques. Laboratory methods have traditionally relied on acidic extraction in aqueous media using heat, whereupon the fucoidans are released into the water phase. Ethanol (up to 60% v/v) is then used to precipitate the fucoidans, relying on the inherent lack of solubility of these molecules in less polar solvents. The resulting precipitate can then be further solubilized and reprecipitated if additional purification is required. The 1952 descriptions by Black remain an excellent guide to these classical methods (Black, Dewar, and Woodward 1952). Currently, much of the commercially available fucoidan, including the research-grade fucoidan (crude Fucus

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vesiculosus extract; Sigma-Aldrich, St. Louis, MO, USA, Code F5631), is produced using variations of this method. Alternatives to the use of ethanol to precipitate the fucoidan include selective membrane filtration, or resin-based chromatography to purify the fucoidans. In general, a preliminary fucoidan extract will contain free salt, as well as coextracts from the plant, which are typically polyphenols, alginates laminarin, and mannitol. Depending on the extent of subsequent purification steps, the purity of the resulting fucoidan can be optimized. An excellent review on extraction methods has recently been presented (Flórez-Fernández, Torres, González-Muñoz et al. 2018). New methods that can enhance extraction yields include enzyme-mediated release and modification, which yields a fucoidan extract or fraction that is, by definition, distinct from the native fucoidan from which it was derived. Endo- and exoenzymes from marine bacteria, marine mollusks, and fungi and even terrestrial fungi and ticks have the ability to digest fucoidans and potentially aid in their extraction. Fucoidanase activity was reviewed in 2003 (Berteau and Mulloy 2003) and more recently in 2015 (Kusaykin, Silchenko, Zakharenko et al. 2016). These comprehensive expert reviews present an excellent overview of the known enzymes active with fucoidans. Enzyme-mediated modification of fucoidans offers the possibility of defined fractions, a way toward obtaining fucoidans with altered biological activities. For example, a reduction in anticoagulant activity could be perhaps achieved by reducing specific sulfate groups. However, the activity of fucoidanase enzymes can be quite specific, with one discovery indicating that a fucoidanase would selectively desulfate at position 2 (Daniel, Berteau, Chevolot et al. 2001), while another found that a scallop derived fucoidanase was able to digest fucoidan from Ascophyllum nodosum, but not from Fucus vesiculosus (Berteau, McCort, Goasdoué et al. 2002). In another example, fucoidan hydrolase, α-l-fucosidase, and arylsulfatase from the marine mollusk Littorina kurila were isolated and described (Kusaykin, Burtseva, Svetasheva et al. 2001; Kusaykin, Bakunina, Sova et al. 2008). It was found that α-l-fucosidase and arylsulfatase cannot hydrolyze natural fucoidan, whereas fucoidan hydrolase cleaves fucoidan to produce sulfated oligosaccharides and fucose. Other investigators isolated an enzyme from a marine bacterium with specific hydrolytic activity for fucoidans extracted from Fucus evanescens and Fucus vesiculosus, which was not replicated in fucoidan from Saccharina cichorioides (Silchenko, Kusaykin, Kurilenko et al. 2013), while a terrestrial fungal enzyme from Aspergillus niger degraded Laminaria japonica fucoidan (Rodriguez-Jasso, Mussatto, Pastrana et al. 2010). Fucosidase enzymes found in many other species do not necessarily include the ability to digest fucoidan. The α-l-fucosidase found in ticks can digest fucoidan (Moreti, Perrella, and Lopes 2013), but a fucosidase recently discovered in spiders cannot (Perrella, Fuzita, Moreti et al. 2018). To date, no specific fucoidanases are available at a commercial scale for the processing of fucoidans from marine algae. One type of fucoidan from Cladosiphon spp. available from a Japanese manufacturer is reported to be an abalone glycosidase–digested fucoidan extract (Shirahata, Zhang, Yoshida et al. 2013); however, to date there have been no other enzymatically altered fucoidans available commercially. Additional techniques for fucoidan extraction include microwave, high-pressure, and ultrasound processing that offer both laboratory-scale and industrial isolation of fucoidans with short extraction times. In future, these technologies might be used commercially, although they will need to be optimized for each type of fucoidan as described above. In a recent study the structural features of fucoidan extracts from Fucus vesiculosus were found to be influenced by the extraction pressure, time, and water: algae ratio (RodriguezJasso, Mussatto, Pastrana et al. 2011). Such structural variations resulting from extraction

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methods have been well described (Ale, Mikkelsen, and Meyer 2011), where the importance of maintaining structural integrity throughout the fucoidan extraction process, in order to achieve consistent biological functionality, is reinforced. A microwave-assisted extract of a fucoidan from Ascophyllum nodosum was effective in yielding “antioxidant” fractions (Yuan and Macquarrie 2015). An important determinant for commercial fucoidan used in food or food supplements is a lack of organic solvents. Regulations require an absence of solvent, and thus nonsolvent methods of extraction are preferred for this application. As mentioned earlier, fucoidan extracts may contain “nonfucoidan” components. The presence of such bioactive compounds should be established as they will interfere with any structure–function study focused on fucoidans. These actives may themselves be complex carbohydrates such as the ubiquitous alginates, or the glucose-rich storage polymers prevalent in the laminariales, such as laminarin (Kadam, O’Donnell, Rai et al. 2015). Additionally, substantial quantities of nonpolysaccharide components, such as the sugar-alcohol, mannitol, and importantly, potent marine antioxidants—polyphloroglucinols—are also routinely observed in fucoidan-containing extracts (Zaragoza, Lopez, Saiz et al. 2008; Parys, Rosenbaum, Kehraus et al. 2007). The presence of these components should be particularly recognized for the contributing effect they may have on antioxidant activity. Thus, without appropriate extraction and purification controls, any conclusions about any observed biological activity should be considered with these impurities in mind. This is particularly true when comparing biological activities of extracts from laboratory-based methods that have not been validated. In the last decade, the availability of commercial-scale, validated fucoidan extracts has allowed a more systematic approach to fucoidan research. Scaleup from the laboratory requires consideration of volumes, availability and selection of biomass, residual solvents, and chemical and microbiological safety. All of these considerations may alter the chosen method of isolation. Chemical analytical techniques have been developed for comprehensive carbohydrate profiling and linkage analysis together with sulfate, acetyl, and counterion quantification— all of which contribute to the overall composition. Together, these analyses provide the data necessary to adequately describe and contrast fucoidan-containing extracts from different sources and species. Examples of composition are summarized in Table 3.1, for fucoidan extracts produced systematically using an aqueous extraction process under controlled pH, temperature, and time conditions. The data presented are modified from a previous review at Marinova Pty Ltd.’s laboratory according to the methods described next (Fitton, Stringer, and Karpiniec 2015). The carbohydrate profile was obtained using a gas chromatography (GC)-based method for the accurate determination of individual monosaccharide ratios in a sample. This method relies on the preparation of acetylated alditol derivatives of the hydrolyzed samples (Morvai-Vitányi, Molnár-Perl, Knausz et al. 1993). The uronic acid content was determined by spectrophotometric analysis of the hydrolyzed compound in the presence of 3-phenylphenol, against glucuronic acid standards, based on a previously described method (Filisetti-Cozzi and Carpita 1991). Sulfate content was analyzed spectrophotometrically using a BaSO4 precipitation method (BaCl2 in gelatin), based on existing work (Dogson 1961). Cations, including Na, K, Ca, and Mg, were determined by standard flame atomic absorption spectroscopy. Molecular weight profiles were determined by gel permeation chromatography, with the aid of a size-exclusion column and are reported relative to dextran standards. The polyphenolic components can be

Species Fucus vesiculosus Sigma Crude Fucus vesiculosus Ascophyllum nodosum Macrocystis pyrifera Cladosiphon spp. Laminaria japonica Ecklonia radiata Durvillaea potatorum Lessonia nigrescens Alaria esculenta Undaria pinnatifida Pelvetia canaliculata

Fucose (%)

Xylose (%)

Galactose (%)

Arabinose (%)

Rhamnose (%)

Uronic acid (%)

Sulfate (%)

Cations (%)

Sulfation ratio

43.1 45.9

8.8 3.3

2.2 4.3

1.2 0

0.2 0

8.7 7

30.1 32

5.7 7.6

0.81 0.92

82.5 20.7

36.1

10.6

2.0

0.0

0.7

4.1

24.2

2.2

0.8

224.0

30.5

2.2

5.6

0

1.7

12.4

32.4

15.1

1.1

176.4

51.2 34.1 19 27.9

2.1 1 11 2.1

1.3 4.2 12 6.2

0 0.3 6.2 0.3

0 1 1.7 0.7

15.5 14.4 25.5 32.4

23 31.7 19.1 21.4

7 13.2 5.4 8.9

0.58 1 0.45 0.57

1927.2 395.4 528.2 336.3

26.2

8.1

13

2

0.9

12.9

29.1

7.8

0.82

491.8

37.5 32.6

3.4 0

16.4 25.2

0.6 0.5

1.3 0.4

12.3 4

20.2 29.6

8.2 7.7

0.5 0.85

147.9 51.7

38.3

3

5.6

0.1

0

43

5.7

1.4

103.9

4.3

Peak average MW (kDa)

Chapter three:  Manufacture, characterization, and uses of fucoidans from macroalgae

Table 3.1  Absolute mass percentages of components of fucoidan extracts from varying species

51

52

Enzymatic Technologies for Marine Polysaccharides

determined spectrophotometrically using the Folin-Ciocalteu reagent (Jiménez-Escrig, Jiménez-Jiménez, Pulido et al. 2001; Zhang, Zhang, Shen et al. 2006; Zoecklein, Fugelsang, Gump et al. 1995). The crude Fucus vesiculosus extract historically provided by Sigma-Aldrich (St. Louis, MO, USA, Code F5631) was also analyzed, and the composition of its fucoidan component is provided for comparative purposes. In addition to the fucoidan components described above, the Sigma-Aldrich extract (Code F5631) also contained 3.9% polyphenols and 0.4% mannitol. Other investigators have fractionated the same Sigma fucoidan and reported on its fractional composition (Nishino, Nishioka, Ura et al. 1994). For the purpose of comparing the fucoidan compositions in Table 3.1, the uronic acid content has been included in the data. While the inclusion of uronates in fucoidans from Cladosiphon spp. has been clearly established (Nagaoka, Shibata, Kimura-Takagi et al. 1999), caution is urged in assuming that the full uronic acid component determined above is contained within the fucoidan polymer, as it can be indicative of alginate. Further linkage analyses would be required to fully distinguish fucoidan uronates from simple “coextracted” alginate uronates. Variations in fucoidan composition, with respect to the source species, are clearly highlighted in Table 3.1, with extremes in fucose content from 51.2% for Cladosiphon spp. to 19.0% for Ecklonia radiata. Similarly, the galactose content varied from the widely recognized, galactose-rich subtidal Alariales, such as Undaria pinnatifida and Alaria esculenta at 25.2% and 16.4%, down to the low-galactose-containing intertidal Cladosiphon spp. and Fucus vesiculosus at 1.3% and 2.2%, respectively. Broad ranges in sulfation ratio are also observed, from lows of 0.50 for Alaria esculenta to the highly sulfated Pelvetia canaliculata at 1.4. Although not analyzed for all samples in the above sample set, fucoidan acetyl content is also known to vary substantially with, for example, as much as 3% by mass acetyl content in Undaria pinnatifida fucoidan and less than 0.5% for Fucus vesiculosus fucoidan, as determined by nuclear magnetic resonance (NMR) spectroscopy techniques (unpublished data). With such complex compositional variations observed between species that are exposed to identical extraction and purification procedures, it is evident that comprehensive chemical analyses are required for any exploration of fucoidan bioactivity. Additional methods for examination of the tertiary structure of fucoidans include NMR spectroscopy and mass spectroscopy. As recently outlined, mass spectroscopy has limitations because the fucoidans tend to desulfate rather than ionize during measurement (Anastyuk, Imbs, Dmitrenok et al. 2014; Anastyuk, Shevchenko, and Gorbach 2015). These authors describe rational examination of different fucoidans using exhaustive hydrolysis followed by matrix-assisted laser desorption/ionization–time-of-flight (MALDI-TOF) and electrospray ionization (ESI) mass spectroscopy, and recently determined some further intricacies of the linking of the sugars within a fucoidan backbone— specifically, previously unseen (1,2) linkages in fucoidan from S. cichorioides as well as C-4 substitution of fragments of fucoidan from F. evanescens (Anastyuk, Shevchenko, Belokozova et al. 2018). The molecular weight profiles of fucoidan extracts are polydisperse and can be highly dependent on the extraction methodology used. This can be seen in the contrasting molecular weights of the aqueous Fucus vesiculosus extract reported here, and that of the crude Fucus vesiculosus extract available from Sigma-Aldrich, which exhibits a markedly lower molecular weight profile, perhaps demonstrating significant hydrolysis of the fucoidan during processing. The molecular weight peak averages of the extracts characterized in Table 3.1 are also summarized in that table.

Chapter three:  Manufacture, characterization, and uses of fucoidans from macroalgae

53

Table 3.2  Absolute mass percentages of components of commercially available fucoidan extracts determined using standard analysis techniques

Sample no.

Description (claimed)

Total carbohydrate (%)

1 2 3 4 5 6 7 8 9 10

90% fucoidan extract 85% fucoidan extract 80% fucoidan extract 85% fucoidan extract 85% fucoidan extracta 85% fucoidan extract 85% low-MW fucoidan 85% fucoidan extract 80% fucoidan extract 19.3% fucoidan extract

52.6 32.1 37.2 20.7 N/D 75.0 42.4 73.8 71.5 38

a

Fucose (%)

Galactose (%)

Uronic acid (%)

Fucoidan (%)

24.2 16.9 16.9 8.7 N/D 7.9 99%

120

> 99%

120 60 w (°)

A

0

(C)

Bnr – Cnr

Cnr – Anr

60

–60

–60

–60

–120

–120

0

60

120

c (°)

(D)

60 0

63% A

24% C

–120 –120

–60

120

0

120

–120

–60

120

0

B 2% –120

–60

0

60

120

c (°)

Cr – A′r 120 60

w (°)

120

98% A

60

–120 60

60

Br – Cr

–60

0

0 c (°)

120

c (°)

(G)

–60

(F)

B 1%

c (°)

B < 1% –120

> 98% A

< 1% C

60

–120 60

60

Dr – Br

–60

0

0 c (°)

120

B 13%

–60

–60

(E)

Anr – Dr 120

–120

w (°)

–60

A

0

–120 –120

> 99%

120

A

0

w (°)

w (°)

60

w (°)

(B)

D′nr – Bnr

w (°)

(A)

0

2% D

–60 –120 –120

88% 8% A C B 2% –60

0

60

120

c (°)

Figure 6.9  Dihedral angle (ψ/φ) distributions of the glycosidic linkages in Lv I. Each dot represents a configuration in the MD simulation of the disaccharide building blocks of Lv I. (A) D′nr−Bnr, (B) Bnr−Cnr, (C) Cnr−Anr, (D) Anr−Dr, (E) Dr−Br, (F) Br−Cr, and (G) Cr−A′r. The letters used were A for 2-sulfation, B for 2,4-disulfation, C for 2-sulfation, D for 4-sulfation, A′ for 2-desulfated, and D′ for the 4-sulfated. The subscripts “nr” and “r” denote, respectively, the “nonreducing end” and the “reducing end.” (Reprinted with permission from Queiroz et al. 2015.)

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Enzymatic Technologies for Marine Polysaccharides (B)

6

RMSD of ring atoms

RMSD of ring atoms

(A)

4

2

50

100

150 200 250 300 350 Simulation time (ns)

400

6

4

2

50

100

150 200 250 300 350 Simulation time (ns)

400

D′ nr – B nr – C nr – Anr – Dr – Br – Cr – A′r Figure 6.10  Comparison of the RMSD between octasaccharide and its composing tetrasaccharide units. (A) The four units belonging to the nonreducing end side (in blue) and (B) the four units belonging to the reducing end side (in red) are denoted by the subscripts “nr” and “r,” which denote, respectively, the “nonreducing end” and the “reducing end.” The black simulation belongs to the entire octasaccharide Lv I. (Reprinted with permission from Queiroz et al. 2015.)

central disaccharide unit between residues A and D can be explained only on the basis of sulfation pattern, since the only difference among all these disaccharide building blocks analyzed by MD is sulfation. The influence of sulfation pattern on dynamics of Lv I can be additionally seen from the MD experiment performed to determine the root-mean-square deviation (RMSD) values of the two component tetrasaccharide units (red and blue data in Figure 6.10) compared to the entire octasaccharide (black data in Figure 6.10). From this experiment, we can observe that the variations and values in the atomic distances in the Lv I octasaccharide are higher than those seen in the tetrasaccharide units analyzed separately. This strongly indicates that the octasaccharide presents enhanced dynamics as compared to the two component tetrasaccharide building blocks. Taking the results of the experiments in Figures 6.9 and 6.10 together, we can conclude that dynamics of the octasaccharide originated primarily from the connection between the 2-sulfated unit (A) and the 4-sulfated unit (D). To support the theoretical results from the computational MD approaches (Figures 6.9 and 6.10), we have measured the longitudinal spin–relaxation (T1) values using a NMR 1D 1H inversion-recovery experiment on the Lv I at three different temperatures (25°C, 37°C, and 50°C (Table 6.3). As expected, the T1 spin–relaxation time values of the 1H anomeric signals from Lv I generally increase as the temperature rises. Relaxation time values of the 1H1 signals from the 2-sulfated unit (A) of the central disaccharide unit and from the 4-sulfated unit (D) of the reducing end terminal are generally longer than the 1H1 signals of the other units, indicating higher dynamics at these units as compared to the other. These NMR results go in favor with the results observed from the MD experiments. A final computational experiment based on MM was carried out to calculate the interatomic distances between the different composing atoms of the octasaccharide Lv I (Queiroz et al. 2015). This experiment indicated that hydrogen bonds are formed between the units inside the two component tetrasaccharides (indicated in blue in Figure 6.11A) and that a repulsive force exists between the units A and D of these two tetrasaccharides

Chapter six:  Hydrolytic mechanisms of defined tetrasaccharide-repeating sulfate fucan

125

Table 6.3  Longitudinal spin–relaxation (T1) of the anomeric 1H signals of the Lv I octasaccharide from L. variegatus. Modified with permission from Queiroz et al. 2015 1

H T1 (s)a

NMR resonance

25°C

37°C

50°C

A H1 (2-sulfated) B H1 (2,4-disulfated) C H1 (2-sulfated) D H1 (4-sulfated) A′H1 (nonsulfated reducing end)

1.13 1.06 1.17 1.26 1.30

1.15 1.07 1.23 1.30 1.36

1.31 1.30 1.31 1.36 1.37

Source: Modified with permission from Queiroz et al. (2015). T1 values were measured via 1H 1D NMR inversion recovery experiment at three temperatures.

OH

(A) O O

O

OSO3 O O OSO3 O

nd s

O

bo en Hy dr og

O OH O O O

OSO3

Br

Dr

O Sulfation-caused repulsion

O

O–

S

O

5.3 Å

O O

OSO3

Cnr OSO3

O O– O S

4.0-4.3 Å

ge

OSO3

nb

on

H

4.7 Å

O

S

O O–

Bnr

OSO3 OH

Anr

OH O

OSO3

OH

O

Hy dro

O

2.7-2.8 Å

OSO3

Cr

OH O

OH

A′r

OH O

(B)

ds

a

OH

D′nr

Figure 6.11 Dynamics and interactomic distances of the fractionated L. variegatus SF-derived octasaccharide obtained by mild-acid hydrolysis. (A) Schematic representation of the sulfation pattern-related hydrogen bonds and repulsive force (dynamic site) in the minimal L. variegatus octasaccharide. The hydrogen bonds are shown in blue and are stabilized by sulfates and hydroxyl groups of the adjacent units. The dynamic site on the central glycosidic bond is driven by the repulsive force between created between the neighboring 2-equatorial and 4-axial sulfations. (B) Calculated interactomic distances of the dynamic and less flexible regions on the L. variegatus octasaccharide (Lv I). (Reprinted with permission from Queiroz et al. 2015.)

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Enzymatic Technologies for Marine Polysaccharides

(indicated in red in Figure 6.11A). This repulsive force is dependent on the adjacent equatorial 2-sulfation of residue A and the axial 4-sulfation of residue D. Representative values of the interatomic distances in these two regions of the Lv 1 octasaccharide are shown in Figure 6.11B. From the results discussed above, we have concluded that while the interresidual hydrogen bonds between hydroxyl and sulfate groups work contributing to decrease dynamics in the tetrasaccharide-repeating units of the L. variegatus SF, the amplified motions produced by the electrostatic repulsive force, caused by the equatorial 2- and axial 4-sulfation located on opposite sides of the central glycosidic bond of the linking disaccharide between the tetrasaccharide units, enhance dynamics. This conclusion has been achieved by advanced conformational and dynamical studies on the octasaccharide Lv I.

Concluding Remarks It is a great coincidence to see that the 2-desulfation and the cleavage of the glycosidic bond during the mild acid hydrolysis of L. variegatus SF investigated in the work of 2005 (Pomin et al. 2005a) occurs exactly at the site of enhanced dynamics in the molecules reinvestigated 10 years later (Queiroz et al. 2015). We believe that the enhanced dynamics on the central glycosidic bond between the tetrasaccharide units of the L. variegatus SF is likely to contribute to the 2-desulfation and consequently to the glycosidic cleavage during the mild-acid hydrolysis.

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Silchenko, Artem S., Nadezhda E. Ustyuzhanina, Mikhail I Kusaykin, Vadim B. Krylov, Alexander S. Shashkov, Andrey S. Dmitrenok, Roza V. Usoltseva, Anastasiya O. Zueva, Nikolay E. Nifantiev, and Tatyana N. Zvyagintseva. 2016. “Expression and Biochemical Characterization and Substrate Specificity of the Fucoidanase from Formosa Algae.” Glycobiology 27(3):254–263 (doi.org/10.1093/glycob/cww138). Sousa, Willer M., Renan O. Silva, Francisco F. Bezerra, Rudy D. Bingana, Francisco Clark N. Barros, Luís E. C. Costa, Venicios G. Sombra, et al. 2016. “Sulfated Polysaccharide Fraction from Marine Algae Solieria Filiformis: Structural Characterization, Gastroprotective and Anti­ oxidant Effects.” Carbohydrate Polymers 152(Nov.):140–148 (doi.org/10.1016/j.carbpol.2016.06.111). Usov, Anatolii I. 2011. “Polysaccharides of the Red Algae.” Advances in Carbohydrate Chemistry and Biochemistry 65:115–217 (doi.org/10.1016/B978-0-12-385520-6.00004-2). Ustyuzhanina, Nadezhda E., Maria I. Bilan, Andrey S. Dmitrenok, Elizaveta Yu. Borodina, Nikolay E. Nifantiev, and Anatolii I. Usov. 2018. “A Highly Regular Fucan Sulfate from the Sea Cucumber Stichopus Horrens.” Carbohydrate Research 456(Feb.):5–9 (doi.org/10.1016/J. CARRES.2017.12.001). Vilela-Silva, Ana-Cristina E. S., Ana-Paula Alves, Ana-Paula Valente, V D Vacquier, and Paulo A. S. Mourão. 1999. “Structure of the Sulfated Alpha-L-Fucan from the Egg Jelly Coat of the Sea Urchin Strongylocentrotus Franciscanus: Patterns of Preferential 2-O- and 4-O-Sulfation Determine Sperm Cell Recognition.” Glycobiology 9(9):927–933 (http://www.ncbi.nlm.nih.gov/ pubmed/10460834). Vilela-Silva, Ana-Cristina E. S., Michelle O. Castro, Ana-Paula Valente, Christiane H. Biermann, and Paulo A. S. Mourão. 2002. “Sulfated Fucans from the Egg Jellies of the Closely Related Sea Urchins Strongylocentrotus Droebachiensis and Strongylocentrotus Pallidus Ensure SpeciesSpecific Fertilization.” Journal of Biological Chemistry 277(1):79–87 (doi.org/10.1074/jbc. M108496200). Vilela-Silva, Ana-Cristina E. S., Noritaka Hirohashi, and Paulo A. S. Mourão. 2008. “The Structure of Sulfated Polysaccharides Ensures a Carbohydrate-Based Mechanism for Species Recognition during Sea Urchin Fertilization.” The International Journal of Developmental Biology 52(5–6): 551–559 (doi.org/10.1387/ijdb.072531av). Wu, Mingyi, Li Xu, Longyan Zhao, Chuang Xiao, Na Gao, Lan Luo, Lian Yang, Zi Li, Lingyun Chen, and Jinhua Zhao. 2015. “Structural Analysis and Anticoagulant Activities of the Novel Sulfated Fucan Possessing a Regular Well-Defined Repeating Unit from Sea Cucumber.” Marine Drugs 13(4):2063–2084 (doi.org/10.3390/md13042063).

chapter seven

Biosynthesis and extrusion of β-chitin nanofibers by diatoms Gregory L. Rorrer

Oregon State University, Corvallis, Oregon

Contents 7.1 Introduction......................................................................................................................... 129 7.2 Morphology of extracellular diatom chitin nanofibers................................................. 131 7.3 Cellular and molecular processes for chitin biosynthesis and nanofiber formation............................................................................................................ 133 7.3.1 Silica biomineralization and frustule cell wall formation................................ 134 7.3.2 Photosynthetic carbon and nitrogen assimilation............................................. 135 7.3.3 Chitin biosynthesis................................................................................................. 136 7.3.4 Chitin nanofiber extrusion.................................................................................... 137 7.3.5 Molecular genetic basis of chitin biosynthesis and degradation in diatoms.......................................................................................... 137 7.3.6 Ecological role of extracellular diatom chitin nanofibers................................. 139 7.4 Bioreactor production of chitin nanofibers..................................................................... 140 7.5 Emerging applications of diatom-derived β-chitin nanofibers.................................... 143 7.5.1 Unique features of β-chitin nanofibers produced by diatoms......................... 143 7.5.2 Biomedical materials derived from β-chitin nanofibers................................... 144 7.5.3 Potential of β-chitin nanofibers as nanowires for protonic devices................ 144 7.6 Future research................................................................................................................... 146 Concluding remarks.................................................................................................................... 146 Acknowledgments....................................................................................................................... 147 References...................................................................................................................................... 147

7.1 Introduction Diatoms are single-celled algae that make cell walls of biogenic silica called frustules that possess intricate pore arrays patterned at the submicron and nanoscale. Consequently, diatoms require dissolved silicon as a required substrate for cell wall biosynthesis and cell division (Martin-Jézéquel et al. 2000). In addition to silicon metabolism, diatoms possess unique carbon-partitioning pathways (Hockin et al. 2012; Kroth et al. 2008; Obata et al. 2013; Smith et al. 2012, 2016), and form various carbohydrates (Gügi et al. 2015), including fibrous materials composed of the biopolymer known as chitin. Chitin is a linear polymer of anhydro-N-acetylglucosamine molecules joined by β-1,4-glycosidic linkages. Through the β-1,4-linkage, each monomer unit is rotated 180° around the axis of the polymer backbone chain relative to the last repeat unit, with the reducing end at carbon C1. Structures of chitin and its derivatives are presented in Figure 7.1. 129

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Enzymatic Technologies for Marine Polysaccharides CH3

Chitin OH C-1 O HO

O O

C-4

NH

O HO

O

NH

OH

O

OH

OH

NH O

O HO

N-acetyl glucosamine

O

OH NH

hydrolysis

O CH3

O

HO HO

O CH3

CH3

deacetylation OH O HO

O

OH

NH2 O HO

O O

NH2

OH

Chitosan

OH O

HO NH2

O

HO HO

O OH NH2

Glucosamine

Figure 7.1  Structures of chitin, chitosan, N-acetylglucosamine, and glucosamine. Dashed-line box denotes the N-acetyl group of chitin.

Chitin can be deacylated to chitosan, or hydrolyzed to N-acetylglucosamine and then deacylated to glucosamine, a well-known nutraceutical. These materials have a bewildering array of applications that are beyond the scope of this chapter. Chitin-derived nanofibers have been the subject of several recent reviews, particularly with respect to their biomedical material applications (Azuma et al. 2014; Ding et al. 2014; Elsabee et al. 2012; Habibi and Lucia 2012; Jayakumar et al. 2010; Muzzarelli et al. 2014; Rolandi and Rolandi 2014). Chitin is the second most abundant biopolymer in nature after cellulose, and is produced by a variety of organisms, including the exoskeletons of arthropods of terrestrial and marine origin (Younes and Rinaudo 2015; Merzendorfer 2011), as well as the cell wall matrix of fungi and yeast (Bowman and Free 2006). The linear polymer chains of chitin are hydrogen-bonded into a para-crystalline structure similar to that of crystalline cellulose. Chitin isolated from these natural sources is in the alpha form (α-chitin), where chains β-1,4 monomer assemble into an antiparallel array, with lateral chains alternatively aligning at the reducing end (C1) and nonreducing end (C4). In this antiparallel arrangement, α-chitin possesses both intramolecular hydrogen bonds between adjacent chains, and intermolecular hydrogen bonds between monomer units within a given linear chain. A much rarer chitin allomorph is β-chitin associated with proteins in squid pens (Cuong et al. 2016; Nagahama et al. 2008), the tube-like structures of pogonophoran and vestimentiferan worms found in the deep sea (Blackwell 1969), and the centric diatoms within order Thalassiosirales. In β-chitin, β-1,4-linked anhydro-N-acetyl glucosamine chains assemble together in a parallel array, with all polymer chains terminating at the reducing end (Blackwell 1969; Imai et al. 2003). β-Chitin possesses intrachain and intrasheet hydrogen bonds, but it is not known to possess intersheet hydrogen bonds between adjacent chains. The β-chitin allomorph has never been recrystallized from α-chitin in vitro, but naturally sourced β-chitin can be recrystallized to α-chitin by NaOH treatment (Noishiki et al. 2003). Diatoms are known to produce only β-chitin. Furthermore, the extracellular extrusion of unique β-chitin nanofibers is known to occur in only two genera of centric diatoms within order Thalassiosirales, namely, Cyclotella and Thalassiosira (McLachlan et al. 1965; Dweltz et al. 1968; Blackwell et al. 1967; Herth 1979; Herth and Barthlott 1979; Herth and Zugenmaier 1977).

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This includes three species within Cyclotella (C. cryptica, C. nana, C. meneghiniana) and seven species within Thalassiosira (T. pseudonana, T. fluviatilis, T. guillardii, T. minuscula, T. oceanica, T. punctigura, T. weissflogii). Most of the initial discoveries of diatom chitin were made over 40 years ago, but their interest as an advanced nanomaterial is fairly recent. Despite the ubiquitous nature of chitin, β-chitin from these diatoms is special, because the β-chitin allomorph itself is rare, and because the β-chitin is extruded from the cell as a pure, uniform nanofiber that is readily isolated from the organism. In contrast, chitin sourced from other organisms is usually associated with other carbohydrates, proteins, and minerals, and must be refined (Aranaz et al. 2009; Ifuku and Saimoto 2012; Younes and Rinaudo 2015). Diatom-derived β-chitin nanofibers may have future utility as advanced biomaterials for a variety of applications, given their unique morphology, crystalline structure, and purity. This chapter overviews the biosynthetic and cellular processes for β-chitin nanofiber formation by the centric diatoms Cyclotella and Thalassiosira. The presentation integrates atomic, molecular, cellular, and bioprocess scales to provide a comprehensive perspective on the unique features of β-chitin nanofibers extruded from diatoms. The chapter also highlights the key enzymes involved in diatom chitin metabolism, complimentary to the overall theme of enzymatic technologies for marine polysaccharides. In this chapter, four topic areas are highlighted: (1) the morphology of extracellular chitin nanofibers, (2) the cellular and molecular processes involved in β-chitin biosynthesis and nanofiber formation, (3) the bioreactor production of diatom-derived β-chitin nanofibers, and (4) the potential of these nanofibers for advanced biomedical and bioelectronic material applications. The chapter concludes with suggested avenues for future research.

7.2  Morphology of extracellular diatom chitin nanofibers Native chitin nanofibers from diatoms can be observed by light microscopy under differential interference (Nomarksi) contrast, but elucidation of morphological details requires scanning electron microscopy (SEM). Representative SEM images of the centric diatoms Cyclotella cryptica and Thalassiosira pseudonana are presented in Figure 7.2, and representative SEM images of chitin nanofibers emanating from the surface of C. cryptica are presented in Figure 7.3. In Figures 7.2 and 7.3, the SEM images were obtained from whole cells following fixation with glutaraldehyde and critical point drying (Chiriboga and Rorrer 2017). C. cryptica is about 8–10 μm in nominal size, whereas T. pseudonana is smaller, at 4–5 μm. The biosilica portion of cell wall (frustule) of a centric diatom contains two thecae per cell, aligned in upper and lower sections that fit together. Each theca contains a valve that forms the top and bottom faces of the cell wall, and a girdle band encircling the valve, which continues to form after the cell division. Lining the rim of each valve are larger pores called fultoportulae. The number of fultoportulae range from about 15 (T. pseudonana) to 20 (C. cryptica). Chitin nanofiber production is induced at the point of silicon starvation (Chiriboga and Rorrer 2017). At silicon starvation, a multitude of fibers emanate from each cell (Figure 7.3A,B). However, only one nanofiber is extruded through each pore (Figure 7.3C). Individual fibers are approximately 50 nm in diameter (Figure 7.3D) and can measure up to 100 μm in length with an aspect ratio of 2000. Ultrastructural analyses of diatom fultoportulae within the order Thalassiosirales reveal that these pores can possess nozzle-like features (Kaczmarska et al. 2005), which are clearly evident in Figures 7.2 and 7.3. It is not known whether the fultoportula diameter determines the nanofiber diameter, but it is interesting to note that the number of fultoportulae lining the rim of the T. weissfogii valve face is proportional to the valve diameter (Johansen and Theriot 1987).

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Enzymatic Technologies for Marine Polysaccharides (A)

(C)

2 mm (B)

2 mm (D)

2 mm

2 mm

Figure 7.2 SEM images of centric diatoms Cyclotella cryptica and Thalassiosira pseudonana: (B,D) Cyclotella cryptica; (A,C) Thalassiosira pseudonana. (Unpublished images provided by the author.)

(A)

(D) 47 nm 55 nm

10 mm (B)

5 mm

1 mm (C)

2 mm

Figure 7.3  SEM image chitin nanofibers from Cyclotella cryptica: (A) network of fibers from two diatoms; (B,C) fibers extruding from valve face, one fiber per fultoportula; (D) individual fibers of approximately 50 nm diameter. (Unpublished images provided by the author.)

Chapter seven:  Biosynthesis and extrusion of β-chitin nanofibers by diatoms a-b plane

133 b-c plane

b

b

a

c

c = 10.38Å

γ = 97.5°

a = 4.85Å

b = 9.26Å

(A)

(B)

Figure 7.4  Molecular structures of paracrystalline β-chitin: (A) monoclinic unit cell; (B) molecular models at 1 Å resolution for a–b and b–c planes obtained from X-ray crystallography data for chitin nanofibers of the diatom Thalassiosira weissflogii (Nishiyama et al. 2011). Dotted lines show intramolecular hydrogen bonds; green denotes carbon (C), blue denotes nitrogen (N), and red denotes oxygen (O) atoms. [Reprinted with permission from Macromolecules, Vol. 44, Y. Nishiyama, Y. Noishiki, and M. Wada, “X-ray Structure of Anhydrous β-Chitin at 1 Å Resolution,” pages 950–957. Copyright (2011) American Chemical Society.]

Extracellular nanofibers extruded from diatoms within Cyclotella and Thalassiosira consist of pure β-chitin. The crystal structure of β-chitin from these diatoms has been deduced by X-ray diffraction (Dweltz et al. 1968; Blackwell et al. 1967; Imai et al. 2003; Nishiyama et al. 2011; Ogawa et al. 2011). Complementary structural analyses have included sum-frequency generation (Ogawa et al. 2016) and computational modeling (Faria et al. 2016). For β-chitin isolated from the centric diatoms Thalassiosira fluviatilis and Cyclotella cryptica, the unit cell of chitin in anhydrous form is monoclinic (space group P21) with a = 4.85 Å, b = 9.26 Å, and c = 10.38 Å (monomer unit repeat length) and angle of γ = 97.5° (Blackwell et al. 1967). Molecular models of the para-crystalline hydrogen bonding in diatom-derived β-chitin are presented in Figure 7.4. The carbonyl oxygen of the N-acetyl group formed intermolecular hydrogen bonds between the primary hydroxyl and amine groups, resulting in a twodimensional hydrogen bonding network in the plane perpendicular to the pyranosyl ring plane (Nishiyama et al. 2011). Given the unit cell size overall fiber diameter of approximately 50 nm, and the a–b planar cross section dimensions of 4.85 × 9.26 Å, approximately 4400 linear poly-N-acetylglucosamine molecules are packed together in a given fiber cross section. The degree of polymerization of β-chitin chains within the diatom chitin nanofiber is not known. However, the β-chitin isolated from squid pens has average molecular weight of 13.4 · 105 g/mol, and average degree of polymerization of 6865 (Fiamingo et al. 2016).

7.3 Cellular and molecular processes for chitin biosynthesis and nanofiber formation A simplified scheme for diatom cell division and chitin nanofiber formation is shown in Figure 7.5, and key metabolic pathways are sketched out in Figure 7.6. Our research has shown that chitin production in Cyclotella cryptica occurs just after cell division, but is elicited only when the diatom cell is in a silicon-starved state (Chiriboga and Rorrer 2017). Dissolved silicon is a required substrate for biosilica frustule biosynthesis and

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valve

upper thecae

girdle band

upper thecae

SDV SDV

new chitin nanofiber formation SiT

Si(OH)4 Si Uptake

Si Starvation & Final Cell Division

Figure 7.5 Simplified scheme for silicon uptake, cell division, and chitin nanofiber formation in Cyclotella and Thalassiosira. [Abbreviations: SDV = silica deposition vesicle; Si(OH)4 = silicic acid (soluble form of Si); SiT = silicon transporter.]

subsequent diatom cell division. Therefore, chitin production is tied to silica biomineralization, photosynthetic carbon assimilation, and carbohydrate biosynthesis pathways in the diatoms Cyclotella and Thalassiorsira. Key cellular and metabolic processes for silica biomineralization, photosynthetic carbon accumulation, and chitin production are reviewed in the paragraphs that follow.

7.3.1  Silica biomineralization and frustule cell wall formation The processes of silica biomineralization in diatoms are described in several review papers that provide valuable insights on silicon uptake into the diatom cell, biosilica assembly into complex nanoporous structures, and the genomic underpinnings of these processes (Martin-Jézéquel et al. 2000; Hildebrand 2008; Kröger and Poulsen 2008; Brunner et al. 2009a; Hildebrand and Lerch 2015). Biosynthesis of the diatom silica cell wall occurs just prior to cell division through the uptake of soluble silicon in the form of Si(OH)4 (silicic acid), and its subsequent condensation into biosilica within the silica deposition vesicle (SDV). The Si(OH)4 enters the cell through a silicon transport (SiT) channel in the cell membrane and then diffuses to the SDV. The SDV forms at the plane of cell division on each side of the dividing cell. The conversion of Si(OH)4 to SiO2 is mediated by “silaffin” proteins within the SDV. Within the SDV, the deposition process is also templated to self-assemble the nanostructured silica precursors into cell wall structures that possess intricate pore arrays. On cell division, each daughter cell possesses a new theca formed at the plane of division, and the old thecae carried over from the parent cell.

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light H2O Pyruvate AcetylCoA

Photosynthetic

PSII

Apparatus

PSI

G3P

e–

NADPH+ATP

O2

Fruc-6-P Calvin Cycle

Gluc-6-P

CO2

glutamate UDP-NAGlc synthesis

GS/ GOGAT glutamine

chitin nanofiber

chitin synthesis

NR NH4+

NO2–

NO3–

Figure 7.6 Simplified scheme of major pathways for photosynthetic carbon assimilation from CO2, nitrogen assimilation, and chitin biosynthesis in Cyclotella cryptica. Silicon metabolism not shown. (Abbreviations: Fruc-6-P = fructose 6-phosphate; G3P = glycerate 3-phosphate; Gluc-6-P = glucose-6-phosphate; GS/GOGAT = glutamine synthetase/glutamine oxoglutarate aminotransferase; NR = nitroreductase; PSI = photosystem I; PSII = photosystem II; UDP = uridine diphosphate.)

7.3.2  Photosynthetic carbon and nitrogen assimilation The silicon metabolism processes described above are downstream of photosynthetic processes in the diatom cell (Huysman et al. 2014). The processes of photosynthetic carbon assimilation and primary metabolic processes in diatoms, and their interactions with silicon metabolism, are well described (Kroth et al. 2008; Smith et al. 2012, 2016; Obata et al. 2013; Traller et al. 2016). In Cyclotella and Thalassiorsira, the major secondary metabolic sinks for carbon are lipids, β-1,3-glucan (chrysolaminarin), and chitin, whereas the major sinks for nitrogen are proteins and chitin. A simplified schematic highlighting the major biosynthetic pathways associated with photosynthetic carbon assimilation, nitrogen assimilation, and chitin biosynthesis is provided in Figure 7.6. The photosynthetic reduction of CO2 to the pyruvate hub provides the intermediates for gluconeogenesis and carbohydrate polymer biosynthesis, leading to the intracellular accumulation of chrysolaminarin and the extracellular extrusion of chitin nanofibers. Carbon flux to fatty-acid biosynthesis competes with carbon flux to carbohydrate biosynthesis. Reducing power provided by photosynthesis also drives the reduction of nitrate (NO3−) to ammonium (NH4+) for assimilation into the glutamine synthase/ glutamine oxoglutarate aminotransferase (GS/GOGAT) pathway for amino acid biosynthesis. The amino acid glutamine, which is produced through this pathway, is used in the biosynthesis of UDP-N-acetylglucosamine, a key intermediate in chitin biosynthesis. Detailed pathways for amino acid biosynthesis in diatoms are provided by Bromke (2013).

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Although cellular compartmentalization is not shown in Figure 7.6, carbon fixation (Calvin cycle), nitrate reduction, the GS/GOGAT cycle, and fatty-acid biosynthesis occur within the chloroplast and periplastid space, whereas the TCA cycle occurs within the mitochondria. In T. pseudonana, two glutamine synthase isoforms employing the GS/ GOGAT cycle are expressed in the plastid and cytosol (Bromke 2013). Gluconeogenesis occurs in the cytosol. Downstream of gluconeogenesis, the carbohydrate biopolymer β-1,3-glucan (chrysolaminarin) is biosynthesized in the cytosol and then sequestered within the vacuole, whereas β-chitin is bioysnthesized in a secretion vesicle on the plasma membrane and then exported out of the cell.

7.3.3  Chitin biosynthesis Chitin biosynthesis and nanofiber self-assembly in the diatoms Cyclotella and Thalassiorsira are presumed to possess biochemical pathways and cellular processes consistent with those in other organisms that produce extracellular chitin (Merzendorfer 2011; Sugiyama et al. 1999). The core biosynthetic pathway for β-1,4-linked N-acetylglucosamine biopolymer formation consists of the following five main steps (shown in Figure 7.7): 1. Fructose-6-P in the intracellular pool is converted to glucosamine-6-P via aminotransferase (EC 2.6.1.16) using l-glutamine, which is supplied from the GS/GOGAT pathway (see Figure 7.6). 2. Glucosamine-6-P is converted to N-acetylglucosamine-6-P via N-acetyltransferase (EC 2.3.1.4) using acetyl co-A (coenzyme A) as the cosubstrate. H3C

chitin

OH O HO

O

OH O

NH

O

OH

NH

O OH

OH

O

O

OH O

O

O

HO

NH

OH

OH

O

O

UDP

O

OH

CH3

O

O

O

O

HO

O NH O CH3

P OH

O

P

O

N

O

OH

HO

UDP-N-Acetyl glucosamine

O

glucosamine-6-P O OH

NH

HO

CH3

N-acetyl transferase

O

NH2 Acetyl-Co-A Co-A

HO

pyrophosphorylase

OH

O

HO O

PPi UTP

OH

P

O

O

OH

OH OH

O

OH NH

OH

HO P

O

chitin synthase O

OH

glutamate OH

aminotransferase

OH

NH O

OH glutamine

HO HO

H3C

OH

HO

O

CH3

UDP O

O

P

HO

NH

O

OH

CH3

HO O

O

fructose-6-P

O

OH mutase P

NH

OH O

OH

O OH NH

O

O CH3

N-Acetyl-glucosamine-1-P

Figure 7.7  Chitin biosynthesis pathway, starting from frustose-6-P.

CH3

N-Acetyl-glucosamine-6-P

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137

3. The phosphate group on N-acetylglucosamine-6-P is changed from the 6-P to the 1-P position by phosphoacetylglucosamine mutase (EC 5.4.2.3). 4. N-Acetylglucosamine-1-P is converted to uridine diphosphate (UDP)-Nacetylglucosamine via pyrophosphorylase (EC 2.7.7.23) using uridine triphosphate (UTP) as the cosubstrate. 5. Chitin synthase (EC 2.4.1.16) catalyzes β-1,4-glycosidic bond formation between UDPN-acetylglucosamine and (N-acetylglucosamine)n at the nonreducing end of the chain, releasing UDP and adding one monomer unit to the nonreducing end of the poly-N-acetylglucosamine chain. Molecular studies have confirmed the genes encoding for all of these enzymes in Cyclotella cryptica (Traller et al. 2016). The production of chitin fibers from both Thalassiosira fluviatilis and C. cryptica were blocked by polyoxin D, a UDP-N-acetylglucosamine analog (Morin et al. 1986), providing early evidence of active chitin synthase in these diatoms.

7.3.4  Chitin nanofiber extrusion The true mechanism of β-chitin nanofiber extrusion by living cells of Cyclotella and Thalassiosira is still under study. It is believed that β-chitin crystallizes at the interface between the cell membrane and the exterior environment by a simultaneous polymerization and crystallization process with the chains aligned in a parallel array. In contrast, for α-chitin assembly in arthropods, polymerization occurs within a vesicle to form a so-called liquid crystalline phase that facilitates antiparallel chain assembly. This vesicle fuses to the plasma membrane and then extrudes α-chitin (Sugiyama et al. 1999). In both α-chitin and β-chitin biosynthesis, chitin synthase is bound to the vesicle membrane. But synthesis of β-chitin is simpler because both polymerization and self-assembly are localized to the plasma membrane. Transmission electron microscope (TEM) analysis of stained cross sections from C. cryptica revealed that the chitin fiber originated at a conical invagination of the plasma membrane just below the fultoportulae (Herth 1979). Herth (1979) asserted that chitin biosynthesis occurred at this conical invagination, which presumably contained membrane-bound chitin synthase. Following the discussion above, the extracellular extrusion of β-chitin nanofibers by Cyclotella and Thalassiorsira is conceptually illustrated in Figure 7.8. Chitin synthase bound to the plasma membrane surrounding the vesicle catalyzes the stepwise polymerization of UDP-N-acetylglucosamine at the nonreducing end of the molecule form β-1,4-linked linear chains, which subsequently undergo intramolecular hydrogen bonding. The reducing end is at the apical tip of the fiber. The pore lumen guides the unidirectional, lateral association and alignment of linear chains as they assemble into the nanofiber. The structure is stabilized in all directions by van der Waals interactions and hydrophobic effects, but as stated earlier, β-chitin has no intermolecular hydrogen bonds between adjacent lateral chains. Therefore, it seems that β-chitin is the only allomorph of chitin that this elegant yet simple “molecular extruder” can accommodate. In higher organisms, α-chitin, with intermolecular bonds between adjacent chitin chains, would be needed for biosynthesis and self-assembly of more complex scaffolded materials.

7.3.5  Molecular genetic basis of chitin biosynthesis and degradation in diatoms Genes encoding for the expression of key enzymes involved in chitin biosynthesis have been identified in Cyclotella cryptica (Traller et al. 2016). The number of enzyme isoforms and their putative locations within the cell, taken from the nuclear genome sequencing

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Enzymatic Technologies for Marine Polysaccharides

fultoportula frustule

plasma membrane UDP-NAGlc UDP

chitin synthase conical invagination

Figure 7.8  Conceptual model of cellular apparatus for polymerization and extracellular extrusion of chitin nanofibers in Cyclotella.

analysis of Traller et al. (2016), are provided in Figure 7.9. From this analysis, the conversion of fructose-6-P to glucosamine-6-P occurs within the mitochondrion, although molecular studies with T. pseudonana suggest that glutamine synthase is expressed within the plasid and cytosol (Bromke 2013). Subsequent conversion steps to UDP-N-acetylglucosamine occur in within cytosol. It is suggested that this chitin synthase may be associated with the plasma membrane or the silica deposition vesicle (Traller et al. 2016). Presumably, glucosamine 6-P must move from the mitochondrion to the cytosol for conversion to UDP-Nacetylglucosamine. The UDP-N-acetylglucosamine then moves to chitin synthase in the membrane of the putative chitin fiber secretion vesicle, where it is ultimately processed into chitin nanofibers and extruded outside of the cell.

glucosamine fructose-6-P aminotransferase

mitochondrion cytosol ER unknown

glucosamine-6-P N-acetyltransferase phosphoacetylglucosamine mutase UDP-N-acetylglucosamine pyrophosphorolase chitin synthase 0

1

2

3 4 5 Number of Genes

6

7

8

Figure 7.9  Genes and their predicted subcellular localization putatively identified for chitin biosynthesis in Cyclotella cryptica. [Data obtained from Traller et al. (2016).]

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Following the availability of the whole-genome sequence for Thalassiosira pseudonana (Armbrust et al. 2004), subsequent genomic analyses have revealed broad and complex roles for chitin-related processes in diatoms, which includes genes that encode for expression of multiple chitin synthases, chitinases, and proteins with chitin-binding domains. Protein domain predictions and differential gene expression patterns provide evidence that chitin synthases have multiple functions within a diatom cell. T. pseudonana possesses six genes encoding three types of chitin synthases (Durkin et al. 2009). Cyclotella cryptica also has six genes that encode for chitin synthases associated with cytosol and the ER that are unique for only the Thalassiosirales diatoms (Traller et al. 2016). In T. pseudonana, transcript abundance of the gene encoding for one of these chitin synthase types increases when cells resume division after short-term silicic acid starvation, and during short-term limitation by silicic acid (Durkin et al. 2009). A common set of 84 genes were induced by both silicon and iron limitations (Mock et al. 2008). Interestingly, all the genes necessary for chitin synthesis found in the centric diatom T. pseudonana were also found in the pennate diatom Phaeodactylum tricornutum, despite the fact that P. tricornutum does not produce extracellular chitin fibers (Durkin et al. 2009). This suggests that chitin biosynthesis may also be internal to the diatom cell wall. Brunner et al. (2009b) showed that cell walls of T. pseudonana contained a network-like, chitin-based scaffold that resembled the size and shape of the biosilica. These scaffolds consisted of interconnected fibers with an average diameter of about 25 nm that contained other as-yet unknown biomolecules apart from chitin, and were thought to play a role in biosynthesis of the silica frustule. However, transcriptome analysis of the silicon response of the diatom T. pseudonana suggested that chitin synthase genes were not upregulated during valve synthesis, and so their role in valve formation remains unclear (Shrestha et al. 2012). Chitinases promote the degradation of chitin by hydrolytic cleavage of the β-1,4 linkage, and have two primary categories: (1) endochitinases (EC 3.2.1.14), which cleave internal linkages, primarily down to chitobiose (N,N’-diacetylchitobiose); and (2) β-1,4-Nacetylglucosaminidases (EC 3.2.1.30), which cleave soluble chitin oligomers (chitobiose, chitotriose, and chitotetraose) at the nonreducing end into N-acetylglucosamine monomer (Patil et al. 2000). Cyclotella cryptica has a nearly complete chitin degradation pathway with 22 chitinases (Traller et al. 2016), and T. pseudonana has at least 20 genes that encode putative chitin-degrading chitinases (Durkin et al. 2009). Most of the chitinases are shared within the Thalassiosirales group and have similarity to either bacterial or fungal chitinases. A detailed description of diatom chitinases is beyond the scope of this chapter. In insects, chitin functions as a scaffold material, and so growth and morphogenesis are strictly dependent on the coordination of chitin synthesis and its degradation (Dahiya 2009). Furthermore, in both fungi and insects, chitin biosynthesis is tightly regulated at the transcriptional level during growth and development (Merzendorfer 2011). In diatoms, since the β-chitin nanofiber is exported from the cell, it is not likely that there is a strong regulatory loop for coordinated action of chitin synthase and chitinase to tailor net production and final morphology. However, if the fiber gets too long, it is speculated that chitinases could trim the fiber length by breaking off the fiber at the point where the fiber exits the cell boundary within the fultoportula. This scenario suggests an ecological role, as suggested below.

7.3.6  Ecological role of extracellular diatom chitin nanofibers From an ecological perspective, the reason for extrusion of chitin nanofibers from the Thalassiosira and Cyclotella frustule surface is unknown. On the basis of laboratory experiments, it is speculated that chitin fibers play an important role in cell separation and cell

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buoyancy. Thalassiosira and Cyclotella diatom cells without chitin fibers sedimented in flask culture, whereas diatom cells with fibers did not (Gooday et al. 1985; Morin et al. 1986). The chitin fibers also facilitated the formation of diatom cell chains, which also increased buoyancy. The removal of chitin fibers from the diatom Thalassiosira fluviatilis by treatment of chitinase sank at rate of more than 1.7 times faster than control cells with attached fibers (Walsby and Xypolyta 1977). These studies suggest that chitin nanofibers may enable these diatoms to move up and down the water column to access light near the surface and nutrients in deeper waters. This ability would provide a competitive advantage relative to other algae with respect to light and nutrient access. From the discussion above, why would the diatoms Thalassiosira and Cyclotella make β-chitin nanofibers in the first place? Unlike single-celled green algae, these diatoms lack the biochemical machinery to store photosynthetic carbon as starch for later conversion to energy. These diatoms do store carbohydrates as β-1,3-glucan, but have also developed an alternative mechanism to export excess photosynthetic carbon as β-chitin without the need for intracellular storage. This process could kick in at silicon limitation, because without silicon, the cells can no longer divide, but remain photosynthetically active. To support this point, previous work has demonstrated that chitin biosynthesis has lower light requirements than lipid biosynthesis (Ozkan and Rorrer 2017a). Therefore, at low light levels deeper in the water column, chitin nanofiber formation could still occur and provide the means to increase the buoyancy of the cell.

7.4  Bioreactor production of chitin nanofibers The scalable and controlled production of extracellular chitin nanofibers cultivation of the diatom Cyclotella has been demonstrated using a bubble-column photobioreactor (Rorrer et al. 2016). The bioreactor provides controlled CO2, light, and nutrient delivery to the diatom cells. In a bubble-columm photobioreactor, the introduction of aeration gas containing CO2 to the bottom of a cylindrical, glass vessel creates rising gas bubbles that provide CO2/O2 gas exchange and mixing of the cell suspension. Light is uniformly delivered to the cell suspension by an array of lamps surrounding the vessel. Medium pumps provide dissolved silicon, nitrate, and nutrients to the vessel. Representative data for the batch cultivation of centric diatom Cyclotella cryptica (UTEX 1269, Cyclotella spp.) are provided in Figure 7.10. Methods are described in our previous work (Ozkan and Rorrer 2017a), with chitin assayed as glucosamine after concentrated acid hydrolysis of whole cell mass. In batch cultivation, dissolved silicon, nitrate, and nutrients were added at the beginning of the process, and cell division proceeded until the cell suspension consumed all of the silicon from the cultivation medium (Figure 7.10A). When all of the silicon was consumed from the medium, then the cells were considered to be in the silicon-starved state, and cell division ceased unless more silicon was added. As shown in Figure 7.10A, the silicon starvation occurred after about 100 hours of cultivation. Although all of the silicon was taken up by the cells, it was not yet processed into frustule biosilica, and so the cells divided one more time, from a cell number density of approximately 1.5 · 106 cells/mL to 3.0 · 106 cells/mL. At the onset of silicon limitation, chitin production began (Figure 7.10B), and the chitin yield per cell dramatically increased at the onset of silicon limitation (Figure 7.10C). Nitrate is also a required substrate for chitin production. Chitin production ended when cell division was complete, and all the nitrate was consumed from the medium. The initial silicon and nitrate concentrations were 0.8 mM and 2.2 mM respectively, for complete consumption of both nutrients; the final chitin concentration was 68 mg/L; the final cell number

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cells Si N

Nutrient conc. (mmol/L)

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Figure 7.10  Batch cultivation of Cyclotella cryptica in a bubble-column photobioreactor for production of chitin nanofibers: (A) cell number density, Si concentration, and nitrate concentration versus time; (B) chitin concentration in bulk cell suspension versus time; (C) chitin yield (mg chitin/109 cells); (D) calculated fiber length and fiber formation rate. Cultivation conditions: total volume 4.0 L, aeration rate (∼400 ppm CO2) 0.47 L air/L • culture • minute, incident light intensity to vessel surface 90 μE/m2 • second (14 hours light/10 hours dark photoperiod), temperature 22°C. Initial silicon and nitrate concentrations were 0.8 mM and 2.2 mM, respectively.

density was 3.0 · 106 cells/mL; and the final chitin yield was 21 mg chitin per 109 cells. About 10% of the nitrogen fed to the cell suspension culture as nitrate found its way into the final chitin product. From the chitin yield, the fiber production rate and cumulative fiber length for a 50-nm-diameter fiber were calculated, assuming 20 fultoportulae per valve with two valves per cell, and the solid chitin density was 1.49 g/cm3 (Dweltz et al. 1968). The rate of fiber extrusion (μm/hour) and cumulative fiber length–time profiles are presented in Figure 7.10D. According to this analysis, the peak fiber formation rate was 2.2 μm/hour, which occurred about 24 hours after silicon starvation. This fiber production rate corresponded to a turnover rate of 5500 β-1,4 linkages formed per secretion vesicle per second. As described above, chitin production occurs after cell division and silicon limitation. Therefore, to enable sustained chitin production, cell division must also be maintained, but in a silicon-limited state. Toward this end, controlled feeding of dissolved silicon to the diatom cell suspension at a rate lower than the intrinsic Si uptake rate needed for Si-replete cell division puts the diatom cell division cycle in a continuously silicon-limited state. In this state, the cell division rate followed the silicon delivery rate. When the silicon

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delivery rate to the diatom cell suspension was constant, the cell number production, and the chitin production per unit volume of culture, were both linear with time (Chiriboga and Rorrer 2017). Multistage addition of dissolved silicon along with nitrate, the external nitrogen source for chitin, enabled sustained linear production of cell mass and chitin at a constant rate of 44 mg chitin/L daily to concentration in the culture broth approaching 700 mg chitin/L, where 19 wt% of the total biomass was chitin (Ozkan and Rorrer 2017b). The optimal nitrogen/silicon ratio for maximum chitin yield per cell was 4 mol N/mol Si, and 10% of nitrate nitrogen fed was incorporated into the final chitin product (Chiriboga and Rorrer 2018). Rates of photosynthetic carbon accumulation and lipid production were strongly dependent on light intensity and CO2 partial pressure in the aeration gas. However, rate of chitin production and the final chitin yield per cell were not (Ozkan and Rorrer 2017a,c). Consequently, control of chitin production is accomplished primarily through controlled feeding of silicon and nitrate to the diatom cell suspension under conditions where light and CO2 delivery are not limiting photosynthetic biomass production. A physical separation process was also developed to isolate and purify chitin nanofibers from the diatom cells (Figure 7.11). Fibers were dislodged from the cells by high-speed blending for 10 seconds. The blended suspension was centrifuged under low speed (1500g) for short duration of 1.0 minutes. Chitin nanofibers collected in the supernatant, whereas the cell debris from high-speed blending were collected as a solid. The supernatant was centrifuged under high speed (11,000g) for 30 minutes. The resulting pellet, rich in diatom chitin, was treated with 1.0 M HCl at 70°C for 30 minutes under stirring to dissolve residual calcium carbonate salts, and then treated with 0.5 wt% sodium dodecyl sulfate (SDS) to remove any proteins adsorbed on the chitin fibers. The treated suspension was passed through a polytetrafluoroethylene (PTFE) filter of 1.0 μm pore size, and the fiber mass was washed with distilled water. The washed fiber mass was then treated 95% ethanol at room temperature for 30 minutes to remove any residual organic impurities, rinsed again in ethanol, and then dried in air at 45°C for 4 hours. Chitin recovery after the low-speed centrifugation and high-speed centrifugation averaged 97% and 92%, respectively, based on the total chitin initially present in the cell mass (Chotyakul 2014). diatom cell suspension

debris

waste liquid

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Drying Filter Cake

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EtOH Washing & Filtration

filtrate

High-Speed Centrifugation

H2O Washing & Filtration

filtrate

chitin-rich pellet

HCI & SDS Treatment

filtrate

Figure 7.11 Flowchart for isolation of chitin nanofibers from bioreactor-cultured Cyclotella cryptica cells.

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7.5  E  merging applications of diatom-derived β-chitin nanofibers Chitin nanofibers have a variety of emerging advanced material and biomedical applications (Ding et al. 2014; Elsabee et al. 2012; Habibi and Lucia 2012; Ifuku and Saimoto 2012; Muzzarelli et al. 2014; Jayakumar et al. 2010; Azuma et al. 2014). This section summarizes the unique features of diatom-derived β-chitin nanofibers from an applications perspective. A comprehensive review on the myriad of applications for chitin and chitosan is beyond the scope of this presentation. However, two emerging material applications for β-chitin nanofibers—nanostructured tissue engineering scaffold materials and protonic device components—provide inspirational examples for future development.

7.5.1  Unique features of β-chitin nanofibers produced by diatoms Table 7.1 summarizes the key differences between chitin sourced from macrophytic marine organisms and single-celled diatom algae, drawn from information mentioned throughout this chapter. Chitin nanofibers being developed for advanced material and biomedical applications are typically isolated shellfish waste, which has many contaminants, including proteins and minerals (Younes and Rinaudo 2015; Aranaz et al. 2009). Crustacean shells typically consist of 30–40% proteins, 30–50% calcium carbonate, and 20–30% chitin, as well as residual lipophilic pigments (Aranaz et al. 2009). Furthermore, only a fraction of this raw chitin can be purified into nanofibers. Chitin nanofibers isolated from this resource are not uniform with diameters ranging from 10 to 20 nm, and exist only in the α-chitin allomorph (Ifuku and Saimoto 2012). Nanofibers of α-chitin or β-chitin isolated by various methods from nondiatom sources ranged from 2 to 20 nm in diameter, with lengths usually less than 1 μm, but typically never greater than 10 μm (Muzzarelli et al. 2014). Chitin produced in cultured fungal cells is imbedded within the cell wall matrix, and constitutes up to 20 wt% of the cell wall fraction of the biomass, depending on the species (Bowman and Free 2006). This fungal chitin is intimately associated with other polysaccharides and proteins, and so would require both chemical and physical methods of isolation. Table 7.1  Comparison of chitin sourced from macrophytic marine organisms versus diatom algae Chitin source Feature

Macrophytic marine organisms

Diatom algae

Allomorphic forms Source

α-Chitin (dominant); β-chitin (rare) α-Chitin: Crustacean—waste from shellfish seafood processing β-Chitin: No readily available source, presently isolated from squid pens or tube worms from deep-sea vents Not pure—associated with other carbohydrates, proteins, and minerals Refining process involving impurity removal and isolation of nanofibers Short, ∼2-5-nm-diameter nanofibers of varying length cold HCl extract > hot H2SO4 extract > cold HCl. Rioux et al. (2010) also extracted crude laminarin and galactofucan from S. longicruris using a 1% (w/v) CaCl2 solution at 85°C for 4 hours. The pellets obtained after ethanol precipitation were hydrated and dialyzed, followed by ultrafiltration to remove galactofucan from the mixture. Je et al. (2009) used carbohydrate and protein-hydrolyzing enzymes for preparation of extracts from Undaria pinnatifida yielding a laminarin. The authors reported that enzymatic digestion of algae produced a high bioactive yield and exhibited enhanced biofunctional activity compared to the solvent extraction method.

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11.2.4 Alginate Alginate, sometimes shortened to “algin,” is present in the cell walls of brown seaweeds, and is partly responsible for the flexibility of the seaweed (McHugh 2003). A high-quality alginate forms strong gels and gives thick aqueous solutions. A good raw material for alginate extraction should also give a high yield of alginate. The brown seaweeds that fulfill these criteria are species of Ascophyllum (Ascophyllum nodosum), Durvillaea (Durvillaea potatorum), Ecklonia, Laminaria (L. digitata, L. japonica, L. hyperborean, and L. Saccharina), Lessonia (Lessonia nigrescens, Lessonia trabeculata), Macrocystis (Macrocystis pyrifera), and Sargassum (Sargassum flavicans), although the last, Sargassum, is used only when nothing else is available: its alginate content is usually borderline quality and the yield usually low. Alginate is the term generally used for the salts of alginic acid, but it can also refer to all the derivatives of alginic acid and to alginic acid itself. Alginate is present in the cell walls of brown algae as the calcium, magnesium, and sodium salts of alginic acid. The goal of the extraction process is to obtain dry, powdered, sodium alginate. The calcium and magnesium salts do not dissolve in water, but the sodium salt does. The rationale behind the extraction of alginate from the seaweed is to convert all the alginate salts to the sodium salt, dissolve this in water, and remove the seaweed residue by filtration. The alginate must then be recovered from the aqueous solution. The solution is very dilute, and evaporation of the water is not economic. There are two different ways to recover the alginate: 1. The first method consists in the addition of acid, which causes alginic acid to form; this does not dissolve in water, and the solid alginic acid is separated from the water. The alginic acid separates as a soft gel, and some of the water must be removed from this. After this has been done, alcohol is added to the alginic acid, followed by sodium carbonate, which converts the alginic acid into sodium alginate. The sodium alginate does not dissolve in the mixture of alcohol and water, so it can be separated from the mixture, dried, and milled to an appropriate particle size that depends on its particular application. 2. The second method consists in recovering the sodium alginate from the initial extraction solution by adding a calcium salt. This causes calcium alginate to form with a fibrous texture; it does not dissolve in water and can be separated from it. The separated calcium alginate is suspended in water, and acid is added to convert it into alginic acid. This fibrous alginic acid is easily separated, placed in a planetarytype mixer with alcohol, and sodium carbonate is gradually added to the paste until all the alginic acid is converted to sodium alginate. The paste of sodium alginate is sometimes extruded into pellets that are then dried and milled. The essentials of these two processes are illustrated in the flow diagram in Figure 11.7. The process appears to be straightforward, and certainly the chemistry is simple: (1) convert the insoluble alginate salts in the seaweed into soluble sodium alginate; (2) precipitate either alginic acid or calcium alginate from the extract solution of sodium alginate; and (3) convert either of these back to sodium alginate, this time in a mixture of alcohol and water, in which the sodium salt does not dissolve. The difficulties lie in handling the materials encountered in the process, and to understand these problems, a little more detailed knowledge of the process is required. To extract the alginate, the seaweed is broken into pieces and stirred with a hot solution of an alkali, usually sodium carbonate. Over a period of about 2 hours, the alginate dissolves as

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Enzymatic Technologies for Marine Polysaccharides Wet chopped seaweed Sodium carbonate Alkaline extract

Separation Add calcium carbonate

Sodium alginate

Calcium alginate fibre

Seaweeds

Add acid

Alginic acid

Add acid Alginic acid

Dewatering alginic acid Add sodium

Add sodium Sodium alginate

Sodium alginate

Figure 11.7  Flowchart for Sodium Alginate extraction.

sodium alginate to give very thick slurry. This slurry also contains the part of the seaweed that does not dissolve, mainly cellulose. This insoluble residue must be removed from the solution. The solution is too thick (viscous) to be filtered and must be diluted with a very large quantity of water. After dilution, the solution is forced through a filter cloth in a filter press. However, the pieces of undissolved residue are very fine and can quickly clog the filter cloth. Therefore, before filtration begins, a filter aid, such as diatomaceous earth, must be added; this holds most of the fine particles away from the surface of the filter cloth and facilitates filtration. However, filter aid is expensive and can significantly contribute to costs. To reduce the quantity of filter aid needed, some processors force air into the extract as it is being diluted with water (the extract and the diluting water are mixed in an inline mixer into which air is forced). Fine air bubbles attach themselves to the particles of residue. The diluted extract is left standing for several hours while the air rises to the top, taking the residue particles with it. This frothy mix of air and residue is removed from the top, and the solution is withdrawn from the bottom and pumped to the filter. The next step consists in precipitation of the alginate from the filtered solution, either as alginic acid or calcium alginate.

11.2.4.1  Alginic acid method When acid is added to the filtered extract, alginic acid forms in soft, gelatinous pieces that must be separated from the water. Again, flotation is often employed; filtration is not possible because of the soft jelly-like nature of the solid. If an excess of sodium carbonate is used in the original extraction, this will still be present in the filtered extract so that when acid is added, carbon dioxide will form. Fine bubbles of this gas attach themselves to the pieces of

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alginic acid and lift them to the surface where they can be continuously scraped away. The processor now has a jelly-like mass of alginic acid that actually contains only 1–2% alginic acid, with 98–99% water. Somehow, this water content must be reduced. It is too soft to allow the use of a screw press. Some processors place the gel in basket-type centrifuges lined with filter cloth. Centrifuging can increase the solid content to 7–8%, and this is sufficient if alcohol is to be used in the next step of converting it to sodium alginate. It is also now sufficiently firm to be squeezed in a screw press. The 7–8% alginic acid is placed in a mixer and alcohol (usually ethanol or isopropanol) is added to give a 50:50 mixture of alcohol and water. Then solid sodium carbonate is added gradually until the resulting paste reaches the desired pH. The paste of sodium alginate can be extruded as pellets, oven dried and milled.

11.2.4.2  Calcium alginate method When a soluble calcium salt, such as calcium chloride, is added to the filtered extract, solid calcium alginate is formed. If the calcium solution and filtered extract are mixed carefully, the calcium alginate can be formed as fibers; a bad mixing gives a gelatinous solid. This fibrous material can be readily separated on a metal screen (sieve) and washed with water to remove excess calcium. It is then stirred in dilute acid and converted to alginic acid, which retains the fibrous characteristics of the calcium alginate. This form of alginic acid can be easily squeezed in a screw press. A screw press with a graduated-pitch screw is generally used; the squeezing action must be applied very gradually; otherwise the material will just move backward and out of the press. The product from the screw press appears relatively solid but still contains only 20–25% alginic acid. However, it is dry enough to form a paste when sodium carbonate is mixed with it to convert it to sodium alginate. Sodium carbonate is added to the alginic acid in a suitable type of mixer until the required pH level is reached; then the paste is extruded as pellets, dried, and milled. The disadvantage of this second method, compared to the alginic acid method, is that an extra step is added to the process. The advantage is that handling of the fibrous calcium alginate and alginic acid is much simpler and alcohol is not needed. Alcohol is expensive, and while it is usually recovered and recycled, recovery is never 100%, so its use adds to the costs. Other important factors in alginate production are color control of the product, water supply, and waste disposal. If the original seaweed is highly colored (e.g., Ascophyllum), the alkaline extract will also be highly colored and the process will eventually yield a dark product that commands only a low price as it is limited to use in technical applications. Lighter-colored seaweeds, such as Macrocystis, yield a lighter-colored alginate suitable for food and other applications. Color can be controlled by the use of bleach; this can be sodium hypochlorite that is added to the filtered alkaline extract or even to the paste at the final conversion stage. Care must be taken, since excessive bleach can lower the viscosity of the alginate, reducing its value. Sometimes the seaweed is soaked in a formalin solution before it is extracted with alkali. The formalin helps bind the colored compounds to the cellulose in the cell walls; so much of the color is left behind in the seaweed residue when the alkaline extract is filtered. Large quantities of water are used in the process, especially when diluting the thick (viscous) initial alkaline extract to a viscosity suitable for filtration. Thus plentiful and reliable water supply is a necessity for an alginate factory to survive. Wastewater from filtration is alkaline; it contains calcium from the calcium precipitation (excess calcium gives a more fibrous calcium alginate) and acid from the acid conversion step. In some countries the waste is pumped out to sea. Where environmental concerns are greater, or when water supplies are limited, recycling is not too difficult, and its costs may be partly offset by the lowering the quantity and cost of water used by the factory. A means of disposing of solid wastes—the

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seaweed residue and used filter aid—must be found. There have been several positive studies reported on the use of this waste to adsorb heavy metals, such as cadmium, zinc, and copper, from industrial liquid wastes (Romero-Gonzalez et al. 2001). Attempts to ferment this waste to produce ethanol from the cellulose content appear to be less economical (Horn et al. 2000).

11.2.5 Ulvan Ulvan, also known as rhamnan sulfate, can be extracted from the edible green seaweed, such as Ulva pertusa, Ulva lactuca, or Ulva rotundata. Ulvan is extracted via two major processes: the acid extraction process and combined chemical and enzymatic process. Ulvan acid extraction process as illustrated in Figure 11.8 is initiated by heating a sizable amount of the algal powder in HCl solution stirred at 250 rpm in a jacketed Algal powder + HCl solution Heated at 80–90°C for 2 hours with stirring Extracted suspension

Filtration

Centrifugation

Supernatant

Solid particles

Filtration

Filtrate

Impurities Centrifugation

Add ethanol (washing) Pellet

Drying

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Ulvan

Figure 11.8  Flowchart for ulvan extraction.

**Repeated washing at different alcohol percentage

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stainless-steel reactor flask with a thermostatic bath in a discontinuous process at 80–90°C for about 2 hours. After extraction, the suspension is filtered through cheesecloth and then allowed to cool down to room temperature. The filtrate is centrifuged at 10,000 rpm at 10°C for 20 minutes in order to remove solid particles. The supernatant is then filtered to remove the impurities. The pH of this extract is adjusted to 3.5 with NaOH. The ulvan is precipitated by the addition of ethanol to the extract. The alcohol precipitate is then separated from the supernatants by centrifugation at 5000 rpm for 20 minutes at 10°C. The alcohol precipitate is washed 3 times with 50%, 75%, and 100% ethanol consecutively and centrifuged at 5000 rpm for 10 minutes’ consistency at 10°C. Finally, it is dried in a vacuum oven at 40°C to a constant weight and then finely ground using a centrifugal milling. Using the combined chemical and enzymatic extraction process, algal powder is suspended in deionised water and brought to pH 5 by the addition of HCl. Cellulose is added, and the suspension is left at 50°C for 2 hours. The final pH is adjusted to 7 with NaOH, and 3 mL of active protease enzyme is added. The suspension is left for 2 hours at 50°C and then for 30 minutes in a boiling-water bath. The suspension is centrifuged at 8000 rpm for 30 minutes, and the pellets obtained are redissolved in deionized water and reextracted for 30 minutes in a boiling-water bath. The suspension is centrifuged as done previously, and the two supernatants are pooled together. The supernatant obtained is then filtered through cheesecloth and then through a sintered-glass filter to remove the impurities. The pH of the extracts is adjusted to 3.5 with NaOH and the suspension is concentrated and desalted using the ultrafiltration process. The retained solution is diafiltered 5 times with deionized water and then precipitated using ethanol. The ethanol precipitate is separated from the supernatants by centrifugation at 5000 rpm for 20 minutes at 10°C. Finally, it is dried in a vacuum oven at 40°C to a constant weight before being finely ground using a centrifugal milling.

11.2.6  Chitin and chitosan Chitin is extracted from the shells of exoskeletal animals such as crab, shrimp, crayfish, and lobster, its derivative, chitosan, is a biomaterial, produced primarily from its alkaline deacetylation using 40–50% NaOH (Al-Manhel 2016). However, N-deacetylation is almost never complete; thus chitosan is considered as a partially N-deacetylated derivative of chitin. It is an abundant natural biopolymer, obtained from the exoskeletons of crustaceans and arthropods, which is a nontoxic copolymer consisting of β-(1,4)-2-acetamido-2deoxy-d-glucose and β-(1,4)-2-anaino-2-deoxy-d-glucose units. The two main processes for chitin extraction are physicomechanical extraction and chemical extraction.

11.2.6.1  Physicomechanical extraction The physical process comprises the step of cutting the shells in a wet state to approximately uniform particles. The resultant mass is then diluted and mixed to form a pumpable slurry. The slurry is separated to remove most unwanted materials, after which the particle size of the concentrated chitinous residue is further reduced. The once-washed, finely cut chitinous slurry is again separated, resulting in a chitin end product as illustrated in Figure 11.9. Figure 11.10 further illustrates the mechanical chitin extraction. Exoskeletons, which have been peeled from crustaceans, are promptly subjected to high-speed cutting, yielding fairly uniform particles, preferably about 1/16 inch to 1/4 inch in size. The cut exoskeletal material is then diluted and mixed with water in a ratio of about 1:5 to about 1:30. The particles of chitinous and other materials are distributed in the dilution solution by simple

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Enzymatic Technologies for Marine Polysaccharides Wet shell

Cutting

Water addition, washing and separation

First waste stream

First chitin extract

Shear cutting

Water addition, washing and separation

Second waste stream

Second chitin extract

Figure 11.9  Schematics for the mechanical extraction of Chitin.

mixing, thus forming pumpable slurry. This pumpable slurry is then pumped through a filtering agitated sieving device with a fine-mesh (∼800-μm) screen, which substantially allows all of the added water to be separated from the solid chitinous residue, thus concentrating the chitin content. By this process, a substantial amount of the unwanted contaminant materials are removed, along with most of the diluting solution. The bacterial load remaining in the resulting material is significantly reduced because it is carried away with the other unwanted materials. The separated slurry, now consisting of partially purified chitinous particles, is once again diluted and mixed with water and to about the same mixture ratios as mentioned above, to form a second concentrated slurry. The particles within this second chitinous slurry are further reduced in size by passing them through a highspeed impact cutting device, which reduces the particle size to an average of approximately 70 μm. This resultant finely cut and once-washed second chitinous slurry is then filtered using a final filtering agitated sieve with a 200-mesh (∼70-μm) screen. This step ensures that the newly added diluting solution and most of the diluting solution remaining from the initial washing of the first slurry are removed. This final filtering also removes most of the remaining increments of the finely dispersed protein and lipids, as well as a majority of the small calcium granulites. This method extracts an approximately 65% pure chitin product. The resultant chitin product can then be further diluted with an aqueous solution and filtered to create a more purified chitin product. The resultant washed and purified chitin is then dried in any suitable drier.

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Peeler Liquid Drained shells

Coarse pre-cutter Water

Dilution tank

Rotary sieve Liquid waste Washed particulate mass

High speed micro-cutter Water

Dilution tank

Rotary sieve

Liquid waste

Chitin

Figure 11.10  Flowchart for the mechanical extraction of Chitin.

11.2.6.2  Chemical extraction Shells of exoskeleton animals, such as shrimp, consist of matrix of chitin, to which protein is covalently bonded (Chang and Tsai 1997). This chitin-protein complex is also associated with unwanted fine granules of calcium carbonate and lipids, which are primarily in the form of a lipoprotein membrane, especially the epicuticle (Roer and Dillamen 1984). The extent of these associations appears to be characteristic for each crustacean species and thus varies. For the chitin to be functionally useful, it must be effectively separated from these unwanted nonchitinous materials, specifically the lipids, protein, and calcium carbonate. However, since chitin is insoluble in virtually everything except highly concentrated acidic solutions, the conventional practice is to remove the unwanted nonchitinous contaminants from the insoluble chitin matrix by using strong, and environmentally hazardous, acidic solutions. Prior efforts to accomplish this separation have utilized different chemical procedures such as boiling lye, alkali with constant electrical current, sulfuric acid, and sodium hydroxide, followed by hydrochloric acid (deproteination by alkali and de-mineralization by acid) (No and Myers 1995).

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Washed shell Waste water Dried shell HCl (Demineralization)

At ambient temp (30°C) for 6 hours Residue 1

NaOH (Deproteination) Residue 2 NaOCl (Decoloration) Residue 3 Distilled water Chitin 50% NaOH (Deacetylation Chitosan

Figure 11.11  Flowchart for the chemical extraction of Chitin and Chitosan.

As illustrated in Figure 11.11, the exoskeletal shells are thoroughly washed with water and dried to remove excess water. Then dried shells are demineralized using HCl at ambient temperature (approximately 30°C) for 6 hours. The residue is then washed with distilled water until pH 6.5–7 is reached and dried afterward. Then, after the demineralized exoskeletal shells are deproteinized using NaOH solutions at 65°C for 2 hours and decolorized using NaOCl, the residue is washed thoroughly with distilled water until the pH reaches 6.5–7.5. The chitin is dried and ground. This chitin could be further deacetylated in 50% NaOH for 5 hours’ consistency at 100°C to produce chitosan. After deacetylation, the chitosan is washed thoroughly with distilled water until pH 6.5–7.5 is reached (Ocloo et al. 2011). After drying, the chitin matrix becomes lithified to the point where it can best be described as vitreous in nature, and thus much more difficult to cut or grind. Thus, if the exoskeletons of marine crustaceans are treated and extracted promptly and most importantly in a wet state, they can easily be cut (rather than ground) into a very small particles. When finely cut in this manner, the inherent protein, fat, and much of the calcium can be separated and removed by simply washing the whole exoskeletal cuts with water, thus forming the first chitin extract. The step ensures no need for any of the harsh and dangerous chemicals used, and thus produces chitin that is low in residual chemical contamination. Furthermore, since the cutting, subsequent dilution, and initial separation of solids takes place early in the process, preferably at or near the shellfish processing site, bacterial loads are not allowed to increase as much as in conventional practice.

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11.2.7 Fucoidan Fucoidan, or fucose-containing sulfated polysaccharides (FCSPs), is present in algae and has numerous functions relating to physiological activity (Ale et al. 2011; Fitton 2011). Fucoidan is classified as a mucopolysaccharide of brown macroalgae. It is a natural, heterogeneous sulfated polysaccharide unique to marine brown macroalgae. The carbon backbone of fucoidan comprises the fucose-repeating chain. Fucoidan has been extracted from marine invertebrates such as the sea cucumber (Fitton 2005; Ribeiro et al. 1994); however, brown macroalgae is the largest source (Hahn et al. 2012). Brown macroalgae that have been used for fucoidan extraction include Sargassum filipendula (Costa et al. 2011), Undaria pinnatifida (Yang et al. 2008), Fucus vesiculosus, Ascophyllum nodosum (Rioux et al. 2007), Laminaria saccharina, and Cladosiphon okamuranus (Cumashi et al. 2007). Following the Lee et al. (2012) fucoidan extraction method as illustrated in Figure 11.12, algal powder is added to deionized water (algal: water ratio of 1:40) and then boiled while stirring for 1 h. After cooling to room temperature, the extract is centrifuged at 3273g for 10 minutes, and the supernatant collected. A fraction of the algal supernatant is lyophilized and referred to as product A. The remaining supernatant is treated with trichloroacetic acid (TCA) to precipitate the protein and then centrifuged at 3273g for 10 minutes. The sediment is lyophilized to yield product B, resultant supernatant is collected, and a portion of this supernatant lyophilized to yield product C. Then 4 volumes of 95% ethanol is added to the remaining supernatant in order to precipitate the polysaccharide, which is recovered by centrifugation at 3273g for 10 minutes, lyophilized to yield fucoidan.

Algal powder Boiled in water for 1 hour with stirring Sediment

Supernatant Lyophilized Residue A Add TCA to precipitate protein Sediment

Supernatant

Lyophilized

Lyophilized Residue C

Residue B

Add Alcohol Sediment

Supernatant

Lyophilized Fucoidan

Figure 11.12 Procedures for extraction of fucoidan from brown macroalgae according to Lee et al. (2012).

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Enzymatic Technologies for Marine Polysaccharides

Algal powder 85% ethanol for 12 hours then washed with acetone and left to dry at room temp overnight

Supernatant

Sediment Add 100 ml of deionized water for 1 hour at 65% Sediment

Supernatant Add CaCl2 and stored at 4°C overnight Supernatant

Sediment

95% ethanol to precipitate fucoidan Fucoidan

Figure 11.13 Procedures for extraction of fucoidan from brown macroalgae described by Yang et al. (2008); Rodriguez-Jasso et al. (2011); and Foley et al. (2011).

Subsequent Fucoidan extraction procedure from brown macro algae is illustrated in Figure 11.13 as described by Yang et al. (2008), Rodriguez-Jasso et al. (2011), and Foley et al. (2011). The algal powder (approximately 1 g) is mixed with 100 mL of 85% ethanol and stirred for at least 12 hours at room temperature to remove lipids and pigments. The solution is then centrifuged at 3273g for 10 minutes and the supernatant removed. The precipitate is then washed with acetone and left to dry at room temperature overnight. The precipitate is then extracted with 200 mL of deionized water by continuous stirring on a hotplate at 65°C for 1 hour. The solution is then centrifuged at 3273g for 10 minutes and 1% CaCl2 added to the supernatant to precipitate the alginate, and the resultant solution is stored in a 4°C refrigerator overnight. The solution is then centrifuged at 3273g for 10 minutes, with the addition of ethanol to the supernatant obtained, giving a final ethanol concentration of 30% (v/v). The solution was then centrifuged at 3273g for 10 minutes to remove unwanted impurities. Then ethanol was added to the supernatant collected, giving a final ethanol concentration of 70% (v/v), and the resultant mixture was stored at 4°C overnight. The fucoidan was then recovered by centrifuging the mixture at 3273g for 10 minutes; the precipitate, which was the fucoidan, was lyophilized and stored at 4°C. The first method clearly extracts the polysaccharide in its most natural state without chemical interference, but this extraction method is greatly limited because it is very expensive, solely because it requires freeze drying. However, the second method utilizes a selective chemical precipitation to remove the impurities, thus leaving an ethanolprecipitated fucoidan. This second method can be said to be more suitable for commercial production because it is cheaper, but for accessing the biological activities of fucoidan, the

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first method should be utilized as fucoidan is extracted in the absence of chemicals that are capable of disrupting its natural structural conformation.

11.2.8 Porphyran Porphyran is obtained mainly from marine red alga Porphyra umbilicalis, and it is structurally composed of mixture of related polysaccharides (Rees and Conway 1962). The three main extraction methods for porphyran are as illustrated in Figure 11.14. The first method (method A in the figure) is devoid of any chemical treatment, as the algal fronds are basically extracted with distilled water. In the second method (B), after filtering the

Dry raw algal fronds of Porphyra yezoensis B

A Add 200 ml of distilled water

C Add 200 ml of distilled water

Dip in 200 ml of HCHO at 30°C for 24 hours Add 200 ml of distilled water

Heat at 100°C with stirring for 4 hours

Filter through a layer of diatomaceous earth

Add 200 ml of TCA and centrifuge at 3000 rpm for 10 minutes

Dialyze against running water overnight

Concentrate under pressure to ¼ of its original volume

Precipitate with 4 volumes of ethanol, then centrifuge at 3000 rpm for 10 minutes

Wash with ethanol and acetone

Air-dry at 30°C for 12 hours

Porphyran

Figure 11.14  Three main protocols for extraction of porphyran from Porphyra yezoensis.

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algal fronds on diatomaceous earth, a chemical treatment with 10% trichloroacetic acid is carried out. In the third method (C), the raw algal fronds are first treated with 3.7% formaldehyde solution in a flask closed with a stopper at 30°C for 24 hours. After addition of 2 × volume of distilled water, the suspension is then heated at 100°C with continuous stirring for 12 hours. The extract is cooled to room temperature and filtered through a layer of diatomaceous earth. The clarified filtrate is dialyzed against running tap water for 24 hours and evaporated to one-fourth of its original volume under reduced pressure, and to this 4 × volume of ethanol is added at room temperature. The gelatinous precipitate collected by centrifugation is washed with ethanol, and acetone, and dried at 30°C for 12 hours (Nishide et al. 1988). The chemical treatment was found to affect the total yield and the galactose content of the resultant porphyran, but it induced the release of more prophyran from the core of the raw algal frond given rise to higher galactose content. Thus, the chemically induced process is advisable for commercial production of porphyran, while the chemically free process can be harnessed for intrinsic biological activity.

11.3  Assessment of intrinsic purity and structural elucidation The different solvents used in precipitation of these polysaccharides and chromatographic separation are basically the first step in the purification of these compounds, resulting in a degree of purity and homogeneity. Marine polysaccharides generally are polydisperse (i.e., chemically homogenous but varying in molecular weight) or polymolecular (i.e., varying in chemical composition and molecular size), and these often pose major challenges during structural analysis. Homogeneity ensures the absence of extraneous contaminants and “discontinuities in molecular size and structure.” Homogeneity of size can be shown by size-exclusion chromatography (SEC), while homogeneity of chemical composition requires the consistency of certain parameters such as sugar ratio, sugar/functional group ratio, physical properties, and spectroscopic data. Before undertaking rigorous structural analyses of isolated polysaccharide, it is necessary to estimate its purity, that is, the proportion of polysaccharide component and the presence of impurities. Figure 11.15 is a graphic representation of the methodologies employed in the fractionation and purification of polysaccharides in order to elucidate their structural compositions, linkage positions, oligomer structures, and molecular weight distribution.

11.3.1  Methods used in assessing intrinsic purity 11.3.1.1  Molecular mass Average molecular weight (MW), average radius of gyration (Rg), intrinsic viscosity ([η]), and other molecular characteristics of soluble polysaccharides are determined by sizeexclusion– gel permeation chromatography (SEC-GPC), laser light scattering (LLS), and viscometry. Individual polysaccharides have their have their specific molecular weight, radius of gyration, and viscosity. These parameters are targeted in samples to ascertain the purity of each polysaccharide. Polysaccharides are commonly dissolved in an aqueous system or are analyzed directly in the extracts (Synytsya and Novak 2014).

11.3.1.2  Chemolytic method The chemolytic method is based on chemical or enzymatic cleavage of polysaccharide followed by separation of degradation products (oligo- or monosaccharides or even smaller fragments). A parent polysaccharide and/or a degradation product can be chemically

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Fractionated Polysaccharide

Complete hydrozylation

HPLC/ derivation-GLC

Complete Methylation

Partial hydroxylation

Complete hydroxylation

Oligomer purification

Reduced to additols and acetylated

HPSEC

Structural analysis

GLC analysis Monomer composition

Position of linkages

Oligomer structure

Molecular weight distribution

Enzymatic specificity or NMR for anomeric configuration determination

Figure 11.15  An overview of the structural analysis of polysaccharides.

modified for analytical purposes. Resistance of glycosidic bonds is not the same for various polysaccharides and depends on anomeric configuration, position, molecular environment, and availability (Synytsya and Novak 2014). Chemolytic methods are still attractive for linkage analysis of complex polysaccharides. Total hydrolysis of polysaccharides can be made under strong acidic conditions by the use of formic, sulfuric, or trifluoroacetic acid at concentrations of 2 mol/L and heating up to 110°C for several hours (Synytsya and Novak 2014). Several steps of the acid hydrolysis in different media can be combined to achieve a better effect. The hydrolysates obtained are neutralized, diluted, and analyzed by an appropriate separation method, usually liquid chromatography. Alternatively, monosaccharide mixtures can undergo reduction and then acetylation or silylation to obtain volatile derivatives for gas chromatography (GC).

11.3.1.3  Methylation analysis (MA) This analysis is employed using the sugar linkages within the polysaccharides following the steps listed below (Lindberg and Lönngren 1978) and produces methylated derivatives that are used to verify the specific constituent in a known purified polysaccharide sample: 1. Complete methylation of all hydroxyls in the polysaccharide. This is confirmed by the disappearance of a broad absorption band at 3200–3700 cm−1 (OH stretching) in the Fourier transform infrared (FTIR) spectrum of methylated glucan. 2. Cleavage of the methylated glucan into the monosaccharide fragments. This treatment leads to breakage of all glycosidic linkages, but not methyl–ether bonds. Strongly acidic hydrolysis, methanolysis, or acetolysis is suitable for this degradation.

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3. Reduction of formed aldehydes followed by acetylation or silylation. This step leads to partially methylated alditols or silans, which are then further acetylated or silylated. Acetolysis at the previous step leads to complete acetylation of free hydroxyls. 4. Separation analysis of the alditol derivatives. This step is carried out using flame ionization (FID) or mass spectrometric (MS) detectors, or gas chromatography (GC). Positions of glycosidic linkages in the parent glucan correspond to nonmethylated hydroxyls in methylated alditols or silans, while all free hydroxyls would have been methylated.

11.3.1.4  Vibration spectroscopy Fourier transform infrared (FTIR) spectroscopy is a powerful tool in the structural analysis of polysaccharides. It is sensitive to the position and anomeric configuration of glycosidic linkages in polysaccharides. It is possible to analyze glucans in crude high-molecularweight fractions isolated from various raw materials. Greater resolution of FTIR exhibited differences in several IR bands assigned mainly to vibrations of groups involved in the system of hydrogen bonds in polysaccharide (Michell 1993). For the obtained dynamic IR signals (in-phase and out-of-phase responses), the dynamic IR cross-correlation is defined. (Synytsya and Novak 2014). Cross-correlation of the bands with the bands from a standard sample shows the degree of purity of the polysaccharide.

11.3.1.5  X-ray diffraction spectrometer (XRD) This method is used to detect crystallinity of polysaccharides. The XRD pattern will show characteristic peaks. The sharper peaks will provide evidence of denser crystalline structure. The purity of the sample can be ascertained by the sharpness of the peaks, while an adulterated sample will not produce such peaks.

11.3.1.6  Degree of deacetylation This index is used basically for chitin and chitosan. Degree of deacetylation is becoming an increasingly important property for chitosan, as it determines how the biopolymer can be utilized. Chitosan is the product of total deacetylation of chitin; hence there is a chance that the two products might be in the mixture in the process of deacylation. DA can be used to ascertain complete deacylation hence showing that the sample of chitin being deacylated is completely deacylated and thus, what we have now is a pure chitosan.

11.3.2  Structural elucidation 11.3.2.1  Desulfation and methylation Structural analysis of sulfated polysaccharides requires determination of the attached sulfate esters along with their backbones and the glycosidic linkage types. Methylation is used to determine linkages between monosaccharides. By comparing the methylation of native polysaccharides to that of their desulfated counterparts, one can determine the positions of the sulfate groups. Thus, hydrolysis of the permethylated, desulfated polysaccharides, yields partially methylated monosaccharides, which following acetylation are separated and identified on gas chromatography/mass spectra (Bjorndal et al. 1976). Accurate structural determination requires desulfation of the polysaccharide without

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cleavage of the polysaccharides chain linkages. Often a solvolytic desulfation procedure is used wherein the polysaccharide as a pyridinium salt is heated in dimethyl sulfoxide (Jiao et al. 2011). Desulfation of fucoidan by methyl sulfoxide pyridine is rapid and complete, resulting in higher yields with little degradation. However, a method using chlorotrimethylsilane (CTMS) for treatment of pyridinium salts is more appropriate for desulfation of sulfated galactans of both the agaran and carrageenan families (Kolender et al. 2004). Other approaches for desulfation that have been used involve methanolic hydrogen chloride, silylating reagents, and pyromellitic acid (Jiao et al. 2011). Chemical desulfation is relatively nonspecific and usually results in a significant loss of sample material. The use of sulfatases represents a more specific approach to desulfation and would be advantageous in structural studies, although such enzymatic approaches do not appear to be frequently used (Kusaykin et al. 2006). The reason for this is unclear but could be due to the lack of commercially available enzymes of the required specificity. The typical methylation procedure is straightforward, involving treatment of the desulfated polysaccharide sample with methyl iodide in the presence of solid base, usually sodium hydroxide, in methyl sulfoxide, and the procedure can be repeated to obtain a complete methylation (Jiao et al. 2011).

11.3.2.2  Nuclear magnetic resonance (NMR) spectroscopy This is a powerful tool for the structural elucidation of sulfated polysaccharides as it can provide structural details such as the monosaccharide components, linkages, anomeric configurations, and positions of branching or sulfations. This can be done by combining various 1D and 2D-NMR techniques. The use of NMR in the structural analysis of polysaccharides has been substantial (Usov et al. 1980). This may be attributed to the relative high proportion of repeating sequences (identical sulfation pattern) in these polysaccharides that make them amenable to analysis by 13C NMR.

11.3.2.3  Mass spectroscopy (MS) This is valuable in the structural analysis of polysaccharides, as it generates accurate molecular mass data for oligosaccharides and can also provide sequence information. Compared with other analytical techniques, MS methods have several advantages, including low sample consumption (e.g., picomole quantities) and short analysis time. While analysis of sulfated polysaccharides by MS can be problematic because of the labile nature of the sulfate groups, approaches based on electrospray ionization collision–induced dissociation (ESI-CID) and matrix-assisted laser desorption/ionization (MALDI) are increasingly being developed to characterize sulfated oligosaccharides (Daniel et al. 2007; Yang et al. 2009; Jiao et al. 2011).

11.3.2.4  Sulfation of algal polysaccharides Structure modification of sulfated polysaccharides, such as desulfation, oversulfation, acetylation, and benzoylation, would allow the development of new and possibly more effective derivatives of naturally occurring polysaccharides (Jiao et al. 2011). Certainly, the most frequent structural modification to sulfate polysaccharides is oversulfation due to the typically strong positive correlation between their sulfate content and biological activity. A number of methods have been developed for polysaccharide oversulfation, such as treatment with sulfuric acid, sulfur trioxide–pyridine, chlorosulfonic acid–pyridine, dimethylformamide, and sulfur trioxide-dimethylamine (Jiao et al. 2011).

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11.3.2.5  Molecular size modification Lower-molecular-weight algal sulfated polysaccharides can be prepared by chemical, physical, or enzymatic means to obtain oligosaccharides with more diverse bioactivities. For example, linear or branched sulfated galactans and fucans can be cleaved by mild acid hydrolysis (Teruya et al. 2010) or by a radical process involving a hydrogen peroxide–cupric redox system (Tao et al. 2010). The chemical methods are easy to perform but lack specificity. Minor changes in temperature or acidity can lead to variations in oligosaccharide sizes and sulfation patterns, while strong acid may alter the sulfation pattern or destroy the polysaccharide chain (Holtkamp et al. 2009). Enzymatic degradation of sulfated polysaccharides can be achieved by selecting enzymes such as hydrolases, fucoidanases, α-l-fucosidases, and galactosidases (Bhattacharyya 2010) to target glycosidic bonds while preserving the sulfate groups (Qiu et al. 2006).

11.3.3  Structural characterization–specific polysaccharide 11.3.3.1 Agar The most important properties to be considered when determining the quality of agar for food, tissue culture, or bacteriological use are gel strength, gelation and melting temperatures, sulfate and methoxyl contents, clarity of the solution, and ash content. Agar possessing high gel strength is always preferred for food. For bacteriological media, gel strength of 300–400 g/cm at a 1% concentration is acceptable. Agar, when dissolved in hot water, should form a clear solution, the clarity of which can be determined by measuring the transmittance of the solution. Proximate composition reveals that agar is composed of nitrogen, carbon, and sulfate contents, which were determined by elemental microanalysis. Protein content was calculated from N% using the correction factor of 6.25, as proposed by Marks et al. (1985). Moisture was obtained by heating 0.5 g of samples at 105°C for 24 hours (Maciel et al. 2008). The peak molar mass (Mpk) was estimated by high-performance size-exclusion chromatography (HPSEC) at room temperature using an ultrahydrogel linear column (7.8 × 300 mm), flow rate of 0.5 mL/minute, 0.5% polysaccharide concentration, and 0.1 M NaNO3 as solvent. A differential refractometer was used as detector and the elution volume was corrected to ethylene glycol at 11.25 mL as an internal marker (Barrosa et al. 2013). One-dimensional (1D) FTIR and two-dimensional (2D) NMR spectroscopy of 13C and 1H were used to determine the chemical structure of the polysaccharide fraction obtained. Infrared (IR) spectroscopy, basically the FTIR spectra, was recorded between 400 and 4000 cm−1. Nuclear magnetic resonance (NMR) spectroscopy 13C and 1H NMR spectra of 2.5% (w/v) solutions in D2O were recorded at 353 K on a FTIR spectrometer with an inverse multinuclear gradient probe head equipped with z-shielded gradient coils, and with silicon graphics. Sodium 2,2-dimethylsilapentane-5-sulfonate (DSS) was used as the internal standard (0.00 ppm for 1 hour). A distortionless enhancement by a polarization transfer spectrum was recorded in order to determine the hydrogenation of each carbon. The FTIR spectrum shows characteristic bands of agar-type polysaccharides (1258, 1075, 930, and 891 cm−1) (Barrosa et al. 2013). As this band possesses a low intensity in pyrolysis gas chromatography (PGC), only a small degree of substitution was expected for this polysaccharide. This result was confirmed by the low degree of substitution (DS) sulfate (0.13) calculated by elemental microanalysis. The absorbance at 930 cm−1 has been assigned to the vibration C-O-C bridge in 3,6-anhydro-l-galactose (Christiaen and Bodard 1983) and the skeletal mode of galactans was attributed to the signal at 1075 cm−1 (Sekkal and Legrand 1993).

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OH CH2OH H

H

O O

H

H

β1

OH H

H

H α1

O

3 O

CH2

O

O

4 H

H

OH

Figure 11.16 Chemical structure of the repeating disaccharide unit of agar molecules (RheinKnudsen et al. 2015).

In agar polysaccharide, the position of sulfate group can be inferred by bands in the 800– 850 cm−1 region. Reports show that bands at 845 and 830 cm−1 can be attributed to respectively the 4-O-sulfate and 2-O-sulfate groups present in D-galactose units, while the signals at 820 and 805 cm−1 are due to sulfation on C6 of l-galactose and C2 of the 3,6-anhydro-l-galactose (Barrosa et al. 2013). Agar has been found to have a linear chain similar to that of carrageenans, consisting of repeating sections of (1→3) linked β-d-galactopyranosyl units joined to 3,6-anhydro-α-l-galactopyranosyl units via (1→4)-linkages (Figure 11.16). It has very few sulfate groups and up to 21% of C-6 carbon on β-d-galactopyranosyl units may contain methoxyl groups, affecting the gelation properties. Unique gelation properties of agar is that gelation occurs at temperatures (30–35°C) far below the gel melting temperature (90–95°C).

11.3.3.2 Carrageenan The purity of carrageenan can be assessed via different methods. Carrageenans extracted under optimum conditions were analyzed for total ash, calcium, iron, sodium, potassium (AOAC 2005), protein (Lowry et al. 1951), and phosphate content (Instituto Adolfo Lutz 2005). Sulfate content was measured turbidimetrically after hydrolyzing 40 mg of carrageenan crude extract in sealed tubes for 2 hours in 0.5 N HCl at 105°C (Jackson and McCandless 1978). Total carbohydrate was determined by the phenol sulfuric acid method (Dubois et al. 1956). Carrageenan is insoluble in ethanol; it is soluble in water at a temperature of about 80°C, forming a viscous, clear, or slightly opalescent solution that flows readily; it disperses in water more readily if first moistened with alcohol, glycerol, or a saturated solution of glucose or sucrose in water. Hydrolysed sulfate groups are precipitated as barium sulfate in carrageenan and must not be less than 15% or more than 40%. For the structural elucidation, Fourier transform infrared spectroscopy (FTIR) of carrageenan was performed using a spectrometer with a resolution of 4 cm−1 in the range of 4000–400 cm−1 (Webber et al. 2012). The spectra were obtained in KBr chips (spectrometric degree). Commercial and κ-carrageenan patterns were used as reference materials for infrared spectroscopy analysis. The study of carrageenans by FTIR spectroscopy shows the presence of very strong absorption bands in the 1210–1260 cm−1 region (due to the S═O of sulfate esters) and the 1010–1080 cm−1 region (ascribed to the glycosidic linkage) for all types of carrageenan (Webber et al. 2012). A particularly intense signal is recorded in all samples at 803–805 cm−1, which is specific to 3,6-anhydrogalactose-2-sulfate. Another signal is observed at 840–850 cm−1 (attributed to d-galactose 4-sulfate). Peaks are also observed at 925–935 cm−1 in all samples (3,6-anhydro-d-galactose) (Pereira 2009; PradoFernández 2003). Since carrageenan is the general name for a family of galactans, which are the most common and abundant cell wall constituents in red algae, the backbone structure is based on linear chains of repeating galactose units in D configuration (d-sugar) and 3,6-anhydro-galactose copolymer, joined by alternating α-(1→3) and

248

Enzymatic Technologies for Marine Polysaccharides ν-carrageenan

µ-carrageenan –

–

OSO3

CH2OH O 3

O

O OSO3–

4

–

OSO3 O

CH2OH

α1

O

OH β1

O

OSO3

–

OSO3 O

3

HO

κ-carrageenan

ι-carrageenan – OSO3

–

OSO3

CH2OH O O

OH

4

HO

3

O

O

OH β1

O

α1

O O

O

OH β1

CH2OH

α1 O

O 3 O

4 OH

O O

O

OH β1

α1 O

4 OSO3–

Figure 11.17  Chemical structures of the various forms of carrageenan (Rhein-Knudsen et al. 2015).

β-(1→4) linkages. In terms of chemical structure, this polygalactan is classified into various types (Figure 11.17), including but not limited to κ-, λ-, μ-, ι-, θ-, β- and ν-carrageenans, all containing 15–40% ester sulfate with the exception of λ-carrageenan, which is devoid of sulfate content (De Ruiter and Rudolph 1997). At least 15 different carrageenan structures have been reported, of which kappa (κ), iota (ι), and lambda (λ) forms are the most industrially relevant. The major difference among the various forms of carrageenan is related to structural characteristics, including the number and position of sulfate groups and the occurrence of 3,6-anhydro-D-galactose in the chain (Lahaye 2001). For instance, κ-, ι-, and λ-carrageenans are distinguished by the presence of one, two, and three ester sulfate groups per repeating disaccharide unit, respectively (Guangling et al. 2011). The chemical structures of carrageenans are thus very heterogeneous and are correlated with the algal sources, the life stage of the seaweed (i.e., gametophyte), and the extraction procedures of the polysaccharide (Hawkes 1990). The structure of κ-carrageenan is an alternating 3-linked-d-galactose and 4-linked anhydrogalactose (AG) unit. It has an ester sulfate content of about 25–30% and a 3,6-AG content of approximately 28–35%. ι-Carrageenan has an additional sulfate group on C2 of the AG residue, resulting in two sulfates per disaccharide-repeating unit. It has an ester sulfate content of 28–30% and approximately 25–30% content of 3,6-AG. λ-Carrageenan has three sulfate groups per disaccharide unit, with the third sulfate group of this form at the C6 position of the 4-linked residue. There is an ester sulfate content of about 32–39% and no 3,6-AG content (Guangling et al. 2011).

11.3.3.3 Laminarin The purity of laminarin is ensured from the procedure of production as the specificity of isopropyl alcohol is limited to laminarin and fucoidan. The fucoidan will separate first and can be filtered from the solution. Ultrafiltration can also be used for the separation,

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249 CH2OH

CH2OH

O

O OH

O O

CH2(CHOH)4CH2OH

OH

Mannitol

OH

OH

n

OH

(A) M chain Laminarin CH2OH

CH2OH

CH2OH

O

O OH

O

OH

OH OH

OH

O OH

O OH n

OH

(B) G chain Laminarin

Figure 11.18  The structures of the M and G chains in Laminarin (Kadam et al. 2015).

as it separates on the basis of size and molecular weight. The membrane chosen will be dependent on the molecular weight of the laminarin and fucoidan and the desired separation level between the two. Viscosity of laminarin can be used to determine its molecular weight. High-molecular-weight laminarin yields high viscous solution (Hjelland et al. 2013). Characterization of laminarin can be carried out by NMR spectroscopy using the 1H–13C heteronuclear single-quantum coherence method (Date et al. 2012), while monosaccharides analysis of laminarin can be carried out using GSMS spectroscopy with a fused-silica capillary column and polar capillary phase (Cheng et al. 2011). Laminarin contains a large amount of neutral sugars and it could be of M-chain of G-chain laminarin (see Figure 11.18). For example, laminarin from S. longicruris contained 99.1% of neutral sugars, while A. nodosum and Fucus vesiculosus have relatively lower concentrations of 89.6% and 84.1%, respectively (Rioux et al. 2007). Also, the β-glucan structure in laminarin can be characterized with respect to refractive index and dielectric constant using terahertz time-domain spectroscopy (Shin et al. 2009). Various techniques used for characterization of laminarin are outlined in Table 11.3. Both evaluation and characterization of compositional variations in seaweed biomass are required in order to extract useful information from the seaweed biomass complexes (Date et al. 2012).

11.3.3.4 Alginate Alginic acid purity can be analyzed by determining the uronic content according to the Blumenkrantz and Asboe-Hansen (1973) protocol as modified by Tullia et al. (1991). This method consists of several steps. First, concentrated sulfuric acid action leads to hydrolysis of osidic bonds and dehydration of the released monosaccharides, resulting in formation of derived furfural compounds, namely, uronic acids. The uronic acids react with 3-phenyl phenol, resulting in a pink coloration in which intensity is measured spectrophotometrically. Coloration is enhanced by addition of borate and interferences due to neutral sugars are minimized owing to sulfamate (added before acid hydrolysis). Purity is calculated as the ratio of the dry weight of uronic acids quantified to the dry weight of alginic powder

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Enzymatic Technologies for Marine Polysaccharides Table 11.3  Techniques employed in the structural elucidation of laminarin

Analysis

Technique

Conditions

Molecular weight

High-performance sizeexclusion chromatography (HPSEC)

Molecular weight

High-performance sizeexclusion chromatography– multiangle laser light scattering (HPSEC-MALLS)

Molecular weight

Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF-MS) Electrospray ionization mass spectrometry (ESI-MS)

Column—TSK gel column G4000PWXL (7.8 mm × 300 mm) and G5000PWXL (7.8 mm × 300 mm); column temperature 25°C; detector—refractive-index detector with response time of 4 seconds; sample concentration 1.5 mg/mL; injection volume 10 µL; efluent—double-distilled water; flow rate 0.5 mL/minute Column—TSK guard column PWXL (6 mm × 40 mm); TSK-G5000 PWXL (7.5 mm × 300 mm) and a TSK-G3000 PWXL (7.5 mm × 300 mm); mobile phase 0.1 M NaCl solution; flow rate 0.5 mL/minute; detector—refractive-index detector Sample preparation—methylated laminarin dissolved in 80:20 (v/v) methanol: water; matrix 2,5-dihydroxybenzoic acid (DHB); external calibrant—insulin B Voltage 2 kV, positive-ion mode; nebulizer pressure 250 kPa; desolvation temperature 210°C Matrix—monothioglycerol; sample—dissolved in methanol Column—fused-silica capillary column (0.32 mm internal diameter, film thickness 0.5 lm); carrier gas—helium; temperature increased at rate of 2°C/minute from 30°C–280°C and maintained for 15 minutes Internal standard 90% deuterium oxide and 1 mM sodium 2,2-dimethyl-2-silapentane5-sulfonate (DSS); temperature 298 K; frequency 700 MHz; probe—inverse gradient 5 mm Cryo 1H/13C/15N Instrument—Nicolet 6700 FTIR spectrometer; wavelength 400–4000 cm−1

Molecular weight

Molecular weight Separation

Fast atomic bombardment mass spectrometry (FABMS) Gas chromatography–mass spectrometry (GC-MS)

Water-soluble components

Nuclear magnetic resonance (NMR) spectrometry

Chemical profile of the solid-state biomass constituents

Attenuated total reflectance Fourier transform infrared (ATR-FTIR) spectroscopy

used for the quantification. Intrinsic viscosity can also be utilized to assess purity by capillary viscosimetry using the elution time. The elution time for each alginate samples is measured for serial dilutions in a 0.44-mm-diameter capillary viscosimeter immersed in a heated circulating water bath maintained at 20°C. Gel-permeation chromatography, strong anion-exchange chromatography, ESIMS and 2D NMR spectral analyzes have been successfully used in the sequential structure determination of alginate-derived oligosaccharides (Zhang et al. 2004). The alginate

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H O

4

COO– H

H

OH O

H

H

α1 O H 4

OH H

OH

α1

H

COO– OH O

O 4

O

HO OH H

H

H

H HO O 4 β1 H

H

H

H

H β1

O

O COO–

OH

H COO–

Figure 11.19  Chemical structure of the repeating units of alginate molecules (Rhein-Knudsen et al. 2015).

oligosaccharides were size-fractioned by low-pressure gel permeation chromatography (LPGPC). The oligosaccharide mixture (100–150 mg/2 mL) was loaded on BioGel P4 and eluted with 0.2 mol/L NH4HCO3 solution at a flow rate of 1.5 mL/hour; the procedure was carried out at room temperature. In each fraction of 1.5 mL/tube collected the uronate content was measured at 235 nm. Strong anion-exchange fast-protein liquid chromatography (SAX-FPLC) was further used to separate the uniform-sized fractions separated from the BioGel P4 column. The column was eluted with a linear gradient from 0 to 0.4 mol/L NaCl solution at a flow rate of 2 mL/minute for 2.5 hours. Each fraction was rotary evaporated and desalted on a Sephadex G-10 column and eluted with double-distilled water. Electrospray–ionization mass spectroscopy was used for determination of the molecular mass of each oligosaccharide. Positive and negative ionization modes were used to get the mass spectra at the same time. The sample was dissolved in 1:1 MeOH:H2O (10 pmol/mL), and it was delivered to the electrospray source using a syringe pump at a flow rate of 5l L/minute. The capillary temperature was kept at 250°C, and nitrogen gas was used as nebulizing and desolvation gas. The pure samples (1–5 mg) were dissolved in 1 mL of D2O (99.96%) and freeze-dried 2 times to remove exchangeable protons, and then dissolved in 0.5 mL of D2O (99.96%) for NMR analysis. All NMR analyses were performed at 298 K (Zhang et al. 2004). Alginate was found to be a linear polysaccharide that is composed of β-dmannuronopyranosyl and α-L-guluronopyranosyl units (Figure 11.19). The units occur in M blocks (containing solely mannuronopyranose residues), G blocks (containing solely guluronopyranose residues), or MG blocks. The proportion of G, M, and MG blocks affects the gel strength and calcium reactivity. Alginate with high G blocks results greater gel strength, while alginate with high M blocks is more calcium-tolerant and less likely to be problematic with syneresis (Haug et al. 1966, 1969).

11.3.3.5 Ulvan Purification of ulvan to eliminate pigments, lipids, amino acids, and peptides has been reported using organic solvents (Chattopadhyay et al. 2007). Generally, a polysaccharide with improved purity is obtained by precipitation with organic solvents, usually ethanol. Afterward, ulvan aqueous extract is concentrated in a rotary evaporator, then hot-air-dried or freeze-dried. The removal of impurities, as well as the drying of ulvan extract, may favor the modification of the polysaccharide conformation and properties (Lahaye et al. 1993). The purity of the polysaccharide is assessed using an anionexchange high-performance liquid chromatogram of its hydrolysate to reflect certain peaks, and a correlation with peaks of l-rhamnose, d-glucose, and d-xylose as well as

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those of d-glucuronic acid and l-idulonic acid indicates that the polysaccharide isolated is an ulvan (Yu et al. 2003). The specific rotation and FTIR spectrum are also efficient in ascertaining ulvan purity. The major absorption in FTIR is attributed to the stretching of —OH, C—H, and C═O groups of uronic acid, the presence of sulfate ester substitutions, C—O—S stretching, vibration of the C—O—C bridge of the glucosides, and also the characteristic FTIR peaks for sulfate groups on position C3 of ulvan’s rhamnose residue (Yu et al. 2003). Ulvan can be methylated according to the procedure described by Ciucanu and Kerek (1984). The permethylated ulvan obtained was subjected to complete acid hydrolysis to produce mixtures of the methylated sugars that were analyzed as the corresponding alditol acetates using gas–liquid chromatography (GC) and combined gas–liquid chromatography/mass spectroscopy (GCMS). In 13C NMR spectra of ulvan, the four sugar moieties are designated as residues A, B, C, and D according to their decreasing anomeric carbon chemical shifts (Tako et al. 2015). The signals A (106.14 ppm), B (105.91), C (103.88), and D (102.82) were assigned to be β-d-glucuronic acid, α-l-idulonic acid, 1,4-linked α-l-rhamnose substituted with sugar, and 1,4-linked α-l-rhamnose, respectively. From 1H spectra of ulvan signals A (4.603 ppm), B (5.264 ppm), C (4.871 ppm), D (4.760 ppm), E (4.573 ppm), and F (5.058 ppm) were assigned to be β-d-glucuronic acid, α-l-idulonic acid, 1,4-linked α-l-rhamnose substituted with sidechain, 1,4-linked α-l-rhamnose, β-d-xylose, and an unknown sugar, respectively (Lahaye 1998). However, Nakamura et al. (2011) reported signal F (5.058 ppm) as 1, 3-linked α-l-rhamnose. Figure 11.20 shows the structural composition of ulvan. The major repeating disaccharide in the ulvan extracted from different ulvan samples comprising two different types of aldobiouronic acid. These were designated as ulvan obiuronic acid 3-sulfate types A and type B (A3s and B3s, respectively). The A3s disaccharide is composed of glucuronic acid and sulfated rhamnose, while type B3s consists of iduronic acid and sulfated rhamnose, associated mainly via (1→4) glycosidic linkages. Rhamnose residues are sulfated mainly at position C3 or at both positions C2 and C3. In some ulvan extracts, xylose or sulfated xylose residues may occur in place of uronic acids. In this case, the disaccharides are called ulvanobiose acids and denoted symbolically as U3s (ulvanobiose acid 3-sulfate) and U2s3s (ulvanobiose acid 2,3-disulfate) (Cunha and Grenha 2016). Ulvan exhibits some of the hydroxyl groups of the sugar residues substituted by sulfate groups. These biopolymers consist of complex highly branched molecules that do not appear to have a defined backbone or simple repeating unit, nor do they appear to have long chains of a single sugar (Percival 1979). The sugar composition of ulvans is extremely variable, and the commons are rhamnose (16.8–45.0%), xylose (2.1–12.0%), glucose (0.5–6.4%), glucuronic acid (6.5–19.0%), and iduronic acid (1.1–9.1%) (Cunha and Grenha 2016). Mannose, galactose, and arabinose have also been found in ulvan from some Ulva species. The heterogeneous chemical composition of ulvan leads to an essentially disordered conformation of the biopolymer. Despite this disordered structure, the local regularity given by the repeating aldobiouronic units, for [3)-α-L-Rhap-(1→4) -α-L-Rhap-(1→4) -α-L-Rhap-(1→4) -α-L-Rhap-(1→4)-β-D-Xylp-(1→4) -α-L-Rhap-(1]n 3 β-D-GlcAp or α-L-IdoAp

2 β-D-GlcAp

Figure 11.20  The structure of ulvan from Ulva fertus (Tako et al. 2015).

3

SO3Na

3

2

SO3Na SO3Na

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instance, is believed to be sufficient for the formation of transient “junction zones” responsible for the formation of the weak gel that ulvan is known to produce in native state (Paradossi et al. 1999). Ulvan extracts have generally demonstrated a pseudoplastic behavior, a viscosity decrease being observed as the shear rate increased (Yaich et al. 2014). Ulvan is considered to have a high charge density, which determines its water solubility. However, it has a certain hydrophobic character, possibly determined by the presence of a high number of methyl groups in the rhamnoserepeating unit. However, the aqueous solubility of the polysaccharides from an ultrastructural analysis revealed the presence of spherical aggregates of ulvan in aqueous solution (Cunha and Grenha 2016). As a polyelectrolyte, both the ionic strength and the pH of the solvent used affected the solubility and morphology of ulvan, since the type and amount of counter ions in solution could contribute to the condensation of the polymer (Robic et al., 2009).

11.3.3.6  Chitin and chitosan The method of chitin extraction is expected to yield pure-quality chitin as the deproteination and demineralization steps are repeated twice (Majekodunmi 2016). This approach makes it possible to obtain higher yields of chitin from the shells. The final deacetylation of chitin at room temperature for up to 3 days gives a longer reaction time, which may result in higher yield of chitosan (Divya et al. 2014). Moisture content and residue on ignition or ash content are analyzed according to the methods described by the Association of Official Analytical Chemists (AOAC). The moisture content may vary depending on season, relative humidity, and intensity of sunlight. The ash content of chitosan is an indication of the effectiveness of the method employed for removing inorganic materials (Islam et al. 2011). The pH measurement of chitosan solutions is carried out with the use of a pH meter, and viscosity is determined using a viscometer. Viscosity of chitosan can be used to determine molecular weight. High-molecular-weight chitosan yields high-viscosity solutions; hence, low viscosity is preferred. The high degree of deacetylation also results in a high amount of protein (Puvvada et al. 2012). The solubility of chitosan in acetic acid is a mark of its purity as it dissolves completely in 1% acetic acid (Majekodunmi 2016). A few grams of chitosan are mixed in 35 mL of 1% acetic acid, and the solution is maintained on a magnetic stirrer for 30 minutes. Sample is taken out and the insolubles are removed by filtration through Whatmann No. 1 filter paper and weighed. For example, if the concentration of chitosan in acetic acid is 7.7 g/L, this indicates that the chitosan obtained is 77% pure. Chitosan, unlike chitin, has a high content of highly protonated free amino groups that attracts ionic compounds (see Figure 11.21). This is why chitosan is soluble in inorganic acid (Mohammed and Williams 2013). X-Ray diffraction (XRD) analysis was used to detect crystallinity of chitosan (Klepka et al. 2006). The XRD pattern shows characteristic peaks at 2θ = 9.28° and 20.18°. The sharper peaks were evidence of denser crystalline structure; however, the characteristics peaks of 2θ = 9.9–10.7 and 19.8–20.7 in range were reported by Majekodunmi (2016). FTIR spectra of chitin and chitosan are usually recorded in the middle infrared (from 4000 cm−1 down to 400 cm−1), with a resolution of 4 cm−1 in the absorbance mode for 8–128 scans at room temperature (Kumirska et al. 2010). The samples for FTIR analysis are prepared by grinding the dry blended powders with powdered KBr, often in the ratio of 1:5 (sample:KBr) and then compressed to form disks. Spectra are then measured using a deuterated triglycerine sulfate detector (DTGS) with a specific detectivity of 1 × 109 cm.Hz(1/2).W−1 (Thanpitcha et al. 2008) or on films using an attenuated total refraction (ATR) method in an IR spectrometer (Kumirska et al. 2010). Diffuse reflectance

254

Enzymatic Technologies for Marine Polysaccharides NHCOCH3

HO HOH2C

HOH2C O

O

HO

O O

HO

NHCOCH3 HOH2C

HO

O

O

NHCOCH3

H

n

Chitin NH2

HO

HOH2C

HOH2C

O

O

HO

O

O

O

HO

H

O

NH2

HO HOH2C

NH2

n

Chitosan

Figure 11.21  The chemical structures of chitin and chitosan (Majekodunmi et al. 2016).

infrared Fourier transform (DRIFT) spectroscopic analysis has also been applied (Urreaga and de la Orden 2006). Depending on the source, chitin (composed majorly of linear chain of acetylglucosamine groups) can occur in α, β, and γ forms, in which their differences depend on the arrangement of chains in the crystalline regions (Jang et al. 2004). The most abundant and stable form is α-chitin, which is obtained mainly from crab and shrimp and is composed of polysaccharide strands aligned in alternating antiparallel fashion that gives rise to strong hydrogen bonding, making it more stable (Minke and Blackwell 1978). The rare β-chitin is composed of parallel strands of polysaccharides, and it is found in association with proteins in squid pens (Rudall and Kenchington 1973; Rudall 1969) The γ form of chitin contains two parallel strands and one antiparallel strand of chitin (Atkins, 1985). γ-chitin could be converted to α-chitin by use of lithium thiocyanate (Rudall, 1973). Conversely, chitosan is a homopolymer of β-(1→4)-linked N-acetyl-dglucosamine; it is soluble in most diluted acids and is obtained by removing acetyl groups (CH3-CO) from chitin (Majekodunmi 2016). Thus the major difference between chitin and chitosan is the content of acetyl in individual polymer.

11.3.3.7 Fucoidan A quantitative method for obtaining the purified form of fucoidan from the crude extract utilizes two methods that are optimized for the detection of fucose: a colorimetric method and an enzymatic method. Both methods are specific for the detection of fucose. In the colorimetric method, harsh acidic conditions are used that depolymerize and desulfate fucoidan into monomeric fucose (Ragan and Jensen 1979), and then fucose is directly measured at specified wavelengths. For the second, enzymatic, method, a commercially

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available l-fucose kit was used to specifically quantify fucose in the presence of other sugars and impurities. In this method, the enzyme l-fucose dehydrogenase reduces NADP+ to NADPH. The amount of NADPH in this reaction is stoichiometrically equal to the amount of l-fucose in the solution. The reduced NADPH in this reaction results in an increase in the absorbance at 340 nm (Talha 2015). A colorimetric method for detection of fucose (methylpentoses) in the presence of other sugars, as developed by Dische and Shettles (1948), involves treatment of the sugars with concentrated sulfuric acid (85%) and heating the solution at 100°C for 10 minutes, and on cooling, cysteine hydrochloric acid is added. This harsh acidic treatment converts the fucose residues into 5-methylfurfural, which on cooling, reacts with cysteine to form a fucose–cysteine complex that gives a pale yellow color. This method is called the Dische method. An enzymatic method that uses a commercially available l-fucose assay kit has also been used to detect l-fucose, and this proceeds by oxidization of the l-fucose in the sample by the enzyme l-fucose dehydrogenase in the presence of nicotinamide–adenine dinucleotide phosphate (NADP+) to form l-fucono-1,5 lactone with formation of reduced nicotinamide–adenine dinucleotide phosphate (NADPH) (Talha 2015). The enzymatic method is more specific toward the detection of fucose than the Dische method because of its simple depolymerization of fucoidan (a fucan) to fucose, which is easily detected. Although the structures of fucoidans may vary from one brown algae species to another, they are composed mainly of fucose and sulfate. For example, fucoidan prepared from Fucus vesiculosus is commercially available and is composed of 44.1% fucose, 26.3% sulfate, 31.1% ash, and a little aminoglucose (Li et al. 2008) After methylation and alkali treatment, the main component fucoidan unit was found to be 1,2-α-fucose, and most of sulfate groups were located at position C4 of the fucose units (Li et al. 2008). The structural model obtained from the GCMS analysis data of this methylation shows that the core region of fucoidan is primarily a polymer of α-(1→3)-linked fucose with sulfate groups substituted at the C4 position on some of the fucose residues; fucose also attaches to the primary chain to form branched points, with one branch at every two to three fucose residues within the primary chain (see Figure 11.22). It has been further shown that fucoidan may not be pure fucan sulfate but the heteropolymer of fucose, galactose, and a trace amount of xylose (Li et al. 2008). Subsequently, other sugars such as mannose, glucose, xylose, and glucuronic acid (GlcA) had been found in fucoidans from different brown seaweeds (see Table 11.4), which increased the difficulty of structural analysis. The position of sulfate groups is significant to the biological activities of fucoidans. The IR spectra showed that most sulfate groups were in axial positions and the remainder were in equatorial positions according to a strong band at 842 cm−1 and a shoulder at Fucα1

3Fucα1 4 Fucα1

3Fucα1

3Fucα1 4 –

SO3

3Fucα1

2 3Fucα1 4 –

SO3

Figure 11.22  Structural model from methylation study of fucoidan (adapted from Li et al. 2008).

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Enzymatic Technologies for Marine Polysaccharides Table 11.4  Chemical compositions of some fucoidans

Brown seaweed

Chemical composition

F. vesiculosus F. evanescens Lessonia vadosa Undaria pinnatifida Ascophyllum nodosum Himanthalia lorea and Bifurcaria bifurcate Padina pavonia Laminaria angustata Dictyota menstrualis Spatoglossum schroederi Sargassum stenophyllum

Fucose, sulfate Fucose, sulfate, acetate Fucose, sulfate Fucose, galactose Fucose, xylose, GlcA, sulfate Fucose, xylose, GlcA, sulfate Fucose, xylose, mannose, glucose, galactose, sulfate Fucose, galactose, sulfate Fucose, xylose, uronic acid, galactose, sulfate Fucose, xylose, galactose, sulfate Fucose, galactose, mannose, GlcA, glucose, xylose, sulfate Fucose, galactose, mannose, xylose, GlcA, sulfate Fucose, galactose, mannose, sulfate Fucose, galactose, mannose, xylose, GlcA, sulfate

Ecklonia kurome Adenocytis utricularis Hizikia fusiforme

820 cm−1 in the spectra. Although this method is commonly used in determining the sulfate position, comparing the stability of sulfate esters IR analysis data with those of alkali and methylation IR analysis in order to determine the sulfate position is also essential so as to avoid wrong conclusions because besides the C—O—S vibration in 820–850 cm−1 region, there is also C—H bending vibration of the sugar-reducing end, which affected the judgment on the position of the sulfate group (Li et al. 2008). Consequently, the use of electrospray ionization trap mass spectrometry (ESI-MS) and capillary electrophoresis has been valuable in determination of the sulfate position as shown experimentally in three isomers: 2-O-, 3-O-, and 4-O-sulfated fucose. Data revealed that it was possible to differentiate between these three positional isomers of sulfated fucose according to their fragmentation pattern (Tissot et al. 2006). Furthermore, the use of NMR to study the conformational behavior of fucoidan fragments as related to the sulfate groups at positions C2 and C4 shows that O-sulfation of (1→3)-linked oligofucosides hampers and alters their conformational flexibility and equilibrium, respectively, when compared with nonsulfated parent oligosaccharides (Li et al. 2008).

11.3.3.8 Porphyran The purification of porphyran oligosaccharides was carried out by preparative SEC (sizeexclusion chromatography) and purity ascertained by comparing with fractions collected with standard oligosachharides (Correc et al. 2011). Gel filtration on Sepharose had also been utilized in porphyran purification (Nishide et al. 1988). The crude polysaccharide preparation is separated using a DEAE ion-exchange column (2.6 cm × 30 cm), followed with a Sephacryl S-300 gel filtration column (1.6 cm × 100 cm). Using combined methods of composition analysis, methylation analysis, IR, and NMR, porphyran has been further purified and characterized (Liu et al. 2013). Porphyran in D2O solution is monomeric, and this form is suggested to be the imino N-protonated porphyran according to the results obtained from 13C NMR chemical shift calculation and prediction (Chuang et al. 2014). It was revealed that porphyrin, although

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257 –

OSO3 OH H

CH2OH H

O H H

H

H 4

O β1

OH

O

H

O O

H HO

H

α1 OH

H

Figure 11.23  Chemical structure of the repeating units of Porphyran molecules (Rhein-Knudsen et al. 2015).

positively charged in the solution, may be negatively charged or carry no charge at all in organisms. Nuclear Overhauser effect spectroscopy (NOESY) was therefore conducted to determine the structural conformations of the ionic and neutral forms of porphyran, to establish whether one or two protons are ionized from the carboxylic acid groups and also to clarify the correlation between the structure and energy interconversion (Liu et al. 2013). For the proton NMR, porphyran polysaccharide spectra were recorded at 70°C using 64 scans. The oligosaccharide spectra were recorded at 25°C using 16 scans. Chemical shifts are expressed in ppm in reference to the external standard TSP (trimethylsilylpropionic acid). The NMR signals of the porphyran disaccharide were fully assigned, using a complete set of correlation spectrums: COSY (double-quantum-filtered correlation spectroscopy), HMQC (heteronuclear multiple quantum correlation), and HMBC (heteronuclear multiple bond correlation). Other oligosaccharides were characterized using the same techniques and compared to the structure obtained for the porphyran disaccharide (Correc et al. 2011). Porphyran was found basically to be composed of D-galactose, L-galactose, 3,6-anhydrol-galactose, 6-0-methyl-d-galactose and ester sulphate, some of the ester being present as 1-4- linked L-galactose 6-sulphate (Turvey and Rees 1961). Predominately, porphyran has half of its sugar content as 3,6-anhydro-l-galactose and l-galactose 6-sulfatetotal, while the other half is made up of d-galactose and 6-O-methyl-d-galactose (Rees and Conway 1962). From 13C NMR spectroscopy, porphyran was found to be made up of sulfated disaccharide units, as shown in Figure 11.23 (Rhein-Knudsen et al. 2015).

11.4  Biological activities The extraction and purification of sulfated polysaccharides (sPSs) from their respective sources are quite expensive processes, and their utilization is justified only for important studies. However, the biological activities that they exhibit have become the major rationale for their continued extraction and purification. Numerous investigations toward utilization of these natural bioactive compounds for therapeutic applications have yielded affirmative results. Thus, sPSs promise effective treatment solutions to the growing number of individuals suffering from different types of ailments and diseases, countering antibiotics resistance, due to identification and isolation of different strains of viruses and bacteria. These research results have reduced the secondary effects usually caused by use of synthetic drugs and other traditional types of treatment used to prevent or control illnesses and conditions such as cancer, hypertension, diabetes, and obesity. Some of the biological activities of sSPs are discussed below.

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11.4.1  Antiviral activities Sulfated polysaccharide (sPS) antiviral activities have had a major impact on herpes simplex virus (HSV), human immunodeficiency virus (HIV), the human papilloma virus (HPV), the encephalomyocarditis virus, the hepatitis types A and B viruses, and the Dengue and yellow fever virus. Inhibitions of infections by most of these viruses were explained by the action of sPSs, which could have blocked the attachment of virions to the host cell surfaces (Baba et al. 1988). Furthermore, molecular weight (MW) could have played an important role in sPS antiviral properties, with the effect increasing as the MW increases; however, other structural features such as sulfation patterns, composition, and distribution of sugar residues along the backbone, and the complexity of the polymers could have been co-responsible for the reinforcement of the antiviral effectiveness (Raposo et al. 2015). The fucoidans from Laminaria japonica have been proven effective in fighting both RNA and DNA viruses, such as poliovirus III, adenovirus III, ECHO6 virus, and coxsackie B3 and A16 viruses, thus protecting host cells by inhibiting the cytopathic activity of these viruses (Li et al. 1995). Some fucoidans, apart from inhibiting attachment of virus particles to host cells, were found to inhibit attachment of human spermatozoids to the zona pellucida of oocytes; thus this property can be employed towards the development of a contraceptive gel with microbicidal activity (Smith 2004). Other action exerted by fucoidans and other sPS is the inhibition of the syncytium formation induced by viruses (Raposo et al. 2015). Furthermore, the antiviral activity of the sPS may also depend on the culture medium, algal strain, and cell line and methodology used for testing and the degree of sulfation, as exemplified in Porphyridium cruentum (Raposo et al. 2015). Viruses must interact with some glycosaminoglycan receptors (GAG), such as heparin sulfate (HS) before they can infect cells (Esko and Selleck 2002). The GAGs are proteins found in the intracellular matrix of various connective and muscle tissues and can covalently bind to other target cell surfaces. sPS may impair the attachment of the virus particles by competing for those GAG receptors (Heaney-Kieras et al. 1977). Virus attachment to host cells could be disrupted by the anionic components of the sPS linking with the basic groups of the glycoproteins of the virus. Thus sPS blocks the interaction of the virus–host cell receptor. Fucoidan (Hidari et al. 2008), ulvan (Cassolato et al. 2008), chitosan (Ikram et al. 2014), and carrageenans (Cunha and Grenha 2016) have all been found to have antiviral properties; thus, antiviral activity of sPS is due largely to interaction between their structural components, which also include content and distribution of sulfate groups along the polysaccharide backbone, molecular weight, sugar residue composition, and stereochemistry.

11.4.2  Antibacterial activities The antimicrobial activity of sPS differs according to whether the bacteria in question are Gram-positive or Gram-negative. For example, in Gram-positive Staphylococcus aureus the antimicrobial activity increases with increasing molecular weight of chitosan; however, in Gram-negative Escherichia coli, the antimicrobial activity increased on decreasing molecular weight (Ikram et al. 2014). Chitosan was shown to have stronger effects on Gram-positive bacteria such as Bacillus cereus, Staphylococcus aureus, Lactobacillus plantarum, Listeria monocytogenes, Bacillus megaterium, L. brevis, and L. bulgaris than for Gram-negative bacteria such as Salmonella typhymurium, E. coli, Pseudomonas fluorescens, and Vibrio parahaemolyticus (Ikram et al. 2014). This antimicrobial activity was due largely to the fact that

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the solubility of chitosan increases in acidic medium, with higher molecular weight and degree of acetylation (Ikram et al. 2014). Other sPS such as fucoidan from L. japonica and sodium alginate were found to inhibit E. coli, by adhering to the bacteria cell consequently killing them as exemplified by Helicobacter pylori, colonies of which, in clinical trial studies, were eradicated, with restoration of stomach mucosa (Raposo et al. 2015). Carrageenans and sulfated exopolysaccharide (sEPS) from the red microalga Porphyridium cruentum showed a significant inhibitory activity against S. enteritidis (Yamashita et al. 2001; Raposo et al. 2014). Laminarans were found to induce antibacterial responses in rats and some resistance to mammal organisms toward infections by E. coli. (Raposo et al. 2015).

11.4.3  Antifungal activities The antifungal mechanism of sPS has been attributed to direct interference with cell growth. The antifungal effects of chitosan, which directly impedes fungal growth, have been shown to be similar to the effects observed in bacteria cells. Microscopic study shows that chitosan oligomers diffuse inside hyphae, interfering with the enzymatic activity responsible for fungus growth (Eweis et al. 2006). Thus, this action is deemed to be fungistatic rather than fungicidal, involving metabolic regulatory changes (Ikram et al. 2014). Laminarans have also been shown to induce antifungal responses in rats (Rice et al. 2005), while fungistatic properties of alginate have been applied in improving fiber quality and wound-dressing products (Rinaudo 2014).

11.4.4  Anticoagulant and antithrombotic activities The sPS not only are potent anticoagulants, acting by directly increasing the clotting time by inhibiting the contact activation pathway (intrinsic pathway), but also show potent antithrombotic bioactivity by inhibiting the heparin cofactor II–mediated action of thrombin (Li et al. 2012). The activities of these sPSs have been connected to their chemical and structural features, the amount and distribution pattern of sulfate, the composition and distribution of monosaccharides and their glycosidic bonds, and also their molecular weight, which may play a major role in the coagulation and platelet aggregation processes (Li et al. 2008; Silva et al. 2010). Thus the higher the content of these factors, the higher the anticoagulant antithrombotic activity (Li et al. 2008; Silva et al. 2010; Li et al. 2012). Fucoidan molecular weight was linked to the anticoagulant activity as high molecular weight (320,000 Da) extracted from Lessonia vadosa showed high anticoagulant activity, while lower molecular weight (32,000 Da) showed low anticoagulant activity (Chandía and Matsuhiro 2008). The sugar composition and sulfate groups attached to the sugar were also shown to regulate fucoidan anticoagulant activity (Nishino et al. 1989; Li et al. 2008). Laminarin, a nonsulphated polysaccharide, was shown to possess blood anticoagulant activity on sulfation (Hoffman et al. 1982; Shanmugam and Mody 2000). Laminarin sulfate was found to exhibit about 30% of the anticoagulant activity of heparin and was therapeutically effective in the prevention and treatment of cerebrovascular diseases (Miao et al. 1995). The anticoagulant action mechanisms of sPS may therefore be attributed to either direct inhibition of thrombin or activation of antithrombin III (AT-III) (Matsubara et al. 2000), or to an increase in the activity of thrombin inhibitors, such as AT-III and/or heparin cofactor II (HC-II), in both the intrinsic (contact activation or normal, measured by APPT test) and extrinsic [tissue factor (TF), measured by PT test] pathways (Raposo et al. 2015). Thrombin

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inhibition could be linked to binding of sPS to thrombin, rendering it inactive, or to inhibit thrombin from linking to the protease activated receptor-1 and GP-1b receptors on human platelets (Raposo et al. 2015).

11.4.5  Antiproliferative, tumor suppressor, apoptotic, and cytotoxicity activities The sPS are excellent antiproliferative, tumor-suppressing, apoptotic, and cytotoxic agents. Sulfated fucoidan from Cladosiphon okamuranus showed antiproliferative activity in myeloid cancer and leukemia cell lines, inducing apoptosis (Cho et al. 2010; Raposo et al. 2015), while that from L. guryanovae inactivated the epidermal growth factor (tyrosine kinase) receptor (EGFR), which is significantly involved in cell transformation, differentiation, and proliferation (Khotimchenko 2010). Thus fucoidan is assumed to be useful as a therapeutic or preventive agent against cancer and other tumors as it was shown to selectively destroy lymphoblastoids cancer cells without damaging the normal cells (Ohigashi et al. 1992; Khotimchenko 2010). This action could have been achieved by boosting cell immunity, inducing apoptosis, and inhibiting angiogenesis processes, cutting the supply of nutrients to the cancer cells (Teas and Zhang 2006). Furthermore, fucoidan has antiadhesive properties that may well clarify their antimetastatic activity in vitro and in vivo, in various animal models, as they inhibit the adhesion of tumor cells to platelets, reducing the tendencies of proliferation of neoplastic cells (Teruya et al. 2009). Chitosan oligomers have also demonstrated antineoplastic activity by inducing apoptosis, inhibiting growth of tumor cells, and inhibiting tumor-induced angiogenesis and metastasis (Murata et al. 1989, 1991; Carreno-Gomez and Duncan 1997; Chen et al. 1997; Sayari et al. 2016). Prophyran and sulphated-galatan porphyran obtained from Porphyra yezoensis and Porphyra spp., respectively, showed antitumor activity (Kwon and Nam 2006; Raposo et al. 2015). While laminarin induces apoptosis in human colon cancer, low-ester-sulfate-content sodium alginate was shown to inhibit the growth of sarcoma 180 in mice and to chemically induce intestinal carcinogenesis in rats (Fujihara et al. 1984; Yamamoto and Maruyama 1985; Miao et al. 1999; Ji et al. 2012). Lymphocytic and macrophage activities are shown to be enhanced by sPS by activation of signaling receptors in their membranes [cluster of differentiation 14 (CD14), competent receptor-3 (CR-3), scavenging receptor (SR), and Toll-like receptor-4 (TLR-4)] and also activation of other intracellular pathways, involving protein kinases, which enhances nitric oxide (NO) production significantly involved in apoptosis (Zhou et al. 2004; Teruya et al. 2009). Sulfonation and molecular weight of sPS also play a major role in antineoplastic processes as oversulfated PS showed great ability to inhibit the growth of leukemia tumors in mice (Yamamoto et al. 1984); likewise, low-molecular-weight PS derivatives enhanced antitumor activity (Yang et al. 2008).

11.4.6  Anti-inflammatory and immunomodulatory activities Fucoidan from D. menstrualis was shown to decrease inflammation by directly binding to the cell surface of leukocytes, in particular polymorphonuclear cells (PMNs). It was shown in injured mouse tissue to totally inhibit the movement of leukocytes into the peritoneal cavity after the tissues were subjected to simulated pain and inflammation, without the production of proinflammatory cytokines (Preobrazhenskaya et al. 1997; Albuquerque et al. 2013). Kang et al. (2011) showed that anti-inflammatory activity was associated with

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the immunomodulatory effect in peritoneal macrophage primary cells, due to the action of fucoidan from E. cava, which inhibits both enzyme nitric oxide synthase and cyclooxygenase-2 (COX-2), consequently preventing the production of nitric oxide (NO) and prostaglandin E2 (PGL2). The immunomodulatory effect of fucoidan was further verified by a study showing that it could be used in the treatment of severe inflammation by reversing the excessive release of cytokines and proteinases, which are involved in rheumatoid arthritis, chronic wounds, or leg ulcers (Senni et al. 2006). Further studies showed that it could act as immunostimulators by activating macrophages and lymphocytes, resulting in effective cancer immunoprevention (Choi et al. 2005). sPS from Ulva rigida, a green seaweed, also stimulates the production of cytokines in macrophage, thus suppressing the growth and proliferation of tumor cells, and are as such referred to as “natural neoplastic-cell killers” (Raposo et al. 2015). Chitin was found to have immunopotentiating activity by enhancing immune responses by increasing phagocytosis, production of osteopontin, leukotriene B (via polymorphonuclear leukocytes stimulation), interferon-γ, interleukin-1 and -8 (via fibroblasts stimulation), transforming growth factor b1, and platelet-derived growth factor (via macrophage stimulation) (Ueno et al. 2001). λ-Carrageenan was shown to stimulate T-helper 1 (Th1)-patterned cytokine in mouse T-cell cultures (Tsuji et al. 2003). It also stimulates the activity of macrophages, enhances production of interferon-γ response in TLR4-deficient mice, and reduces allergic reactions in mice immunized with ovalbumin by reducing ovalbumin-specific IgE and serum histamine release (Tsuji et al. 2003; Prajapati et al. 2014; Raposo et al. 2015). Laminarin was shown to alter systemic inflammation response through the release of inflammatory mediators such as calcium, nitric oxide, hydrogen peroxide, vascular endothelial growth factor, monocyte chemotactic protein-1, leukaemia inhibitory factor, and granulocyte-colony-stimulating factor, enhancing signal transducer expression and activation of transcription and cyclooxygenase-2 gene (Neyrinck et al. 2007; Lee et al. 2012).

11.4.7 Antilipidemic (hypocholesterolemic and hypotriglyceridemic), hypoglycemic, and hypotensive activities The inhibitory effect of seaweeds sPS of human pancreatic cholesterol esterase, an enzyme that promotes intestinal cholesterol and fatty-acid absorption, is potentiated by degree of sulfation and higher molecular weights (Laurienzo 2010). Sulfated ulvan from U. pertusa using a mouse model study regulated the HDL : LDL ratio and reduced the levels of triglycerides (TG) in serum (Yu et al. 2003). Antiperoxidative activity has also been reported for sulfated ulvan animal models as it protects liver tissues from hyperlipidaemia, reduces oxidative stress, and improves antioxidant activities (Raposo et al. 2015). Native or low-molecular-weight derivative fucoidan from L. japonica showed hypolipidaemic activity by decreasing total and LDL cholesterol and TG in rat serum and prevented hypercholesterolaemia in mice (Raposo et al. 2015). It was shown that fucoidan prevents the artery-clogging buildup of fatty acid in the blood vessels, and thus reduces hypertension and heart-related illness (Li et al. 2008; Huang et al. 2010). Porphyran from P. yezoensis was shown to proffer antihyperlipidaemic activity in human liver cultured cells by both reducing the release of apolipoprotein-B100 (apoB100) and synthesis of lipids. It therefore has the potential to be used as a therapeutic agent for treatment of cardiovascular diseases (CVDs) (Raposo et al. 2015). The κ- and

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λ-carrageenans from G. radula also were shown to reduce serum cholesterol and triglyceride levels in humans and rats fed κ-/λ-carrageenan-enriched diets (Raposo et al. 2015). Laminarin has been shown to reduce total and LDL cholesterol, TG, and systolic blood pressure levels, and act against irradiation in liver cells (Holdt and Kraan 2011). Chitin and chitosan were also shown to be excellent cholesterol-lowering and fat blockers as they increased the HDL cholesterol: LDL cholesterol ratio (Koide 1998). Administration of chitosan to genetically obese diabetic male KK-Ay mice resulted in lowering of the serum glucose levels and reduced their overdrinking; thus chitosan is assumed to be useful in the treatment of obesity-related diabetes mellitus (Do et al. 2008). Likewise, ulvans showed antilipedemic activity, reduced total serum cholesterol and triglycerides, but elevated the HDL cholesterol : LDL cholesterol ratio (Pengzhan et al. 2003; Jiao et al. 2011; Qia et al. 2012).

11.4.8  Antioxidant activity Typically marine sPSs from seaweed exert their primary antioxidant action by scavenging free-radicals and reactive chemical species such as superoxide ion (O2−), hydroxyl ion (OH−), and hydrogen peroxide (H2O2), which act as strong reducing agents and as metal chelators (Wang et al. 2014). Fucoidan has been shown to be a natural antioxidant with a high potential to reduce free-radical-associated illness (Li et al. 2008; Wang et al. 2008). Its antioxidant activity is higher than that of vitamin C and κ-carrageenan (Wang et al. 2008); however, the lower-molecular-weight derivative has demonstrated antioxidant activity higher than that of the higher-molecular-weight derivative, and this is likely due to its ability to easily integrate with the cell and donate protons (Wang et al. 2008; Wijesekara et al. 2011). The sulfate level is another important factor, as higher sulfate content gives higher antioxidant activity (Wang et al. 2008). It has been proposed that fucoidan could be useful in the food industry or in medicine as a therapeutic agent for managing free-radical-induced illnesses (Li et al. 2002; Rupérez et al. 2002). Laminarin has also be shown to be an excellent antioxidant, as incorporation into animal feed results in reduction in lipid oxidation in laminarin-fed pigs (Moroney et al. 2012). Highly acetylated chitosan shows high scavenging activity toward hydroxyl and superoxide radicals (Park et al. 2004a,b; Limam et al. 2011), likewise as the lowmolecular-weight derivative (Kim and Thomas 2007). High sulfate content is shown to influence the antioxidant potentials of ulvans (Qi et al. 2005) and carrageenans (Rocha de Souza et al. 2007).

Conclusion The heterogeneity in these marine polysaccharides requires divergent extraction protocols, which, in turn, gives rise to different purity levels. This is a challenge that should be thoroughly investigated, as purity level is synonymous with the type and quality of biological effects that these marine polysaccharides they possess. Hence, there is a need for protocols that will qualitatively and quantitatively extract these polysaccharides from seaweed and yet retain their functional moieties, ensuring that their optimal biological activities will be harnessed. Also, specific structural modifications that could potentially increase their biological effects should also be exploited. As an increasing number of these polysaccharides are discovered, extracted, and purified to the highest degree, their applications to human issues should be more involved in areas of medical, pharmaceutical, food, and other industrial applications.

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chapter twelve

Fucoidan A tool for molecular diagnosis and targeted therapy of cardiovascular diseases Murielle Maire1, Lucas Chollet1,2 , Lydia Rolland2 , Didier Letourneur3, Cédric Chauvierre3, and Frédéric Chaubet1

Galilée Institute, Paris 13 University, Villetaneuse, France Algues & Mer, Kernigou, Ouessant, France 3Inserm, U1148, LVTS, Paris Diderot University, Paris 13 University, Paris, France 1 2

Contents 12.1 Introduction....................................................................................................................... 273 12.2 A brief overview of atherosclerosis................................................................................ 274 12.3 Medical imaging of cardiovascular diseases: reality and challenges....................... 275 12.4 Fucoidan and P-selectin: toward a biospecific contrast agent for CVD imaging............................................................................................................. 276 12.4.1 P-selectin............................................................................................................... 276 12.4.2 Fucoidan: origin and structure......................................................................... 276 12.4.3 Biological activities of fucoidan........................................................................ 279 12.4.4 P-selectin/fucoidan interaction......................................................................... 279 12.4.5 A fucoidan-based contrast agent for vascular imaging................................ 280 12.5 A fucoidan fraction for the clinical development of a scintigraphic contrast agent of vascular pathologies..........................................................................................284 12.5.1 Polysaccharides cannot yet be fully synthesized...........................................284 12.5.2 Fucoidan extraction from brown algae must be secured.............................. 285 12.6 How fucoidan helps to fight stroke................................................................................ 287 Conclusion.................................................................................................................................... 291 Acknowledgments....................................................................................................................... 292 References...................................................................................................................................... 292

12.1 Introduction According to the World Health Organization, cardiovascular diseases (CVDs), and their consequences, are responsible of more than 25% of the deaths in the world (Finegold et al. 2013; Go et al. 2014). Atherosclerosis is the most widespread CVD. Lipid-rich plaques build up inside large and medium arteries. The process begins as soon as childhood and evolves according to environmental and genetic factors (Jackson 2011; Lusis 2000; Bentzon et al. 2014; Falk et al. 2013). Massive plaque rupture or arterial intraplaque hemorrhage may induce a thrombus formation (namely, atherothrombosis) with dramatic consequences such as acute coronary syndrome, stroke, and limb ischemia 273

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Enzymatic Technologies for Marine Polysaccharides

(Willerson and Ridker 2004; Kaperonis et al. 2006; Libby 2002). The silent and asymptomatic progression of atherosclerosis remains very difficult to diagnose without the help of imaging techniques associated with contrast agents to improve the quality and the specificity of the images. The large amount of P-selectin, a glycoprotein overexpressed at the surface of activated platelets in thrombi and on inflamed endothelium in the pathological area, makes it a relevant marker for the molecular imaging of the vascular inflammation and of venous and arterial thrombosis as well as of transitory episodes of ischemia and for the treatment of CVD with drug-targeting systems (Ley 2003; Choudhury and Fisher 2009). So far, no biospecific clinical product is available for diagnostic imaging of thrombosis or vascular inflammation in relation to the development of CVD. Sulfated polysaccharides are able to bind to P-selectin, including fucoidan, a sulfated poly-l-fucose extracted mainly from brown seaweeds (Varki 1994). For the last 10 years a fucoidan from the brown seaweed Ascophyllum nodosum (An) allowed imaging of thrombosis and endothelial inflammation in animal models by scintigraphy (Rouzet et al. 2011; Bonnard et al. 2014), Magnetic resonance imaging (MRI) (Suzuki et al. 2015) and ultrasonography (Li et al. 2017). A scintigraphic kit for 99mTc-SPECT imaging of atherothrombosis and ischemia developed with a low-molecular-weight fraction of this fucoidan is currently in Phase I clinical trial in the framework of the large-scale European project Nanoathero (Chauvierre and Letourneur 2015). Finally, some therapeutic advances could also be obtained by using fucoidan to potentiate the effect of tissue plasminogen activator, a drug currently used in the emergency treatment of stroke (Juenet et al. 2018; Ghebouli et al. 2018).

12.2  A brief overview of atherosclerosis The vascular wall consists of a monolayer of endothelial cells in contact with blood, anchored on a collagen matrix: the intima. A sleeve of smooth muscle cells, the media, surrounds the intima, ensuring in particular the mechanical integrity of the vessel. At the periphery of the intima, fibroblastic cells form the feeding adventice. Atherosclerosis causes a remodeling of the wall of large and medium-sized arteries from the infiltration of blood-circulating lipids between the intima and the media. A necrotic core is formed, the atheroma, covered with a fibrous cap of connective tissue isolating the core from the arterial lumen, the sclerosis (Libby et al. 2002; Newby and Zaltsman 1999). The whole, so-called the atherosclerotic plaque, is a complex biological object in permanent evolution. The protease activity that develops within the plaque degrades the collagen, increasing the risk of rupture according to hemodynamic conditions (Sukhova et al. 1999). When a plaque ruptures, its content is exposed to blood circulation leading to the formation of a thrombus mainly composed of an aggregate of activated platelets consolidated by a network of long fibrin chains (Davies et al. 1993; Michel et al. 2014). This thrombus can decrease the arterial lumen (stenosis) or completely obstruct the artery (occlusive thrombus). The stenosis or the occlusion of an artery leads to an ischemic situation for the tissues downstream because they are no longer irrigated by the blood (Pelisek et al. 2012). Numerous studies describe this process, from atherogenesis to plaque rupture and thrombosis (Choudhury et al. 2004; Stary et al. 1992, 1994, 1995; Libby 2002; Libby et al. 2002). Inflammatory processes within the intima activate the endothelial cells that express cellular adhesion proteins on their surfaces (Libby 2002; Furie and Furie 2004), including P-selectin. These proteins have the role of capturing circulating leukocytes via interaction with their ligands present on their membrane, allowing their infiltration under the media and the maintenance of the process.

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12.3 Medical imaging of cardiovascular diseases: reality and challenges Medical imaging is the most widespread tool for the diagnosis of atherothrombosis. The various imaging techniques of the vascular space essentially allow visualization of the circulation of the blood flow in the arterial lumen in order to identify stenoses and aneurysms (consequences of a mechanical weakening of the artery wall leading to a dramatic increase in its diameter) or to evaluate the perfusion of tissues to identify ischemic or infarcted zones. However, they do not allow detecting the presence of biological markers, nor the early diagnosis of atherothrombotic lesions, and multiple examinations may be necessary to delineate the pathological areas. All the modalities are not adapted to the different parts of the body possibly involved, and the imaging of anatomical evolution of the plaques is not necessarily representative of their state of vulnerability. Figure 12.1 summarizes the advantages and disadvantages of the main imaging modalities (Patel et al. 2014; Rahmim and Zaidi 2008; Quillard and Libby 2012; Mankoff 2007; Sanz and Fayad 2008; Wildgruber et al. 2013; Orbay et al. 2013). Among them MRI is by far the most widely used and is considered a gold standard in clinical settings for detecting atherothrombosis. Molecular imaging is a promising development dedicated to visualization of the biological processes at the cellular and molecular levels owing to the use of probes designed to recognize a local target and bind it specifically (Herrmann et al. 2012; Smith and Gambhir 2017; Spatial resolution of imagery

1mm

SPECT

PET

MRI

Ultrasonography Scanner Fluorescence

2mm

4mm

Detection sensitivity of contrast agents Fluorescence PET SPECT

nmolar Technique Computed tomography (CT scan) Scanner

Modality

Ultrasonography

µmolar

Usual contrast agents

X Ray

Scintigraphy (single-photon emission computed tomography SPECT) and positron emission tomography (PET))

Radioactivity

Magnetic resonance imaging

Nuclear magnetic resonance

Ultrasonography

Ultrasounds

Fluorescence imaging

Fluorescence

Scanner

MRI

PET : Fluorine-18 (18F) Gallium - 68 (68Ga) SPECT: technetium-99m (99mTc) Chelates of gadolinium (III) -Iron oxide based particles Small gas bubbles in albumin or fatty acids caps Perfluorooctylbromide Fluorophore (FITC, Alexa ...)

mmolar Strength

Limitations

High resolution - Fast

Ionizing radiation

Sensitivity

Ionizing radiation - Low resolution

High resolution - no ionizing radiation

Low sensitivity

Low cost - Availability Transportable

Low quality of image

Sensitivity - High resolution

Limited penetration in tissues

Figure 12.1  The main medical imaging modalities with a comparison of their characteristics.

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Enzymatic Technologies for Marine Polysaccharides

Chen et al. 2016). Therefore, the main challenge of medical imaging in the cardiovascular field is the appreciation of molecular and cellular processes of atherosclerosis, thrombosis, and vascular inflammation with the use of contrast agents able to target a relevant biomarker of the pathology. More generally, molecular imaging faces challenges in both the diagnostic and therapeutic fields when a drug is linked to the contrast agent, which makes it possible to monitor and understand the biological mechanisms involved (Choudhury and Fisher 2009; Lobatto et al. 2011).

12.4 Fucoidan and P-selectin: toward a biospecific contrast agent for CVD imaging Many studies have focused on the development of contrast agents targeting atheromatous plaque at different stages of the pathology (Jacobin-Valat et al. 2011; Kaufmann et al. 2007; Choudhury and Fisher 2009; McAteer and Choudhury 2013). However, it seems more clinically relevant to develop a thrombus diagnostic technique because 70% of deaths due to myocardial infarction are caused by atherothrombotic development (Patel et al. 2014). Two key components of the thrombus are promising for the development of a molecular imaging probe: fibrin and activated platelets (de Haas et al. 2014; Stratton et al. 1981). However, in the case of transient ischemic attacks, a thrombus can disappear rapidly after having formed with symptoms lasting only a few minutes to 24 hours (Easton et al. 2009). These ischemic accidents induce an inflammatory reaction and activate the surrounding endothelial cells. P-selectin, expressed on the surfaces of both activated platelets and activated endothelium, would therefore be a relevant target for the molecular imaging of thrombosis as well as of ischemic situations as it persists on the surface of the endothelium for hours after disappearance of the transient thrombus (Porter 2007).

12.4.1 P-selectin Protein ligands have been developed to target P-selectin. Iron oxide microparticles for MRI (McAteer et al. 2012) and microbubbles for ultrasonography (Kaufmann et al. 2007) were coupled to murine antibodies, to P-selectin glycoprotein ligand-1 (PSGL-1), the main natural ligand of P-selectin (Bettinger et al. 2012), or a P-selectin-binding peptide (Molenaar et al. 2002) to diagnose endothelial inflammation after stroke with MRI (Jin et al. 2009). Saccharide ligands have also been studied, in particular the tetrasaccharide sialyl Lewis X (SLeX) involved in the interaction between P-selectin and PSGL-1 (Ferrante et al. 2009; Jin et al. 2010). The participation of sulfated tyrosine carried by the peptide chain of PSGL-1 is, however, necessary for an optimal interaction (Vestweber and Blanks 1999; Brandley et al. 1993; Ramachandran et al. 1999) (Figure 12.2). Varki has shown that several sulfated polysaccharides can recognize selectins and more particularly P-selectin, by mimicking the action of SLeX: heparin, heparan sulfate, fucoidan, or even dextran sulfate (Varki 1994).

12.4.2  Fucoidan: origin and structure Fucoidan belongs to a family of marine sulfated polysaccharides termed fucans. The fucoidan molecule was identified for the first time in 1913 by Kylin as “fucoidin,” but has since been formally named fucoidan, according to the International Union of Pure

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Cytoplasmic domain

Cell membrane Peptide chain OSO3 OSO3

NH2

OSO3

Tyrosine sulfate Sialyl Lewis X

(A) P-selectin

Ca2+ OH HO OH

H3C OH +

HN3

Lys 113 OH HO

OOC

O O

OH O O

O OH

NHAc O O OH

O

Tyr

NaO3SO

HO AcNH

PSGL-1

OH

Sialyl Lewis X

(B) Figure 12.2  (A) Schematic representation of structure of the glycoprotein PSGL-1, natural ligand of P-selectin. The chemical structures necessary for the interaction between PSGL-1 and P-selectin are SLeX and sulfate groups born by the peptide chain [adapted from Ley (2003) with permission). (B) Scheme depicting interaction between P-selectin and PSGL-1 implicating SLeX and sulfated tyrosines (Ramphal et al. 1994; Leppanen et al. 2000).

and Applied Chemistry (IUPAC) recommendations (Kylin 1913). The fucan family also includes ascophyllans (or xylofucoglycuronan and xylofucomanuronan) polysaccharides made up of a skeleton of polyuronic acids (glucuronic or mannuronic acid) with ramifications of xylose and fucose (Larsen et al. 1966; Leite et al. 1998; Bilan et al. 2014) and sargassans (or glycuronofucogalactan) other polysaccharides made up of galactose, fucose, and uronic acids (Medcalf et al. 1978; Mabeau et al. 1990). Fucoidans were found in more than 70 species of brown algae (Phaeophyceae) (Usov and Bilan 2009). Similar fucans were isolated also from about 10 species of marine invertebrates, for example, in

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Enzymatic Technologies for Marine Polysaccharides

R1 = SO3– ou H ou COCH3

R2 = SO3– ou H R3 =

O CH3 1 R O

O

CH3

O

O

R2O

R2O

O

R2O

CH3 OH

HO O

O

(A) Chorda fium

CH3

HO O

O

CH3

R1O

R2O O CH3

CH3 R2O

O

R1O

O3SO

HO

O

O

O

O

CH3 – O3SO

R1O

O

O

O

–O SO 3

O3SO

O

O OH

(D) Strongylocentrotus droebachiensis

HO

O CH3 – O3SO

O

CH3 –

CH3 –

R2O

O CH3 – O3SO

O

O

HO O

O

–O SO 3

CH3 –O SO 3

R3O

R2O O

CH3

(C) Ludwigothuria grisea

O

HO O

(E) Strongylocentrotus franciscanus

OSO3–

(B) Ascophyllum nodosum/ Fucus vesiculosis/Fucus evanescens

Figure 12.3  Repeating chemicals structures of some fucoidans from brown algae: (A) Chorda filum (Chizhov et al. 1999); (B) Ascophyllum nodosum, Fucus vesiculosis, and Fucus evanescens (Chevolot et al. 2001; Bilan et al. 2002; Chevolot et al. 1999) and from marine invertebrates; (C) Ludwigothuria grisea (Mulloy et al. 1994) (Holothuroidea); (D) Strongylocentrotus droebachiensis (Vilela-Silva et al. 2002); (E) Strongylocentrotus franciscanus (Echinoidea) (Vilela-Silva et al. 1999) [from Chollet et al. (2016)].

the jelly coat of sea urchin eggs (Echinoidea) (Vasseur et al. 1948; Alves et al. 1997) and in the body wall of sea cucumbers (Holothuriidae) (Mourão and Bastos 1987; Berteau and Mulloy 2003). To our knowledge, fucoidan was never isolated from other classes of marine algae (Chlorophyceae, Rhodophyceae), nor from freshwater algae or terrestrial plants. In the brown algae, fucoidans are contained in the amorphous phase of the cellular wall and likely play a key role in the adaptation to osmotic stress (Deniaud-Bouet et al. 2014; Mourão and Bastos 1987). In fact, the environmental conditions around the algae can strongly vary over very short durations, particularly in the case of salinity, which varies sharply with the tides and the temperature of seawater. It is thus necessary that organisms subjected to such conditions be able to quickly adjust their osmotic potential (Kirst 1990; Deniaud-Bouet et al. 2017). For the last hundred years fucoidan structures from different brown seaweeds have been widely investigated, highlighting their large variability, which is closely related not only to species (Figure 12.3) but also to ecophysiological parameters, location of culture, harvesting season, and even the method of extraction from the crude algae. The first structure was elucidated in 1950 by Conchie and Percival from a fucoidan from Fucus vesiculosus, a mainchain of α(1→2)-l-fucose with sulfate substitution on ring position C4 and occasionally an α(1→4) linkage (Conchie and Percival 1950). This model was accepted until 1993, when Patankar and coworkers published a revised structure: α(1→3)l-fucose linear backbone with sulfate substitution on C4 and some fucose branched at C4 or C2, one for every two or three fucose residues within the chain (Patankar et al. 1993). Fucoidans extracted from sea cucumbers or sea urchins have a more regular chemical structure: linear tri- or tetrasaccharides connected with the same glycosidic linkage, with each repeating unit defined by a distinctive pattern of sulfate substitution (Pomin 2012; Pomin and Mourao 2008; Mourão and Bastos 1987; Aquino et al. 2005). Besides fucose and sulfate, fucoidan may also contain some additional carbohydrates such as mannose, galactose, glucose, xylose, and also uronic acids, in much smaller amounts, sometimes substituted

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279

with acetyl groups (Duarte et al. 2001; Ponce et al. 2003; Bilan et al. 2002; Li et al. 2006; Cunha and Grenha 2016).

12.4.3  Biological activities of fucoidan The main interest in fucoidan lies within the large spectrum of biological activity, including anticoagulant (Nardella et al. 1996; Zaporozhets and Besednova 2016), antithrombotic (Pomin 2012; Zaporozhets and Besednova 2016), anti-inflammatory (Cumashi et al. 2007; Berteau and Mulloy 2003), anti- or proangiogenic (Liu et al. 2012; Ustyuzhanina et al. 2014), antitumor (Kwak 2014; Sanjeewa et al. 2017), anticomplementary (Blondin et al. 1994; Tissot and Daniel 2003), antidiabetic (Shan et al. 2015) and antiviral properties (Berteau and Mulloy 2003; Ponce et al. 2003; Wang et al. 2012; Shi et al. 2017). It is now widely accepted that levels of fucose and sulfate as well as molecular weight (MW) are major structural parameters whose variation affects the biological properties. One of the most striking examples is the anti/proangiogenic activity. Ustyuzhanina and coworkers reviewed numerous studies on the angiogenic activities of fucoidans from different brown algae to highlight structure–activity relationships. They could only conclude that fucoidan from An with MW over 30 kDa exhibited antiangiogenic activity, whereas fucoidan with MW lower than 30 kDa exhibited proangiogenic activity (Ustyuzhanina et al. 2014). So far, multiple targets have been identified in blood and tissues to explain these activities. For example, the anticoagulant activity, one of the most frequently studied with reference to heparin, can be explained by the interactions of fucoidan toward natural thrombin inhibitors, serpins antithrombin and heparin cofactor II, potentiating their activity (Pomin 2012).

12.4.4  P-selectin/fucoidan interaction The affinity for P-selectin of a low-molecular-weight fucoidan (LMWF) from An (7–10 kDa) was evaluated in purified systems with surface plasmon resonance (SPR) and surface acoustic wave (SAW) (Figure 12.4). With these technologies, the recorded signals are roughly proportional to the weight increase of the layer at the surface of the sensors. The difference with the response in absence of circulating species allows researchers to establish the kinetic parameters of the interaction and, following a simple Langmuir model, obtain the affinity constant. Fucoidan interacted strongly with P-selectin, independently of the technique used, and the signals were much more intense when LMWF (7–10 kDa) was immobilized and P-selectin (100 kDa) was injected in the running buffer. The interaction was first quantified by Bachelet et al. (2009) with P-selectin linked to the sensor chip providing a nanomolar affinity (Figure 12.4A), much higher than that of a heparin (577 nM) and a dextran sulfate (118 nM) of similar molecular weights. Moreover, LMWF inhibited the binding of SLeX and PSGL-1 to P-selectin with a half-maximal inhibitory concentration (IC50) of 20 nM as compared with heparin (400 nM) and dextran sulfate (>25,000 nM). Comparisons have been performed with L- and E-selectins and with VEGF165, a proangiogenic growth factor strongly potentiated by LMWF (Lake et al. 2006). The interaction of fucoidan-coated ultrasmall superparamagnetic iron oxide (USPIO-FUCO) with P-selectin was also demonstrated by SPR in purified system (Figure 12.4B) (Suzuki et al. 2015). The data clearly showed a dose-dependent binding of USPIO-FUCO to P-selectin, whereas no binding could be observed with carboxymethyldextran-coated USPIO, ruling out a simple effect of size or charge (carboxymethyldextran and fucoidan are both anionic polysaccharides, and coated USPIOs were of similar sizes).

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Enzymatic Technologies for Marine Polysaccharides

Response Unit

10 8 4

0 0

100

200

300

400

Response Unit (0=baseline)

30 5 20 4 10 0 1

–10

160

Time (s)

320

640

Time (s)

(A)

(B) 2.0

1200 1000

1.5

800 Phase (°)

Response (0–baseline)

480

3 2

VEGF P-selectin

600 400 200

0.5

L-selectin E-selectin

0 200

400

600

P-selectin

1.0

L and E-selectins

800

1000

0.0

0

50

100 150 200 250 300 350 400

Time (s)

Time (s)

(C)

(D)

Figure 12.4  (A) Surface plasmon resonance (SPR) sensorgrams showing the interaction of fucoidan with P-selectin (black) or with an immobilized IgG (gray = nonspecific control) (Bachelet et al. 2009). (B) SPR sensorgrams depicting the interaction between carboxymethyldextran-coated USPIO (USPIO-CMD) and fucoidan-coated USPIO (USPIO-FUCO) at 0.1 or 1 µM with immobilized P-selectin (1—buffer; 2—0.1 µM USPIO-CMD; 3—1 µM USPIO-CMD; 4—0.1 µM USPIO-FUCO; 5—1 µM USPIO-FUCO) [adapted from Suzuki et al. (2015) with permission n]. (C) SPR sensorgrams recorded with immobilized LMWF and circulating selectins and VEGF (D) SAW sensorgrams showing the interaction of immobilized LMWF with circulating selectins. Selectins were injected at a concentration of 100 nM.

12.4.5  A fucoidan-based contrast agent for vascular imaging Although MRI be considered as a gold standard for the examination of the vascular space, because of its great sensitivity, scintigraphy remains the first method of choice for molecular imaging used in clinical routine. Thus, Rouzet et al. radiolabeled the LMWF with technetium 99m (99mTc), and the 99mTc-fucoidan complex was validated as a SPECT molecular imaging agent of atherothrombosis and heart ischemia in different animal models expressing P-selectin: endocarditis, heart ischemia, and abdominal aorta aneurysm (Rouzet et al. 2011; Michel et al. 2010). Surprisingly, a simple combination of 99mTcO4 with LMWF in the presence of a reducing agent produced a stable radiolabeled complex for the duration of the examinations, specifically, for several hours.

Chapter twelve:  Fucoidans in medicine

281

With starting fucoidan

With Ca-purified fucoidan TLC front

TLC front

50

500 40

200

30

0.1%

20

97.1% c/KeV * 1000

300

c/KeV * 1000

99.9% 400

2.9%

10

100

0

0 (A)

(B)

Figure 12.5  Radiochromatogram of 99mTc-fucoidan according to the protocol of Rouzet et al. (2011). After 1 hour of incubation, the radiolabeling mixture is eluted with methyl ethyl ketone on silica gel. The percentages indicate the proportion of 99mTc-fucoidan retained at the deposit point (left peak) compared to the total activity added in the solution, the small right signal corresponding to remaining free starting technetium salt.

Moreover, the radiochemical purity was greater than 99.5% (Figure 12.5). These observations demonstrated the capacity of LMWF to strongly bind technetium ions without the need to graft the carbohydrate backbone with a metal ion chelating structure such as tetraazacyclododecane tetra(acetic acid) (DOTA) or 1,4,7-triazacyclononane-N,N′,N″tri(acetic acid) (NOTA). Otherwise, the capacity of fucoidan to aim contrastophores at P-selectin in vitro and in vivo was confirmed with other modalities; in 2014, USPIO coated with this fucoidan allowed MR imaging of aneurysmal thrombi in rats (Suzuki et al. 2015), and a strong interaction of these nanoparticles (NPs) with human activated platelets was also demonstrated in human whole blood (Bachelet-Violette et al. 2014). Bonnard et al. prepared microparticles (MP) of 4 µm diameter by co-crosslinking dextran, pullulan, and fucoidan (Bonnard et al. 2014). Following the protocol of Rouzet et al., 99mTc-MP were prepared and injected in a rat model of aneurysm allowing SPECT imaging of aneurysmal thrombus (Rouzet et al. 2011). Li et al. worked on MP obtained from polymerization of isobutyl cyanoacrylate in the presence of perfluorooctyl bromide as a contrast agent for ultrasonography, coated with polysaccharides and functionalized with fucoidan. Flow cytometry tests showed that fucoidan-coated MPs have a strong affinity for human activated platelets (Li et al. 2017). Anyway, after the initial results of Rouzet’s group, important questions have emerged: 1. What structures of LMWF are responsible for the technetium complexation? 2. Since LMWF extracted from An is a mixture of macromolecular species, are the structures involved in the recognition of P-selectin and the technetium complexing groups carried by the same polymer chains? 3. Is it possible to obtain LMWF in a reproducible way, making it possible to envisage the clinical development of a scintigraphic kit for the imaging of thrombosis and ischemia?

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Enzymatic Technologies for Marine Polysaccharides

Beyond the complete determination of the structures responsible for the interaction of fucoidan with P-selectin, the development of a biospecific probe for clinical molecular imaging of vascular pathologies is required to assess the overall fate of fucoidan within tissues and organs. Because of their natural origin and nonstandardized methods of purification, fucoidans form a complex family of polysaccharides varying in molecular weight, structure, and composition. In order to minimize this natural heterogeneity, all studies from our laboratory have been performed with the same LMWF (Ascophyscient™) provided by Algues & Mer company (Bachelet et al. 2009; Rouzet et al. 2011; Suzuki et al. 2015; Bachelet-Violette et al. 2014). Ascophyscient™ is a commercially available LMWF produced from An according to a confidential industrial process. In addition, nitrogen analyzes highlighted the absence of residual proteins (data not shown). Saboural et al. treated Ascophyscient™ with calcium ions to obtain a purified extract. The molecular weight, structure, and composition of the purified LMWF were determined and compared to those of the starting compound. After radiolabeling with 99mTc according to a previously described method (Rouzet et al. 2011), the biodistribution of both fucoidans was assessed in healthy rats. SPECT imaging was then performed in an ischemia—reperfusion model to evaluate the efficiency of the purified LMWF for in vivo detection of ischemic events in rat myocardium. Uronic acids are frequently recovered in fucoidans from algae either as coextracted polysaccharidic structures, or as discrete short branchings (Ale and Meyer 2013). Eventually, repeated simple fucoidan structures can be obtained but always after several purification steps from crude fucoidan extracts (Nishino et al. 1994; Chizhov et al. 1999; Chevolot et al. 2001; Bilan et al. 2002). To date, only sulfated fucose-rich species have been reported to be responsible for the biological properties of fucoidans (Pomin 2012; Berteau and Mulloy 2003). However, sulfate and fucose represent less than 50% of the dry weight of Ascophyscient™ with a remaining high proportion of other sugars (see Table 12.1 in Section 12.5.2). Thus the extract would be a mixture of bioactive sulfated fucose–based polymers and some polyuronic acids, likely ascophyllan, a sulfated polysaccharide with a uronic acid backbone and sulfated polyfucose branchings (Mabeau and Kloareg 1987; Leite et al. 1998), that Table 12.1  Carbohydrate composition of Ascophyscient™ batches (molar ratios) determined by HPLC assay with comparison to fucose Batch

Fucosea,b

Sulfatea,c

Xylosea,b

Galactosea,b

Mannosea,b

Gluc. Ac.a,b

Glucosea,b

1 2 3 4 5 6 7 8 9 10

1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00

0.87 0.90 0.73 0.65 0.84 0.70 0.89 0.70 0.73 0.66

0.34 0.38 0.43 0.48 0.40 0.33 0.31 0.33 0.37 0.33

0.06 0.08 0.06 0.07 0.07 0.05 0.07 0.05 0.07 0.05

0.03 0.05 0.06 0.05 0.07 0.04 0.04 0.04 0.04 0.03

0.03 0.02 0.02 0.02 0.02 0.04 0.07 0.05 0.06 0.02

0.06 0.04 0.03 0.07 0.03 0.01 0.07 0.02 0.07 0.05

Average

1.00

0.77

0.37

0.06

0.04

0.04

0.04

a b c

mol/mol. HPLC (Zhang et al. 2009). Colorimetric assay (Bachelet-Violette et al. 2014).

Chapter twelve:  Fucoidans in medicine 1

a dRI LS

c

b

e dRI

Relative Scale

Relative Scale

1

283

f

LS

c

d 0 10

12.5

15

17.5 Volume (mL)

20

22.5

25

0 10

12.5

15

17.5

20

22.5

25

Volume (mL)

Figure 12.6 Refractive index (DRI) and light scattering (LS) chromatograms of crude LMWF (Ascophyscient™) (left) and purified LMWF (right) eluted in 0.10 M LiNO3 at 0.5 mL/minute with Shodex SB-802.5 and SB-803 columns. Peak a—high-molecular-weight population of Ascophyscient™ (MW = 12.3 kDa); peak b—low-molecular-weight population of Ascophyscient™ (MW = 2.8 kDa); peak c—total volume of the column; peak d—aggregates observed with LS (no dRI signal); peak e—high-molecular-weight population of purified LMWF (MW = 10.0 kDa); peak f—low-molecularweight population of purified LMWF (MW = 3.1 kDa).

could be coextracted, since alginates are removed by the industrial extraction process (data not shown). Are ascophyllan structures entangled with fucoidan ones responsible for the complexation of technetium? The purification with calcium ions was intended to remove all these remaining uronic acid–containing species. The uronic acid backbone of ascophyllans would form complexes with calcium ions and then precipitate. Our findings showed that FTIR and 1H NMR spectra of crude and purified LMWF presented typical features of a sulfated polysaccharide with sulfation at some O2, O3, and O4 positions of l-fucose as previously demonstrated with fucoidan extracts from An (Saboural et al. 2014). After calcium ion treatment, the purified fraction was enriched in fucose. Moreover, the molecular weight distribution of purified LMWF was shifted to slightly higher molecular weights as small molecules were also removed during the end dialysis (Figure 12.6). Radiolabeling with 99mTc was achieved for both species with a radiochemical purity higher than 95%, similar to that of the 99mTc-heparin (Kulkarni et al. 1980). Biodistribution of the two radiolabeled LMWFs was performed in healthy rats, by quantifying the activities from SPECT images within volumes of interest for two groups of organs: (1) the hepatosplenic system—liver and spleen, and (2) the urinary system—kidneys and bladder (Figure 12.7). Moreover, the presence of 99mTc-fucoidan in the other organs was evaluated by subtracting the two previous measurements from activity of the whole body. Saboural noticed that starting radiolabeled LMWF was massively caught by the liver, while for purified LMWF was mainly eliminated from the body by the kidneys. Saboural et al. hypothesized that the higher accumulation of 99mTc-crude LMWF in the liver and spleen compared to that of 99mTc-purified LMWF could be explained by the higher quantities of uronic acids present in the first one (Saboural et al. 2014). However, knowing that the actual amount of uronic acid is much lower than measured by colorimetry, no simple explanation could be proposed to highlight the differences. Anyway, biodistribution of the purified LMWF is much more satisfactory than that of Ascophyscient™.

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Enzymatic Technologies for Marine Polysaccharides Head

Liver Spleen Kidneys

Figure 12.7 Representative whole rat body SPECT images of the 99mTc-radiolabeled crude LMWF (left) and purified LMWF (right). The rats (5 for crude LMWF and 4 for purified LMWF) were imaged 2 hours after injection of 2 μg of the samples [from Saboural et al. (2014) with permission].

Bladder

Tail

12.5 A fucoidan fraction for the clinical development of a scintigraphic contrast agent of vascular pathologies 12.5.1  Polysaccharides cannot yet be fully synthesized Thus, fucoidans are abundant and cheap marine polysaccharides with a wide spectrum of properties that are potentially interesting in various medical fields, especially in cardiovascular medicine. Nevertheless, one wonders why they are not more widely developed. Despite intensive researches during the last 100 years, the chemical structures of all fucoidans are still not fully determined. The great variability of structures, depending on algal species, areas and seasons of harvesting, and extraction and purification methods incontestably slow down their adoption. Thus it is also mandatory to obtain robust structure–activity relationships. As a consequence, a standard method to obtain a reproducible well characterized fucoidan still remains to be developed. The fucoidans available from well-known suppliers (such as the Sigma-Aldrich Company) are heterogeneous mixtures in terms of composition and molecular weight distribution, with limited availabilities of the batches and a strong interbatch variability. A promising way would be the full synthesis of fucoidan. However, polysaccharides, in contrast to polynucleotides and polypeptides, cannot yet be readily synthesized. Nifantiev’s group succeeded in obtaining synthetic oligofucoidans, up to 16 residues, by reproducing the various structures of the compounds extracted from brown algae (Gerbst et al. 2003; Krylov et al. 2011). This way, using only the tools for organic synthesis, is still extremely complex, expensive, and not at all transposable for an industrial production. At the moment, the only relevant strategy is to select a bioactive fucoidan in the field of interest and to “secure” it from an industrial production perspective, that is, from the cultivation of algae to the production of a defined fraction, through the establishment of an efficient process of extraction and purification.

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12.5.2  Fucoidan extraction from brown algae must be secured Following the type of algae, the location and season of harvesting, and the process of extraction and purification (such as pH, time, temperature, and pressure), the physicochemical characteristics of the final product are highly variable (Ale et al. 2011a; Hahn et al. 2012; Lorbeer et al. 2015; Garcia-Vaquero et al. 2017). So far, a vast number of methods of extraction and purification of fucoidans have been described in publications following three main steps: 1. After harvesting, algae are washed with water to remove sand and other solid impurities. This cleaning is frequently followed by a step of drying (oven drying or freeze drying). The dried biomass is then milled to obtain the highest surface-to-volume ratio to optimize extraction during the next procedures (Garcia-Vaquero et al. 2017; Imbs et al. 2009; Hahn et al. 2012). 2. A second washing often follows this first step with organic solvents such as mixtures of methanol, chloroform, and water (4: 2: 1; v/v/v) or acetone alone (Ale et al. 2011b) to remove lipids, proteins, and more specifically enzymes, dyes, and polyphenols. The polysaccharides are solubilized in water and precipitated with organic solvents miscible with water or with surfactants, and extracted by centrifugation or extensive dialysis (Garcia-Vaquero et al. 2017; Hahn et al. 2012; Duarte et al. 2001). Additional purification steps are sometimes necessary to enrich the mixtures such as ion-exchange chromatography, size-exclusion chromatography, or membrane filtration. 3. The extraction of fucoidan is generally carried out mainly using water or ethanol, slightly acidified, at temperatures ranging from room temperature to 120°C, for several hours (Cong et al. 2016; Shan et al. 2015; Wang et al. 2012; Foley et al. 2011; Imbs et al. 2009; Garcia-Vaquero et al. 2017; Lorbeer et al. 2015). Some other techniques such as ultrasound-assisted, microwave-assisted, or enzyme-assisted extraction have also been used (Michalak and Chojnacka 2014; Garcia-Vaquero et al. 2017). Alginates, which are frequently coextracted with fucans, can be removed by precipitation with calcium chloride or sometimes by acidification of the extracts. In the crude extracts (after step 3) molecular weights are often higher than 106 g/mol with a very large size distribution and a heterogeneous chemical composition. Depolymerization methods are necessary to obtain fractions with at least tighter molecular weight distribution, allowing easier structure–activity relationship studies (Nardella et al. 1996). Thus, acid or radical hydrolysis (Ale et al. 2011b; Nardella et al. 1996), enzymatic degradation with fucoidanases extracted from marine microorganisms or marine invertebrates (Kusaykin et al. 2008), and γ-irradiation (Kim et al. 2009; Choi et al. 2014; Choi and Kim 2013) are the most widespread techniques. However, some discrepancies regarding the reproducibility of the final structures and composition preclude the determination of relevant structure-activity relationships (Ale et al. 2011b) and, as a consequence, clinical developments. Algues & Mer developed and optimized a process of extraction starting from An to the production of Ascophyscient™. The reliability and reproducibility of the process were validated by the analysis of 10 batches of Ascophyscient™ harvested at different times between 2012 and 2017. Monosaccharide composition and sulfate content are essential characteristics that are provided infrequently in scientific articles dealing

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Enzymatic Technologies for Marine Polysaccharides 8.99

100 % 0

Ribose

525.1985

118.1228

300

100 % 118.1233 235.2389 0 200 300 100 % 118.1237 235.2393 0 200 300

8.67

526.2017

481.2075

235.2382

200

Xylose

Fucose

100

400

500

661.1219

600

700

893.4725 937.4975

800

900

m/z 1000

481.2096 482.2122

579.1776

481.1434

400

500

600

735.2449

700

863.2414

800

900

999.3655

m/z 1000

495.2257 496.2285 495.1629

400

500

593.1935

600

749.2621 853.3099 891.2744

700

800

900

m/z 1000

Mannose

%

?

6.70 6.87 7.25

Ac. Gluc. Glucose Galactose

7.35

OH

8.50

8.08

6.38

0

8.30

HO 9.54 9.64

6.00 6.50 7.00 7.50 8.00 8.50 9.00 9.50

N

O OH

CH3

HO

OH H3C NH

Time (min)

NH

O N

Glu-MPP m/z = 511.2193

Figure 12.8  RP-HPLC analysis of an Ascophyscient™ hydrolyzate with online mass spectrometry. After the acid hydrolysis of the polysaccharide in 2 M trifluoroacetic acid at 110°C for 4 hours, the sugars were derived with 3-methyl-1-phenyl-2-pyrazolin-5-one (MPP) (insert) (maximum absorbance at 245 nm) and eluted with mixtures of acetonitrile, ammonium acetate and triethylamine at pH 6.3 (Zhang et al. 2009). Examples of mass spectra are provided for glucuronic acid, xylose, and fucose. The amount of each monosaccharide was then determined using a reference solution and an internal reference composed of ribose.

with fucoidans. In particular, sulfate levels are a key factor in their biological activities (Koyanagi et al. 2003; Soeda et al. 2000; Oliveira et al. 2017; Kim et al. 2015). A precolumn HPLC derivation method described by Zhang et al. (2009) was adapted to establish the carbohydrate composition of Ascophyscient™. Figure 12.8 shows the chromatogram of an Ascophyscient™ hydrolyzate. Comparison with standard chromatograms and peak analysis by mass spectrometry allowed the identification of the different species. The 10 batches of Ascophyscient™ were analyzed by this method. The results are presented in Table 12.1. Fucose residues represent the main carbohydrate units followed by xylose and galactose residues. The molar ratios indicate the presence of sulfates on approximately 80% of fucoses. Although ranging from 0.65 to 0.90, the sulfate: fucose ratio is considered reproducible for two reasons: 1. Algae cultivation is controlled and optimized within the limits of a protected natural environment chosen to guarantee the quality of the extracts; note that the reproducibility of the Ascophyscient™ monosaccharide composition demonstrates that the harvest period has virtually no effect on its final composition. 2. The activity of fucoidans is more directly related to the presence and number of particular bioactive structures than to the average number of sulfate groups per fucose

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Table 12.2  Composition of Ascophyscient™ and FUCO* after purification Fucoidans Ascophyscienta FUCOb

Mw (kDa)

% Fucosec,d

% Sulfatec,e

% Other neutral osesc,d

% Ashesc,f

6.2 ± 0.3 7.5 ± 0.2

29.2% ± 0.6 43.0% ± 0.2

14.7% ± 1.6 26.5% ± 0.2

31.0% ± 3.0 20.1% ± 3.0

11.5 7.5

* FUCO: low molecular weight fucoidan obtained from an optimized process a Average values on 10 batches of Ascophyscient™. b Average values on the batch of Ascophyscient™ developed for clinical studies in Nanoathero framework (Chauvierre and Letourneur 2015). c w/w. d HPLC (Zhang et al. 2009). e Colorimetric assay (Bachelet-Violette et al. 2014). f Values obtained from thermogravimetric analysis (data not shown).

unit. As an example, Chevolot et al. (1999) evidenced that anticoagulant activity of fucoidan from An is closely related to the position of sulfate groups on the polysaccharide (Chevolot et al. 1999). For the clinical development of therapeutic or diagnostic agents, all components administered to humans must be of pharmaceutical grade; that is, they must have been produced following Good Manufacturing Practices (GMP) as described in the European pharmacopoeia. In this context, after almost 10 years of a tight collaboration, a joint laboratory was created in 2013 between LVTS and Algues & Mer Company to secure the production of reproducible LMWF with well-defined composition and molecular weight, from the process developed by the company to produce Ascophyscient™. The treatment of Ascophyscient™ with calcium acetate previously described by Saboural et al. (2014) was optimized, allowing to get a new LMWF, (termed FUCO), from the starting product (Table 12.2). Not only did FUCO contain higher amounts of fucose residues and sulfate groups, but uronic acid–containing species had been completely eliminated (data not s shown). In 2015, FUCO was labeled by the French authorities as “raw materials for pharmaceutical uses.” Today, it is part of the development of a 99mTc-SPECT imaging of human atherothrombosis, and clinical trials started in 2018 in the framework of the large-scale European project Nanoathero (Chauvierre and Letourneur 2015).

12.6  How fucoidan helps to fight stroke The treatment of acute thrombosis depends on its severity and on localization. Two approaches are considered: surgery and/or drugs. If occlusion is not total, the treatment prioritizes administration of anticoagulants and platelet inhibitors, which prevent the growth of the thrombus and facilitate its disappearance (Bonaventura et al. 2016; Piechowski-Jozwiak and Bogousslavsky 2013). In case of total occlusion of an artery, the intravenous injection of recombinant tissue plasminogen activator (rtPA) remains the most common noninvasive treatment to recanalize vessels (Marler et al. 1995; Bonaventura et al. 2016) (Figure 12.9). For the past 20 years, thrombolysis by rtPA has been the only clinically approved drug treatment for ischaemic stroke (Hacke et al. 2008). However, its use is limited to a very short therapeutic window of 4–5 hours after the first clinical signs, due to the high risk of haemorrhagic transformation (Marler et al. 1995; Sussman and Connolly 2013). In addition, since rtPA is administered intravenously, it is also

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Mechanism of action of rtPA

Plasminogen activator inhibitor-1 (PAI-1)

Ternary complex rt-PA/plasminogen/fibrin

Site of interaction with fibrin (Lys)

Platelet

rtPA Thrombus

Fibrin

Site of interaction with fibrin (Lys) Plasminogen Thrombus disintegration

Fibrin degradation

Figure 12.9  Scheme of action of tissue plasminogen activator (tPA). tPA is the key serine protease that catalyzes the conversion of plasminogen to plasmin. Then plasmin degrades the fibrin network of the thrombus, thereby promoting thrombolysis. Plasminogen and tPA (or rtPA) bind to lysine residues in the fibrin network, forming tPA–fibrin–plasminogen ternary complexes (Longstaff et al. 2011). tPA binds to the cationic amino groups of lysine sidechains by a specific domain (Silva et al. 2012), while plasminogen binds mainly to the carboxy-terminal lysines. These bindings allow tPA–plasminogen interaction and protect tPA from its circulating inhibitors, in particular PAI-1 (Kaneko et al. 1992), therefore initiating the local conversion of plasminogen into plasmin.

quickly neutralized and inhibited by circulating plasminogen activator inhibitor (PAI-1) (Cesarman-Maus and Hajjar 2005; Medcalf 2007; Mutch et al. 2007). Besides, rtPA has some neurological toxicity related to its local concentration (Chapman et al. 2000). Eventually fewer than 10% of patients end up receiving treatment (Jaffer et al. 2011). The association of rtPA treatment with the use of endovascular devices constituted an improvement for the management of stroke (Fiehler et al. 2016), although it was limited to particular situations such as the accessibility of the thrombi and needing specific clinical facilities (Wahlgren et al. 2016). In addition to the development of other fibrinolytic agents (Bonaventura et al. 2016; Piechowski-Jozwiak and Bogousslavsky 2013), nanomedical approaches for local or targeted delivery of thrombolytic drugs arouse a growing interest (Cicha 2015; Silva et al. 2014; Varna et al. 2015). This strategy, combined with active targeting approaches, allowed the accumulation of nanocarriers onto the thrombosed area, improving thrombolytic efficiency while reducing hemorrhagic complications, in preclinical models of thrombosis (Ma et al. 2009; Bi et al. 2009; Yang et al. 2012; Chen et al. 2016; Hu et al. 2016; Marsh et al. 2011; McCarthy et al. 2012; Kim et al. 2015; Fredman et al. 2015; Absar et al. 2013; Vaidya et al. 2011; Zhou et al. 2014). P-Selectin, which has been poorly explored with respect to targeted thrombolysis, was chosen as the molecular target. Since fucoidan strongly binds to P-selectin, we hypothesized that an association of rtPA to fucoidan would allow a more specific delivery to the thrombus and, as a consequence, would increase fibrinolytic efficacy.

Chapter twelve:  Fucoidans in medicine (i) H2N

(A) Fucoidan

C

F-NH2

H O

+

289

F-NH2

NH2

CH2 NH

Fucoidan

(ii) Reduction H2N

H2N

(iii) EDC/NHS

HO

F–NH

NH2

O

Fucoidan-dilysine/rt-PA complex

NH2

O

2

NH2 2

= rt-PA

(B) CH2 OH OH

CH2

O O OH CH2 OH OH

Ce4+ pH1

OH OH

O O OH CH2

OH 50% dextran OH 40% dextran-NH2 10% fucoidan

N O

O

O O OH CH2 OH OH

O

O O OH CH2 OH OH

O OH CH2

O

O CN CH2 C CH2 COOR

OH

Vectorization of rt-PA onto activated platelets

CN C COOR

n

Polymer nanoparticle

136 ± 4 nm

More rapid recanalization

#A #B Activated endothelial cells

Activated platelets

Thrombus Fibrin

P-selectin

Figure 12.10  Strategies of vectorization of rtPA with fucoidan [adapted from Juenet et al. (2018) and from Ghebouli et al. (2018) with permission]. (A) Representative intravital images of mouse mesenteric vessel reperfusion after a partial occlusion induced by ferric chloride 10%. The accumulation of Rhodamine 6G stained platelets (in red), and Fibrinogen Alexa (purple) was recorded. Monitoring of thrombolysis postadministration of rtPA (1 mg/kg) +/− DLF (0.5 mg/kg). Reperfusion of vessels was also observed for animals following rtPA/DLF treatment as compared with rtPA alone. Bar ¼ 100 μm. (B) Quantification or the rate of recanalized vessels within 1 hour following thrombosis induction. Groups were compared using the Pearson chi-squared test; *** p < 0.001 (n ¼ 32 mice in each group).

Two strategies of vectorization have been tested (Figure 12.10): 1. The first strategy explored the possibility to use fucoidan as a chaperon molecule able to improve the rtPA bioavailability to thrombus. For this purpose, oligolysines were linked to a LMWF from A&M (MW = 6.2 kDa) by their carboxy-terminal function to expose their NH 2 residues. In vitro, rtPA mixed to fucoidan-dilysine showed a greater fibrinolytic effect than rtPA alone, both on platelet-rich thrombus and in whole blood. In vivo, occluded mesenteric vessels, carotid artery and vena cava were more efficiently recanalized by fucoidan-dilysine complexed to rtPA than by rtPA alone (Ghebouli et al. 2018) (Figure 12.11A,B).

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Enzymatic Technologies for Marine Polysaccharides

(A)

+15 min

+rtPA

+20 min

(B)

+60 min

60 Number of mice

40

3/32

27/32

rtPA

rtPA/DLF

10% FeCI3 2min30

20 +rtPA/DLF

0

Occluded 10% FeCI3 2min30

(D)

60 40 20

ns

0

Fu c

rtPAFuco-NPs

o-

Bu

ffe r

rtPA-Control-NPs

80

0

5

10 15 20 Time after injection (min)

25

30

PA rt-C PA on tro rtl-N PA Ps -F uc oN Ps

rtPA

100

Ps

Thombus density (%)

Fuco-NPs

N

Vein walls Blood flow

Buffer

rt-

(C)

Recanalized

Figure 12.11  (A) Representative intravital images of mouse mesenteric vessel reperfusion after a partial occlusion induced by FeCl3 10% (n = 32). The accumulation of Rhodamine 6G stained platelets (red), and Alexa stained fibrinogen (purple) was recorded. The thrombolysis postadministration of fluorescent rtPA (green) (1 mg/kg) ± fucoidan-dilysine (FDL) (0.5 mg/kg) was monitored. Reperfusion of vessels was also observed for animals following rtPA/FDL treatment as compared with rtPA alone. (B) Quantification of the rate of recanalized vessels. 27 mice on 32 had a fully recanalized vessel 60 minutes after rtPA/DLF injection [adapted from Ghebouli et al. with permission (Ghebouli et al. 2018)]. (C) Evaluation of thrombolysis efficiency in a mouse model of venous thrombosis (n = 41) by fluorescence imaging of Rhodamine 6G labeled platelets at site of thrombosis over time. Representative examples of thrombus evolution as determined by platelet accumulation at different times after injection of buffer, fucoidan-containing NPs (Fuco-NPs), rtPA, rtPA-Control-NPs, and rtPA-Fuco-NPs, respectively. Prior to injection, circulating platelets were fluorescently labeled with Rhodamine 6G and thrombosis was generated on a mesenteric vein by applying a FeCl3-soaked paper for 1 minute. (D) Thrombus density at 30 minutes after injection. Five groups were analyzed. [Adapted from Juenet et al. with permission (Juenet et al. 2018).]

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2. In a second strategy solid spherical fluorescent polysaccharide-poly(isobutyl­ cyanoacrylate) NPs with a hydrodynamic diameter of 136 ± 4 nm were produced by redox radical emulsion polymerization (Chauvierre et al. 2003). rtPA was loaded by adsorption on the polysaccharide shell to promote its direct availability at the thrombus site and the full retention of its activity. To mimic the free primary amines present in the natural fibrin binding sites dextran modified with amino groups was incorporated into the NP formulation (Krylov et al. 2011). For this strategy a medium-molecular-weight fucoidan also provided by A&M was used (MW = 104 kDa). After demonstrating that this fucoidan can effectively improve in vitro the interaction of fucoidan-functionalized NPs with P-selectin under flow, the thrombolysis efficiency was confirmed in a mouse model of venous thrombosis by monitoring the platelet density with intravital microscopy (Figure 12.11C,D) (Juenet et al. 2018). The chemical feasibility and in vitro and in vivo proof-of-concept efficacy of lysine-bearing fucoidan as a thrombus-targeted vector for rtPA as well as that of local delivery of rtPA loaded by adsorption on aminated NPs functionalized with fucoidan were explored. An interesting point to consider is that the fucoidan molecules used for these studies were produced by A&M from Ascophyllum nodosum from similar processes.

Conclusion The biology and biochemistry of the 20th century revolutionized the life sciences and medicine through an integrated approach to phenomena. In parallel, the need to miniaturize electronic systems to improve their performance gave birth to nanotechnologies allowing tremendous advances in imaging. Smaller and smaller objects could be examined, gradually giving access to the observation of interactions at cellular and molecular scales. Finally, chemistry has allowed the emergence of synthetic materials, in particular polymer materials, which we have constantly tried to associate with the living world. In this context, proteins, nucleic acids, and lipids have been rapidly synthesized and associated with pharmaceutical and biomaterials research. Except polysaccharides, the last family of natural macromolecules, are still now extracted more or less laboriously from terrestrial and marine organisms. Anyway, no one can dispute today their essential role in biological systems. The history of fucoidan is edifying on this point since its remarkable properties in the healthcare field are being increasingly (Zaporozhets and Besednova 2016). Over the next 15–20 years, there is no doubt that organic chemists are finally able to overcome the synthesis difficulties inherent in the particular structures of polysaccharides. In the meantime, it is still possible to exploit the clinical potential of molecules such as fucoidans extracted from algae, as long as the necessary efforts are made to secure their production. After about 15 years of a tight partnership with our laboratory, the GMP production of a LMWF by Algues & Mer Company (a subsidiary of the Solabia Group since 2017) allowed us to develop a kit for human scintigraphy of atherothrombosis. We are confident in the success of the clinical trials that started in 2018 in the framework of the European project Nanoathero since (1) the very tiny amount of fucoidan necessary for 1 kit and (2) no potential adverse effect has ever been identified in the use of fucoidan in the health field. The next step will be the development of the extracts to build novel therapeutics/ theranostics dedicated to the diagnosis, the treatment and the follow-up of cardiovascular diseases.

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Acknowledgments This work was supported by Inserm and University Paris 13 and the competitiveness cluster Medicen Paris Region. Lucas Chollet is a recipient of a CIFRE grant from ANRT (ANR-13-RPIB-0006 “FucoThrombo”). The authors acknowledge the financial support from FP7 NMP-LA-2012-309820 “Nanoathero,” ANR-13-LAB1-0005-01 “FucoChem,” and ANR-13-RPIB-0006 “FucoThrombo.”

References Absar, S., K. Nahar, Y. M. Kwon, et al. 2013. Thrombus-targeted nanocarrier attenuates bleeding complications associated with conventional thrombolytic therapy. Pharm. Res. 30(6):1663–1676. Ale, M. T., H. Maruyama, H. Tamauchi, et al. 2011a. Fucose-containing sulfated polysaccharides from brown seaweeds inhibit proliferation of melanoma cells and induce apoptosis by activation of caspase-3 in vitro. Mar. Drugs 9(12):2605–2621. Ale, M. T., and A. S. Meyer. 2013. Fucoidans from brown seaweeds: an update on structures, extraction techniques and use of enzymes as tools for structural elucidation. RSC Adv. 3(22):8131–8141. Ale, M. T., J. D. Mikkelsen, and A. S. Meyer. 2011b. Important determinants for fucoidan bioactivity: A critical review of structure-function relations and extraction methods for fucose-containing sulfated polysaccharides from brown seaweeds. Mar. Drugs 9(10):2106–2130. Alves, A. P., B. Mulloy, J. A. Diniz, et al. 1997. Sulfated polysaccharides from the egg jelly layer are species-specific inducers of acrosomal reaction in sperms of sea urchins. J. Biol. Chem. 272:6965–6971. Aquino, R. S., A. M. Landeira-Fernandez, A. P. Valente, et al. 2005. Occurrence of sulfated galactans in marine angiosperms: evolutionary implications. Glycobiology 15(1):11–20. Bachelet-Violette, L., A. K. A. Silva, M. Maire, et al. 2014. Strong and specific interaction of ultra small superparamagnetic iron oxide nanoparticles and human activated platelets mediated by fucoidan coating. RSC Adv. 4(10):4864. Bachelet, L., I. Bertholon, D. Lavigne, et al. 2009. Affinity of low molecular weight fucoidan for P-selectin triggers its binding to activated human platelets. Biochim. Biophys. Acta 1790 (2):141–146. Bentzon, J. F., F. Otsuka, R. Virmani, et al. 2014. Mechanisms of plaque formation and rupture. Circ. Res. 114(12):1852–1866. Berteau, O., and B. Mulloy. 2003. Sulfated fucans, fresh perspectives: structures, functions, and biological properties of sulfated fucans and an overview of enzymes active toward this class of polysaccharide. Glycobiology 13(6):29R–40R. Bettinger, T., P. Bussat, I. Tardy, et al. 2012. Ultrasound molecular imaging contrast agent binding to both E- and P-selectin in different species. Invest. Radiol. 47(9):516–523. Bi, F., J. Zhang, Y. J. Su, et al. 2009. Chemical conjugation of urokinase to magnetic nanoparticles for targeted thrombolysis. Biomaterials 30(28):5125–5130. Bilan, M. I., A. A. Grachev, N. E. Ustuzhanina, et al. 2002. Structure of a fucoidan from the brown seaweed Fucus evanescens C.Ag. Carbohydr. Res. 337:719–730. Bilan, M. I., A. S. Shashkov, and A. I. Usov. 2014. Structure of a sulfated xylofucan from the brown alga Punctaria plantaginea. Carbohydr. Res. 393:1–8. Blondin, C., E. Fischer, C. Boisson-Vidal, et al. 1994. Inhibition of complement activation by natural sulfated polysaccharides (fucans) from brown seaweed. Mol. Immunol. 31:247–253. Bonaventura, A., F. Montecucco, and F. Dallegri. 2016. Update on the effects of treatment with recombinant tissue-type plasminogen activator (rt-PA) in acute ischemic stroke. Expert Opin Biol. Theor. 16(11):1323–1340. Bonnard, T., G. Yang, A. Petiet, et al. 2014. Abdominal aortic aneurysms targeted by functionalized polysaccharide microparticles: A new tool for SPECT imaging. Theranostics 4:592–603.

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Wang, W., S.-X. Wang, and H.-S. Guan. 2012. The antiviral activities and mechanisms of marine polysaccharides: An overview. Mar. Drugs 10:2795–2816. Wildgruber, M., F. K. Swirski, and A. Zernecke. 2013. Molecular imaging of inflammation in atherosclerosis. Theranostics 3(11):865–884. Willerson, J. T., and P. M. Ridker. 2004. Inflammation as a cardiovascular risk factor. Circulation 109(21):2–10. Yang, H. W., M. Y. Hua, K. J. Lin, et al. 2012. Bioconjugation of recombinant tissue plasminogen activator to magnetic nanocarriers for targeted thrombolysis. Int. J. Nanomed. 7:5159–5173. Zaporozhets, T., and N. Besednova. 2016. Prospects for the therapeutic application of sulfated polysaccharides of brown algae in diseases of the cardiovascular system: Review. Pharm. Biol. 5 (12):3126–3135. Zhang, J., Q. Zhang, J. Wang, et al. 2009. Analysis of the monosaccharide composition of fucoidan by precolumn derivation HPLC. Chin. J. Oceanol. Limn. 27(3):578–582. Zhou, J., D. J. Guo, Y. Zhang, et al. 2014. Construction and evaluation of Fe3O4-based PLGA nanoparticles carrying rtPA used in the detection of thrombosis and in targeted thrombolysis. Acs Appl. Mater. Inter. 6(10):7961–7961.

chapter thirteen

Marine polysaccharides as promising source of biological activities Extraction and purification technologies, structure, and activities A. Mzibra1,2 , I. Meftah Kadmiri2 , and H. El Arroussi2

Hassan II Institute of Agronomy and Veterinary Medicine (IAV), Rabat, Morocco Moroccan Foundation for Advanced Science, Innovation and Research (MASCIR), Rabat, Morocco 1 2

Contents 13.1 Introduction....................................................................................................................... 301 13.2 Extraction techniques, structural determination, and methodologies to assess biological activities........................................................................................................... 302 13.2.1 Extraction technologies of marine polysaccharides....................................... 302 13.2.2 Purification of marine polysaccharides...........................................................306 13.3 Marine polysaccharides: structures, function, and biological activities studies..... 309 13.3.1 Biological activities of marine polysaccharides..............................................309 13.3.2 Linking chemical composition and structure to biological activities of marine polysaccharide����������������������������������������������������������������� 311 Conclusion.................................................................................................................................... 314 References...................................................................................................................................... 315

13.1 Introduction Marine polysaccharides can be extracted from different sources such as seaweeds, microalgae, microorganisms, and marine vertebrates and invertebrates (mollusks, sponges, arthropods– crustaceans) (Senni et al. 2011; Wang et al. 2012a; Trincone 2014). Among this vast marine organism diversity, polysaccharides from the marine algae represent the most abundant polysaccharides in the sea (Wang et al. 2012a). Generally, polysaccharides are present with various other components such as proteins, polynucleotides, lipids, lignin, and some inorganic minerals. The biological activities of marine polysaccharides can be undermined by other compounds, which can even cause antagonistic effects or undesirable toxicity (Colegate and Molyneux 2008). The pure bioactive polysaccharides must allow a safe, reproducible, strict dosage and accurate concentrations for experimental or therapeutic applications, and allow the study of the structure–activity relationship, facilitating the development of new compounds with similar or higher bioactivities (Colegate and Molyneux 2008). Thus, the extraction of polysaccharides from complex matrix networks, while minimizing any loss of desired bioactivity, is one of 301

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the most challenging tasks to deal with (Wijesekara et al. 2011). There are now a variety of well-established methods for the extraction of marine polysaccharides such as supercritical fluid extraction (SFE), microwave-assisted extraction (MAE), and ultrasoundassisted extraction (UAE). This chapter focuses on the recent literature and describes the best extraction conditions and purification procedures employed to obtain highvalue marine polysaccharides. It summarizes the common practices desired for the 21st century to convince the world of research and industry to take a step forward and replace the conventional extractions by appropriate, selective, cost-effective, ecofriendly extraction technologies. Moreover, a bibliographical synthesis of the structure–function relationships of these polysaccharides and their importance in new applications, as well as an update on the methods and modes utilized to investigate the effect of these macro­ molecules are given.

13.2 Extraction techniques, structural determination, and methodologies to assess biological activities 13.2.1  Extraction technologies of marine polysaccharides To extract polysaccharides from marine sources, specific methods of preparing the material are selected depending on the raw material and future use of the polysaccharide. They include pretreatment methods, extraction, separation, and purification.

13.2.1.1  Preparation of the biomass The procedure of extracting marine polysaccharides involves reducing the water content by drying the biomass, two methods can be used: freeze drying (sample is washed with water and freeze-dried at temperature of 253.15 K for several days) or drying at low temperature (close to 35°C) in order to avoid the degradation of thermolabile compounds (Crampon et al. 2011). Grinding of the dried biomass is necessary in order to ensure uniform distributed mass as well as a higher surface-to-volume ratio (Hahn et al. 2012; Dore et al. 2013; Imbs et al. 2016). Before appropriate extraction process, and in order to increase the effectiveness of polysaccharide extraction, two types of pretreatments are applied to the biomass. The first one is vital for preventing the coextraction of other bioactive compounds from marine biomass that have similar solubility properties (Hahn et al. 2012). Therefore, these treatments are applied to remove lipids, proteins and phenols, but also mannitol and chlorophyll compounds that are highly bound to the polysaccharides, contaminating the target compounds (Hahn et al. 2012). For example, defatting of the marine biomass with a mixture of methanol, chloroform, and water at 4: 2: 1 (v/v/v) is found to be advantageous to prevent the coextraction of total lipids during the fucoidan extraction (Hahn et al. 2012). Pretreatment can also be done with acetone alone to eliminate lipids and pigments (Dore et al. 2013) or a mixture of acetone and ethanol (Dinesh et al. 2016). Recently, a number of alternative pretreatment methods at different temperatures have been explored, including pretreatment by ethanol (96%, 40°C, 24 hours) and acetone washes (Imbs et al. 2016). Ethanol pretreatment (80%, room temperature, 18 hour and repeated at 70°C, 4 hours) or ethanol (95%, 80°C, 4 hours, 2 times) (Yuan and Macquarrie 2015; Shan et al. 2016). A novel pretreatment described by Huang et al. (2016) includes the compressional puffing hydrothermal process, which consists of heating at atmospheric pressure (140°C, 180°C, and 220°C). When the chamber reached the target temperature, the separable cylindrical chamber automatically moved up and down 3 times and provided a

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mechanical compression force of approximately 5 kg/cm2 to the sample. Then the separable cylindrical quickly opened to induce puffing by rapid steam pressure release. The total reaction time for each operation was about 10 seconds. The corresponding pressure levels inside the chamber were 1.7, 10.0, and 18.3 kg/cm2, and the temperature reached 140°C, 180°C, and 220°C, respectively. The reduction of pressure allows modification of the cellular structure of the seaweeds prior to polysaccharides extraction. The second type of pretreatment improves extraction yield, because at this stage, cells are disrupted, which makes bioactive compounds more available (Harun et al. 2014). This is true especially in the case of microalgae, whose size is only a few micrometers. Hence, finding an effective and inexpensive method to destroy the cell wall is particularly important (Samarasinghe et al. 2012). Several methods of cell disruption are described in the literature, including mechanical and physical methods using ball milling, high-pressure extrusion, and, to a lesser extent, ultrasonication, which is a repetitive compression and a rarefaction of the waves that cause cavities and/or microbubbles to collapse and create shear forces that disrupt cell walls and membranes (Harun et al. 2014). Microwave pretreatment has also been proven to be effective method, but requires high-energy input. Thermal treatment can be performed in a natural gas drum dryer. The disadvantage is that even if the heat treatment serves to lyse the cells while removing the residual water, there is always a risk of cell rupture. As well as the shear, forces created by a mechanical disturbance can be used to further degrade or denature the desired products (Huang et al. 2014). Alternative chemical and biological cell disruption techniques are attracting attention. Chemical agents such as acid or alkaline (i.e., hydrochloric or sulfuric acid, sodium hydroxide) can be added to the biomass in order to hydrolyze it into constituent molecules (Lee et al. 2010). Enzymatic cell wall degradation is not widely practiced in the industry now, because cell-lysing enzymes are costly (Sander and Murthy 2009).

13.2.1.2  Extraction methods 13.2.1.2.1  Conventional solvent extraction (CSE)  The extraction of marine polysaccharides is carried out mainly by the classical extraction procedure consisting of a solvent extraction, by using pretreated dried or defatted biomass with different solvents at different temperatures ranging from room temperature to 120°C, different incubation time, and different numbers of extraction cycles. The most commonly used solvents are water (Wang et al. 2012b,c; Cong et al. 2016; Shan et al. 2016) and ethanol (Foley et al. 2011; Huang et al. 2016). Solutions with low HCl molarity were also used in several studies (Anastyuk et al. 2012; Dinesh et al. 2016; Imbs et al. 2016; Lorbeer et al. 2015; Menshova et al. 2015). The precipitation of polysaccharide is performed after the initial solvent extraction, by an application of organic solvents such as ethanol or tensids like cetrylmethylammonium bromide (CTAB) that interact specifically with sulfated polysaccharides (Hahn et al. 2012; Kadam et al. 2015a). The precipitated polysaccharides can then be dialyzed or precipitated further with ethanol in one or several steps to remove salts and other compounds from the enriched extracts (Ale et al. 2011a; Cong et al. 2016). Furthermore, conventional techniques are usually manual processes and reproducibility is challenging. The organic solvents required in these processes are harmful to the environment. For example, n-hexane is ranked the highest out of 189 hazardous air pollutants (HAPs) by the US Environmental Protection Agency (EPA) (Shen et al. 2005). Thus, because of these limitations, combined with the significant increase in the demand for marine organisms, novel techniques should be developed (not based on heat and solvent

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use, which can damage bioactive polysaccharides). They should be appropriate, selective, cost-effective, and ecofriendly extraction technologies. Such efficient technologies produce higher yields and comply with relevant legislation (Mamidipally et al. 2004). 13.2.1.2.2  Ultrasound-assisted extraction (UAE)  Ultrasound waves are high-­frequency sound waves above the range for human hearing capacity (>20 kHz). They are physical waves that cross solid, gaseous, and liquid media propagating by rarefaction and compression, and cause a negative pressure in the liquid. If the pressure exceeds the tensile strength of the liquid, vapor bubbles are formed. Implosion of cavitation bubbles generates macroturbulence, high-speed interparticle collisions, and perturbation in microporous particles of biomass (Shirsath et al. 2012). The circulation near the liquid-solid interfaces directs a flow of liquid moving rapidly through the cavity to the surface. The impact of these microjets results in surface peeling, erosion, and particle decomposition, which facilitates the release of bioactive substances from the biological matrices. This effect increases the efficiency of the extraction by increasing the mass transfer by the internal diffusion mechanisms (Vilkhu et al. 2011). Thus, it appears that application of ultrasound allows target compounds to dissolve in the solvent, thereby boosting yield with shorter time through the disruption of the cell wall. Ultrasound-assisted extraction was used to obtain polysaccharides from Ascophyllum nodosum (Wijesekara et al. 2011) and laminarin from Laminaria hyperborea in combination with weak-acid solutions (Kadam et al. 2015b). In some cases, UAE is also employed as a pretreatment method before extraction, which could be performed in an ultrasonic bath or ultrasound probe instrument. Despite being an energy input extraction method, UAE has certain advantages over conventional solvent extraction methods. UAE is a simple, cost-effective, and efficient alternative in comparison with other novel extraction techniques and has high potential to scale up to industry. Ultrasound facilitates the extraction of heat-sensitive compounds with minimal damage. Benefits of using ultrasound in solid–liquid extraction include an increase of extraction yield and faster kinetics, and the equipment costs are lower than with other novel extraction techniques (Chemat et al. 2011). Other advantages of UAE include low solvent consumption, high level of automation, and possibilities of combining this technique with others, such as superfluid-assisted extraction or microwave-assisted extraction (Ibañez et al. 2012; Michalak and Chojnacka 2014). Two main types of ultrasonic equipment can be used for extraction purposes: an ultrasonic water bath and an ultrasonic probe system (Ibañez et al. 2012). 13.2.1.2.3  Microwave-assisted extraction (MAE)  The wall of plant cells consists mainly of cellulose, which hinders the isolation process of polysaccharides from marine biomass. The use of physical methods, such as microwaves and ultrasounds, can accelerate and facilitate the extraction of polysaccharides and even increase performance (Li et al. 2012). Microwaves are nonionizing electromagnetic radiation with a frequency ranging from 300 MHz to 300 GHz. In the MAE, microwaves induce the vibration of water molecules within the cell. Vibrations cause the increase in temperature of the intracellular liquids that activates the evaporation of water and exerts pressure on the cell walls. As the cell wall breaks down, its constituents, along with the intracellular active substances, including polysaccharides, are released into the medium in a facilitated manner (Michalak and Chojnacka 2014). In MAE hydrogen bonds are disrupted, and migration of dissolved ions increase the penetration of solvent into the matrix, which facilitates the extraction of active compounds (Routray et al. 2012).

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The use of microwaves for extraction of various compounds of interest from both plants (Mendes et al. 2016) and algae (Gilbert-López et al. 2016; Rodriguez et al. 2011; Yuan and Macquarrie 2015). MAE advantages include improved extraction rate, lighter use of solvents, and improved extraction yield. Compared to ultrasounds, MAE is an energyassisted extraction method, uses lower amounts of solvents, and improves the extraction yields of some intracellular compounds. However, the heat generated during the extraction process may damage the heat-sensitive compounds (Michalak and Chojnacka 2014) and degrade the polysaccharide preparations (Ebringerová and Hromádková 2009). 13.2.1.2.4  Enzyme-assisted extraction (EAE)  Marine organisms’ cell walls and cuticles are made up of chemically complex and heterogeneous biomolecules, that is, sulfated and branched polysaccharides associated with proteins and various bound ions, including calcium and potassium (Kadam et al. 2013). In order to extract polysaccharides from the biomass, application of a well-defined mixture of enzyme is, in some cases, necessary for extraction. Enzyme-assisted extraction of polysaccharides is performed under moderate conditions that protect the biologically active compounds from their degradation (Hahn et al. 2012). Protocols for EAE emphasize the importance of maintaining optimum treatment time, pH, and temperature conditions for enzymes to maximize extraction yields (Wijesinghe et al. 2012). Optimum conditions of pH ranging from 3.8 to 8 and temperature ranging from 40°C to 60°C must be maintained for enzymes such as amyloglucosidase (AMG), agarase, alcalase, carrageenanase, cellulase, cellulose protamex, kojizyme, neutrase, termamyl, ultraflo, umamizyme, xylanase, and viscozyme, which are most commonly used in EAE (Jeon et al. 2011). Enzyme-assisted extraction is ecofriendly and nontoxic as it obviates the use of solvents in the process. The technology can also be applied for large-scale operations (Michalak and Chojnacka 2014). However, the usage of enzymes is limited because of their high price in industrial applications (Hahn et al. 2012; Michalak and Chojnacka 2014). This innovative technology relies on the application of one or several enzymes that interact with its substrates under particular conditions defined by each enzyme, which facilitate the extraction of particular compounds by degrading the cell wall. 13.2.1.2.5  Supercritical fluid extraction (SFE)  In supercritical fluid extraction, solvents at temperatures and pressures above their critical points are used (Ibañez et al. 2012). In this state, the density of the fluid is similar to that of a liquid. The viscosity of the supercritical fluid is similar to that of a gas, and its diffusivity is intermediate between those of a liquid and a gas (Ibañez et al. 2012). Thus, because of their low viscosity and high diffusivity, supercritical fluids possess better transport properties than liquid. An important characteristic of SFE is that fluid density can be altered by changing temperature and pressure of the fluid (Kadam et al. 2013); this means that the dissolving power of a fluid can be altered by temperature and pressure changes, as it is dependent on the density of fluid (Herrero et al. 2010). The effects of the SFE process depend largely on a properly prepared raw material. Dried and ground marine biomass is subjected to a pressure between 100 and 1000 bar generally at a temperature of 20°C–45°C. The process should not be carried out at very high temperatures, which can result in the loss of biological activity of extracted compounds. SFE allows the operator to obtain a high yield in a relatively short time. Carbon dioxide (CO2) is frequently used as a solvent, as it is considered to be a “green,” nontoxic, and noncorrosive solvent, easily separated from the extract (Pan et al. 2012). Moreover, CO2 is cheap, available, inert to the product, nonflammable, and shows great

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affinity to volatile compounds (Ibañez et al. 2012; Halim et al. 2012). Additionally, CO2 can be easily and completely removed from the extract (leave no harmful residues) when compared to agents used in other conventional extraction techniques (Crampon et al. 2011). The main drawback of supercritical CO2 is its low polarity (it will not extract any polar substance). This problem can be solved by the addition of polar modifiers or cosolvents that change the polarity of the supercritical fluid and increase its solvating power toward the extracted compound (Ibañez et al. 2012; Crampon et al. 2011). Because of recent advances in SFE technology, increased applications in research and industry have been reported. SFE is a suitable technology for extraction of nutraceuticals. Bioactive compounds can also be extracted without any loss of volatility. SFE offers a rapid extraction rate and high yield and is an ecofriendly technology with minimal or no use of organic solvents. 13.2.1.2.6  Ionic liquids (ILs)  Ionic liquids are low-melting-point organic salts, thus forming liquids that consist only of cations and anions designed to serve as liquids at room temperature or even at temperatures lower than a boiling point of water (Kadokawa 2011). Interest in interleukins (ILs) has been extended to consider their potential use as solvents for biopolymers such as naturally occurring polysaccharides, for which ILs have specific good affinities (Seoud et al. 2007). The processing of cellulose and other polysaccharides in ILs is a step toward feasible and economic energy conversion systems such as biofuel cells. Three steps are involved: (1) the extraction of polysaccharides from marine biomass; (2) hydrolysis of polysaccharides into mono-, di-, or oligosaccharides; (3) conversion of chemical energy, involving the resulting sugars, into electric energy. Biofuel cells based on ordinary molecular liquids, including water, have drawbacks of solvents volatility, short lifetime of enzymes, and even algae growth (Abe et al. 2010). Although ILs overcomes these problems, with low temperature and short time extraction, they present others such as high viscosity (Abe et al. 2010).

13.2.2  Purification of marine polysaccharides After the extraction process, polysaccharides are dissolved in a rich mixture of small amounts of proteins and phenolic compounds that could also have several beneficial biological activities (Ale et al. 2011b). Purification of bioactive marine polysaccharides from crude extracts is of great importance to obtain various homogeneous active polysaccharides fractions and still one of the major obstacles of polysaccharide research and development (Shi. 2016). There are many methods and approaches for isolation, separation, and purification of polysaccharides, such as ethanol precipitation, fractional precipitation, ion-exchange chromatography (IEC), size-exclusion chromatography (SEC), affinity chromatography, and ultrafiltration method by taking advantages of particular properties of the desired compound such as acidity, polarity, and molecular size (Jin et al. 2013).

13.2.2.1  Ion-exchange chromatography (IEC) Column chromatography is currently the most widely used method for marine polysaccharide purification due to its good purification effect and simple operation. Ion exchange chromatography is based on ionic (or electrostatic) interactions between ionic and polar analytes, ions present in the eluent and ionic functional groups fixed to the chromatographic support. Two distinct mechanisms play a role in the separation in ion chromatography as follows: (1) ion exchange due to competitive ionic binding (attraction) and (2) ion exclusion due to repulsion between similarly charged analyte ions and the ions

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fixed on the chromatographic support. The separation is based on the formation of ionic bonds between the charged groups of biomolecules and an ion-exchange gel/support carrying the opposite charge (Acikara 2013). Positively charged ion-exchange resin with affinity for molecules with net negative surface charges [anion-exchange chromatography (AEC)] are commonly used in sulfated polysaccharide purification. Fucoidans exhibit high anionic charges, even at low pH values, due to the sulfate ester groups linked to the carbohydrate backbone. Different fucoidan fractions could be further separated by AEC based on the degree of sulfates groups. In general, highly sulfated fucoidan fractions showed stronger interactions with the resins, which require higher salt concentrations to elute these compounds. Thus, AEC could be used to purify different fucoidan fractions from seaweeds that exhibit distinct structural and chemical properties (Hahn et al. 2012). The reasons for the success of ion exchange are its relatively low average cost, high resolving power, high capacity, large sample handling capacity, and automation. Consequently, ion-exchange chromatography is one of the most widely used liquid chromatography technique for purification, and it continues to expand with development of new technologies (Acikara 2013).

13.2.2.2  Gel permeation chromatography (GPC) Another routinely used technique in the purification of marine polysaccharides is gel permeation chromatography (GPC), where polysaccharide molecules pass through a gel filtration medium packed in a column at rates varying with their size and shape of; thus GPC operates according to the principle of a molecular sieve. A key difference from ion-exchange chromatography or affinity chromatography is that in GPC, molecules do not bind to the medium (Spurr 2014). The commonly used gels are various types of Sephacryl, Superdex, and Superose (Shi 2016). The application of gel permeation chromatography following anion-exchange chromatography is typical for the purification of a highly valuable product. Thus, the salts used for the elution of the target compound from the anion-exchange resin are removed. Dialysis and ethanol precipitation are also used for the desalting and removal of low-molecular-weight compounds. This technique allows for fractionation of the fucoidans with respect to their molecular weights (Hahn et al. 2012). Neither gel permeation chromatography nor anion-exchange chromatography influence the sulfate ester groups of the polysaccharide; however, the strong changes in ionic strength may affect the spatial pattern of the target compounds. These changes in molecular geometry may also affect the bioactivity of the resulting sulfated polysaccharide (Hahn et al. 2012).

13.2.2.3  Size-exclusion chromatography (SEC) Size-exclusion chromatography is used to separate the molecules according to their size as they pass through a porous matrix of particles with chemical and physical stability and inertness. This method overcomes some of the problems associated with the traditional methods, namely, the disposal of solvents and the lengthy and complex procedures (Zhang and Row 2014). Subsequently, a significant advantage of SEC is that it can be used directly following IEC or other purification techniques using the buffer that best suits the type of sample in terms of preservation or further purification (Mori and Barth 2013). The application of SEC after IEC becomes a common procedure in the purification of polysaccharide. SEC is a relatively easy and commonly used solution for molecular weight characterization. For SEC, the molecular weight range of the column should cover the molecular

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weight distributions of the chosen polysaccharides. Other factors, such as high resolution, low adsorption, and stability over a wide pH range are also important (Zhang and Row 2014). The high-performance size-exclusion chromatography method is a suitable procedure for separating soluble polysaccharides in brown seaweeds on the basis of molecular weight (Zhang and Row 2014).

13.2.2.4  Affinity chromatography Affinity chromatography is one of the most diverse and powerful chromatographic methods for purification of a specific molecule or a group of molecules from a complex mixture. This method relis on a highly specific biological reversible interaction between the molecule and a specific ligand based on their biological function or individual chemical structure for purification process (Urh et al. 2009). The ligand is covalently attached to a matrix, which must be chemically inert, and porous, and must possess a variety of functional groups suitable for coupling with diverse ligands (Pohleven et al. 2012). The process is described simply as follows. First, an affinity column should be prepared in advance. The affinity column is eluted using the polysaccharide solution as mobile phase. The polysaccharide solution is a mixture of polysaccharide fractions. During the elution process, only the polysaccharide fraction that can bind with the ligand would be adsorbed to the column, and other polysaccharide fractions that cannot bind with the ligand would flow out of the column. Then the ionic strength and pH value of the mobile phase can be properly changed to dissociate the polysaccharide fraction combined with the ligand. In the case of polysaccharides, fucose-binding lectins were used effectively to purify fucoidan from crude solutions; however, problems could arise if the sulfate content of the polysaccharide blocks the fucose units that interact with the lectins (Hahn et al. 2016). Strong interactions have been described between the anionic sulfate esters of the sulfated polysaccharides and different cationic dyes (i.e., toluidine and methylene blue), forming a strong donor–acceptor complex (Hahn et al. 2016). A new dye affinity chromatography method has been developed using modified amino-derivatized sepabeads with toluidine blue. This method is showing promising results in the purification of fucoidan from brown seaweeds with higher purity levels than the commercially available purified (95%) fucoidan from Sigma-Aldrich (Hahn et al. 2016). Affinity chromatography is a high-efficiency, user-friendly, and unique method in purification technology since it is the only technique that enables the purification of a biomolecule on the basis of its biological function or individual chemical structure. However, the disadvantage is that it is difficult to find a proper ligand for a given polysaccharide molecule. Therefore, affinity chromatography is rarely applied in polysaccharide purification.

13.2.2.5  Ultrafiltration method Since the marine polysaccharide molecules have different sizes and shapes in a solution, they can be separated by passing them through the ultrafiltration membrane under pressure because this membrane can only allow a certain MW range of polysaccharides. In fact, the principle of the ultrafiltration method is also a molecular sieve. Nevertheless, practically, most ultrafiltration membranes can adsorb polysaccharides, which lead to the large decrease of polysaccharide yield. For example, a hollow-fiber ultrafiltration membrane can greatly adsorb polysaccharides. In addition, the ultrafiltration speed is much lower and the process is too time-consuming because many polysaccharide solutions are very viscous (Shi 2016).

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13.3 Marine polysaccharides: structures, function, and biological activities studies 13.3.1  Biological activities of marine polysaccharides Marine polysaccharides have promising prospects and a very important potential that could contribute to the development of a marine bioindustry. Specifically, polysaccharides obtained from microalgae and seaweeds have attracted a great deal of attention in the last few years, in view of their diversity and biological activities. These biomolecules have a wide variety of industrial applications including health, nutraceuticals, environment, and agriculture, due to their broad range of biological activities. In the field of health, Sulfated polysaccharides from algae have several pharmacological activities such as anticoagulant, antioxidant, antiproliferative, antitumoral, anticomplementary, anti-inflammatory, antiviral, antiseptic and antiadhesive activities (Cumashi et al. 2007). Anticoagulant activity is among the most widely studied properties of sulfated polysaccharides (Liu et al. 2018), especially those with low molecular weight, and these sulfated polysaccharides have been described to possess anticoagulant activity similar to that of heparin (de Souza et al. 2007). Sulfated polysaccharides from Catathelasma ventricosum are known to possess excellent antioxidant, anti-inflammation, and anticoagulant activities (Khemakhem et al. 2018; Liu et al. 2018). The polysaccharides recovered from the microalgae Navicula sp. presented antioxidant activity, which could be related to the molecular structural characteristics such as molecular weight and sulfate content. Analysis of these polysaccharides using gas chromatography showed that they contain glucose, galactose, rhamnose, xylose, and mannose as main neutral sugars and with higher content of sulfate content (Fimbres-Olivarria et al. 2018). The polysaccharides also showed a very significant anticancer effect. Exopolysaccharides from marine bacterium Bacillus thuringiensis inhibited the growth of cancer cells in a dosedependent manner, and maximum anticancer activity was found to be 76% against liver cancer. At 1000 μg/mL these polysaccharides showed potent antioxidant properties, which were investigated against DPPH, hydroxyl, and superoxide free radicals (Ramamoorthy et al. 2018). In the same sense, Sulfated polysaccharides isolated from Cerastoderma edule were shown to have antiproliferative activity on chronic myelogenous leukemia and relapsed acute lymphoblastic leukemia cell lines. Analysis of these marine sulfated polysaccharides confirmed the presence of glycosaminoglycan-like structures that were enriched in ion-exchange purified fractions containing antiproliferative activity. The antiproliferative activity of these glycosaminoglycan-like marine polysaccharides was shown to be susceptible to heparinase (Aldairi et al. 2018). In recent years, polysaccharides of microalgae have attracted the attention of researchers for their biological activities. Polysaccharides from Dictyota dichotoma microalgae that have high sulfate, glucose, fructose, manose, galactose, and 1,3- and 1,6 -β-d-glucan levels have shown a potent anticancer effect against colon cancer cells HCT-116, HT-29, and DLD-1 (Usoltseva et al. 2018). Many extracellular microalgal polysaccharides (EPSs) have been reported in recent decades, and their composition, structure, biosynthesis, and functional properties were documented. Several microalgae such as Chlamydomonas reinhardtii, Botryococcus braunii, Dunaliella tertiolecta, Porphyridium cruentum, Spirulina sp., Isochrysis galbana, and Dunaliella salina, can be a source of exopolysaccharides. In the agricultural field, microalgae exopolysaccharides can be a new sustainable source of plant biostimulants for crop growth improvement and protection against biotic and abiotic stress, including salt stress. Salt stress is one of the major abiotic

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stress factors that adversely affect the physiological and biochemical mechanisms of plants and cause severe losses in plant productivity worldwide (Aman et al. 2017). Salt tolerance in plants depends mainly on the capacity of roots to restrict or control the uptake of sodium (Na+) and chloride (Cl−) ions, and to uptake essential elements, particularly K+ and NO3− (Jeschke et al. 1988). A study by Ashraf (2004) showed that the use of bacterial exopolysaccharides extracts attenuates the effect of salt stress on germination and growth of plant wheat by regulating the assimilation of Na+ and K+ ions. Another study showed that exopolysaccharides extracted from Phaeodactylum tricornutum and D. salina stimulated the germination of pepper under salt stress conditions (Guzmán-Murillo et al. 2013). The structural features of polysaccharides, including their molecular size, frequency of branching, monosaccharide composition, conformation, types and sequence of linkages, determine their physical and physiological properties (Ferrierra et al. 2015; Jin et al. 2012). Uronic acids are a class of sugar acids with both carbonyl and carboxylic acid functional groups. A subclass of these oligosaccharides, called oligosaccharins, possess “hormone-like” activity and may contribute to the natural control of plant metabolism, growth, and development. For example, oligosaccharides, generated in vitro by hydrolysis of pectins –which is rich in galacturonic acid-, promote ripening of tomato (Lycopersicon esculentum L) and other fruits (Dumville et al. 2000). El Arroussi et al. (2018) showed that the D. salina exopolysaccharides significantly improve length and weight on both the shoot and root system of tomato plants (Figure 13.1). In the same sense, Mzibra et al. (2018) showed that the polysaccharides obtained from Moroccan seaweeds induced an increase in plant biomass and in chlorophyll levels. The biological activity of polysaccharides is related to their structure, sulfate content, uronic acid, neutral sugars, and other functions (Ribera et al. 2017). Hence, the positive effect of polysaccharides on plant growth may possibly be due to the presence of sugars such as glucose, galactose, and maltose. Castellanos-Barriga et al. (2017) showed that the Ulvan

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from Ulva lactuca stimulated seedling growth and chlorophyll content in mung bean (Vigna radiata). Similarly, Alghamdi (2017) showed that the application of Ascophyllum nodosum oligosaccharides have a biostimulant effect on plant growth and induced resistance against biotic and abiotic stresses. Polysaccharides from U. lactuca and P. gymnospora have also a stimulatory effect on tomato and mung bean plant physiology (HernándezHerrera et al. 2016). Carbohydrates can be recognized in the plant cell wall and act as signaling molecules, inducing production of phytochemical compounds that can be active in root induction and growth processes (Klarzynski et al. 2003; Chandía et al. 2004; Pardee et al. 2004). Furthermore, sugars are known to improve plant growth in a similar way to hormones (Rolland et al. 2002). Several modes of application have been studied to evaluate the effect of polysaccharides on plants. To investigate the effect on germination and growth, polysaccharides were used as a seed coating or seed priming. Foliar application is widely described as a mode of use as a biostimulant of growth and development (Ertani et al. 2018, Khan et al. 2009) and elicitor of the natural plant defense against abiotic and biotic stresses (Mzibra et al. 2018; Vera et al. 2011). Soil application is the most used mode for stimulation of the plant root development, stimulation of the soil microbial population, and stimulation of the absorption and assimilation of nutrients.

13.3.2 Linking chemical composition and structure to biological activities of marine polysaccharide The overall research on marine polysaccharides and their biotechnological applications emphasizes the diversity and the richness of their sources. Such diversity contributes to the tremendous variability in the structure of marine polysaccharides and their bioactivities. According to Singh et al. (2017), the collective study of the physical, chemical and biological properties of marine polysaccharides is called characterization. This definition highlights the complexity of the structural elucidation of polysaccharides from marine sources and the assessment of their biological activities. It underlines the importance of multidisciplinary approaches in order to improve industrial applications of marine polysaccharides. The research on marine polysaccharides is one of the most dynamic biological research areas in the last few decades. Publications related to marine polysaccharides have increased more than threefolds between 1994 and 2017, and since 2013 more than hundred papers are published yearly. (Source: Science Direct Database, research done in May 2018 using “marine polysaccharides” as keywords.) Marine polysaccharides derived mainly from seaweeds, which are probably considered as the most abundant organic molecules in the oceans. They have a great molecular diversity which is not yet well understood and analogs of some marine polysaccharides, like fucoidans, have not yet been found in land sources (Ermakova et al. 2015). It was demonstrated that the biological activities of marine polysaccharides are strongly related to their chemical composition, polymer structure, and environmental conditions (Mzibra et al. 2018; Xu et al. 2017). However, it was noted that little information is available about the chemical structure of marine polysaccharides and their biological activities (Praiboon et al. 2017). Bioactivity and biocompatibility of some marine polysaccharides, and their derivatives is related to several phenomena such as structural features, distribution of some functions in the polymer chain, porosity, charge, crystallinity, type of existing hazards, and products of their degradability (Prabaharan 2015).

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13.3.2.1 Chitosan Chitosan and its derivatives such as chitooligosaccharides and low-molecular-weight chitosan (LMWC) were described for many biological activities. Zheng et al. (2016) demonstrated that the immunostimulative activity of chitosan depends of its molecular weight; they suggested the molecular mechanism responsible for regulating immune response by LMWCs. Molecular weight of chitosan might affect its structure and its physicochemical properties, which enhance or reduce its biological activities. Chitooligosaccharides (COSs) are produced by partial hydrolysis of chitosan. Two COS mixtures (1.2 and 5.3 kDa) were investigated for their anti-inflammatory activity in vivo using the Edema induced by phlogistic agents (Fernandes et al. 2010). Results of this study showed that both concentration and molecular weight of COS impact the intensity of mice paw edema volume decrease. The COS 1.2 kDa administrated at 500 mg/kg showed the best results (Fernandes et al. 2010). COSs are the products of chitosan degradation and are produced by several methods, but mainly enzymatic hydrolysis (Trincone 2017; Aung et al. 2017; Aljawish et al. 2015). Hence, the production methods yield different chitosan derivatives with particular structures and physicochemical properties that are strongly correlated with biological activities (Xia et al. 2010). Novel bioactivities may result from the improvement of efficient synthetic routes and the selective modification of these biopolymers that is paving the way for improved properties and tailored activities (Carvalho et al. 2016). The effects of structural modifications of chitosan on antimicrobial activity and toxicity reviewed an update of the current state-of-the-art by Sahariah and Masson (2017). Factors affecting the antimicrobial activity and the toxicological profile of chitosan may include the structure, chemical properties, degree of substitution for substituents attached to the polymer backbone, molecular weight, and degree of acetylation. Figure 13.2 illustrates the effects of such factors on the Structure-activity relationship R1

Effect of degree of substitution O

R2

-----R2 = --NH

n

Effect of spacer length

O ---NH X Spacer = 2 ----X Spacer = 0

where N

(5p)

H N

NH2

NH (5r)

N H

N H

-----R1 = O

NH2HCl

O

-----R1 =

-----R2 =

DS = 0.69

-----R2 =

H N

NH2

-----R2 =

NH DS = 0.11-1.0 (5k) -----R2 =

H N

N H

N

HO DS = 0.22-0.42

(5g)

N

O DS = 0.13-1.0 (5n)

O

-----R2 =

O

and

N

-----X = N

(5p, 5s)

Increasing degree of substitution

Decreasing antibacterial activity

Increasing spacer length

---NH Spacer = 6

NH NH

-----R2 = O HN

DS = 0.14-0.39 (5b) OH O -----R2 = NH NH N N H H NH2HCl N N H DS = 0.16-0.44 H DS = 0.10-0.71 (5d) (5e)

-----R2 =

where -----R2 =

5 X

OH

O DS = 0.10-0.53 (5a)

Chitosan derivative

O

3

H N

NH N H

NH2

O DS = 0.07-0.48 (5c) -----R1 = -----R2 = OH and ---NH

---O O DS = 0-0.85

-----R2 =

H N

(5f)

-----R2 = N

NH

N OH DS = 0.57-0.76

N HO DS = 0.06-0.44 (5j)

3N NH2 NH2 H DS = 0.06-0.30 (5h, 5i)

O DS = 0.14-1.0 (5l) -----R2 =

OH

O

NH 5N H

NH2

O DS = 0.12-1.0 (5m)

Increasing antibacterial activity

O

O HO

H N

5N O DS = 0.13-1.0 (5o)

Figure 13.2  Effect of degree of substitution and the positioning of cationic charge on antimicrobial activity of chitosan and derivatives [reproduced with permission from Sahariah and Masson (2017)].

Chapter thirteen:  Marine polysaccharides as promising source of biological activities

Algal Species, Location, Season, Maturity stage

Seaweeds

Green

Brown

313

Red Extraction Method: Duration, sample volume, concentration of extraction solution...

Carrageenan Kappa, iota & lamda type

Ulvan

Monomeric constituents, molecular size, sulfates groupment and degree of branching

Fucoidan

Figure 13.3 Sulfated polysaccharides obtained from marine algae and sources of variability [Adapted from Patel (2012)].

biological activities of chitosan and derivatives (Sahariah and Masson 2017). However, it was concluded that more studies considering more than one factor at the same time and their synergistic effects are needed.

13.3.2.2  Sulfated polysaccharides from marine algae Algae sulfated polysaccharides have attracted great attention in recent years. Several relevant research reports are published each year and are devoted to extraction methods of algae polysaccharide, structural diversity elucidation, biological activities, and importance in various fields (Jiao et al. 2011; Patel 2012; Cunha and Grenha 2016; GarciaVaquero et al. 2017). The main sulfated polysaccharide extracted from marine seaweeds are ulvans from green algae, fucans from brown algae and carrageenan from red algae (Figure 13.3). According to Figure 13.3, the main sources of variability that impact the biological activities of sulfated polysaccharides are related to the algal species, location and season of sampling, the maturity stage, and the extraction and purification methods. Furthermore, the structural modifications of sulfated polysaccharides may modify the structural and biological properties (Gurpilhares et al. 2016). Such variability influences the monomeric constituents, molecular size, sulfate groups, and degree of branching. In fact, the chemical structure of sulfated polysaccharide is the driving force for their described biological activities. For example, fucoidan extracted from different algal species can vary in their chemical structure (Jiao et al. 2011). The seasonal variation of sulfated polysaccharide is another parameter that is poorly considered in the literature. Recently, Fletcher et al. (2017) investigated the seasonal variation of fucoidan extracted monthly from three different species of brown seaweeds in the UK over one year. The authors suggested that the extracted fucoidan from the three species contains a higher sulfate rate in the winter compared to the summer and hence its functionality varies over the year. This study can be considered as the beginning of an understanding of the seasonal variation of sulfated polysaccharide extracted from

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seaweeds. Further studies are required to understand the complexity of variation of sulfated polysaccharide extracted from different seasons and locations and from different maturity stages of the algal species. The extraction and purification methods of sulfated polysaccharides from marine algae need to maintain their structural integrity. Several conventional and modern extraction methods are available and were reviewed earlier in this chapter. The impact of the extraction technologies on key structural components of polysaccharides needs to be investigated in order to ensure a more informed selection of the extraction method with a possible upscale to the industrial level. Xu et al. (2017) analyzed and summarized relationships between structure and biological activities of polysaccharide extracted from marine algae. Even if more than 25 works were analyzed in this recent review, we are still lacking an investigation of the influence of the extraction methods on the structural properties and activities of polysaccharides from marine algae. The extraction conditions (lysis time, sample volume, concentration of extraction solution, probe size, amplitude, etc.) are diverse and may influence the structural properties of the extracted polysaccharide. Ale et al. (2011a) recommended the development of standard extraction procedures that will be able to generate a better common basis for analysis and understanding of bioactivities. However, the extraction conditions may not be optimal in all cases depending on seasonal change, location, and sources, even if the same species is of interest (Okolie 2018). Hence, the optimization of these parameters is of great interest and should allow researchers to obtain the best fit for the seaweeds of interest.

Conclusion Biological diversity of marine organisms is continually surprising scientific communities because of its vast diversity and the complexity of marine habitats. Such diversity is considered as an untapped source of marine natural products that encompass a wide variety of chemical classes. Marine polysaccharides have a great molecular diversity, which is not yet well understood, and analogs of some marine polysaccharides, like fucoidans, have not yet been found in land sources. They derived mainly from seaweeds and have attracted attention of scientific and economic communities. In this chapter, we tried to identify methods of extraction and purification of these macromolecules while accounting for the influence of these methods on the chemical structure and the biological activities. Thus, we have focused on the relationship between the structure of these molecules and their biological activities before describing the different industrial applications of these bioactive macromolecules while focusing on new industrial applications. Several methods of extraction and purification were described in this chapter. Advantages and disadvantages of each method and their effects on the yield and quality, as well as the biological activities, of these macromolecules are discussed. The review presented in this chapter showed that the structure as well as the activity of these macromolecules can be influenced by the extraction and purification conditions. Marine biomass pretreatments are necessary to improve the extraction yield and/ or remove maximum impurities that may affect the activity of these molecules. According to this bibliographic analysis, it appears that, besides the known applications of marine polysaccharides, such as therapeutic, cosmetic, and food applications, other applications have drawn the attention of researchers in the last several years. Novel applications such as plant biostimulant and elicitors are emerging. In fact, these molecules have shown a good track of industrial valorization as plant biostimulants by promoting nutrients and

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water uptake and by stimulating root development and the photosynthetic capacity of plants. They also act as elicitors of the natural defense of plants against biotic and abiotic aggression. The technical and scientific challenges for a good industrialization of these marine polysaccharides are the choice of extraction and purification methods with regard to the application of these macromolecules. This choice should be based on the biological sources of these polysaccharides, the sensitivity of the structure of these molecules, as well as the peculiarity and complexity of the desired application. Economically and technically reliable industrial processes can broaden the spectrum of application of these bioactive molecules. Moreover, in-depth studies of their chemical structures and behavior in the environmental conditions of extraction and purification are needed. Such studies need to be multidisciplinary because of the complexity of marine polysaccharides and the number of variability sources that are mentioned in this chapter. Thus, studies of the mechanisms of action of this bioactivity in the different applications will allow the control of the structure–­f unction relationship of these molecules, and the technical precautions to be taken during the extraction and purification of these polysaccharides.

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chapter fourteen

Microwave-assisted conversion of marine polysaccharides Shuntaro Tsubaki1, Ayumu Onda2 , Tadaharu Ueda3, Masanori Hiraoka4 , Satoshi Fujii1,5, and Yuji Wada1

Department of Chemical Science and Engineering, School of Materials and Chemical Technology, Tokyo Institute of Technology, Ookayama, Meguro, Tokyo, Japan 2Research Laboratory of Hydrothermal Chemistry, Faculty of Science, Kochi University, Kochi, Japan 3Department of Marine Resource Science, Faculty of Agriculture and Marine Science, Kochi University, Kochi, Japan 4Usa Marine Biological Institute, Kochi University, Inoshiri, Usa, Tosa, Kochi, Japan 5Department of Information and Communication Systems Engineering, Okinawa National College of Technology, Okinawa, Japan 1

Contents 14.1 Introduction....................................................................................................................... 321 14.2 MW heating as a fractionation of biomass components............................................. 323 14.2.1 How microwaves heat materials....................................................................... 323 14.2.2 MW heating mechanisms and dielectric property of materials.................. 323 14.2.3 MW apparatus..................................................................................................... 324 14.3 Extraction of marine polysaccharides from seaweed biomass by MWs................... 326 14.4 Hydrolysis of marine polysaccharides under MWs.................................................... 327 14.5 Dielectric properties of marine polysaccharides in water.......................................... 329 Conclusion.................................................................................................................................... 331 References...................................................................................................................................... 331

14.1 Introduction A wide variety of marine polysaccharides are expected as high-value-added chemicals to use it for rheological materials as well as biologically active materials (Kerton, Liu, Omari, et al. 2013). Polysaccharides in seaweeds can be classified as either structural polysaccharides, matrix polysaccharides, or storage polysaccharides (Wei, Quaterman, and Jin 2013). In general, structural polysaccharides contribute to mechanical strength of the algal body, while matrix polysaccharides contribute to viscoelasticity. Structural and storage polysaccharides (cellulose, starch, and laminalan) can be employed in bioenergy applications (Figure 14.1). In contrast, matrix polysaccharides, which are characterized as having a unique chemical variation of marine polysaccharides in seaweeds, are available for food and medical applications (Figure 14.1). Chemical properties of marine polysaccharides vary depending on the species of seaweeds. Brown algae accumulate alginate and fucoidan; Red algae accumulate agar and carrageenan; green algae accumulate ulvan and rhamnan sulfate. 321

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Starch HO

Extraction

HO HO OH O

OH O

Cellulose

OH O

OH O HO

10~15%

O OH O HO

HO O

10~20%

OH O

OH

O O HO

HO

OH O OH

HO O

OH

O O HO

OH O

HO

OH OH

n

OH O

Bioenergy application

Phycocolloids Extraction Fast-growing Ulva spp.

Rare sugars

30~50%

Ulvan

Rhamnose OH

+

Na–OOC O HO

O OH

H C 3 O HO

Hydrolysis

O OH

H C 3 HO HO

O OH

Food and medical application

Figure 14.1  Schematic illustration of fractionation and hydrolysis of polysaccharides from seaweed biomass. [The figure was reproduced with permission from John Wiley & Sons Ltd. (Tsubaki, Zhu, and Hiraoka 2017).]

Alginate, carrageenan, and agar are widely used for food additives by exploiting their viscoelastic property. Sulfated polysaccharides (fucoidan, ulvan, and rhamnan sulfate) exhibit various biological properties such as anticoagulant, antithrombotic, antivirus, antitumor, immunomodulatory, and anti-inflammatory effects (Smit 2004; Lahaye and Robic 2007; Li, Lu, Wei, et al. 2008; Alves, Sousa, and Reis 2013). The extracted polysaccharides can be further depolymerized to produce oligomeric and monomeric sugars, which should also be biological active chemicals (Jiao, Yu, Zhang, et al. Ewart 2011). The green separation process of chemical components with less energy consumption is an important key step in the utilization of seaweed biomass. Several extraction processes based on physical and physicochemical treatments such as supercritical and subcritical fluid extraction, ultrasonic extraction, and microwave-assisted extraction have been used to improve extraction efficiency (Flórez, Conde, and Domínguez 2015). For instance, ultrasonication increases mass transfer of chemicals from plant bodies by breaking cell walls by rupture of bubbles generated by ultrasonic. Supercritical fluids with very low viscosity and low surface tension penetrate into plant bodies to enhance mass transfer of useful extracts trapped in the plant body. In addition, subcritical water with a high dissociation coefficient (Kw = [H+][OH−]) initiates autohydrolysis of plant cell wall to enhance extraction of cell wall polysaccharides (Peterson, Vogel, Lachance, et al. 2008; Savage 2009). Microwaves generate heat from within due to dielectric loss to enable rapid and selective heating of materials, which generates nonequilibrium thermal distribution in completely different manner from conventional heating. Since MW heating can contribute to low environmental impact, due to decrease in extraction time and energy consumption, MW-assisted extraction is regarded as a green extraction technology. The number of publications on this topic has been increasing recently. Therefore, MWs have been used to initiate fractionation of chemical components from not only marine biomass but also terrestrial biomass (woody biomass and herbaceous biomass) (Azuma, Tanaka, and Koshijima 1984; Tsubaki and Azuma 2011). The MW distillation is one of the most practical MW-assisted extraction processes

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(Chan, Yusoff, Ngoh, and Kung 2011; Kokolakis and Golfinopoulos 2013; Li, Fabiano-Tixier, Vian and Chemat 2013; Chemat and Cravotto 2013), which have been already in practical use. The MW distillation facilitates extraction of volatile compounds such as aroma chemicals by directly heating water trapped inside the plant body without addition of any water, while conventional hydrodistillation require a large amount of energy to generate steam (Flórez, Conde, and Domínguez, 2015). Therefore, the duration of extraction as well as energy consumption can be reduced by using MW heating. MWs are also used for solid – liquid extraction of oils and pigments from microalgae biomass (Reddy, Muppaneni, Sun, et al. 2014) and polysaccharides from various kinds of biomass including seaweeds (Rodriguez-Jassp, Mussato, Pastrana, et al. 2011; Cheong, Wang, Wu, et al. 2016; Kumar, Sivakumar, and Ruckmani 2016; Smiderle, Morales, Gil-Ramírez, de Jesus, et al. 2017). In this chapter, the fundamentals of MW heating mechanism and MW apparatus is illustrated in the first section. Then, practical applications of MWs are summarized on extraction and hydrolysis of marine polysaccharides from seaweed biomass. Finally, the key dielectric properties of marine polysaccharides are discussed to understand the heating mechanism of marine polysaccharides in water.

14.2  MW heating as a fractionation of biomass components 14.2.1  How microwaves heat materials Microwaves are electromagnetic waves in a frequency range between 300 MHz and 300 GHz, which corresponds to the wavelength range between 1 mm and 1 m in wavelength. MWs are used mostly for telecommunications and radars. Selected MW frequencies (915 MHz, 2.45 GHz, 5.8 GHz, 24.125 GHz) and radiofrequencies (13.56 MHz, 27.12 MHz, 40.68 MHz) can be used for dielectric heating materials as industrial, scientific, and medical (ISM) bands. MW heating at 2.45 GHz has been widely used for heating devices for cooking. Lower MW frequencies at 915 MHz and radiofrequency have been used for food pasteurization and sterilization (Piyasena, Dussault, Koutchma, et al. 2003) and timber drying (Elustondo, Avramidis, and Shida 2004).

14.2.2  MW heating mechanisms and dielectric property of materials Microwave heating is strongly dependent on dielectric property of materials as well as intensity of electric and magnetic field. Equation (14.1) describes the MW power converted to heat



P=

1 2 2 2 σ E + pfε 0 ε r′′ E + p ′fµ 0 µ r′′ H 2

where P = energy loss at unit volume E = electric field intensity H = magnetic field intensity σ = electric conductivity f = frequency ε0 = permittivity of free space εr ′′ = dielectric loss μ0 = permeability of free space μr ′′ = magnetic loss p, p′ = coefficients

(14.1)

324

Enzymatic Technologies for Marine Polysaccharides Liquids

Electrolytes

Solid catalysts

Ionic conduction

Joule heating Magnetic loss

δ– δ+ Dipole rotation

Figure 14.2  Schematic illustration of MW heating mechanisms.

Dielectric loss is related to orientation and relaxation of dipolar molecules such as water under an oscillating electric field (Figure 14.2) (Gabriel, Gabriel, Grant, et al. 1998). Water exhibits highest dielectric loss at approximately 18 GHz at 20°C and gradually decreases at lower frequencies. Electric field also affects electrolytes, due to an ionic conduction mechanism (Figure 14.2). For instance, saline water exhibits higher MW absorption than does pure water (Tanaka and Sato 2008). Ionic conduction mechanism becomes prominent at lower MW frequencies (i.e., 20 GHz), corresponding to the cooperative relaxation of long-range H-bond-mediated dipole–dipole interactions of free water (Fukasawa, Sato, Watanabe, et al. 2005). On the other hand, both sodium alginates and κ-carrageenan gave prominent dielectric loss at lower frequencies (